Thank you to Dr. Amy Cash-Ahmed, Nathanael J. Beach, and Alison L. Sankey for their assistance in carrying out this work. Mention of trade names or commercial products in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the U.S. Department of Agriculture. USDA is an equal opportunity provider and employer. This research was supported by a grant from the Defense Advanced Research Projects Agency # HR0011-16-2-0019 to Gene E. Robinson and Huimin Zhao, USDA project 2030-21000-001-00-D, and the Phenotypic Plasticity Research Experience for Community College Students at the University of Illinois at Urbana Champaign.

1. QMC assembly
<ol>
	<li>Assemble QMCs from parts (Figure 1A) with a single egg laying plate (ELP) inserted as shown in Figure 1B. Do not add feeder tubes until after the workers have been added to the cage. Temporarily cover the 4 feeder holes with laboratory grade tape.</li>
	<li>Insert the queen excluder and the feeding chamber door over the feeding chamber to keep the queen from entering the feeding chamber and contacting the treated diet. See Fine et al.32 for further assembly details.</li>
	<li>Collect the wax comb frames containing the capped worker brood from honeybee colonies 24 h prior to adult eclosion and place them in an incubator (34.5 &#176;C) inside a brood box. 24 h later, brush the eclosed bees off the frames and into an open container that has been lined with an insect barrier paint (e.g., Fluon) to prevent the bees from crawling out.</li>
	<li>Add at least 50 bees by weight (5 g &#8776; 50 bees44,45) to the egg laying chamber of each QMC. To ensure that a diverse genetic pool of workers is represented in the experiment, obtain an approximately equal number of worker bees from at least three colonies and mix them prior to adding them to the QMCs. 
	NOTE: Newly eclosed worker bees less than 1 day old cannot fly or sting due to their underdeveloped flight muscles and unhardened cuticle. If they are added at this age, there is no need to anesthetize them prior to handling. They can be weighed by gently scooping bees from the container using a small &#188; cup volume measuring cup and placing them into a second container (lined with insect barrier paint e.g., Fluon) that has been tared on a scale. The area of the frames covered by capped brood should be roughly equal to ensure that source colonies are equally represented in the QMC worker populations. Homogenization of worker bees can be achieved by brushing newly eclosed bees from frames taken from all colonies into the same container and allowing them to mix for 5 min prior to adding them to QMCs.</li>
	<li>Add the feeders containing sucrose solution, water, and pollen supplement (See section 2).</li>
	<li>Expose the individual mated queens to CO2 gas to stimulate egg laying46 and to ease transfer into QMC.
	<ol>
		<li>Use queens purchased from a commercial breeder within 48 hours of receipt. While the queen is still inside the shipping cage, place it in a clear plastic bag. Place one end of a plastic tube connected to a CO2 gas cannister inside the bag and gently open the cannister valve to allow the CO2 gas to flow.</li>
		<li>When the bag has been inflated with gas, simultaneously close the cannister valve and hold the bag closed to trap the gas inside. Keep the bag closed for 30 s or until the queen has stopped moving. Remove the queen and open the shipment cage once she is observed to be unconscious.</li>
	</ol>
	</li>
	<li>Partially open the door to the egg laying chamber, gently place the unconscious queen inside and close the lid, taking care not to crush the queen or workers inside. Add the second egg laying plate to each QMC as shown in Figure 1C. Place a piece of laboratory tape across the top of the two ELPs to keep them from separating from the QMC frame and prevent workers from exiting the cage.</li>
	<li>Place the cages in a dark incubator with stable environmental conditions of 34 &#177; 0.5 &#176;C and 60% &#177; 10% relative humidity, like the conditions inside a normal colony.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62316/62316fig01.jpg" /> 
Figure 1. A: Disassembled QMC. B: Partially assembled QMC with 1 ELP inserted. C: Fully assembled QMC with 2 ELPs. <a href="https://www.jove.com/files/ftp_upload/62316/62316fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Preparing and administering diets laced with agrochemicals
<ol>
	<li>To prepare 1000 g of 50% (g/g) sucrose solution, place a stir bar in the bottom of a clean 1 L glass reagent bottle. Add 500 g sucrose and 500 mL of deionized water. Unscrew the lid of the bottle and use a heated stir plate set to low heat to mix the solution until all the sucrose has dissolved. Allow the solution to cool to room temperature before adding the agrochemical stock solutions.</li>
	<li>Prepare the stock solutions of agrochemicals in an appropriate solvent, such as acetone, at a concentration that can be added to diet to achieve the desired final concentration of the agrochemical of interest. 
