This work was supported by grants DA041613 and GM128974 from the National Institutes of Health to TFCM and RRHA.

1. Construction of the testing apparatus
<ol>
	<li>Create a cardboard template the size of a 24-well cell culture plate by tracing around the plate on cardboard and cutting out the designated area.</li>
	<li>Cut a piece of small insect screen mesh the size of the cell culture plate using the cardboard template from step 1.1.</li>
	<li>Prepare a 24-well cell culture plate by placing a small line of hot glue around the perimeter of the top of the plate using a hot glue gun and affixing the screen mesh on top of the open wells.</li>
	<li>Secure a wooden craft stick to each of three sides of the same cell culture plate from step 1.3 using a hot glue gun. The modified cell culture plate should now resemble the plate diagram shown in Figure 1A and the experimental setup shown in Figure 2. 
	NOTE: Prepare at least as many cell culture plates as will fit in the filming chambers (see below).</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61108/61108fig1.jpg" /> 
Figure 1: Diagram of the testing apparatus and filming chamber. (A) Upper Diagrams. The top, side, and front views of the testing apparatus are shown, respectively. A screen mesh lays flat on top of a 24-well cell culture plate. The wooden craft sticks, represented by the arrowheads, are attached to three adjacent sides for stability and alignment aid, two on the side of the well plate with six wells and one on the side of the plate with four wells. All attachments are hot glued onto the apparatus. (B) Lower Diagrams. The top, side, and front views of the assay set-up are shown, respectively. A slit is cut in the right side of the box, from the opening for the lid to the back of the opening, with the bottom of the slit level to the inner surface. The hole on the top of the box, the surface parallel to the ground, is centered for maximum video exposure. The shaded box represents the video camera. <a href="https://www.jove.com/files/ftp_upload/61108/61108fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61108/61108fig2.jpg" /> 
Figure 2: Photograph of the assay system. The video camera is placed on top of the polystyrene chamber, with the lens inserted in the cut-out hole, illustrated in the diagrams of Figure 1B. Two sets of modified 24-well cell culture plates rest on top of an illumination pad that is inserted in a slit through the side of the chamber. <a href="https://www.jove.com/files/ftp_upload/61108/61108fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Construction of the filming chamber
<ol>
	<li>Create a filming chamber by cutting a hole the size of the video camera lens on the side of a polystyrene box. Cut an additional slit the width of the illumination pad in the opposite side of the polystyrene box. The filming chamber should resemble the filming chamber shown in Figure 1B and Figure 2.</li>
	<li>Prepare the filming chamber for use by inserting the illumination pad into the slit and positioning the camera in the lens hole above the illumination pad.</li>
	<li>Place all materials and perform all subsequent testing in a controlled environment, preferably a behavioral chamber with approximately 30% humidity, 25 &#176;C temperature, uniform airflow, and noise levels less than 65 dB.</li>
</ol>
3. Preparation of the testing apparatus and flies
<ol>
	<li>Pipette 1 mL of 100% ethanol through the screen mesh into each well.</li>
	<li>Dry the screen mesh with a piece of cheesecloth.</li>
	<li>Cut two pieces of cheesecloth the dimensions of the cell culture plate using the cardboard template created in step 1.1. Place them on top of the dry screen mesh of the modified cell culture plate containing ethanol from step 3.2.</li>
	<li>Create a small piece of thin, flexible plastic cutting board by tracing around the cardboard template created in step 1.1 as a general guide and expanding the traced area by 1&#8211;2 cm on one of the short sides. Cut out the expanded traced area from the thin, flexible plastic cutting board. After cutting, ensure that the plastic still fits between the three wooden craft sticks on the testing apparatus, but hangs off one end by 1&#8211;2 cm.</li>
	<li>(Optional) If an aspirator needs to be created, assemble an aspirator like the one shown in Figure 3 by first cutting a P1000 pipette tip in half. Insert the piece with a larger diameter into one end of a ~30 cm piece of flexible tubing to serve as a mouthpiece.</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61108/61108fig3.jpg" /> 
Figure 3: A fly aspirator in which flies are collected with an interchangeable mouthpiece attached to flexible tubing and a wide bore serological pipette with a cotton gauze stopper. The operator can aspirate a single fly into the pipette for transfer without anesthesia. <a href="https://www.jove.com/files/ftp_upload/61108/61108fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="6">
	<li>(Optional) To complete the aspirator assembly, cover the wide end of a 10 cm piece of serological pipette with gauze to prevent flies from getting into the tubing and insert the pipette, gauze first, into the open end of the tubing to serve as a fly chamber. The aspirator should resemble that shown in Figure 3.</li>
	<li>Using an aspirator (Figure 3, steps 3.5 and 3.6), aspirate one fly per well into a separate 24-well cell culture plate. Use the flexible plastic to cover any wells containing previously aspirated flies. Record the well position and any relevant genotype or phenotype information of each fly.</li>
	<li>Hold the flexible plastic flush with the top of the cell culture plate containing the flies to prevent their escape and invert the plate onto the top of the modified cell culture plate with the ethanol. The sheet of flexible plastic should be resting on top of the sheets of cheesecloth. Align the inverted cell culture plate containing flies using the craft sticks to ensure each well with ethanol aligns with each well containing a fly.</li>
	<li>The experimental setup should resemble Figure 2.</li>
</ol>
4. Testing the flies
<ol>
	<li>Ensure the illumination pad is lit at full brightness for maximum visual contrast. Start recording with the video camera.</li>
	<li>To expose the flies to ethanol, carefully remove the plastic from between the well plate and testing apparatus, taking care not to dislodge the cheesecloth.</li>
	<li>Terminate the video recording once all flies have lost postural control. Once it is suspected that all flies have lost postural control, tap firmly in the center of the plate to ensure that all flies have complete loss of postural control. If there is movement, continue to record. Continue to tap periodically (every 1&#8211;2 min) until no movement occurs.</li>
	<li>(Optional) To quickly recover the flies, remove only the top plate from the testing apparatus, revealing sedated flies resting on the cheesecloth. Aspirate individual flies into chosen containers for recovery.</li>
	<li>Replace the ethanol in the modified cell culture plates with 1 mL of fresh 100% ethanol at least 1x every hour to control for evaporation and humidification of the ethanol and to maintain consistent ethanol exposure throughout the assay. Dry the screen mesh with cheesecloth.</li>
	<li>Repeat for as many samples as desired. 
	NOTE: For highest throughput, aspirate the next round of flies into new cell culture plates during the video recording. The protocol can be paused here, as the video recording can be reviewed later.</li>
</ol>
5. Determination of fly sedation time
<ol>
	<li>Record sedation time for each individual fly by watching the video recording. Sedation time is defined as the moment a fly loses complete postural control and locomotor ability. It is recommended to watch the film in reverse and record the time that the fly begins to move to ensure accuracy.</li>
</ol>

