This work was supported in part by grants from the Intramural support research program (IRSP) from UMMC, AHA Scientist Development Grant (13SDG14000006) awarded to Mallikarjuna R. Pabbidi.

The male rats were housed in the Animal Care Facility at UMMC, which is approved by the Association for Assessment and Accreditation of Laboratory Animal Care (AAALAC). Animals had free access to food and water throughout the study. Animals were maintained in a controlled environment with temperature at 24 &#177; 2 &#176;C, humidity levels of 60&#8211;80% and 12 h light/dark cycles. All protocols were approved by the Animal Care and Use Committee of UMMC.
1. Preparation of Equipment
<ol>
	<li>Place a dual channel differential electrometer amplifier (see the Table of Materials) close to the vessel chamber and at the desired location.</li>
	<li>Connect the output of the amplifier channel A or B to the channel input of the digitizer with a BNC-BNC cable.</li>
	<li>Mount the probe in the micromanipulator and place it near the microscope and the myograph. The recording setup must be installed on a vibration-free table.</li>
	<li>Place the knobs and switches on the front of the amplifier in positions that configure it for this experiment as described in the manual.</li>
	<li>Connect the bath ground to the circuit ground of the amplifier via with an appropriate electrode. Similarly, ensure that the cage is grounded to the chassis of the amplifier.</li>
</ol>
2. Preparation of Microelectrodes and Assembly
<ol>
	<li>Use borosilicate glass microelectrodes (see the Table of Materials) and pull the glass tip to have a 8&#8211;10 mm taper, diameter of &#60;1 &#181;m and resistance of 80&#8211;120 M&#8486; when filled with 3 M KCl.
	<ol>
		<li>Use a standard puller to achieve a short gradual taper using the following settings: heat = 650; velocity = 20; pull = 25; time = 250 and loop twice for higher resistances and smaller tips. See the Table of Materials as the settings are instrument-specific. 
		NOTE: Tip diameters &#60;1 &#181;m will cause minimal damage to the cell when impaled.</li>
	</ol>
	</li>
	<li>Fill the microelectrode with 3 M KCl using a microfiber syringe (see the Table of Materials).
	<ol>
		<li>Slowly pull the plunger of the microfiber syringe up while injecting the 3 M KCl into the microelectrode to allow space for the fluid to fill and to prevent the formation of air bubbles inside the microelectrode.</li>
		<li>Fill the microelectrode until full and ensure that there are no air bubbles before placing it in the microelectrode holder. If bubbles are present, gently tap the microelectrode with a finger to remove the bubbles.</li>
	</ol>
	</li>
	<li>Exercising care, firmly push the electrode shank into the holder through the bored hole. If excess fluid is present, remove it with a tissue.</li>
	<li>Connect the electrode holder assembly to the amplifier probe. Conduct an electrode test, adjust the input offset, verify zero setting and check the probe input leakage as per the amplifier manual.</li>
	<li>Measure electrode resistance using an electrode test as shown in Table 1.</li>
	<li>Note that a working electrode displays a positive DC voltage shift of 1 mV/M&#937; at the channel output. On the other hand, if a large voltage appears at the channel output and on the meter, this indicates a blocked or broken electrode.</li>
	<li>Open the recording software, assign a name to the file and save it for future analysis in a storing software.</li>
</ol>
3. Isolation and Cannulation of the Middle Cerebral Artery
<ol>
	<li>Preparation the reagents.
	<ol>
		<li>Prepare normal and low calcium physiological salt solution (PSS) as described in Table 2.</li>
	</ol>
	</li>
	<li>Prepare the myograph.
	<ol>
		<li>Rinse the myograph chamber (see the Table of Materials) with distilled water multiple times to keep it free of debris. Load the chamber with 5 mL of normal PSS.</li>
		<li>Fill both glass cannulas with filtered normal PSS using a 5&#8211;10 mL syringe. Carefully fill the entire cannula and the attached tubing without introducing any air bubbles.</li>
		<li>Prepare two monofilament nylon sutures (10-0, 0.02 mm) with a half-knot each using blunt forceps.</li>
		<li>Place the partially closed suture knots on both cannulas slightly away from the tip using dissection forceps under a dissection microscope. Later these knots will be slid off and tied carefully onto the cannulated arterial ends to secure the vessel.</li>
	</ol>
	</li>
	<li>Isolate and cannulate the middle cerebral artery.
	<ol>
		<li>Induce deep anesthesia in a Sprague Dawley rat by using 2&#8211;4% inhaled isoflurane.</li>
		<li>Decapitate the rat using guillotine under deep anesthesia.</li>
		<li>Carefully remove the skull using a bone cutter and a scissor.</li>
		<li>Remove the brain from the skull and place it in 5 mL of low calcium PSS on ice.</li>
		<li>Identify and dissect out an unbranched segment of rat middle cerebral artery (MCA) with an inner diameter of 100&#8211;200 &#956;m from the brain using spring scissors and forceps.</li>
		<li>Mount the MCA onto the glass cannulas using fine forceps and secure by tightening the sutures in the myograph containing normal PSS.</li>
		<li>Close off the distal cannula so that there will be no flow within the MCAs.</li>
		<li>Connect the inflow pipette to a reservoir holding PSS to allow for control of intraluminal pressure which will be monitored with an in-line pressure transducer.</li>
		<li>Visualize the cannulated MCAs using a charge-coupled device camera (see the Table of Materials) mounted on an inverted microscope and an imaging software.</li>
		<li>Set the axial length of the MCA to an approximate length where it should appear neither rigid nor flaccid.</li>
		<li>Equilibrate the bath solution with O2 (95%) and CO2 (5%) at 37 &#176;C to provide adequate oxygenation, temperature and to maintain pH at 7.4.</li>
	</ol>
	</li>
	<li>Impale (penetrate the cell plasma membrane) the vascular smooth muscle cells.
	<ol>
		<li>Connect the ground electrode and keep it immersed in the PSS of the myograph.</li>
		<li>Illuminate the vessel chamber and look through the microscope to visualize the tip of the microelectrode in the bath solution. 
		NOTE: Alternatively, one can visualize the MCA and microelectrode on a computer having an imaging software.</li>
		<li>Use the controls of the micromanipulator to move the tip of the microelectrode close to the outer wall of the blood vessel. The micromanipulator and the tip of the microelectrode must be in a stable position in relation to the tissue. 
		NOTE: Before beginning experiments, confirm that the membrane voltage has stabilized. If the measured voltage is unstable, the connection between the electrode and the cell is not sealed, indicating a leak.</li>
		<li>Begin the recording.</li>
		<li>Slowly move the tip of the microelectrode towards the vessel, aiming for the center of the vessel using course or fine control of the micromanipulator. 
		NOTE: Occasionally, a small deflection in the recording may be observed when the microelectrode tip contacts a muscle fiber membrane.</li>
		<li>When the tip comes in close proximity to the vessel, advance the electrode forward in one rapid motion using the micromanipulator to impale the membrane of the muscle.</li>
		<li>At this point, one can begin observing the changes in Vm being recorded. Do not touch the micromanipulator when the microelectrode impales the membrane of the cell. 
		NOTE: The difference in voltage between the recording and reference electrode decreases from 0 mV to between -40 mV and -75 mV depending on the level of intravascular pressure or other excitatory or inhibitory stimuli. These readings characterize the transmembrane potential difference of the current cell.</li>
		<li>Perform multiple impalements on a single vessel in different areas of the vessel without damaging VSMCs in order to get accurate measurements.</li>
		<li>After recording, use the manipulator to remove the microelectrode in one rapid movement.</li>
		<li>Stop the recording and save data files for further analysis.</li>
	</ol>
	</li>
</ol>

