We thank Fox Avery for testing these protocols and reviewing the manuscript and Eric Hall for programming assistance. Funding for the methods presented herein was provided in part by the National Science Foundation 1724026 (D.R.A.).

1. Neural imaging equipment
NOTE: See Lawler and Albrecht15 for detailed instructions on building the imaging and stimulation system, which controls the microscope illumination timing, image acquisition, and stimulus delivery (Figure 1). An inexpensive Arduino Nano stimulus controller actuates the fluidic valves through digital signals to a valve controller and controls the optogenetic illumination through analog voltage signals to an LED controller. Other stimuli, such as vibration motors and thermal heaters, can be controlled using digital or analog signals. The stimulus controller synchronizes the stimulation and image recording via camera signals, as specified by the open-source Micro-Manager microscope control software (&#181;Manager)16. See the Table of Materials for details related to all the materials, reagents, equipment, and organisms used in this protocol.
<ol>
	<li>Set up the neural imaging equipment, including an epifluorescence microscope with GFP optics, an sCMOS camera with a digital exposure output signal, and a stimulus controller15.</li>
	<li>Connect the stimulus controller to the desired systems via digital or analog signals. For the examples presented below, the following systems are used:
	<ol>
		<li>Use a valve controller for chemical stimulation. Connect the valve controller to fluidic solenoid or pinch valves (Figure 1)15.</li>
		<li>Use a 615 nm red LED and controller for optogenetic stimulation, mounted above the microscope stage.</li>
	</ol>
	</li>
	<li>Set up the computer with the microscope control software, create a Configuration Preset with equipment settings,&#160;and ensure proper operation of the stimulus control15.</li>
</ol>
2. Microfluidic device fabrication
NOTE: See Lagoy et al.17 for detailed information about obtaining or fabricating the master molds and the production, use, and cleaning of the microfluidic devices. These steps are summarized below.
<ol>
	<li>Obtain or fabricate a master mold using the microfluidic design file provided (albrechtlab.github.io/microfluidics)17. For young adult C. elegans, ensure that the channel height is 55-70 &#181;m.</li>
	<li>Combine the PDMS base and curing agent at a 10:1 ratio by weight and mix thoroughly with transfer pipettes.</li>
	<li>Degas for 30-60 min in a vacuum desiccator until the bubbles disappear.</li>
	<li>Place the master mold into a large (150 mm diameter) Petri dish, and pour the degassed PDMS up to a depth of 4-5 mm (~100 g). Inspect and remove any dust or bubbles with a transfer pipette.</li>
	<li>Bake at 65 &#176;C on a level oven shelf for 3 h to overnight.</li>
	<li>Once cured, use a scalpel to cut the PDMS from the mold and a straight razor blade to separate the devices.</li>
	<li>Punch inlet and outlet holes using a 1 mm dermal punch, and clean them with dH2O, ethanol, and again with dH2O. Dry the device in an airstream (see Figure 2A).</li>
	<li>Clean both sides of the PDMS device with adhesive tape, removing any dust or debris.</li>
	<li>Prepare the glass slides to complete the microfluidic device as described17. Drill inlet holes in the top slide using a diamond bit, and render the bottom slide hydrophobic by exposure to TFOCS vapors or by applying water-repellant glass treatment (Figure 2B).</li>
	<li>Assemble the glass-PDMS device sandwich into a clamp (Figure 2C).</li>
</ol>
3. Animal preparation
<ol>
	<li>Obtain or create animals with genetically encoded calcium indicators expressed in the neurons of interest. 
	NOTE: For example, line NZ1091 (kyIs587 [gpa-6p::GCaMP2.2b; unc-122p::dsRed]; kyIs5662 [odr-7p::Chrimson:SL2:mCherry; elt-2p::mCherry]) expresses GCaMP and the red light-sensitive cation channel Chrimson in the AWA sensory neuron pair6. Both transgenes are integrated into the genome for stable expression in every animal.</li>
	<li>One day before experimentation, place at least 20 L4 larval stage C. elegans per experiment onto a nematode growth medium (NGM) agar plate seeded with an OP50 E. coli lawn. When maintained at 20 &#176;C, this will synchronize the wild-type animals at the young adult stage the next day. 
	NOTE: For array transgenes, pick animals using a fluorescence stereoscope to ensure transgene expression in the chosen animals.</li>
</ol>
4. Solution preparation
<ol>
	<li>Prepare 1x S Basal buffer (100 mM NaCl and 0.05 M KPO4, pH 6.0) from a 10x stock solution.</li>
	<li>Prepare 1 mM tetramisole buffer by diluting 1 M stock in 1x S Basal. Use this paralytic buffer to prepare all the stimuli. An experiment typically uses about 150 mL.</li>
	<li>Add 0.1-1 &#181;g/mL fluorescein to the &#34;control&#34; buffer reservoirs to visualize the flow.</li>
	<li>Create stimulus solutions by serial dilution to the desired final concentration. For example, create a 10&#8722;7 dilution of diacetyl attractant by first creating a 10&#8722;3 stock. 
	NOTE:&#160;A small amount of fluorescein (0.1&#8211;1 &#181;g/mL) may be added to the stimulus or buffer solution to verify stimulus timing, but concentration should be minimal to avoid artifacts in neural fluorescence.</li>
</ol>
5. Microfluidic device preparation
NOTE: See Reilly et al.9 for a video protocol showing the reservoir generation, device setup, and the loading of the animals. See also Lagoy et al.17 for a written protocol including many helpful tips.
<ol>
	<li>Prepare three or more fluid reservoirs. For each, attach a 30 mL or 60 mL syringe reservoir, a 3 mL priming syringe, and a needle stub to a three-way Luer valve (Figure 2D). Connect the needle to microbore tubing fitted with a metal tube at the end. Label the reservoirs, mount them onto a rack attached to a ring stand, and fill them with the corresponding buffer or stimulus fluids (Figure 2E).</li>
	<li>Degas the assembled microfluidic device in a vacuum desiccator for ~1 h.</li>
	<li>Fill and remove the air bubbles from the reservoir tubing using the priming syringe. Fill the outflow tubing with buffer.</li>
	<li>Remove the microfluidic device from the vacuum, and quickly insert the outlet tubing and inject the fluid through the device until a droplet emerges from one inlet.</li>
	<li>At this inlet, use a &#34;drop-to-drop&#34; connection17 to insert the corresponding fluidic inlet tube (Figure 2F). Ensure that liquid drops are present on both the inlet tubing and the device port hole to avoid introducing a bubble.</li>
	<li>Inject more fluid from the outlet, connect the next inlet tube, and repeat until all the inlets are filled. Insert a solid blocking pin at any unused inlets and the worm loading port.</li>
	<li>Initiate flow by opening the inlet and outlet Luer valves. Inspect the device for leaks at the inlets and glass base. Inspect the device for any bubbles within the flow channels or inlets using the microscope image capture software in live mode. 
	&#8203;NOTE: If bubbles are present, wait for them to absorb into the PDMS material.</li>
</ol>
6. Animal loading
NOTE: See Lagoy et al.17.
<ol>
	<li>Transfer young adult animals onto an unseeded NGM agar plate using a wire-tipped &#34;pick&#34;.</li>
	<li>Flood the plate with approximately 5 mL of 1x S Basal buffer such that the animals are swimming.</li>
	<li>Draw the worms into a loading syringe (1 mL or 3 mL syringe with attached tubing that has been prefilled with 1x S Basal).