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The authors have nothing to disclose.

Here, we present a simple, inexpensive, and high-throughput method for assessing sedation time due to ethanol exposure in Drosophila melanogaster. Unlike many current methods, which require group analyses, this assay enables a single person to collect individual sedation time data for ~2,000 flies within an 8 h work period. We found that a single person can score 48 flies for sedation time in about 5 min. At this rate, 2,000 flies can be scored in approximately 4 h, though scoring can be conducted later. With our assay, the recorded sedation time for most flies ranges from 5&#8211;15 min at an exposure to 1 mL of 100% ethanol. Lower concentrations of ethanol or smaller delivery volumes will result in longer sedation times.
Current methods for assessing sedation time require testing large numbers of flies without readily enabling measurements on single individuals15,16,17,18,19,20,21,22,23,24,25,26. Many current sedation and sensitivity assays rely upon ST5022,23,24, the timepoint at which 50% of the flies are sedated as a result of ethanol exposure. Although obtaining the ST50 for groups of flies was not the primary motivation for developing this assay, the video recordings demonstrate higher utility compared to current methods, as the recordings can be used to ascertain the ST50 for groups of individually tested flies and to measure the percentage of flies that satisfy a given criterion (e.g., loss of postural control) at any time point. It should be noted that such video analyses would require additional time.
Unlike current inebriometer assays, the method we describe does not require specialized tools to set up and can be performed in any laboratory using common materials. Using this method, we have obtained reliable and consistent sedation times for individual flies. The assay can, in principle, be extended to assess the effects of exposure to any volatile substance. The assay can also be applied to measure effects of acute toxicity of volatiles on other insects, including other fly species. Individual sedation time data can be used to assess the extent of phenotypic variation within a population, such as the DGRP.
We used small insect screen mesh to prevent direct contact with the ethanol solution while allowing adequate quantities of ethanol vapors to reach the fly. The layer of white cheesecloth on top of the screen mesh provides visual contrast between the fly and the surface below and ensures that flies do not get caught in the screen mesh, which could lead to ambiguous determination of loss of postural control. Commercially available membranes that are porous to water and air gave inconsistent results and were insufficiently penetrable to ethanol vapors. We intentionally used small insect screen mesh because it is a uniformly porous material that minimizes variation in ethanol exposure as a result of fly position within a well. Modifications can be made to this protocol based on available materials, although we recommend a controlled behavioral chamber, access to 90%&#8211;100% ethanol close to the fly, and uniform ethanol exposure.
Fly position within the cell culture plates should be randomized between replicates to avoid positional bias. For larger experiments that require use of this assay across multiple days and are therefore subject to environmental variation that could influence assay results (e.g., changes in barometric pressure)27, we strongly recommend that flies be tested at the same time each day and randomized both within and across days, especially if different lines and/or sexes are to be compared against one another.
The method we developed is best suited for measuring the effect of acute alcohol exposure but is not suitable for obtaining consumption data or modeling addiction. Alcohol sedation sensitivity data obtained from this assay can, however, be integrated with other measures of alcohol-related phenotypes. One limitation of the system is that the vertical height of standard cell culture plates allows for vertical fly movement that cannot be readily tracked by video for detailed assessment of overall activity or locomotion. However, this limitation does not affect accurate assessment of sedation time. When using flies of different genotypes (e.g., in DGRP-derived outbred populations28), this assay also enables retrieval of individual flies to collect pools of flies with contrasting phenotypes for bulk DNA sequencing and extreme QTL mapping29,30. Overall, this assay permits rapid, inexpensive collection of alcohol sedation data on large numbers of single flies.

