The authors would like thank the Bloomington stock center, Liqun Luo, Charles Zuker, and Lily and Yuh Nung Jan for the generous sharing of fly stocks and the following for funding: SOMAS-URM (to JD and SW), Bachelor Ford Faculty Summer Fellowship (to SW), BD Corporation Summer Research Fellowship (to CL and DL), The Renee and Anthony M. Marlon, M.D. &#39;63 Summer Research Fellowship (to DL)James C. &#39;75 and Jane Colihan Summer Research Fellowship (TO) through the Alumni/Parent Summer Research Fund of the College of the Holy Cross and the Stransky Foundation Summer Research Fellowship (to TO). Special thanks to the Department of Biology and the Dean&#39;s Office at College of the Holy Cross for the support of all the work in the lab.

1. Crosses to Generate MARCM Flies
<ol>
	<li>Maintain flies on cornmeal molasses media (6.5 g/L agar; 23.5 g/L yeast; 60 g/L cornmeal; 60 ml/L molasses; 4 ml/L acid mix; 0.13% Tegosept) or standard Drosophila media at 22 degrees Celsius in a 12-hr light/dark cycle. 
	Note: A Lower than standard temperature is used to keep MARCM-stocks healthy. This extends the developmental time between egg and adulthood to 14 days.</li>
	<li>Use virgin females (approximately 1 - 8 days old) from a MARCM-ready stock containing the necessary genetic elements (GFP under the control of an upstream-activating sequence (UAS-GFP), the Gal80 repressor protein, the Gal4 transcription factor, a heat-shock driven recombinase, and a FRT recombination site). 
	Note: Our MARCM ready-stocks contain the following genotypes: elavGal4, UAS-CD8-GFP, hs-flipase; FRT40A, tubGal80/CyO; tubGal4/TM3Sb. 
	Note: The specific FRT site will depend on the location of the mutation in the genome. For example, FRT40A is used to test mutations on the left arm of chromosome 2, such as NOMPC (see10 for more details on MARCM protocols).</li>
	<li>Cross MARCM-ready virgin females to males that contain a mutation of interest and its corresponding FRT site (for example, w-; NOMPC3, FRT40A/CyO; +/+). To control for effective MARCM induction, set a second cross using male flies containing the same FRT site, but without a mutation. Use an approximate ratio of 5 females: 1 male in crosses.</li>
</ol>
2. Timed Egg Laying
<ol>
	<li>After allowing flies to mate for at least one night, move adults into new vials with fresh media. Allow flies to lay eggs in a dark environment for approximately 4 hr before again transferring the crosses into new vials. Note the start and end times of the egg laying period. 
	Note: Shorter intervals are acceptable when there appears to be a high number of eggs on the media before the end of the 4-hr interval.
	<ol>
		<li>Keep vials with a sufficient number of eggs (approximately greater than 20) and store in a 12-hr light/dark cycle at 22 degrees Celsius to be heat shocked later. 
		Note: Vials with a low number of eggs on the media (less than approximately 20) are unlikely to yield sufficient numbers of adult flies for behavioral testing and in the interest of efficiency it is recommended they not be retained for further testing. 
		Note: Our flies are raised with the lights on from 8 AM to 8 PM. We have found the evening time points lead to higher egg densities, therefore usually perform timed crosses between 4 - 8 PM and 8 PM-midnight. The flies lay their eggs at RT&#160;in a dark cabinet and then are returned to the incubator the following day so as not to disturb the regular light-dark cycle in the incubator.</li>
	</ol>
	</li>
</ol>
3. Perform Heat Shock to Induce Mitotic Recombination
<ol>
	<li>Approximately 85 - 100 hr after egg laying, place vials into a water bath at 37 degrees Celsius for 1 hr. Ensure that the water level reaches above the height of the media in the vials, but does not fully submerge the vials to prevent the larvae from drowning.</li>
	<li>Remove vials from water bath and allow a 1 hr recovery period at RT.</li>
	<li>Place vials back into 37 degrees Celsius water bath for 1 hr.</li>
	<li>Remove vials from water bath and store in incubator on 12-hr light/dark cycle at 22 degrees Celsius until eclosion.</li>
</ol>
4. Preparation for Behavioral Testing
<ol>
	<li>Following eclosion, anesthetize flies on ice. Select flies with genotypes that contain all the genetic elements for MARCM and indicate GFP expression-marked clones will likely be observed based on the observable phenotype markers used in the parent stocks. Decapitate flies with iridectomy scissors. 
	Note: For example, we selected for flies without the genetic markers CyO (curly wings) and Sb (stubble bristle) to identify flies that would contain all the genetic elements necessary to create mosaic regions marked with GFP. 
	Note: Decapitation is necessary to prevent the adults from flying away when stimulated.</li>
	<li>Place headless flies in a closed, moist environment and allow approximately 10 - 20 hr of recovery. Only use flies that right themselves when perturbed for further testing. Test flies within 30 hr of decapitation.</li>
</ol>
5. Behavioral Testing
<ol>
	<li>By observing decapitated flies under a fluorescence dissecting microscope, identify homozygous clones marked with GFP at the bristle external sensory organs. Record the bristle name and left or right side. To eliminate any potential bias, the lab member performing the bristle stimulation should be blind to the genotype of the bristle, wild-type or mutant.</li>
	<li>Elicit a grooming reflex by deflecting the GFP marked bristle towards the fly body with a stiff hair or fine forceps and observe the leg response. Give a score of one to flies that lift their leg in response to bristle stimulation and a score of zero to flies that do not move their legs. 
	Note: Previous studies6,7 have characterized the leg response that is elicited by stimulating a particular bristle. This includes the leg that responds to each bristle and the proportion of flies that respond to stimulation of that particular bristle. We only scored for movements of the leg that we expected to respond based on Vandervorst&#39;s classification6.</li>
	<li>Conduct five trials spaced 2 min&#160;apart.</li>
</ol>