	NOTE: When using acetone as a vehicle solvent, the Organization for Economic Cooperation and Development (OECD) guidelines stipulate that the final concentration of acetone in diet must be &#8804; 5% for chronic oral toxicity tests on adult honey bees47. However, some solvents such as n-methyl-2-pyrrolidone5,31 and dimethyl sulfoxide25 can exert toxic effects below this concentration, so it is recommended to keep solvent concentrations as low as possible in treatment diet. Depending on the volume and type of solvent used, it may be necessary to include both a solvent control group and a negative control group to ensure that potential effects due to solvent toxicity are detected. When using formulated products, the amount of the product used must be adjusted based on the concentration present in the formulation. Depending on the stability of the agrochemical of interest in the solvent, stock solutions can be kept for up to 2 weeks at -20 &#176;C.</li>
	<li>Select sublethal doses based on the results of OECD Test No. 245: Honey Bee (Apis mellifera L.), Chronic Oral Toxicity Test (10-Day Feeding)47, and identify the relevant literature by querying the Ecotox knowledgebase48.</li>
	<li>Administer the agrochemical treatments in a sucrose solution, a commercial pollen supplement (if available as a powder), or both. Prepare the experimental diet for use the same day by adding an appropriate amount of stock solution to chilled/room temperature 50% sucrose solution (w/w). Mix thoroughly by vortexing or with a stir bar set to medium speed. For pollen supplements, add the agrochemical laced sucrose solution to the powdered supplement instead of the syrup according to manufacturer protocols, making sure to adjust the volume of stock solution used according to the final weight of the pollen diet. See Table 1 for example calculations.</li>
	<li>Prepare the feeder tubes from 2 mL microcentrifuge tubes.
	<ol>
		<li>For liquid diet feeders, heat the tip of a 20-gauge needle on a hot plate/stovetop and puncture the bottom of the tube twice. Close the tube lid and pipette approximately 1.5 mL of sucrose solution or water through one of the puncture holes. Set the tube down with the punctured side up until it is added to the QMC.</li>
		<li>For pollen supplement feeders, use a razor blade to slice off the bottom of the tube. Close the lid and a push a 1-2 g ball of pollen supplement into the tube until it touches the lid.</li>
	</ol>
	</li>
	<li>Record the feeder weights prior to placing them in the QMCs. Do not keep unused diet at 4 &#176;C for over 48 hours.</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>Desired Concentration</td>
			<td>Desired Solvent Vehicle Concentration</td>
			<td>Desired final volume/mass of sucrose solution</td>
			<td>Voume of stock solution&#160;</td>
			<td>Imidacloprid in stock solution</td>
			<td>Suggested stock solution recipe</td>
		</tr>
		<tr>
			<td>Sucrose solution</td>
			<td>10 ppb (w/w)</td>
			<td>0.05% (v/v)</td>
			<td>81.45 mL/100 g*</td>
			<td>40.7 &#181;L</td>
			<td>0.001 mg/40.7 &#181;L</td>
			<td>0.02 mg/814 &#181;L</td>
		</tr>
		<tr>
			<td>Pollen supplement</td>
			<td>10 ppb (w/w)</td>
			<td></td>
			<td>10 g**</td>
			<td>4.07 &#181;L</td>
			<td>0.0001 mg/4.07 &#181;L</td>
			<td>0.02 mg/814 &#181;L</td>
		</tr>
		<tr>
			<td>Sucrose solution</td>
			<td>50 ppb (w/w)</td>
			<td>0.05% (v/v)</td>
			<td>78.5 mL/100 g*</td>
			<td>40.7 &#181;L</td>
			<td>0.005 mg/40.7 &#181;L</td>
			<td>0.1 mg/814 &#181;L</td>
		</tr>
		<tr>
			<td>Pollen supplement</td>
			<td>50 ppb (w/w)</td>
			<td></td>
			<td>10 g**</td>
			<td>4.07 &#181;L</td>
			<td>0.0005 mg/4.07 &#181;L</td>
			<td>0.1 mg/814 &#181;L</td>
		</tr>
	</tbody>
</table>
Table 1: Example recipes for treated sucrose solution, pollen supplement, and stock solution. *Volume based on the density of 50% (w/w) sucrose solution (1.228 g/mL). **The density of the pollen supplement will vary depending on what product is used, but if this solvent volume is used, the final solvent concentration in pollen supplement will be within the desired range of &#8804; 5% by volume.
3. Monitoring - Egg Production Rate
<ol>
	<li>Quantify the egg laying 1 to 2 times per day in the morning and/or evening. Begin by removing QMCs from the incubator to check for eggs. 
	NOTE: In a successful experiment, egg production will commence in most of the control QMCs within 3 days of initial cage assembly. Only take as many QMCs out of the incubator at one time that can be checked and fed within 10 min. Longer periods outside the incubator may disrupt egg production.</li>
	<li>Examine the backs of the clear ELPs for eggs. If eggs are present, remove the door panel in front of the plate of interest. Remove the tape from across the ELPs and carefully slide the door panel between the ELP and the bees inside the QMC, taking care not to crush any bees that might be cleaning the cells in the ELPs.</li>
