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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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-E)29,30. However, in neuronal classes with four radially symmetric neurons, such as CEM, all four neurons are visible only one-third of the time. Another third of worms tested have only three of the four neurons visible, and for the remaining third, the only distinguishable difference is between the dorsal and ventral cell bodies, not the left-right asymmetry (data not shown). The narrower port is combined with a lower channel height to prevent worm fluctuation across the z-axis. This design allows for the imaging of males in future studies, which, when combined with the constantly increasing knowledge of the male connectome25,31, will allow for a better understanding of sex-specific neural function.
The analysis performed in this protocol uses the free software ImageJ to measure changes in fluorescence in the neuron of interest. With the current design, 1 mM tetramisole in the buffer effectively paralyzes the worms and prevents movement of the neurons being imaged. If movement is not preventable, or if the user wishes to avoid the use of a paralytic, more complex tracking scripts must be written that track the neurons as they move7. However, in this protocol, male worms only move when they were too small to be effectively constrained by the loading port and when presented with an extremely aversive stimulus, such as glycerol. Even in these instances, the movement is brief and does not require large amounts of tracking adjustments&#8212;setting an ROI around the new neuron location alleviates the incorrect fluorescent readouts (Figure 2).
A limitation of single-worm, trap-based imaging is that only one worm can be imaged at a time1. Another limitation of these traps is that worms can get stuck within the device, causing devices to be clogged and &#34;used up&#34; after imaging only a few worms. However, the quick turnaround time for the fabrication of new devices from a master mold alleviates this downside. Extended blue-light illumination has also been shown to induce photodamage in C. elegans32,33. The relatively short experimental time frame of this protocol (30 s) allows for imaging without measurable photobleaching. However, to avoid photobleaching and photodamage in longer experiments, the light source can be pulsed7. For example, during each 100-ms exposure, the light can be pulsed for 10 ms. This has been shown to eliminate increased body autofluorescence over time7.
In order to properly test males for their responses to ascarosides, larval-stage 4 (L4) males must first be isolated from hermaphrodites, for at least 5 h, in order to achieve a near-na&#239;ve response to the pheromones23. Isolation for less than this length of time may cause animals to fail to respond to the ascarosides. However, this isolation is not necessary when testing non-ascaroside cues, such as glycerol. For the sake of consistency, however, animals were always isolated at least 5 h prior to calcium imaging. In some neurons, such as the CEM, not every stimulation will elicit a neuronal response, and each CEM that does responds does so to generate a certain &#34;code&#34; of neural representation. This phenomenon in CEM has been observed via electrophysiology studies, as well as with calcium imaging studies like the ones described here23. Thus, measurable calcium transients in CEM neurons in every ascaroside-exposed animal are not guaranteed23. In fact, many of the ascarosides investigated to date do not elicit measurable calcium transients13,15,34,35. The successful elicitation of measurable transients was observed during only one of three pulses of pheromone in two of the five animals exposed to the ascaroside of interest (Figure 3). This matches the rate of success previously observed in other labs23. This variability is a limitation when studying pheromone response and is not due to the male-based focus of this protocol.
When investigating calcium transients elicited in response to ascarosides, one should not dismiss a lack of consistent response without further investigation. This can be tested through experiments such as electrophysiology studies to confirm the variable response within a neuronal class. For neurons, such as ASH, that respond reliably, a lack of consistent response could be indicative of larger experimental problems, such as errors in stimulus control. The peak intensity of the responses can also be investigated if variability is expected in the response. The traces can be plotted with the standard deviation or standard error (as in Figure 2F). If the standard deviation is small, the traces can be plotted and analyzed as such. If there is noticeable amount of variation leading to moderate standard deviations, the data can be plotted the same, with accompanying heatmaps sorted by response &#34;type&#34; to show the response-by-response variation. If there is significant variation (Figure 3), wherein the peaks cannot be distributed in a Gaussian manner, responses can be categorized into response &#34;types&#34; (e.g., depolarizing, hyperpolarizing, or non-polarizing)23. Responses that fall into a certain &#34;type&#34; can be plotted and analyzed together. Similarly, heatmaps should accompany this analysis as well.
Moving forward, this device can be adapted to allow for the imaging of larval-staged nematodes by narrowing the loading port even further. Further narrowing of the end of the loading port will allow for the constraint of the animal to allow for imaging of just the cilia of the sensory neurons, as opposed to the cell body. While other devices are designed for the more commonly studied hermaphrodite, this adapted olfactory chip allows for the imaging of neural activity in male neural circuits. As the connectome of the male is still being elucidated, being able to measure neural dynamics in sex-specific networks is critical to fully understanding neuronal signaling. Differences between hermaphroditic and male responses can now be tested and measured using this device.