The National Institute on Alcohol Abuse and Alcoholism reports that in 2015 excessive alcohol consumption, designated as &#34;alcohol use disorder&#34;, affected an estimated 16 million people in the United States. Alcohol abuse causes a wide range of adverse physiological effects and is a major cause of death in the U.S. In humans, decreased sensitivity, or a low level of response to alcohol, has a strong genetic component and is associated with a higher risk of developing alcohol use disorders1,2,3,4. Genetic risk studies on human populations are challenging because of population admixture, diverse developmental histories and environmental exposures, and reliance on self-reported questionnaires to quantify alcohol-related phenotypes, which are often confounded with other neuropsychiatric conditions.
Drosophila melanogaster provides an excellent model to study the genetic underpinnings of alcohol sensitivity5,6,7,8. The Drosophila model allows strict control over genetic background, and virtually unlimited numbers of individuals of the same genotype can be reared rapidly under well-controlled environmental conditions without regulatory restrictions and at relatively low cost. In addition to publicly available mutations and RNAi lines that target a majority of genes in the genome, the availability of the Drosophila melanogaster Genetic Reference Panel (DGRP), a population of 205 inbred wild-derived lines with complete genome sequences, has enabled genome-wide association studies9,10. Such studies have identified genetic networks associated with effects on development time and viability upon developmental exposure to ethanol11,12. Evolutionary conservation of fundamental biological processes enables translational inferences to be drawn by superimposing human orthologs on their fly counterparts.
Flies exposed to ethanol undergo physiological and behavioral changes that resemble human alcohol intoxication, including loss of postural control8, sedation, and development of tolerance13,14,15. Alcohol induced sedation in Drosophila can be quantified using inebriometers. These are 122 cm long vertical glass columns with slanted mesh partitions to which flies can attach16,17,18. A group of at least 50 flies (sexes can be analyzed separately) are introduced in the top of the column and exposed to ethanol vapors. Flies that lose postural control fall through the column and are collected at 1 min intervals. The mean elution time serves as a measure of sensitivity to alcohol intoxication. When flies are exposed to alcohol a second time after recovering from the first exposure, they can develop tolerance, as evident from a shift in mean elution time13,15,19,20. Whereas inebriometer assays have led to identification of genes, genetic networks, and cellular pathways associated with alcohol sedation sensitivity and development of tolerance12,13,14,21, the assay is time consuming, low-throughput, and ineffective for measuring alcohol sensitivity in single flies.
Alternative ethanol sedation assays that do not require the elaborate inebriometer set-up allow for more convenient measurements but are still limited in throughput and generally require analyses of groups of flies rather than individuals21,22,23,24,25. Assessing single flies minimizes the potential for confounding effects due to group interactions, such as those stemming from social behaviors. Here, we present a simple, low-cost, high-throughput assay for assessing alcohol sedation sensitivity in large numbers of single flies.

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		<tr height="21">
			<td height="21" style="height:21px;width:289px;">24-well Cell Culture Plates</td>
			<td style="width:207px;">Corning</td>
			<td style="width:128px;">3526</td>
			<td style="width:772px;">Flat-bottomed; will house flies throughout assay</td>
		</tr>
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			<td height="21" style="height:21px;width:289px;">Aspirator</td>
			<td style="width:207px;"></td>
			<td style="width:128px;"></td>
			<td style="width:772px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:289px;">Cheesecloth</td>
			<td style="width:207px;">Genesee Scientific</td>
			<td style="width:128px;">53-100</td>
			<td>Widely available.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:289px;">Ethanol</td>
			<td style="width:207px;">Decon Labs</td>
			<td style="width:128px;">V1001</td>
			<td>Widely available.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:289px;">Flexible Plastic Cutting Board (Plate Cover)</td>
			<td style="width:207px;">Walmart</td>
			<td style="width:128px;">550098612</td>
			<td>Any flat plastic that can slide easily and cover a 24-well plate completely. Flexible plastic cutting board works well.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:289px;">Gauze (for aspirator)</td>
			<td style="width:207px;">Honeywell North</td>
			<td style="width:128px;">67622</td>
			<td>Widely available.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:289px;">Illumination Pad</td>
			<td style="width:207px;">Amazon (AGPtek)</td>
			<td style="width:128px;">ASIN B00YA9GP0G</td>
			<td>Any light pad to provide contrast is suitable.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:289px;">Jumbo Craft Sticks</td>
			<td style="width:207px;">Michaels</td>
			<td style="width:128px;">10334892</td>
			<td>Any craft stick at least 7 cm long is suitable.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:289px;">P1000 Pipette Tip (for aspirator)</td>
			<td style="width:207px;">Genesee Scientific</td>
			<td style="width:128px;">24-165RL</td>
			<td>Any P1000 pipette tip is suitable.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:289px;">Serological Pipette (for aspirator)</td>
			<td style="width:207px;">Genesee Scientific</td>
			<td style="width:128px;">12-104</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:289px;">Small Insect Screen Mesh</td>
			<td style="width:207px;">Lowe&#39;s (Saint-Gobain ADFORS)</td>
			<td style="width:128px;">89322</td>
			<td>Any small insect screen mesh is suitable.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:289px;">Testing Chamber</td>
			<td style="width:207px;"></td>
			<td style="width:128px;"></td>
			<td>Interior space dimension big enough to encompass light pad. Can be constructed from a polystyrene box.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:289px;">Tygon Tubing (for aspirator)</td>
			<td style="width:207px;">Grainger</td>
			<td style="width:128px;">9CUG7</td>
			<td>Widely available.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:289px;">Video Camera</td>
			<td style="width:207px;">Canon</td>
			<td>1959C001AA</td>
			<td>Any video camera is suitable.</td>
		</tr>
	</tbody>
</table>

high-throughput method for measuring alcohol sedation time of individual drosophila melanogaster

Drosophila melanogaster provides an excellent model to study the genetic underpinnings of alcohol sensitivity. In contrast to studies in human populations, the Drosophila model allows strict control over genetic background, and virtually unlimited numbers of individuals of the same genotype can be reared rapidly under well-controlled environmental conditions without regulatory restrictions and at relatively low cost. Flies exposed to ethanol undergo physiological and behavioral changes that resemble human alcohol intoxication, including loss of postural control, sedation, and development of tolerance. Here, we describe a simple, low-cost, high-throughput assay for assessing alcohol sedation sensitivity in large numbers of single flies. The assay is based on video recording of single flies introduced without anesthesia in 24-well cell culture plates in a set-up that enables synchronous initiation of alcohol exposure. The system enables a single person to collect individual ethanol sedation data on as many as 2,000 flies within an 8 h work period. The assay can, in principle, be extended to assess the effects of exposure to any volatile substance and applied to measure effects of acute toxicity of volatiles on other insects, including other fly species.

