These studies were supported by grants from the National Institutes of Health, National Institute for Alcoholism and Alcohol Abuse to M.G. (P20AA017828, R01AA020634, P50 AA022537). The authors thank Jill Bettinger for helpful discussions and Jacqueline DeLoyht for technical assistance.

1. Day Before Assay
<ol>
	<li>Collect flies into fresh food vials in groups of 11 (single sex) under brief (1-5 min) CO2.</li>
	<li>Allow flies to recover O/N in food vials in an environmentally controlled space (typically 25 &#176;C, 60% relative humidity, 12 hr light/dark cycle).</li>
	<li>Prepare ethanol solution(s) by diluting pure (100%) ethanol in purified (&#8805;18 M&#937;) water to final concentration(s) appropriate for the planned experiment. Allow solution(s) to return to RT O/N. 
	Note: Dilution of ethanol is exothermic.</li>
</ol>
2. Day of Assay
<ol>
	<li>For each vial of flies to be tested, prepare (i) a clean, empty food vial; i.e., testing vial, (ii) a new cellulose acetate plug, (iii) a silicone stopper and (iv) 1 ml of ethanol solution (see Table 3).</li>
	<li>Prepare testing room by adjusting temperature to 20-25 &#176;C and relative humidity to 55-65%.</li>
	<li>Have another worker assign a unique code to each group of vials and record the code for later. Place coded vials with flies in testing room to acclimate for a few min.</li>
	<li>Label empty testing vials to match codes on fly vials from 2.3.</li>
	<li>Construct a hard copy testing log by entering the codes into columns (one column/vial) in a spreadsheet similar to Table 1.</li>
	<li>Using the testing log as a guide, arrange coded food vials with flies and empty testing vials into matching arrays in the testing room. 
	Note: A reasonable maximum number of vials to test is 24 (i.e.,&#160;6 sets of 4 vials each).</li>
	<li>Transfer flies from food vials into matched/labeled empty testing vials and immediately insert cellulose acetate plugs into testing vials until cellulose acetate plugs are 2 cm below the vial tops.</li>
	<li>Hereafter, handle each row of four vials as a set at staggered 1 min&#160;intervals.</li>
	<li>For the time 0 assessment, grasp each vial individually with thumb and forefinger, tap gently on the table three times to knock flies to the bottom of the vial, wait 30 sec and then count the number of flies that are immobile/dead. Record the number of immobile/dead flies for each vial at time 0 min in the hardcopy testing log.</li>
	<li>Start timer counting up continuously at time 0 and immediately begin adding 1 ml of ethanol to cellulose acetate plugs in the vials for the first row/set of 4 vials. Add 1 ml of ethanol to the cellulose acetate plugs in the vials at 5 sec intervals in the order they will be tested. Add ethanol to the cellulose acetate plugs in a circular motion so that the ethanol is absorbed evenly throughout the cellulose acetate plugs. When ethanol has been added to all 4 testing vials in the set, insert a silicone plug in each vial to seal it.</li>
	<li>At times 1, 2, 3, 4 and 5 min, add 1 ml of ethanol to the second, third, fourth, fifth and sixth sets of 4 vials, respectively. Continue inserting silicone stoppers after adding ethanol to each set of 4 vials.</li>
	<li>At time 6 min, test the first set of 4 vials by grasping each vial with thumb and forefinger, tapping gently on the table three times to knock flies to the bottom of the vial, waiting 30 sec and then counting and recording the total number of flies that are sedated. Score flies as sedated if they (i) stand on the floor of the vial but do not walk or (ii) lie on their backs with or without flapping their wings.</li>
	<li>Handle each vial within the set at 5 sec intervals using the schedule in Table 2.</li>
	<li>At times 7, 8, 9, 10 and 11 min, test the second, third, fourth, fifth and sixth sets of vials, respectively, as done for the first set.</li>
	<li>At time 12 min, test the first set of 4 vials again as described in 2.12 and continue testing the second, third, fourth, fifth and sixth sets of vials at 13, 14, 15, 16 and 17 min, respectively.</li>
	<li>Continue testing flies as described in 2.12 until all flies are sedated.</li>
	<li>Enter the total number of flies in each vial in the hard copy testing log. Censor immobile/dead flies at time 0 from the total number of flies.</li>
	<li>Calculate the percent active flies at each assessment time-point and plot data as % active flies (y-axis) vs. time (x-axis). Quantitate ethanol sedation by interpolating Sedation Time 50 values (ST50, time to 50% sedation) from third-order polynomial or sigmoidal curve fits or calculating area under the curve.</li>
	<li>Compile data from decoded vials and perform statistical analyses (e.g.,&#160;one-way ANOVA with Bonferroni multiple comparison test) as appropriate for the experimental design.</li>
</ol>

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The authors have nothing to disclose.