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><tbody><tr><td>Silicon Wafer</td><td>University Wafer</td><td>452</td><td></td></tr><tr><td>SU-8 2035</td><td>MicroChem</td><td>Y111070-0500L1GL</td><td></td></tr><tr><td>Developer</td><td>MicroChem</td><td>Y020100-4000L1PE</td><td></td></tr><tr><td>Wafer Mask</td><td>Cad/Art Services</td><td>-</td><td>Custom order. Printed at 25,000 dpi.</td></tr><tr><td>Sylgard-184</td><td>Ellsworth Adhesives</td><td>184 SIL ELAST KIT 0.5KG</td><td></td></tr><tr><td>1.0 mm Dermal Punches</td><td>Acuderm Inc.</td><td>P150</td><td></td></tr><tr><td>Soft Tubing</td><td>Cole-Palmer</td><td>EW-06419-01</td><td></td></tr><tr><td>Hard Tubing</td><td>IDEX Health &amp;amp; Science</td><td>1622</td><td></td></tr><tr><td>Pins</td><td>New England Small Tube</td><td>NE-1027-12</td><td></td></tr><tr><td>Blocking Pins</td><td>New England Small Tube</td><td>0.415/0.425&quot; OD x .500 Long</td><td>Batch PB07027</td></tr><tr><td>3 mL syringes</td><td>BD</td><td>309657</td><td></td></tr><tr><td>30 mL syringes</td><td>Vitality Medical</td><td>302832</td><td>Used as buffer reservoirs.</td></tr><tr><td>Stainless Steel Blunt Needle 23 Gauge, Polyprolylene Luer</td><td>Component Supply Company</td><td>NE-231PL-50</td><td></td></tr><tr><td>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><td>Fisherfinest Premium Cover Glass</td><td>Fisher Scientific</td><td>12-548-5M</td><td></td></tr><tr><td>Mercator Control System LF-5 Plasma System</td><td>Mercator</td><td>LF-5</td><td></td></tr><tr><td>Scotch Tape</td><td>Scotch</td><td>BSN43575</td><td></td></tr><tr><td>Series 20 Chamber</td><td>Warner Instruments</td><td>P-2</td><td></td></tr><tr><td>Vacuum Desicator</td><td>Bel-Art Scienceware</td><td>420250000</td><td>24 cm inner diameter.</td></tr><tr><td>Weigh Boats</td><td>Cole-Palmer</td><td>EW-01017-27</td><td></td></tr><tr><td>Classic Plus Balance</td><td>Mettler Toledo</td><td>PB1501-S/FACT</td><td></td></tr><tr><td>Glass Pasteur Pipettes</td><td>Cole-Palmer</td><td>EW-25554-06</td><td></td></tr><tr><td>Transfer pipettes</td><td>Genesee Scientific</td><td>30-202</td><td></td></tr><tr><td>Oven</td><td>Sheldon Manufacturing Inc</td><td>9120993</td><td>Model Number: 1500E.</td></tr><tr><td>60 mm, non-vented, sharp edge Petri dishes</td><td>TriTech Research</td><td>T3308</td><td></td></tr><tr><td>Zeiss Axio Observer.A1</td><td>Zeiss</td><td>-</td><td></td></tr><tr><td>Hammamatsu Orca Flash 4.0 Digital CMOS</td><td>Hammamatsu</td><td>C11440-22CU</td><td></td></tr><tr><td>Blue Fluorescent Light</td><td>Lumencor</td><td>SOLA SM6-LCR-SA</td><td>24-30V/7.9A DC.</td></tr><tr><td>Illumination Adaptor</td><td>Zeiss</td><td>423302-0000</td><td></td></tr><tr><td>Series 1 and 2 Miniature Inert PTFE Isolation Valve</td><td>Parker</td><td>001-0017-900</td><td>3-way valve for controlling flow.</td></tr><tr><td>ValveLink8.2&amp;reg;</td><td>AutoMate Scientific</td><td>01-18</td><td>Flow Switch Controller</td></tr><tr><td>Micro Manager</td><td>Micro-Manager</td><td>-</td><td>Free software, can be downloaded at: https://www.micro-manager.org/wiki/Download_Micro-Manager_Latest_Release</td></tr><tr><td>ImageJ</td><td>ImageJ</td><td>-</td><td>Free software, can be downloaded at: https://imagej.nih.gov/ij/download.html</td></tr><tr><td>Agar, Bacteriological Grade</td><td>Apex</td><td>9012-36-6</td><td></td></tr><tr><td>Peptone</td><td>Apex</td><td>20-260</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>VWR</td><td>BDH0224-1KG</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>230391-1kg</td><td></td></tr><tr><td>Cholesterol</td><td>Alfa Aesar</td><td>A11470</td><td></td></tr><tr><td>Ethanol</td><td>Sigma-Aldrich</td><td>270741-4L</td><td></td></tr><tr><td>Tetramisole</td><td>Sigma-Aldrich</td><td>L9756-10(G)</td><td>Store at 4 &amp;deg;C.