We would like to thank Manuel Zimmer for providing us with the initial design file that was adapted for use with males; Frank Schroeder for the synthesis and supply of ascr#3; Ross Lagoy for the insight and assistance with imaging and analysis; and Laura Aurilio for the master fabrication and who, alongside Christopher Chute, contributed to the review of this manuscript. Funding for this work was provided under the National Institutes of Health grant 1R01DC016058-01 (J.S.), the National Science Foundation grant CBET 1605679 (D.R.A.), and the Burroughs Wellcome Career Award at the Scientific Interface (D.R.A.).

1. Device Fabrication
NOTE: See reference1.
NOTE: Silicon master molds were fabricated using standard photolithographic techniques for patterning SU-8 photoresist on a silicon master1,7. Photomasks for wafer patterning were printed at 25,000 dpi. The male-adapted device features a Chronis Olfactory Chip design19 with a change in the worm loading port, adapting a design obtained from M. Zimmer (personal correspondence, 2016). A turn is included to control the rotation of the animals. The width of the worm loading port channel is narrowed to 50 &#956;m. All channels are 32 &#956;m tall. Once a silicon master mold is available to the user, the user can follow the subsequent protocol, as described previously1.
<ol>
	<li>Mix PDMS base and curing agent at a 10:1 ratio by weight.</li>
	<li>Mix thoroughly with transfer pipettes.</li>
	<li>Degas the mixture in a vacuum desiccator for 1 h, until all visible bubbles are removed.</li>
	<li>Pour the mixture onto a silicon mold master in a 150 mm diameter dish until it is 5 mm thick (100 g). Use a Pasteur pipette to remove any bubbles or dust that have been introduced to the mixture.</li>
	<li>Bake at 65 &#176;C for at least 3 h, or overnight.</li>
	<li>Cut the PDMS away from the mold using a scalpel and cut the separate devices apart using a razor blade.</li>
	<li>Punch inlet and outlet holes with a 1 mm dermal punch.</li>
	<li>Flush the holes with dH2O, ethanol, and again with dH2O to remove particles from the punches. Dry the device in an air stream pulse.</li>
	<li>Clean both channel sides and the top side of the device with adhesive tape, removing any dust or debris remaining on the device to allow for successful bonding.</li>
	<li>Plasma-bond the device, channel-side down, to a no. 1 cover glass.
	<ol>
		<li>Expose cover glass and device (channel-side up) to air plasma using conditions that allow for proper bonding, such as 100 W for 30 s or 24 W for 60 s. 
		NOTE: Settings can be adjusted to improve the bonding efficiency. The plasma-bonding conditions are not as critical as proper cleaning when attempting to improve the bonding efficiency. An insufficiently cleaned device will not bond, even under ideal plasma conditions.</li>
		<li>Invert the cover glass onto the channel side of the device and press down with the thumb for 5 s.</li>
	</ol>
	</li>
</ol>
2. Buffer Preparation
<ol>
	<li>Dilute 1x S Basal (100 mM NaCl and 0.05 M KPO4, pH 6.0) from a sterile 10x stock.</li>
	<li>Dilute 1 M tetramisole stock to a final concentration of 1 mM in 1x S Basal for all buffer solutions.</li>
	<li>Add fluorescein to both the &#34;flow control&#34; and &#34;buffer&#34; reservoirs.
	<ol>
		<li>Create a 100-mg/mL stock of fluorescein in 1x S Basal.</li>
		<li>Dilute the stock to final concentrations of 1 &#181;g/mL in the flow control and 0.1 &#181;g/mL in the buffer.</li>
	</ol>
	</li>
	<li>Create the stimuli.
	<ol>
		<li>Dilute glycerol to a final concentration of 1 M in 1X S Basal.</li>
		<li>Dilute ascaroside #3 (ascr#3) to a final concentration of 1 &#181;M into 1X S Basal.</li>
	</ol>
	</li>
</ol>
3. Device Setup
NOTE: See1.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56026/56026fig1.jpg" /> 
Figure 1. Microfluidic device setup. (A) Reservoirs and tubing. A 30 mL syringe without a plunger serves as the &#34;reservoir.&#34; This is attached to a Luer valve with three flow options. One outlet is connected to a 3 mL syringe with a plunger, while the other is connected to a needle (orange) that is inserted into the tubing that connects to the microfluidic device. (B) The overall setup of the microfluidic imaging experiment. The device is placed on a stage of an inverted epifluorescence microscope, above the objective lenses. The &#34;flow control&#34; buffer travels through a 3-way valve that is controlled by a unit on the shelf above the setup. Lines containing buffers are then inserted into the appropriate device ports. (C) The ports of the microfluidic device. The &#34;flow control&#34; ports flank the other inlet ports: the &#34;stimulus&#34; and &#34;buffer&#34; ports. The &#34;outlet&#34; port is the right-most port. Due to the location of the worm loading arena, the &#34;worm loading&#34; port is the central-most port on the device. <a href="//cloudfront.jove.com/files/ftp_upload/56026/56026fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Prepare three fluid reservoirs by attaching a 30 or 60 mL syringe to 3-way Luer valve, with a 3 mL syringe and needle attached to the Luer valve as well (as in Figure 1A). Connect the needle to tubing that extends to the microfluidic device (as in Figure 1A-B).</li>
	<li>Remove the air bubbles from the reservoir and tubing.</li>
	<li>Fill the 3 mL syringe with attached tubing with 1x S Basal and insert it into the outlet port.</li>
	<li>Gently apply pressure to the syringe until the buffer appears at the top of the inlet holes.</li>
	<li>Connect the flow control, buffer, and stimulus tubing to appropriate inlet holes (as in Figure 1B-C), ensuring that liquid drops are present on both the loading port hole and the buffer tubing to be attached.</li>
	<li>Again, gently apply pressure to the syringe that is connected to the outlet port until droplets appear in the worm loading port inlet.</li>
	<li>Insert a solid blocking pin into the worm loading port.</li>
	<li>Remove the syringe from the outlet port and attach the outlet line connected to the house vacuum (-670 Torr).</li>
	<li>Inspect the device for any bubbles in the flow channels, visually and through video confirmation via a software compatible with the camera used, such as the open-source software Micro-Manger. See step 6 for tips on using Micro-Manager.
	<ol>
		<li>If any bubbles are present, wait for them to dislodge or be absorbed into the PDMS wall prior to loading any animals; the presence of bubbles will disturb the proper flow of fluids through the device.</li>
	</ol>
	</li>
	<li>Using a GFP filter, confirm proper flow dynamics within the device prior to worm loading by actuating the 3-way valve and observing the switching of buffers.
	<ol>
		<li>Determine the proper flow dynamics: observe the fluorescein present in the flow control and buffer solutions (Figure 2D-2E) changing when the flow control value is changed by pressing the control button corresponding to the 3-way valve on the perfusion controller software (Figure 1B).</li>
		<li>After opening Micro-Manager, click on &#34;Live&#34; to observe a live image of the device. Turn on the fluorescent light source to observe the flow of buffers in the device (Figure 2D-2E).</li>
	</ol>
	</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56026/56026fig2.jpg" /> 