<ol>
	<li>Gr&#252;nert, U., Gnatzy, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=K++and+Ca+++in+the+receptor+lymph+of+arthropod+cuticular+mechanoreceptors.">K+ and Ca++ in the receptor lymph of arthropod cuticular mechanoreceptors.</a> J Comp Physiol A. 161, 329-333 (1987).</li><li>Kernan, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanotransduction+and+auditory+transduction+in+Drosophila.">Mechanotransduction and auditory transduction in Drosophila.</a> Pflugers Arch. 454, 703-720 (2007).</li><li>Gillespie, P. G., Walker, R. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+basis+of+mechanosensory+transduction.">Molecular basis of mechanosensory transduction.</a> Nature. 413, 194-202 (2001).</li><li>Corfas, G., Dudai, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adaptation+and+fatigue+of+a+mechanosensory+neuron+in+wild-type+Drosophila+and+in+memory+mutants.">Adaptation and fatigue of a mechanosensory neuron in wild-type Drosophila and in memory mutants.</a> J Neurosci. 10, 491-499 (1990).</li><li>Kernan, M., Cowan, D., Zuker, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+dissection+of+mechanosensory+transduction:+mechanoreception-defective+mutations+of+Drosophila.">Genetic dissection of mechanosensory transduction: mechanoreception-defective mutations of Drosophila.</a> Neuron. 12, 1195-1206 (1994).</li><li>Vandervorst, P., Ghysen, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+control+of+sensory+connections+in+Drosophila.">Genetic control of sensory connections in Drosophila.</a> Nature. 286, 65-67 (1980).</li><li>Corfas, G., Dudai, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Habituation+and+dishabituation+of+a+cleaning+reflex+in+normal+and+mutant+Drosophila.">Habituation and dishabituation of a cleaning reflex in normal and mutant Drosophila.</a> J Neurosci. 9, 56-62 (1989).</li><li>Chen, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Discovery-based+science+education:+functional+genomic+dissection+in+Drosophila+by+undergraduate+researchers.">Discovery-based science education: functional genomic dissection in Drosophila by undergraduate researchers.</a> PLoS Biol. 3, e59 (2005).</li><li>Lee, T., Luo, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mosaic+analysis+with+a+repressible+cell+marker+for+studies+of+gene+function+in+neuronal+morphogenesis.">Mosaic analysis with a repressible cell marker for studies of gene function in neuronal morphogenesis.</a> Neuron. 22, 451-461 (1999).</li><li>Wu, J. S., Luo, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+protocol+for+mosaic+analysis+with+a+repressible+cell+marker+(MARCM)+in+Drosophila.">A protocol for mosaic analysis with a repressible cell marker (MARCM) in Drosophila.</a> Nature Protocols. 1, 2583-2589 (2006).</li><li>Eberl, D. F., Hardy, R. W., Kernan, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetically+similar+transduction+mechanisms+for+touch+and+hearing+in+Drosophila.">Genetically similar transduction mechanisms for touch and hearing in Drosophila.</a> J Neurosci. 20, 5981-5988 (2000).</li><li>Walker, R. G., Willingham, A. T., Zuker, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Drosophila+mechanosensory+transduction+channel.">A Drosophila mechanosensory transduction channel.</a> Science. 287, 2229-2234 (2000).</li><li>Yan, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+NOMPC+is+a+mechanotransduction+channel+subunit+for+gentle-touch+sensation.">Drosophila NOMPC is a mechanotransduction channel subunit for gentle-touch sensation.</a> Nature. 493, 221-225 (2013).</li><li>Arnadottir, J., Chalfie, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Eukaryotic+Mechanosensitive+Channels.">Eukaryotic Mechanosensitive Channels.</a> Annual Review of Biophysics. 39, 111-137 (2010).</li><li>Christensen, A. P., Corey, D. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=TRP+channels+in+mechanosensation:+direct+or+indirect+activation?.">TRP channels in mechanosensation: direct or indirect activation?.</a> Nature Reviews Neuroscience. 8, 510-521 (2007).</li><li>Cheng, L. E., Song, W., Looger, L. L., Jan, L. Y., Jan, Y. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Role+of+the+TRP+Channel+NompC+in+Drosophila+Larval+and+Adult+Locomotion.">The Role of the TRP Channel NompC in Drosophila Larval and Adult Locomotion.</a> Neuron. 67, 373-380 (2010).</li><li>Lee, J., Moon, S., Cha, Y., Chung, Y. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+TRPN(=NOMPC)+channel+localizes+to+the+distal+end+of+mechanosensory+cilia.">Drosophila TRPN(=NOMPC) channel localizes to the distal end of mechanosensory cilia.</a> PLoS One. 5, e11012 (2010).</li><li>Liang, X., Madrid, J., Saleh, H. S., Howard, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NOMPC,+a+Member+of+the+TRP+Channel+Family,+Localizes+to+the+Tubular+Body+and+Distal+Cilium+of+Drosophila+Campaniform+and+Chordotonal+Receptor+Cells.">NOMPC, a Member of the TRP Channel Family, Localizes to the Tubular Body and Distal Cilium of Drosophila Campaniform and Chordotonal Receptor Cells.</a> Cytoskeleton. 68, 1-7 (2011).</li><li>de Vries, S. E., Clandinin, T. 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The authors have no competing financial interests.