Straightforward assays that reproducibly quantitate meaningful phenotypes are of great value for the analysis of behavior. The work described here addresses several practical aspects of an assay for measuring ethanol sedation sensitivity and rapid tolerance in Drosophila. Although not a focus of this work, behavioral analyses are facilitated by maintaining the environment and genetic background constant for test subjects within a study. Furthermore, comparisons should typically be made between groups of flies reared and tested side-by-side. To this end, all flies within individual experiments in this work had the same genetic background and were reared side by side under uniform environmental conditions (25 &#176;C, 60% relative humidity, 12 hr light/dark cycle) with standard food medium (10% sucrose, 3.3% cornmeal, 2% yeast and 1% agar) except for those experiments in which the food medium was explicitly manipulated.
The ethanol sedation assay described here requires very few materials (plastic vials, cellulose acetate plugs, silicone stoppers and ethanol), all of which are inexpensive and readily available from commercial sources (Table 3). This ethanol sedation assay is similar to and to a large extent based on previously reported methods (e.g., 6,20,25-27, comprehensively reviewed in 3,28). Although all of these assays have utility for measuring ethanol sedation sensitivity, advantages of the assay described here include (i) flies are not exposed to liquid ethanol during the assay, (ii) the reduced possibility that flies can become entangled in cotton or other material in the testing vial during the assay, (iii) the ability to test many groups of flies in parallel, and (iv) three options for the objective quantification of ethanol sensitivity from primary data sets. Together, these advantages largely eliminate the possibility that flies drink ethanol, that ethanol could wet the external surfaces of flies thereby inhibiting their overall locomotor abilities, and that performance of flies in the assay could be confounded by difficulties associated with entangling material in the testing environment. Additionally, these advantages increase the overall throughput and reproducibility of the assay. Furthermore, ethanol sensitivity measured in this assay is not influenced by expression of endogenous white or the transgenic marker mini-white18, making it suitable for studies with transgenes and genetic backgrounds commonly used in Drosophila genetic analyses.
Several key parameters should be controlled to obtain reliable results from ethanol sedation assays. In addition to controlling the growth environment and genetic background of strains being tested (see above), the volume and concentration of ethanol are of course extremely important to the quantitation of ethanol sensitivity. Note that the dilution of ethanol in water is exothermic. Consequently, diluted ethanol solutions should be allowed to equilibrate to RT prior to the initiation of sedation assays. An additional parameter that should be made uniform across the groups being tested is the length of time that flies are housed in empty food vials prior to starting the sedation assay. Two additional parameters should also be controlled. The number of flies per vial should ideally be (a) the same in all vials and groups, (b)
a number that can be easily counted quickly, and (c) a number large enough to allow for relatively smooth sedation time-courses. Eleven flies/vial works well in our laboratory and is a suggested starting point. A final parameter to consider is the age of the flies used in sedation assays. Although no reproducible effects of age on ethanol sedation sensitivity in 1-10 day old flies has been found (data not shown), the use of young age-matched animals seems justified given the large literature on age-related behavioral changes in flies 29,30.
Other parameters do not seem as critical in sedation assays, at least when testing control flies using the ranges of parameters described here. Sex, strength of tapping, recovery time from tapping, and supplementation of fly food with antibiotics (ampicillin, tetracycline and chloramphenicol) or live yeast do not alter ethanol sensitivity measured as described here. Similarly, quantitating ethanol sensitivity by interpolation from third-order polynomial curves, interpolation from sigmoidal curves, or determination of AUC from the primary sedation time-course data leads to essentially indistinguishable interpretations. Although no effect of sex was consistently observed in the studies described here, effects of sex on ethanol sensitivity in several Drosophila genetic backgrounds have been reported 31. Thus, it is possible that sex effects were simply masked by genetic variance in the w[A] background used here. Alternatively, it is possible that measuring sex effects on behavioral responses in Drosophila require as yet unidentified assay or growth conditions. In any case, the recommendation is to make all parameters as uniform as possible in sedation assays, including the sex of any groups being explicitly compared.
Given that live yeast produce sizeable amounts of ethanol in fly food, it was somewhat surprising that supplementation of food with live yeast did not increase internal ethanol in flies. One possible explanation is that the ethanol is not produced uniformly across the surface of the food and flies might therefore selectively ingest food with no or relatively low concentrations of ethanol. Additional studies will be required to address this and other possible explanations for this finding.
A large number of modifications to the assay described here are possible depending on the goals of the experiment being performed. For example, the number of vials tested in an experiment, the number of flies tested in each vial, the concentration and volume of ethanol used, the duration of ethanol exposure, the time interval between sedation assessments, the age and sex of the flies tested, and the criteria for sedation can all be modified to suit the needs of an individual laboratory. The recommendation is to start with one or two groups of four vials when initially learning the assay and then scale up to using the number of vials that provides the throughput required for the project. Should unexpectedly short or long ST50s be observed, it would be good practice to ensure that test flies were subjected to short (1-5 min) anesthesia times, were allowed to recover from anesthesia O/N, were less than 10 days old, were not physically damaged during handling, were not wetted by ethanol solution, and did not have locomotor impairments.
The assay measures ethanol sedation sensitivity only and therefore is not designed or suited for assessing other behavioral responses to ethanol. Like all known ethanol behavioral paradigms in flies, this assay requires flies to have largely normal locomotor abilities and therefore genotypes with impaired locomotion should not be tested. Another limitation of the assay is that the concentration of ethanol vapor in the vial is presumably rising continuously as the drug volatilizes from the cellulose acetate plug. Although the flies&#8217; internal ethanol rises progressively with time in all fly ethanol behavioral assays, it seems possible that delivering a fixed concentration of ethanol vapor to flies could improve the consistency of results from this assay. A method for delivering a fixed concentration of ethanol vapor has not yet been identified that would allow the assay to be used at the scale described here.
The ethanol sedation assay described here is well-suited for genetic analysis of ethanol sensitivity and rapid tolerance 18. Growth environment, genetic background, the concentration and volume of ethanol, the amount of time flies spend in empty food vials, and the number and age of flies used should all be controlled. Assuming these parameters are adequately controlled, the sedation assay should be suitable for both reverse and forward genetic approaches that investigate the molecular basis for behavioral responses to ethanol in Drosophila.