</td></tr><tr><td>Fluorescein</td><td>Sigma-Aldrich</td><td>FD2000S-250mg</td><td>Light Sensitive. Store in photoprotective vials.</td></tr><tr><td>Glycerol</td><td>Sigma-Aldrich</td><td>G6279-1L</td><td></td></tr><tr><td>Ascaroside #3</td><td>-</td><td>-</td><td>Synthesized in the Schroeder Lab (Cornell University).</td></tr><tr><td>NaCl</td><td>Genesee Scientific</td><td>18-215</td><td></td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>BDH</td><td>BDH9268.25</td><td></td></tr><tr><td>K&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>J.T. Baker</td><td>3252-025</td><td></td></tr><tr><td>ASH GCaMP3 line</td><td>-</td><td>-</td><td>CX10979 (KyEx2865 [p&lt;em&gt;sra&lt;/em&gt;-6::GCAMP3 @ 100 ng/uL]). Developed in Bargmann lab. Provided from Albrecht Lab library.</td></tr><tr><td>CEM GCaMP6 line</td><td>-</td><td>-</td><td>JSR49 (FkEx98[p&lt;em&gt;pkd&lt;/em&gt;-2::GCaMP::SL2::dsRED + pBX-1]; &lt;em&gt;pha&lt;/em&gt;-1(e2123ts); &lt;em&gt;him&lt;/em&gt;-5(e1490); &lt;em&gt;lite&lt;/em&gt;-1(ce314)). Developed by Robyn Lints. Provided from Srinivasan Lab library.</td></tr><tr><td>&lt;em&gt;E. coli&lt;/em&gt; (OP50)</td><td>&lt;em&gt;Caenorhabditis&lt;/em&gt; Genetics Center</td><td>OP50</td><td></td></tr><tr><td>&quot;Reservoir&quot;</td><td>-</td><td>-</td><td>To create a Reservoir: A &quot;30 mL syringe&quot;, is connected to a &quot;Stopcock with Luer connections; 3-way; male lock; 5 flow pattern; non-sterile&quot;, which is connected to a &quot;3 mL syringe&quot; and a &quot;Stainless Steel Blunt Needle 23 Gauge, Polyprolylene Luer&quot;. The &quot;Stainless Steel Blunt Needle 23 Gauge, Polyprolylene Luer&quot; is then inserted into &quot;Soft Tubing&quot; 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 design of the male-adapted microfluidic device contains a curve in the loading port, but flow dynamics are identical to the device designed by Chronis et al.19(Figure 2A-2C). The flow of buffers can be controlled by altering which flow control valve is open (Figure 2A-2B). The measurements of the device as fabricated vary from the designed file. The measurements provided in Figure 2C are &#34;as fabricated&#34; measurements.
After loading male C. elegans into the male-adapted olfactory device, their placement and orientation, as well as channel flow dynamics, can be verified via both bright-field and fluorescent imaging (Figure 2D-2E). The exposure of worms expressing GCaMP3 in the nociceptive neuron, ASH, to 1 M glycerol results in visible changes in fluorescence within the ASH neuron, indicative of neural activity (Figure 2F). Subtle changes in fluorescence may not be visible by eye, but software can be used to quantify these changes. The free ImageJ software can be used to analyze and quantify the fluorescent intensity of ASH neurons upon exposure to 1 M glycerol over time (Figure 2F). This is similar to what is observed in hermaphrodites27 and, due to the robustness of the ASH neuronal response to glycerol, this is observed in all animals tested.
A small amount of axial or rotational movement is expected in unparalyzed animals, often necessitating a neuron-tracking algorithm during video analysis (Figure 3A). The addition of a paralytic in the buffers (e.g., 1 mM tetramisole) nearly eliminates this effect, although some animals (~10%) still move during the trials. This can be circumvented by: (a) using older males, which are more efficiently trapped; (b) decreasing the width or thickness of the worm loading port even further; or (c) either increasing the concentration of the paralytic used or using another paralytic. This will also ensure that there is not too much of the head past the end of the trap and exposed to the odor channel. If a trial with worm movement occurs, the area of analysis can be moved and reread from the new neuronal location, starting at the frame after which the movement occurs (Figure 3B). Manual reconstruction of the neural traces by the user is required in this instance. Scripts that analyze the fluorescent changes within the neuron and that follow the neuron&#39;s center as it moves can also be written19.
