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. Preparing Electrodes
Note: A three-channel transmitter is used for surgical implantation, which enables 2 electroretinogram (ERG) and 1 visual evoked potential (VEP) recording to be conducted simultaneously. The three active and three inactive electrodes need to be pre-fashioned into a ring shape before implantation in order to attach to the eye. For identification purposes, the manufacturer has enclosed active electrodes in half-white, half-colored plastic sheaths, while inactive electrodes are covered in full-colored sheaths. The ground electrode (clear plastic sheath) is left unaltered for all active and inactive electrodes to conduct steps 1.1, 1.2, 1.3, and 1.7.
<ol>
	<li>Untwist the double-stranded stainless-steel electrode with two fine-tipped pliers.</li>
	<li>Trim one of the stainless-steel strands (approximately 1 cm from the tip), leaving a single longer straight strand remaining to shape the ring electrode.</li>
	<li>Fold the single stainless-steel strand back onto itself and twist, forming a smooth ring at the tip of the electrode.</li>
	<li>For the ERG active electrodes, fashion this loop ~ 0.2 - 0.5 mm in diameter by twisting the base of the loop (for the purpose described here, shape two active electrodes in this way to record ERG from both eyes), and for the ERG inactive and VEP electrodes make the loop diameter ~ 0.8 mm diameter (in this example, do this for one active VEP electrode and all three inactive electrodes).</li>
	<li>Hook the circular VEP active electrode around a stainless-steel screw (diameter 0.7 mm, length 3 mm) so the electrode rests against the screw head.</li>
	<li>Hook the 3 inactive electrodes (2 ERG, 1 VEP) around a second stainless steel screw (diameter 0.7 mm, length 3 mm).</li>
	<li>Pull the plastic sleeve forward over the sharp ends of the two stainless-steel strands to reduce irritation.</li>
	<li>Sterilize the telemetry transmitters by soaking them in 2% glutaraldehyde for more than 10 hours at approximately 25 &#176;C. Then, rinse the transmitter with sterile saline three times.</li>
</ol>
2. Transmitter Implantation
<ol>
	<li>Animal Preparation&#8203;
	<ol>
		<li>Disinfect the surgical area prior to experimentation by cleaning it with 70% ethanol. Autoclave all surgical equipment before use and maintain equipment in chlorhexidine when not in use during surgery. Cover the animal with a surgical drape during surgery to maintain a sterile environment. Ensure all experimenters wear surgical masks, sterile gloves, and gowns.</li>
		<li>Induce anesthesia with 1.5 - 2% isoflurane at a flow rate of 3 L/min and maintained at 1.5 - 2 % at 2 L/min throughout the surgery. Confirm sufficient depth of anesthesia by the absence of a pedal reflex upon pinching the muscle between the toes.</li>
		<li>Shave a 40 mm x 30 mm area over the abdomen from above the groin to the sternum.</li>
		<li>Shave a 30 mm x 20 mm area over the forehead, posterior to the eyes and anterior to the ears.</li>
		<li>Disinfect the two shaved areas. For the forehead area disinfect with 10% povidone-iodine three times (avoid the use of alcohol-based antiseptics for area near the eye, being consistent with the Standard of Practice set out by the Association of Surgical Technologists). Over the abdomen, disinfect with 10% povidone-iodine and 70% ethanol.</li>
		<li>Apply 1 drop of proxymetacaine to the cornea for additional topical anesthesia.</li>
		<li>Apply 1 drop of carboxymethylcellulose sodium to the cornea to prevent drying of the eyes.</li>
	</ol>
	</li>
	<li>Surgical implantation
	<ol>
		<li>Make a 10 mm incision on the head along the vertical midline between the ears with a surgical scalpel.</li>
		<li>Make a 5 mm incision on the abdomen through the skin layer along the midline below the sternum.</li>
		<li>Tunnel a 5 mm diameter cannula subcutaneously from the abdomen incision to the head incision.</li>
		<li>Feed the electrode wires (3 active and 3 inactive) of the transmitter through the cannula from the abdomen to the head.</li>