This protocol utilizes an adult behavioral assay to screen for mutations that affect mechanosensation in Drosophila. Because the collection of mutants contains lethal p-element mutations that would preclude screening as adults, this protocol makes use of a complex genetic technique first described by Lee and Luo, (1999) and detailed as a protocol by Wu and Luo, (2006) to circumvent adult lethality. MARCM induces mitotic recombination between homologous chromosomes to generate clonal regions of homozygous cells. ...

Humans rely on the ability to convert mechanical stimuli from their environment, such as touch, pressure, vibration, or sound waves, into sensory information that can be processed by the nervous system, in a process termed mechanotransduction. Many of the overall mechanistic features of mechanotransduction between humans and invertebrates are the same1, making Drosophila a useful model to study the molecular mechanisms of mechanotransduction. Drosophila melanogaster contain two sets of specialized sensory organs (Type I and II) that are capable of converting mechanical stimuli into action potentials. Type I mechanoreceptors have a neuron with a single dendrite or sensory process, surrounded by three support cells2. The Type I mechanoreceptors include bristle mechanoreceptors, hearing sensitive chordotonal organs (the Johnston's organ), and campaniform sensilla that convey information about wing beats3. The bristles that cover the dorsal side of the fly are the most abundant and easily accessible of the Type I organs. Similar to the extracellular environment of hair cells associated with hearing and balance in vertebrates, the support cells surrounding fly mechanosensitive neurons secrete a high potassium endolymph that creates an unusual concentration gradient for potassium1. Mechanosensitive neurons, of both the mammalian and fly systems, utilize this high extracellular potassium to depolarize the cell. In response to mechanical stimulation of the macrochaete bristles towards the body wall, the sensory neurons respond with a burst of action potentials driven by this potassium depolarization of the cell membrane4. The Drosophila sensory neurons that innervate bristles resemble the mechanosensitive cells of other organisms, including vertebrate hair cells, in both structure and function1,5. The accessibility of Drosophila external sense organs to experimental manipulation and the abundance of genetic techniques available to researchers make Drosophila an excellent model system to investigate the molecular underpinnings of mechanosensation. 
In the fly, stimulation of a single sensory neuron that innervates a bristle leads to an observable behavioral response. Stimulation of different bristles evokes specific, reproducible behavioral responses, depending on the bristle that is stimulated. Upon tactile stimulation, decapitated wild-type flies exhibit a complex grooming reflex wherein they clean the area near the stimulated bristle with a patterned set of leg movements5-7. When homozygous for known single-gene mutations that interfere with learning7 or coordination and locomotor activity5, flies respond abnormally to mechanical stimulation. This grooming reflex is therefore a useful tool to study the effects of single-gene mutations on a specific, replicable behavior.
