We thank U. Heberlein and A. Devineni for long-lasting discussions and technical advice. We also thank the Shohat-Ophir lab members, A. Benzur, L. Kazaz, and O. Shalom, for the help with demonstrating the method. Special appreciation goes to Eliezer Costi for establishing the fly systems in the lab. This work was supported by the Israel Science Foundation (384/14) and the Marie Curie Career Integration Grants (CIG 631127).

Note: General overview of the experimental design: The experimental design includes an adapted protocol for courtship suppression7-9 in which male flies are exposed to rewarding and nonrewarding experiences in 3 consecutive training sessions over the course of 4 d. At the end of the experience phase, the flies are tested in a two-choice voluntary ethanol consumption assay for 3&#160;- 4 d. The protocol herein includes several preparatory steps, some of which can be done in advance to be used in more than one experiment, while others should take place in a timely manner before the beginning of the experiment (Table 1).
1. Preparatory Steps
<ol>
	<li>Preparing the capillary feeder system
	<ol>
		<li>Heat a 27&#160;G needle and use it to puncture small holes along a vial (Figure 1).</li>
		<li>Cut the vial plug in half (crosswise) using a razor blade. Mark four points on the surface of the plug, creating a square shape to position the capillary holders into. Use an 18&#160;G needle to make 4 holes through the plug (Figure 1b).</li>
		<li>Use a razor blade to cut micropipets so that they can firmly hold 5 &#181;L glass capillaries, which serve as adaptors.</li>
		<li>Insert 4 capillary adaptors into the prepared plugs.</li>
		<li>Mark the position of the 2 ethanol- and&#160;two nonethanol-containing capillaries on the plug. 
		Note: It is recommended to keep the positions of the 2 types of capillaries consistent across all vials. Pay attention to the size of the holes, as large holes allow the escape of flies out of the vials. In addition, it is recommended to use the smoother side of the plug (usually opposite the cut side) as the side that faces down when inserted into the vial, as cutting the plug creates some small, loose fibers that the flies can get tangled in.</li>
		<li>Prepare a large number of vials and adaptors to be used for the capillary feeder system (CAFE) experiments described in step 2.3.</li>
	</ol>
	</li>
	<li>Preparing small glass food vials
	<ol>
		<li>A day before collecting the flies (step 1.4), arrange 200 glass vials on microcentrifuge tube racks. Melt fly food (according to the cornmeal molasses recipe, see the Table of Materials) using a microwave. Pour 2 mL of fly food into the glass vials (try not to smear the food on the vial walls). Let the food harden, and cover the vials with plastic wrap. 
		NOTE: The glass vials containing the food can be kept at 4 &#176;C for several weeks.</li>
		<li>On the day of fly collection, take the vials out of the refrigerator and let them reach RT before using them to house flies. 
		NOTE: If food vials are several weeks old, make sure that the food is still intact and not detached from the vial, as flies can be trapped in the cracks between the food and the glass.</li>
	</ol>
	</li>
	<li>Making stock food solution for the capillary feeder assay
	<ol>
		<li>To prepare the stock food solution (5% sucrose and 5% yeast extract), dissolve 5.88 g of yeast extract and 5.88 g of sucrose in 100 mL of distilled water. 
		NOTE: The stock solution is highly prone to contamination. Therefore, it is recommended to autoclave it (121 &#176;C, 30 min, and without a drying cycle) and aliquot it into 850&#160;&#181;L working aliquots to be kept at 4 &#176;C until further use. The concentration of the ingredients in the stock solution is 5.88% and will reach the final concentration of 5% after the addition of 15% ethanol or water (150 &#181;L into 850 &#181;L of premade food solution) just before its use in step 2.3.</li>
	</ol>
	</li>
	<li>Collecting experimental flies
	<ol>
		<li>Collect 200 na&#239;ve male flies using a CO2 pad. 
		Note: Use any lab strain relevant for the experimental question. 250 flies can be easily collected from five bottles of flies at their eclosion pick phase. When collecting the flies, pay careful attention to collect flies that have recently eclosed (virgin-looking flies: bright flies with a meconium spot on their abdomen). Experimental flies should be kept singly in small glass vials containing food and aged to 3 - 4 d&#160;old before being used in the experiment.</li>
		<li>Use soft plugs to close the vials. 
		NOTE: The same soft plugs will be very useful for the training phase when using mouth aspiration to insert females into and out of the vials during the experience phase.</li>
	</ol>
	</li>
	<li>Collecting trainer flies 
	NOTE: All collected flies should be kept in a 12&#160;h/12&#160;h light/dark incubator at 25 &#176;C and 70% Relative Humidity (RH). The 12&#160;h/12&#160;h light/dark cycle can be changed to a convenient starting time.
	<ol>
		<li>Virgin female trainer flies
		<ol>
			<li>Collect 500 virgin female trainer flies from a WT stock or a virginator stock for use as virgin receptive females. Keep the female flies in groups of 25 per regular-size food vial, and allow them to age to 3 - 4 d old before using them in the experiment.</li>
		</ol>
		</li>
		<li>Mated female trainer flies
		<ol>
			<li>Collect 300 virgin female flies from a WT stock or a virginator stock to undergo mating before the training phase (step 1.6.1). 
			NOTE: Female flies are kept in groups of 10 females per regular-size food vial and aged to 3 - 4 d old before being used in the experiment.</li>
		</ol>
		</li>
		<li>Male flies used to generate mated female trainer flies
		<ol>
			<li>Collect 150 male flies (not necessarily virgin). 
			NOTE: Flies are kept in groups of 15 males per regular-size food vial and aged to 3 - 4 d old before being paired with virgin female flies to create the mated trainer females.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Generating mated female trainer flies
	<ol>
		<li>To generate mated female flies to be used as trainers, pair groups of 10 virgin female flies with 15 male flies by flipping one male-containing vial into a female-containing vial 16 - 18 h before the training phase (step 2.1)10. 
		NOTE: This should normally take place the evening before the training session. The 1.5:1 ratio between male flies to female flies assures that all females will be mated.</li>