		<li>Leave the reference electrode with the transmitter base and cover the electrode tip with aseptic gauze.</li>
		<li>Cover the electrode tips (3 active and 3 inactive) with aseptic gauze.</li>
		<li>Secure the rat&#39;s head to a stereotaxic platform.</li>
		<li>Extend the forehead incision to 30mm in length with surgical scissors.</li>
		<li>Expose the surgical area by retracting loose skin with 2 sutures (3 - 0) at ~ 3 and 9 o&#39;clock.</li>
		<li>Scrape off the periosteum overlying the skull using sterilized gauze to expose bregma, lambda, and midline sutures.</li>
		<li>Drill two holes through the skull at the VEP active (7 mm ventral to bregma, 3 mm lateral to midline) and inactive (5 mm rostral to bregma on the midline) stereotaxic coordinates.</li>
		<li>Attach VEP active and inactive electrodes with pre-attached stainless-steel screws (diameter 0.7 mm, length 3 mm) to the skull with a small screwdriver to ~ 1 mm in depth into the premade holes. This anchors the screw to the bone without damaging the underlying cortical tissue.</li>
		<li>To implant the ERG active electrodes, use an 8 - 0 suture to retract the upper eyelid temporarily.</li>
		<li>Insert a 16 to 21 G cannula subcutaneously from behind the eye through to the superior conjunctival fornix.</li>
		<li>Remove the guiding needle.</li>
		<li>Feed the active electrode through the shortened plastic catheter from the forehead towards the eye. Then, remove the plastic catheter.</li>
		<li>Use a temporary suture (8 - 0), which is threaded through the electrode loop, to prevent the electrode from retracting back into the tunnel.</li>
		<li>Make a 0.5 mm incision on the superior conjunctiva at 12 o&#39;clock, 1 mm behind the limbus. Use blunt dissection to expose the underlying sclera.</li>
		<li>Implant an 8 - 0 or 9 - 0 suture immediately behind the limbus at half scleral thickness.</li>
		<li>Remove temporary suture from ERG active electrode.</li>
		<li>Anchor the ERG active electrode to the half scleral thickness suture by tying 3 consecutive knots, ensuring the electrode tip is situated close to the limbus.</li>
		<li>Close the conjunctival flap using 1 to 2 interrupted sutures (8 - 0 to 9 - 0). Ensure that the conjunctiva completely covers the ERG electrode to improve comfort.</li>
		<li>Remove the eyelid retracting suture.</li>
		<li>Repeat the procedure for the contralateral eye.</li>
		<li>Apply cyanoacrylate gel over the skull to secure all stainless screws and electrode wires. Ensure the ERG active electrodes are not pulled too tight before securing them to enable eye movements.</li>
		<li>Close the head wound using a non-absorbable 3 - 0 suture.</li>
		<li>Rotate the rodent to expose the abdominal area. Lengthen the abdominal dermal incision to 40 mm along the Linea alba with surgical scissors.</li>
		<li>Make a 35 mm incision through the inner muscle wall to expose the inner abdominal cavity.</li>
		<li>Using two sutures (3 - 0), the transmitter body is attached to the animal&#39;s right-hand side inner abdominal wall. Avoid contacting the liver.</li>
		<li>Loop the ground electrode and secure it in this shape with a suture (3 - 0). Then, place it free-floating in the abdominal cavity.</li>
		<li>Close the peritoneum using a continuous suture (3 - 0).</li>
		<li>Close the skin incision using interrupted sutures (3 - 0).</li>
	</ol>
	</li>
	<li>Post-operative care&#8203;
	<ol>
		<li>Monitor the animal until it has regained sufficient consciousness to maintain sternal recumbency. House the animal singly following the surgery.</li>
		<li>Administer carprofen subcutaneously for analgesia (5 mg/kg) once a day for 4 days.</li>
		<li>Add prophylactic oral antibiotics (Enrofloxin, 5 mg/kg) to the drinking water for 7 days post-surgery.</li>
		<li>Apply an anti-inflammatory ointment to skin incision sites to reduce irritation for the first 7 days post-surgery.</li>
	</ol>
	</li>
</ol>
3. Conduct ERG and VEP Recordings in Conscious Rats&#8203;