Male C. elegans sense attractive biogenic pheromones called ascarosides via the four sex-specific CEM neurons23. When calcium transients are observed in males, the responses are variable in shape, sign, and magnitude between both neurons and animals (Figure 4A-B). However, male response to pheromones is not as reliably observed as calcium transients in many animals (Figure 4C). This is not discouraging, as most ascarosides do not elicit calcium transients upon sensation13,14,15.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56026/56026fig3.jpg" /> 
Figure 3: Addressing worm movement during the image acquisition period. (A) &#916;F/F0 (change in fluorescent intensity divided by the average background fluorescent intensity for first 1 s of acquisition) was calculated using ImageJ by defining an area where the neuron of interest was reliably static for 1 s. During the stimulus pulse (blue area), the animal moved, as seen in the images above the calcium trace, resulting in the neuron no longer being contained within the analyzed area (yellow box). The scale bars denote 42 &#181;m. the dashed lines show the region of the trace that corresponds to the location of the worm in each image. (B) If the area of analysis is moved to follow the neuron during the trial and the individual traces are rebuilt to convey the actual neural dynamics, the trace appears as if the animal did not move. The dashed lines show the region of the trace that was corrected for worm movement. Traces for both ASH neurons, left and right (ASHL (red) and ASHR (blue), respectively) are shown. The Y-axes show &#916;F/F0. The scale bar denotes 5 s. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56026/56026fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56026/56026fig4.jpg" /> 
Figure 4: Male C. elegans CEM response to 1 &#956;M ascr#3 is variable. The male-specific CEM neurons display unique patterns of response to biogenic pheromones. (A) The responses observed in each CEM neuron for one pulse in one responsive animal are shown. There are four CEM neurons: dorsal right (CEM DR), dorsal left (CEM DL), ventral right (CEM VR), and ventral left (CEM VL). Three of the four neurons respond with depolarizations of varying shape and magnitude, with the fourth not responding to the ascaroside. (B) Approximately one-third of the trapped animals (2/7 tested in this study) result in only three neurons able to be imaged. The responses of a worm in this orientation resulted in CEM calcium transients that were different than those observed in the first worm. This was not due to the change in orientation of the worm, as CEM DR is visible in both orientations and exhibits variable response between animals. (C) Many worms do not respond with any detectable calcium transients (5/7 tested in this study). Individual traces shown are representative of the single animals shown in the images above the plots. The scale bars denote 42 &#181;m. Traces: The blue area denotes the time of 1 &#956;M ascr#3 exposure. The red traces denote the depolarizing response. The black traces denote no observed response. The Y-axes show &#916;F/F0. The scale bar denotes 5 s. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56026/56026fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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 ...