	<ol>
		<li>Using a stereoscope, move the tubing end below the liquid surface with one hand to each desired animal, and draw it into the tubing using the syringe held in the other hand.</li>
		<li>Draw the worms only into the tubing, not into the syringe. 
		NOTE: The tubing typically holds only about 100 &#181;L. The animals can be expelled into a local area and then drawn again into the tubing with a small volume.</li>
	</ol>
	</li>
	<li>Close the outlet line, remove the worm loading pin, and connect the worm loading syringe to the device using a drop-to-drop connection.</li>
	<li>Gently flow the animals into the arena, establish buffer flow, and allow up to 1 h for immobilization by tetramisole. 
	&#8203;NOTE: During the immobilization period, ensure that only buffer fluid enters the arena. The stimulus and control reservoirs can be turned off during this period but open them and verify the correct flow before running the stimulation trials.</li>
</ol>
7. Automated stimulation and neuronal recording
<ol>
	<li>Create a stimulus definition text file called &#34;User Defined Acquisition Settings.txt&#34; with a text editor (e.g., Notepad) containing the stimulation settings for the automated image acquisition (see examples in Figure 3). The settings are divided into two sections:
	<ol>
		<li>Microscope acquisition settings: Define the experiment type (Single-Stimulus or Multi-Pattern), exposure and excitation timing, trial duration and intervals, and save directory.</li>
		<li>Stimulation settings: Define the stimulus control parameters. A &#34;stimulation command&#34; specifies the actions occurring at certain video frames with the syntax &#60;letter code&#62;&#60;frame number&#62;, where the letter codes are A = valve1, B = valve2, C = valve3, L = LED light; additionally, uppercase = on and lowercase = off. The LED intensity is set with the letter code &#34;i&#34; and a value of 0 (off) to 255 (maximum brightness). 
		NOTE: The LED intensity value from 0 to 255 sets the output analog voltage from 0 V to 5 V. The current controller used here has a linear intensity scaling, and others should be calibrated with a light power meter.
		<ol>
			<li>For a Single-Stimulus experiment with only one repeated stimulation command, use the format in Figure 3A.</li>
			<li>For a Multi-Pattern experiment with multiple stimulation commands, use the format in Figure 3B. Include a &#34;Pattern Sequence&#34; of digits that represents the order of the stimulus patterns. For each pattern, enter a stimulation command on a separate line. 
			NOTE: A pseudorandom sequence or m-sequence can be useful to investigate stimulus history dependence.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Run the microscope control software. Verify that all the fluidic inlets are open, the flow is as desired within the arena (see Figure 2H), and the neurons of interest are in focus within the live window.</li>
	<li>Close the live window and run the script &#34;MultiPattern_RunScript.bsh&#34; within the software. 
	NOTE: It is useful to run a test experiment without animals to verify proper flow and stimulation. Substituting a different fluorescein concentration for each stimulus fluid can help visualize and document new stimulus patterns.</li>
	<li>After experimentation, disassemble the microfluidic device, and rinse all the surfaces, tubing, and reservoirs with water. 
	&#8203;NOTE: All the components can be reused dozens of times if kept clean. Ensure that reservoirs, tubing, and microfluidic devices are maintained wet or fully dried in an air stream to avoid salt crystallization, which can clog and is difficult to remove. The devices can be kept in ethanol to sterilize but should be fully dried before reuse (&#62;1 h at 65 &#176;C)17.</li>
</ol>
8. Data analysis using NeuroTracker
NOTE: NeuroTracker4,18,19 is an ImageJ/FIJI20 software plugin for tracking the fluorescence intensity of multiple neurons and animals, even as they move during trials. This plugin saves data as text files with each neuron&#39;s position and background-corrected fluorescence intensity (F). The fluorescence data are normalized to the baseline fluorescence (F0), for example, the average of several seconds prior to stimulation, as &#916;F/F0 = (F -F0)/F0, which can be averaged across populations.
<ol>
	<li>Install NeuroTracker scripts as instructed (github.com/albrechtLab/Neurotracker).</li>
	<li>Run NeuroTracker by clicking on Plugins | Tracking | NeuroTracker.</li>
	<li>Select the folder containing the .TIF video files to be tracked, and select the desired settings. 
	NOTE: If only a subset of the video files are to be tracked, select the numerical range of the files to analyze at the prompt. Default tracking settings are appropriate for nonmotile animals at 250 pix/mm resolution. Adjust the parameters proportionally for other resolutions. See the User Guide19 for further settings information.</li>
	<li>Set the intensity threshold such that the neurons are visible for selection (Figure 4).</li>
	<li>Open the Brightness/Contrast (B/C) and Threshold control windows using the Image | Adjust menu.</li>
	<li>In the B/C window, adjust the Minimum and Maximum sliders until the neurons are clearly distinguishable (Figure 4A).</li>
	<li>In the Threshold window, check Dark background and Don&#39;t Reset Range.</li>
	<li>Slide the frame slider to observe the neuron movement and intensity changes, noting any animals to exclude from tracking, such as due to overlap with other animals.</li>
	<li>Identify the neurons for tracking.</li>
	<li>For each animal, adjust the threshold level such that the red threshold area above the neuron is visible in every frame before, during, and after stimulation.</li>
	<li>Click on the neuron to record its position and threshold level.</li>
	<li>Repeat the threshold adjustment and selection for each animal and neuron to track. See Figure 4C for a good threshold level example and Figure 4D,E for over-threshold and under-threshold examples. Previously selected neurons are indicated by a small box.</li>
	<li>When all the neurons are selected, press the spacebar to begin tracking. 
	NOTE: For additional NeuroTracker examples, see previous references4,19.</li>
	<li>Monitor the tracking process for each animal, and make any necessary corrections.</li>
	<li>If NeuroTracker pauses, it has lost the neuron. Reclick on the neuron, adjusting the threshold level as needed.</li>
	<li>If the integration box jumps to another nearby animal or non-neuronal structure, press the spacebar to pause, move the slider back to the first erroneous frame, and reclick on the correct neuron location.</li>
</ol>
9. Data exploration and visualization
NOTE: The MATLAB analysis script is used for data processing and visualization and to generate summary PDFs for each analysis.
<ol>
	<li>Run the &#34;NeuroTrackerSummary_pdf.m&#34; file in MATLAB, and select the folder containing the NeuroTracker data text files.</li>
	<li>Wait for a summary PDF to be produced, allowing the verification of the tracking process (Figure 5). Animals are identified by a number (Figure 5A), and the neural responses from each animal and trial can be viewed to assess the population variability (Figure 5B).</li>
	<li>Use the function &#34;databrowse.m&#34; to explore the neural data, for example, grouping by trial number (Figure 5C), by animal number (Figure 5D), by stimulation pattern, or by another category.</li>
</ol>