	<li>With the door panel in place, remove the ELP, and count and record the number of eggs inside the ELP cells. Remove the eggs by tapping the edge of the ELP, open cell-side down, on a hard surface (such as the lip of a waste receptacle). Once the eggs fall out, replace the empty ELP in the QMC. Gently remove and replace the door panel behind the ELP on the outside of the QMC. Repeat as necessary with the second ELP and replace the tape across the QMC when finished. 
	&#8203;NOTE: Egg production generally declines and mortality increases in QMCs after 2 weeks32,33, therefore it is recommended to conclude experiments after 14 days.</li>
</ol>
4. Monitoring - Food Consumption
<ol>
	<li>Replace all the food remaining in QMC feeders with freshly prepared diet every two days. Prepare new feeder tubes (including water) and weigh them before removing QMCs from the incubator for monitoring. Swap all old tubes with new ones and weigh old tubes before disposing of unconsumed diet. Compare the final weight of the feeder tube and unconsumed diet to the weight of the same feeder tube prior to placing it in the QMC to estimate diet consumption.</li>
	<li>Between days when feeders are scheduled to be replaced, check diet consumption once per day (at the same time when QMCs are monitored for egg production) to ensure that feeders are never empty. If a feeder tube is empty or near empty, remove it, refill it, record the weight of the tube before and after and add the difference to the 2-day diet consumption total for the QMC.</li>
</ol>
5. Monitoring - Embryo Viability
<ol>
	<li>At a selected point during a QMC experiment, remove ELPs containing freshly laid eggs from the QMC according to step 3, but do not dislodge eggs from the ELP.</li>
	<li>Cover the ELP with a universal microplate lid and place it inside a desiccator with a saturated K2SO4 solution (150 g K2SO4 in 1 L of water, kept in a shallow dish). 
	NOTE: Some salt should be visible on the bottom of the dish after the mixture comes to temperature in the incubator.</li>
	<li>Keep the desiccator in an incubator set to 34.5 &#176;C, resulting in a relative humidity of 95% inside the desiccator, similar to the conditions used by Collins49. 
	NOTE: Almost all eggs will hatch within 72 &#177; 6 hours of when they were laid49, hence hatching rates can be assessed as early as 78 hours after the ELPs were removed from the QMC. A &#34;C&#34; shape larva in the bottom of the cell is indicative of a successful hatching event. Some variation in this timing is possible if, for example, the eggs are drones and not workers50.</li>
</ol>
6. Worker Sampling
<ol>
	<li>If the QMCs have been populated with excess workers, sample the worker bees at a selected time point during the experiment for assessment of treatment induced changes in their physiology. Perform the collections in conjunction with daily feeding and egg counting activities to minimize the time for which the QMCs are outside of the incubator.</li>
	<li>Before sampling, place a door panel between an ELP and the interior of the QMC, and remove the ELP. Carefully lift the door panel approximately 0.5 cm from the base of the cage and remove a worker bee from inside the QMC using featherweight tweezers. To prevent bees from escaping, cover portions of the 0.5 cm opening with a gloved finger or piece of cotton as necessary.</li>
	<li>Preserve the collected bee for follow-up analysis and repeat this process until the desired number of samples have been collected. For gene expression analysis, snap freezing bees in liquid nitrogen and immediate storage at -80 &#176;C is strongly recommended51.</li>
</ol>
7. Worker Mortality
<ol>
	<li>Assess worker mortality during the experiment by counting the number of dead bees at the bottom of the feeding chamber and the egg laying chamber. Perform this assessment in conjunction with daily egg laying quantification.</li>
	<li>Using forceps, carefully remove the dead bees through the feeder holes, covering the hole with a gloved finger or piece of cotton while the forceps are not inserted.</li>
	<li>Remove the dead bees from the egg laying chamber by carefully lifting the door panel approximately 0.5 cm from the base of the cage and inserting forceps. To prevent bees from escaping, cover portions of the 0.5 cm opening with a gloved finger or piece of cotton as necessary.</li>
	<li>Assess the worker mortality at the conclusion of the experiment by removing and counting all the dead bees from the QMCs using the previously described methods prior to euthanizing the remaining bees. 
	NOTE: In the absence of worker bees, queens will not produce eggs and will starve within 24 hours. Therefore, if all the workers in a QMC are observed to be dead, the QMC should be removed from the experiment. Likewise, if a queen dies during the experiment, the QMC should be removed, and the data should be appropriately censored.</li>
</ol>

