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Podsumowanie

This protocol aims to describe a new methodology to measure intrinsic cardiac firing rate using microelectrode array recording of the whole sinoatrial node tissue to identify pacemaking defects in mice. Pharmacological agents can also be introduced in this method to study their effects on intrinsic pacemaking.

Abstract

The sinoatrial node (SAN), located in the right atrium, contains the pacemaker cells of the heart, and dysfunction of this region can cause tachycardia or bradycardia. Reliable identification of cardiac pacemaking defects requires the measurement of intrinsic heart rates by largely preventing the influence of the autonomic nervous system, which can mask rate deficits. Traditional methods to analyze intrinsic cardiac pacemaker function include drug-induced autonomic blockade to measure in vivo heart rates, isolated heart recordings to measure intrinsic heart rates, and sinoatrial strip or single-cell patch-clamp recordings of sinoatrial pacemaker cells to measure spontaneous action potential firing rates. However, these more traditional techniques can be technically challenging and difficult to perform. Here, we present a new methodology to measure intrinsic cardiac firing rate by performing microelectrode array (MEA) recordings of whole-mount sinoatrial node preparations from mice. MEAs are composed of multiple microelectrodes arranged in a grid-like pattern for recording in vitro extracellular field potentials. The method described herein has the combined advantage of being relatively faster, simpler, and more precise than previous approaches for recording intrinsic heart rates, while also allowing easy pharmacological interrogation.

Introduction

The heart is a complex organ governed by both cardiac-intrinsic and extrinsic influences such as those that originate in the brain. The sinoatrial node (SAN) is a defined region in the heart that houses the pacemaker cells (also referred to as sinoatrial cells, or SA cells) responsible for the initiation and perpetuation of the mammalian heartbeat^1,2. The intrinsic heart rate is the rate driven by the pacemaker cells without influence by other cardiac or neuro-humoral influences, but traditional measures of heart rate in humans and live animals, such as electrocardiograms, reflect both the pacemaker and neural influences on the heart. The most notable neural influence on SA cells is from the autonomic nervous system, which constantly modulates firing patterns to meet the physiological requirements of the body³. Supporting this idea, both sympathetic and parasympathetic projections can be found near the SAN⁴. The intrinsic cardiac nervous system (ICNS) is another important neural influence where ganglionated plexi, specifically in the right atria, innervate and regulate the activity of the SAN^5,6.

Understanding pacemaking deficits is clinically important, as dysfunction can underlie many cardiac disorders, as well as contribute to the risk of other complications. Sick sinus syndrome (SSS) is a category of diseases characterized by dysfunction of the sinoatrial node which impedes proper pacemaking^7,8. SSS can present with sinus bradycardia, sinus pauses, sinus arrest, sinoatrial exit block, and alternating bradyarrythmias and tachyarrhythmias⁹ and can lead to complications including increased risk of embolic stroke and sudden death^8,10. Those with Brugada syndrome, a cardiac disorder marked by ventricular fibrillation with an increased risk of sudden cardiac death, are at greater risk for arrhythmogenic events if they also have comorbid SAN dysfunction^11,12. Sinoatrial dysfunction may also have physiological consequences beyond the heart. For example, SSS has been observed to trigger seizures in a patient due to cerebral hypoperfusion¹³.