Alcohol use disorder (AUD) is an enormous health problem worldwide (reviewed in 1). Although the mechanisms driving the development of AUD are complex, these disorders have a major genetic component (e.g., 2). The large heritability of AUD&#160;and the conserved behavioral responses to ethanol across many species (reviewed in 3,4) have generated strong interest in using genetic model organisms to investigate the involvement of specific genes in ethanol-related behaviors toward better understanding the molecular basis of AUD. The fruit fly, Drosophila melanogaster, has emerged as a leading model organism for exploring molecular-genetic mechanisms of ethanol-related behaviors (reviewed in 3,4). Studies in flies have highlighted roles for several signaling pathways in behavioral responses to ethanol (reviewed in 5). Intriguingly, some of the genes and pathways that influence behavioral responses to ethanol in flies have also been implicated in rodent ethanol-related behaviors and/or human AUD&#160;(e.g.,&#160;6-14). The conservation of mechanisms driving ethanol-related behaviors across species, coupled with the suite of genetic tools available in the Drosophila model system, underscore the utility of the fruit fly model for investigating the genetics of behavioral responses to ethanol.
Sensitivity 15,16 and tolerance (reviewed in 17) to ethanol in humans is linked to the development of AUD. Both of these behavioral responses to ethanol can be modeled in flies via a variety of laboratory assays (reviewed in 3,4). All of the fly assays known to the authors are based on either time-dependent ethanol-induced sedation/incoordination or time-dependent recovery from ethanol sedation.
In a previous article from our group on the genetics of ethanol sensitivity and rapid tolerance in Drosophila, a behavioral assay based on ethanol vapor-induced sedation of flies was used 18. Testing in this assay was initiated by transferring live adult flies without anesthesia to empty food vials, trapping the flies in the vials with a cellulose acetate plug, adding ethanol to the top (i.e.,&#160;non-fly side) of the cellulose acetate plug, and sealing the vial containing flies, cellulose acetate plug and ethanol with a silicone stopper (see schematic in Figure S3, reference 18). Multiple vials representing different groups of flies were assessed in parallel, increasing throughput of this assay. Vials were given an anonymous code and experimenters were blinded to treatment group to prevent unintended bias in the assessment of sedation. In a standard experiment, flies in vials were tapped gently at 6 min intervals and, after a 30 sec recovery, the number of sedated flies in each vial was counted and converted to percent active flies. Flies absorbed ethanol vapor from the cellulose acetate plug in a time-dependent fashion, causing progressive increases in internal ethanol18 and sedation (c.f. reference 18 and Figure 1A and 1B in this report). Sedation in this assay was operationally defined as flies (i) standing in the absence of walking or (ii) lying on their backs with or without flapping their wings. Here, this ethanol sedation assay is described in detail, further operational optimization relevant to using it is provided, and the assay is used to address the contribution of food supplementation options on fly sedation sensitivity.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>food vials</td>
			<td>VWR</td>
			<td>89092-772</td>
			<td>narrow</td>
		</tr>
		<tr>
			<td>Flugs</td>
			<td>Genesee/flystuff.com</td>
			<td>49-102</td>
			<td>narrow</td>
		</tr>
		<tr>
			<td>silicone stopper</td>
			<td>Fisher Scientific</td>
			<td>09-704-1l</td>
			<td>#4</td>
		</tr>
		<tr>
			<td>ethanol</td>
			<td>Pharmaco-Aaper</td>
			<td>111000200</td>
			<td>200 proof</td>
		</tr>
	</tbody>
</table>

an inexpensive, scalable behavioral assay for measuring ethanol sedation sensitivity and rapid tolerance in drosophila

Alcohol use disorder (AUD) is a serious health challenge. Despite a large hereditary component to AUD, few genes have been unambiguously implicated in their etiology. The fruit fly, Drosophila melanogaster, is a powerful model for exploring molecular-genetic mechanisms underlying alcohol-related behaviors and therefore holds great promise for identifying and understanding the function of genes that influence AUD. The use of the Drosophila model for these types of studies depends on the availability of assays that reliably measure behavioral responses to ethanol. This report describes an assay suitable for assessing ethanol sensitivity and rapid tolerance in flies. Ethanol sensitivity measured in this assay is influenced by the volume and concentration of ethanol used, a variety of previously reported genetic manipulations, and also the length of time the flies are housed without food immediately prior to testing. In contrast, ethanol sensitivity measured in this assay is not affected by the vigor of fly handling, sex of the flies, and supplementation of growth medium with antibiotics or live yeast. Three different methods for quantitating ethanol sensitivity are described, all leading to essentially indistinguishable ethanol sensitivity results. The scalable nature of this assay, combined with its overall simplicity to set-up and relatively low expense, make it suitable for small and large scale genetic analysis of ethanol sensitivity and rapid tolerance in Drosophila.