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<li>Hughes, A. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Calcium+channels+in+vascular+smooth+muscle+cells%5BTitle%5D)+AND+%22Journal+of+Vascular+Research%22%5BJournal%5D)">Calcium channels in vascular smooth muscle cells.</a> Journal of Vascular Research. 32 (6), 353-370 (1995).</li>
<li>Large, W. A., Wang, Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Characteristics+and+physiological+role+of+the+Ca%282%2B%29-activated+Cl-+conductance+in+smooth+muscle%5BTitle%5D)+AND+%22American+Journal+of+Physiology%22%5BJournal%5D)">Characteristics and physiological role of the Ca(2+)-activated Cl- conductance in smooth muscle.</a> American Journal of Physiology. 271 (2 Pt 1), C435-C454 (1996).</li>
<li>Nelson, M. T., Conway, M. A., Knot, H. J., Brayden, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Chloride+channel+blockers+inhibit+myogenic+tone+in+rat+cerebral+arteries%5BTitle%5D)+AND+%22Journal+of+Physiology%22%5BJournal%5D)">Chloride channel blockers inhibit myogenic tone in rat cerebral arteries.</a> Journal of Physiology. 502 (Pt 2), 259-264 (1997).</li>
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<li>Gibson, A., McFadzean, I., Wallace, P., Wayman, C. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Capacitative+Ca2%2B+entry+and+the+regulation+of+smooth+muscle+tone%5BTitle%5D)+AND+%22Trends+in+Pharmacol+Sciences%22%5BJournal%5D)">Capacitative Ca2+ entry and the regulation of smooth muscle tone.</a> Trends in Pharmacol Sciences. 19 (7), 266-269 (1998).</li>
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<li>Shelly, D. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Na%28%2B%29+pump+alpha+2-isoform+specifically+couples+to+contractility+in+vascular+smooth+muscle%3A+evidence+from+gene-targeted+neonatal+mice%5BTitle%5D)+AND+%22American+Journal+of+Physiology-Cell+Physiology%22%5BJournal%5D)">Na(+) pump alpha 2-isoform specifically couples to contractility in vascular smooth muscle: evidence from gene-targeted neonatal mice.</a> American Journal of Physiology-Cell Physiology. 286 (4), C813-C820 (2004).</li>
<li>Coca, A., Garay, R. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aCoca%2C+A+%22Ionic+Transport+in+Hypertension+New+Perspectives%22">Ionic Transport in Hypertension New Perspectives</a>. , Taylor & Francis. (1993).</li>
<li>Harder, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Pressure-dependent+membrane+depolarization+in+cat+middle+cerebral+artery%5BTitle%5D)+AND+%22Circulation+Research%22%5BJournal%5D)">Pressure-dependent membrane depolarization in cat middle cerebral artery.</a> Circulation Research. 55 (2), 197-202 (1984).</li>
<li>Knot, H. J., Nelson, M. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Regulation+of+arterial+diameter+and+wall+%5BCa2%2B%5D+in+cerebral+arteries+of+rat+by+membrane+potential+and+intravascular+pressure%5BTitle%5D)+AND+%22Journal+of+Physiology%22%5BJournal%5D)">Regulation of arterial diameter and wall [Ca2+] in cerebral arteries of rat by membrane potential and intravascular pressure.</a> Journal of Physiology. 508 (Pt 1), 199-209 (1998).</li>
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<li>Harder, D. R., Sperelakis, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Action+potentials+induced+in+guinea+pig+arterial+smooth+muscle+by+tetraethylammonium%5BTitle%5D)+AND+%22American+Journal+of+Physiology%22%5BJournal%5D)">Action potentials induced in guinea pig arterial smooth muscle by tetraethylammonium.</a> American Journal of Physiology. 237 (1), C75-C80 (1979).</li>
<li>Nystoriak, M. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Fundamental+increase+in+pressure-dependent+constriction+of+brain+parenchymal+arterioles+from+subarachnoid+hemorrhage+model+rats+due+to+membrane+depolarization%5BTitle%5D)+AND+%22American+Journal+of+Physiology-Heart+and+Circulatory+Physiology%22%5BJournal%5D)">Fundamental increase in pressure-dependent constriction of brain parenchymal arterioles from subarachnoid hemorrhage model rats due to membrane depolarization.</a> American Journal of Physiology-Heart and Circulatory Physiology. 300 (3), H803-H812 (2011).</li>
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</ol>