To identify sinoatrial pacemaking deficits, intrinsic heart rates need to be determined by measuring the activity of the SAN without the influence of the autonomic nervous system or humoral factors. Clinically, this can be approximated by pharmacological autonomic blockade¹⁴, but this same technique can also be applied in mammalian models to study intrinsic cardiac function^15,16. While this approach blocks a large portion of contributing neural influences and allows for in vivo cardiac examination, it does not completely eliminate all extrinsic influences on the heart. Another research technique used to study intrinsic cardiac function in animal models is isolated heart recordings using Langendorff-perfused hearts, which typically involve measurements using electrograms, pacing, or epicardial multielectrode arrays^17,18,19,20. While this technique is more specific to cardiac function since it involves removing the heart from the body, the measurements may still be influenced by mechano-electric autoregulatory mechanisms that could influence intrinsic heart rate measurements²¹. The isolated heart recordings may also still be influenced by autonomic regulation through the ICNS^5,6,22,23. Furthermore, maintaining a physiologically relevant temperature of the heart, which is critical for cardiac function measurements, can be difficult in isolated heart approaches²⁰. A more direct method to study SAN function is to specifically isolate SAN tissue and measure its activity. This can be accomplished through SAN strips (isolated SAN tissue) or isolated SAN pacemaker cells^24,25. Both require a high degree of technical training, as the SAN is a very small and highly defined region, and cell isolation poses an even greater challenge as dissociation can damage the overall health of the cell if not performed correctly. Furthermore, these techniques require expert electrophysiological skills in order to successfully record from the tissue or cells using individual recording microelectrodes.

In this protocol, we describe a technique to record the SAN in vitro by using a microelectrode array (MEA) to obtain intrinsic heart rate measurements. This approach has the advantage of making highly specific electrophysiological recordings accessible to researchers lacking intensive electrophysiological skillsets. MEAs have previously been used to study cardiomyocyte function in primary cardiomyocyte cultures^{26,27,28,29,30,31,32}, cardiac sheets^{33,34,35,36,37,38,39}, and tissue whole mounts^{40,41,42,43,44,45,46,47}. Previous work has also been done to examine field potentials in SAN tissue^41,42. Here, we provide a methodology to use the MEA to record and analyze murine intrinsic SAN firing rates. We also describe how this technique can be used to test pharmacological effects of drugs on SAN intrinsic firing rates by providing a sample experiment showing the effects of 4-aminopyridine (4-AP), a voltage-gated K⁺ channel blocker. Using defined anatomical landmarks, we can accurately record the SAN without having to perform the extensive tissue dissections or cell isolations required in other methods. While the MEA can be cost-prohibitive, the recordings provide highly specific and reliable measures of pacemaking that can be used in a vast array of clinical and physiological research applications.