Figure 2: A male-adapted microfluidic olfactory chip. (A) The flow patterns of the device when the worm is exposed to buffer. Buffer (B) is shown in brown, and flow control (FC) is shown in yellow, with stimulus (S) in white. The worm loading port has been adapted to include a curve, which allows for better control of worm orientation. (B) The flow patterns of the device when the worm is exposed to stimulus. Buffer (B) is shown in brown, and flow control (FC) is shown in yellow, with stimulus (S) in white. (C) Measurements of the adapted device as fabricated. The worm loading port ends in a 42 &#181;m opening, with a 50 &#181;m channel designed for the male width. The measured height of the channels is 32 &#181;m, despite a target of 25 &#181;m in the design. (D-E) A trapped male expressing psra-6::GCaMP3. The sra-6 promoter is not ASH-specific, and some expression may be observed in the ASI neuron, although no calcium transients were observed in ASI. The image is (D) a combination of bright-field and fluorescent illumination, while (E) is fluorescent only. The scale bars denote 42 &#181;m. (F) The ASH neuron responds to 1 M glycerol stimulation with robust neural activity. The blue area denotes the time of the 1 M glycerol stimulus. The shaded region denotes the standard error, with n = 20 pulses from seven worms. The red traces denote depolarizing responses. The Y-axes show &#916;F/F0. The scale bar denotes 5 s. <a href="//cloudfront.jove.com/files/ftp_upload/56026/56026fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Animal Preparation
NOTE: See reference23.
<ol>
	<li>Imaging ASH responses to 1 M glycerol.
	<ol>
		<li>Place approximately 20 C. elegans males that are positive for psra-6::GCaMP3 array expression onto a nematode growth medium (NGM) agar plate seeded with a lawn of OP50 E. coli. Use the expression of fluorescent GECI and/or a co-injection marker for the identification of array-positive animals. 
		NOTE: Array positive animals will fluoresce according to the GECI used (i.e., animals expressing GCaMP will fluoresce green under blue-light stimulation, while RCaMP animals will fluoresce red under green-light stimulation). Co-injection markers can range from other fluorescent proteins, such as GFP and RFP, to phenotypic markers, such as rol-6, or can rescue a dominant phenotype, such as the pha-1 mutation28.
		<ol>
			<li>If picking immediately prior to the assay, pick young adult males. If picking the day prior to the assay, pick L4 larval males.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Imaging the CEM responses to 1 &#181;M ascr#3.
	<ol>
		<li>Pick approximately 20 L4 C. elegans males (fkEx98[ppkd-2::GCaMP::SL2::dsRED + pBX-1]; pha-1(e2123ts); him-5(e1490); lite-1(ce314)) that are positive for dsRed co-injection marker expression. 
		NOTE: dsRed expression within the ray neurons of the male tale is easier to observe and confirm than GCaMP expression within the four CEM neurons.</li>
		<li>Isolate these males from hermaphrodites on an NGM agar plate seeded with a lawn of OP50 E. coli for 5-14 h before performing the imaging experiment. 
		NOTE: Males not isolated for a minimum of 5 h do not behaviorally respond to ascr#3 and therefore may exhibit even fewer calcium transients to the ascaroside than observed here.</li>
	</ol>
	</li>
</ol>
5. Animal Loading
NOTE: See refefence1.
<ol>
	<li>Pick one worm onto an unseeded NGM agar plate using standard worm maintenance techniques.
	<ol>
		<li>Pick worms by flaming a pick (made of flattened platinum wire), picking bacteria onto the pick, and &#34;dabbing&#34; a worm to pick it up. Gently place the worm onto the new plate, allowing it to crawl off on its own.</li>
	</ol>
	</li>
	<li>Add approximately 5 mL of 1x S Basal to the unseeded plate, such that plate is flooded.</li>
	<li>Draw the worm into a loading syringe (i.e., 3 mL syringe with attached tubing) that has been pre-filled with 1x S Basal.
	<ol>
		<li>Be sure to suck the worm only into the tubing, not all the way into the syringe. 
		NOTE: If the worm travels into the syringe, it is near impossible to get it back into the tubing.</li>
	</ol>
	</li>
	<li>Turn off the vacuum to stop the flow by turning the outlet Luer valve.</li>
	<li>Remove the solid pin blocking the worm loading port.</li>
	<li>Turn the Luer valve connected to the outlet port (Figure 1B) so that it is venting. 
	NOTE: Use a live video feed while loading the worm to confirm the location and orientation of the animal (steps 5.8-5.13).</li>
	<li>Insert the worm loading tube into the worm loading port.</li>
	<li>Gently apply pressure to the syringe until the worm appears in the loading channel.</li>
	<li>If the worm starts to enter the channel tail-first, pull on the syringe plunger to prevent the worm from entering the channel.</li>
	<li>Switch between applying and reversing pressure until the head enters the channel first.</li>
	<li>Open the vacuum by turning the 3-way Luer valve connected to the outlet port to open it to vacuum instead of atmosphere.</li>
	<li>Manually apply pressure by depressing the syringe plunger to orient and place the worm head such that it is exposed to the buffer flow channel, but not so far that the head can move around freely (Figure 2D-2E).</li>
</ol>
6. Stimulus and Acquisition
<ol>
	<li>Using an open-source microscopy software, such as Micro-Manager, record by capturing images as a TIFF stack at 10 frames/s using blue-light excitation (470 nm) for 30 s.</li>
	<li>Set the exposure on the main menu to 100 ms.
	<ol>
		<li>Open &#34;Multi-D Acq.&#34; from the main menu of the software. Set the &#34;number&#34; to &#34;300,&#34; and the &#34;interval&#34; to &#34;0.&#34; Click &#34;Acquire!&#34; to acquire the video.</li>
	</ol>
	</li>
	<li>Apply a 10 s pulse of the stimulus 5 s after initiating acquisition. Adjust the duration of stimulus application as desired.
	<ol>
		<li>After acquiring 5 s of video, change the 3-way valve controlling the flow control buffer to apply the stimulus to the animal being tested. Click the left-most button on the perfusion controller software (Figure 1B).</li>
		<li>After 10 s of stimulus exposure (this time can be adjusted as desired by the user), alter the flow of buffers by again pressing the left-most button on the perfusion controller software.</li>
	</ol>
	</li>
	<li>Record under buffer only until the 30-s window is complete to allow the GECI fluorescence to return to baseline.</li>
	<li>Repeat as desired. Wait 30 s between the end of acquisition and the initiation of the next trial.</li>
</ol>
7. Image Analysis
<ol>
	<li>Open the TIFF stack with the open-source software, ImageJ, by dragging file into the ImageJ window.</li>
	<li>Click using the cursor and drag to set the region of interest (ROI) around the neuron of interest. Set the region to include the soma of the neuron of interest (as in Figure 3A).</li>
	<li>Plot the z-stack of the fluorescence intensity of the ROI across stacks by clicking Open -&#62; Image -&#62; Stacks -&#62; Plot Z-axis Profile.</li>
	<li>Click &#34;List&#34; in the window that opens. Click Edit -&#62; Copy to copy the values. Paste the values into a spreadsheet program.</li>
	<li>Analyze the background fluorescence for each pulse by dragging the ROI to a region of the worm that does not contain GCaMP expression.</li>
	<li>Perform background subtraction for each pulse by subtracting the background fluorescence value from the neuron fluorescence intensity value.</li>
	<li>Calculate &#916;F/F0 for each frame of each pulse.
	<ol>
		<li>Calculate F0 as the average intensity value of the ROI for first 1 s of acquisition (e.g., frames 1-10).</li>
		<li>Calculate &#916;F/F0 by dividing the background-subtracted value for the frame of interest by the calculated F0 value.</li>
	</ol>
	</li>
	<li>Repeat for every neuron imaged and every stimulus pulse.</li>
	<li>For neurons with consistent response profiles, such as ASH, average all pulses for each neuron and calculate the SEM (as in Figure 2F).</li>
	<li>Plot the average &#916;F/F0 with SEM over time for each neuron. 
	NOTE: In this instance, it is common practice to include heatmaps of the individual neuronal responses of each trial as well. In neurons that do not exhibit consistent changes in calcium transients upon exposure to stimuli across repeated stimulations, or in different individuals23, it may be more applicable to show individual pulse traces (as in Figure 4). See the Discussion for details on determining how to display the data.</li>
</ol>