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S., DeLuca, J., Caplan, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temporal+Inhibition.">Temporal Inhibition.</a> Encyclopedia of Clinical Neuropsychology. , 2480-2481 (2011).</li><li>Lawler, D. E., Albrecht, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Monitoring+neural+activity+during+sleep/wake+events+in+adult+C.+elegans+by+automated+sleep+detection+and+stimulation.">Monitoring neural activity during sleep/wake events in adult C. elegans by automated sleep detection and stimulation.</a> STAR Protocols. 3 (3), 101532 (2022).</li><li>Edelstein, A. 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=Advanced+methods+of+microscope+control+using+&#181;Manager+software.">Advanced methods of microscope control using &#181;Manager software.</a> Journal of Biological Methods. 1 (2), 10 (2014).</li><li>Lagoy, R. C., Larsen, E., Lawler, D., White, H., Albrecht, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microfluidic+devices+for+behavioral+analysis,+microscopy,+and+neuronal+imaging+in+Caenorhabditis+elegans.">Microfluidic devices for behavioral analysis, microscopy, and neuronal imaging in Caenorhabditis elegans.</a> Methods in Molecular Biology. 2468, 293-318 (2022).</li><li>NeuroTracker. GitHub Available from: <a target="_blank" href="https://github.com/albrechtLab/Neurotracker">https://github.com/albrechtLab/Neurotracker</a> (2022)</li><li>NeuroTracker User Guide. GitHub Available from: <a target="_blank" href="https://github.com/albrechtLab/Neurotracker">https://github.com/albrechtLab/Neurotracker</a> (2022)</li><li>Schindelin, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fiji:+An+open-source+platform+for+biological-image+analysis.">Fiji: An open-source platform for biological-image analysis.</a> Nature Methods. 9 (7), 676-682 (2012).</li><li>Galotto, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chitin+triggers+calcium-mediated+immune+response+in+the+plant+model+Physcomitrella+patens.">Chitin triggers calcium-mediated immune response in the plant model Physcomitrella patens.</a> Molecular Plant-Microbe Interactions. 33 (7), 911-920 (2020).</li><li>Qian, X., Song, H., Ming, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Brain+organoids:+Advances,+applications+and+challenges.">Brain organoids: Advances, applications and challenges.</a> Development. 146 (8), 166074 (2019).</li><li>Ventimiglia, D., Bargmann, C. 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The authors have no conflicts of interest to disclose.

In this protocol, we describe an open-access microscopy system for the assessment of neural activity phenomena using the temporally precise delivery of different stimulus patterns. The microfluidic platform delivers repeatable stimuli while keeping tens of animals in the microscope field of view. Few commercial microscopy software packages allow for the easy programming of various stimulus timing patterns, and those that do often require the manual entry of each pattern or proprietary file formats. In contrast, experimen...