		<li>On the day of the experiment, separate the mated females from the males by aspirating the males out of the vial using a mouth aspirator. Perform this right before the beginning of each training day.</li>
	</ol>
	</li>
</ol>
2. Experimental Steps
<ol>
	<li>Setting up the experience phase
	<ol>
		<li>Start the experiment as close as possible to the beginning of the light phase in the incubator. Place the trainer females (both virgin and mated) in the behavior chamber. 
		NOTE: It is recommended to perform the experiment in a behavior chamber, or any secluded environment that is quiet and set to 25 &#176;C and 60 - 65% RH. Keeping humidity over 50% is very important, since courtship and mating behaviors depend on pheromone sensing, which is known to be sensitive to humidity.</li>
	</ol>
	</li>
	<li>Exposing male flies to rewarding and nonrewarding experiences
	<ol>
		<li>Arrange the glass vials containing the single male flies on microcentrifuge racks, positioning the vials along the margins of the rack. 
		NOTE: This arrangement allows the experimenter to easily observe the pairs of flies that undergo copulation without taking the vials out of the rack.</li>
		<li>Prepare (as described in step 2.2.1) 2 groups of 100 glass vials containing single male flies on separate racks for males undergoing rewarding and nonrewarding mating experiences (i.e.&#160;rejected and mated).</li>
		<li>Aspirate trainer females and add them to each vial containing the male flies. Set the timer for 1 h. Starting with the rejected group, add a mated female to each vial. Then, move on to the males to be mated and add a virgin female to each vial.</li>
		<li>Watch the mating encounters closely to make sure that the males interacting with virgin females mate successfully within the first hour and that the males interacting with the previously-mated females do not mate. To monitor the rejected cohort over the course of 1 h, set a second timer for 15 min and watch the males every 15 min.</li>
		<li>Discharge the males from the rejected cohort that managed to mate and males from the mated cohort that did not end up mating by the end of the first training session.</li>
		<li>Finish the first training session by aspirating out the previously-mated female trainers into a regular fly food vial, and keep them for the next conditioning session. Follow this step by gently aspirating the females from the mated male group.</li>
		<li>Let the flies rest for 1 h.</li>
		<li>Repeat steps 2.2.3 - 2.2.7 twice.</li>
		<li>Repeat the training session in steps 2.2.3 - 2.2.8 each day for 3 more d.</li>
	</ol>
	</li>
	<li>Recording consumption (CAFE)
	<ol>
		<li>At the end of the last conditioning session (fourth day), aspirate the single males and group them into the capillary feeder vials. Use a soft plug to cover the capillary feeder vial while inserting the flies, and replace it quickly with the ready-made capillary feeder plug after all the flies have been inserted. Group 6 - 8 flies from each of the experimental groups into a vial, creating no less than 8 vials per condition.</li>
		<li>Wet the plugs by gently adding 5 mL of water to the top of the plug. Be careful while adding the water to avoid spillage into the vials. Add equal amount of water to each vial. 
		NOTE: This step is done to reduce evaporation from the capillaries. Therefore, it is repeated every day for the rest of the experiment. Depending on the relative humidity, the following days may require less water to be added to the plugs.</li>
		<li>Prepare two separate food solutions by adding ethanol or water to the stock food aliquots to create a final concentration of 15%.</li>
		<li>To fill the capillaries with the two food solutions, create 5&#160;&#181;L liquid food drops on a laboratory film strip and allow the solution to fill the capillary until it reaches the 5&#160;&#181;L mark. Make sure to use the clear end of the capillary, which is close to the 5&#160;&#181;L black mark line. 
		NOTE: To fill several capillaries at once, it is possible to connect 6 - 8 capillary adaptors together and use them as an insert to hold the capillaries.</li>
		<li>Tap the capillaries gently on the paraffin strip to position the food solution exactly at the black mark. Seal the capillaries by dipping the end of each capillary into small drops of mineral oil. This step minimizes potential evaporation of the food solution.</li>
		<li>Insert two ethanol-containing capillaries and two nonethanol containing capillaries through the 4 adaptors/holders in the plug until the exposed sides of the capillaries barely peek from the interior of the vial (1 mm or less from the plug surface). 
		NOTE: Keeping the opening of the capillaries close to the wet plug reduces evaporation and allows easy access to the food, as flies normally sit on the plugs and climb down on the exposed capillary to feed.</li>
		<li>In addition to vials that contain experimental flies, prepare a mock vial without flies. This control serves to assess natural evaporation. 
		NOTE: Try to maintain relatively equal positioning of the capillary opening compared to the plugs, as differences can lead to variable evaporation and to misleading consumption data. Maintain adequate humidity during the experiment and position the capillary openings as closely as possible to the wet plug, as low humidity and large distance from the wet plug can both result in uncontrolled evaporation.</li>
		<li>Record the relative position of the liquid in each of the capillaries with respect to the black mark line in mm using a ruler, and write these values down. 
		NOTE: If the solution is positioned exactly at the black line, the value should equal zero, and if it is above or below, it should be positive or negative values, respectively. The initial levels of the liquid set the reference points for the final O/N consumption values.</li>
		<li>Set the vials back in the incubator for 24 h. Replace the capillaries at a constant time throughout the entire experiment.</li>
		<li>Remove the capillaries gently from the vials and measure the levels of liquid with respect to the black marking line.</li>
		<li>Carefully check the liquid level in the mock vial. 
		NOTE: If the levels dropped more than 2 mm, it indicates that evaporation is high and the consumption levels are not reliable.</li>
		<li>Dampen the plugs gently with 2 mL of water and insert a new set of capillaries, as indicated in steps 2.3.3 - 2.3.10. Set the vials back in the incubator for 24 h.</li>
	</ol>
	</li>
	<li>Determining preference
	<ol>
		<li>Calculate the total consumption of each capillary by subtracting the final level from the initial level at the starting point.</li>