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E., Albertson, D. G., Thomson, J. 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+Caenorhabditis+elegans+male:+postembryonic+development+of+nongonadal+structures.">The Caenorhabditis elegans male: postembryonic development of nongonadal structures.</a> Dev Biol. 78 (2), 542-576 (1980).</li><li>Hilliard, M. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+imaging+of+C.+elegans+ASH+neurons:+cellular+response+and+adaptation+to+chemical+repellents.">In vivo imaging of C. elegans ASH neurons: cellular response and adaptation to chemical repellents.</a> The EMBO Journal. 24 (1), 63-72 (2005).</li><li>Evans, T. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transformation+and+microinjection.">Transformation and microinjection.</a> WormBook. , (2006).</li><li>C&#225;ceres, I. d. C., Valmas, N., Hilliard, M. A., Lu, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Laterally+Orienting+C.+elegans+Using+Geometry+at+Microscale+for+High-Throughput+Visual+Screens+in+Neurodegeneration+and+Neuronal+Development+Studies.">Laterally Orienting C. elegans Using Geometry at Microscale for High-Throughput Visual Screens in Neurodegeneration and Neuronal Development Studies.</a> PLoS ONE. 7 (4), e35037 (2012).</li><li>Schrodel, T., Prevedel, R., Aumayr, K., Zimmer, M., Vaziri, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Brain-wide+3D+imaging+of+neuronal+activity+in+Caenorhabditis+elegans+with+sculpted+light.">Brain-wide 3D imaging of neuronal activity in Caenorhabditis elegans with sculpted light.</a> Nat Methods. 10 (10), 1013-1020 (2013).</li><li>Garc&#237;a, L. R., Portman, D. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neural+circuits+for+sexually+dimorphic+and+sexually+divergent+behaviors+in+Caenorhabditis+elegans.">Neural circuits for sexually dimorphic and sexually divergent behaviors in Caenorhabditis elegans.</a> Curr Opin Neurobiol. 38, 46-52 (2016).</li><li>Clokey, G. V., Jacobson, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+autofluorescent+&amp;#34;lipofuscin+granules&amp;#34;+in+the+intestinal+cells+of+Caenorhabditis+elegans+are+secondary+lysosomes.">The autofluorescent &amp;#34;lipofuscin granules&amp;#34; in the intestinal cells of Caenorhabditis elegans are secondary lysosomes.</a> Mech Ageing Dev. 35 (1), 79-94 (1986).</li><li>Coburn, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anthranilate+Fluorescence+Marks+a+Calcium-Propagated+Necrotic+Wave+That+Promotes+Organismal+Death+in+C.+elegans.">Anthranilate Fluorescence Marks a Calcium-Propagated Necrotic Wave That Promotes Organismal Death in C. elegans.</a> PLoS Biology. 11 (7), e1001613 (2013).</li><li>Macosko, E. 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=A+hub-and-spoke+circuit+drives+pheromone+attraction+and+social+behaviour+in+C.+elegans.">A hub-and-spoke circuit drives pheromone attraction and social behaviour in C. elegans.</a> Nature. 458 (7242), 1171-1175 (2009).</li><li>Park, D., 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=Interaction+of+structure-specific+and+promiscuous+G-protein-coupled+receptors+mediates+small-molecule+signaling+in+Caenorhabditis+elegans.">Interaction of structure-specific and promiscuous G-protein-coupled receptors mediates small-molecule signaling in Caenorhabditis elegans.</a> Proc Natl Acad Sci U S A. 109 (25), 9917-9922 (2012).</li></ol>

The authors have nothing to disclose.