Calcium imaging techniques have allowed the noninvasive recording of in vivo neural dynamics in real time using fluorescence microscopy and genetically encoded calcium indicators expressed in target cells1,2,3. These sensors typically use a green fluorescent protein (GFP), such as the GFP-calmodulin-M13 peptide (GCaMP) family, to increase the fluorescence intensity upon neuronal activation and elevated intracellular calcium levels. Calcium imaging has been especially powerful in the nematode C. elegans for examining how neurons and neural circuits function in living, behaving animals4,5,6,7,8,9,10, as their transparent nature means no surgical process is required for optical access, and cell-specific gene promoters target expression to the cells of interest. These techniques often make use of microfluidic devices, which provide precisely controlled environments to study biological, chemical, and physical phenomena at a small physical scale11,12. Microfluidic devices abound for measuring neural activity, with new designs continually under development, and they are readily fabricated in the research lab. However, many designs trap a single animal at a time, limiting the experimental throughput7,9,13. Neural responses often vary substantially across animals due to differences in prior experience, internal states such as stress or hunger, or stochastic factors such as gene expression levels. These differences establish a need for methods that can simultaneously stimulate and observe many animals and extract information from individuals4.
In addition, certain neuromodulatory phenomena become apparent only under specific stimulation conditions, such as temporal inhibition14, which refers to the brief suppression of responses when stimulation occurs in rapid succession. Electrophysiological systems can drive neural activity across a broad stimulus space for this purpose, modulating, for example, the electrical pulse current, voltage, frequency, waveform, duty cycle, and timing of periodic stimulus trains. Indirect stimulation by naturally detected stimuli or optogenetic systems would benefit from a similar breadth of control mechanisms. Currently, many natural stimuli are presented in a simple &#34;on-off&#34; manner, such as odor presentation and removal, using commercial systems that have been slow to add flexibility. However, inexpensive microcontrollers can now automate the delivery of several types of stimuli in a manner that is customizable to the researchers&#39; needs. Combined with microfluidics, these systems have achieved the goal of increased experimental throughput and flexibility, allowing neural responses to a variety of precise stimuli to be measured simultaneously in many animals4,6. Multimodal stimulation can be used to further interrogate the neuronal circuitry, such as by monitoring changes in neural excitability when consistently stimulating before, during, and after an orthogonal perturbation such as drug exposure4. The benefits of inexpensive, open microscopy systems are clear for advancing scientific research, yet in practice, the need for part sourcing, construction, and performance validation can impede the adoption of these techniques.
This protocol aims to alleviate some of these technical challenges. Whereas previous protocols have focused on microfluidic device use and basic stimulation9,15,17, we describe here the construction and use of a flexible, automated, multimodal stimulus delivery system for neural imaging in C. elegans or other small organisms utilizing previously described microfluidic devices4. The open-source system is programmed via simple text files to define the experiments, and the NeuroTracker data analysis program semi-automatically extracts the neural activity data from the microscope videos. We demonstrate this system with examples of assessing temporal inhibition, disinhibition, and stimulus crosstalk using the chemosensory neuron AWA, which depolarizes in response to different food odors or in response to light when expressing optogenetic light-sensitive ion channels5,6.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Bacterial strains</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>E.&#160;coli (OP50)</td>
			<td>Caenorhabditis Genetics Center (CGC)</td>
			<td>Cat# OP50</td>
			<td></td>
		</tr>
		<tr>
			<td>Experimental models: Organisms/strains</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>C.&#160;elegans strains expressing GCaMP (and optionally, Chrimson) in desired neurons</td>
			<td>Caenorhabditis Genetics Center (CGC) or corresponding authors of published work</td>
			<td>NZ1091, for example</td>
			<td></td>
		</tr>
		<tr>
			<td>Chemicals, Treatments, and Worm Preparation Supplies</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>2,3-Butanedione</td>
			<td>Sigma-Aldrich</td>
			<td>Cat# B85307</td>
			<td>diacetyl, example chemical stimulus</td>
		</tr>
		<tr>
			<td>Calcium chloride, CaCl2</td>
			<td>Sigma-Aldrich</td>
			<td>Cat# C3881</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescein, Sodium salt</td>
			<td>Sigma-Aldrich</td>
			<td>Cat# F6377</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass water repellant</td>
