This work was financed by FEDER - Fundo Europeu de Desenvolvimento Regional funds through the COMPETE 2020 - Operacional Programme for Competitiveness and Internationalisation (POCI), Portugal 2020, and by Portuguese funds through FCT - Funda&#231;&#227;o para a Ci&#234;ncia e a Tecnologia/ Minist&#233;rio da Ci&#234;ncia, Tecnologia e Ensino Superior in the framework of the project PTDC/CTM-NAN/3146/2014 (POCI-01-0145-FEDER-016623). Jos&#233; C Mateus was supported by FCT (PD/BD/135491/2018). Paulo Aguiar was supported by Programa Ci&#234;ncia - Programa Operacional Potencial Humano (POPH) - Promotion of Scientific Employment, ESF and MCTES and program Investigador FCT, POPH and Fundo Social Europeu. The microfluidic devices were fabricated at INESC - Microsystems and Nanotechnologies, Portugal, under the supervision of Jo&#227;o Pedro Conde and Virginia Chu.

All procedures involving animals were performed according to the European Union (EU) Directive 2010/63/EU (transposed to Portuguese legislation by Decreto-Lei 113/2013). The experimental protocol (0421/000/000/2017) was approved by the ethics committee of both the Portuguese Official Authority on animal welfare and experimentation (Dire&#231;&#227;o-Geral de Alimenta&#231;&#227;o e Veterin&#225;ria - DGAV) and of the host Institution.
1. Preparation of Culture Media and Other Solutions
NOTE: Freshly prepare the coating solutions on the day of its use. The unused coating solutions and culture media may be stored at -20 &#176;C or 4 &#176;C as detailed below.
<ol>
	<li>&#160;Prepare 10 mL of a 0.05% (v/v) Poly(ethylene imine) (PEI) working solution.
	<ol>
		<li>Prepare 20 mL of 0.25% (w/v) PEI stock solution by dissolving purified PEI (see Table of Materials) in sterile water.</li>
		<li>Dilute 2 mL of 0.25% (w/v) PEI stock solution in 8 mL of sterile distilled water. Mix well and use. Unused solution can be stored at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Prepare 1 mL of a 5 &#181;g/mL laminin solution.
	<ol>
		<li>Dilute 5 &#181;L of 1 mg/mL laminin stock solution in 995 &#181;L of basal medium (see Table of Materials). Mix well and use. Unused solution can be stored frozen at -20&#176;C for up to 1-2 weeks.</li>
	</ol>
	</li>
	<li>Prepare 1 L of HEPES-buffered Hank&#39;s Balanced Salt Solution (H-HBSS).
	<ol>
		<li>Prepare 1 M HEPES solution by dissolving 11.9 g of HEPES powder in 50 mL of ultrapure water. Adjust pH to 7.2.</li>
		<li>Prepare H-HBSS solution (without calcium and magnesium) by dissolving 5.33 mM potassium chloride, 0.44 mM potassium phosphate monobasic, 138 mM sodium chloride, 0.3 mM sodium phosphate dibasic and 5.6 mM glucose in 800 mL of ultrapure water.</li>
		<li>Add 15 mL of 1 M HEPES solution (final concentration of 15 mM).</li>
		<li>Adjust pH to 0.2-0.3 units below the desired pH 7.4 using 1 N HCl or 1 N NaOH. Add ultrapure water to make up the final solution volume. 
		NOTE pH tends to rise during filtration.</li>
		<li>Sterilize by filtration using a 0.22 &#181;m porosity poly(ether sulfone) (PES) membrane.</li>
	</ol>
	</li>
	<li>Prepare 10 mL of H-HBSS containing 10% heat-inactivated Fetal Bovine Serum (hiFBS).
	<ol>
		<li>Dilute 1 mL of hiFBS in 9 mL of H-HBSS. Mix well and use. Unused solution can be stored at 4 &#176;C for 3-4 weeks.</li>
	</ol>
	</li>
	<li>Prepare 5 mL of 1.5 mg/mL trypsin solution.
	<ol>
		<li>Immediately before use, dissolve 7.5 mg of trypsin in 5 mL of H-HBSS. Sterilize by filtration using a 0.22 &#181;m porosity PES membrane.</li>
	</ol>
	</li>
	<li>Prepare 50 mL of supplemented medium.
	<ol>
		<li>In a 50 mL conical tube, mix 48.5 mL of basal medium (see Table of Materials), 2% (v/v) B-27 supplement, 0.5 mM L-glutamine and 1% Pen/Strep (10,000 U/mL penicillin and 10,000 &#181;g/mL streptomycin). Supplemented medium can be stored at 4 &#176;C for 3-4 weeks.</li>
	</ol>
	</li>
</ol>
2. Microfluidic and MEA Devices Preparation
NOTE: Perform steps 2.1-2.11 on the day before cell seeding.
<ol>
	<li>Clean microfluidic devices to remove fabrication and other debris using vinyl tape. Gently press the tape against the device to reach all areas of the device.</li>
	<li>Air plasma-clean the MEA for 3 min (0.3 mbar) to clean the surface and make it more hydrophilic.</li>
	<li>Briefly submerge microfluidic devices in 70% ethanol and allow them to air-dry inside a laminar flow hood.</li>
	<li>Place each MEA in a sterile 90 mm Petri dish and sterilize them by 15 min of ultraviolet (UV) light exposure.</li>
	<li>Coat the MEA central surface area by incubating with 500 &#181;L of 0.05% (v/v) PEI solution for at least 1 h at 37 &#176;C. 
	NOTE: As described by others17, PEI coating of MEAs results in less cell clustering than using poly(lysine) coating.</li>
	<li>Aspirate the PEI coating solution from the MEA surface and wash MEAs 4x with 1 mL of sterile distilled water. Be careful to not touch the MEA surface during washing steps as this can damage the electrodes.</li>
	<li>Guided by a stereomicroscope inside a laminar flow hood, center the MEA chip and add 1 mL of sterile water to the central area of MEA.</li>
	<li>Place the microfluidic device on top of the MEA surface.</li>
	<li>Carefully align the microgrooves of microfluidic device with the microelectrode grid of the MEA.</li>
	<li>When correctly aligned, carefully aspirate the excess water without touching the MEA or the microfluidic device.</li>
	<li>Incubate the &#181;EFs overnight at 37 &#176;C to dry and allow complete attachment. To facilitate the attachment, gently press the microfluidic device against the MEA. 
	NOTE: Perform step 2.12 on the day of cell seeding.</li>
	<li>Once the &#181;EFs have completely dried, load the microfluidic wells with 150 &#181;L of 5 &#181;g/mL laminin coating solution and incubate at 37 &#176;C for at least 2 h before cell seeding. 
	NOTE: When loading &#181;EF with laminin solution, ensure that the solution fills the microgrooves. use a vacuum to remove any air bubble that might be present within the microgrooves.</li>
</ol>
3. Prefrontal Cortex Dissection
<ol>
	<li>Prepare the dissection area for embryo removal.
	<ol>
		<li>Disinfect the working surface and the surgical instruments with 70% ethanol. 
		NOTE: Though this dissection is not a sterile procedure, disinfection helps to reduce chances of contamination.</li>
		<li>Use a clean disposable diaper to confine the dissection area and lay out the surgical instruments for embryo removal.</li>
		<li>Prepare 4 90 mm Petri dishes filled with cold H-HBSS and place them on a tray with ice near the dissection area.</li>
	</ol>
	</li>
	<li>Euthanize an E-18 time-pregnant Wistar rat by carbon dioxide (CO2) inhalation.</li>
	<li>Upon completion of the procedure, remove the animal from the CO2 chamber and confirm death by a secondary method of euthanasia, such as exsanguination.</li>
	<li>Place the animal ventral side up on the disposable diaper, spray the lower abdomen with 70% ethanol and, with the help of forceps and scissor, make a U-shape cut through the skin to expose the uterus.</li>
	<li>Carefully remove the embryos from the uterus by cutting away any connective tissue and place them in one of the Petri dishes with cold H-HBSS.</li>
	<li>Remove each embryo from its embryonic sac and, with the help of forceps and scissor, decapitate the embryo.</li>
	<li>Transfer the embryo&#180;s head to a second Petri dish filled with cold H-HBSS.</li>
	<li>Repeat steps 3.6 - 3.7 for each embryo.</li>
	<li>Dissect the prefrontal cortex.
	<ol>
		<li>Transfer an embryo&#180;s head to a third Petri dish.</li>
		<li>Use a pair of forceps to expose the brain.</li>
		<li>Remove the brain from its cavity and cover it with cold H-HBSS.</li>
		<li>If present, remove and discard the olfactory bulbs. Dissect the prefrontal cortex and remove meninges.</li>
		<li>Transfer the prefrontal cortex fragments to a fourth Petri dish filled with cold H-HBSS.</li>
		<li>Repeat the steps 3.9.1 - 3.9.5 for each embryo.</li>
	</ol>
	</li>
</ol>
4. Cortical Neurons Dissociation and Culture on &#181;EF
NOTE: The following procedures were performed inside a laminar flow hood after a 15 min cycle of UV sterilization. Working surface area as well as all the materials placed inside the hood should be previously disinfected with 70% ethanol.