The male-adapted olfactory chip incorporates a turn into a narrower loading port, which allows for more control of the orientation and for the efficient trapping of male C. elegans. This allows for the visualization of both the left and right members of neuronal bilateral pairs, without the need for z-stacking. This curve leads to an orientation away from vertical 100% of the time in worms where only one bilateral pair is targeted with a fluorescent marker, such as ASH (Figure 2D<st...

Microfluidic devices provide increased access to precisely controlled environments, wherein animals, such as the nematode C. elegans, can be experimentally manipulated1. These studies include behavioral assays, calcium imaging studies, or even screenings for specific phenotypes, resulting in more exact measurements of experimental outcomes1,2,3,4,5,6. Microfluidics provide small-scale liquid conditions through which detailed experiments can be run while utilizing minimal amounts of reagents. There is a constant production of new microfluidic device designs, and the use of each varies, from arenas that allow for the natural sinusoidal motion of C. elegans in behavioral assays and neural imaging studies, to trap devices used in neural imaging and olfactory studies, to devices that allow for high-throughput phenotypic analysis in genetic screens4,5,6,7. Following the fabrication of a master mold, microfluidic devices are inexpensive to construct&#8212;given the reusability of the master&#8212;and easy to use, allowing for rapid data generation via high-throughput studies. The fabrication of devices using polymers such as polydimethylsiloxane (PDMS) allows for the creation of new devices within hours.
Calcium imaging studies use genetically encoded calcium indicators (GECIs) expressed in target cells to measure the neural dynamics of those cells in real time8,9,10,11. The transparent nature of C. elegans allows for the recording of the fluorescent levels of these proteins in live animals. Traditionally, GECIs rely on the green fluorescent protein (GFP)-based sensor GFP-Calmodulin-M13 Peptide (GCaMP), although more recent studies have adapted these sensors to allow for better signal-to-noise ratios and red-shifted excitation profiles. Following the development of GCaMP3, proteins with these specifications have varied, including sensors such as GCaMP6s and GCaMP6f (slow and fast fluorescence off-rates, respectively), as well as RFP-Calmodulin-M13 Peptide (RCaMP), which has a red-shifted activation profile. The combination of these GECIs with C. elegans cell-specific gene promoter sequences can target cells of interest, particularly sensory neurons12,13,14,15,16.
While the ease of C. elegans use in microfluidic studies is apparent, almost all studies have focused on hermaphrodites. Despite males only accounting for 0.01-0.02% of the wild type population, invaluable findings can arise from their characterization. While the physical connectome of the hermaphrodite nervous system has been fully mapped for decades17, the male connectome remains incomplete, especially in the head region of the animal18. The use of calcium imaging in males will help to generate an understanding of the male nervous system and the differences that arise between the two sexes. The smaller size of C. elegans adult males prevents effective and reliable trapping in the loading ports of traditional olfactory devices designed for larger hermaphrodites. To address this, a modified version of the Chronis Olfactory Chip19 was developed with a narrower loading port, a lower channel height, and turns in the worm loading port (which rotate the animal), allowing for the visualization of bilateral left/right neuronal pairs. This design permits: (1) the effective trapping of young adult males, (2) a more reliable orientation of the animal for the visualization of both members of bilateral paired neurons, and (3) the precise imaging of neural activity in male neurons.
Increasingly, studies show that C. elegans males respond differently than hermaphrodites to a variety of ascarosides (ascr), or nematode pheromones20,21,22,23,24. Therefore, developing an understanding of the neural dynamics and representations within the male connectome has become even more pertinent. Male C. elegans contain 87 sex-specific neurons not present in the hermaphrodite25,26, altering the connectome in as-yet undetermined ways. Being able to image these unique neural dynamics will allow us to better understand sex-specific responses and neural representations.
This protocol describes the use of a male-adapted olfactory chip for the neural imaging of male C. elegans chemosensation. The nociceptive neuron ASH responds reliably to 1 M glycerol in males, consistent with previous hermaphroditic studies27. Exposure to ascarosides may elicit responses that are variable from animal to animal, requiring a larger number of animals to be tested. The response of the male-specific CEM neurons has previously been shown, through both electrophysiology and calcium imaging studies, to respond variably to ascaroside #323.