			<td>Rain-X</td>
			<td>Cat #800002250</td>
			<td>glass hydrophobic treatment (single-use)</td>
		</tr>
		<tr>
			<td>Magnesium chloride, MgCl2</td>
			<td>Sigma-Aldrich</td>
			<td>Cat# M2393</td>
			<td></td>
		</tr>
		<tr>
			<td>Nematode Growth Medium (NGM) agar</td>
			<td>Genesee</td>
			<td>Cat #: 20-273NGM</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dishes (60&#160;mm)</td>
			<td>Tritech</td>
			<td>Cat #T3305</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly(dimethyl siloxane) (PDMS): Sylgard 184</td>
			<td>Dow Chemical</td>
			<td>Cat# 1673921</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium phosphate monobasic</td>
			<td>Sigma-Aldrich</td>
			<td>Cat# P5655</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium phosphate dibasic</td>
			<td>Sigma-Aldrich</td>
			<td>Cat# P8281</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride, NaCl</td>
			<td>Sigma-Aldrich</td>
			<td>Cat# S7653</td>
			<td></td>
		</tr>
		<tr>
			<td>(tridecafluoro-1,1,2,2-tetrahydrooctyl)trichlorosilane (TFOCS)</td>
			<td>Gelest</td>
			<td>CAS# 78560-45-9</td>
			<td>glass hydrophobic treatment (durable)</td>
		</tr>
		<tr>
			<td>Software and algorithms</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Arduino IDE</td>
			<td>Arduino</td>
			<td>https://www.arduino.cc/en/software</td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td>NIH</td>
			<td>https://imagej.nih.gov/ij/</td>
			<td></td>
		</tr>
		<tr>
			<td>MATLAB</td>
			<td>MathWorks</td>
			<td>https://www.mathworks.com/products/matlab.html</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro-manager</td>
			<td>Micro-manager</td>
			<td>https://micro-manager.org/</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope control software</td>
			<td>Albrecht Lab</td>
			<td>https://github.com/albrechtLab/MicroscopeControl</td>
			<td></td>
		</tr>
		<tr>
			<td>Neurotracker data analysis software</td>
			<td>Albrecht Lab</td>
			<td>https://github.com/albrechtLab/Neurotracker</td>
			<td></td>
		</tr>
		<tr>
			<td>Automated Microscope and Stimulation System</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Axio Observer.A1 inverted microscope set up for epifluorescence (GFP filter cubes, 5&#215; objective or similar)</td>
			<td>Zeiss</td>
			<td>Cat #491237-0012-000</td>
			<td></td>
		</tr>
		<tr>
			<td>Excelitas X-cite XYLIS LED illuminator</td>
			<td>Excelitas</td>
			<td>Cat #XYLIS</td>
			<td></td>
		</tr>
		<tr>
			<td>Orca Flash 4.0 Digital sCMOS camera</td>
			<td>Hamamatsu</td>
			<td>Cat #C11440-22CU</td>
			<td></td>
		</tr>
		<tr>
			<td>Arduino nano</td>
			<td>Arduino</td>
			<td>Cat #A000005</td>
			<td></td>
		</tr>
		<tr>
			<td>3-way Miniature Diapragm Isolation Valve (LQX12)</td>
			<td>Parker</td>
			<td>Cat #LQX12-3W24FF48-000</td>
			<td>Valve 1: Control</td>
		</tr>
		<tr>
			<td>2-way normally-closed (NC) Pinch Valve</td>
			<td>Bio-Chem Valve Inc</td>
			<td>Cat #075P2-S432</td>
			<td>Valve 2: Outflow</td>
		</tr>
		<tr>
			<td>3-way Pinch Valve</td>
			<td>NResearch</td>
			<td>Cat #161P091</td>
			<td>Valve 3: Stimulus selection</td>
		</tr>
		<tr>
			<td>Optogenetic stimulation LED and controller (615 nm)</td>
			<td>Mightex</td>
			<td>Cat #PLS-0625-030-S and #SLA-1200-2</td>
			<td></td>
		</tr>
		<tr>
			<td>ValveLink 8.2 digital/manual valve controller</td>
			<td>AutoMate Scientific</td>
			<td>Cat #01-18</td>
			<td></td>
		</tr>
		<tr>
			<td>Wires and connectors</td>
			<td>various</td>
			<td>See Fig. 2 of Cell STARS Protocol (Lawler, 2021)</td>
			<td></td>
		</tr>
		<tr>
			<td>Microfluidic Device Preparation</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dremel variable speed rotary cutter 4000&#160;</td>
			<td>Dremel</td>
			<td>Cat #F0134000AB</td>
			<td>Set speed to 5k RPM for cutting glass</td>
		</tr>
		<tr>
			<td>Dremel drill press rotary tool workstation</td>
			<td>Dremel</td>
			<td>Cat #220-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Diamond drill bit</td>
			<td>Dremel</td>
			<td>Cat #7134</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass slide, 1&#160;mm thick</td>
			<td>VWR</td>
			<td>Cat #75799-268</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass scribe (Diamond scriber)</td>
			<td>Ted Pella</td>
			<td>Cat #54468</td>
			<td></td>
		</tr>
		<tr>
			<td>Luer 3-way stopcock</td>
			<td>Cole-Parmer</td>
			<td>Cat #EW-30600-07</td>
			<td></td>
		</tr>
		<tr>
			<td>Luer 23 G blunt needle</td>
			<td>VWR</td>
			<td>Cat #89134-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Microfluidic device</td>
			<td>Corresponing author or fabricate from CAD files associated with this article</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Microfluidic device clamp</td>
			<td>Warner Instruments (or machine shop)</td>
			<td>P-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Microfluidic tubing, 0.02&#8243; ID</td>
			<td>Cole-Parmer</td>
			<td>Cat #EW-06419-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Tube 19 G, 0.5&#8243;</td>
			<td>New England Small Tube</td>
			<td>Cat #NE-1027-12</td>
			<td></td>
		</tr>
	</tbody>
</table>