Two 24-well microtiter plates could generate data simultaneously on 48 individual flies within as little as 10 min. Table 1 lists measurements of ethanol sedation times for 48 individual flies, males and females separately, of two DGRP lines with different sensitivities to alcohol exposure on development time and viability13. Flies of line RAL_555 were less sensitive than line RAL_177 (Figure 4, Table 2; p &#60; 0.0001, ANOVA). Males and females of RAL_177 showed no sexually dimorphic effect (Figure 4, Table 2; p &#62; 0.1, ANOVA), whereas females of line RAL_555 were less sensitive to ethanol exposure than the males (Figure 4, Table 2; p &#60; 0.006, ANOVA). The large number of flies that can be measured simultaneously and the ability to measure sexes and different lines contemporaneously can increase accuracy by reducing error due to environmental variation.
<table border="1" fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>A.</td>
			<td colspan="6">Ethanol Sedation Time (s)</td>
			<td>B.</td>
			<td colspan="6">Ethanol Sedation Time (s)</td>
		</tr>
		<tr>
			<td></td>
			<td colspan="3">Females</td>
			<td colspan="3">Males</td>
			<td></td>
			<td colspan="3">Females</td>
			<td colspan="3">Males</td>
		</tr>
		<tr>
			<td></td>
			<td>414</td>
			<td>365</td>
			<td>477</td>
			<td>423</td>
			<td>568</td>
			<td>309</td>
			<td></td>
			<td>937</td>
			<td>742</td>
			<td>622</td>
			<td>460</td>
			<td>331</td>
			<td>498</td>
		</tr>
		<tr>
			<td></td>
			<td>201</td>
			<td>384</td>
			<td>498</td>
			<td>411</td>
			<td>523</td>
			<td>626</td>
			<td></td>
			<td>791</td>
			<td>619</td>
			<td>197</td>
			<td>467</td>
			<td>455</td>
			<td>562</td>
		</tr>
		<tr>
			<td></td>
			<td>228</td>
			<td>364</td>
			<td>333</td>
			<td>440</td>
			<td>403</td>
			<td>267</td>
			<td></td>
			<td>504</td>
			<td>744</td>
			<td>513</td>
			<td>570</td>
			<td>582</td>
			<td>506</td>
		</tr>
		<tr>
			<td></td>
			<td>440</td>
			<td>416</td>
			<td>404</td>
			<td>408</td>
			<td>422</td>
			<td>384</td>
			<td></td>
			<td>970</td>
			<td>540</td>
			<td>369</td>
			<td>865</td>
			<td>533</td>
			<td>492</td>
		</tr>
		<tr>
			<td></td>
			<td>888</td>
			<td>283</td>
			<td>285</td>
			<td>322</td>
			<td>369</td>
			<td>287</td>
			<td></td>
			<td>595</td>
			<td>550</td>
			<td>606</td>
			<td>392</td>
			<td>544</td>
			<td>345</td>
		</tr>
		<tr>
			<td></td>
			<td>1079</td>
			<td>519</td>
			<td>315</td>
			<td>393</td>
			<td>376</td>
			<td>284</td>
			<td></td>
			<td>418</td>
			<td>709</td>
			<td>553</td>
			<td>308</td>
			<td>477</td>
			<td>388</td>
		</tr>
		<tr>
			<td></td>
			<td>718</td>
			<td>287</td>
			<td>432</td>
			<td>275</td>
			<td>206</td>
			<td>411</td>
			<td></td>
			<td>366</td>
			<td>564</td>
			<td>558</td>
			<td>385</td>
			<td>576</td>
			<td>377</td>
		</tr>
		<tr>
			<td></td>
			<td>598</td>
			<td>337</td>
			<td>398</td>
			<td>279</td>
			<td>631</td>
			<td>372</td>
			<td></td>
			<td>437</td>
			<td>692</td>
			<td>578</td>
			<td>460</td>
			<td>511</td>
			<td>412</td>
		</tr>
		<tr>
			<td></td>
			<td>241</td>
			<td>398</td>
			<td>364</td>
			<td>347</td>
			<td>374</td>
			<td>808</td>
			<td></td>
			<td>665</td>
			<td>729</td>
			<td>484</td>
			<td>532</td>
			<td>425</td>
			<td>354</td>
		</tr>
		<tr>
			<td></td>
			<td>229</td>
			<td>423</td>
			<td>534</td>
			<td>386</td>
			<td>396</td>
			<td>628</td>
			<td></td>
			<td>312</td>
			<td>576</td>
			<td>305</td>
			<td>334</td>
			<td>531</td>
			<td>506</td>
		</tr>
		<tr>
			<td></td>
			<td>388</td>
			<td>488</td>
			<td>451</td>
			<td>523</td>
			<td>322</td>
			<td>533</td>
			<td></td>
			<td>682</td>
			<td>638</td>
			<td>420</td>
			<td>560</td>
			<td>548</td>
			<td>379</td>
		</tr>
		<tr>
			<td></td>
			<td>252</td>
			<td>529</td>
			<td>375</td>
			<td>427</td>
			<td>330</td>
			<td>540</td>
			<td></td>
			<td>1045</td>
			<td>741</td>
			<td>708</td>
			<td>832</td>