<ol>
	<li>Dark adapt animal for 12 hours prior to ERG and VEP recordings.</li>
	<li>Conduct all experimental manipulations under dim red illumination (17.4 cd.m-2, &#955;max = 600 nm)</li>
	<li>Apply topical anesthesia (0.5% proxymetacaine) and dilating (0.5% tropicamide) drops to the cornea.</li>
	<li>Guide the conscious rodent into a custom-made, clear restrainer. 
	Note: The length of this plastic tube can be adjusted to accommodate different-sized rats, with the overall diameter fixed at 60 mm. The front end of the device is tapered to minimize head movement and contains perforations to enable normal breathing. This tapered front allows alignment and stabilization of the rat&#39;s head and eyes to the opening of the Ganzfeld sphere. Note that the rodent has been acclimatized to the restrainer (3 to 5 occasions) prior to surgery.</li>
	<li>Place the rodent in front of the Ganzfeld bowl with the eyes aligned with the opening of the bowl.</li>
	<li>Turn on the indwelling transmitter by passing a magnet within ~ 5 cm of it. Then, verify that the transmitter is on by checking the LED status light on the receiver base.</li>
	<li>Collect signals over a range of luminous energies (i.e., -5.6 to 1.52 log cd.s.m2).</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Bioamplifier</td>
			<td>ADInstruments</td>
			<td>ML 135</td>
			<td>Amplifies ERG and VEP signals</td>
		</tr>
		<tr>
			<td>Carboxymethylcellulose sodium 1.0%</td>
			<td>Allergan</td>
			<td>CAS 0009000-11-7</td>
			<td>Maintain corneal hydration during surgery</td>
		</tr>
		<tr>
			<td>Carprofen 0.5%</td>
			<td>Pfizer Animal Health Group</td>
			<td>CAS 53716-49-7</td>
			<td>Post-surgery analgesia, given with injectable saline for fluid replenishment</td>
		</tr>
		<tr>
			<td>Chlorhexadine 0.5%</td>
			<td>Orion Laboratories</td>
			<td>27411, 80085</td>
			<td>Disinfection of surgical instrument</td>
		</tr>
		<tr>
			<td>Cyanoacrylate gel activator</td>
			<td>RS components</td>
			<td>473-439</td>
			<td>Quickly dries cyanoacrylate gel</td>
		</tr>
		<tr>
			<td>Cyanocrylate gel</td>
			<td>RS components</td>
			<td>473-423</td>
			<td>Fix stainless screws to skull</td>
		</tr>
		<tr>
			<td>Dental burr</td>
			<td>Storz Instruments, Bausch and Lomb</td>
			<td>E0824A</td>
			<td>Miniature drill head of ~0.7mm diameter for making a small hole in the skull over each hemisphere to implant VEP screws</td>
		</tr>
		<tr>
			<td>Drill</td>
			<td>Bosch</td>
			<td>Dremel 300 series</td>
			<td>Automatic drill for trepanning</td>
		</tr>
		<tr>
			<td>Enrofloxin</td>
			<td>Troy Laboratories</td>
			<td></td>
			<td>Prophylactic antibiotic post surgey</td>
		</tr>
		<tr>
			<td>Ganzfeld integrating sphere</td>
			<td>Photometric Solutions International</td>
			<td></td>
			<td>Custom designed light stimulator: 36 mm diameter, 13 cm aperture size</td>
		</tr>
		<tr>
			<td>Gauze swabs</td>
			<td>Multigate Medical Products Pty Ltd</td>
			<td>57-100B</td>
			<td>Dries surgical incision and exposed skull surface during surgery</td>
		</tr>
		<tr>
			<td>Isoflurane 99.9%</td>
			<td>Abbott Australasia Pty Ltd</td>
			<td>CAS 26675-46-7</td>
			<td>Proprietory Name: Isoflo(TM) Inhalation anaaesthetic. Pharmaceutical-grade inhalation anesthetic mixed with oxygen gas for VEP electrode implant surgery</td>
		</tr>
		<tr>
			<td>Kenacomb ointment</td>
			<td>Aspen Pharma Pty Ltd</td>
			<td></td>
			<td>To reduce skin irritation and itching after surgery</td>
		</tr>
		<tr>
			<td>Luxeon LEDs</td>
			<td>Phillips Lighting Co.</td>
			<td></td>
			<td>For light stimulation, twenty 5 watt and one 1 watt LEDs, controlled by Scope software</td>
		</tr>
		<tr>
			<td>Needle (macrosurgery)</td>
			<td>World Precision Instruments</td>
			<td>501959</td>
			<td>for suturing abdominal and head surgery, used with 3-0 suture, eye needle, cutting edge 5/16 circle Size 1, 15mm</td>
		</tr>
		<tr>
			<td>Needle holder (macrosurgery)</td>
			<td>World Precision Instruments</td>
			<td>500224</td>