The authors have nothing to disclose.

This article provides the necessary steps on how to use a sharp microelectrode impalement method to record Vm from a cannulated vessel preparation. This method is widely used, and offers high-quality, consistent recordings of Vm that answer a wide range of experimental questions.
Some critical considerations and troubleshooting steps are described here to ensure success of the method. The quality of the microelectrode (its sharpness and resistance) and the cellular proces...

The membrane potential (Vm) of a cell refers to the relative difference of ionic charge across the plasma membrane and the relative permeability of the membrane to these ions. The Vm is generated by the differential distribution of ions and is maintained by ion channels and pumps. Ion channels such as K+, Na+, and Cl&#8722; contribute substantially to the resting Vm. Vascular smooth muscle cells (VSMCs) express more than four different types of K+ channels1, two types of voltage-gated Ca2+ channels (VGCC)2, more than two types of Cl&#8722; channels3,4,5, store-operated Ca2+ channels6, stretch-activated cation channels7,8, and electrogenic sodium-potassium ATPase pumps9 in their plasma membranes, all of which may be involved in the regulation of Vm.
The Vm of VSMCs depends on lumen pressure. In non-pressurized vessels, Vm varies from -50 to -65 mV, however, in pressurized arterial segments, Vm ranges from -37 to -47 mV10. Elevation of intravascular pressure causes VSMCs to depolarize11, decreases the threshold for VGCC opening, and increases calcium influx contributing to the development of myogenic tone12. On the contrary, in passive or non-pressurized vessels, membrane hyperpolarization, due to high K+ channel activity, will prevent VGCC from opening, resulting in limited calcium entry and a decrease in intracellular calcium, contributing to less vascular tone13. Thus, Vm due to changes in lumen pressure appears to play an essential role in vascular tone development, and both VGCC and K+ channels play a crucial role in the regulation of Vm.
Vm varies between vessel type and species. Vm is -54 &#177; 1.3 mV in guinea pig superior mesenteric arterial strips14, -45 &#177; 1 mV in the rat middle cerebral arteries at 60 mmHg lumen pressure12, and -35 &#177; 1 mV in rat parenchymal arteries at 40 mmHg lumen pressure15. The resting Vm recorded in unstretched rat lymphatic muscle is -48 &#177; 2 mV16. Vm of cerebral VSMCs is more negative than in peripheral arteries. In comparison, feline middle cerebral arteries were reported to have a Vm of approximately -70 mV, while mesenteric and coronary arteries were reported to have -49 and -58 mV, respectively17,18. Differences in the Vm across vascular beds may reflect the differences in the expression and function of ion channels and electrogenic sodium-potassium pumps.
Increases and decreases in Vm are referred to as membrane depolarization and hyperpolarization, respectively. These alterations in Vm play a central role in many physiological processes, including ion-channel gating, cell signaling, muscle contraction, and action potential propagation. At a fixed pressure, many endogenous and synthetic vasodilator compounds that activate K+ channels cause membrane hyperpolarization, resulting in vasodilation1,13. Conversely, sustained membrane depolarization is vital in agonist-induced or receptor-mediated vasoconstriction19. Vm is a critical variable that not only regulates Ca2+ influx through VGCC13 but also influences the release of Ca2+ from internal stores20,21 and Ca2+-sensitivity of the contractile apparatus22.
While there are several methods to record Vm from different cell types, data collected from the microelectrode impalement method of cannulated vessels appears to be more physiological than data obtained from isolated VSMCs. When recorded from isolated VSMC using current clamp methods, Vm is seen as spontaneous transient hyperpolarizations in VSMCs24. Isolated VSMCs are not in the syncytium, and the changes in the series resistance may contribute to the oscillatory behavior of Vm. On the other hand, oscillatory behavior is not observed when Vm is recorded from intact vessels, probably due to cell-cell contact between VSMCs that are in syncytium in the artery and are summated throughout the vessel leading to a stable Vm24. Thus, measurement of Vm from pressurized vessels using standard microelectrode impalement technique is relatively close to the physiological conditions.
Recording Vm from cannulated vessels could provide vital information, since Vm of VSMCs that are in syncytium is one of the major determinants of vascular tone and blood flow, and modulation of the Vm could provide a way to dilate or constrict blood vessels. Thus, it is essential to understand the methodology involved in recording Vm. This article describes intracellular recording of Vm from cannulated middle cerebral arteries (MCAs) using a microelectrode impalement method. This protocol will describe how to prepare MCAs, microelectrodes, set up the electrometer and perform the impalement method to record Vm. Also, representative data, common issues that were encountered when using this method and potential issues are discussed.