			<td>509</td>
			<td>472</td>
		</tr>
		<tr>
			<td></td>
			<td>674</td>
			<td>401</td>
			<td>303</td>
			<td>401</td>
			<td>307</td>
			<td>311</td>
			<td></td>
			<td>394</td>
			<td>675</td>
			<td>381</td>
			<td>477</td>
			<td>449</td>
			<td>784</td>
		</tr>
		<tr>
			<td></td>
			<td>303</td>
			<td>453</td>
			<td>351</td>
			<td>429</td>
			<td>525</td>
			<td>262</td>
			<td></td>
			<td>540</td>
			<td>690</td>
			<td>520</td>
			<td>556</td>
			<td>495</td>
			<td>226</td>
		</tr>
		<tr>
			<td></td>
			<td>258</td>
			<td>483</td>
			<td>302</td>
			<td>389</td>
			<td>562</td>
			<td>319</td>
			<td></td>
			<td>356</td>
			<td>615</td>
			<td>336</td>
			<td>454</td>
			<td>524</td>
			<td>590</td>
		</tr>
		<tr>
			<td></td>
			<td>346</td>
			<td>426</td>
			<td>385</td>
			<td>416</td>
			<td>596</td>
			<td>287</td>
			<td></td>
			<td>626</td>
			<td>678</td>
			<td>840</td>
			<td>634</td>
			<td>677</td>
			<td>509</td>
		</tr>
	</tbody>
</table>
Table 1: Measurements of ethanol sedation times (s) of individual flies of (A) DGRP lines RAL_177 and (B) RAL_555 for separate sexes (n = 48). See also Table 2, Figure 4.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61108/61108fig4.jpg" /> 
Figure 4: Alcohol sedation times of DGRP lines RAL_177 and RAL_555. The bars represent means and the error bars SEM (n = 48). Sedation times for RAL_177 flies were less than those for RAL_55 flies (p &#60; 0.0001, ANOVA). Individual data points are indicated in Table 1. Additional statistically significant differences between sexes and lines are indicated in the text and in Table 2. <a href="https://www.jove.com/files/ftp_upload/61108/61108fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Analysis</td>
			<td>Source of Variation</td>
			<td>df</td>
			<td>SS</td>
			<td>F-Value</td>
			<td>P-value</td>
		</tr>
		<tr>
			<td rowspan="4">Full Model Pooled</td>
			<td>Line</td>
			<td>1</td>
			<td>769627</td>
			<td>34.869</td>
			<td>&#60;0.0001</td>
		</tr>
		<tr>
			<td>Sex</td>
			<td>1</td>
			<td>105001</td>
			<td>4.757</td>
			<td>0.0304</td>
		</tr>
		<tr>
			<td>Line x Sex</td>
			<td>1</td>
			<td>86021</td>
			<td>3.897</td>
			<td>0.0498</td>
		</tr>
		<tr>
			<td>Error</td>
			<td>188</td>
			<td>4149491</td>
			<td colspan="2"></td>
		</tr>
		<tr>
			<td rowspan="2">Reduced Model Females</td>
			<td>Line</td>
			<td>1</td>
			<td>685126</td>
			<td>23.58</td>
			<td>&#60;0.0001</td>
		</tr>
		<tr>
			<td>Error</td>
			<td>94</td>
			<td>2730718</td>
			<td colspan="2"></td>
		</tr>
		<tr>
			<td rowspan="2">Reduced Model Males</td>
			<td>Line</td>
			<td>1</td>
			<td>170522</td>
			<td>11.3</td>
			<td>0.0011</td>
		</tr>
		<tr>
			<td>Error</td>
			<td>94</td>
			<td>1418774</td>
			<td colspan="2"></td>
		</tr>
		<tr>
			<td rowspan="2">Reduced Model RAL_177</td>
			<td>Sex</td>
			<td>1</td>
			<td>473</td>
			<td>0.023</td>
			<td>0.8800</td>
		</tr>
		<tr>
			<td>Error</td>
			<td>94</td>
			<td>1943741</td>
			<td colspan="2"></td>
		</tr>
		<tr>
			<td rowspan="2">Reduced Model RAL_555</td>
			<td>Sex</td>
			<td>1</td>
			<td>190549</td>
			<td>8.12</td>
			<td>0.0054</td>
		</tr>
		<tr>
			<td>Error</td>
			<td>94</td>
			<td>2205751</td>
			<td colspan="2"></td>
		</tr>
	</tbody>
</table>
Table 2: Analyses of variance for sedation time across sex and DGRP line. The model used was Y = &#181; + L + S + LxS + &#949;, where &#181; is the overall mean, L is the fixed effect of the DGRP line (RAL_177, RAL_555), S is the fixed effect of sex (male, female), LxS is the interaction term (fixed), and &#949; is the error term. The models Y = &#181; + L + &#949; and Y = &#181; + S + &#949; were used for the reduced models. Line, Sex, and the Line x Sex interaction term were all significant in the full model at &#945; &#60; 0.05. Reduced models by sex and DGRP line RAL_555 were also significant at &#945; &#60; 0.01. See also Table 1, Figure 4. df = degrees of freedom, SS = Type I Sums of Squares.