Protokół

All experimental procedures described here have been carried out in accordance with the guidelines of the National Institutes of Health (NIH), as approved by the Institutional Animal Care and Use Committee (IACUC) at Southern Methodist University. 1. Coating the multielectrode array (MEA) for recording Make 25 mM borate buffer. Dissolve 0.953 g of Na2B4O7·10 H2O in 80 mL of distilled water. Adjust the pH to 8.4 with HCl and then add distilled water to a final volume of 100 mL. Make a 0.1% stock solution of polyethyleneimine (PEI). Add 100 µL of 50% (w/v) PEI to 4.9 mL of distilled water to make a 1% PEI solution. Dilute the 1% PEI solution to 0.1% in borate buffer by adding 1 mL of the 1% PEI solution to 9 mL of the 25 mM borate buffer. Pipette ~1 mL of the 0.1% PEI solution into the microelectrode array (MEA) dish so that the electrodes are completely covered (Figure 1A and 1B). NOTE: The microelectrodes of the MEA are typically composed of platinum black or carbon nanotube and insulated with polyimide (or acrylic); both materials are hydrophobic. By coating the MEA with a cationic polymer such as PEI, the hydrophobic MEA surface is made more hydrophilic, allowing tissue samples to make better contact with the MEA surface (Figure 1A1). Cover the MEA dish with thermoplastic film to reduce evaporation and leave the MEA overnight at room temperature (Figure 1C). Aspirate the PEI solution from the MEA dish using a pipette, being careful not to touch the electrode grid which can damage the electrodes, and then rinse ≥4 times with distilled water (Figure 1D). Store the PEI-coated MEA under 1-2 mL of ultrapure water and sealed with thermoplastic film at 4 °C until needed. Alternatively, store the coated MEA by submerging it in a beaker filled with ultrapure water (Figure 1E). NOTE: The PEI coating process only needs to be performed once for the MEA before its first use, and after each recording session, the MEA should be stored submerged in ultrapure water. 2. Preparing complete Tyrode's solution for tissue dissection Make 1,000 mL of complete Tyrode's solution for dissection; first, add 8.1816 g of NaCl to 800 mL of ultrapure water. Add the following amount of chemicals to the solution: 0.4025 g of KCl; 0.1633 g of KH2PO4; 1.1915 g of HEPES; 0.9999 g of glucose; 0.0952 g of MgCl2; 0.2646 g of CaCl2·2H2O. Adjust the pH to 7.4 with NaOH and then add ultrapure water until the total volume is 1,000 mL. NOTE: The final composition of the Complete Tyrode's solution will be the following (in mM): 140 NaCl, 5.4 KCl, 1.2 KH2PO4, 5 HEPES, 5.55 glucose, 1 MgCl2, 1.8 CaCl2. 3. Preparing oxygenated Tyrode's solution for recording Make 500 mL of Tyrode's solution; add 4.003 g of NaCl to 400 mL of ultrapure water. Add the following amounts of chemicals to the solution: 0.651 g of NaHCO3; 0.042 g of NaH2PO4; 0.132 g of CaCl2·2H2O; 0.149 g of KCl; 0.0476 g of MgCl2; 0.999 g of glucose. Adjust the pH to 7.4 with HCl and then add ultrapure water until the total volume is 500 mL. Oxygenate the solution with carbogen for at least 30 min at room temperature before starting the recording. ​NOTE: The final composition of the Tyrode's solution will be the following (in mM): 137 NaCl, 15.5 NaHCO3, 0.7 NaH2PO4, 1.8 CaCl2, 4 KCl, 1 MgCl2, 11.1 glucose. This Tyrode's solution has a slightly different composition from the Complete Tyrode's solution used for dissection. 4. Preparing 4-aminopyridine (4-AP) solution for pharmacological modulation Make a 1 mM working solution of 4-AP; add 18.82 mg of 4-AP to 200 mL of the Tyrode's solution from step 3. Oxygenate the 4-AP solution for at least 30 min before the experiment. 5. Preparing the Petri dish for dissection Mix silicone elastomer components in a 10:1 ratio (by weight) of the base to curing agent. Pour ~15 mL of silicone elastomer mixture into a 60 mm diameter Petri dish. Allow elastomer to cure at room temperature for 48 h before use. ​NOTE: The siliconized Petri dish can be reused for future dissections. 6. Dissecting the sinoatrial node (SAN) Prepare heparinized Complete Tyrode's solution for SAN dissection. Add 400 µL of heparin (1,000 USP/mL) to 40 mL of Complete Tyrode's solution and warm in a 37 °C water bath. Inject the mouse intraperitoneally with 200-300 μL of heparin (1000 USP/mL) and allow the animal to sit for 10 min. Euthanize the heparinized mouse by isoflurane overdose. Place the mouse in a small glass chamber that contains isoflurane vapors generated by adding 200-300 μL of liquid isoflurane to a filter paper inside a perforated plastic tube. NOTE: Because isoflurane can cause skin irritation and can also be absorbed through the skin, the liquid should not contact the mouse directly. Therefore, the isoflurane-soaked wipe is placed in a