		<li>Combine the values of both capillaries with the same food (i.e.&#160;nonethanol capillaries separately from ethanol-containing capillaries).</li>
		<li>Calculate ethanol preference by subtracting the amount consumed from the non-ethanol capillaries from the amount consumed from the ethanol-containing capillaries and divide the value by the total consumption values from all the capillaries of the same vial.</li>
		<li>Average the preference indexes of all vials of the same experimental group. 
		<img alt="Equation1" src="/files/ftp_upload/54910/54910equation1.jpg" /> 
		NOTE: Nonethanol 1 (NE1), Nonethanol 2 (NE2), Ethanol 1 (E1), Ethanol 2 (E2), Total nonethanol (Total NE) = NE1 + NE2, Total ethanol (Total E) = E1 + E2, Total sum consumption (Total Sum) = Total NE + Total E, and Ethanol preference Index (PI)</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Robinson, G. E., Fernald, R. D., Clayton, D. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genes+and+social+behavior.">Genes and social behavior.</a> Science. 322, 896-900 (2008).</li><li>Koob, G. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neurobiological+substrates+for+the+dark+side+of+compulsivity+in+addiction.">Neurobiological substrates for the dark side of compulsivity in addiction.</a> Neuropharmacol. 56, 18-31 (2009).</li><li>Kaun, K. R., Devineni, A. V., Heberlein, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+melanogaster+as+a+model+to+study+drug+addiction.">Drosophila melanogaster as a model to study drug addiction.</a> Hum Genet. 131, 959-975 (2012).</li><li>Devineni, A. V., Heberlein, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Addiction-like+behavior+in+Drosophila.">Addiction-like behavior in Drosophila.</a> Commun Integr Biol. 3, 357-359 (2010).</li><li>Shohat-Ophir, G., Kaun, K. R., Azanchi, R., Mohammed, H., Heberlein, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sexual+deprivation+increases+ethanol+intake+in+Drosophila.">Sexual deprivation increases ethanol intake in Drosophila.</a> Science. 335, 1351-1355 (2012).</li><li>Devineni, A. V., Heberlein, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preferential+ethanol+consumption+in+Drosophila+models+features+of+addiction.">Preferential ethanol consumption in Drosophila models features of addiction.</a> Curr Biol : CB. 19, 2126-2132 (2009).</li><li>McBride, S. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mushroom+body+ablation+impairs+short-term+memory+and+long-term+memory+of+courtship+conditioning+in+Drosophila+melanogaster.">Mushroom body ablation impairs short-term memory and long-term memory of courtship conditioning in Drosophila melanogaster.</a> Neuron. 24, 967-977 (1999).</li><li>Ejima, A., Smith, B. P., Lucas, C., Levine, J. D., Griffith, L. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sequential+learning+of+pheromonal+cues+modulates+memory+consolidation+in+trainer-specific+associative+courtship+conditioning.">Sequential learning of pheromonal cues modulates memory consolidation in trainer-specific associative courtship conditioning.</a> Curr Biol. 15, 194-206 (2005).</li><li>Ejima, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generalization+of+courtship+learning+in+Drosophila+is+mediated+by+cis-vaccenyl+acetate.">Generalization of courtship learning in Drosophila is mediated by cis-vaccenyl acetate.</a> Curr Biol. 17, 599-605 (2007).</li><li>Keleman, K., Kruttner, S., Alenius, M., Dickson, B. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Function+of+the+Drosophila+CPEB+protein+Orb2+in+long-term+courtship+memory.">Function of the Drosophila CPEB protein Orb2 in long-term courtship memory.</a> Nat Neurosci. 10, 1587-1593 (2007).</li><li>Kaun, K. R., Azanchi, R., Maung, Z., Hirsh, J., Heberlein, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Drosophila+model+for+alcohol+reward.">A Drosophila model for alcohol reward.</a> Nat Neurosci. 14, 612-619 (2011).</li><li>Xu, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+propensity+for+consuming+ethanol+in+Drosophila+requires+rutabaga+adenylyl+cyclase+expression+within+mushroom+body+neurons.">The propensity for consuming ethanol in Drosophila requires rutabaga adenylyl cyclase expression within mushroom body neurons.</a> Genes Brain Behav. 11, 727-739 (2012).</li><li>Devineni, A. V., Heberlein, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+evolution+of+Drosophila+melanogaster+as+a+model+for+alcohol+research.">The evolution of Drosophila melanogaster as a model for alcohol research.</a> Annu Rev Neurosci. 36, 121-138 (2013).</li><li>Ro, J., Harvanek, Z. M., Pletcher, S. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=FLIC:+high-throughput,+continuous+analysis+of+feeding+behaviors+in+Drosophila.">FLIC: high-throughput, continuous analysis of feeding behaviors in Drosophila.</a> PloS one. 9, 101107 (2014).</li><li>Itskov, P. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Automated+monitoring+and+quantitative+analysis+of+feeding+behaviour+in+Drosophila.">Automated monitoring and quantitative analysis of feeding behaviour in Drosophila.</a> Nat Commun. 5, 4560 (2014).</li><li>Yapici, N., Cohn, R., Schusterreiter, C., Ruta, V., Vosshall, L. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Taste+Circuit+that+Regulates+Ingestion+by+Integrating+Food+and+Hunger+Signals.">A Taste Circuit that Regulates Ingestion by Integrating Food and Hunger Signals.</a> Cell. 165, 715-729 (2016).</li><li>Peru, Y. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-lasting,+experience-dependent+alcohol+preference+in+Drosophila.">Long-lasting, experience-dependent alcohol preference in Drosophila.</a> Addict Biol. 19, 392-401 (2014).</li><li>Dus, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nutrient+Sensor+in+the+Brain+Directs+the+Action+of+the+Brain-Gut+Axis+in+Drosophila.">Nutrient Sensor in the Brain Directs the Action of the Brain-Gut Axis in Drosophila.</a> Neuron. 87, 139-151 (2015).</li><li>Anderson, D. J., Adolphs, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+framework+for+studying+emotions+across+species.">A framework for studying emotions across species.</a> Cell. 157, 187-200 (2014).</li><li>Gibson, W. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Behavioral+responses+to+a+repetitive+visual+threat+stimulus+express+a+persistent+state+of+defensive+arousal+in+Drosophila.">Behavioral responses to a repetitive visual threat stimulus express a persistent state of defensive arousal in Drosophila.</a> Curr Biol. 25, 1401-1415 (2015).</li></ol>