<table width="2211">
	<tbody>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Silicon Wafer</td>
			<td style="width:219px;">University Wafer</td>
			<td style="width:205px;">452</td>
			<td style="width:1260px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">SU-8 2035</td>
			<td>MicroChem</td>
			<td>Y111070-0500L1GL</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Developer</td>
			<td>MicroChem</td>
			<td>Y020100-4000L1PE</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Wafer Mask</td>
			<td>Cad/Art Services</td>
			<td>-</td>
			<td>Custom order. Printed at 25,000 dpi.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sylgard-184</td>
			<td>Ellsworth Adhesives</td>
			<td>184 SIL ELAST KIT 0.5KG</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">1.0 mm Dermal Punches</td>
			<td>Acuderm Inc.</td>
			<td>P150</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Soft Tubing</td>
			<td>Cole-Palmer</td>
			<td>EW-06419-01</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Hard Tubing</td>
			<td>IDEX Health &#38; Science</td>
			<td>1622</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Pins</td>
			<td>New England Small Tube</td>
			<td>NE-1027-12</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Blocking Pins</td>
			<td>New England Small Tube</td>
			<td>0.415/0.425&#34; OD x .500 Long</td>
			<td>Batch PB07027</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">3 mL syringes</td>
			<td>BD</td>
			<td>309657</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">30 mL syringes</td>
			<td>Vitality Medical</td>
			<td>302832</td>
			<td>Used as buffer reservoirs.</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Stainless Steel Blunt Needle 23 Gauge, Polyprolylene Luer</td>
			<td>Component Supply Company</td>
			<td>NE-231PL-50</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Stopcocks with Luer connections; 3-way; male lock; 5 flow pattern; non-sterile</td>
			<td>Cole-Palmer</td>
			<td>EW-30600-07</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Fisherfinest Premium Cover Glass</td>
			<td>Fisher Scientific</td>
			<td>12-548-5M</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Mercator Control System LF-5 Plasma System</td>
			<td>Mercator</td>
			<td>LF-5</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Scotch Tape</td>
			<td>Scotch</td>
			<td>BSN43575</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Series 20 Chamber</td>
			<td>Warner Instruments</td>
			<td>P-2</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Vacuum Desicator</td>
			<td>Bel-Art Scienceware</td>
			<td style="width:205px;">420250000</td>
			<td>24 cm inner diameter.</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Weigh Boats</td>
			<td>Cole-Palmer</td>
			<td style="width:205px;">EW-01017-27</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Classic Plus Balance</td>
			<td>Mettler Toledo</td>
			<td style="width:205px;">PB1501-S/FACT</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Glass Pasteur Pipettes</td>
			<td>Cole-Palmer</td>
			<td style="width:205px;">EW-25554-06</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Transfer pipettes</td>
			<td>Genesee Scientific</td>
			<td style="width:205px;">30-202</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Oven</td>
			<td>Sheldon Manufacturing Inc</td>
			<td style="width:205px;">9120993</td>
			<td>Model Number: 1500E.</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">60 mm, non-vented, sharp edge Petri dishes</td>
			<td>TriTech Research</td>
			<td>T3308</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Zeiss Axio Observer.A1</td>
			<td>Zeiss</td>
			<td style="width:205px;">-</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Hammamatsu Orca Flash 4.0 Digital CMOS</td>
			<td>Hammamatsu</td>
			<td style="width:205px;">C11440-22CU</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Blue Fluorescent Light</td>
			<td>Lumencor</td>
			<td style="width:205px;">SOLA SM6-LCR-SA</td>
			<td>24-30V/7.9A DC.</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Illumination Adaptor</td>
			<td>Zeiss</td>
			<td style="width:205px;">423302-0000</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Series 1 and 2 Miniature Inert PTFE Isolation Valve</td>
			<td>Parker</td>
			<td style="width:205px;">001-0017-900</td>
			<td>3-way valve for controlling flow.</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">ValveLink8.2&#174;</td>
			<td>AutoMate Scientific</td>
			<td style="width:205px;">01-18</td>
			<td>Flow Switch Controller</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Micro Manager</td>
			<td>Micro-Manager</td>
			<td style="width:205px;">-</td>
			<td>Free software, can be downloaded at: https://www.micro-manager.org/wiki/Download_Micro-Manager_Latest_Release</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">ImageJ</td>
			<td>ImageJ</td>
			<td style="width:205px;">-</td>
			<td>Free software, can be downloaded at: https://imagej.nih.gov/ij/download.html</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Agar, Bacteriological Grade</td>
			<td>Apex</td>
			<td>9012-36-6</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Peptone</td>
			<td>Apex</td>
			<td>20-260</td>
			<td></td>
		</tr>
		<tr height="25">
			<td height="25" style="width:527px;height:25px;">CaCl2</td>
			<td>VWR</td>
			<td>BDH0224-1KG</td>
			<td></td>
		</tr>
		<tr height="25">
			<td height="25" style="width:527px;height:25px;">MgSO4</td>
			<td>Sigma-Aldrich</td>
			<td>230391-1kg</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Cholesterol</td>
			<td>Alfa Aesar</td>
			<td>A11470</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Ethanol</td>
			<td>Sigma-Aldrich</td>
			<td>270741-4L</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Tetramisole</td>
			<td>Sigma-Aldrich</td>
			<td style="width:205px;">L9756-10(G)</td>
			<td>Store at 4 &#176;C.</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Fluorescein</td>
			<td>Sigma-Aldrich</td>
			<td style="width:205px;">FD2000S-250mg</td>
			<td>Light Sensitive. Store in photoprotective vials.</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Glycerol</td>
			<td>Sigma-Aldrich</td>
			<td style="width:205px;">G6279-1L</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">Ascaroside #3</td>
			<td>-</td>
			<td style="width:205px;">-</td>
			<td>Synthesized in the Schroeder Lab (Cornell University).</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">NaCl</td>
			<td>Genesee Scientific</td>
			<td style="width:205px;">18-215</td>
			<td></td>
		</tr>
		<tr height="25">
			<td height="25" style="width:527px;height:25px;">KH2PO4</td>
			<td>BDH</td>
			<td>BDH9268.25</td>
			<td></td>
		</tr>
		<tr height="25">
			<td height="25" style="width:527px;height:25px;">K2HPO4</td>
			<td>J.T. Baker</td>
			<td>3252-025</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">ASH GCaMP3 line</td>
			<td>-</td>
			<td style="width:205px;">-</td>
			<td>CX10979 (KyEx2865 [psra-6::GCAMP3 @ 100 ng/uL]). Developed in Bargmann lab. Provided from Albrecht Lab library.</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">CEM GCaMP6 line</td>
			<td>-</td>
			<td style="width:205px;">-</td>
			<td>JSR49 (FkEx98[ppkd-2::GCaMP::SL2::dsRED + pBX-1]; pha-1(e2123ts); him-5(e1490); lite-1(ce314)). Developed by Robyn Lints. Provided from Srinivasan Lab library.</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">E. coli (OP50)</td>
			<td>Caenorhabditis Genetics Center</td>
			<td>OP50</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="width:527px;height:21px;">&#34;Reservoir&#34;</td>
			<td>-</td>
			<td>-</td>
			<td style="width:1260px;">To create a Reservoir: A &#34;30 mL syringe&#34;, is connected to a &#34;Stopcock with Luer connections; 3-way; male lock; 5 flow pattern; non-sterile&#34;, which is connected to a &#34;3 mL syringe&#34; and a &#34;Stainless Steel Blunt Needle 23 Gauge, Polyprolylene Luer&#34;. The &#34;Stainless Steel Blunt Needle 23 Gauge, Polyprolylene Luer&#34; is then inserted into &#34;Soft Tubing&#34; approximately 1/3 of the way down the needle.</td>
		</tr>
	</tbody>
</table>

using an adapted microfluidic olfactory chip for the imaging of neuronal activity in response to pheromones in male c. elegans head neurons

The use of calcium indicators has greatly enhanced our understanding of neural dynamics and regulation. The nematode Caenorhabditis elegans, with its completely mapped nervous system and transparent anatomy, presents an ideal model for understanding real-time neural dynamics using calcium indicators. In combination with microfluidic technologies and experimental designs, calcium-imaging studies using these indicators are performed in both free-moving and trapped animals. However, most previous studies utilizing trapping devices, such as the olfactory chip described in Chronis et al., have devices designed for use in the more common hermaphrodite, as the less common male is both morphologically and structurally dissimilar. An adapted olfactory chip was designed and fabricated for increased efficiency in male neuronal imaging with using young adult animals. A turn was incorporated into the worm loading port to rotate the animals and to allow for the separation of the individual neurons within a bilateral pair in 2D imaging. Worms are exposed to a controlled flow of odorant within the microfluidic device, as described in previous hermaphrodite studies. Calcium transients are then analyzed using the open-source software ImageJ. The procedure described herein should allow for an increased amount of male-based C. elegans calcium imaging studies, deepening our understanding of the mechanisms of sex-specific neuronal signaling.