perforated tube for administration. Verify death by cessation of movement and breathing effort and by the absence of a toe pinch reflex. Death usually takes about 1-2 min following placement into the chamber. NOTE: Death is usually accompanied by urination. Place the mouse in a supine position on a dissection board with paws outstretched and fix the paws to the board using 1 inch long, 23-gauge syringe needles. Then remove the fur in the vicinity of the bottom of the rib cage by using surgical scissors and cutting the fur at the roots. NOTE: For a dissection board, polystyrene cooler lids can be used. While holding the skin with a hemostat, use surgical scissors to make a transverse incision in the skin just beneath the bottom of the rib cage from about the left costal arch to the right costal arch (Figure 3A). Cut open the peritoneum with surgical scissors and carefully separate the liver from the diaphragm, being careful not to nick the liver, which will cause excessive bleeding (Figure 3B). Incise the diaphragm along the thorax to expose the thoracic cavity (Figure 3C-D). Using surgical scissors, cut the lateral walls of the rib cage from the edges of the costal arches up to the clavicles to expose the heart, being careful to avoid damaging the heart (Figure 3D). Then use a 23-gauge syringe needle to pin the rib cage over the shoulder, holding it in place and out of the way of the surgical field. Use a transfer pipette to drip warm (37 °C) heparinized Complete Tyrode's solution onto the heart to keep it moist. NOTE: Do not allow the heart to dry out. Remove the lungs by holding them with extra fine Graefe forceps and severing the trachea with surgical scissors (Figure 3E). Hold the apex of the heart with extra fine Graefe forceps and remove it by cutting the aorta and venae cavae with surgical scissors. Transfer the heart to a Petri dish containing cured silicone elastomer (Figure 4A) and use a transfer pipette to bathe the heart with 2-3 mL of warm (37 °C) heparinized Complete Tyrode's solution. NOTE: Be careful not to damage the delicate posterior wall of the right atria, which contains the SAN, and the connected right atrial veins. Bathing the heart with Complete Tyrode's solution keeps the heart from drying out but do not fully submerge the heart in solution as it will impair visibility during dissection. Orient the heart with the right atrium on the experimenter's right and the left atrium on the experimenter's left. NOTE: Dissection of the SAN tissue should be done quickly in order to prevent ischemia-related injury. Attach the apex of the heart to the dish with a dissection pin. Then, while holding the inferior vena cava with Dumont #2 laminectomy forceps, insert a 22 G syringe needle through the inferior and superior vena cava to locate their position in the right atrium, which also identifies the approximate position of the SAN (located in the patch of tissue between the inferior and superior vena cava (Figure 4B). Using small dissection pins, pin the left and right atrial appendages to the dish. While holding the left atrial appendage with Dumont #2 laminectomy forceps, put a dissection pin through the left atrial appendage to hold it in place. While holding the right atrial appendage with Dumont #55 forceps, put a dissection pin through the right atrial appendage to hold it in place. NOTE: The same type of forceps can be used to hold the left and right atrial appendages if desired. Remove the syringe needle that spans the venae cavae. To release blood from the heart, use Castroviejo scissors to remove the apex of the heart (i.e., the bottom half) by making a transverse incision across the ventricles (Figure 4C). Then, wash the heart by adding warm (37 °C) heparinized Complete Tyrode's solution with a transfer pipette. Use Castroviejo scissors to cut along the atrioventricular septum keeping the incision closer to the ventricle than the atria. Continue cutting along the atrioventricular septum until the atria are separated from the ventricles. Cut along the interatrial septum to remove the left atrium. Place dissection pins in the periphery of the right atrium to make it lay flat (Figure 4D). Remove any remaining fat, vessels, or tissue from the atrium using the Castroviejo scissors. Locate the SAN in the right atrium, which in this orientation is approximately bordered by the superior vena cava (on the top), inferior vena cava (on the bottom), and cristae terminalis (on the left) (Figure 4D). ​NOTE: The crista terminalis appears as a dark muscular ridge between the right atrial appendage and the SAN. Often the SAN artery can also be seen coursing through the SAN (Figure 4D). 