			<td>To hold needle during abdominal and head surgery</td>
		</tr>
		<tr>
			<td>Needle holder (microsurgery)</td>
			<td>World Precision Instruments</td>
			<td>555419NT</td>
			<td>To hold needle during ocular surgery</td>
		</tr>
		<tr>
			<td>Optiva catheter</td>
			<td>Smiths Medical International LTD</td>
			<td>16 or 21 G</td>
			<td>Guide corneal active electrodes from skull to conjunctiva</td>
		</tr>
		<tr>
			<td>Povidone iodine 10%</td>
			<td>Sanofi-Aventis</td>
			<td>CAS 25655-41-8</td>
			<td>Proprietory name: Betadine, Antiseptic to prepare the shaved skin for surgery 10%, 500 mL</td>
		</tr>
		<tr>
			<td>Powerlab data acquisition system</td>
			<td>ADInstruments</td>
			<td>ML 785</td>
			<td>Acquire signal from telemetry transmitter, paired to telemetry data converter</td>
		</tr>
		<tr>
			<td>Proxymetacaine 0.5%</td>
			<td>Alcon Laboratories</td>
			<td>CAS 5875-06-9</td>
			<td>Topical ocular analgesia</td>
		</tr>
		<tr>
			<td>Restrainer</td>
			<td>cutom made</td>
			<td></td>
			<td>Front of the restrainer is tapered to minimize head movement, length can be adjusted to accommodate different rat length, overall diameter is 60 mm.</td>
		</tr>
		<tr>
			<td>Scapel blade</td>
			<td>R.G. Medical Supplies</td>
			<td>SNSM0206</td>
			<td>For surgical incision</td>
		</tr>
		<tr>
			<td>Scissors (macrosurgery)</td>
			<td>World Precision Instruments</td>
			<td>501225</td>
			<td>for cutting tissue on the abodmen and forhead</td>
		</tr>
		<tr>
			<td>Scissors (microsurgery)</td>
			<td>World Precision Instruments</td>
			<td>501232</td>
			<td>To dissect the conjunctiva for electrode attachment</td>
		</tr>
		<tr>
			<td>Scope Software</td>
			<td>ADInstruments</td>
			<td>version 3.7.6</td>
			<td>Simultaneously triggers the stimulus via the ADI Powerlab system and collects data</td>
		</tr>
		<tr>
			<td>Shaver</td>
			<td>Oster</td>
			<td>Golden A5</td>
			<td>Shave fur from surgical areas</td>
		</tr>
		<tr>
			<td>Stainless streel screws</td>
			<td>MicroFasteners</td>
			<td>L001.003CS304</td>
			<td>0.7 mm shaft diameter, 3 mm in length</td>
		</tr>
		<tr>
			<td>Stereotaxic frame</td>
			<td>David Kopf</td>
			<td>Model 900</td>
			<td>A small animal stereotaxic instrument for locating the implantation landmarks on the skull</td>
		</tr>
		<tr>
			<td>Surgical drape</td>
			<td>Vital Medical Supplies</td>
			<td>GM29-612EE</td>
			<td>Ensure sterile enviornment during surgery</td>
		</tr>
		<tr>
			<td>Suture (macrosurgery)</td>
			<td>Ninbo medical needles</td>
			<td>3-0</td>
			<td>for suturing abdominal and head surgery, sterile silk braided, 60cm</td>
		</tr>
		<tr>
			<td>Suture needle (microsurgery)</td>
			<td>Ninbo medical needles</td>
			<td>8-0 or 9-0</td>
			<td>for ocular surgery including, suturing electrode to sclera and closing conjunctival wound, nylon suture, 3/8 circle 1&#215;5, 30cm</td>
		</tr>
		<tr>
			<td>Telemetry data converter</td>
			<td>DataSciences International</td>
			<td>R08</td>
			<td>allows telemetry signal to interface with data collection software</td>
		</tr>
		<tr>
			<td>Telemetry Data Exchange Matrix</td>
			<td>DataSciences International</td>
			<td></td>
			<td>Gathers data from transmitters, pair with receiver</td>
		</tr>
		<tr>
			<td>Telemetry data receiver</td>
			<td>DataSciences International</td>
			<td>RPC-1</td>
			<td>Receives telemetry data from transmitter</td>
		</tr>
		<tr>
			<td>Telemetry transmitter</td>
			<td>DataSciences International</td>
			<td>F50-EEE</td>
			<td>3 channel telemetry transmitter</td>
		</tr>
		<tr>
			<td>Tropicamide 0.5%</td>
			<td>Alcon Laboratories</td>
			<td></td>
			<td>Iris dilation</td>
		</tr>
		<tr>
			<td>Tweezers (macrosurgery)</td>
			<td>World Precision Instruments</td>
			<td>500092</td>
			<td>Manipulate tissues during abdominal and head surgery</td>
		</tr>
		<tr>
			<td>Tweezers (microsurgery)</td>
			<td>World Precision Instruments</td>
			<td>500342</td>
			<td>Manipulate tissues during ocular surgery</td>
		</tr>
	</tbody>
</table>