using-an-adapted-microfluidic-olfactory-chip-for-imaging-neuronal

Nanyang Technological University

Research

JoVE Journal

Neuroscience

8.1K Views.  Worcester Polytechnic Institute. Het gebruik van een aangepast "olfactorische chip" voor de efficiënte calcium beeldvorming van C. elegans mannetjes wordt hier beschreven. Studies van mannelijke Gasbedwelming met behulp van glycerol en een feromoon worden ook weergegeven.

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

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are available for microfluidic devices using soft lithography techniques 1-3. Microfluidic devices have been used for sub-cellular imaging 4,5, in vivo laser microsurgery 2,6 and cellular imaging 4,7. In vivo imaging requires immobilization of organisms. This has been achieved using suction 5,8, tapered channels 6,7,9, deformable membranes 2-4,10, suction with additional cooling 5, anesthetic gas 11, temperature sensitive gels 12, cyanoacrylate glue 13 and anesthetics such as levamisole 14,15. Commonly used anesthetics influence synaptic transmission 16,17 and are known to have detrimental effects on sub-cellular neuronal transport 4. In this study we demonstrate a membrane based poly-dimethyl-siloxane (PDMS) device that allows anesthetic free immobilization of intact genetic model organisms such as Caenorhabditis elegans (C. elegans), Drosophila larvae and zebrafish larvae. These model organisms are suitable for in vivo studies in microfluidic devices because of their small diameters and optically transparent or translucent bodies. Body diameters range from ~10 &mu;m to ~800 &mu;m for early larval stages of C. elegans and zebrafish larvae and require microfluidic devices of different sizes to achieve complete immobilization for high resolution time-lapse imaging. These organisms are immobilized using pressure applied by compressed nitrogen gas through a liquid column and imaged using an inverted microscope. Animals released from the trap return to normal locomotion within 10 min.
We demonstrate four applications of time-lapse imaging in C. elegans namely, imaging mitochondrial transport in neurons, pre-synaptic vesicle transport in a transport-defective mutant, glutamate receptor transport and Q neuroblast cell division. Data obtained from such movies show that microfluidic immobilization is a useful and accurate means of acquiring in vivo data of cellular and sub-cellular events when compared to anesthetized animals (Figure 1J and 3C-F 4).
Device dimensions were altered to allow time-lapse imaging of different stages of C. elegans, first instar Drosophila larvae and zebrafish larvae. Transport of vesicles marked with synaptotagmin tagged with GFP (syt.eGFP) in sensory neurons shows directed motion of synaptic vesicle markers expressed in cholinergic sensory neurons in intact first instar Drosophila larvae. A similar device has been used to carry out time-lapse imaging of heartbeat in ~30 hr post fertilization (hpf) zebrafish larvae. These data show that the simple devices we have developed can be applied to a variety of model systems to study several cell biological and developmental phenomena in vivo.

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

A simple microfluidic device has been developed to perform anesthetic free in vivo imaging of C. elegans, intact Drosophila larvae and zebrafish larvae. The device utilizes a deformable PDMS membrane to immobilize these model organisms in order to perform time lapse imaging of numerous processes such as heart beat, cell division and sub-cellular neuronal transport. We demonstrate the use of this device and show examples of different types of data collected from different model systems.