The raw data from this ethanol sedation assay are the numbers of flies that are sedated as a function of ethanol vapor exposure time. Raw data are converted to the percent active flies as a function of time (primary data, Figures 1A, B, D-F). Sensitivity to ethanol sedation from the primary data can be quantitated as Sedation Time 50 (ST50), the time required for 50% of flies to become sedated or aea under the curve (AUC), via interpolation from curve fits. Previously reported18 third-order polynomial curves fit the primary data well as expected (average R2 = 0.934, n = 24, Figure 1A), although sigmoidal curves fit the primary data somewhat better (average R2 = 0.997, n = 24; Figure 1B). As an alternative to determining ST50 values from curve fits, ethanol sedation sensitivity from the primary data can also be quantitated as area under the curve (AUC, % active flies x time) (Figure 1C). The results from these three methods of quantitation correlate extremely well with each other (Pearson r2 &#8805;0.998, n = 24) and are essentially indistinguishable (Figure 1C), although the units are necessarily different for ST50 (min) and AUC (% active flies x time). Details regarding all calculations, spreadsheets used, etc. are available from the corresponding author. Note that higher and lower values of ST50 (or AUC) indicate blunted and enhanced sensitivity to ethanol sedation, respectively. For consistency with the previous report18, ST50 values derived from third-order polynomial curve fits were used throughout the remainder of this study.Importantly, the assay detects the effects of RNAi against Cnx14D (Figure 1D) and mutations in scb (Figure 1E), aru and hppy (Figure 1F), Clic and NPFR1 (data not shown) on ethanol sensitivity as reported in 18 and the articles originally describing the effects of these genetic manipulations 7,19-22.
During routine use of this assay, tapping of vials with cellulose acetate plugs wetted with 2 ml of ethanol caused the cellulose acetate plugs to travel toward the bottoms of some vials in a few individual experiments (not shown), thereby potentially changing the flies&#8217; free space and the concentration of ethanol vapor. Additionally, approximately 0.25 ml of ethanol reached the bottom (i.e.,&#160;fly side) of the cellulose acetate plugs when they were wetted with 2 ml of ethanol (Figure 2A), raising the possibility that flies in these assays were exposed to ethanol liquid in addition to ethanol vapor. Decreasing the volume of ethanol in the cellulose acetate plug from 2 ml to 1 ml eliminated the downward migration of the cellulose acetate plugs during the assay even after vigorous tapping (not shown) and also eliminated measureable amounts of ethanol solution on the bottoms of cellulose acetate plugs (Figure 2A). Sedation of flies was sensitive to the volume of ethanol used even with volumes less than 1 ml (Figure 1A-C). Using 1 ml of ethanol therefore helps maintain a constant open space for flies during the assay and ensures that liquid ethanol is not being ingested. Flies are therefore presumably exposed to ethanol solely as a vapor in this assay. One ml of 85% ethanol (i.e.,&#160;vapor from 1 ml of 85% (vol/vol) ethanol) and 1-5 day old w[A] flies were used in all subsequent studies reported here unless indicated otherwise.
In experiments with 24 vials, flies are tapped gently at 6 min intervals to allow the experimenter to assess (i.e.,&#160;cycle through) all vials on a reasonable, prescribed schedule. ST50s from studies with control flies using fewer vials and 5 min intervals were comparable to ST50s from studies with 6 min intervals (data not shown). Flies are assessed for sedation 30 sec after tapping to allow the experimenter time to tap a set of four vials in sequence prior to determining the sedation status of the flies in each vial. To determine if differences in tapping vigor or recovery time after tapping impact ethanol sedation sensitivity, ST50 values in male and female control flies were measured while being tapped normally (Regular) or very sharply (Hard) and after a standard 30 sec or longer 60 sec recovery. Tapping vigor (Figure 2B) and recovery time (Figure 2C) had no measurable effect on ST50 values in control males or females.
Antibiotics can be used to suppress bacterial growth in fly food medium. To determine if supplementation of the food medium with antibiotics affects ethanol sedation sensitivity, ST50 values were measured in flies reared for one generation in the presence or absence of three antibiotics (ampicillin, 100 &#181;g/ml; tetracycline, 20 &#181;g/ml; chloramphenicol, 125 &#181;g/ml). Rearing flies on these three antibiotics had no effect on ethanol sedation sensitivity (Figure 2D).
Flies are transferred to empty food vials&#8212;and are therefore deprived of food and water for ~5 min&#8212;immediately prior to being exposed to ethanol vapor in the sedation assay. To determine if the amount of time flies are kept in empty food vials before initiating the assay impacts ethanol sedation sensitivity, flies were starved for 6 hr (at least 70-fold of normal) and then ST50 values were measured. Starvation for 6 hr significantly decreased ST50s in both male and female control flies (Figure 2E). Although the effect of starvation on ST50 was relatively small (9-14% in these experiments), the amount of time flies spend in empty vials prior to being exposed to ethanol vapor in the assay should be held uniform across groups within an experiment and also between experiments.
Laboratory fly food can be supplemented with live yeast (e.g., 18,20,23,24) to promote production of progeny. For example, food vials supplemented with live yeast (Saccharomyces cerevisiae) produced adults earlier than those with heat-killed yeast or no live yeast (compare d10-d11, white bars, Figure 3A), although the total number of progeny produced over 20 days was indistinguishable in vials supplemented with live and heat-killed yeast (Figure 3A). To address the possibility that live yeast produce meaningful amounts of ethanol on fly food, the effects of live yeast on (i) ethanol in the food medium, (ii) ethanol in flies, and (iii) ethanol sedation sensitivity in flies were measured. Food medium supplemented with live yeast contained substantial amounts of ethanol compared to food supplemented with heat-killed or no yeast (Figure 3B). Nevertheless, control flies grown on medium supplemented with live, killed or no yeast had indistinguishably low concentrations of internal ethanol (Figure 3C). Furthermore, ethanol sedation sensitivity was indistinguishable in control flies grown on medium supplemented with heat-killed or live yeast (Figure 3D).
<img alt="Figure 1" src="/files/ftp_upload/52676/52676fig1.jpg" /> 