7. Preparing the MEA system for recording Add Tyrode's solution (from step 3) to the input solution bottle (Figure 5C) and oxygenate it by turning on the flow of carbogen gas (Figure 5A) to the system. NOTE: The Tyrode's solution used for the recording is slightly different in composition from the Complete Tyrode's solution used for the dissection. Verify the flow of carbogen by observing bubbles in the conical flask, which is used to humidify the gas (Figure 5B), and the input solution bottle (Figure 5C) . Insert the peristaltic pump inflow tubing (Figure 5D) into the Tyrode's recording solution (Figure 5C). Then, insert the peristaltic pump outflow tubing into the collection bottle (Figure 5I). Set the peristaltic pump to 25 rpm, which gives a flow rate of 2 mL/min and start the pump. Check the system for any buffer leakage or overflow. Set the temperature controller to 37 °C, the physiological temperature of mice (Figure 5E). 8. Placing the heart tissue on the MEA grid Transfer the dissected SAN tissue with the help of a paint brush (Figure 6A) from the dissecting Petri dish onto the MEA grid (Figure 1A1). While looking under an inverted microscope, gently position the tissue with a soft paint brush so that the SAN region overlays the electrode grid. Re-position the tissue as necessary to ensure it lays flat on the electrode grid, making good contact with the electrodes. NOTE: A soft paint brush is required for moving the tissue to avoid damaging the electrode grid. Once the tissue is correctly positioned, use bone forceps (or any curved forceps) to place the mesh over the tissue (Figure 6A) . Then use the bone forceps to place the harp anchor (Figure 6A) on the mesh to hold everything in place (Figure 6B). Take a picture of the positioning of the tissue on the MEA so that the activity of individual electrodes can be correlated with their anatomical location during the recording. This can be done by holding a smartphone up to the inverted microscope objective or by using an attached microscope camera. NOTE: If the orientation of the MEA is not changed after taking the picture, the top left electrode will appear as the first channel (Ch1) during recording. Place the MEA dish on the connector plate (Figure 5F and 6C) and carefully place the perfusion cap (Figure 6C) on the MEA dish without disturbing the harp slice anchor. The perfusion cap can be further secured using a piece of lab tape (Figure 6C). NOTE: In addition to having adjustable solution inflow and outflow pipes, the cap also has a port for the delivery of gas (Figure 6C). In addition, the reference electrode ring runs through the cap (Figure 6C). Allow the tissue to recover from handling and to acclimate to the chamber for 15-20 min prior to recording. 9. Setting the data acquisition protocol for recording NOTE: The following steps describe opening the software protocol for spontaneous beat recording and defining the recording conditions. The specifics of these steps may vary depending on the specific software being used, but the general outline should remain the same. Turn on the amplifier (Figure 5G), and set up a workflow for the recording in the software on the computer (Figure 5H). Open the software and click on the Workflow. Select Open New Folder. Open the From Templates folder. Select 64MD1-1920X1080 (depending on the resolution of your desktop). Open the QT folder. Open the Spontaneous recording folder. Select Beat_recording.moflo template and open it (Figure 7A). Set the recording parameters to specify the number of traces, trace duration, trace interval, input voltage, sampling rate, etc., according to the desired recording conditions (Figure 7B). NOTE: For beat frequency and interspike data acquisition, typically use an input range voltage of 2.9 mV, a 1-Hz high pass filter, a 1000-Hz low pass filter, and a sampling rate of 20 kHz. To mark different phases or conditions of the experiment, such as before and after drug administration, click on the Annotations tab to add the desired notations (Figure 7C). To specify the file destination for the data to be collected, select the Enable Storage box and enter the desired file name in the File name modifier box. 10. Performing the recording and collecting data Click the Record and Play button on the topmost menu bar of the acquisition software to start the recording. Acquire data for 10 traces of 1 min duration with intervals of 2 min between traces. From these initial traces, verify that the recorded waveforms are consistent with a healthy and high-quality tissue preparation by confirming that the majority of the recording channels exhibit signal amplitudes of ≥ 0.5 mV and identical inter-spike intervals (Figure 8). NOTE: An initial assessment of the activity and waveforms of the individual microelectrodes corresponding to their anatomical locations can