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are ...

Bioengineering

Calcium Imaging of Odor-evoked Responses in the Drosophila Antennal Lobe

The antennal lobe is the primary olfactory center in the insect brain and represents the anatomical and functional equivalent of the vertebrate olfactory bulb1-5. Olfactory information in the external world is transmitted to the antennal lobe by olfactory sensory neurons (OSNs), which segregate to distinct regions of neuropil called glomeruli according to the specific olfactory receptor they express. Here, OSN axons synapse with both local interneurons (LNs), whose processes can innervate many different glomeruli, and projection neurons (PNs), which convey olfactory information to higher olfactory brain regions.
Optical imaging of the activity of OSNs, LNs and PNs in the antennal lobe - traditionally using synthetic calcium indicators (e.g. calcium green, FURA-2) or voltage-sensitive dyes (e.g. RH414) - has long been an important technique to understand how olfactory stimuli are represented as spatial and temporal patterns of glomerular activity in many species of insects6-10. Development of genetically-encoded neural activity reporters, such as the fluorescent calcium indicators G-CaMP11,12 and Cameleon13, the bioluminescent calcium indicator GFP-aequorin14,15, or a reporter of synaptic transmission, synapto-pHluorin16 has made the olfactory system of the fruitfly, Drosophila melanogaster, particularly accessible to neurophysiological imaging, complementing its comprehensively-described molecular, electrophysiological and neuroanatomical properties2,4,17. These reporters can be selectively expressed via binary transcriptional control systems (e.g. GAL4/UAS18, LexA/LexAop19,20, Q system21) in defined populations of neurons within the olfactory circuitry to dissect with high spatial and temporal resolution how odor-evoked neural activity is represented, modulated and transformed22-24.
Here we describe the preparation and analysis methods to measure odor-evoked responses in the Drosophila antennal lobe using G-CaMP25-27. The animal preparation is minimally invasive and can be adapted to imaging using wide-field fluorescence, confocal and two-photon microscopes.

Calcium Imaging of Odor-evoked Responses in the Drosophila Antennal Lobe

We describe an established technique to measure and analyze odor-evoked calcium responses in the antennal lobe of living Drosophila melanogaster.

The antennal lobe is the primary olfactory center in the insect brain and represents the anatomical and functional equivalent of the vertebrate olfactory ...

Imaging Neuronal Responses in Slice Preparations of Vomeronasal Organ Expressing a Genetically Encoded Calcium Sensor

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within the same species 1,2. These intraspecies signals, the pheromones, as well as signals from some predators 3, activate the vomeronasal sensory neurons (VSNs) with high levels of specificity and sensitivity 4. At least three distinct families of G-protein coupled receptors, V1R, V2R and FPR 5-14, are expressed in VNO neurons to mediate the detection of the chemosensory cues. To understand how pheromone information is encoded by the VNO, it is critical to analyze the response profiles of individual VSNs to various stimuli and identify the specific receptors that mediate these responses.
The neuroepithelia of VNO are enclosed in a pair of vomer bones. The semi-blind tubular structure of VNO has one open end (the vomeronasal duct) connecting to the nasal cavity. VSNs extend their dendrites to the lumen part of the VNO, where the pheromone cues are in contact with the receptors expressed at the dendritic knobs. The cell bodies of the VSNs form pseudo-stratified layers with V1R and V2R expressed in the apical and basal layers respectively 6-8. Several techniques have been utilized to monitor responses of VSNs to sensory stimuli 4,12,15-19. Among these techniques, acute slice preparation offers several advantages. First, compared to dissociated VSNs 3,17, slice preparations maintain the neurons in their native morphology and the dendrites of the cells stay relatively intact. Second, the cell bodies of the VSNs are easily accessible in coronal slice of the VNO to allow electrophysiology studies and imaging experiments as compared to whole epithelium and whole-mount preparations 12,20. Third, this method can be combined with molecular cloning techniques to allow receptor identification.
Sensory stimulation elicits strong Ca2+ influx in VSNs that is indicative of receptor activation 4,21. We thus develop transgenic mice that express G-CaMP2 in the olfactory sensory neurons, including the VSNs 15,22. The sensitivity and the genetic nature of the probe greatly facilitate Ca2+ imaging experiments. This method has eliminated the dye loading process used in previous studies 4,21. We also employ a ligand delivery system that enables application of various stimuli to the VNO slices. The combination of the two techniques allows us to monitor multiple neurons simultaneously in response to large numbers of stimuli. Finally, we have established a semi-automated analysis pipeline to assist image processing.