Figure 1.&#160;(A-C) Sedation assay data analysis options. Control w[A] females (w1118 isogenic, Bloomington Indiana Drosophila Stock Center stock #5905) were exposed to vapor from the indicated volumes of 85% (vol/vol) ethanol. (A) Sedation time-course data fit with third-order polynomials. (B) Sedation time-course data fit with sigmoidal curves. (C) Ethanol sedation sensitivity quantitated as (left Y-axis) ST50 values interpolated from third-order polynomials and sigmoidal curves or (right Y-axis) area under the curve (AUC, % active flies x time). Ethanol volume, but not analysis method, impacted ethanol sensitivity (two-way ANOVA; effect of volume, p &#60;0.0001; effect of analysis method, n.s.; n = 8/group; AUC data transformed 1/100 to account for the difference in magnitude of the values). (D-F) Representative time-course data from. (D) Expression of Cnx14D RNAi in the nervous system (elav-Gal4/v5597) blunted ethanol sensitivity relative to controls (v5597/+ and elav-Gal4/+). (E and F) Mutation of scb (scbVol2) and aru (aru8.128) enhanced ethanol sedation sensitivity and mutation of hppy (hppyKG5537) blunted ethanol sedation sensitivity compared to controls. Data in E-F were previously reported as ST50 values 18.
<img alt="Figure 2" src="/files/ftp_upload/52676/52676fig2.jpg" /> 
Figure 2.&#160;Assessment of operational parameters in the sedation assay. (A) Amount of ethanol in the bottom 2 mm of cellulose acetate plugs. The volume of ethanol added to the tops of cellulose acetate plugs had a significant effect on the volume of ethanol that moved into the bottom 2 mm of cellulose acetate plugs during a mock 60 min experiment (one-way ANOVA, p &#60;0.0001; *Bonferroni multiple comparison test, 2 ml vs 0 and 1 ml, p &#60;0.05 n = 4). Ethanol was quantitated as a change in mass of the cellulose acetate plugs. (B) Neither vigor of tapping the vials during testing (Regular, Hard) nor sex of the flies tested affected ST50s (two-way ANOVA; effect of tapping, n.s.; effect of sex, n.s.; n = 12). (C) Neither recovery time after tapping nor sex had significant effects on ST50s (two-way ANOVA; effect of recovery time, n.s.; effect of sex, n.s.; n = 6). (D) Inclusion of antibiotics (ATC; ampicillin, tetracycline and chloramphenicol) in the growth medium had no overall effect on ST50s, but there was an effect of sex on ST50 (two-way ANOVA; effect of ATC, n.s.; effect of sex, p = 0.001; interaction, n.s.; n = 6). (E) Effect of starvation on ethanol sedation sensitivity. ST50s were significantly lower in flies deprived of food and water for 6 hr (Starved) compared to normally fed (Fed) flies; ST50s in males and females were indistinguishable overall (two-way ANOVA; effect of starvation, p &#60;0.0001; effect of sex, n.s.; n = 12; *Bonferroni multiple comparison tests, effect of starvation in males and females, p &#60;0.05).
<img alt="Figure 3" src="/files/ftp_upload/52676/52676fig3.jpg" /> 
Figure 3.&#160;Effect of live yeast on growth, ethanol content and sedation sensitivity. (A) Adult progeny from vials containing food medium supplemented with no yeast (No Y), heat killed yeast (Killed Y) or live yeast (Live Y). White bars, adult progeny emerging during days 10-11; grey bars, days 12-20. Overall, yeast treatment had a significant effect on progeny production (one-way ANOVAs; all (10-20) days, p &#60;0.0001; days 10-11, p &#60;0.0001; days 12-20, p = 0.0002; n = 5). Total number of progeny produced during all (10-20) days was greater from vials with Killed Y and Live Y than No Y (Bonferroni multiple comparison test, p&#60;0.05). During days 10-11, vials with Live Y produced more progeny than Killed Y and vials with Killed Y produced more progeny than No Y (white bars, Bonferroni multiple comparison tests, p&#60;0.05). During days 12-20, vials with Killed Y produced more progeny than vials with No Y or Live Y (grey bars, Bonferroni multiple comparison test, p &#60;0.05). (B) Ethanol in the food medium was significantly higher in vials supplemented with Live Y compared to vials with No Y and Killed Y (one-way ANOVA, p &#60;0.0001, n = 5; Bonferroni multiple comparison test, p &#60;0.05). (C) Internal ethanol in w[A] female flies was not significantly affected by supplementation with yeast (one-way ANOVA, n.s., n = 5-10). Ethanol content in B and C determined as described18. (D) ST50 values were not significantly affected by the supplementation of the growth medium with yeast in male or female w[A] (two-way ANOVA; effect of yeast, n.s.; effect of sex, n.s.; n = 12).
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>vial &#8594;</td>
			<td>1</td>
			<td>2</td>
			<td>3</td>
			<td>4</td>
			<td>5</td>
			<td>6</td>
			<td>7</td>
			<td>8</td>
		</tr>
		<tr>
			<td>code &#8594;</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>total # &#8594;</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>0 min</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>6 min</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>12 min</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>18 min</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>24 min</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>30 min</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>36 min</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>42 min</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>48 min</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>54 min</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>60 min</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 1: Typical testing log. See protocol step 2.5.
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Vial</td>
			<td>Tap</td>
			<td>Assess</td>
		</tr>
		<tr>
			<td>1</td>
			<td>6 min 0 sec</td>
			<td>6 min 30 sec</td>
		</tr>
		<tr>
			<td>2</td>
			<td>6 min 5 sec</td>
			<td>6 min 35 sec</td>
		</tr>
		<tr>
			<td>3</td>
			<td>6 min 10 sec</td>
			<td>6 min 40 sec</td>
		</tr>
		<tr>
			<td>4</td>
			<td>6 min 15 sec</td>
			<td>6 min 45 sec</td>
		</tr>
	</tbody>
</table>
Table 2: Typical tapping and testing schedule. See protocol step 2.13.
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Name of Material/ Equipment</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments/Description</td>
		</tr>
		<tr>
			<td>food vials</td>
			<td>VWR</td>
			<td>89092-772</td>
			<td>narrow</td>
		</tr>
		<tr>
			<td>Flugs</td>
			<td>Genesee/flystuff.com</td>
			<td>49-102</td>
			<td>narrow</td>
		</tr>
		<tr>
			<td>silicone stopper</td>
			<td>Fisher Scientific</td>
			<td>09-704-1l</td>
			<td>#4</td>
		</tr>
		<tr>
			<td>ethanol</td>
			<td>Pharmaco-Aaper</td>
			<td>111000200</td>
			<td>200 proof</td>
		</tr>
	</tbody>
</table>
Table 3: Materials.