An example of the overall device setup can be seen in Figure 1A-B. Figure 1A depicts the proper reservoir construction and setup. Figure 1B shows the connections of the reservoirs to the microfluidic device. Figure 1C depicts a microfluidic device with individual ports labeled for clarity.
The ...

Watch this Scientific Journal Video about Using an Adapted Microfluidic Olfactory Chip for the Imaging of Neuronal Activity in Response to Pheromones in Male C. Elegans Head Neurons at JoVE.com

Using an Adapted Microfluidic Olfactory Chip for the Imaging of Neuronal Activity in Response to Pheromones in Male C. Elegans Head Neurons

The use of an adapted &#34;olfactory chip&#34; for the efficient calcium imaging of C. elegans males is described here. Studies of male exposure to glycerol and a pheromone are also shown.

Using an Adapted Microfluidic Olfactory Chip for the Imaging of Neuronal Activity in Response to Pheromones in Male C. Elegans Head Neurons

The use of calcium indicators has greatly enhanced our understanding of neural dynamics and regulation. The nematode Caenorhabditis elegans, with its ...

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8.0K Views.  Worcester Polytechnic Institute. The use of an adapted "olfactory chip" for the efficient calcium imaging of C. elegans males is described here. Studies of male exposure to glycerol and a pheromone are also shown.

Video: Using an Adapted Microfluidic Olfactory Chip for the Imaging of Neuronal Activity in Response to Pheromones in Male C. Elegans Head Neurons

In vivo Laser Axotomy in C. elegans

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron is damaged by injury or disease, it may regenerate. Cell-intrinsic and extrinsic factors influence the ability of a neuron to regenerate and restore function. Recently, the nematode C. elegans has emerged as an excellent model organism to identify genes and signaling pathways that influence the regeneration of neurons1-6. The main way to initiate neuronal regeneration in C. elegans is laser-mediated cutting, or axotomy. During axotomy, a fluorescently-labeled neuronal process is severed using high-energy pulses. Initially, neuronal regeneration in C. elegans was examined using an amplified femtosecond laser5. However, subsequent regeneration studies have shown that a conventional pulsed laser can be used to accurately sever neurons in vivo and elicit a similar regenerative response1,3,7.
We present a protocol for performing in vivo laser axotomy in the worm using a MicroPoint pulsed laser, a turnkey system that is readily available and that has been widely used for targeted cell ablation. We describe aligning the laser, mounting the worms, cutting specific neurons, and assessing subsequent regeneration. The system provides the ability to cut large numbers of neurons in multiple worms during one experiment. Thus, laser axotomy as described herein is an efficient system for initiating and analyzing the process of regeneration.

In vivo Laser Axotomy in C. elegans

A protocol to cut neurons in C. elegans with a MicroPoint pulsed laser is presented. We describe setting up the system, immobilizing worms, and severing labeled neurons. Advantages include a relatively low-cost system and the ability to sever neuronal processes or ablate cells in vivo.

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron ...

A Molecular Readout of Long-term Olfactory Adaptation in C. elegans

During sustained stimulation most sensory neurons will adapt their response by decreasing their sensitivity to the signal. The adaptation response helps shape attention and also protects cells from over-stimulation. Adaptation within the olfactory circuit of C. elegans was first described by Colbert and Bargmann1,2. Here, the authors defined parameters of the olfactory adaptation paradigm, which they used to design a genetic screen to isolate mutants defective in their ability to adapt to volatile odors sensed by the Amphid Wing cells type C (AWC) sensory neurons. When wildtype C. elegans animals are exposed to an attractive AWC-sensed odor3 for 30 min they will adapt their responsiveness to the odor and will then ignore the adapting odor in a chemotaxis behavioral assay for ~1 hr. When wildtype C. elegans animals are exposed to an attractive AWC-sensed odor for ~1 hr they will then ignore the adapting odor in a chemotaxis behavioral assay for ~3 hr. These two phases of olfactory adaptation in C. elegans were described as short-term olfactory adaptation (induced after 30 min odor exposure), and long-term olfactory adaptation (induced after 60 min odor exposure). Later work from L'Etoile et al.,4 uncovered a Protein Kinase G (PKG) called EGL-4 that is required for both the short-term and long-term olfactory adaptation in AWC neurons. The EGL-4 protein contains a nuclear localization sequence that is necessary for long-term olfactory adaptation responses but dispensable for short-term olfactory adaptation responses in the AWC4. By tagging EGL-4 with a green fluorescent protein, it was possible to visualize the localization of EGL-4 in the AWC during prolonged odor exposure. Using this fully functional GFP-tagged EGL-4 (GFP::EGL-4) molecule we have been able to develop a molecular readout of long-term olfactory adaptation in the AWC5. Using this molecular readout of olfactory adaptation we have been able to perform both forward and reverse genetic screens to identify mutant animals that exhibit defective subcellular localization patterns of GFP::EGL-4 in the AWC6,7. Here we describe: 1) the construction of GFP::EGL-4 expressing animals; 2) the protocol for cultivation of animals for long-term odor-induced nuclear translocation assays; and 3) the scoring of the long-term odor-induced nuclear translocation event and recovery (re-sensitization) from the nuclear GFP::EGL-4 state.

A Molecular Readout of Long-term Olfactory Adaptation in C. elegans

Here we describe a molecular readout of long-term olfactory adaptation in Caenorhabditis elegans. The Protein Kinase G, EGL-4, is necessary for stable adaptation responses in the primary sensory neuron pair called AWC. During prolonged odor exposure EGL-4 translocates from the cytosol to nucleus of the AWC.

During sustained stimulation most sensory neurons will adapt their response by decreasing their sensitivity to the signal. The adaptation response helps ...