The vomeronasal organ (VNO) detects intraspecies chemical signals that convey social and reproductive information. We have performed Ca2+ imaging experiments using transgenic mice expressing G-CaMP2 in VNO tissue. This approach allows us to analyze the complicated response patterns of the vomeronasal neurons to large numbers of pheromone stimuli.

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within ...

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, ...

Extraction and Quantification of Volatile Sex Pheromone Responses in C. elegans

Volatile Sex Pheromone Extraction and Chemoattraction Assay in Caenorhabditis elegans

Chemical communication is vital in organismal health, reproduction, and overall well-being. Understanding the molecular pathways, neural processes, and computations governing these signals remains an active area of research. The nematode Caenorhabditis elegans provides a powerful model for studying these processes as it produces a volatile sex pheromone. This pheromone is synthesized by virgin females or sperm-depleted hermaphrodites and serves as an attractant for males.
This protocol describes a detailed method for isolating the volatile sex pheromone from several C. elegans strains (WT strain N2, daf-22, and fog-2) and C. remanei. We also provide a protocol for quantifying the male chemotaxis response to the volatile sex pheromone. Our analysis utilizes measurements such as chemotaxis index (C.I.), arrival time (A.T.), and a trajectory plot to compare male responses under various conditions accurately. This method allows for robust comparisons between males of different genetic backgrounds or developmental stages. Furthermore, the experimental setup outlined here is adaptable to investigating other chemoattraction chemicals.

Author Spotlight: Examining Volatile Sex Pheromone Influence on Male C. elegans Behavior

This protocol establishes methods for extracting and quantifying responses to the volatile sex pheromone in C. elegans, providing tools to study chemical communication and navigation trajectory.

Chemical communication is vital in organismal health, reproduction, and overall well-being. Understanding the molecular pathways, neural processes, and ...

Behavior

Using a Microfluidics Device for Mechanical Stimulation and High Resolution Imaging of C. elegans

One central goal of mechanobiology is to understand the reciprocal effect of mechanical stress on proteins and cells. Despite its importance, the influence of mechanical stress on cellular function is still poorly understood. In part, this knowledge gap exists because few tools enable simultaneous deformation of tissue and cells, imaging of cellular activity in live animals, and efficient restriction of motility in otherwise highly mobile model organisms, such as the nematode Caenorhabditis elegans. The small size of C. elegans makes them an excellent match to microfluidics-based research devices, and solutions for immobilization have been presented using microfluidic devices. Although these devices allow for high-resolution imaging, the animal is fully encased in polydimethylsiloxane (PDMS) and glass, limiting physical access for delivery of mechanical force or electrophysiological recordings. Recently, we created a device that integrates pneumatic actuators with a trapping design that is compatible with high-resolution fluorescence microscopy. The actuation channel is separated from the worm-trapping channel by a thin PDMS diaphragm. This diaphragm is deflected into the side of a worm by applying pressure from an external source. The device can target individual mechanosensitive neurons. The activation of these neurons is imaged at high-resolution with genetically-encoded calcium indicators. This article presents the general method using C. elegans strains expressing calcium-sensitive activity indicator (GCaMP6s) in their touch receptor neurons (TRNs). The method, however, is not limited to TRNs nor to calcium sensors as a probe, but can be expanded to other mechanically-sensitive cells or sensors.