<ol>
	<li>As described in step 1.5, dissolve the previously weighted trypsin in 5 mL of H-HBSS and filter the solution.</li>
	<li>Collect the cortex pieces previously dissected in a total of 5 mL of H-HBSS. Transfer them to a sterile 15 mL conical tube.</li>
	<li>Add 2.3 mL of freshly prepared 1.5 mg/mL trypsin solution to the tube containing the cortex fragments.</li>
	<li>Mix the suspension and incubate at 37 &#176;C for 15 min (briefly agitate every 5 min).</li>
	<li>Stop tissue digestion and wash out trypsin.
	<ol>
		<li>Discard the supernatant containing the trypsin. Add 5 mL of H-HBSS containing 10% hiFBS to inactivate the remaining trypsin; gently agitate.</li>
		<li>Let cortex fragments settle down and then discard the supernatant.</li>
		<li>Add 5 mL of H-HBSS to remove the hiFBS residues. Let cortex fragments settle down and discard the supernatant.</li>
		<li>Wash again the cortex fragments with 5 mL of H-HBSS.</li>
		<li>Let cortex fragments settle down and discard the supernatant. Add 5 mL of supplemented neuronal medium.</li>
	</ol>
	</li>
	<li>Mechanically dissociate cortex fragments with a 5 mL serologic pipette by slowly pipetting up and down for 10-15 times. If necessary, repeat the procedure using a small caliber pipette until the cell suspension become homogenous.</li>
	<li>Discard the remaining undissociated tissues by filtering the cell suspension trough a 40 &#181;m cell strainer.</li>
	<li>Count viable cells and determine the cell density.
	<ol>
		<li>In a microtube, add 20 &#181;L of 0.4% (w/v) Trypan Blue to a 20 &#181;L cell suspension. Mix well to homogenate the solution and transfer 10 &#181;L to a Neubauer counting chamber.</li>
		<li>Count viable cells and determine the cell density of viable cells.</li>
	</ol>
	</li>
	<li>Adjust cell density to 3.6 x 107 cells/mL (if needed, centrifuge cell suspension).</li>
	<li>Immediately before cell seeding, remove laminin coating from all wells of the &#181;EF. Pipette 50 &#181;L of supplemented medium to each well of the &#181;EF axonal compartment.</li>
	<li>Seed 5 &#181;L of cell suspension in the &#181;EF somal compartment. Incubate at 37 &#176;C for 1 h, to allow cells attachment to the substrate.</li>
	<li>Gently fill each well of &#181;EF with warmed supplemented medium. Transfer the &#181;EF to a humidifier incubator at 37 &#176;C supplied with 5% CO2. 
	NOTE: To avoid quick culture medium evaporation, place an opened microtube filled with water inside the Petri dish carrying the &#181;EF.</li>
	<li>Maintain cultures for the required period by replacing 50% of culture medium every second to third day of culture. 
	NOTE: After the experiments, the microfluidics can be detached from MEAs by immersing the &#181;EF in water overnight. If needed, gently peel the microfluidics with the help of forceps. MEAs can be cleaned by overnight incubation with a 1% (v/v) commercial enzymatic detergent (see Table of Materials).</li>
</ol>
5. Recording Spontaneous Neuronal Activity
NOTE: Embryonic cortical neuron cultures on MEAs typically exhibit spontaneous activity as soon as 7 days in vitro (DIV), but take 2-3 weeks (14-21 DIV) to mature and exhibit stable activity. It is up to the experimenter to decide when to start the electrophysiological measurements based on the objectives of the study. In this protocol, the recording of neuronal activity from &#181;EF is demonstrated by using a commercial recording system (see Table of Materials for hardware details), with a heating module incorporated. For performing recordings, we used a freely available software (see Table of Materials for software details).
<ol>
	<li>Turn on the MEA recording system and the temperature controller and allow the heated base plate of the preamplifier to reach 37 &#176;C. 
	NOTE: For long-term recordings (&#62;30 min at a time) one should also have a system that maintains a CO2 atmosphere.</li>
	<li>Place the &#181;EF on the preamplifier (headstage). 
	NOTE: Ensure that the MEA chip is correctly aligned with the reference number in the upper left edge. The MEA chip we use is rotationally symmetrical, but this alignment matches the correct orientation of the pin layout of the electrodes and the hardware IDs.</li>
	<li>Close and latch the preamplifier lid.</li>
	<li>To record, open the recording software and define the recording parameters and the path of the recorded data streams (see the software manual for details on how to set-up a recording scheme). 
	NOTE: An acquisition with a sampling rate of 20 kHz is generally enough for most applications. A higher sampling frequency is advisable if more precision is required in spike timings. If one intends only to record spikes, set a high-pass filter at 300 Hz, which will attenuate the lower frequency components of the signal. If raw and filtered data are recorded simultaneously, this will significantly increase the data file size. It is always possible to do offline filtering of the raw data. To reduce data file size one can also limit microelectrodes from which to record (e.g., only microelectrodes within microgrooves).</li>
	<li>Click Start DAQ to start data acquisition. Check if the display shows running traces of activity and start recording. 
	NOTE: The noise level is usually slightly higher in the microelectrodes corresponding to the microgrooves. If the noise level is too high (peak-to-peak amplitude &#62;50 &#181;V) or unsteady in several microelectrodes, it could be due to dirt hindering a good pin-pad contact. This problem is usually solved by gently cleaning the MEA contact pads and the preamplifier pins using a cotton swab moistened with 70% ethanol.</li>
	<li>Open the recorded file in the analysis software (see Table of Materials) to quickly explore the data.</li>
</ol>
6. Data Analysis
NOTE: The following steps show how to use the &#181;SpikeHunter software, a computational tool developed at Aguiar&#39;s lab (freely available upon email request to pauloaguiar@ineb.up.pt), to analyze data recorded with &#181;EF. A graphical user interface (Figure 2) is used to load the data, identify propagating spike waves, determine their direction (anterograde or retrograde) and estimate propagation velocities. &#181;SpikeHunter is compatible with HDF5 files generated from recordings obtained using MEA2100-family systems, which can be used in conjunction with 60-, 120-, and 256-electrode MEAs. The recordings obtained using the multi-channel experimenter can be easily converted to HDF5 files using freely available software (see Table of Materials).
<ol>
	<li>Click Browse to select the recording file. Then, click the File Info button to list the sampling rate, the recording duration as well as a list of the recorded data streams (e.g., raw data, filtered data). 
	NOTE: For true propagation velocity estimations, it is recommended to select the raw data stream for analysis.</li>
	<li>Select microelectrodes (single row or column associated with microgroove) for analysis and set the spike detection threshold value (positive or negative phase) at 6x the standard deviation (SD) of the signal noise. Click Read Data to apply the event detector and obtain the identified propagation sequences. 
	NOTE: If the criteria are met, a list of detected propagation sequences will populate the analysis panel. The user can then view and interact with several analysis tools for the propagation sequence, including voltage traces, inter-microelectrode cross-correlations, single-sequence propagation velocities, a kymograph, and an audio playback tool. Although the propagation velocity estimation can be obtained for any pair of microelectrodes or as the average of the estimates for all pairs, this estimation is more reliable if the farthest pair is selected.</li>
	<li>Repeat the previous step for the microelectrode sets of interest. Each time click Save All Events to save the time and amplitude of the spikes (on each of the microelectrodes) for all identified propagation sequences to a CSV file.</li>
	<li>Import the CSV files to data analysis environments for further analysis of the patterns of activity of the culture (e.g., firing rate, inter-spike intervals).</li>
</ol>