be performed by referencing the picture acquired after positioning the tissue on the MEA. To measure the effects of drugs on the tissue, pause the recording after acquiring initial baseline data by clicking the pause button on the topmost menu bar. NOTE: The drug response phase of the experiment can be notated in the recording by clicking on the Annotations tab and adding the desired notation as described above (Figure 7C). Pause the pump and switch the pump inflow tubing from the normal recording solution to Tyrode's solution containing the desired drug of choice. NOTE: In the example experiment, Tyrode's solution was used with 1 mM 4-aminopyridine (4-AP). Restart the pump and un-pause the recording to begin collecting data again. Once the drug-infused Tyrode's solution has reached the tissue, record 10 traces in the same manner as done previously for the baseline recordings. NOTE: The traces will take some time to stabilize as the drug infuses into the recording chamber. The mechanism of action of the drug may also affect recording stability. For drugs that have reversible mechanisms of action, a washout period should also be recorded to confirm restoration of activity to baseline levels, which is an indicator of healthy tissue. Click Stop to conclude the recordings. Take a final picture of the positioning of the tissue on the MEA under the microscope in case the tissue has shifted following the initial recording setup procedure. 11. Cleaning the setup after the recording Clean the MEA. After finishing recording, gently remove the recording solution from the MEA dish using a 1 mL micropipette. NOTE: Be careful not to contact the MEA electrodes which can damage them. Remove the mesh and harp anchor with bone forceps (or any curved forceps). Then use a paint brush to dislodge the tissue from the MEA surface always being careful not to touch the individual microelectrodes. Using a wash bottle, gently rinse the MEA dish with ultrapure water about 3 to 4 times. Store the cleaned MEA immersed in ultrapure water at 4 °C. Rinse the system tubing by running ultrapure water through it for at least 5 min using the maximum speed setting on the peristaltic pump. ​NOTE: To prevent fungal growth, no water or buffer solution should be left inside the tubing after cleaning. 12. Analyzing the MEA recordings to measure SAN beat frequency Open the saved recorded data file in the "Beat_frequency_analysis" template of the analysis software (Figure 9) . Click on the Play button and allow the entire recording to run to visualize the data set and assign appropriate analysis parameters. Select the binning window for the desired display format of the data, whether it is displayed as an average per trace or average per time (Figure 10A). Select the channels to be included in the analysis and set the desired amplitude maxima or amplitude minima threshold values for automated waveform peak identification (Figure 10B). NOTE: An individual channel, a combination of channels, or all 64 channels can be selected for analysis at this step (Figure 9). If the threshold values selected are too close to the waveform's maxima and minima values, some waveform peaks may not be identified by the analysis software. Set the amount of pre-spike and post-spike time to be included in the analysis. NOTE: Settings of 50 ms pre-spike and 100 ms post-spike usually work well (Figure 10B). After setting the analysis conditions, click on the Play button again to rerun the data set and confirm that the analysis parameters are appropriate for spike extraction. For analysis, identify the three most stable consecutive traces that exhibit stable beating rate for each trace across the majority of channels both during the baseline period of the experiment and another three consecutive stable traces during the drug exposure period (Figure 10A). Specify the start and end traces for analysis and enter the time duration of each trace to be analyzed (Figure 9). Before starting the analysis, select the enable boxes for both Save beat per minute and Save interspike interval (Figure 10B). Enter the desired file name in the File name modifier box (Figure 10B). The analyzed data for beat frequency and interspike interval will be saved in the form of ASCII (text) format. NOTE: To analyze different conditions (such as baseline and drug response), analysis must be run separately for each condition. Click the Play and Record button on the top tab bar to start the analysis. To export the data for other applications, select the boxes for Save beat per minute and Save interspike interval (Figure 10B). Enter the desired file name in the File name modifier box (Figure 10B) and click Save to save the analyzed data in ASCII text format in the selected folder.