The robust behavioral response to stimulation of a single macrochaete bristle holds the potential to assist in identifying new genes involved in mechanotransduction. This protocol being used to test a collection of mutant flies for the absence of a behavioral response to indicate that the mutation interferes with mechanosensation. In the mutant collection selected for screening, the mutations cause lethality before adulthood, and therefore would be impossible to test using traditional adult behavior screens. Originally, this collection of lethal p-elements was combined with FRT recombination sites to test cell growth defects in clones. The clones were made specifically in the eye because adult flies can survive in the lab setting without functional vision8. However, removal of all mechanosensation can cause adults to be severely uncoordinated or die before eclosion5. This protocol uses a mosaic approach to circumvent the lethality of the mutations and allow for adult stage testing. A genetic technique called Mosaic Analysis with a Repressible Cell Marker (MARCM)9 is used to generate homozygous mutant cells in a limited number of adult fly sensory organs, while the rest of the organism remains heterozygous. These MARCM flies readily survive until adulthood, yet the bristles are homozygous for the lethal genes of interest.
MARCM allows for regions of homozygous mutant cells to be generated and marked by the expression of green fluorescent protein (GFP), while the rest of the organism remains heterozygous at that particular locus and unmarked9. MARCM combines individual p-element mutations with five common genetic elements: GFP under the control of an upstream-activating sequence (UAS-GFP), Gal80 repressor protein under control of a general promoter (tub-Gal80), Gal4 transcription factor under control of a general promoter (tub-Gal4), the FRT recombinase enzyme expressed through a heat-shock controlled promoter, and a FRT recombination site10. By driving mitotic recombination through a heat-shock activated recombinase, a limited number of cells are made homozygous for the mutation and marked with GFP. GFP expression is repressed in heterozygous cells by the presence of Gal80 on the wild-type copy of the chromosome. 
A heat-shock protocol for MARCM was optimized to induce recombination in the fly bristle external sense organs, while much of the organism remains heterozygous, and thus unmarked with GFP. Mosaic flies generated using this protocol contained homozygous mutations most frequently in the post alar or dorsal central bristles on the surface of the notum. 
We have tested the utility of this combination of MARCM and the grooming behavior screen with a known mechanosensitive mutant, NOMPC. The ion channel, no mechanoreceptor potential C (NOMPC), is an essential component of the mechanotransduction pathway in Drosophila5,11-13. NOMPC belongs to the transient receptor potential (TRP) superfamily of cation channels5 and satisfies all of the criteria to qualify as a mechanosensitive channel in Drosophila14,15: 1) NOMPC is expressed in the ciliate tips of type 1 sensory neurons of Drosophila 13,16-18, 2) NOMPC null larvae do not have an electrical response to tactile stimulation13, 3) Ectopic expression of NOMPC in touch insensitive cells can induce sensitivity to mechanical stimulation13, 4) heterologous expression of NOMPC in Schneider 2 cells yields a mechanosensitive channel13, and 5) NOMPC adult mutants display defects in their response to mechanical stimulation5. Given this evidence, we predicted that NOMPC mutant clones would show an altered or inhibited grooming response in response to mechanical stimulation of bristles.
A MARCM-stock containing the NOMPC mutation was developed for use in a proof of principle experiment of the grooming assay. Mosaic flies were stimulated at macrochaete bristles containing homozygous NOMPC mutant cells. We expected an inhibition of the grooming response following stimulation of the macrochaete bristle. We found that only 2 of 14 mutant bristle flies tested gave a single response to repeated stimulation; most did not respond to stimulation of the homozygous mutant bristle. Having confirmed that this MARCM-based behavioral assay produces an abnormal grooming reflex in a known mechanosensitive mosaic mutant, this technique can be used in a screen for additional mechanosensitive mutations.