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The authors have nothing to disclose.

The protocol presented here shows how to assemble a &#181;EF, comprised of a microfluidic device and a MEA with standard commercially-available designs, and how to analyze the recorded data.
When designing an experiment, researchers must take into account that the in vitro model is limited by the MEA fixed grid, which constrains microgroove arrangements. The use of a particular microfluidic or MEA design will depend on the specific experimental needs but, in general, the same procedure steps should apply to different &#181;EF configurations.
A critical decision to be made before &#181;EF assembly is whether the user intends to reuse the assembled &#181;EF in future experiments. Treatment with oxygen plasma on both surfaces can be done to covalently bond microfluidic devices to MEAs20. However, this sealing often makes the MEA chips unusable for further experiments, as detaching the microfluidic device irreversibly damages the passivation layer. To circumvent this problem, research groups tend to reuse the mounted &#181;EF, despite the possible drawbacks (e.g., debris from previous experiments). By carefully following the steps outlined in this protocol, from experiment to experiment, one can safely attach and detach microfluidic devices without damaging the MEA.
The use of &#181;EF allows the isolation of a set of axons within the microgrooves. The time required for a reasonable number of axons to cross the microgrooves will greatly depend on the cell seeding procedure, namely seeded cell density. Using the density specified in this protocol, neurons use to cross the whole microgroove within 3 to 5 DIV.
Depending on the culture environment (i.e., incubator and humidity level), it may be required to replace media every 2 to 3 days of culture. When renewing media, remove the media from &#181;EF wells while maintaining media within the main channels. Then, gently add media to the &#181;EF wells allowing it to slowly flow through the main channels before filling up the wells with final media volume. Following these recommendations, we are able to maintain these cultures in healthy conditions for at least a month.
A key advantage of this platform is the ability to isolate, measure and track axonal signals. Although the &#181;EF recording is simple, the great amount of data that can be generated is cumbersome to handle and requires good knowledge of data analysis. The analysis software used here, &#181;SpikeHunter21, is an advanced yet intuitive computational tool, which allows for the detailed quantification of several related measures (e.g., propagating events, propagation velocity etc.) in a few steps. Although data analysis is not the focus of this protocol, the information here provided already allows the extraction of meaningful data from a &#181;EF recording. However, it is important to note that the reliable isolation of a single axon per microgroove remains impracticable. Thus, very complex spike waveforms (due to multiple axons passing through the microgroove) may arise and affect the spike timings and propagation velocity calculations. This limitation can be attenuated by using very narrow microgrooves (below 5 &#181;m)13 and/or through spike sorting techniques21, which help distinguish different signal sources.
Even though in vitro cultures cannot recapitulate the full in vivo complexity, their combination with &#181;EF enables well-controlled bottom-up research approaches. We hope this protocol will help both beginners and proficient MEA users establishing new and reliable models for studying electrophysiological communication in neuronal circuits.