Multi-unit Recording Methods to Characterize Neural Activity in the Locust (Schistocerca Americana) Olfactory Circuits

Detection and interpretation of olfactory cues are critical for the survival of many organisms. Remarkably, species across phyla have strikingly similar olfactory systems suggesting that the biological approach to chemical sensing has been optimized over evolutionary time1. In the insect olfactory system, odorants are transduced by olfactory receptor neurons (ORN) in the antenna, which convert chemical stimuli into trains of action potentials. Sensory input from the ORNs is then relayed to the antennal lobe (AL; a structure analogous to the vertebrate olfactory bulb). In the AL, neural representations for odors take the form of spatiotemporal firing patterns distributed across ensembles of principal neurons (PNs; also referred to as projection neurons)2,3. The AL output is subsequently processed by Kenyon cells (KCs) in the downstream mushroom body (MB), a structure associated with olfactory memory and learning4,5. Here, we present electrophysiological recording techniques to monitor odor-evoked neural responses in these olfactory circuits.
First, we present a single sensillum recording method to study odor-evoked responses at the level of populations of ORNs6,7. We discuss the use of saline filled sharpened glass pipettes as electrodes to extracellularly monitor ORN responses. Next, we present a method to extracellularly monitor PN responses using a commercial 16-channel electrode3. A similar approach using a custom-made 8-channel twisted wire tetrode is demonstrated for Kenyon cell recordings8. We provide details of our experimental setup and present representative recording traces for each of these techniques.

Multi-unit Recording Methods to Characterize Neural Activity in the Locust Schistocerca Americana Olfactory Circuits

We demonstrate variations of the extracellular multi-unit recording technique to characterize odor-evoked responses in the first three stages of the invertebrate olfactory pathway. These techniques can easily be adapted to examine ensemble activity in other neural systems as well.

Detection and interpretation of olfactory cues are critical for the survival  of many organisms. Remarkably, species across phyla have strikingly similar ...

In vivo Neuronal Calcium Imaging in C. elegans

The nematode worm C. elegans is an ideal model organism for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, well-characterized nervous system allows identification and fluorescence imaging of any neuron within the intact animal. Simple immobilization techniques with minimal impact on the animal's physiology allow extended time-lapse imaging. The development of genetically-encoded calcium sensitive fluorophores such as cameleon 1 and GCaMP 2 allow in vivo imaging of neuronal calcium relating both cell physiology and neuronal activity. Numerous transgenic strains expressing these fluorophores in specific neurons are readily available or can be constructed using well-established techniques. Here, we describe detailed procedures for measuring calcium dynamics within a single neuron in vivo using both GCaMP and cameleon. We discuss advantages and disadvantages of both as well as various methods of sample preparation (animal immobilization) and image analysis. Finally, we present results from two experiments: 1) Using GCaMP to measure the sensory response of a specific neuron to an external electrical field and 2) Using cameleon to measure the physiological calcium response of a neuron to traumatic laser damage. Calcium imaging techniques such as these are used extensively in C. elegans and have been extended to measurements in freely moving animals, multiple neurons simultaneously and comparison across genetic backgrounds. C. elegans presents a robust and flexible system for in vivo neuronal imaging with advantages over other model systems in technical simplicity and cost.

In vivo Neuronal Calcium Imaging in C. elegans

With its small transparent body, well-documented neuroanatomy and a host of amenable genetic techniques and reagents, C. elegans makes an ideal model organism for in vivo neuronal imaging using relatively simple, low-cost techniques. Here we describe single neuron imaging within intact adult animals using genetically encoded fluorescent calcium indicators.

The nematode worm C. elegans is an ideal model organism  for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, ...

Transplantation of Olfactory Ensheathing Cells to Evaluate Functional Recovery after Peripheral Nerve Injury

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth and regrowth of the primary olfactory neurons. Indeed, the primary olfactory system is characterized by its ability to give rise to new neurons even in adult animals. This particular ability is partly due to the presence of OECs which create a favorable microenvironment for neurogenesis. This property of OECs has been used for cellular transplantation such as in spinal cord injury models. Although the peripheral nervous system has a greater capacity to regenerate after nerve injury than the central nervous system, complete sections induce misrouting during axonal regrowth in particular after facial of laryngeal nerve transection. Specifically, full sectioning of the recurrent laryngeal nerve (RLN) induces aberrant axonal regrowth resulting in synkinesis of the vocal cords. In this specific model, we showed that OECs transplantation efficiently increases axonal regrowth.
OECs are constituted of several subpopulations present in both the olfactory mucosa (OM-OECs) and the olfactory bulbs (OB-OECs). We present here a model of cellular transplantation based on the use of these different subpopulations of OECs in a RLN injury model. Using this paradigm, primary cultures of OB-OECs and OM-OECs were transplanted in Matrigel after section and anastomosis of the RLN. Two months after surgery, we evaluated transplanted animals by complementary analyses based on videolaryngoscopy, electromyography (EMG), and histological studies. First, videolaryngoscopy allowed us to evaluate laryngeal functions, in particular muscular cocontractions phenomena. Then, EMG analyses demonstrated richness and synchronization of muscular activities. Finally, histological studies based on toluidine blue staining allowed the quantification of the number and profile of myelinated fibers.
All together, we describe here how to isolate, culture, identify and transplant OECs from OM and OB after RLN section-anastomosis and how to evaluate and analyze the efficiency of these transplanted cells on axonal regrowth and laryngeal functions.

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth of the primary olfactory neurons. This specific property can be used for cellular transplantation. We present here a model of cellular transplantation based on the use of OECs in a laryngeal nerve injury model.

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth and regrowth of the primary olfactory neurons. Indeed, the primary olfactory ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

The mechanisms controlling stress-induced phenotypic plasticity in animals are frequently complex and difficult to study in vivo. A classic example of stress-induced plasticity is the dauer stage of C. elegans. Dauers are an alternative developmental larval stage formed under conditions of low concentrations of bacterial food and high concentrations of a dauer pheromone. Dauers display extensive developmental and behavioral plasticity. For example, a set of four inner-labial quadrant (IL2Q) neurons undergo extensive reversible remodeling during dauer formation. Utilizing the well-known environmental pathways regulating dauer entry, a previously established method for the production of crude dauer pheromone from large-scale liquid nematode cultures is demonstrated. With this method, a concentration of 50,000 - 75,000 nematodes/ml of liquid culture is sufficient to produce a highly potent crude dauer pheromone. The crude pheromone potency is determined by a dose-response bioassay. Finally, the methods used for in vivo time-lapse imaging of the IL2Qs during dauer formation are described.