The authors have no conflicts of interest to declare.

The fecundity of female solitary insects as well as queens in eusocial insect colonies can be influenced by abiotic stressors such as agrochemicals25,28,29,30,33. In honey bees, the effects of agrochemicals on queens may be indirect, as they can occur via changes in their care and feeding by worker bees. Our representative results, which are similar to those r...

Due to increased global demand for agricultural products, modern farming practices often require the use of agrochemicals to control numerous pests known to reduce or harm crop yields1. Simultaneously, the growers of many fruit, vegetable, and nut crops rely on the pollination services provided by commercial honey bee colonies to ensure abundant crop yields2. These practices may result in pollinators, including honey bees (Apis mellifera), being exposed to harmful levels of pesticide residues3. At the same time, the widespread presence of parasitic Varroa destructor mite infestations in honey bee colonies frequently require beekeepers to treat their hives with miticides, which may also exert negative effects on the health and longevity of the colony4,5,6. To reduce and mitigate harmful effects of agrochemical products, it is necessary to fully evaluate their safety to honey bees prior to their implementation so that recommendations for their use can be made to protect beneficial insects.
Currently, the Environmental Protection Agency (EPA) relies upon a tiered risk-assessment strategy for honey bee pesticide exposure, which involves laboratory tests on adult bees and sometimes honey bee larvae7. If lower tier laboratory tests fail to alleviate concerns of toxicity, higher tier field and semi-field testing may be recommended. While these laboratory tests provide valuable insight into the potential effects of agrochemicals on worker longevity, they are not necessarily predictive of their effects on queens, which differ significantly from workers biologically8 and behaviorally9. Furthermore, there are numerous potential effects of agrochemicals on insects beyond mortality, which can have considerable consequences for social insects that rely on coordinated behaviors to function as a colony unit10,11.
Although mortality is the most commonly considered effect of agrochemical pesticides12, these products can have a wide range of effects on both target and non-target arthropods including altered behavior13,14,15,16, repellency or attractancy17,18,19, changes in feeding patterns20,21,22, and increased or decreased fecundity20,21,22,23,24,25. For social insects, these effects can systemically disrupt colony interactions and functions11. Of these functions, reproduction, which is heavily reliant on a single egg-laying queen supported by the rest of the of the colony unit9, may be particularly vulnerable to perturbation due to pesticide exposure.
Studies performed on immature queens have demonstrated that developmental exposure to miticides can affect adult queen behavior, physiology, survival26,27. Similarly, studies using full or reduced sized colonies have demonstrated that agrochemicals can affect adult honey bee queens by decreasing mating success28, decreasing oviposition29, and decreasing the viability of the eggs produced25,30,31. These phenomena have previously been difficult to observe without the use of whole colonies, due largely to a lack of available laboratory methods. However, a method to study queen oviposition under tightly controlled laboratory conditions using Queen Monitoring Cages (QMC)32 has recently been adapted to examine the effects of agrochemicals on queen fecundity33. Here, these techniques are described in detail along with additional methods to measure and track worker diet consumption in QMCs.
These methods are more advantageous than experiments requiring full sized colonies because they allow for the administration of precise doses of agrochemicals to a greatly reduced number of workers relative to the tens of thousands typically present inside a colony34, which then provision the queen. This exposure technique mirrors the second-hand exposure that queens would experience in real-world scenarios because, within a colony, queens do not feed themselves and rely upon workers to provision them with diet9. Similarly, queens do not generally leave the hive except during colony reproduction (swarming) for mating flights35. Mated honey bee queens can be purchased from commercial queen breeders and shipped overnight. Typically, queen breeders sell queens directly after confirming that they have started to lay eggs, which is taken as an indication of successful mating. If more precise information on queen age or relatedness is needed, researchers may consult with the queen breeder before placing an order.
QMCs allow for precise observation and quantification of honey bee queen oviposition and egg hatching rates32,33, yielding valuable data related to the effects of agrochemical exposure on queen fecundity. The representative results presented here describe an experiment quantifying oviposition, diet consumption, and embryo viability in QMCs under chronic exposure to field relevant concentrations of the systemic neurotoxicant neonicotinoid pesticide imidacloprid36. Once applied, imidacloprid translocates to plant tissues37, and residues have been detected the pollen and nectar of numerous bee pollinated plants38,39,40. Exposure to imidacloprid can have a broad range of detrimental effects on honey bees including impaired foraging performance16, impaired immune function41, and decreased rates of colony expansion and survival42,43. Here, imidacloprid was selected for use as a test substance because field experiments have shown that it can affect honey bee queen oviposition29