Watch this Scientific Journal Video about High-Throughput Method for Measuring Alcohol Sedation Time of Individual Drosophila melanogaster at JoVE.com

High-Throughput Method for Measuring Alcohol Sedation Time of Individual Drosophila melanogaster

Current methods to measure alcohol sensitivity in Drosophila are designed to test groups of flies. We present a simple, low-cost, high-throughput assay for assessing alcohol sedation sensitivity in large numbers of single flies. The method does not require specialized tools and can be performed in any laboratory using common materials.

High-Throughput Method for Measuring Alcohol Sedation Time of Individual Drosophila melanogaster

Drosophila melanogaster provides an excellent model to study the genetic underpinnings of alcohol sensitivity. In contrast to studies in human ...

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Anxiolytics, Sedatives and Hypnotics

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7.0K 조회수.  Clemson University. Current methods to measure alcohol sensitivity in Drosophila are designed to test groups of flies. We present a simple, low-cost, high-throughput assay for assessing alcohol sedation sensitivity in large numbers of single flies. The method does not require specialized tools and can be performed in any laboratory using common materials.

비디오: High-Throughput Method for Measuring Alcohol Sedation Time of Individual Drosophila melanogaster

Rodent Brain Microinjection to Study Molecular Substrates of Motivated Behavior

Brain microinjection can aid elucidation of the molecular substrates of complex behaviors, such as motivation. For this purpose rodents can serve as appropriate models, partly because the response to behaviorally relevant stimuli and the circuitry parsing stimulus-action outcomes is astonishingly similar between humans and rodents. In studying molecular substrates of complex behaviors, the microinjection of reagents that modify, augment, or silence specific systems is an invaluable technique. However, it is crucial that the microinjection site is precisely targeted in order to aid interpretation of the results. We present a method for the manufacture of surgical implements and microinjection needles that enables accurate microinjection and unlimited customizability with minimal cost. Importantly, this technique can be successfully completed in awake rodents if conducted in conjunction with other JoVE articles that covered requisite surgical procedures. Additionally, there are many behavioral paradigms that are well suited for measuring motivation. The progressive ratio is a commonly used method that quantifies the efficacy of a reinforcer to maintain responding despite an (often exponentially) increasing work requirement. This assay is sensitive to reinforcer magnitude and pharmacological manipulations, which allows reinforcing efficacy and/ or motivation to be determined. We also present a straightforward approach to program operant software to accommodate a progressive ratio reinforcement schedule.

Rodents are an appropriate model to investigate the molecular substrates of behavior and complex psychiatric disorders. Brain microinjection in awake rodents can be used to elucidate disease substrates. An efficient and customizable brain microinjection method as well as the execution of an operant paradigm that quantifies motivation is presented.

쥐 뇌 미세 주입은 분자 기판 동기를 행동을 연구하기 위해

Brain microinjection can aid elucidation of the molecular substrates of complex behaviors, such as motivation. For this purpose rodents can serve as ...

연구

행동분석학

A Method for Evaluating the Reinforcing Properties of Ethanol in Rats without Water Deprivation, Saccharin Fading or Extended Access Training

Operant oral self-administration methods are commonly used to study the reinforcing properties of ethanol in animals. However, the standard methods require saccharin/sucrose fading, water deprivation and/or extended training to initiate operant responding in rats. This paper describes a novel and efficient method to quickly initiate operant responding for ethanol that is convenient for experimenters and does not require water deprivation or saccharin/sucrose fading, thus eliminating the potential confound of using sweeteners in ethanol operant self-administration studies. With this method, Wistar rats typically acquire and maintain self-administration of a 20% ethanol solution in less than two weeks of training. Furthermore, blood ethanol concentrations and rewards are positively correlated for a 30 min self-administration session. Moreover, naltrexone, an FDA-approved medication for alcohol dependence that has been shown to suppress ethanol self-administration in rodents, dose-dependently decreases alcohol intake and motivation to consume alcohol for rats self-administering 20% ethanol, thus validating the use of this new method to study the reinforcing properties of alcohol in rats.

This protocol describes a novel and efficient method to quickly initiate operant responding for ethanol in rats that, contrary to standard methods, does not require water deprivation or saccharin/sucrose fading to initiate responding.

물 박탈, 사카린 페이딩 또는 확장 액세스 교육을받지 않은 쥐에 에탄올의 강화 속성을 평가하는 방법

Operant oral self-administration methods are commonly used to study the reinforcing properties of ethanol in animals. However, the standard methods ...

The Treadmill Fatigue Test: A Simple, High-throughput Assay of Fatigue-like Behavior for the Mouse

Fatigue is a prominent symptom in many diseases and disorders and reduces quality of life for many people. The lack of clear pathogenesis and failure of current interventions to adequately treat fatigue in all patients leaves a need for new treatment options. Despite the therapeutic need and importance of preclinical research in helping identify promising novel treatments, few preclinical assays of fatigue are available. Moreover, the most common preclinical assay used to assess fatigue-like behavior, voluntary wheel running, is not suitable for use with some strains of mice, may not be sensitive to drugs that reduce fatigue, and has relatively low throughput. The current protocol describes a novel, non-voluntary preclinical assay of fatigue-like behavior, the treadmill fatigue test, and provides evidence of its efficacy in detecting fatigue-like behavior in mice treated with a chemotherapy drug known to cause fatigue in humans and fatigue-like behavior in animals. This assay may be a beneficial alternative to wheel running, as fatigue-like behavior and potential interventions can be assessed in a greater number of mice over a shorter time frame, thus permitting faster discovery of new therapeutic options.

Fatigue is a common, undertreated and frequently poorly-understood symptom in many diseases and disorders. New preclinical assays of fatigue may help to improve current understanding and future treatment of fatigue. To that end, the current protocol provides a novel means of measuring fatigue-like behavior in the mouse.

디딜 방아 피로 시험 : 마우스에 대한 피로와 같은 행동 간단한, 높은 처리량 분석

Fatigue is a prominent symptom in many diseases and disorders and reduces quality of life for many people. The lack of clear pathogenesis and failure of ...

A Method to Test the Effect of Environmental Cues on Mating Behavior in Drosophila melanogaster

An individual's sexual drive is influenced by genotype, experience and environmental conditions. How these factors interact to modulate sexual behaviors remains poorly understood. In Drosophila melanogaster, environmental cues, such as food availability, affect mating activity offering a tractable system to investigate the mechanisms modulating sexual behavior. In D. melanogaster, environmental cues are often sensed via the chemosensory gustatory and olfactory systems. Here, we present a method to test the effect of environmental chemical cues on mating behavior. The assay consists of a small mating arena containing food medium and a mating couple. The mating frequency for each couple is continuously monitored for 24 h. Here we present the applicability of this assay to test environmental compounds from an external source through a pressurized air system as well as manipulation of the environmental components directly in the mating arena. The use of a pressurized air system is especially useful to test the effect of very volatile compounds, while manipulating components directly in the mating arena can be of value to ascertain a compound's presence. This assay can be adapted to answer questions about the influence of genetic and environmental cues on mating behavior and fecundity as well as other male and female reproductive behaviors.