The authors have nothing to disclose.

Here, we illustrate the details of an integrated approach to measure alterations in reward-seeking behavior, based on previous work described by Devineni et al.6 and Shohat-Ophir et al5. The first section of the protocol uses different types of sexual interactions as the experience input, and the second section uses a two-choice feeding assay to assess the effect of experience on the preference to consume ethanol.
As shown by Devineni et al.<sup...

Modification of behavior in response to experience allows animals to adjust their behavior to changes in their environment1. During this process, animals integrate their internal physiological state with the changing conditions of the external environment and subsequently choose one action over another to increase their chances of survival and reproduction. Reward systems evolved to motivate behaviors that are required for the survival of individuals and species by reinforcing behaviors that enhance immediate survival, such as eating or drinking, or those that ensure long-term survival, such as sexual behavior or caring for offspring2. Artificial compounds such as drugs of abuse also affect reward systems by co-opting neural pathways that mediate natural rewards2.
During the last two decades, the fruit fly Drosophila melanogaster has been established as a promising model for studying the molecular and neuronal mechanisms that are shaping the effects of ethanol on behavior3,4.
Previously, we have identified a subset of peptidergic neurons in flies (NPF/NPF receptor (R) neurons) that couple natural rewards, such as sexual experience, to the motivation of obtaining drug rewards5. NPF expression is sensitive to both sexual experiences and to drug rewards, such as ethanol intoxication. Changes in NPF expression levels are converted to alterations in ethanol self-administration5, where high NPF reduces and low NPF increases the preference to consume ethanol. Activating NPF neurons is rewarding for flies, as they display strong preference for an odor paired with the activation, which is also reflected by reduced ethanol consumption. More importantly, activation of NPF neurons interferes with the ability of flies to form a positive association between ethanol intoxication and an odor cue. The causal link between the NPF/R system, reward memory, and ethanol consumption suggests that one can use ethanol self-administration as a measure for changes in reward states5.
In this publication we demonstrate an integrated approach for tapping into the fly natural reward system and assaying changes in reward states. The approach consists of two separate parts, a training protocol for manipulating natural reward-related experiences, followed by a two-choice capillary feeder assay (CAFE) to assess ethanol self-administration as an estimate for changes in reward states. The CAFE assay is analogous to the two-bottle choice assays used in rodent studies for drug self-administration and has been shown to reflect certain properties of addiction-like behavior in flies6.