automated multimodal stimulation and simultaneous neuronal recording from multiple small organisms

Fluorescent genetically encoded calcium indicators have contributed greatly to our understanding of neural dynamics from the level of individual neurons to entire brain circuits. However, neural responses may vary due to prior experience, internal states, or stochastic factors, thus generating the need for methods that can assess neural function across many individuals at once. Whereas most recording techniques examine a single animal at a time, we describe the use of wide-field microscopy to scale up neuronal recordings to dozens of Caenorhabditis elegans or other sub-millimeter-scale organisms at once. Open-source hardware and software allow great flexibility in programming fully automated experiments that control the intensity and timing of various stimulus types, including chemical, optical, mechanical, thermal, and electromagnetic stimuli. In particular, microfluidic flow devices provide precise, repeatable, and quantitative control of chemosensory stimuli with sub-second time resolution. The NeuroTracker semi-automated data analysis pipeline then extracts individual and population-wide neural responses to uncover functional changes in neural excitability and dynamics. This paper presents examples of measuring neuronal adaptation, temporal inhibition, and stimulus crosstalk. These techniques increase the precision and repeatability of stimulation, allow the exploration of population variability, and are generalizable to other dynamic fluorescent signals in small biosystems from cells and organoids to whole organisms and plants.

We present several examples of stimulus patterns that assess different neural phenomena, including temporal inhibition, adaptation, and disinhibition. Temporal inhibition is the momentary suppression of a neural response to a second stimulus presentation occurring shortly after the initial presentation14. To test this phenomenon, in a paired-pulse experiment, eight patterns consisting of two 1 s odorant pulses separated by an interval ranging from 0 s to 20 s were presented (<stro...

Watch this Scientific Journal Video about Automated Multimodal Stimulation and Simultaneous Neuronal Recording from Multiple Small Organisms at JoVE.com

Automated Multimodal Stimulation and Simultaneous Neuronal Recording from Multiple Small Organisms

We present a method for the flexible chemical and multimodal stimulation and recording of simultaneous neural activity from many Caenorhabditis elegans worms. This method uses microfluidics, open-source hardware and software, and supervised automated data analysis to enable the measurement of neuronal phenomena such as adaptation, temporal inhibition, and stimulus crosstalk.

Fluorescent genetically encoded calcium indicators have contributed greatly to our understanding of neural dynamics from the level of individual neurons ...

automated-multimodal-stimulation-simultaneous-neuronal-recording-from

Research

JoVE Journal

Neuroscience

890 Views.  Worcester Polytechnic Institute. We present a method for the flexible chemical and multimodal stimulation and recording of simultaneous neural activity from many Caenorhabditis elegans worms. This method uses microfluidics, open-source hardware and software, and supervised automated data analysis to enable the measurement of neuronal phenomena such as adaptation, temporal inhibition, and stimulus crosstalk.

Video: Automated Multimodal Stimulation and Simultaneous Neuronal Recording from Multiple Small Organisms

Design and Construction of a Cost Effective Headstage for Simultaneous Neural Stimulation and Recording in the Water Maze

Headstage preamplifiers and source followers are commonly used to study neural activity in behavioral neurophysiology experiments. Available commercial products are often expensive, not easily customized, and not submersible. Here we describe a method to design and build a customized, integrated circuit headstage for simultaneous 4-channel neural recording and 2-channel simulation in awake, behaving animals. The headstage is designed using a free, commercially available CAD-type design package, and can be modified easily to accommodate different scales (e.g. to add channels). A customized printed circuit board is built using surface mount resistors, capacitors and operational amplifiers to construct the unity gain source follower circuit. The headstage is made water-proof with a combination of epoxy, parafilm and a synthetic rubber putty. We have successfully used this device to record local field potentials and stimulate different brain regions simultaneously via independent channels in rats swimming in a water maze. The total cost is &lt; $30/unit and can be manufactured readily.

We present a low-cost method to design and construct a light headstage pre-amplifier system with simultaneous neural recording and stimulation capability. This device can be waterproofed for use in swimming animals.

Headstage preamplifiers and source followers are commonly used to study neural activity in behavioral neurophysiology experiments. Available commercial ...

Automated Sholl Analysis of Digitized Neuronal Morphology at Multiple Scales

Neuronal morphology plays a significant role in determining how neurons function and communicate1-3. Specifically, it affects the ability of neurons to receive inputs from other cells2 and contributes to the propagation of action potentials4,5. The morphology of the neurites also affects how information is processed. The diversity of dendrite morphologies facilitate local and long range signaling and allow individual neurons or groups of neurons to carry out specialized functions within the neuronal network6,7. Alterations in dendrite morphology, including fragmentation of dendrites and changes in branching patterns, have been observed in a number of disease states, including Alzheimer's disease8, schizophrenia9,10, and mental retardation11. The ability to both understand the factors that shape dendrite morphologies and to identify changes in dendrite morphologies is essential in the understanding of nervous system function and dysfunction.
Neurite morphology is often analyzed by Sholl analysis and by counting the number of neurites and the number of branch tips. This analysis is generally applied to dendrites, but it can also be applied to axons. Performing this analysis by hand is both time consuming and inevitably introduces variability due to experimenter bias and inconsistency. The Bonfire program is a semi-automated approach to the analysis of dendrite and axon morphology that builds upon available open-source morphological analysis tools. Our program enables the detection of local changes in dendrite and axon branching behaviors by performing Sholl analysis on subregions of the neuritic arbor. For example, Sholl analysis is performed on both the neuron as a whole as well as on each subset of processes (primary, secondary, terminal, root, etc.) Dendrite and axon patterning is influenced by a number of intracellular and extracellular factors, many acting locally. Thus, the resulting arbor morphology is a result of specific processes acting on specific neurites, making it necessary to perform morphological analysis on a smaller scale in order to observe these local variations12.
The Bonfire program requires the use of two open-source analysis tools, the NeuronJ plugin to ImageJ and NeuronStudio. Neurons are traced in ImageJ, and NeuronStudio is used to define the connectivity between neurites. Bonfire contains a number of custom scripts written in MATLAB (MathWorks) that are used to convert the data into the appropriate format for further analysis, check for user errors, and ultimately perform Sholl analysis. Finally, data are exported into Excel for statistical analysis. A flow chart of the Bonfire program is shown in Figure 1.

We have developed a computer program to analyze neuronal morphology. In combination with two existing open source analysis tools, our program performs Sholl analysis and determines the number of neurites, branch points, and neurite tips. The analyses are performed so that local changes in neurite morphology can be observed.

Neuronal morphology plays a significant role in determining how neurons function and communicate1-3.  Specifically, it affects the ability of neurons to ...