<table><tbody><tr><td>&lt;strong&gt;Dissection instruments&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Aneshetic Vaporiser</td><td>Parkland scientific</td><td>V3000PK</td><td></td></tr><tr><td>Dissection microscope</td><td>Nikon Instruments Inc., NY</td><td>Eclipse Ti-S</td><td></td></tr><tr><td>Kleine Guillotine Type 7575</td><td>Harvard Apparatus, MA</td><td>73-198</td><td></td></tr><tr><td>Littauer Bone Cutter</td><td>Fine science tools</td><td>16152-15</td><td></td></tr><tr><td>Moria MC40 Ultra Fine Forceps</td><td>Fine science tools</td><td>11370-40</td><td></td></tr><tr><td>Surgical scissors Sharp-Blunt</td><td>Fine science tools</td><td>14008-14</td><td></td></tr><tr><td>Suture</td><td>Harvard Apparatus</td><td>72-3287</td><td></td></tr><tr><td>Vannas Spring Scissors</td><td>Fine science tools</td><td>15018-10</td><td></td></tr><tr><td>&lt;strong&gt;Electrophysiology Instruments&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Charge-coupled device camera</td><td>Qimaging, , BC</td><td>Retiga 2000R</td><td></td></tr><tr><td>Differential electrometer amplifier</td><td>WPI</td><td>FD223A</td><td></td></tr><tr><td>In-line pressure transducer</td><td>Harvard Apparatus, MA</td><td>MA1 72-4496</td><td></td></tr><tr><td>Micromanipulator</td><td>Thor labs</td><td>PCS-5400</td><td></td></tr><tr><td>Microelectrodes</td><td>Warner Instruments LLC, CT</td><td>G200-6,</td><td></td></tr><tr><td>Micro Fil (Microfiber syringe)</td><td>WPI</td><td>MF28G67-5</td><td></td></tr><tr><td>Microelectrode holder</td><td>WPI</td><td>MEH1SF</td><td></td></tr><tr><td>Myograph</td><td>Living Systems Instrumentation, VT</td><td>CH-1-SH</td><td></td></tr><tr><td>Puller</td><td>Sutter Instrument, San Rafael, CA</td><td>P-97</td><td></td></tr><tr><td>Vibration-free table</td><td>TMC</td><td>3435-14</td><td></td></tr><tr><td>&lt;strong&gt;Softwares&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Clampex 10</td><td>Molecular devices</td><td></td><td></td></tr><tr><td>p Clamp 10</td><td>Molecular devices</td><td></td><td></td></tr><tr><td>Imaging software</td><td>Nikon, NY</td><td>NIS-elements</td><td></td></tr><tr><td>&lt;strong&gt;Chemicals&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>NaCl</td><td>Sigma</td><td>S7653</td><td></td></tr><tr><td>KCl</td><td>Sigma</td><td>P4504</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>M7506</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2 &lt;/sub&gt;</td><td>Sigma</td><td>C3881</td><td></td></tr><tr><td>HEPES</td><td>Sigma</td><td>H7006</td><td></td></tr><tr><td>Glucose</td><td>Sigma</td><td>G7021</td><td></td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>S0751</td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Sigma</td><td>S5761</td><td></td></tr></tbody></table>

microelectrode impalement method to record membrane potential from a cannulated middle cerebral artery