A Method to Test the Effect of Environmental Cues on Mating Behavior in Drosophila melanogaster

We demonstrate an assay to analyze the environmental and genetic cues that influence mating behavior in the fruit fly Drosophila melanogaster.

환경에서 신호의 결합 효과에 미치는 영향을 시험하는 방법<em&gt; Drosophila melanogaster</em

An individual's sexual drive is influenced by genotype, experience and environmental conditions. How these factors interact to modulate sexual behaviors ...

A Burrowing/Tunneling Assay for Detection of Hypoxia in Drosophila melanogaster Larvae

Oxygen deprivation in animals can result from exposure to low atmospheric oxygen levels or from internal tissue damage that interferes with oxygen distribution. It is also possible that aberrant behavior of oxygen-sensing neurons could induce hypoxia-like behavior in the presence of normal oxygen levels. In D. melanogaster, development at low oxygen levels results in inhibition of growth and sluggish behavior during the larval phases. However, these established manifestations of oxygen deficit overlap considerably with the phenotypes of many mutations that regulate growth, stress responses or locomotion. As result, there is currently no assay available to identify i) cellular hypoxia induced by a mutation or ii) hypoxia-like behavior when induced by abnormal neuronal behavior.
We have recently identified two distinctive behaviors in D. melanogaster larvae that occur at normal oxygen levels in response to internal detection of hypoxia. First, at all stages, such larvae avoid burrowing into food, often straying far away from a food source. Second, tunneling into a soft substratum, which normally occurs during the wandering third instar stage is completely abolished if larvae are hypoxic. The assay described here is designed to detect and quantitate these behaviors and thus to provide a way to detect hypoxia induced by internal damage rather than low external oxygen. Assay plates with an agar substratum and a central plug of yeast paste are used to support animals through larval life. The positions and state of the larvae are tracked daily as they proceed from first to third instar. The extent of tunneling into the agar substratum during wandering phase is quantitated after pupation using NIH ImageJ. The assay will be of value in determining when hypoxia is a component of a mutant phenotype and thus provide insight into possible sites of action of the gene in question.

A Burrowing/Tunneling Assay for Detection of Hypoxia in Drosophila melanogaster Larvae

The protocol describes a simple assay to identify Drosophila melanogaster larvae that are experiencing hypoxia under normal atmospheric oxygen levels. This protocol allows hypoxic larvae to be distinguished from other mutants that show overlapping phenotypes such as sluggishness or slow growth.

초파리 melanogaster 애벌레에서 Hypoxia의 검출에 대 한 분석 결과 내/터널링

Oxygen deprivation in animals can result from exposure to low atmospheric oxygen levels or from internal tissue damage that interferes with oxygen ...

Noninvasive, High-throughput Determination of Sleep Duration in Rodents

Traditionally, sleep is monitored by an electroencephalogram (EEG). EEG studies in rodents require surgical implantation of the electrodes followed by a long recovery period. To perform an EEG recording, the animal is connected to a receiver, creating an unnatural tether to the head-mount. EEG monitoring is time consuming, carries risk to the animal, and is not a completely natural setting for the measurement of sleep. Alternative methods to detect sleep, particularly in a high-throughput fashion, would greatly advance the field of sleep research. Here, we describe a validated method for detecting sleep via activity-based home-cage monitoring. Previous studies have shown that sleep assessed via this method has a high degree of agreement with sleep defined by traditional EEG-based measures. Whereas this method is validated for total sleep time, it is important to note that sleep bout duration should be assessed by an EEG which has better temporal resolution. The EEG can also differentiate rapid eye movement (REM) and non-REM sleep, giving more detail about the exact nature of sleep. Nevertheless, activity-based sleep determination can be used to analyze multiple days of undisturbed sleep and to assess sleep as a response to an acute event (like stress). Here, we show the power of this system to detect the response of mice to daily intraperitoneal injections.

We describe a high-throughput method of measuring sleep by means of activity-based home-cage monitoring. This method offers advantages over traditional EEG-based methods. It is well validated for the determination of total sleep duration and can be a powerful tool to monitor sleep in rodent models of human disease.

수 면 기간 설치류의 비 침범 성, 높은 처리량 결정

Traditionally, sleep is monitored by an electroencephalogram (EEG). EEG studies in rodents require surgical implantation of the electrodes followed by a ...

An Automated Method to Determine the Performance of Drosophila in Response to Temperature Changes in Space and Time

Temperature is a ubiquitous environmental factor that affects how species distribute and behave. Different species of Drosophila fruit flies have specific responses to changing temperatures according to their physiological tolerance and adaptability. Drosophila flies also possess a temperature sensing system that has become fundamental to understanding the neural basis of temperature processing in ectotherms. We present here a temperature-controlled arena that permits fast and precise temperature changes with temporal and spatial control to explore the response of individual flies to changing temperatures. Individual flies are placed in the arena and exposed to pre-programmed temperature challenges, such as uniform gradual increases in temperature to determine reaction norms or spatially distributed temperatures at the same time to determine preferences. Individuals are automatically tracked, allowing the quantification of speed or location preference. This method can be used to rapidly quantify the response over a large range of temperatures to determine temperature performance curves in Drosophila or other insects of similar size. In addition, it can be used for genetic studies to quantify temperature preferences and reactions of mutants or wild-type flies. This method can help uncover the basis of thermal speciation and adaptation, as well as the neural mechanisms behind temperature processing.

An Automated Method to Determine the Performance of Drosophila in Response to Temperature Changes in Space and Time

Here we present a protocol to automatically determine the locomotor performance of Drosophila at changing temperatures using a programmable temperature-controlled arena that produces fast and accurate temperature changes in time and space.

시간과 공간에서 온도 변화에 대 한 응답에서 초파리 의 성능을 결정 하는 자동화 된 방법

Temperature is a ubiquitous environmental factor that affects how species distribute and behave. Different species of Drosophila fruit flies have specific ...