Watch this Scientific Journal Video about An Inexpensive, Scalable Behavioral Assay for Measuring Ethanol Sedation Sensitivity and Rapid Tolerance in Drosophila at JoVE.com

An Inexpensive, Scalable Behavioral Assay for Measuring Ethanol Sedation Sensitivity and Rapid Tolerance in Drosophila

Straightforward assays for measuring ethanol sensitivity and rapid tolerance in Drosophila facilitate the use of this model organism for investigating these important ethanol-related behaviors. Here, a relatively simple, scalable assay for measuring ethanol sensitivity and rapid tolerance in flies is described.

An Inexpensive, Scalable Behavioral Assay for Measuring Ethanol Sedation Sensitivity and Rapid Tolerance in Drosophila

Alcohol use disorder (AUD) is a serious health challenge. Despite a large hereditary component to AUD, few genes have been unambiguously implicated in ...

an-inexpensive-scalable-behavioral-assay-for-measuring-ethanol

Nanyang Technological University

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JoVE Journal

Neuroscience

10.2K Views.  Virginia Commonwealth University. Straightforward assays for measuring ethanol sensitivity and rapid tolerance in Drosophila facilitate the use of this model organism for investigating these important ethanol-related behaviors. Here, a relatively simple, scalable assay for measuring ethanol sensitivity and rapid tolerance in flies is described.

Video: An Inexpensive, Scalable Behavioral Assay for Measuring Ethanol Sedation Sensitivity and Rapid Tolerance in Drosophila

Combining Behavioral Endocrinology and Experimental Economics: Testosterone and Social Decision Making

Behavioral endocrinological research in humans as well as in animals suggests that testosterone plays a key role in social interactions. Studies in rodents have shown a direct link between testosterone and aggressive behavior1 and folk wisdom adapts these findings to humans, suggesting that testosterone induces antisocial, egoistic or even aggressive behavior2. However, many researchers doubt a direct testosterone-aggression link in humans, arguing instead that testosterone is primarily involved in status-related behavior3,4. As a high status can also be achieved by aggressive and antisocial means it can be difficult to distinguish between anti-social and status seeking behavior.
We therefore set up an experimental environment, in which status can only be achieved by prosocial means. In a double-blind and placebo-controlled experiment, we administered a single sublingual dose of 0.5 mg of testosterone (with a hydroxypropyl-&beta;-cyclodextrin carrier) to 121 women and investigated their social interaction behavior in an economic bargaining paradigm. Real monetary incentives are at stake in this paradigm; every player A receives a certain amount of money and has to make an offer to another player B on how to share the money. If B accepts, she gets what was offered and player A keeps the rest. If B refuses the offer, nobody gets anything. A status seeking player A is expected to avoid being rejected by behaving in a prosocial way, i.e. by making higher offers.
The results show that if expectations about the hormone are controlled for, testosterone administration leads to a significant increase in fair bargaining offers compared to placebo. The role of expectations is reflected in the fact that subjects who report that they believe to have received testosterone make lower offers than those who say they believe that they were treated with a placebo. These findings suggest that the experimental economics approach is sensitive for detecting neurobiological effects as subtle as those achieved by administration of hormones. Moreover, the findings point towards the importance of both psychosocial as well as neuroendocrine factors in determining the influence of testosterone on human social behavior.

The procedure described in this protocol shows that testosterone administration and folk beliefs about testosterone may be associated with directly opposed social behaviors.

Behavioral endocrinological research in humans as well as in animals suggests that testosterone plays a key role in social interactions. Studies in ...

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 ...

A General Method for Evaluating Incubation of Sucrose Craving in Rats

For someone on a food-restricted diet, food craving in response to food-paired cues may serve as a key behavioral transition point between abstinence and relapse to food taking 1. Food craving conceptualized in this way is akin to drug craving in response to drug-paired cues. A rich literature has been developed around understanding the behavioral and neurobiological determinants of drug craving; we and others have been focusing recently on translating techniques from basic addiction research to better understand addiction-like behaviors related to food 2-4.
As done in previous studies of drug craving, we examine sucrose craving behavior by utilizing a rat model of relapse. In this model, rats self-administer either drug or food in sessions over several days. In a session, lever responding delivers the reward along with a tone+light stimulus. Craving behavior is then operationally defined as responding in a subsequent session where the reward is not available. Rats will reliably respond for the tone+light stimulus, likely due to its acquired conditioned reinforcing properties 5. This behavior is sometimes referred to as sucrose seeking or cue reactivity. In the present discussion we will use the term "sucrose craving" to subsume both of these constructs.
In the past decade, we have focused on how the length of time following reward self-administration influences reward craving. Interestingly, rats increase responding for the reward-paired cue over the course of several weeks of a period of forced-abstinence. This "incubation of craving" is observed in rats that have self-administered either food or drugs of abuse 4,6. This time-dependent increase in craving we have identified in the animal model may have great potential relevance to human drug and food addiction behaviors.
Here we present a protocol for assessing incubation of sucrose craving in rats. Variants of the procedure will be indicated where craving is assessed as responding for a discrete sucrose-paired cue following extinction of lever pressing within the sucrose self-administration context (Extinction without cues) or as responding for sucrose-paired cues in a general extinction context (Extinction with cues).