Membrane potential (Vm) of vascular smooth muscle cells determines vessel tone and thus blood flow to an organ. Changes in the expression and function of ion channels and electrogenic pumps that regulate Vm in disease conditions could potentially alter Vm, vascular tone, and blood flow. Thus, a basic understanding of electrophysiology and the methods necessary to accurately record Vm in healthy and diseased states are essential. This method will allow modulating Vm using different pharmacological agents to restore Vm. Although there are several methods, each with its advantages and disadvantages, this article provides protocols to record Vm from cannulated resistance vessels such as the middle cerebral artery using the microelectrode impalement method. Middle cerebral arteries are allowed to gain myogenic tone in a myograph chamber, and the vessel wall is impaled using high resistance microelectrodes. The Vm signal is collected through an electrometer, digitized, and analyzed. This method provides an accurate reading of the Vm of a vessel wall without damaging the cells and without changing the membrane resistance.

The presented method can be reliably used to record Vm in cannulated vessels. A brief procedure describing how to isolate MCA from the brain is presented in Figure 1A. After separating the brain from the skull, the MCA was dissected out and placed in a Petri dish containing low calcium PSS. Part of the connective tissue that was attached was also dissected along with MCA using spring scissors and forceps to prevent damage to MCA during the isolatio...

Watch this Scientific Journal Video about Microelectrode Impalement Method to Record Membrane Potential from a Cannulated Middle Cerebral Artery at JoVE.com

Microelectrode Impalement Method to Record Membrane Potential from a Cannulated Middle Cerebral Artery

The primary goal of this article is to provide details of how to record membrane potential (Vm) from the middle cerebral artery using the microelectrode impalement method. The cannulated middle cerebral artery is equilibrated to gain myogenic tone, and the vessel wall is impaled using high resistance microelectrodes.

Membrane potential (Vm) of vascular smooth muscle cells determines vessel tone and thus blood flow to an organ. Changes in the expression and function of ...

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Research

JoVE Journal

Medicine

6.7K Views.  University of Mississippi Medical Center. The primary goal of this article is to provide details of how to record membrane potential (Vm) from the middle cerebral artery using the microelectrode impalement method. The cannulated middle cerebral artery is equilibrated to gain myogenic tone, and the vessel wall is impaled using high resistance microelectrodes.

Video: Microelectrode Impalement Method to Record Membrane Potential from a Cannulated Middle Cerebral Artery

Multi-electrode Array Recordings of Human Epileptic Postoperative Cortical Tissue

Epilepsy, affecting about 1% of the population, comprises a group of neurological disorders characterized by the periodic occurrence of seizures, which disrupt normal brain function. Despite treatment with currently available antiepileptic drugs targeting neuronal functions, one third of patients with epilepsy are pharmacoresistant. In this condition, surgical resection of the brain area generating seizures remains the only alternative treatment. Studying human epileptic tissues has contributed to understand new epileptogenic mechanisms during the last 10 years. Indeed, these tissues generate spontaneous interictal epileptic discharges as well as pharmacologically-induced ictal events which can be recorded with classical electrophysiology techniques. Remarkably, multi-electrode arrays (MEAs), which are microfabricated devices embedding an array of spatially arranged microelectrodes, provide the unique opportunity to simultaneously stimulate and record field potentials, as well as action potentials of multiple neurons from different areas of the tissue. Thus MEAs recordings offer an excellent approach to study the spatio-temporal patterns of spontaneous interictal and evoked seizure-like events and the mechanisms underlying seizure onset and propagation. Here we describe how to prepare human cortical slices from surgically resected tissue and to record with MEAs interictal and ictal-like events ex vivo.

We here describe how to perform multi-electrode array recordings of human epileptic cortical tissue. Epileptic tissue resection, slice preparation and multi-electrode array recordings of interictal and ictal events are demonstrated in detail.

Epilepsy, affecting about 1% of the population, comprises a group of neurological disorders characterized by the periodic occurrence of seizures, which ...

Direct-current Stimulation and Multi-electrode Array Recording of Seizure-like Activity in Mice Brain Slice Preparation

Cathodal transcranial direct-current stimulation (tDCS) induces suppressive effects on drug-resistant seizures. To perform effective actions, the stimulation parameters (e.g., orientation, field strength, and stimulation duration) need to be examined in mice brain slice preparations. Testing and arranging the orientation of the electrode relative to the position of the mice brain slice are feasible. The present method preserves the thalamocingulate pathway to evaluate the effect of DCS on anterior cingulate cortex seizure-like activities. The results of the multichannel array recordings indicated that cathodal DCS significantly decreased the amplitude of the stimulation-evoked responses and duration of 4-aminopyridine and bicuculline-induced seizure-like activity. This study also found that cathodal DCS applications at 15 min caused long-term depression in the thalamocingulate pathway. The present study investigates the effects of DCS on thalamocingulate synaptic plasticity and acute seizure-like activities. The current procedure can test the optimal stimulation parameters including orientation, field strength, and stimulation duration in an in vitro mouse model. Also, the method can evaluate the effects of DCS on cortical seizure-like activities at both the cellular and network levels.