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated. Thus, simultaneous recordings of neuronal activity and behavior are essential to correlate brain activity to behavior. For such behavioral analyses, the fruit fly, Drosophila melanogaster, allows us to incorporate genetically encoded calcium indicators such as GCaMP1, to monitor neuronal activity, and to use sophisticated genetic manipulations for optogenetic or thermogenetic techniques to specifically activate identified neurons2-5. Use of a thermogenetic technique has led us to find critical neurons for feeding behavior (Flood et al., under revision). As a main part of feeding behavior, a Drosophila adult extends its proboscis for feeding6 (proboscis extension response; PER), responding to a sweet stimulus from sensory cells on its proboscis or tarsi. Combining the protocol for PER7 with a calcium imaging technique8 using GCaMP3.01, 9, I have established an experimental system, where we can monitor activity of neurons in the feeding center &#x2013; the suboesophageal ganglion (SOG), simultaneously with behavioral observation of the proboscis. I have designed an apparatus ("Fly brain Live Imaging and Electrophysiology Stage": "FLIES") to accommodate a Drosophila adult, allowing its proboscis to freely move while its brain is exposed to the bath for Ca2+ imaging through a water immersion lens. The FLIES is also appropriate for many types of live experiments on fly brains such as electrophysiological recording or time lapse imaging of synaptic morphology. Because the results from live imaging can be directly correlated with the simultaneous PER behavior, this methodology can provide an excellent experimental system to study information processing of neuronal networks, and how this cellular activity is coupled to plastic processes and memory.

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

The fruit fly, Drosophila melanogaster, extends its proboscis for feeding, responding to a sugar stimulus from its proboscis or tarsus. I have combined observations of the proboscis extension response (PER) with a calcium imaging technique, allowing us to monitor the activity of neurons in the brain, simultaneously with behavioral observation.

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated.  Thus, ...

A Fully Automated and Highly Versatile System for Testing Multi-cognitive Functions and Recording Neuronal Activities in Rodents

We have developed a fully automated system for operant behavior testing and neuronal activity recording by which multiple cognitive brain functions can be investigated in a single task sequence. The unique feature of this system is a custom-made, acoustically transparent chamber that eliminates many of the issues associated with auditory cue control in most commercially available chambers. The ease with which operant devices can be added or replaced makes this system quite versatile, allowing for the implementation of a variety of auditory, visual, and olfactory behavioral tasks. Automation of the system allows fine temporal (10 ms) control and precise time-stamping of each event in a predesigned behavioral sequence. When combined with a multi-channel electrophysiology recording system, multiple cognitive brain functions, such as motivation, attention, decision-making, patience, and rewards, can be examined sequentially or independently.

In this report, we present a fully automated and highly versatile system capable of simultaneously testing multi-cognitive behaviors and recording neuronal activities for rodents.

We have developed a fully automated system for operant behavior testing and neuronal activity recording by which multiple cognitive brain functions can be ...

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

Much information about the coupling of presynaptic ionic currents with the release of neurotransmitter has been obtained from invertebrate preparations, most notably the squid giant synapse1. However, except for the preparation described here, few vertebrate preparations exist in which it is possible to make simultaneous measurements of neurotransmitter release and presynaptic ionic currents. Embryonic Xenopus motoneurons and muscle cells can be grown together in simple culture medium at room temperature; they will form functional synapses within twelve to twenty-four hours, and can be used to study nerve and muscle cell development and synaptic interactions for several days (until overgrowth occurs). Some advantages of these co-cultures over other vertebrate preparations include the simplicity of preparation, the ability to maintain the cultures and work at room temperature, and the ready accessibility of the synapses formed2-4. The preparation has been used widely to study the biophysical properties of presynaptic ion channels and the regulation of transmitter release5-8. In addition, the preparation has lent itself to other uses including the study of neurite outgrowth and synaptogenesis9-12, molecular mechanisms of neurotransmitter release13-15, the role of diffusible messengers in neuromodulation16,17, and in vitro synaptic plasticity18-19.

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

This video demonstrates the procedures used to grow primary cultures of embryonic Xenopus nerve and muscle cells and the usefulness of this preparation for making simultaneous pre- and post-synaptic patch clamp recordings.

Much information about the coupling of  presynaptic ionic currents with the release of neurotransmitter has been obtained  from invertebrate preparations, ...

Simultaneous Electrophysiological Recording and Calcium Imaging of Suprachiasmatic Nucleus Neurons

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in regulating the intracellular calcium concentration. Changing environmental conditions, such as the light-dark cycle, can modify neuronal and neural network activity and the expression of a family of circadian clock genes within the suprachiasmatic nucleus (SCN), the location of the master circadian clock in the mammalian brain. Excitatory synaptic transmission leads to an increase in the postsynaptic Ca2+ concentration that is believed to activate the signaling pathways that shifts the rhythmic expression of circadian clock genes. Hypothalamic slices containing the SCN were patch clamped using microelectrodes filled with an internal solution containing the calcium indicator bis-fura-2. After a seal was formed between the microelectrode and the SCN neuronal membrane, the membrane was ruptured using gentle suction and the calcium probe diffused into the neuron filling both the soma and dendrites. Quantitative ratiometric measurements of the intracellular calcium concentration were recorded simultaneously with membrane potential or current. Using these methods it is possible to study the role of changes of the intracellular calcium concentration produced by synaptic activity and action potential firing of individual neurons. In this presentation we demonstrate the methods to simultaneously record electrophysiological activity along with intracellular calcium from individual SCN neurons maintained in brain slices.