Multi-Modal Signals for Analyzing Pain Responses to Thermal and Electrical Stimuli

The assessment of pain relies mostly on methods that require a person to communicate. However, for people with cognitive and verbal impairments, existing methods are not sufficient as they lack reliability and validity. To approach this problem, recent research focuses on an objective pain assessment facilitated by parameters of responses derived from physiology, and video and audio signals. To develop reliable automated pain recognition systems, efforts have been made in creating multimodal databases in order to analyze pain and detect valid pain patterns. While the results are promising, they only focus on discriminating pain or pain intensities versus no pain. In order to advance this, research should also consider the quality and duration of pain as they provide additional valuable information for more advanced pain management. To complement existing databases and the analysis of pain regarding quality and length, this paper proposes a psychophysiological experiment to elicit, measure, and collect valid pain reactions. Participants are subjected to painful stimuli that differ in intensity (low, medium, and high), duration (5 s / 1 min), and modality (heat / electric pain) while audio, video (e.g., facial expressions, body gestures, facial skin temperature), and physiological signals (e.g., electrocardiogram [ECG], skin conductance level [SCL], facial electromyography [EMG], and EMG of M. trapezius) are being recorded. The study consists of a calibration phase to determine a subject&#8217;s individual pain range (from low to intolerable pain) and a stimulation phase in which pain stimuli, depending on the calibrated range, are applied. The obtained data may allow refining, improving, and evaluating automated recognition systems in terms of an objective pain assessment. For further development of such systems and to investigate pain reactions in more detail, additional pain modalities such as pressure, chemical, or cold pain should be included in future studies. Recorded data of this study will be released as the &#8220;X-ITE Pain Database&#8221;.

This article focuses on the experimental elicitation of pain through heat (thermal) and electrical stimulation while recording physiological, visual, and paralinguistic responses. It aims at collecting valid multimodal data for analyzing pain based on its intensity, quality, and duration.&#160;

열 및 전기 자극에 고통 응답을 분석 하기 위한 다중 모드 신호

The assessment of pain relies mostly on methods that require a person to communicate. However, for people with cognitive and verbal impairments, existing ...

An Improved Assay and Tools for Measuring Mechanical Nociception in Drosophila Larvae

Published assays for mechanical nociception in Drosophila have led to variable assessments of behavior. Here, we fabricated, for use with Drosophila larvae, customized metal nickel-titanium alloy (nitinol) filaments. These mechanical probes are similar to the von Frey filaments used in vertebrates to measure mechanical nociception. Here, we demonstrate how to make and calibrate these mechanical probes and how to generate a full behavioral dose-response from subthreshold (innocuous or non-noxious range) to suprathreshold (low to high noxious range) stimuli. To demonstrate the utility of the probes, we investigated tissue damage-induced hypersensitivity in Drosophila larvae. Mechanical allodynia (hypersensitivity to a normally innocuous mechanical stimulus) and hyperalgesia (exaggerated responsiveness to a noxious mechanical stimulus) have not yet been established in Drosophila larvae. Using mechanical probes that are normally innocuous or probes that typically elicit an aversive behavior, we found that Drosophila larvae develop mechanical hypersensitization (both allodynia and hyperalgesia) after tissue damage. Thus, the mechanical probes and assay that we illustrate here will likely be important tools to dissect the fundamental molecular/genetic mechanisms of mechanical hypersensitivity.

An Improved Assay and Tools for Measuring Mechanical Nociception in Drosophila Larvae

The goal of this protocol is to show how to perform an improved assay for mechanical nociception in Drosophila larvae. We use the assay here to demonstrate that mechanical hypersensitivity (allodynia and hyperalgesia) exists in Drosophila larvae.

드로소필라 애벌레의 기계적 고치절 측정을 위한 향상된 분석 및 도구

Published assays for mechanical nociception in Drosophila have led to variable assessments of behavior. Here, we fabricated, for use with Drosophila ...

A High Throughput Microplate Feeder Assay for Quantification of Consumption in Drosophila

Quantifying food intake in Drosophila is used to study the genetic and physiological underpinnings of consumption-associated traits, their environmental factors, and the toxicological and pharmacological effects of numerous substances. Few methods currently implemented are amenable to high throughput measurement. The Microplate Feeder Assay (MFA) was developed for quantifying the consumption of liquid food for individual flies using absorbance. In this assay, flies consume liquid food medium from select wells of a 1536-well microplate. By incorporating a dilute tracer dye into the liquid food medium and loading a known volume into each well, absorbance measurements of the well acquired before and after consumption reflect the resulting change in volume (i.e., volume consumed). To enable high throughput analysis with this method, a 3D-printed coupler was designed that allows flies to be sorted individually into 96-well microplates. This device precisely orients 96- and 1536-well microplates to give each fly access to up to 4 wells for consumption, thus enabling food preference quantification in addition to regular consumption. Furthermore, the device has barrier strips that toggle between open and closed positions to allow for controlled containment and release of a column of samples at a time. This method enables high throughput measurements of consumption of aqueous solutions by many flies simultaneously. It also has the potential to be adapted to other insects and to screen consumption of nutrients, toxins, or pharmaceuticals.

A High Throughput Microplate Feeder Assay for Quantification of Consumption in Drosophila

The microplate feeder assay offers an economical, high throughput method for quantifying liquid food consumption in Drosophila. A 3D-printed device connects a 96-well microplate in which flies are housed to a 1536-well microplate from which flies consume a feeding solution with a tracer dye. The solution volume decline is measured spectrophotometrically.

드로소필라에서 소비의 정량화를 위한 높은 처리량 마이크로 플레이트 피더 분석

Quantifying food intake in Drosophila is used to study the genetic and physiological underpinnings of consumption-associated traits, their environmental ...