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Fluon</td>
			<td>BioQuip, Rancho Dominguez, CA</td>
			<td>2871A</td>
			<td></td>
		</tr>
		<tr>
			<td>Honey bee queens</td>
			<td>Olivarez Honey Bees, Orland, CA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Imidacloprid</td>
			<td>Sigma-Aldritch, St. Louis, MO</td>
			<td>37894</td>
			<td></td>
		</tr>
		<tr>
			<td>MegaBee Powder</td>
			<td>MegaBee, San Dieago, CA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microcentrifuge tubes 2 mL</td>
			<td>ThermoFisher Scientific, Waltham, MA</td>
			<td>02-682-004</td>
			<td></td>
		</tr>
		<tr>
			<td>Needles 20 gauge</td>
			<td>W. W. Grainger, Lake Forest, IL</td>
			<td>5FVK4</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Sulfate</td>
			<td>Sigma-Aldritch, St. Louis, MO</td>
			<td>P0772</td>
			<td></td>
		</tr>
		<tr>
			<td>Queen Monitoring Cages</td>
			<td>University of Illinois Urbana-Champaign</td>
			<td></td>
			<td>Patent application number: 20190350175</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma-Aldritch, St. Louis, MO</td>
			<td>S8501</td>
			<td></td>
		</tr>
		<tr>
			<td>Universal Microplate Lids</td>
			<td>ThermoFisher Scientific, Waltham, MA</td>
			<td>5500</td>
			<td></td>
		</tr>
	</tbody>
</table>

assessing agrochemical risk to mated honey bee queens

Current risk assessment strategies for honey bees rely heavily upon laboratory tests performed on adult or immature worker bees, but these methods may not accurately capture the effects of agrochemical exposure on honey bee queens. As the sole producer of fertilized eggs inside a honeybee colony, the queen is arguably the most important single member of a functioning colony unit. Therefore, understanding how agrochemicals affect queen health and productivity should be considered a critical aspect of pesticide risk assessment. Here, an adapted method is presented to expose honey bee queens and worker queen attendants to agrochemical stressors administered through a worker diet, followed by tracking egg production in the laboratory and assessing first instar eclosion using a specialized cage, referred to as a Queen Monitoring Cage. To illustrate the method&#39;s intended use, results of an experiment in which worker queen attendants were fed diet containing sublethal doses of imidacloprid and effects on queens were monitored are described.

The production of eggs was monitored in QMCs assembled and maintained as described above with once daily observations of egg production and 15 cages per treatment group. Newly mated queens of primarily Carniolan stock were purchased and shipped overnight from a queen breeder, and honey bee workers were obtained from 3 colonies maintained according to standard commercial methods at The Bee Research Facility at the University of Illinois Urbana-Champaign. Here, 4 dietary treatment groups were used: 1) 50 ppb (g/g) imidaclo...

Watch this Scientific Journal Video about Assessing Agrochemical Risk to Mated Honey Bee Queens at JoVE.com

Assessing Agrochemical Risk to Mated Honey Bee Queens

This protocol was developed to enhance the understanding of how agrochemicals affect honey bee (Apis mellifera) reproduction by establishing methods to expose honey bee queens and their worker caretakers to agrochemicals in a controlled, laboratory setting and carefully monitoring their relevant responses.

Current risk assessment strategies for honey bees rely heavily upon laboratory tests performed on adult or immature worker bees, but these methods may not ...

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2.6K Views.  USDA-ARS. This protocol was developed to enhance the understanding of how agrochemicals affect honey bee (Apis mellifera) reproduction by establishing methods to expose honey bee queens and their worker caretakers to agrochemicals in a controlled, laboratory setting and carefully monitoring their relevant responses. 

Video: Assessing Agrochemical Risk to Mated Honey Bee Queens

In vivo Ca2+- Imaging of Mushroom Body Neurons During Olfactory Learning in the Honey Bee

The in vivo and semi-in vivo preparation for Calcium imaging has been developed in our lab by Joerges, K&uuml;ttner and Galizia over ten years ago, to measure odor evoked activity in the antennal lobe1. From then on, it has been continuously refined and applied to different neuropiles in the bee brain. Here, we describe the preparation currently used in the lab to measure activity in mushroom body neurons using a dextran coupled calcium-sensitive dye (Fura-2). We retrogradely stain mushroom body neurons by injecting dye into their axons or soma region. We focus on reducing the invasiveness, to achieve a preparation in which it is still possible to train the bee using PER conditioning. We are able to monitor and quantify the behavioral response by recording electro-myograms from the muscle which controls the PER (M17)2.
After the physiological experiment the imaged structures are investigated in greater detail using confocal scanning microscopy to address the identity of the neurons.