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		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:163px;">Ice bucket</td>
			<td style="width:163px;"></td>
			<td style="width:163px;"></td>
			<td style="width:163px;">for cold anthesia</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:163px;">Homemade cold anthesia tray</td>
			<td style="width:163px;"></td>
			<td style="width:163px;"></td>
			<td style="width:163px;">for cold anthesia decapitation</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:163px;">Plastic boxes</td>
			<td style="width:163px;"></td>
			<td style="width:163px;"></td>
			<td style="width:163px;">for recovery of decaptitated flies in humid environment</td>
		</tr>
	</tbody>
</table>

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.

The success of this protocol largely depends on the overall yield of testable flies, the effectiveness of the heat shock to induce mitotic recombination, and the ability to obtain a robust behavioral response in flies stimulated at 2 min intervals. Given that an estimated 12.5% of embryos contain the necessary MARCM elements and an estimated 16.67% of adults that eclose have the potential for GFP expression, it is critical to obtain a high number of eggs during timed lays. An example of a...

Watch this Scientific Journal Video about A Behavioral Assay for Mechanosensation of MARCM-based Clones in Drosophila melanogaster at JoVE.com

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.

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 ...

a-behavioral-assay-for-mechanosensation-marcm-based-clones-drosophila

Research

JoVE Journal

Neuroscience

10.1K Views.  College of the Holy Cross. 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.

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

Magnetic Resonance Spectroscopy of live Drosophila melanogaster using Magic Angle Spinning

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

Magnetic Resonance Spectroscopy of live Drosophila melanogaster using Magic Angle Spinning

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

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

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 ...

Paired Nanoinjection and Electrophysiology Assay to Screen for Bioactivity of Compounds using the Drosophila melanogaster Giant Fiber System

Screening compounds for in vivo activity can be used as a first step to identify candidates that may be developed into pharmacological agents1,2. We developed a novel nanoinjection/electrophysiology assay that allows the detection of bioactive modulatory effects of compounds on the function of a neuronal circuit that mediates the escape response in Drosophila melanogaster3,4. Our in vivo assay, which uses the Drosophila Giant Fiber System (GFS, Figure 1) allows screening of different types of compounds, such as small molecules or peptides, and requires only minimal quantities to elicit an effect. In addition, the Drosophila GFS offers a large variety of potential molecular targets on neurons or muscles. The Giant Fibers (GFs) synapse electrically (Gap Junctions) as well as chemically (cholinergic) onto a Peripheral Synapsing Interneuron (PSI) and the Tergo Trochanteral Muscle neuron (TTMn)5. The PSI to DLMn (Dorsal Longitudinal Muscle neuron) connection is dependent on D&alpha;7 nicotinic acetylcholine receptors (nAChRs)6. Finally, the neuromuscular junctions (NMJ) of the TTMn and the DLMn with the jump (TTM) and flight muscles (DLM) are glutamatergic7-12. Here, we demonstrate how to inject nanoliter quantities of a compound, while obtaining electrophysiological intracellular recordings from the Giant Fiber System13 and how to monitor the effects of the compound on the function of this circuit. We show specificity of the assay with methyllycaconitine citrate (MLA), a nAChR antagonist, which disrupts the PSI to DLMn connection but not the GF to TTMn connection or the function of the NMJ at the jump or flight muscles.
Before beginning this video it is critical that you carefully watch and become familiar with the JoVE video titled "Electrophysiological Recordings from the Giant Fiber Pathway of D. melanogaster " from Augustin et al7, as the video presented here is intended as an expansion to this existing technique. Here we use the electrophysiological recordings method and focus in detail only on the addition of the paired nanoinjections and monitoring technique.