Using a Microfluidics Device for Mechanical Stimulation and High Resolution Imaging of C. elegans

New tools for mechanobiology research are needed to understand how mechanical stress activates biochemical pathways and elicits biological responses. Here, we showcase a new method for selective mechanical stimulation of immobilized animals with a microfluidic trap allowing high-resolution imaging of cellular responses.

One central goal of mechanobiology is to understand the reciprocal effect of mechanical stress on proteins and cells. Despite its importance, the ...

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

It has become increasingly clear that neural circuit activity in behaving animals differs substantially from that seen in anesthetized or immobilized animals. Highly sensitive, genetically encoded fluorescent reporters of Ca2+ have revolutionized the recording of cell and synaptic activity using non-invasive optical approaches in behaving animals. When combined with genetic and optogenetic techniques, the molecular mechanisms that modulate cell and circuit activity during different behavior states can be identified.
Here we describe methods for ratiometric Ca2+ imaging of single neurons in freely behaving Caenorhabditis elegans worms. We demonstrate a simple mounting technique that gently overlays worms growing on a standard Nematode Growth Media (NGM) agar block with a glass coverslip, permitting animals to be recorded at high-resolution during unrestricted movement and behavior. With this technique, we use the sensitive Ca2+ reporter GCaMP5 to record changes in intracellular Ca2+ in the serotonergic Hermaphrodite Specific Neurons (HSNs) as they drive egg-laying behavior. By co-expressing mCherry, a Ca2+-insensitive fluorescent protein, we can track the position of the HSN within ~ 1 &#181;m and correct for fluctuations in fluorescence caused by changes in focus or movement. Simultaneous, infrared brightfield imaging allows for behavior recording and animal tracking using a motorized stage. By integrating these microscopic techniques and data streams, we can record Ca2+ activity in the C. elegans egg-laying circuit as it progresses between inactive and active behavior states over tens of minutes.

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

This protocol describes the use of genetically encoded Ca2+ reporters to record changes in neural activity in behaving Caenorhabditis elegans worms. &#8195;

It has become increasingly clear that neural circuit activity in behaving animals differs substantially from that seen in anesthetized or immobilized ...

A Simple Microfluidic Chip for Long-Term Growth and Imaging of Caenorhabditis elegans

Caenorhabditis elegans (C. elegans) have proved to be a valuable model system for studying developmental and cell biological processes. Understanding these biological processes often requires long-term and repeated imaging of the same animal. Long recovery times associated with conventional immobilization methods done on agar pads have detrimental effects on animal health making it inappropriate to repeatedly image the same animal over long periods of time. This paper describes a microfluidic chip design, fabrication method, on-chip C. elegans culturing protocol, and three examples of long-term imaging to study developmental processes in individual animals. The chip, fabricated with polydimethylsiloxane and bonded on a cover glass, immobilizes animals on a glass substrate using an elastomeric membrane that is deflected using nitrogen gas. Complete immobilization of C. elegans enables robust time-lapse imaging of cellular and sub-cellular events in an anesthetic-free manner. A channel geometry with a large cross-section allows the animal to move freely within two partially sealed isolation membranes permitting growth in the channel with a continuous food supply. Using this simple chip, imaging of developmental phenomena such as neuronal process growth, vulval development, and dendritic arborization in the PVD sensory neurons, as the animal grows inside the channel, can be performed. The long-term growth and imaging chip operates with a single pressure line, no external valves, inexpensive fluidic consumables, and utilizes standard worm handling protocols that can easily be adapted by other laboratories using C. elegans.

A Simple Microfluidic Chip for Long-Term Growth and Imaging of Caenorhabditis elegans

The protocol describes a simple microfluidic chip design and microfabrication methodology used to grow C. elegans in presence of a continuous food supply for up to 36 h. The growth and imaging device also enables intermittent long-term high-resolution imaging of cellular and sub-cellular processes during development for several days.

Caenorhabditis elegans (C. elegans) have proved to be a valuable model system for studying developmental and cell biological processes. Understanding ...

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

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