<table border="0" cellpadding="0" cellspacing="0" width="1176">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">Polystyrene 25 x 95mm Vials</td>
			<td style="width:352px;">FlyStuff</td>
			<td style="width:251px;">32-109</td>
			<td style="width:285px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">narrow plastic vials flugs</td>
			<td style="width:352px;">FlyStuff</td>
			<td style="width:251px;">42-102</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Disposable Sterile Needle 18G and 27G&#160;</td>
			<td>can be acquired by any company</td>
			<td style="width:251px;"></td>
			<td>1.20 X 38mm (18Gx1 1/2&#34;) , 0.40 X 13mm (27Gx1/2&#34;)</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:288px;">10x75mm Borosilicate Glass Disposable Culture Tubes</td>
			<td style="width:352px;">kimble chase</td>
			<td style="width:251px;">73500-1075</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">calibrated pipets 5ul (microliter)</td>
			<td style="width:352px;">VWR</td>
			<td style="width:251px;">53432-706</td>
			<td>color coded white to contain 5 microliters</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Mineral Oil&#160;</td>
			<td style="width:352px;">Sigma-Aldrich&#160;</td>
			<td style="width:251px;">M5904</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">Sucrose, Molecular Biology Grade</td>
			<td style="width:352px;">CALBIOCHEM</td>
			<td dir="LTR" style="width:251px;">573113</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">Yeast extract Powder for microbiology</td>
			<td>can be acquired by any company</td>
			<td style="width:251px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">Ethanol</td>
			<td style="width:352px;">Sigma-Aldrich&#160;</td>
			<td>32221</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:288px;">standard pipette Tips (micro-pipets)</td>
			<td style="width:352px;">ThermScientific</td>
			<td style="width:251px;">T114R-Q</td>
			<td>volume- 0.1-20 ul Ultra micro</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:288px;">IDENTI-PLUGS (Foam Tube Plugs)</td>
			<td>Jaece</td>
			<td>L800-A</td>
			<td>fits opening 6 to 13mm</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:288px;">IDENTI-PLUGS (Foam Tube Plugs)</td>
			<td>Jaece</td>
			<td>L800-D</td>
			<td>fits opening 35 to 45mm</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">virginator fly stock&#160;</td>
			<td style="width:352px;">bloomington drosophila stock center</td>
			<td style="width:251px;">#24638</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Narrow Vials, Tray Pack (PS)</td>
			<td>Genesee Scientific Corporation</td>
			<td>&#160;# 32-109BR</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;"></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Drosophila Media Recipes and Methods</td>
			<td>Bloomington Drosophila Stock Center</td>
			<td colspan="2">http://flystocks.bio.indiana.edu/Fly_Work/media-recipes/molassesfood.htm</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">propionic acid</td>
			<td style="width:352px;">Sigma-Aldrich&#160;</td>
			<td style="width:251px;">P5561</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">phosphoric acid</td>
			<td style="width:352px;">Sigma-Aldrich&#160;</td>
			<td style="width:251px;">W290017</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">Methl 4-Hydroxybenzoate</td>
			<td style="width:352px;">Sigma-Aldrich&#160;</td>
			<td style="width:251px;">H3647</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">Agar Agar</td>
			<td>can be acquired by any company</td>
			<td style="width:251px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">corn meal</td>
			<td>can be acquired by any company</td>
			<td style="width:251px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">Grandma&#39;s molasses</td>
			<td>B&#38;G Foods, Inc</td>
			<td style="width:251px;">not indicated</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">instant dry yeast</td>
			<td>can be acquired by any company</td>
			<td style="width:251px;"></td>
			<td></td>
		</tr>
	</tbody>
</table>

a simple way to measure alterations in reward-seeking behavior using drosophila melanogaster

We describe a protocol for measuring ethanol self-administration in fruit flies (Drosophila melanogaster) as a proxy for changes in reward states. We demonstrate a simple way to tap into the fly reward system, modify experiences related to natural reward, and use voluntary ethanol consumption as a measure for changes in reward states. The approach serves as a relevant tool to study the neurons and genes that play a role in experience-mediated changes of internal state. The protocol is composed of two discrete parts: exposing the flies to rewarding and nonrewarding experiences, and assaying voluntary ethanol consumption as a measure of the motivation to obtain a drug reward. The two parts can be used independently to induce the modulation of experience as an initial step for further downstream assays or as an independent two-choice feeding assay, respectively. The protocol does not require a complicated setup and can therefore be applied in any laboratory with basic fly culture tools.

FPreviously, Devineni et al. showed that when fruit flies are given the choice to consume ethanol-containing food, they display a strong preference for ethanol-containing food over nonethanol containing food6. Shown here are some representative results we obtained when assaying the innate ethanol preference of na&#239;ve male flies that did not undergo the training protocol.
Na&#239;ve Canton S male flies wer...