Procedures are described to perform simultaneous recordings of membrane potential or current and changes of intracellular calcium concentration. Suprachiasmatic nucleus neurons are filled with the calcium indicator bis-fura-2 using a patch clamp electrode in the whole cell patch clamp configuration.

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in ...

Deep Brain Stimulation with Simultaneous fMRI in Rodents

In order to visualize the global and downstream neuronal responses to deep brain stimulation (DBS) at various targets, we have developed a protocol for using blood oxygen level dependent (BOLD) functional magnetic resonance imaging (fMRI) to image rodents with simultaneous DBS. DBS fMRI presents a number of technical challenges, including accuracy of electrode implantation, MR artifacts created by the electrode, choice of anesthesia and paralytic to minimize any neuronal effects while simultaneously eliminating animal motion, and maintenance of physiological parameters, deviation from which can confound the BOLD signal.&#160;Our laboratory has developed a set of procedures that are capable of overcoming most of these possible issues. For electrical stimulation, a homemade tungsten bipolar microelectrode is used, inserted stereotactically at the stimulation site in the anesthetized subject. In preparation for imaging, rodents are fixed on a plastic headpiece and transferred to the magnet bore. For sedation and paralysis during scanning, a cocktail of dexmedetomidine and pancuronium is continuously infused, along with a minimal dose of isoflurane; this preparation minimizes the BOLD ceiling effect of volatile anesthetics. In this example experiment, stimulation of the subthalamic nucleus (STN) produces BOLD responses which are observed primarily in ipsilateral cortical regions, centered in motor cortex. Simultaneous DBS and fMRI allows the unambiguous modulation of neural circuits dependent on stimulation location and stimulation parameters, and permits observation of neuronal modulations free of regional bias. This technique may be used to explore the downstream effects of modulating neural circuitry at nearly any brain region, with implications for both experimental and clinical DBS.

This protocol describes a standard method for simultaneous functional magnetic resonance imaging and deep brain stimulation in the rodent. The combined use of these experimental tools allows for the exploration of global downstream activity in response to electrical stimulation at virtually any brain target.

In order to visualize the global and downstream neuronal responses to deep brain stimulation (DBS) at various targets, we have developed a protocol for ...

Gold Nanorod-assisted Optical Stimulation of Neuronal Cells

Recent studies have demonstrated that nerves can be stimulated in a variety of ways by the transient heating associated with the absorption of infrared light by water in neuronal tissue. This technique holds great potential for replacing or complementing standard stimulation techniques, due to the potential for increased localization of the stimulus and minimization of mechanical contact with the tissue. However, optical approaches are limited by the inability of visible light to penetrate deep into tissues. Moreover, thermal modelling suggests that cumulative heating effects might be potentially hazardous when multiple stimulus sites or high laser repetition rates are used. The protocol outlined below describes an enhanced approach to the infrared stimulation of neuronal cells. The underlying mechanism is based on the transient heating associated with the optical absorption of gold nanorods, which can cause triggering of neuronal cell differentiation and increased levels of intracellular calcium activity. These results demonstrate that nanoparticle absorbers can enhance and/or replace the process of infrared neural stimulation based on water absorption, with potential for future applications in neural prostheses and cell therapies.

This protocol outlines how to use the transient heating associated with the optical absorption of gold nanorods to stimulate differentiation and intracellular calcium activity in neuronal cells. These results potentially open up new applications in neural prostheses and fundamental studies in neuroscience.

Recent studies have demonstrated that nerves can be stimulated in a variety of ways by the transient heating associated with the absorption of infrared ...

Simultaneous Recording of Electroretinography and Visual Evoked Potentials in Anesthetized Rats

The electroretinogram (ERG) and visual evoked potential (VEP) are commonly used to assess the integrity of the visual pathway. The ERG measures the electrical responses of the retina to light stimulation, while the VEP measures the corresponding functional integrity of the visual pathways from the retina to the primary visual cortex following the same light event. The ERG waveform can be broken down into components that reflect responses from different retinal neuronal and glial cell classes. The early components of the VEP waveform represent the integrity of the optic nerve and higher cortical centers. These recordings can be conducted in isolation or together, depending on the application. The methodology described in this paper allows simultaneous assessment of retinal and cortical visual evoked electrophysiology from both eyes and both hemispheres. This is a useful way to more comprehensively assess retinal function and the upstream effects that changes in retinal function can have on visual evoked cortical function.

This protocol describes simultaneous measurement of electroretinogram and visual evoked potentials in anesthetized rats.

The electroretinogram (ERG) and visual evoked potential (VEP) are commonly used to assess the integrity of the visual pathway. The ERG measures the ...

Chronic Transcranial Electrical Stimulation and Intracortical Recording in Rats

Transcranial electrical stimulation (TES) is a powerful and relatively simple approach to diffusely influence brain activity either randomly or in a closed-loop event-triggered manner. Although many studies are focusing on the possible benefits and side-effects of TES in healthy and pathologic brains, there are still many fundamental open questions regarding the mechanism of action of the stimulation. Therefore, there is a clear need for a robust and reproducible method to test the acute and the chronic effects of TES in rodents. TES can be combined with regular behavioral, electrophysiological, and imaging techniques to investigate neuronal networks in vivo. The implantation of transcranial stimulation electrodes does not impose extra constraints on the experimental design while it offers a versatile, flexible tool to manipulate brain activity. Here we provide a detailed, step-by-step protocol to fabricate and implant transcranial stimulation electrodes to influence brain activity in a temporally constrained manner for months.

This detailed protocol describes transcranial stimulation electrode placement on the temporal bone in order to investigate the short- and long-term effects of transcranial electrical stimulation in freely moving rats.

Transcranial electrical stimulation (TES) is a powerful and relatively simple approach to diffusely influence brain activity either randomly or in a ...