In vivo Ca2+- Imaging of Mushroom Body Neurons During Olfactory Learning in the Honey Bee

Bees can be conditioned in an appetitive olfactory learning paradigm (PER-conditioning). Using odors as stimuli, we established a method in which behavior is recorded while simultaneously Calcium Imaging is used to measure odor evoked activity in mushroom body neurons in vivo.

The in vivo and semi-in vivo preparation for Calcium imaging has been developed in our lab by Joerges, K&uuml;ttner and Galizia over ten years ago, to ...

A Noninvasive Hair Sampling Technique to Obtain High Quality DNA from Elusive Small Mammals

Noninvasive genetic sampling approaches are becoming increasingly important to study wildlife populations. A number of studies have reported using noninvasive sampling techniques to investigate population genetics and demography of wild populations1. This approach has proven to be especially useful when dealing with rare or elusive species2. While a number of these methods have been developed to sample hair, feces and other biological material from carnivores and medium-sized mammals, they have largely remained untested in elusive small mammals. In this video, we present a novel, inexpensive and noninvasive hair snare targeted at an elusive small mammal, the American pika (Ochotona princeps). We describe the general set-up of the hair snare, which consists of strips of packing tape arranged in a web-like fashion and placed along travelling routes in the pikas&#x2019; habitat. We illustrate the efficiency of the snare at collecting a large quantity of hair that can then be collected and brought back to the lab. We then demonstrate the use of the DNA IQ system (Promega) to isolate DNA and showcase the utility of this method to amplify commonly used molecular markers including nuclear microsatellites, amplified fragment length polymorphisms (AFLPs), mitochondrial sequences (800bp) as well as a molecular sexing marker. Overall, we demonstrate the utility of this novel noninvasive hair snare as a sampling technique for wildlife population biologists. We anticipate that this approach will be applicable to a variety of small mammals, opening up areas of investigation within natural populations, while minimizing impact to study organisms.

We present a noninvasive sampling approach to efficiently collect hair samples from elusive small mammals, as shown for the American pika. We demonstrate the utility of this method by extracting DNA from sampled hair and amplifying several types of molecular markers commonly used in studies of wildlife ecology and conservation.

Noninvasive genetic sampling approaches are becoming increasingly important to study wildlife populations. A number of studies have reported using ...

Heart Dissection in Larval, Juvenile and Adult Zebrafish, Danio rerio

Zebrafish have become a beneficial and practical model organism for the study of embryonic heart development (see recent reviews1-6), however, work examining post-embryonic through adult cardiac development has been limited7-10. Examining the changing morphology of the maturing and aging heart are restricted by the lack of techniques available for staging and isolating juvenile and adult hearts. In order to analyze heart development over the fish's lifespan, we dissect zebrafish hearts at numerous stages and photograph them for further analysis11. The morphological features of the heart can easily be quantified and individual hearts can be further analyzed by a host of standard methods. Zebrafish grow at variable rates and maturation correlates better with fish size than age, thus, post-fixation, we photograph and measure fish length as a gauge of fish maturation. This protocol explains two distinct, size dependent dissection techniques for zebrafish, ranging from larvae 3.5mm standard length (SL) with hearts of 100&mu;m ventricle length (VL), to adults, with SL of 30mm and VL 1mm or larger. Larval and adult fish have quite distinct body and organ morphology. Larvae are not only significantly smaller, they have less pigment and each organ is visually very difficult to identify. For this reason, we use distinct dissection techniques.
We used pre-dissection fixation procedures, as we discovered that hearts dissected directly after euthanization have a more variable morphology, with very loose and balloon like atria compared with hearts removed following fixation. The fish fixed prior to dissection, retain in vivo morphology and chamber position (data not shown). In addition, for demonstration purposes, we take advantage of the heart (myocardial) specific GFP transgenic Tg(myl7:GFP)twu34 (12), which allows us to visualize the entire heart and is particularly useful at early stages in development when the cardiac morphology is less distinct from surrounding tissues. Dissection of the heart makes further analysis of the cell and molecular biology underlying heart development and maturation using in situ hybridization, immunohistochemistry, RNA extraction or other analytical methods easier in post-embryonic zebrafish. This protocol will provide a valuable technique for the study of cardiac development maturation and aging.

Heart Dissection in Larval, Juvenile and Adult Zebrafish, Danio rerio

A clear, standardized method for dissection and isolation of the zebrafish heart at multiple developmental stages are described. Annotation and quantification techniques are also discussed.

Zebrafish have become a beneficial and practical model organism for the study of embryonic heart development (see recent reviews1-6), however, work ...