Understanding electrical communication in neuronal circuits is a fundamental step to reveal normal function, and devise therapeutic strategies to address dysfunction. Neurons integrate, compute and relay action potentials (APs) which propagate along their thin axons. Traditional electrophysiological techniques (e.g., patch clamp) are powerful techniques to study neuronal activity but are often limited to the larger cellular structures, such as the soma or the dendrites. Imaging techniques offer an alternative to study axonal signals with high spatial resolution, but they are technically difficult to perform and do not allow long-term measurements1. In this context, the combination of microelectrode arrays (MEAs) and microfluidics can make a powerful contribution in disclosing the fundamental properties of neuron&#180;s activity and signal transmission within neuronal networks in vitro2,3.
MEA technology relies on extracellular recordings of neuronal cultures. The main advantages of this electrophysiological methodology are its ability to support long-term, simultaneous stimulation and recording at multiple sites and in a non-invasive way3. MEAs are made of biocompatible, high conductive and corrosion resistant microelectrodes embedded in a glass wafer substrate. They are compatible with conventional cell culture bio-coatings, which by promoting cell adhesion significantly increase the sealing resistance between the substrate and cells3,4. Moreover, they are versatile in design and may vary in microelectrodes size, geometry and density. Overall, MEAs work as conventional cell culture vessels with the advantage of allowing concurrent live-imaging and electrophysiological recordings/stimulation.
The use of MEA technology has contributed to the study of important features of neural networks5. However, there are inherent features that limit the performance of MEAs for studying communication and APs propagation in a neuronal circuit. MEAs enable recordings from single cells and even subcellular structures like axons, but when compared to somal signals, axonal signals have a very low signal-to-noise ratio (SNR)6. Moreover, the characteristic of sensing extracellular field potentials from all sources around every microelectrode hampers the tracking of signal propagation in a neuronal circuit.
Recent studies have demonstrated, however, that better recording conditions can be achieved by having the microelectrodes aligned within narrow microchannels into which axons can grow. This configuration provides a significant increase in the SNR such that propagating axonal signals can be easily detected7,8,9,10,11,12,13. The strategy of allying microfluidic devices with MEA technology creates an electrically isolated microenvironment suitable to amplify axonal signals11. Moreover, the presence of multiple sensing microelectrodes along a microgroove is fundamental for detection and characterization of axonal signal propagation.
Such in vitro platforms with highly controllable neuronal network topographies can be adapted to many research questions14. These platforms are suitable to be used in the context of neuronal cultures but can be expanded to engineer co-culture configurations, where the communication between neurons and other cell types can be monitored and assessed. This setup thus provides very interesting conditions to explore a number of neural-related studies such as neurodevelopment, neuronal circuits, information coding, neurodegeneration and neuroregeneration. Furthermore, its combination with emerging models of human induced pluripotent stem cells15,16 can open new avenues in the development of potential therapies for human diseases that affect the nervous system.
Our lab is using this platform combining microelectrodes with microfluidics (&#181;EF) to understand neuronal processes at the cellular and network level and their implication in the physio- and pathologic nervous system. Given the value of such platform in the field of neuroscience, the purpose of this protocol is to demonstrate how to create a compartmentalized neuronal culture over substrate-integrated MEAs, how to culture neurons in this platform and how to successfully record, analyze and interpret the results from such experiments. This protocol will certainly enrich the experimental toolbox for neuronal cultures in the study of neural communication.

<table><tbody><tr><td>B-27 Suplement (50X)</td><td>Thermo Fisher Scientific</td><td>LTI17504-044</td><td></td></tr><tr><td>Branched poly(ethylene imine) (PEI), 25 kDa</td><td>Sigma-Aldrich</td><td>408727</td><td>Purify branched PEI by dialysis using a 2.5 kDa cut-off membrane for 3 days at 4&amp;deg;C against a 5 mM HCl solution (renewed daily). Freeze-dry the purified PEI.</td></tr><tr><td>Cell strainer (40 &amp;micro;m)&amp;nbsp;</td><td>Falcon</td><td>352340</td><td></td></tr><tr><td>Conical microtubes (1.5 ml)</td><td>VWR</td><td>211-0015</td><td></td></tr><tr><td>Disposable diaper, 60x40 cm</td><td>Bastos Viegas SA</td><td>455-019</td><td></td></tr><tr><td>Forceps Dumont #5, straight</td><td>Fine Science Tools</td><td>91150-20</td><td></td></tr><tr><td>Forceps Dumont #5/45</td><td>Fine Science Tools</td><td>11251-35</td><td></td></tr><tr><td>Forceps Dumont #7, curved</td><td>Fine Science Tools</td><td>91197-00</td><td></td></tr><tr><td>Heat Inactivated Fetal Bovine Serum Premium</td><td>Biowest</td><td>S181BH-500ML</td><td></td></tr><tr><td>Laminin from Engelbreth-Holm-Swarm&amp;nbsp;</td><td>Sigma-Aldrich</td><td>L2020-1MG</td><td>Prepare laminin stock solution at 1 mg/mL by dissolving the powder in the respective volume of non-supplemented medium. Store laminin solution at -20 &amp;deg;C in small aliquots (20 &amp;micro;L) to avoid repeated freeze/thaw cycles.</td></tr><tr><td>L-Glutamine 200mM&amp;nbsp;</td><td>Thermo Fisher Scientific</td><td>LTID25030-024</td><td></td></tr><tr><td>Neubauer improved counting chamber (hemocytometer)</td><td>Marienfeld</td><td>630010</td><td></td></tr><tr><td>Neurobasal Medium (1X)</td><td>Thermo Fisher Scientific</td><td>21103-049</td><td>Basal medium used for neuronal cultures</td></tr><tr><td>PDMS microfluidic devices&amp;nbsp;</td><td>not applicable</td><td>not applicable</td><td>Composed of two cell seeding compartments interconnected by 20 microgrooves with 450 &amp;mu;m length &amp;times; 10 &amp;mu;m height &amp;times; 14 &amp;mu;m width dimensions and separated by 86 &amp;micro;m (total interspace of 100 &amp;mu;m).</td></tr><tr><td>Penicillin-streptomycin (P/S) solution (100X)</td><td>Biowest</td><td>L0022-100</td><td></td></tr><tr><td>PES syringe filter unit (&amp;Oslash; 30 mm), 0.22 &amp;micro;m&amp;nbsp;</td><td>Frilabo</td><td>1730012</td><td></td></tr><tr><td>Polypropylene conical tubes, 15 ml</td><td>Thermo Fisher Scientific</td><td>07-200-886</td><td></td></tr><tr><td>Polypropylene conical tubes, 50 ml</td><td>Thermo Fisher Scientific</td><td>05-539-13</td><td></td></tr><tr><td>Polystyrene disposabel serological pipets, 10 ml&amp;nbsp;</td><td>Thermo Fisher Scientific</td><td>1367811D</td><td></td></tr><tr><td>Polystyrene disposabel serological pipets, 5 ml&amp;nbsp;</td><td>Thermo Fisher Scientific</td><td>1367811D</td><td></td></tr><tr><td>Standard Regenerated cellulose membrane (2 kDa)&amp;nbsp;&amp;nbsp;</td><td>Spectrum labs</td><td>132107</td><td></td></tr><tr><td>Standard surgical scissor</td><td>Fine Science Tools</td><td>91401-14</td><td></td></tr><tr><td>Substrate-integrated planar MEAs (256 microelectrodes)</td><td>Multi Channel Systems</td><td>256MEA100/30iR-ITO</td><td>252 titanium nitride (TiN) recording electrodes and 4 internal reference electrodes organized in a 16 by 16 square grid. Each recording electrode is 30 &amp;micro;m in diameter and interspaced by 100 &amp;micro;m.</td></tr><tr><td>Syringe luer-lock tip, 10 ml&amp;nbsp;</td><td>Terumo Europe</td><td>5100-X00V0</td><td></td></tr><tr><td>Syringe luer-lock tip, 50 ml&amp;nbsp;</td><td>Terumo Europe</td><td>8300006682</td><td></td></tr><tr><td>Terg-A-Zyme</td><td>Sigma-Aldrich</td><td>Z273287&amp;nbsp;</td><td>Enzyme-active powdered detergent used for MEAs cleaning</td></tr><tr><td>Tissue culture plates, 35 mm&amp;nbsp;</td><td>StemCell Technologies</td><td>27150</td><td></td></tr><tr><td>Tissue culture plates, 90 mm</td><td>Frilabo</td><td>900095</td><td></td></tr><tr><td>Trypan Blue solution (0.4%)&amp;nbsp;</td><td>Sigma-Aldrich</td><td>T8154</td><td></td></tr><tr><td>Trypsin (1:250)</td><td>Thermo Fisher Scientific</td><td>27250018</td><td></td></tr><tr><td>Vinyl tape 471</td><td>3M</td><td>B40071909</td><td></td></tr></tbody></table>