Responding for food or drug-paired cues increases over the course of a period of abstinence and this may relate to an increased susceptibility to relapse behaviors. Here we detail a procedure for evaluating this "incubation of craving" in rats that have self-administered sucrose.

For someone on a food-restricted diet, food craving in response to food-paired cues may serve as a key behavioral transition point between abstinence and ...

Recording and Analysis of Circadian Rhythms in Running-wheel Activity in Rodents

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, including rats, hamsters, and mice, are active during the night and relatively inactive during the day. Many other behavioral and physiological measures also exhibit daily rhythms, but in rodents, running-wheel activity serves as a particularly reliable and convenient measure of the output of the master circadian clock, the suprachiasmatic nucleus (SCN) of the hypothalamus. In general, through a process called entrainment, the daily pattern of running-wheel activity will naturally align with the environmental light-dark cycle (LD cycle; e.g. 12 hr-light:12 hr-dark). However circadian rhythms are endogenously generated patterns in behavior that exhibit a ~24 hr period, and persist in constant darkness. Thus, in the absence of an LD cycle, the recording and analysis of running-wheel activity can be used to determine the subjective time-of-day. Because these rhythms are directed by the circadian clock the subjective time-of-day is referred to as the circadian time (CT). In contrast, when an LD cycle is present, the time-of-day that is determined by the environmental LD cycle is called the zeitgeber time (ZT).
Although circadian rhythms in running-wheel activity are typically linked to the SCN clock6-8, circadian oscillators in many other regions of the brain and body9-14 could also be involved in the regulation of daily activity rhythms. For instance, daily rhythms in food-anticipatory activity do not require the SCN15,16 and instead, are correlated with changes in the activity of extra-SCN oscillators17-20. Thus, running-wheel activity recordings can provide important behavioral information not only about the output of the master SCN clock, but also on the activity of extra-SCN oscillators. Below we describe the equipment and methods used to record, analyze and display circadian locomotor activity rhythms in laboratory rodents.

Circadian rhythms in voluntary wheel-running activity in mammals are tightly coupled to the molecular oscillations of a master clock in the brain. As such, these daily rhythms in behavior can be used to study the influence of genetic, pharmacological, and environmental factors on the functioning of this circadian clock.

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, ...

Measuring Changes in Tactile Sensitivity in the Hind Paw of Mice Using an Electronic von Frey Apparatus

Measuring inflammation-induced changes in thresholds of hind paw withdrawal from mechanical pressure is a useful technique to assess changes in pain perception in rodents. Withdrawal thresholds can be measured first at baseline and then following drug, venom, injury, allergen,&#160;or otherwise evoked inflammation by applying an accurate force on very specific areas of the skin. An electronic von Frey apparatus allows precise assessment of mouse hind paw withdrawal thresholds that are not limited by the available filament sizes in contrast to classical von Frey measurements. The ease and rapidity of measurements allow for incorporation of assessment of tactile sensitivity outcomes in diverse models of rapid-onset inflammatory and neuropathic pain as multiple measurements can be taken within a short time period. Experimental measurements for individual rodent subjects can be internally controlled against individual baseline responses and exclusion criteria easily established to standardize baseline responses within and across experimental groups. Thus, measurements using an electronic von Frey apparatus represent a useful modification of the well-established classical von Frey filament-based assays for rodent mechanical allodynia that may also be applied to other nonhuman mammalian models.

Electronic von Frey measurements are an effective and improved alternative to classical von Frey filaments, as they allow rapid and precise measurement of changes in rodent mechanical withdrawal thresholds to continuously applied pressure stimuli before and after an inflammatory event.

Measuring inflammation-induced changes in thresholds of hind paw withdrawal from mechanical pressure is a useful technique to assess changes in pain ...

A Simple and Inexpensive Method for Determining Cold Sensitivity and Adaptation in Mice

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life. The cold plantar assay allows the objective and inexpensive assessment of cold sensitivity in mice, and can quantify both analgesia and hypersensitivity. Mice are acclimated on a glass plate, and a compressed dry ice pellet is held against the glass surface underneath the hindpaw. The latency to withdrawal from the cooling glass is used as a measure of cold sensitivity.
Cold sensation is also important for survival in regions with seasonal temperature shifts, and in order to maintain sensitivity animals must be able to adjust their thermal response thresholds to match the ambient temperature. The Cold Plantar Assay (CPA) also allows the study of adaptation to changes in ambient temperature by testing the cold sensitivity of mice at temperatures ranging from 30 &#176;C to 5 &#176;C. Mice are acclimated as described above, but the glass plate is cooled to the desired starting temperature using aluminum boxes (or aluminum foil packets) filled with hot water, wet ice, or dry ice. The temperature of the plate is measured at the center using a filament T-type thermocouple probe. Once the plate has reached the desired starting temperature, the animals are tested as described above.
This assay allows testing of mice at temperatures ranging from innocuous to noxious. The CPA yields unambiguous and consistent behavioral responses in uninjured mice and can be used to quantify both hypersensitivity and analgesia. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

The Cold Plantar Assay (CPA) measures cold responsiveness between 30 &#176;C and 5 &#176;C, and can also measure cold adaptation. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life.  The cold ...