Paired Nanoinjection and Electrophysiology Assay to Screen for Bioactivity of Compounds using the Drosophila melanogaster Giant Fiber System

A rapid in vivo assay to test for neuromodulatory compounds using the Giant Fiber System (GFS) of Drosophila melanogaster is described. Nanoinjections in the head of the animal along with electrophysiological recordings of the GFS can reveal bioactivity of compounds on neurons or muscles.

Screening compounds for in vivo activity can be used as a first step to identify candidates that may be developed into pharmacological agents1,2. We ...

A Lightweight, Headphones-based System for Manipulating Auditory Feedback in Songbirds

Experimental manipulations of sensory feedback during complex behavior have provided valuable insights into the computations underlying motor control and sensorimotor plasticity1. Consistent sensory perturbations result in compensatory changes in motor output, reflecting changes in feedforward motor control that reduce the experienced feedback error. By quantifying how different sensory feedback errors affect human behavior, prior studies have explored how visual signals are used to recalibrate arm movements2,3 and auditory feedback is used to modify speech production4-7. The strength of this approach rests on the ability to mimic naturalistic errors in behavior, allowing the experimenter to observe how experienced errors in production are used to recalibrate motor output.
Songbirds provide an excellent animal model for investigating the neural basis of sensorimotor control and plasticity8,9. The songbird brain provides a well-defined circuit in which the areas necessary for song learning are spatially separated from those required for song production, and neural recording and lesion studies have made significant advances in understanding how different brain areas contribute to vocal behavior9-12. However, the lack of a naturalistic error-correction paradigm - in which a known acoustic parameter is perturbed by the experimenter and then corrected by the songbird - has made it difficult to understand the computations underlying vocal learning or how different elements of the neural circuit contribute to the correction of vocal errors13.
The technique described here gives the experimenter precise control over auditory feedback errors in singing birds, allowing the introduction of arbitrary sensory errors that can be used to drive vocal learning. Online sound-processing equipment is used to introduce a known perturbation to the acoustics of song, and a miniaturized headphones apparatus is used to replace a songbird's natural auditory feedback with the perturbed signal in real time. We have used this paradigm to perturb the fundamental frequency (pitch) of auditory feedback in adult songbirds, providing the first demonstration that adult birds maintain vocal performance using error correction14. The present protocol can be used to implement a wide range of sensory feedback perturbations (including but not limited to pitch shifts) to investigate the computational and neurophysiological basis of vocal learning.

We describe the design and assembly of miniaturized headphones suitable for replacing a songbird&#x2019;s natural auditory feedback with a manipulated acoustic signal. Online sound processing hardware is used to manipulate song output, introduce real-time errors in auditory feedback via the headphones, and drive vocal motor learning.

Experimental manipulations of sensory feedback during complex behavior have provided valuable insights into the computations underlying motor control and ...