interfacing microfluidics with microelectrode arrays for studying neuronal communication and axonal signal propagation

Microelectrode arrays (MEAs) are widely used to study neuronal function in vitro. These devices allow concurrent non-invasive recording/stimulation of electrophysiological activity for long periods. However, the property of sensing signals from all sources around every microelectrode can become unfavorable when trying to understand communication and signal propagation in neuronal circuits. In a neuronal network, several neurons can be simultaneously activated and can generate overlapping action potentials, making it difficult to discriminate and track signal propagation. Considering this limitation, we have established an in vitro setup focused on assessing electrophysiological communication, which is able to isolate and amplify axonal signals with high spatial and temporal resolution. By interfacing microfluidic devices and MEAs, we are able to compartmentalize neuronal cultures with a well-controlled alignment of the axons and microelectrodes. This setup allows recordings of spike propagation with a high signal-to-noise ratio over the course of several weeks. Combined with specialized data analysis algorithms, it provides detailed quantification of several communication related properties such as propagation velocity, conduction failure, firing rate, anterograde spikes, and coding mechanisms.
This protocol demonstrates how to create a compartmentalized neuronal culture setup over substrate-integrated MEAs, how to culture neurons in this setup, and how to successfully record, analyze and interpret the results from such experiments. Here, we show how the established setup simplifies the understanding of neuronal communication and axonal signal propagation. These platforms pave the way for new in vitro models with engineered and controllable neuronal network topographies. They can be used in the context of homogeneous neuronal cultures, or with co-culture configurations where, for example, communication between sensory neurons and other cell types is monitored and assessed. This setup provides very interesting conditions to study, for example, neurodevelopment, neuronal circuits, information coding, neurodegeneration and neuroregeneration approaches.

Using the protocol described here, E-18 rat cortical neurons seeded on &#181;EF are able to develop and remain healthy in these culture conditions for over a month. As soon as 3 to 5 days in culture, cortical neurons grow their axons through microgrooves towards the axonal compartment of &#181;EF (Figure 1). After 15 days in culture, cortical neurons cultured on &#181;EF are expected to exhibit steady levels of activity and propagation of action potentials along the microgrooves is expected. Older cultures (&#62;14 DIV) tend to be dominated by bursting activity as in conventional MEA recordings18,19.
Recordings and data analysis
Raw data analysis with &#181;SpikeHunter (Figure 2) permitted the detection and extraction of propagating spike waves along sets of 4 microelectrodes (within microgrooves). Figure 3 displays one of these events. &#181;SpikeHunter allowed for the estimation of propagation velocities, based on the cross-correlations between the voltage waveforms of selected pairs of&#160;microelectrodes (providing a time delay) and the associated known inter-electrode distance.
The extracted data was further analyzed using custom designed code in MATLAB. A representative raster plot of the propagating spike waves along 11 (out of 16) microgrooves is shown in Figure 4a. The instantaneous firing rate for a high- and a low-activity microgroove is shown in Figure 4b.
&#181;EF recordings exhibit varying levels of activity per microelectrode. Frequently, several microelectrodes in the somal compartment are &#34;silent&#34;. However, most microelectrodes within microgrooves tend to be active (Figure 5a). It is well described that the microgrooves function as signal amplifiers8. The amplitude of a recorded signal depends not only on the size of the source currents, but also on the resistivity of the surrounding media. The resistance along a microgroove is particularly high, which greatly increases the amplitude of the measured signals in comparison to those of the microelectrodes in the somal compartment (Figure 5b).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58878/58878fig1v2.jpg" /> 
Figure 1. Embryonic rat cortical neurons cultured on &#181;EF. (a) Photograph of &#181;EF. Scale bar: 1 cm. (b) Representative image of cortical neurons cultured in the &#181;EF for 5 days, showing several axons crossing the microgrooves and reaching the axonal compartment (arrows). Scale bar: 100 &#181;m. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58878/58878fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58878/58878fig2v2.jpg" /> 
Figure 2.&#160;Screen capture of the &#181;SpikeHunter main graphical user interface. The user can load the data recorded with &#181;EF, identify propagating spike waves, determine their direction (anterograde or retrograde) and estimate propagation velocities. A kymograph tool allows the user to manually estimate the propagation velocity based on a line drawn on the kymograph. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58878/58878fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58878/58878fig3v2.jpg" /> 
Figure 3.&#160;Anterograde propagating spike wave sensed by 4 microelectrodes (E8-E11) within a microgroove. Each trace represents one microelectrode raw recording for 3 ms. After analysis with &#181;SpikeHunter, the cross-correlation between the farthest microelectrodes (E8 and E11) permitted the calculation of a propagation velocity of 0.52 m/s. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58878/58878fig3v2large.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/58878/58878fig4v2.jpg" /> 
Figure 4.&#160;Information barrage through the microgrooves.(a) Representative raster plot of 2 minutes of spontaneous activity recorded along 11 microgrooves. Each dot represents a propagating spike wave (unit) sensed by 4 microelectrodes and identified through the analysis with &#181;SpikeHunter. A high- and a low-activity microgroove are highlighted in yellow and red, respectively. (b) Evolution of the instantaneous firing rate (as in rate-coding) for the two highlighted microgrooves along 10 minutes. Only propagating units were considered for the calculation of the firing rate. Note the activity synchronization, despite the different firing rate levels. The instantaneous firing rate was calculated convolving the spike events with a Gaussian kernel with a standard deviation of 100 ms. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58878/58878fig4v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/58878/58878fig5v2.jpg" /> 
Figure 5. Quality of extracellular recordings in &#181;EF. (a) Time window (1 second) of a &#181;EF recording (rat cortical neurons at 15 days in vitro) with a sampling rate of 20 kHz and a high-pass filter at 300 Hz. Electrode rows 1-7 correspond to the somal compartment and rows 8-11 to the microgrooves. (b) Box and whisker plot of the full range of spike amplitudes extracted (spikes detected using a threshold method, with negative phase only, set at 6x standard deviation) from the specified rows (total recording time of 10 minutes). The spikes&#39; amplitudes are significantly larger in the microelectrodes within microgrooves (rows 8-11; 16 microgrooves), when compared to the microelectrodes of the somal compartment (rows 1-7; 16 active microelectrodes). One-way ANOVA, Dunn&#39;s multiple comparison tests, ****p &#60;0.0001.&#160;<a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58878/58878fig5v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Interfacing Microfluidics with Microelectrode Arrays for Studying Neuronal Communication and Axonal Signal Propagation at JoVE.com