Using RNA-mediated Interference Feeding Strategy to Screen for Genes Involved in Body Size Regulation in the Nematode C. elegans

Double-strand RNA-mediated interference (RNAi) is an effective strategy to knock down target gene expression1-3. It has been applied to many model systems including plants, invertebrates and vertebrates. There are various methods to achieve RNAi in vivo4,5. For example, the target gene may be transformed into an RNAi vector, and then either permanently or transiently transformed into cell lines or primary cells to achieve gene knockdown effects; alternatively synthesized double-strand oligonucleotides from specific target genes (RNAi oligos) may be transiently transformed into cell lines or primary cells to silence target genes; or synthesized double-strand RNA molecules may be microinjected into an organism. Since the nematode C. elegans uses bacteria as a food source, feeding the animals with bacteria expressing double-strand RNA against target genes provides a viable strategy6. Here we present an RNAi feeding method to score body size phenotype. Body size in C. elegans is regulated primarily by the TGF- &beta; - like ligand DBL-1, so this assay is appropriate for identification of TGF-&beta; signaling components7. We used different strains including two RNAi hypersensitive strains to repeat the RNAi feeding experiments. Our results showed that rrf-3 strain gave us the best expected RNAi phenotype. The method is easy to perform, reproducible, and easily quantified. Furthermore, our protocol minimizes the use of specialized equipment, so it is suitable for smaller laboratories or those at predominantly undergraduate institutions.

Using RNA-mediated Interference Feeding Strategy to Screen for Genes Involved in Body Size Regulation in the Nematode C. elegans

We demonstrate how to use the RNAi feeding technique to knock down target genes and score body size phenotype in C. elegans. This method could be used for a large scale screen to identify potential genetic components of interest, such as those involved in body size regulation by DBL-1/TGF-&beta; signaling.

Double-strand RNA-mediated interference (RNAi) is an effective strategy to knock down target gene expression1-3. It has been applied to many model systems ...

Obtaining Specimens with Slowed, Accelerated and Reversed Aging in the Honey Bee Model

Societies of highly social animals feature vast lifespan differences between closely related individuals. Among social insects, the honey bee is the best established model to study how plasticity in lifespan and aging is explained by social factors.
The worker caste of honey bees includes nurse bees, which tend the brood, and forager bees, which collect nectar and pollen. Previous work has shown that brain functions and flight performance senesce more rapidly in foragers than in nurses. However, brain functions can recover, when foragers revert back to nursing tasks. Such patterns of accelerated and reversed functional senescence are linked to changed metabolic resource levels, to alterations in protein abundance and to immune function. Vitellogenin, a yolk protein with adapted functions in hormonal control and cellular defense, may serve as a major regulatory element in a network that controls the different aging dynamics in workers.
Here we describe how the emergence of nurses and foragers can be monitored, and manipulated, including the reversal from typically short-lived foragers into longer-lived nurses. Our representative results show how individuals with similar chronological age differentiate into foragers and nurse bees under experimental conditions. We exemplify how behavioral reversal from foragers back to nurses can be validated. Last, we show how different cellular senescence can be assessed by measuring the accumulation of lipofuscin, a universal biomarker of senescence.
For studying mechanisms that may link social influences and aging plasticity, this protocol provides a standardized tool set to acquire relevant sample material, and to improve data comparability among future studies.

In honey bee workers, aging depends on social behaviors rather than on chronological age. Here we show how worker-types with very different aging patterns can be obtained and analyzed for cellular senescence.

Societies of highly social animals feature vast lifespan differences between closely related individuals. Among social insects, the honey bee is the best ...

Methods for Comparing Nutrients in Beebread Made by Africanized and European Honey Bees and the Effects on Hemolymph Protein Titers

Honey bees obtain nutrients from pollen they collect and store in the hive as beebread. We developed methods to control the pollen source that bees collect and convert to beebread by placing colonies in a specially constructed enclosed flight area. Methods were developed to analyze the protein and amino acid composition of the pollen and beebread. We also describe how consumption of the beebread was measured and methods used to determine adult worker bee hemolymph protein titers after feeding on beebread for 4, 7 and 11 days after emergence. Methods were applied to determine if genotype affects the conversion of pollen to beebread and the rate that bees consume and acquire protein from it. Two subspecies (European and Africanized honey bees; EHB and AHB respectively) were provided with the same pollen source. Based on the developed methods, beebread made by both subspecies had lower protein concentrations and pH values than the pollen. In general, amino acid concentrations in beebread made by either EHB or AHB were similar and occurred at higher levels in beebread than in pollen. Both AHB and EHB consumed significantly more of the beebread made by AHB than by EHB. Though EHB and AHB consumed similar amounts of each type of beebread, hemolymph protein concentrations in AHB were higher than in EHB. Differences in protein acquisition between AHB and EHB might reflect environmental adaptations related to the geographic region where each subspecies evolved. These differences could contribute to the successful establishment of AHB populations in the New World because of the effects on brood rearing and colony growth.