Studies have shown that cathodal transcranial direct-current stimulation can produce suppressive effects on drug-resistant seizures. In this study, an in vitro experimental setup was devised in which the direct-current stimulation and multielectrode array recording of seizure-like activity were evaluated in mice brain slice preparation. The direct-current stimulation parameters were evaluated.

Cathodal transcranial direct-current stimulation (tDCS) induces suppressive effects on drug-resistant seizures. To perform effective actions, the ...

Neuroscience

Recording and Modulation of Epileptiform Activity in Rodent Brain Slices Coupled to Microelectrode Arrays

Temporal lobe epilepsy (TLE) is the most common partial complex epileptic syndrome and the least responsive to medications. Deep brain stimulation (DBS) is a promising approach when pharmacological treatment fails or neurosurgery is not recommended. Acute brain slices coupled to microelectrode arrays (MEAs) represent a valuable tool to study neuronal network interactions and their modulation by electrical stimulation. As compared to conventional extracellular recording techniques, they provide the added advantages of a greater number of observation points and a known inter-electrode distance, which allow studying the propagation path and speed of electrophysiological signals. However, tissue oxygenation may be greatly impaired during MEA recording, requiring a high perfusion rate, which comes at the cost of decreased signal-to-noise ratio and higher oscillations in the experimental temperature. Electrical stimulation further stresses the brain tissue, making it difficult to pursue prolonged recording/stimulation epochs. Moreover, electrical modulation of brain slice activity needs to target specific structures/pathways within the brain slice, requiring that electrode mapping be easily and quickly performed live during the experiment. Here, we illustrate how to perform the recording and electrical modulation of 4-aminopyridine (4AP)-induced epileptiform activity in rodent brain slices using planar MEAs. We show that the brain tissue obtained from mice outperforms rat brain tissue and is thus better suited for MEA experiments. This protocol guarantees the generation and maintenance of a stable epileptiform pattern that faithfully reproduces the electrophysiological features observed with conventional field potential recording, persists for several hours, and outlasts sustained electrical stimulation for prolonged epochs. Tissue viability throughout the experiment is achieved thanks to the use of a small-volume custom recording chamber allowing for laminar flow and quick solution exchange even at low (1 mL/min) perfusion rates. Quick MEA mapping for real-time monitoring and selection of stimulating electrodes is performed by a custom graphic user interface (GUI).

We illustrate how to perform recording and electrical modulation of 4-aminopyridine-induced epileptiform activity in rodent brain slices using microelectrode arrays. A custom recording chamber maintains tissue viability throughout prolonged experimental sessions. Live electrode mapping and selection of stimulating pairs are performed by a custom graphical user interface.

Temporal lobe epilepsy (TLE) is the most common partial complex epileptic syndrome and the least responsive to medications. Deep brain stimulation (DBS) ...

Electrocorticographic Recording of Cerebral Cortex Areas Manipulated Using an Adeno-Associated Virus Targeting Cofilin in Mice

The use of electrocorticographic (ECoG) recordings in rodents is relevant to sleep research and to the study of a wide range of neurological conditions. Adeno-associated viruses (AAVs) are increasingly used to improve understanding of brain circuits and their functions. The AAV-mediated manipulation of specific cell populations and/or of precise molecular components has been tremendously useful to identify new sleep regulatory circuits/molecules and key proteins contributing to the adverse effects of sleep loss. For instance, inhibiting activity of the filamentous actin-severing protein&#160;cofilin&#160;using AAV prevents sleep deprivation-induced memory impairment. Here, a protocol is&#160;described that combines the manipulation of cofilin function in a cerebral cortex area with the recording of ECoG activity to examine whether cortical cofilin modulates the wakefulness and sleep ECoG signals. AAV injection is performed during the same surgical procedure as the implantation of ECoG and electromyographic (EMG) electrodes in adult male and female mice. Mice are anesthetized, and their heads are shaved. After skin cleaning and incision, stereotaxic coordinates of the motor cortex are determined, and the skull is pierced at this location. A cannula prefilled with an AAV expressing cofilinS3D, an inactive form of cofilin, is slowly positioned in the cortical tissue. After AAV infusion, gold-covered screws (ECoG electrodes) are screwed through the skull and cemented to the skull with gold wires inserted in the neck muscles (EMG electrodes). The animals are allowed three weeks to recover and to ensure sufficient expression of cofilinS3D. The infected area and cell type are verified using immunohistochemistry, and the ECoG is analyzed using visual identification of vigilance states and spectral analysis. In summary, this combined methodological approach allows the investigation of the precise contribution of molecular components regulating neuronal morphology and connectivity to the regulation of synchronized cerebral cortex activity during wakefulness and sleep.