Interfacing Microfluidics with Microelectrode Arrays for Studying Neuronal Communication and Axonal Signal Propagation

This protocol aims to demonstrate how to combine in vitro microelectrode arrays with microfluidic devices for studying action potential transmission in neuronal cultures. Data analysis, namely the detection and characterization of propagating action potentials, is performed using a new advanced, yet user-friendly and freely available, computational tool.

Microelectrode arrays (MEAs) are widely used to study neuronal function in vitro. These devices allow concurrent non-invasive recording/stimulation of ...

interfacing-microfluidics-with-microelectrode-arrays-for-studying

Universidad San Sebastian

Research

JoVE Journal

Bioengineering

8.0K Views.  Universidade do Porto. This protocol aims to demonstrate how to combine in vitro microelectrode arrays with microfluidic devices for studying action potential transmission in neuronal cultures. Data analysis, namely the detection and characterization of propagating action potentials, is performed using a new advanced, yet user-friendly and freely available, computational tool.

Video: Interfacing Microfluidics with Microelectrode Arrays for Studying Neuronal Communication and Axonal Signal Propagation

Applying Microfluidics to Electrophysiology

Microfluidics can be integrated with standard electrophysiology techniques to allow new experimental modalities.  Specifically, the motivation for the microfluidic brain slice device is discussed including how the device docks to standard perfusion chambers and the technique of passive pumping which is used to deliver boluses of neuromodulators to the brain slice.  By simplifying the device design, we are able to achieve a practical solution to the current unmet electrophysiology need of applying multiple neuromodulators across multiple regions of the brain slice.  This is achieved by substituting the standard coverglass substrate of the perfusion chamber with a thin microfluidic device bonded to the coverglass substrate.  This was then attached to the perfusion chamber and small holes connect the open-well of the perfusion chamber to the microfluidic channels buried within the microfluidic substrate.  These microfluidic channels are interfaced with ports drilled into the edge of the perfusion chamber to access and deliver stimulants.  This project represents how the field of microfluidics is transitioning away from proof-of concept device demonstrations and into practical solutions for unmet experimental and clinical needs.

Microfluidics can be integrated with standard electrophysiology techniques to allow new experimental modalities.  Specifically, the motivation for the ...

Biology

BioMEMS: Forging New Collaborations Between Biologists and Engineers

This video describes the fabrication and use of a microfluidic device to culture central nervous system (CNS) neurons.  This device is compatible with live-cell optical microscopy (DIC and phase contrast), as well as confocal and two photon microscopy approaches.  This method uses precision-molded polymer parts to create miniature multi-compartment cell culture with fluidic isolation. The compartments are made of tiny channels with dimensions that are large enough to culture neurons in well-controlled fluidic microenvironments.  Neurons can be cultured for 2-3 weeks within the device, after which they can be fixed and stained for immunocytochemistry. Axonal and somal compartments can be maintained fluidically isolated from each other by using a small hydrostatic pressure difference; this feature can be used to localize soluble insults to one compartment for up to 20 h after each medium change. Fluidic isolation enables collection of pure axonal fraction and biochemical analysis by PCR. The microfluidic device provides a highly adaptable platform for neuroscience research and may find applications in modeling CNS injury and neurodegeneration.

This video describes the fabrication and use of a microfluidic device to culture central nervous system (CNS) neurons.  This device is compatible with ...

Imaging Analysis of Neuron to Glia Interaction in Microfluidic Culture Platform (MCP)-based Neuronal Axon and Glia Co-culture System

Proper neuron to glia interaction is critical to physiological function of the central nervous system (CNS). This bidirectional communication is sophisticatedly mediated by specific signaling pathways between neuron and glia1,2 . Identification and characterization of these signaling pathways is essential to the understanding of how neuron to glia interaction shapes CNS physiology. Previously, neuron and glia mixed cultures have been widely utilized for testing and characterizing signaling pathways between neuron and glia. What we have learned from these preparations and other in vivo tools, however, has suggested that mutual signaling between neuron and glia often occurred in specific compartments within neurons (i.e., axon, dendrite, or soma)3. This makes it important to develop a new culture system that allows separation of neuronal compartments and specifically examines the interaction between glia and neuronal axons/dendrites. In addition, the conventional mixed culture system is not capable of differentiating the soluble factors and direct membrane contact signals between neuron and glia. Furthermore, the large quantity of neurons and glial cells in the conventional co-culture system lacks the resolution necessary to observe the interaction between a single axon and a glial cell.
	In this study, we describe a novel axon and glia co-culture system with the use of a microfluidic culture platform (MCP). In this co-culture system, neurons and glial cells are cultured in two separate chambers that are connected through multiple central channels. In this microfluidic culture platform, only neuronal processes (especially axons) can enter the glial side through the central channels. In combination with powerful fluorescent protein labeling, this system allows direct examination of signaling pathways between axonal/dendritic and glial interactions, such as axon-mediated transcriptional regulation in glia, glia-mediated receptor trafficking in neuronal terminals, and glia-mediated axon growth. The narrow diameter of the chamber also significantly prohibits the flow of the neuron-enriched medium into the glial chamber, facilitating probing of the direct membrane-protein interaction between axons/dendrites and glial surfaces.

