eoe sub articles

EoE Sub Articles

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Fabrication of the Stimulation Electrodes
<ol>
	<li>To make one stimulation electrode, cut six 10 cm long pieces of miniature hook-up wires, remove 2 cm of the cladding on one end, and 1 cm on the other end.</li>
	<li>Twist two cables together and solder them at the shorter peeled side.</li>
	<li>Acquire a strip of three-tape packaging of any integrated circuit with the SOT-353 (Small Outline Transistor-353) case.</li>
	<li>Stitch the twisted wires through the three holes of the package. The longer peeled segment of the wires should be in the package&#39;s cavities. Position the cables so that the end of the insulation is in line with the edge of the holes.</li>
	<li>Put the dental cement in the holes to fix the wires. 
	CAUTION: The dental cement must be dense enough to avoid leaking into the cavity of the package.</li>
	<li>Once the cement is solidified (minimum 10 min), flip the package and put a thin layer of superglue into the hole to form a watertight seal toward the porous cement. Avoid gluing the peeled wires.</li>
	<li>Twist the cables together on the shorter peeled side and solder them (Figure 1A).</li>
	<li>Grab the longer peeled segment of the wires with the tip of fine tweezers, roll them around, and wrap them into the cavities (Figure 1B). If necessary, cut the excess cable.</li>
</ol>
2. Fabrication of the Recording Electrodes
<ol>
	<li>To make one recording triplet electrode (Figure 1C-D), cut a one-centimeter-long piece of stainless steel tube using a high-speed rotary saw. Remove the fin from the ends of the tube with a sharp needle and ensure that the tube is clean of debris and completely permeable. 
	NOTE: Do not use pliers, as they will distort the tube.</li>
	<li>Bend a gold-plated board spacer pin with a minimum length of 3 cm to form a J-shape holder. Cut the longer leg of the &#39;J&#39; shape and solder it together to get back the original shape to make a detachable joint between the long linear part, which will be held by the stereotaxic device, and the &#39;U&#39; shape, which will hold the electrode, and serve as an anchoring point. Glue the tube and the shorter leg of the &#39;J&#39; holder together. Glue some additional 1 cm long bars, if necessary, for stability.</li>
	<li>Cut 3 pieces of 2.5 cm long electrode wires with stainless steel ultrafine scissors. Make sure that the cut is clear and sharp and that the insulation is intact at the circular surface of the electrode wire. Also, make sure that the cut tip is not bent due to the force applied. Bend 1 or 2 mm on one end of each of the wires at different angles (e.g., one left straight, one 45&#176;, and one right angle) to make them distinguishable.</li>
	<li>Fill the tube with the wires. Choose an appropriate spacing of the electrodes for the experiment (e.g., for cortical local field potential (LFP) and current source density (CSD) registration, the recommended protrusion of the tips of the wires from the opening of the tube is 4.5 mm, 4.1 mm, and 3.7 mm, respectively). Note which wire corresponds to which depth for maintaining a proper channel order.</li>
	<li>Once the wires align right next to each other and are parallel with the tube, to fix the electrode wires, place a single drop of liquid superglue at both ends of the tube with a sharp needle. Make sure that the glue is flowing into the tube due to the capillary effect, but not towards the recording sites at the protruding ends.</li>
	<li>Prepare an electrode interface board with a proper micro-connector compatible with the recording system to be used.</li>
	<li>Introduce the non-recording, bent ends of the wires into the holes of the electrode interface board corresponding to the desired recording channel. When all the wires are in position, push the gold pins into the holes with tweezers. 
	NOTE: Gold pins may be fixed in place using dental cement, but in most cases, they are stable enough without this safety step.</li>
</ol>
3. Anesthesia
<ol>
	<li>Place the animal in a sealed anesthesia induction chamber and fill it with 4 - 5% of isoflurane in 2 L/min medical air.</li>