Watch this Scientific Journal Video about A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster at JoVE.com

A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

We describe a protocol for inducing rewarding and nonrewarding experiences in fruit flies (Drosophila melanogaster) using voluntary ethanol consumption as a measure for changes in reward states.

A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

We describe a protocol for measuring ethanol self-administration in fruit flies (Drosophila melanogaster) as a proxy for changes in reward states. We ...

a-simple-way-to-measure-alterations-reward-seeking-behavior-using

Research

JoVE Journal

Neuroscience

8.3K Views.  Bar-Ilan University. We describe a protocol for inducing rewarding and nonrewarding experiences in fruit flies (Drosophila melanogaster) using voluntary ethanol consumption as a measure for changes in reward states.

Video: A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

Magnetic Resonance Spectroscopy of live Drosophila melanogaster using Magic Angle Spinning

High-Resolution Magic Angle Spinning (HRMAS) proton magnetic resonance spectroscopy (1H-MRS) is a novel non-destructive technique that improves spectral line-widths and allows high-resolution spectra to be obtained from extracts, intact cells, cell cultures, and more importantly intact tissue to investigate relationships between metabolites and cellular processes. In vivo HRMAS 1H-MRS studies have yet to be reported in the live fruit fly Drosophila melanogaster. Drosophila, as a simpler genetic organism, allows the multiple biological functions and various evolutionarily conserved signaling pathways to be examined at the whole organism level and it is a useful model for investigating genetics and physiology. To this end, we developed and implemented an in vivo HRMAS 1H-MRS method to investigate live Drosophila at 14.1 T. Here, we outline an HRMAS 1H-MRS protocol for the molecular characterization of Drosophila with a conventional MR spectrometer equipped with an HRMAS probe. This technique is a novel, in vivo, non-destructive Drosophila metabolite measurement approach, which enables the identification of disease biomarkers and thus may contribute to novel therapeutic development.

Magnetic Resonance Spectroscopy of live Drosophila melanogaster using Magic Angle Spinning

This technique enables the use of high-resolution magic angle spinning proton MR spectroscopy (HRMAS 1H-MRS) for molecular characterization of live Drosophila melanogaster with a conventional 14.1 tesla spectrometer equipped with an HRMAS probe.

High-Resolution Magic Angle Spinning (HRMAS) proton magnetic resonance spectroscopy (1H-MRS) is a novel non-destructive technique that improves spectral ...

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed in sensory hairs called sensilla, which cover the surface of olfactory organs. The surface of each sensillum is covered with tiny pores, through which odorants pass and dissolve in a fluid called sensillum lymph, which bathes the sensory dendrites of the OSNs housed in a given sensillum. The OSN dendrites express odorant receptor (OR) proteins, which in insects function as odor-gated ion channels4, 5. The interaction of odorants with ORs either increases or decreases the basal firing rate of the OSN. This neuronal activity in the form of action potentials embodies the first representation of the quality, intensity, and temporal characteristics of the odorant6, 7.
Given the easy access to these sensory hairs, it is possible to perform extracellular recordings from single OSNs by introducing a recording electrode into the sensillum lymph, while the reference electrode is placed in the lymph of the eye or body of the insect. In Drosophila, sensilla house between one and four OSNs, but each OSN typically displays a characteristic spike amplitude. Spike sorting techniques make it possible to assign spiking responses to individual OSNs. This single sensillum recording (SSR) technique monitors the difference in potential between the sensillum lymph and the reference electrode as electrical spikes that are generated by the receptor activity on OSNs1, 2, 8. Changes in the number of spikes in response to the odorant represent the cellular basis of odor coding in insects. Here, we describe the preparation method currently used in our lab to perform SSR on Drosophila melanogaster and Anopheles gambiae, and show representative traces induced by the odorants in a sensillum-specific manner.

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Electrophysiological responses of olfactory sensory neurons to odorants can be measured in insects using single sensillum recordings. In this video article we will demonstrate how to perform single sensillum recordings in the antennae of the vinegar fly (Drosophila melanogaster) and the maxillary palps of the malaria mosquito (Anopheles gambiae).

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed ...

A Simple Way to Measure Ethanol Sensitivity in Flies

Low doses of ethanol cause flies to become hyperactive, while high doses are sedating. The sensitivity to ethanol-induced sedation of a given fly strain is correlated with that strain s ethanol preference, and therefore sedation is a highly relevant measure to study the genetics of alcohol responses and drinking. We demonstrate a simple way to expose flies to ethanol and measure its intoxicating effects. The assay we describe can determine acute sensitivity, as well as ethanol tolerance induced by repeat exposure. It does not require a technically involved setup, and can therefore be applied in any laboratory with basic fly culture tools.

A simple assay to measure the sedating effects of ethanol on Drosophila flies, based on the loss of righting reflex, is described.

Low doses of ethanol cause flies to become hyperactive, while high doses are sedating. The sensitivity to ethanol-induced sedation of a given fly strain ...