This article describes a protocol for the manipulation of molecular targets in the cerebral cortex using adeno-associated viruses and for monitoring the effects of this manipulation during wakefulness and sleep using electrocorticographic recordings.

The use of electrocorticographic (ECoG) recordings in rodents is relevant to sleep research and to the study of a wide range of neurological conditions. ...

Mapping Cerebral Blood Flow and Electrical Fields: Electrode Placement in Mice

Placement of Extracranial Stimulating Electrodes and Measurement of Cerebral Blood Flow and Intracranial Electrical Fields in Anesthetized Mice

The detection of cerebral blood flow (CBF) responses to various forms of neuronal activation is critical for understanding dynamic brain function and variations in the substrate supply to the brain. This paper describes a protocol for measuring CBF responses to transcranial alternating current stimulation (tACS). Dose-response curves are estimated both from the CBF change occurring with tACS (mA) and from the intracranial electric field (mV/mm). We estimate the intracranial electrical field based on the different amplitudes measured by glass microelectrodes within each side of the brain. In this paper, we describe the experimental setup, which involves using either bilateral laser Doppler (LD) probes or laser speckle imaging (LSI) to measure the CBF; as a result, this setup requires anesthesia for the electrode placement and stability. We present a correlation between the CBF response and the current as a function of age, showing a significantly larger response at higher currents (1.5 mA and 2.0 mA) in young control animals (12-14 weeks) compared to older animals (28-32 weeks) (p &lt; 0.005 difference). We also demonstrate a significant CBF response at electrical field strengths &lt;5 mV/mm, which is an important consideration for eventual human studies. These CBF responses are also strongly influenced by the use of anesthesia compared to awake animals, the respiration control (i.e., intubated vs. spontaneous breathing), systemic factors (i.e., CO2), and local conduction within the blood vessels, which is mediated by pericytes and endothelial cells. Likewise, more detailed imaging/recording techniques may limit the field size from the entire brain to only a small region. We describe the use of extracranial electrodes for applying tACS stimulation, including both homemade and commercial electrode designs for rodents, the concurrent measurement of the CBF and intracranial electrical field using bilateral glass DC recording electrodes, and the imaging approaches. We are currently applying these techniques to implement a closed-loop format for augmenting the CBF in animal models of Alzheimer's disease and stroke.

Author Spotlight: Stimulation-Based Approach to Improve Cerebral Blood Flow in Alzheimer's Model 

We describe a protocol for assessing dose-response curves for extracranial stimulation in terms of brain electrical field measurements and a relevant biomarker-cerebral blood flow. Since this protocol involves invasive electrode placement into the brain, general anesthesia is needed, with spontaneous breathing preferred rather than controlled respirations.

The detection of cerebral blood flow (CBF) responses to various forms of neuronal activation is critical for understanding dynamic brain function and ...

In Vivo Recording of Intracellular Signals from a Single Neuron

Source: Altwegg-Boussac, T., et al. <a href="https://app.jove.com/v/53576">Induction of an Isoelectric Brain State to Investigate the Impact of Endogenous Synaptic Activity on Neuronal Excitability In Vivo.</a> J. Vis. Exp. (2016).
This video demonstrates the recording of electrical signals from single brain neurons using a fine-tipped glass micropipette and a specialized setup. The micropipette is carefully inserted into the rat&#39;s brain and is buzzed to penetrate the neuron to record internal neuronal electrical signals.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

JoVE Encyclopedia of Experiments

Neurophysiology

Electrophysiology Techniques

Recording Neural Activity of Unfolded Hippocampal Tissue Using Penetrating Microelectrode Array

Source: Zhang, M., et al. <a href="https://app.jove.com/v/52601">Neural Activity Propagation in an Unfolded Hippocampal Preparation with a Penetrating Micro-electrode Array.</a> J. Vis. Exp. (2015).
This video demonstrates the procedure to set up a recording chamber with a penetrating microelectrode array (PMEA) to capture neural signals from an unfolded hippocampus. This method allows us to study the speed and direction of propagation of neural activity within the unfolded hippocampus in vitro.

All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review.
1. Solutions for Surgery and ...

Impact of Direct Current Stimulation on Evoked Epileptic Seizures in Mouse Brain Slices

Source: Lu, H., et al. <a href="https://app.jove.com/v/53709">Direct-current Stimulation and Multi-electrode Array Recording of Seizure-like Activity in Mice Brain Slice Preparation.</a> J. Vis. Exp. (2016)
The video demonstrates a study for assessing the impact of direct current stimulation (DCS) on epileptic seizures in mouse brain slices. The brain slices containing the connected medial thalamus (MT) and anterior cingulate cortex (ACC) regions are placed in a multielectrode array (MEA). Electrical stimuli to the MT and seizure-inducing drugs are applied to induce epileptic seizures in the ACC. DCS is then applied to assess its impact on evoked seizures.

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Preparing Experimental Solution ...