	<li>When the rat is recumbent, remove the animal from the chamber and place it in a stereotaxic frame with an appropriate ventilation nosepiece.</li>
	<li>Set the isoflurane level to 2%, though the rate of airflow should remain the same. Check the level of anesthesia. If there is an absence of the paw withdrawal reflex in response to pinching, continue, otherwise increase the depth of anesthesia.</li>
	<li>Monitor and maintain the rat&#39;s body temperature at 37 &#177; 0.5 &#176;C with a homeothermic monitoring system. Apply a droplet of paraffin ointment to the eyes to prevent cornea desiccation. Repeat this procedure several times during the surgery, if necessary.</li>
	<li>Subcutaneously inject 0.3 mg/kg atropine to avoid mucus formation in the airways. For longer surgeries, repeat this every four hours. Monitor breathing and adjust the vaporizer, if needed.</li>
</ol>
4. Implantation of the Stimulation and Recording Electrodes
NOTE: Autoclave all the necessary surgical instruments and carefully follow the general rules of asepsis and antisepsis during the whole procedure. Avoid touching nonsterile areas outside of the surgical area. Immerse the electrodes in ethyl alcohol (70%) for 30 min before implantation.
<ol>
	<li>Remove most of the hair from the scalp with a hair clipper. Apply depilatory cream on the scalp, spread it evenly on the surface, and wait a few minutes. Use a spatula to gently remove the cream and the remaining hair. Rinse the skin with water, then with disinfectant.</li>
	<li>Inject 1 - 2% lidocaine (do not exceed a total dose of 7 mg/kg) subcutaneously to numb the skin. Apply a single drop of vet ointment for the eyes (e.g., paraffin).</li>
	<li>Make a thorough and long (~2 cm) sagittal incision in the midline with a scalpel from the forehead to the neck. Dissect the tissues, including the periosteum, from the skull. Then, using a chisel or tooth tweezers, clean the area between the cristae of the two temporal bones. Retract the dissected skin using four bulldogs to keep the skull exposed.</li>
	<li>Gently place fine tweezers between the steep edge of the temporal bone and the muscles and separate them. Make jigging movements to expose as much of the large surface of the temporal bone as possible, preferably from the edge of the occipital bone to the plane of the coronal sutures, without damaging the muscles.</li>
	<li>Place retractors bitemporally to keep the temporal bones exposed.</li>
	<li>Rinse the surface of the skull with 1 - 2 mL 3% hydrogen peroxide (H2O2), then wash it with 1 - 2 mL water.</li>
	<li>Air-dry the surface of the temporal bones very carefully, mopping up the moisture with ocular sticks. Test if the stimulation electrodes fit onto the cleaned vertical skull surface (the upper edge of the stimulation electrode should be in line with the edge of the crista of the temporal bone). If necessary, re-adjust the bulldogs and the retractors or shape the stimulation electrodes with scissors.</li>
	<li>Fill the cavities of the stimulation electrodes with electroconductive gel and put a thin layer of glue onto the rim of the electrodes.</li>
	<li>Place the stimulation electrode on the dry surface of the temporal bone with one accurate movement and hold it firmly in place for one minute with fine tweezers. Make sure that no moisture is in contact with the glue. Mop up with ocular sticks, if necessary. 
	CAUTION: If the electrode becomes unstable, remove it. After cleaning the bone, repeat this step with a new stimulation electrode.</li>
	<li>Put dental cement over the edges of the stimulation electrode, continuously drying up the leaking moisture in the tissues with ocular sticks. Then, cover the whole stimulation electrode with cement.</li>
	<li>After the cement is completely hardened, repeat these steps on the contralateral side.</li>
	<li>Drill some holes all over the skull for anchoring screws. Drive miniature screws into the holes and put dental cement over them. Use a ~10% smaller diameter drill head compared to the screw diameter.</li>