Methods to Assay Drosophila Behavior

Drosophila melanogaster, the fruit fly, has been used to study molecular mechanisms of a wide range of human diseases such as cancer, cardiovascular disease and various neurological diseases1. We have optimized simple and robust behavioral assays for determining larval locomotion, adult climbing ability (RING assay), and courtship behaviors of Drosophila. These behavioral assays are widely applicable for studying the role of genetic and environmental factors on fly behavior. Larval crawling ability can be reliably used for determining early stage changes in the crawling abilities of Drosophila larvae and also for examining effect of drugs or human disease genes (in transgenic flies) on their locomotion. The larval crawling assay becomes more applicable if expression or abolition of a gene causes lethality in pupal or adult stages, as these flies do not survive to adulthood where they otherwise could be assessed. This basic assay can also be used in conjunction with bright light or stress to examine additional behavioral responses in Drosophila larvae. Courtship behavior has been widely used to investigate genetic basis of sexual behavior, and can also be used to examine activity and coordination, as well as learning and memory. Drosophila courtship behavior involves the exchange of various sensory stimuli including visual, auditory, and chemosensory signals between males and females that lead to a complex series of well characterized motor behaviors culminating in successful copulation. Traditional adult climbing assays (negative geotaxis) are tedious, labor intensive, and time consuming, with significant variation between different trials2-4. The rapid iterative negative geotaxis (RING) assay5 has many advantages over more widely employed protocols, providing a reproducible, sensitive, and high throughput approach to quantify adult locomotor and negative geotaxis behaviors. In the RING assay, several genotypes or drug treatments can be tested simultaneously using large number of animals, with the high-throughput approach making it more amenable for screening experiments.

Methods to Assay Drosophila Behavior

Drosophila melanogaster is a genetically and behaviorally tractable model system that has been used to understand the molecular and cellular basis of many important biological processes for over a century 1. Drosophila has been well exploited to gain insights into the genetic basis of fly behavior.

Drosophila melanogaster, the fruit fly, has been used to study molecular mechanisms of a wide range of human diseases such as cancer, cardiovascular ...

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

A growing number of genetically encoded tools are becoming available that allow non-invasive manipulation of the neural activity of specific neurons in Drosophila melanogaster1. Chief among these are optogenetic tools, which enable the activation or silencing of specific neurons in the intact and freely moving animal using bright light. Channelrhodopsin (ChR2) is a light-activated cation channel that, when activated by blue light, causes depolarization of neurons that express it. ChR2 has been effective for identifying neurons critical for specific behaviors, such as CO2 avoidance, proboscis extension and giant-fiber mediated startle response2-4. However, as the intense light sources used to stimulate ChR2 also stimulate photoreceptors, these optogenetic techniques have not previously been used in the visual system. Here, we combine an optogenetic approach with a mutation that impairs phototransduction to demonstrate that activation of a cluster of loom-sensitive neurons in the fly's optic lobe, Foma-1 neurons, can drive an escape behavior used to avoid collision. We used a null allele of a critical component of the phototransduction cascade, phospholipase C-&beta;, encoded by the norpA gene, to render the flies blind and also use the Gal4-UAS transcriptional activator system to drive expression of ChR2 in the Foma-1 neurons. Individual flies are placed on a small platform surrounded by blue LEDs. When the LEDs are illuminated, the flies quickly take-off into flight, in a manner similar to visually driven loom-escape behavior. We believe that this technique can be easily adapted to examine other behaviors in freely moving flies.

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

Genetically encoded optogenetic tools enable noninvasive manipulation of specific neurons in the Drosophila brain. Such tools can identify neurons whose activation is sufficient to elicit or suppress particular behaviors. Here we present a method for activating Channelrhodopsin2 that is expressed in targeted neurons in freely walking flies.

A  growing number of genetically encoded tools are becoming available that allow non-invasive  manipulation of the neural activity of specific neurons in ...

A Single-fly Assay for Foraging Behavior in Drosophila

For many animals, hunger promotes changes in the olfactory system in a manner that facilitates the search for appropriate food sources. In this video article, we describe an automated assay to measure the effect of hunger or satiety on olfactory dependent food search behavior in the adult fruit fly Drosophila melanogaster. In a light-tight box illuminated by red light that is invisible to fruit flies, a camera linked to custom data acquisition software monitors the position of six flies simultaneously. Each fly is confined to walk in individual arenas containing a food odor at the center. The testing arenas rest on a porous floor that functions to prevent odor accumulation. Latency to locate the odor source, a metric that reflects olfactory sensitivity under different physiological states, is determined by software analysis. Here, we discuss the critical mechanics of running this behavioral paradigm and cover specific issues regarding fly loading, odor contamination, assay temperature, data quality, and statistical analysis.

A Single-fly Assay for Foraging Behavior in Drosophila

In this video article, we describe an automated assay to measure the effect of hunger or satiety on olfactory dependent food search behavior in the adult fruit fly Drosophila melanogaster.

For many animals, hunger promotes changes in the olfactory system in a manner that facilitates the search for appropriate food sources. In this video ...