Stimulation and Modulation Techniques

Monitoring Seizure-Induced Neural Electrical Activity in Brain Slices Using Microelectrode Arrays

Once the 4-AP aCSF level and the recording temperature are stabilized as desired, turn the perfusion and the suction stopcocks to the off position to temporarily stop them. Quickly transfer one brain slice onto the MEA recording chamber using an inverted glass Pasteur pipette.
Adjust its position on the MEA recording area as needed, using a fire-polished curled Pasteur pipette or a soft, compact small brush. Place the hold-down anchor on the brain slice. Restart the perfusion and the suction by turning their stopcocks back to the on position.
Take a picture of the brain slice using a camera mounted on an inverted microscope stage. Then, run the script mapMEA on the computer software to start the GUI to map the electrodes. Click the Browse button to load the picture of the brain slice. Make sure that the reference electrode appears in the upper column of the left half side of the MEA. Then, click the Activate Pointer button.
Select the top and bottom electrodes in the leftmost column of the array to mark the x, y-coordinates for image straightening and electrode mapping. From the Slice Type dropdown menu, select Horizontal. Tick the default structures checkbox.
Using the numbered push buttons below the brain slice picture, select the electrodes corresponding to the ROI, and click the corresponding push buttons in the structure panel to assign them. Repeat this step for each ROI. Afterward, press the Save button. The software will generate a result folder containing a table, reporting the selected electrodes and ROIs.
For electrophysiology study, turn on the stimulus unit at least 10 minutes prior to the stimulation protocol to allow self-calibration and stabilization. Then, start the stimulus control software and verify that the stimulator and MEA amplifier are correctly connected, as indicated by a green LED in the main panel of the stimulus control software.
Set up the stimulation in bipolar configuration. Connect the stimulus unit to the ground. Then, connect the electrode pairs in contact with the pyramidal cell layer of the CA1 proximal subiculum among the ones mapped with the script mapMEA. To acquire data, press the Play button in the main panel of the recording software. Record at least four ictal discharges.
Next, run a fast input/output test to identify the best stimulus intensity. To do so, use the main panel to design a square biphasic positive-negative current pulse with a duration of 100 microseconds per phase.
In the Pulse Amplitude tab, enter an initial pulse amplitude of 100 microamperes per phase. Set the stimulation pulse at 0.2 hertz or lower by adding an interpulse interval of 5 seconds or longer. Increase the stimulus amplitude by 50 to 100-microampere steps at each trial until the stimulation can reliably evoke interictal-like events in the parahippocampal cortices.
For electrical modulation of limbic ectogenesis, program the stimulus unit to deliver the stimulation protocol of interest, and use the stimulus amplitude identified during the input/output test.
After the stimulation stops, verify the network recovery to prestimulus condition by recording at least four ictal discharges.

EoE Sub Articles

Concurrent Electroencephalographic and Local Field Potential Recordings in an Anesthetized Rat

In this procedure, clean and dry the skull surrounding the burr hole using a cotton swab. Carefully apply the conductive EEG paste on a flat side of an EEG spider electrode. Leave a small hole clear of EEG paste on the spider electrode to allow a multilaminar microelectrode to pass through the hole without contacting the paste and the spider electrode.
Align the spider electrode with the burr hole in the skull with the EEG paste facing the skull. Carefully press the spider electrode onto the skull, making firm contact with the skull via the EEG paste.
Remove any paste obscuring the burr hole using a needle on a syringe and remove excessive EEG paste beyond the periphery of the spider electrode, so that the contact between the spider electrode and the skull is spatially constrained to the size of the electrode. Then, smear EEG paste onto the reference electrode for the EEG, and place it securely inside the incision at the back of the rat&#39;s neck.
Next, connect the EEG electrodes to the pre-amplifier via a passive signal splitter for low-impedance signals. At this stage, test the resistance of the EEG probe to make sure it is below five kiloohms. If not, add more EEG paste and make sure the spider electrode makes a good contact with the skull.
Subsequently, mount a micromanipulator arm on the stereotactic frame. Connect a linear 16-channel microelectrode to a 16-channel acute head stage clipped securely onto the micromanipulator arm. Then, smear EEG paste onto the reference electrode for the microelectrode, and then place it securely inside the incision next to the reference electrode for the EEG. Adjust the angle of the micromanipulator arm so that the microelectrode is perpendicular to the cortical surface.
Now, lower the microelectrode under a microscope so that the tip of the microelectrode is aimed at the tiny opening at the bottom of the burr hole, until the uppermost electrode just penetrates the cortical surface. Care must be taken to avoid forcing the microelectrode onto the surface of the dura, as this would break the electrode.
Insert the microelectrode to the cortical surface at a depth of 1500 micrometers. Microadjust the depth by applying a train of stimulus to the whisker pad, and observing the 16-channel evoked LFP on a PC monitor. Carefully turn the z-axis knob on the micromanipulator until the highest amplitude of the evoked LFP occurs as this coincides with the layer 4 in the cortex.