Imaging Analysis of Neuron to Glia Interaction in Microfluidic Culture Platform MCP-based Neuronal Axon and Glia Co-culture System

This study describes the procedures of setting up a novel neuronal axon and (astro)glia co-culture platform. In this co-culture system, manipulation of direct interaction between a single axon (and single glial cell) becomes feasible, allowing mechanistic analysis of the mutual neuron to glial signaling.

Proper  neuron to glia interaction is critical to physiological function of the central  nervous system (CNS). This bidirectional communication is ...

Neuroscience

Co-culture of Glutamatergic Neurons and Pediatric High-Grade Glioma Cells Into Microfluidic Devices to Assess Electrical Interactions

Pediatric high-grade gliomas (pHGG) represent childhood and adolescent brain cancers that carry a rapid dismal prognosis. Since there is a need to overcome the resistance to current treatments and find a new way of cure, modeling the disease as close as possible in an in vitro setting to test new drugs and therapeutic procedures is highly demanding. Studying their fundamental pathobiological processes, including glutamatergic neuron hyperexcitability, will be a real advance in understanding interactions between the environmental brain and pHGG cells. Therefore, to recreate neurons/pHGG cell interactions, this work shows the development of a functional in vitro model co-culturing human-induced Pluripotent Stem (hiPS)-derived cortical glutamatergic neurons pHGG cells into compartmentalized microfluidic devices and a process to record their electrophysiological modifications. The first step was to differentiate and characterize human glutamatergic neurons. Secondly, the cells were cultured in microfluidic devices with pHGG derived cell lines. Brain microenvironment and neuronal activity were then included in this model to analyze the electrical impact of pHGG cells on these micro-environmental neurons. Electrophysiological recordings are coupled using multielectrode arrays (MEA) to these microfluidic devices to mimic physiological conditions and to record the electrical activity of the entire neural network. A significant increase in neuron excitability was underlined in the presence of tumor cells.

Recent works uncover the neuronal impact on high-grade pediatric glioma (pHGG) cells and their reciprocal interactions. The present work shows the development of an in vitro model co-culturing pHGG cells and glutamatergic neurons and recorded their electrophysiological interactions to mimic those interactivities.

Pediatric high-grade gliomas (pHGG) represent childhood and adolescent brain cancers that carry a rapid dismal prognosis. Since there is a need to ...

Cancer Research

Preparation of Neuronal Co-cultures with Single Cell Precision

Microfluidic embodiments of the Campenot chamber have attracted great interest from the neuroscience community. These interconnected co-culture platforms can be used to investigate a variety of questions, spanning developmental and functional neurobiology to infection and disease propagation. However, conventional systems require significant cellular inputs (many thousands per compartment), inadequate for studying low abundance cells, such as primary dopaminergic substantia nigra, spiral ganglia, and Drosophilia melanogaster neurons, and impractical for high throughput experimentation. The dense cultures are also highly locally entangled, with few outgrowths (&#60;10%) interconnecting the two cultures. In this paper straightforward microfluidic and patterning protocols are described which address these challenges: (i) a microfluidic single neuron arraying method, and (ii) a water masking method for plasma patterning biomaterial coatings to register neurons and promote outgrowth between compartments. Minimalistic neuronal co-cultures were prepared with high-level (&#62;85%) intercompartment connectivity and can be used for high throughput neurobiology experiments with single cell precision.

Protocols for single neuron microfluidic arraying and water masking for the in-chip plasma patterning of biomaterial coatings are described. Highly interconnected co-cultures can be prepared using minimal cell inputs.

Microfluidic embodiments of the Campenot chamber have attracted great interest from the neuroscience community. These interconnected co-culture platforms ...

Design, Surface Treatment, Cellular Plating, and Culturing of Modular Neuronal Networks Composed of Functionally Inter-connected Circuits

The brain operates through the coordinated activation and the dynamic communication of neuronal assemblies. A major open question is how a vast repertoire of dynamical motifs, which underlie most diverse brain functions, can emerge out of a fixed topological and modular organization of brain circuits. Compared to in vivo studies of neuronal circuits which present intrinsic experimental difficulties, in vitro preparations offer a much larger possibility to manipulate and probe the structural, dynamical and chemical properties of experimental neuronal systems. This work describes an in vitro experimental methodology which allows growing of modular networks composed by spatially distinct, functionally interconnected neuronal assemblies. The protocol allows controlling the two-dimensional (2D) architecture of the neuronal network at different levels of topological complexity.
A desired network patterning can be achieved both on regular cover slips and substrate embedded micro electrode arrays. Micromachined structures are embossed on a silicon wafer and used to create biocompatible polymeric stencils, which incorporate the negative features of the desired network architecture. The stencils are placed on the culturing substrates during the surface coating procedure with a molecular layer for promoting cellular adhesion. After removal of the stencils, neurons are plated and they spontaneously redirected to the coated areas. By decreasing the inter-compartment distance, it is possible to obtain either isolated or interconnected neuronal circuits. To promote cell survival, cells are co-cultured with a supporting neuronal network which is located at the periphery of the culture dish. Electrophysiological and optical recordings of the activity of modular networks obtained respectively by using substrate embedded micro electrode arrays and calcium imaging are presented. While each module shows spontaneous global synchronizations, the occurrence of inter-module synchronization is regulated by the density of connection among the circuits.

This manuscript describes a protocol to grow in vitro modular networks consisting of spatially confined, functionally inter-connected neuronal circuits. A polymeric mask is used to pattern a protein layer to promote cellular adhesion over the culturing substrate. Plated neurons grow on coated areas establishing spontaneous connections and exhibiting electrophysiological activity.

The brain operates through the coordinated activation and the dynamic communication of neuronal assemblies. A major open question is how a vast repertoire ...