An Assay for Measuring the Effects of Ethanol on the Locomotion Speed of Caenorhabditis elegans

Alcohol use disorders are a significant public health concern, for which there are few effective treatment strategies. One difficulty that has delayed the development of more effective treatments is the relative lack of understanding of the molecular underpinnings of the effects of ethanol on behavior. The nematode, Caenorhabditis elegans (C.&#160;elegans), provides a useful model in which to generate and test hypotheses about the molecular effects of ethanol. Here, we describe an assay that has been developed and used to examine the roles of particular genes and environmental factors in behavioral responses to ethanol, in which locomotion is the behavioral output. Ethanol dose-dependently causes an acute depression of crawling on an agar surface. The effects are dynamic; animals exposed to a high concentration demonstrate an initial strong depression of crawling, referred to here as initial sensitivity, and then partially recover locomotion speed despite the continued presence of the drug. This ethanol-induced behavioral plasticity is referred to here as the development of acute functional tolerance. This assay has been used to demonstrate that these two phenotypes are distinct and genetically separable. The straightforward locomotion assay described here is suitable for examining the effects of both genetic and environmental manipulations on these acute behavioral responses to ethanol in C.&#160;elegans.

An Assay for Measuring the Effects of Ethanol on the Locomotion Speed of Caenorhabditis elegans

C. elegans is a useful model for studying the effects of ethanol on behavior. We present a behavioral assay that quantifies the effects of ethanol on the locomotion speed of crawling worms; both initial sensitivity and the development of acute functional tolerance to ethanol can be measured with this assay.

Alcohol use disorders are a significant public health concern, for which there are few effective treatment strategies. One difficulty that has delayed the ...

A Behavioral Assay for Mechanosensation of MARCM-based Clones in Drosophila melanogaster

Because of the structural and functional homology to the hair cells of the mammalian inner ear, the neurons that innervate the Drosophila external sense organs provide an excellent model system for the study of mechanosensation. This protocol describes a simple touch behavior in fruit flies which can be used to identify mutations that interfere with mechanosensation. The tactile stimulation of a macrochaete bristle on the thorax of flies elicits a grooming reflex from either the first or third leg. Mutations that interfere with mechanotransduction (such as NOMPC), or with other aspects of the reflex arc, can inhibit the grooming response. A traditional screen of adult behaviors would have missed mutants that have essential roles during development. Instead, this protocol combines the touch screen with mosaic analysis with a repressible cell marker (MARCM) to allow for only limited regions of homozygous mutant cells to be generated and marked by the expression of green fluorescent protein (GFP). By testing MARCM clones for abnormal behavioral responses, it is possible to screen a collection of lethal p-element mutations to search for new genes involved in mechanosensation that would have been missed by more traditional methods.

A Behavioral Assay for Mechanosensation of MARCM-based Clones in Drosophila melanogaster

In order to identify novel mutations affecting mechanosensation, we designed an assay that measures the behavioral response to tactile stimulation of fly bristles in mutant clones generated by the MARCM method. The combination of techniques allows for the identification of mechanosensitive mutations that would otherwise be lethal.

Because of the structural and functional homology to the hair cells of the mammalian inner ear, the neurons that innervate the Drosophila external sense ...

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method described here enables the investigator to measure long-lasting (from minutes to hours) high-quality intracellular responses from single Drosophila R1-R6 photoreceptors and Large Monopolar Cells (LMCs) to light stimuli. Because the recording system has low noise, it can be used to study variability among individual cells in the fly eye, and how their outputs reflect the physical properties of the visual environment. We outline all key steps in performing this technique. The basic steps in constructing an appropriate electrophysiology set-up for recording, such as design and selection of the experimental equipment are described. We also explain how to prepare for recording by making appropriate (sharp) recording and (blunt) reference electrodes. Details are given on how to fix an intact fly in a bespoke fly-holder, prepare a small window in its eye and insert a recording electrode through this hole with minimal damage. We explain how to localize the center of a cell's receptive field, dark- or light-adapt the studied cell, and to record its voltage responses to dynamic light stimuli. Finally, we describe the criteria for stable normal recordings, show characteristic high-quality voltage responses of individual cells to different light stimuli, and briefly define how to quantify their signaling performance. Many aspects of the method are technically challenging and require practice and patience to master. But once learned and optimized for the investigator's experimental objectives, it grants outstanding in vivo neurophysiological data.

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

Taste Preference Assay for Adult Drosophila

Olfactory and gustatory perception of the environment is vital for animal survival. The most obvious application of these chemosenses is to be able to distinguish good food sources from potentially dangerous food sources. Gustation requires physical contact with a chemical compound which is able to signal through taste receptors that are expressed on the surface of neurons. In insects, these gustatory neurons can be located across the animal's body allowing taste to play an important role in many different behaviors. Insects typically prefer compounds containing sugars, while compounds that are considered bitter tasting are avoided. Given the basic biological importance of taste, there is intense interest in understanding the molecular mechanisms underlying this sensory modality. We describe an adult Drosophila taste assay which reflects the preference of the animals for a given tastant compound. This assay may be applied to animals of any genetic background to examine the taste preference for a desired soluble compound.

Taste Preference Assay for Adult Drosophila

Taste is an important sensory process which facilitates attraction to beneficial substances and avoidance of toxic substances. This protocol describes a simple ingestion assay for determining Drosophila gustatory preference for a given chemical compound.

Olfactory and gustatory perception of the environment is vital for animal survival.  The most obvious application of these chemosenses is to be able to ...