	<li>Depending on the purpose of the experiments, either solder a connector to the ends of the cables, place it on the surface, and cover it with cement, or continue implanting the recording electrodes and optical fibers, or prepare a cranial window over the skull. In this latter case, solder the connectors and anchor them to the construct only at the end. In case of the need for long-term simultaneous TES and LFP recordings, implant the aforementioned electrode triplets instead of single wires. 
	NOTE: This setup allows for the removal of common mode stimulation artefacts8. For the details of recording electrode implantation, see previous protocols.</li>
	<li>Wash the exposed tissues abundantly with disinfectant. Subcutaneously inject 1 - 2% lidocaine (do not exceed a total dose of 7 mg/kg).</li>
	<li>Debride the wound edges and close them with simple interrupted sutures around the connector/implant. Then, disinfect the wound with povidone-iodine.</li>
	<li>Inject 5 mg/kg carprofene subcutaneously. Repeat this if necessary. 
	NOTE: Any other analgesics may be used, matching the requirements set by the local ethical permission board.</li>
	<li>Suspend anesthesia, release the earbars and nosepiece, and put the animal in a postoperative recovery cage to regain consciousness.</li>
	<li>Ensure that the animal is under close supervision during the first postoperative hours. Continue to monitor the implant area periodically. 
	NOTE: The animal should not be left unattended until it has regained sufficient consciousness to maintain sternal recumbency. House the animal individually until it fully recovers from the procedure.</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Cyanoacrylate liquid</td>
			<td>Henkel</td>
			<td>Loctite 401</td>
			<td></td>
		</tr>
		<tr>
			<td>Cyanoacrylate gel</td>
			<td>Henkel</td>
			<td>Loctite 454</td>
			<td></td>
		</tr>
		<tr>
			<td>Wire for stimulation electrodes</td>
			<td>Phoenix Wire Inc.</td>
			<td>36744MHW - PTFE Microminiature Hook-Up Wire</td>
			<td></td>
		</tr>
		<tr>
			<td>Board spacer</td>
			<td>E-tec Interconnect</td>
			<td>SP1-020-S378/01-55</td>
			<td></td>
		</tr>
		<tr>
			<td>Connector</td>
			<td>E-tec Interconnect</td>
			<td>P2510I-02</td>
			<td></td>
		</tr>
		<tr>
			<td>Tape packaging for stimulation electrodes</td>
			<td>Nexperia</td>
			<td>74HC1G00GW</td>
			<td>Tape packaging of any integrated circuit with SOT-353 case can be used</td>
		</tr>
		<tr>
			<td>Grip Cement Industrial Grade</td>
			<td>Caulk Dentsply</td>
			<td>675571 (powder) 675572 (solvent)</td>
			<td></td>
		</tr>
		<tr>
			<td>Electroconductive gel</td>
			<td>Rextra</td>
			<td>ECG Gel</td>
			<td></td>
		</tr>
		<tr>
			<td>Recording electrode wire</td>
			<td>California Fine Wire Co.</td>
			<td>.002 (50 micron) Tungsten 99.95% (CFW Material #: 100-211), HMl-Natural, cut to 3.0 inch pieces, Round, Cut length piece wire</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrafine scissors</td>
			<td>Hammacher Instrumente</td>
			<td>Stainless HSB 544-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Stainless steel tube</td>
			<td>Vita Needle Company</td>
			<td>29 RW, 304SS Tubing, T.I.G. Welded and Plug</td>
			<td></td>
		</tr>
		<tr>
			<td>High speed rotary saw</td>
			<td>Dremel</td>
			<td>Model # 395</td>
			<td></td>
		</tr>
		<tr>
			<td>Rotary saw holder</td>
			<td>Dremel</td>
			<td>Model # 220</td>
			<td></td>
		</tr>
		<tr>
			<td>Rotary saw cut-off wheel</td>
			<td>Dremel</td>
			<td>Model # 409</td>
			<td></td>
		</tr>
		<tr>
			<td>Ocular sticks</td>
			<td>Lohmann-Rauscher</td>
			<td>Pro-ophta Ocular Sticks</td>
			<td></td>
		</tr>
		<tr>
			<td>Wet disinfectant</td>
			<td>Egis</td>
			<td>Betadine</td>
			<td></td>
		</tr>
		<tr>
			<td>Dry disinfectant</td>
			<td>Wagner Pharma</td>
			<td>Reseptyl-urea</td>
			<td></td>
		</tr>
		<tr>
			<td>Drilling machine</td>
			<td>NSK-Nakanishi United Kingdom</td>
			<td>Vmax35RV Pack</td>
			<td></td>
		</tr>
		<tr>
			<td>Anchoring screws</td>
			<td>Antrin Miniature Specialties, Inc.</td>
			<td>000-120x1/16 SL BIND MS SS</td>
			<td></td>
		</tr>
	</tbody>
</table>