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

Following exposure to specific environmental stressors, the nematode Caenorhabditis elegans undergoes extensive phenotypic plasticity to enter into a stress-resistant &#8216;dauer&#8217; juvenile stage. We present methods for the controlled induction and imaging of neuroplasticity during dauer.

The mechanisms controlling stress-induced phenotypic plasticity in animals are frequently complex and difficult to study in vivo. A classic example of ...

Olfactory Neurons Obtained through Nasal Biopsy Combined with Laser-Capture Microdissection: A Potential Approach to Study Treatment Response in Mental Disorders

Bipolar disorder (BD) is a severe neuropsychiatric disorder with poorly understood pathophysiology and typically treated with the mood stabilizer, lithium carbonate. Animal studies as well as human genetic studies indicate that lithium affects molecular targets that are involved in neuronal growth, survival and maturation, and notably molecules involved in Wnt signaling. Given the ethical challenge to obtaining brain biopsies for investigating dynamic molecular changes associated with lithium-response in the central nervous system (CNS), one may consider the use of neurons obtained from olfactory tissues to achieve this goal.The olfactory epithelium contains olfactory receptor neurons at different stages of development and glial-like supporting cells. This provides a unique opportunity to study dynamic changes in the CNS of patients with neuropsychiatric diseases, using olfactory tissue safely obtained from nasal biopsies. To overcome the drawback posed by substantial contamination of biopsied olfactory tissue with non-neuronal cells, a novel approach to obtain enriched neuronal cell populations was developed by combining nasal biopsies with laser-capture microdissection. In this study, a system for investigating treatment-associated dynamic molecular changes in neuronal tissue was developed and validated, using a small pilot sample of BD patients recruited for the study of the molecular mechanisms of lithium treatment response.

In this study, a novel platform to investigate intraneuronal molecular signatures of treatment response in bipolar disorder (BD) was developed and validated. Olfactory epithelium from BD patients was obtained through nasal biopsies. Then laser-capture microdissection was combined with Real Time RT PCR to investigate the molecular signature of lithium response in BD.

Bipolar disorder (BD) is a severe neuropsychiatric disorder with poorly understood pathophysiology and typically treated with the mood stabilizer, lithium ...

Utilizing Combined Methodologies to Define the Role of Plasma Membrane Delivery During Axon Branching and Neuronal Morphogenesis

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular guidance cues like netrin-1 and results in dramatic increases to the surface area of the axonal plasma membrane. Netrin-1-dependent axon branching likely involves temporal and spatial control of plasma membrane expansion, the components of which are supplied through exocytic vesicle fusion. These fusion events are preceded by formation of SNARE complexes, comprising a v-SNARE, such as VAMP2 (vesicle-associated membrane protein 2), and plasma membrane t-SNAREs, syntaxin-1 and SNAP25 (synaptosomal-associated protein 25). Detailed herein isa multi-pronged approach used to examine the role of SNARE mediated exocytosis in axon branching. The strength of the combined approach is data acquisition at a range of spatial and temporal resolutions, spanning from the dynamics of single vesicle fusion events in individual neurons to SNARE complex formation and axon branching in populations of cultured neurons. This protocol takes advantage of established biochemical approaches to assay levels of endogenous SNARE complexes and Total Internal Reflection Fluorescence (TIRF) microscopy of cortical neurons expressing VAMP2 tagged with a pH-sensitive GFP (VAMP2-pHlourin) to identify netrin-1 dependent changes in exocytic activity in individual neurons. To elucidate the timing of netrin-1-dependent branching, time-lapse differential interference contrast (DIC) microscopy of single neurons over the order of hours is utilized. Fixed cell immunofluorescence paired with botulinum neurotoxins that cleave SNARE machinery and block exocytosis demonstrates that netrin-1 dependent axon branching requires SNARE-mediated exocytic activity.

Light microscopy techniques coupled with biochemical assays elucidate the involvement of SNARE-mediated exocytosis in netrin-dependent axon branching. This combination of techniques permits identification of molecular mechanisms controlling axon branching and cell shape change.

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular ...

Recording Temperature-induced Neuronal Activity through Monitoring Calcium Changes in the Olfactory Bulb of Xenopus laevis

The olfactory system, specialized in the detection, integration and processing of chemical molecules is likely the most thoroughly studied sensory system. However, there is piling evidence that olfaction is not solely limited to chemical sensitivity, but also includes temperature sensitivity. Premetamorphic Xenopus laevis are translucent animals, with protruding nasal cavities deprived of the cribriform plate separating the nose and the olfactory bulb. These characteristics make them well suited for studying olfaction, and particularly thermosensitivity. The present article describes the complete procedure for measuring temperature responses in the olfactory bulb of X. laevis larvae. Firstly, the electroporation of olfactory receptor neurons (ORNs) is performed with spectrally distinct dyes loaded into the nasal cavities in order to stain their axon terminals in the bulbar neuropil. The differential staining between left and right receptor neurons serves to identify the &#947;-glomerulus as the only structure innervated by contralateral presynaptic afferents. Secondly, the electroporation is combined with focal bolus loading in the olfactory bulb in order to stain mitral cells and their dendrites. The 3D brain volume is then scanned under line-illumination microscopy for the acquisition of fast calcium imaging data while small temperature drops are induced at the olfactory epithelium. Lastly, the post-acquisition analysis allows the morphological reconstruction of the thermosensitive network comprising the &#947;-glomerulus and its innervating mitral cells, based on specific temperature-induced Ca2+ traces. Using chemical odorants as stimuli in addition to temperature jumps enables the comparison between thermosensitive and chemosensitive networks in the olfactory bulb.

Recording Temperature-induced Neuronal Activity through Monitoring Calcium Changes in the Olfactory Bulb of Xenopus laevis

Here we describe a protocol for measuring and analyzing temperature responses in the olfactory bulb of Xenopus laevis. Olfactory receptor neurons and mitral cells are differentially stained, after which calcium changes are recorded, reflecting a sensitivity of some neural networks in the bulb to temperature drops induced at the nose.

The olfactory system, specialized in the detection, integration and processing of chemical molecules is likely the most thoroughly studied sensory system. ...