Representative Results

After allowing the tissue to acclimate in the dish for 15 min, 10 one-min traces are recorded. Our current protocol records activity for over an hour, but we have recorded stable firing patterns for ≥4 h in unpublished data not shown here. If an experimental preparation is good for data collection, each recording channel should exhibit consistent and evenly spaced recurring waveforms (i.e., spikes) of uniform shape for a given channel (Figure 11D). These waveforms correspond to individ…

Discussion

Practicing and mastering the SAN dissection process is imperative since the tissue is fragile and healthy tissue is necessary for a successful recording. During the SAN dissection, correct orientation is essential to obtain the proper region of tissue. However, the original orientation of the heart can be easily lost during the dissection process, which complicates this endeavor. Therefore, to ensure the proper left-right orientation, the atria should be visually inspected. Typically, the right atrium tends to be more tr…

Materials

4-Aminopyridine	Sigma	A78403-25G
22 gauge syringe needle	Fisher Scientific	14-826-5A	Used for dissection
23 gauge syringe needle	Fisher Scientific	14-826-6C	Used for dissection
60mm Petri Dishes	Genesee Scientific	32-105G
500mL Pyrex Bottle	Fisher Scientific	06-414-1C	Used to store solutions
1000 mL Pyrex Bottle	Fisher Scientific	06-414-1D	Used to store solutions
Bone Forceps	Fine Science Tools	16060-11
Calcium chloride dihydrate (CaCl2·2H2O)	Sigma-Aldrich	C5080-500G
Carbogen (95% O2, 5% CO2)
Castroviejo Scissors, 4"	Fine Science Tools	15024-10
D-(+)-Glucose	Sigma-Aldrich	G7021-1KG
Data Acquisition PC			CPU: Intel Xeon or Intel Core i7, Memory: 8GB, HDD: 1TB, Graphic Card: NVIDIA or On-board, Screen: 1920×1080
Dissection Microscope	Jenco
Dissecting Pins	Fine Science Tools	26002-20
Dumont #2 Laminectomy Forceps	Fine Science Tools	11223-20
Dumont #55 Forceps	Fine Science Tools	11295-51
Extra Fine Graefe Forceps	Fine Science Tools	11152-10
Glass Chamber	Grainger	49WF30	Used for mouse euthanization
Harp Anchor Kit	Warner Instruments	SHD-22CL/15 WI 64-0247
HCl	Fisher Chemicals	SA48-4	Used for pH balancing
Hemostat	Fine Science Tools	13013-14
Heparin	Aurobindo Pharma Limited IDA, Pashamylaram	NDC 63739-953-25
HEPES	Sigma-Aldrich	H3375-250G
Inverted Microscope	Motic	AE2000
Isoflurane	Patterson Veterinary	07-893-1389
Lab Tape	Fisher Scientific	15-950
Light for Dissection Microscope	Dolan-Jenner	MI150DG 660000391014
Magesium chloride (MgCl2)	Sigma-Aldrich	208337-100G
MED64 Head Amplifier	MED64	MED-A64HE1S
MED64 Main Amplifier	MED64	MED-A64MD1A
MED64 Perfusion Cap	MED64	MED-KCAP01
MED64 Perfusion Pipe Holder Kit	MED64	MED-KPK02
MED64 ThermoConnector	MED64	MED-CP04
Mesh	Warner Instruments	640246
Microelectrode array (MEA)	Alpha Med Scientific	MED-R515A
Mobius Software	WitWerx Inc.		Specific software for the MED64
NaOH	Fisher Chemicals	S320-500	Used for pH balancing
Normal Saline	Ultigiene	NDC 50989-885-17
Paint Brush	Fisher Scientific	NC1751733
Parafilm	Genesee Scientific	PM-996
Peristaltic Pump	Gilson	F155009
Peristaltic Pump Tubing	Fisher Scientific	14-171-298	1/8'' Interior Diameter
Polyethyleneimine	Sigma	P3143
Potassium chloride (KCl)	Sigma-Aldrich	P9333-500G
Potassium phosphate monobasic (KH2PO4)	Sigma-Aldrich	P5655-500G
Sodium Bicarbonate	Sigma	S6297
Sodium chloride (NaCl)	Fisher Scientific	S671-3
Sylgruard Elastomer Kit	Dow Corning	184 SIL ELAST KIT 0.5KG
Sodium Phosphate Monobasic	Sigma	S6566
Sodium tetraborate	Sigma	S9640
Surgical Scissors	Fine Science Tools	14074-09
Transfer Pipets (3mL graduated)	Samco Scientific	225