implanting an on-head electrode system in a rat model for real-time seizure detection

Source: Koz&#225;k, G., et al. <a href="https://appjove.unicartagenaproxy.elogim.com/v/56669">Chronic Transcranial Electrical Stimulation and Intracortical Recording in Rats.</a> J. Vis. Exp. (2018).
This video demonstrates the method of implanting an on-head electrode system in an anesthetized rat for real-time seizure detection. The procedure includes placing stimulation and recording electrodes, securing them with dental cement, and connecting them to a head-mounted amplifier.

<img alt="Figure 1" src="/files/ftp_upload/22833/22833_Figure1.jpg" /> 
Figure 1: Steps of stimulation and recording electrode fabrication. (A) Twisted wires were stitched through the holes and fixed to the packaging before wrapping the peeled wires into the cavities. (B) The final form of the stimulation electrodes. Inset: wrapped wires inside the packaging; (C) Side view of the recording electrodes; (D</str...

Implanting an On-Head Electrode System in a Rat Model for Real-Time Seizure Detection

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

Take an anesthetized rat with the exposed, cleaned temporal bones situated bilaterally to the intracortical neurons-rich parietal cortex.
Place a stimulation electrode filled with an electroconductive gel on one temporal bone for electrical contact and secure it with dental cement.
Fix another stimulation electrode on the opposite temporal bone for uniform stimulation of the parietal region.
Drill holes in the skull and insert anchor screws to stabilize the implant.
Place the recording electrodes in the parietal cortex to capture signals from the intracortical neurons which are involved in the development of seizures, the abnormal electrical activity.
Secure the screws and electrodes with dental cement.
Connect the electrode wires to a head-mounted amplifier and attach a mesh protector around it to protect the amplifier and the electrodes.
Wash the exposed tissues with disinfectant, inject local anesthesia, and suture the wound.
The rat with the implanted on-head electrode system is ready for real-time seizure detection.

implanting-an-on-head-electrode-system-rat-model-for-real-time

Universidad de Cartagena UDEC

Research

JoVE Encyclopedia of Experiments

Neuroscience

Neurophysiology

Electrophysiology Techniques

55 Views. Source: Kozák, G., et al. Chronic Transcranial Electrical Stimulation and Intracortical Recording in Rats. J. Vis. Exp. (2018). This video demonstrates the method of implanting an on-head electrode system in an anesthetized rat for real-time seizure detection. The procedure includes placing stimulation and recording electrodes, securing them with dental cement, and connecting them to a head-mounted amplifier. 

Video: Implanting an On-Head Electrode System in a Rat Model for Real-Time Seizure Detection - Concept