Here are methods to quantify nutrients in pollen before and after its conversion to beebread by two subspecies of honeybees. We describe techniques to measure beebread consumption and resulting protein titers in both subspecies.

Honey bees obtain nutrients from pollen they collect and store in the hive as beebread. We developed methods to control the pollen source that bees ...

Dissection and Observation of Honey Bee Dorsal Vessel for Studies of Cardiac Function

The European honey bee, Apis mellifera L., is a valuable agricultural and commercial resource noted for producing honey and providing crop pollination services, as well as an important model social insect used to study memory and learning, aging, and more. Here we describe a detailed protocol for the dissection of the dorsal abdominal wall of a bee in order to visualize its dorsal vessel, which serves the role of the heart in the insect. A successful dissection will expose a functional heart that, under the proper conditions, can maintain a steady heartbeat for an extended period of time. This allows the investigator to manipulate heart rate through the application of cardiomodulatory compounds to the dorsal vessel. By using either a digital microscope or a microscope equipped with a digital camera, the investigator can make video recordings of the dorsal vessel before and after treatment with test compounds. The videos can then be scored at a time convenient to the user in order to determine changes in heart rate, as well as changes in the pattern of heartbeats, following treatment. The advantages of this protocol are that it is relatively inexpensive to set up, easy to learn, requires little space or equipment, and takes very little time to conduct.

The abdominal dorsal vessel of the honey bee and other insects serves as the functional equivalent of the mammalian heart and plays an important role in nutrient transport, waste removal, immune function, and more. Here we describe a protocol for the visualization and pharmacological manipulation of bee heart rate.

The European honey bee, Apis mellifera L., is a valuable agricultural and commercial resource noted for producing honey and providing crop pollination ...

A Protocol for Using Gene Set Enrichment Analysis to Identify the Appropriate Animal Model for Translational Research

Recent studies that compared transcriptomic datasets of human diseases with datasets from mouse models using traditional gene-to-gene comparison techniques resulted in contradictory conclusions regarding the relevance of animal models for translational research. A major reason for the discrepancies between different gene expression analyses is the arbitrary filtering of differentially expressed genes. Furthermore, the comparison of single genes between different species and platforms often is limited by technical variance, leading to misinterpretation of the con/discordance between data from human and animal models. Thus, standardized approaches for systematic data analysis are needed. To overcome subjective gene filtering and ineffective gene-to-gene comparisons, we recently demonstrated that gene set enrichment analysis (GSEA) has the potential to avoid these problems. Therefore, we developed a standardized protocol for the use of GSEA to distinguish between appropriate and inappropriate animal models for translational research. This protocol is not suitable to predict how to design new model systems a-priori, as it requires existing experimental omics data. However, the protocol describes how to interpret existing data in a standardized manner in order to select the most suitable animal model, thus avoiding unnecessary animal experiments and misleading translational studies.

We provide a standardized protocol for the use of gene set enrichment analysis of transcriptomic data to identify an ideal mouse model for translational research. 
This protocol can be used with DNA microarray and RNA sequencing data and can further be extended to other omics data if data are available.

Recent studies that compared transcriptomic datasets of human diseases with datasets from mouse models using traditional gene-to-gene comparison ...

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 ...

At-Risk Butterfly Captive Propagation Programs to Enhance Life History Knowledge and Effective Ex Situ Conservation Techniques

Improving knowledge of ex situ best practices for at-risk butterflies is important for generating successful conservation and recovery program outcomes. Research on such captive populations can also yield valuable data to address key information gaps about the behavior, life history, and ecology of the target taxa. We describe a protocol for captive propagation of the federally endangered Cyclargus thomasi bethunebakeri that can be used as a model for other at-risk butterfly ex situ programs, especially those in the family Lycaenidae. We further provide a simple and straightforward protocol for recording various life history metrics that can be useful for informing ex situ methodologies as well as adapted for laboratory studies of other lepidoptera.

Here, we present protocols for 1) the laboratory captive propagation of the federally endangered Miami blue butterfly (Cyclargus thomasi bethunebakeri), and 2) assessing basic life history information such as immature development time and number of larval stadia. Both methods can be adapted for use with other ex situ conservation programs.

Improving knowledge of ex situ best practices for at-risk butterflies is important for generating successful conservation and recovery program outcomes. ...