Conditions Affecting Social Space in Drosophila melanogaster

The social space assay described here can be used to quantify social interactions of Drosophila melanogaster &#8212; or other small insects &#8212; in a straightforward manner. As we previously demonstrated 1, in a two-dimensional chamber, we first force the flies to form a tight group, subsequently allowing them to take their preferred distance from each other. After the flies have settled, we measure the distance to the closest neighbor (or social space), processing a static picture with free online software (ImageJ). The analysis of the distance to the closest neighbor allows researchers to determine the effects of genetic and environmental factors on social interaction, while controlling for potential confounding factors. Diverse factors such as climbing ability, time of day, sex, and number of flies, can modify social spacing of flies. We thus propose a series of experimental controls to mitigate these confounding effects. This assay can be used for at least two purposes. First, researchers can determine how their favorite environmental shift (such as isolation, temperature, stress or toxins) will impact social spacing 1,2. Second, researchers can dissect the genetic and neural underpinnings of this basic form of social behavior 1,3. Specifically, we used it as a diagnostic tool to study the role of orthologous genes thought to be involved in social behavior in other organisms, such as candidate genes for autism in humans 4.

Conditions Affecting Social Space in Drosophila melanogaster

The effect of genes and environment on social space of Drosophila melanogaster can be quantified through a powerful but straightforward analytical paradigm. We show here different factors that can affect this social space, and thus need to be taken into consideration when designing experiments in this paradigm.

The social space assay described here can be used to quantify social interactions of Drosophila melanogaster &#8212; or other small insects &#8212; in a ...

High-resolution Quantification of Odor-guided Behavior in Drosophila melanogaster Using the Flywalk Paradigm

In their natural environment, insects such as the vinegar fly Drosophila melanogaster are bombarded with a huge amount of chemically distinct odorants. To complicate matters even further, the odors detected by the insect nervous system usually are not single compounds but mixtures whose composition and concentration ratios vary. This leads to an almost infinite amount of different olfactory stimuli which have to be evaluated by the nervous system.
To understand which aspects of an odor stimulus determine its evaluation by the fly, it is therefore desirable to efficiently examine odor-guided behavior towards many odorants and odor mixtures. To directly correlate behavior to neuronal activity, behavior should be quantified in a comparable time frame and under identical stimulus conditions as in neurophysiological experiments. However, many currently used olfactory bioassays in Drosophila neuroethology are rather specialized either towards efficiency or towards resolution.
Flywalk, an automated odor delivery and tracking system, bridges the gap between efficiency and resolution. It allows the determination of exactly when an odor packet stimulated a freely walking fly, and to determine the animal&#180;s dynamic behavioral reaction.

High-resolution Quantification of Odor-guided Behavior in Drosophila melanogaster Using the Flywalk Paradigm

The automated tracking system Flywalk is used for high-resolution quantification of odor-guided behavior in Drosophila melanogaster.

In their natural environment, insects such as the vinegar fly Drosophila melanogaster are bombarded with a huge amount of chemically distinct odorants. To ...

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat open surfaces is complex,&#160;including: pausing, turning, and meandering. This complexity in the repertoire of movement hinders detailed analysis of the events occurring during a single crawl stride cycle. To overcome this obstacle, linear agarose channels were made that constrain larval behavior to straight, sustained, rhythmic crawling. In principle, because agarose channels and the Drosophila larval body are both optically clear, the movement of larval structures labeled by genetically-encoded fluorescent probes can be monitored in intact, freely-moving larvae. In the past, larvae were placed in linear channels and crawling at the level of whole organism, segment, and muscle were analyzed1. In the future, larvae crawling in channels can be used for calcium imaging to monitor neuronal activity. Moreover, these methods can be used with larvae of any genotype and with any researcher-designed channel. Thus the protocol presented below is widely applicable for studies using the Drosophila larva as a model to understand motor control.

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

The Drosophila larva is a powerful model system to study neural control of behavior. This publication describes the use of linear agarose channels to elicit sustained bouts of linear crawling and methods to quantify the dynamics of larval structures during repetitive crawling behavior.

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat ...

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a considerable amount of information has been generated regarding ORN responses to odorants, the role of specific ORNs in driving behavioral responses remains poorly understood. Complications in behavior analyses arise due to different volatilities of odorants that activate individual ORNs, multiple ORNs activated by single odorants, and the difficulty in replicating naturally observed temporal variations in olfactory stimuli using conventional odor-delivery methods in the laboratory. Here, we describe a protocol that analyzes Drosophila larval behavior in response to simultaneous optogenetic stimulation of its ORNs. The optogenetic technology used here allows for specificity of ORN activation and precise control of temporal patterns of ORN activation. Corresponding larval movement is tracked, digitally recorded, and analyzed using custom written software. By replacing odor stimuli with light stimuli, this method allows for a more precise control of individual ORN activation in order to study its impact on larval behavior. Our method could be further extended to study the impact of second-order projection neurons (PNs) as well as local neurons (LNs) on larval behavior. This method will thus enable a comprehensive dissection of olfactory circuit function and complement studies on how olfactory neuron activities translate in to behavior responses.

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

This protocol analyzes navigational behavior of Drosophila larva in response to simultaneous optogenetic stimulation of its olfactory neurons. Light of 630 nm wavelength is used to activate individual olfactory neurons expressing a red-shifted channel rhodopsin. Larval movement is simultaneously tracked, digitally recorded, and analyzed using custom-written software.