Syringe-injectable Mesh Electronics for Stable Chronic Rodent Electrophysiology

Implantable brain electrophysiology probes are valuable tools in neuroscience due to their ability to record neural activity with high spatiotemporal resolution from shallow and deep brain regions. Their use has been hindered, however, by mechanical and structural mismatches between the probes and brain tissue that commonly lead to micromotion and gliosis with resulting signal instability in chronic recording experiments. In contrast, following the implantation of ultraflexible mesh electronics via syringe injection, the mesh probes form a seamless, gliosis-free interface with the surrounding brain tissue that enables stable tracking of individual neurons on at least a year timescale. This protocol details the key steps in a typical mouse neural recording experiment using syringe-injectable mesh electronics, including the fabrication of mesh electronics in a standard photolithography-based process possible at many universities, loading mesh electronics into standard capillary needles, stereotaxic injection in vivo, connection of the mesh input/output to standard instrumentation interfaces, restrained or freely moving recording sessions, and histological sectioning of brain tissue containing mesh electronics. Representative neural recordings and histology data are presented. Investigators familiar with this protocol will have the knowledge necessary to incorporate mesh electronics into their own experiments and take advantage of the unique opportunities afforded by long-term stable neural interfacing, such as studies of aging processes, brain development, and the pathogenesis of brain disease.

Mesh electronics probes seamlessly integrate and provide stable, long-term, single-neuron level recording within the brain. This protocol uses mesh electronics for in vivo experiments, involving the fabrication of mesh electronics, loading into needles, stereotaxic injection, input/output interfacing, recording experiments, and histology of tissue containing mesh probes.

Implantable brain electrophysiology probes are valuable tools in neuroscience due to their ability to record neural activity with high spatiotemporal ...

JoVE Journal

Bioengineering

Updated Technique for Reliable, Easy, and Tolerated Transcranial Electrical Stimulation Including Transcranial Direct Current Stimulation

Transcranial direct current stimulation (tDCS) is a noninvasive method of neuromodulation using low-intensity direct electrical currents. This method of brain stimulation presents several potential advantages compared to other techniques, as it is noninvasive, cost-effective, broadly deployable, and well-tolerated provided proper equipment and protocols are administered. Even though tDCS is apparently simple to perform, correct administration of the tDCS session, especially the electrode positioning and preparation, is vital for ensuring reproducibility and tolerability. The electrode positioning and preparation steps are traditionally also the most time consuming and error-prone. To address these challenges, modern tDCS techniques, using fixed-position headgear and pre-assembled sponge electrodes, reduce complexity and setup time while also ensuring that the electrodes are consistently placed as intended. These modern tDCS methods present advantages for research, clinic, and remote-supervised (at home) settings. This article provides a comprehensive step-by-step guide for administering a tDCS session using fixed-position headgear and pre-assembled sponge electrodes. This guide demonstrates tDCS using commonly applied montages intended for motor cortex and dorsolateral prefrontal cortex (DLPFC) stimulation. As described, selection of the head size and montage-specific headgear automates electrode positioning. Fully assembled pre-saturated snap-electrodes are simply affixed to the set position snap-connectors on the headgear. The modern tDCS method is shown to reduce setup time and reduce errors for both novice and expert operators. The methods outlined in this article can be adapted to different applications of tDCS as well as other forms of transcranial electrical stimulation (tES) such as transcranial alternating current stimulation (tACS) and transcranial random noise stimulation (tRNS). However, since tES is application specific, as appropriate, any methods recipe is customized to accommodate subject, indication, environment, and outcome specific features.

When administering transcranial direct current stimulation (tDCS), reproducible electrode preparation and placement are vital for a tolerated and effective session. The purpose of this article is to demonstrate updated modern setup procedures for the administration of tDCS and related transcranial electrical stimulation techniques, such as transcranial alternating current stimulation (tACS).

Transcranial direct current stimulation (tDCS) is a noninvasive method of neuromodulation using low-intensity direct electrical currents. This method of ...

Medicine

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. ...

Implanting an On-Head Electrode System in a Rat Model for Real-Time Seizure Detection

Transcript

Implanting an On-Head Electrode System in a Rat Model for Real-Time Seizure Detection

Syringe-injectable Mesh Electronics for Stable Chronic Rodent Electrophysiology

Updated Technique for Reliable, Easy, and Tolerated Transcranial Electrical Stimulation Including Transcranial Direct Current Stimulation

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