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

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

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

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

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

A Simple Behavioral Assay for Testing Visual Function in Xenopus laevis

Measurement of the visual function in the tadpoles of the frog, Xenopus laevis, allows screening for blindness in live animals. The optokinetic response is a vision-based, reflexive behavior that has been observed in all vertebrates tested. Tadpole eyes are small so the tail flip response was used as alternative measure, which requires a trained technician to record the subtle response. We developed an alternative behavior assay based on the fact that tadpoles prefer to swim on the white side of a tank when placed in a tank with both black and white sides. The assay presented here is an inexpensive, simple alternative that creates a response that is easily measured. The setup consists of a tripod, webcam and nested testing tanks, readily available in most Xenopus laboratories. This article includes a movie showing the behavior of tadpoles, before and after severing the optic nerve. In order to test the function of one eye, we also include representative results of a tadpole in which each eye underwent retinal axotomy on consecutive days. Future studies could develop an automated version of this assay for testing the vision of many tadpoles at once.

A Simple Behavioral Assay for Testing Visual Function in Xenopus laevis

Xenopus laevis tadpoles prefer swimming on the white side of a black/white tank. This behavior is guided by their vision. Based on this behavior, we present a simple assay to test the visual function of tadpoles.

Measurement of the visual function in the tadpoles of the frog, Xenopus laevis, allows screening for blindness in live animals. The optokinetic response ...

Straightforward Assay for Quantification of Social Avoidance in Drosophila melanogaster

Drosophila melanogaster is an emerging model to study different aspects of social interactions. For example, flies avoid areas previously occupied by stressed conspecifics due to an odorant released during stress known as the Drosophila stress odorant (dSO). Through the use of the T-maze apparatus, one can quantify the avoidance of the dSO by responder flies in a very affordable and robust assay. Conditions necessary to obtain a strong performance are presented here. A stressful experience is necessary for the flies to emit dSO, as well as enough emitter flies to cause a robust avoidance response to the presence of dSO. Genetic background, but not their group size, strongly altered the avoidance of the dSO by the responder flies. Canton-S and Elwood display a higher performance in avoiding the dSO than Oregon and Samarkand strains. This behavioral assay will allow identification of mechanisms underlying this social behavior, and the assessment of the influence of genes and environmental conditions on both emission and avoidance of the dSO. Such an assay can be included in batteries of simple diagnostic tests used to identify social deficiencies of mutants or environmental conditions of interest.

Straightforward Assay for Quantification of Social Avoidance in Drosophila melanogaster

Here, we present a protocol to quantify the avoidance of stressed individuals. This paradigm is powerful yet user-friendly and can be used to assess the influence of genes and environment on one kind of social interaction in Drosophila melanogaster.

Drosophila melanogaster is an emerging model to study different aspects of social interactions. For example, flies avoid areas previously occupied by ...

Dyeing Insects for Behavioral Assays: the Mating Behavior of Anesthetized Drosophila

Mating experiments using Drosophila have contributed greatly to the understanding of sexual selection and behavior. Experiments often require simple, easy and cheap methods to distinguish between individuals in a trial. A standard technique for this is CO2 anaesthesia and then labelling or wing clipping each fly. However, this is invasive and has been shown to affect behavior. Other techniques have used coloration to identify flies. This article presents a simple and non-invasive method for labelling Drosophila that allows them to be individually identified within experiments, using food coloring. This method is used in trials where two males compete to mate with a female. Dyeing allowed quick and easy identification. There was, however, some difference in the strength of the coloration across the three species tested. Data is presented showing the dye has a lower impact on mating behavior than CO2 in Drosophila melanogaster. The impact of CO2 anaesthesia is shown to depend on the species of Drosophila, with D. pseudoobscura and D. subobscura showing no impact, whereas D. melanogaster males had reduced mating success. The dye method presented is applicable to a wide range of experimental designs.

Dyeing Insects for Behavioral Assays: the Mating Behavior of Anesthetized Drosophila

This protocol describes a simple, cost effective way to individually identify Drosophila or other insects. Demonstration data investigating mating success across three species of Drosophila show that this method is comparable or better than the use of CO2 anaesthesia.

Mating experiments using Drosophila have contributed greatly to the understanding of sexual selection and behavior. Experiments often require simple, easy ...

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.

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.

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

Conditions Affecting Social Space in Drosophila melanogaster

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

Conditions Affecting Social Space in Drosophila melanogaster

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

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