In Vitro Multiparametric Cellular Analysis by Micro Organic Charge-modulated Field-effect Transistor Arrays

Modern electrophysiology has been constantly fueled by the parallel development of increasingly sophisticated tools and materials. In turn, discoveries in this field have driven technological progress in a back-and-forth process that ultimately determined the impressive achievements of the past 50 years. However, the most employed devices used for cellular interfacing (namely, the microelectrode arrays and microelectronic devices based on transistors) still present several limitations such as high cost, the rigidity of the materials, and the presence of an external reference electrode. To partially overcome these issues, there have been developments in a new scientific field called organic bioelectronics, resulting in advantages such as lower cost, more convenient materials, and innovative fabrication techniques.
Several interesting new organic devices have been proposed during the past decade to conveniently interface with cell cultures.&#160;This paper presents the protocol for the fabrication of devices for cellular interfacing based on the organic charge-modulated field-effect transistor (OCMFET). These devices, called micro OCMFET arrays (MOAs), combine the advantages of organic electronics and the peculiar features of the OCMFET to prepare transparent, flexible, and reference-less tools with which it is possible to monitor both the electrical and the metabolic activities of cardiomyocytes and neurons in vitro, thus allowing a multiparametric evaluation of electrogenic cell models.

In Vitro Multiparametric Cellular Analysis by Micro Organic Charge-modulated Field-effect Transistor Arrays

Here, we present the fabrication protocol of an organic charge-modulated field-effect transistor (OCMFET)-based device for in vitro cellular interfacing. The device, called a micro OCMFET array, is a flexible, low-cost, and reference-less device, which will enable the monitoring of the electrical and metabolic activities of electroactive cell cultures.

Modern electrophysiology has been constantly fueled by the parallel development of increasingly sophisticated tools and materials. In turn, discoveries in ...

Microdialysis of Excitatory Amino Acids During EEG Recordings in Freely Moving Rats

Microdialysis is a well-established neuroscience technique that correlates the changes of neurologically active substances diffusing into the brain interstitial space with the behavior and/or with the specific outcome of a pathology (e.g., seizures for epilepsy). When studying epilepsy, the microdialysis technique is often combined with short-term or even long-term video-electroencephalography (EEG) monitoring to assess spontaneous seizure frequency, severity, progression and clustering. The combined microdialysis-EEG is based on the use of several methods and instruments. Here, we performed in vivo microdialysis and continuous video-EEG recording to monitor glutamate and aspartate outflow over time, in different phases of the natural history of epilepsy in a rat model. This combined approach allows the pairing of changes in the neurotransmitter release with specific stages of the disease development and progression. The amino acid concentration in the dialysate was determined by liquid chromatography. Here, we describe the methods and outline the principal precautionary measures one should take during in vivo microdialysis-EEG, with particular attention to the stereotaxic surgery, basal and high potassium stimulation during microdialysis, depth electrode EEG recording and high-performance liquid chromatography analysis of aspartate and glutamate in the dialysate. This approach may be adapted to test a variety of drug or disease induced changes of the physiological concentrations of aspartate and glutamate in the brain. Depending on the availability of an appropriate analytical assay, it may be further used to test different soluble molecules when employing EEG recording at the same time.

Here, we describe a method for in vivo microdialysis to analyze aspartate and glutamate release in the ventral hippocampus of epileptic and non-epileptic rats, in combination with EEG recordings. Extracellular concentrations of aspartate and glutamate may be correlated with the different phases of the disease.

Microdialysis is a well-established neuroscience technique that correlates the changes of neurologically active substances diffusing into the brain ...

Microinjectrode System for Combined Drug Infusion and Electrophysiology

This microinjectrode system is designed for drug infusion, electrophysiology, and delivery and retrieval of experimental probes, such as microelectrodes and nanosensors, optimized for repeated use in awake, behaving animals. The microinjectrode system can be configured for multiple purposes: (1) simple arrangement of the cannula for placement of an experimental probe that would otherwise be too fragile to penetrate the dura mater, (2) microfluidic infusion of a drug, either independently or coupled to a cannula containing an experimental probe (i.e., microelectrode, nanosensor). In this protocol we explain the step by step construction of the microinjectrode, its coupling to microfluidic components, and the protocol for use of the system in vivo. The microfluidic components of this system allow for delivery of volumes on the nanoliter scale, with minimal penetration damage. Drug infusion can be performed independently or simultaneously with experimental probes such as microelectrodes or nanosensors in an awake, behaving animal. Applications of this system range from measuring the effects of a drug on cortical electrical activity and behavior, to understanding the function of a specific region of cortex in the context of behavioral performance based on probe or nanosensor measurements. To demonstrate some of the capabilities of this system, we present an example of muscimol infusion for reversible inactivation of the frontal eye field (FEF) in rhesus macaque during a working memory task.

We present a microinjectrode system designed for electrophysiology and assisted delivery of experimental probes (i.e., nanosensors, microelectrodes), with optional drug infusion. Widely available microfluidic components are coupled to a cannula containing the probe. A step-by-step protocol for microinjectrode construction is included, with results during muscimol infusion in macaque cortex.

This microinjectrode system is designed for drug infusion, electrophysiology, and delivery and retrieval of experimental probes, such as microelectrodes ...

Microfluidics-Assisted Selective Depolarization of Axonal Mitochondria

Mitochondria are the primary suppliers of ATP (adenosine triphosphate) in neurons. Mitochondrial dysfunction is a common phenotype in many neurodegenerative diseases. Given some axons' elaborate architecture and extreme length, it is not surprising that mitochondria in axons can experience different environments compared to their cell body counterparts. Interestingly, dysfunction of axonal mitochondria often precedes effects on the cell body. To model axonal mitochondrial dysfunction in vitro, microfluidic devices allow treatment of axonal mitochondria without affecting the somal mitochondria. The fluidic pressure gradient in these chambers prevents diffusion of molecules against the gradient, thus allowing for analysis of mitochondrial properties in response to local pharmacological challenges within axons. The current protocol describes the seeding of dissociated hippocampal neurons in microfluidic devices, staining with a membrane-potential sensitive dye, treatment with a mitochondrial toxin, and the subsequent microscopic analysis. This versatile method to study axonal biology can be applied to many pharmacological perturbations and imaging readouts, and is suitable for several neuronal subtypes.

The present protocol describes the seeding and staining of neuronal mitochondria in microfluidic chambers. The fluidic pressure gradient in these chambers allows for the selective treatment of mitochondria in axons to analyze their properties in response to pharmacological challenges without affecting the cell body compartment.

Interfacing Microfluidics with Microelectrode Arrays for Studying Neuronal Communication and Axonal Signal Propagation

Summary

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Microdialysis of Excitatory Amino Acids During EEG Recordings in Freely Moving Rats

Microinjectrode System for Combined Drug Infusion and Electrophysiology

Microfluidics-Assisted Selective Depolarization of Axonal Mitochondria