recording of electroretinogram and visual evoked potential in a rat

Source: Charng, J. et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/54160">Implantation and Recording of Wireless Electroretinogram and Visual Evoked Potential in Conscious Rats.</a> J. Vis. Exp. (2016).
This video demonstrates how to record electroretinogram (ERG) and visual evoked potential (VEP) in a conscious rat. The rat, with surgically implanted electrodes, is first dark-adapted. After applying dilating drops, the animal is placed in a restrainer and positioned in a Ganzfeld bowl. The ERG and VEP signals are then recorded.

Recording of Electroretinogram and Visual Evoked Potential in a Rat

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

Begin with a rat with surgically implanted electrodes in its eye and brain.
The electroretinogram, or ERG electrode, measures the electrical activity of the retina, while the visual evoked potential, or VEP electrodes, record brain signals.
These electrodes are connected to a transmitter implanted in the rat&#39;s abdomen.
Dark-adapt the animal before the recordings.
Perform the procedure under dim red light.
Apply anesthetic and dilating drops to the cornea.
Guide the animal into a restrainer to minimize head movement.
Position the animal in front of the Ganzfeld bowl, which ensures uniform exposure of the retina under controlled illumination.
Align the animal&#39;s eyes with the opening of the bowl.
Pass a magnet over the implanted transmitter to turn it on. Confirm its activation on the receiver.
Record ERG and VEP signals under different illumination conditions.

recording-of-electroretinogram-and-visual-evoked-potential-in-a-rat

Nanyang Technological University

Research

JoVE Encyclopedia of Experiments

Neuroscience

Neurophysiology

Electrophysiology Techniques

76 vues. Source : Charng, J. et al. Implantation et enregistrement de l’électrorétinogramme sans fil et du potentiel évoqué visuel chez les rats conscients. J. Vis. Exp. (2016). Cette vidéo montre comment enregistrer l’électrorétinogramme (ERG) et le potentiel évoqué visuel (VEP) chez un rat conscient. Le rat, avec des électrodes implantées chirurgicalement, est d’abord adapté à l’obscurité. Après avoir appliqué des gouttes dilatantes, l’animal est placé dans une contention et...

Vidéo: Enregistrement d’un électrorétinogramme et d’un potentiel évoqué visuel chez un rat - Concept

Direct-Coupled Electroretinogram (DC-ERG) for Recording the Light-Evoked Electrical Responses of the Mouse Retinal Pigment Epithelium

The retinal pigment epithelium (RPE) is a specialized monolayer of cells strategically located between the retina and the choriocapillaris that maintain the overall health and structural integrity of the photoreceptors. The RPE is polarized, exhibiting apically and basally located receptors or channels, and performs vectoral transport of water, ions, metabolites, and secretes several cytokines.
In vivo noninvasive measurements of RPE function can be made using direct-coupled ERGs (DC-ERGs). The methodology behind the DC-ERG was pioneered by Marmorstein, Peachey, and colleagues using a custom-built stimulation recording system and later demonstrated using a commercially available system. The DC-ERG technique uses glass capillaries filled with Hank&#8217;s buffered salt solution (HBSS) to measure the slower electrical responses of the RPE elicited from light-evoked concentration changes in the subretinal space due to photoreceptor activity. The prolonged light stimulus and length of the DC-ERG recording make it vulnerable to drift and noise resulting in a low yield of useable recordings. Here, we present a fast, reliable method for improving the stability of the recordings while reducing noise by using vacuum pressure to reduce/eliminate bubbles that result from outgassing of the HBSS and electrode holder. Additionally, power line artifacts are attenuated using a voltage regulator/power conditioner. We include the necessary light stimulation protocols for a commercially available ERG system as well as scripts for analysis of the DC-ERG components: c-wave, fast oscillation, light peak, and off response. Due to the improved ease of recordings and rapid analysis workflow, this simplified protocol is particularly useful in measuring age-related changes in RPE function, disease progression, and in the assessment of pharmacological intervention.

Direct-Coupled Electroretinogram DC-ERG for Recording the Light-Evoked Electrical Responses of the Mouse Retinal Pigment Epithelium

Here, we present a method for recording light-evoked electrical responses of the retinal pigment epithelium (RPE) in mice using a technique known as DC-ERGs first described by Marmorstein, Peachey, and colleagues in the early 2000s.

Électrorétinogramme couplé direct (DC-ERG) pour l’enregistrement des réponses électriques évoquées par la lumière de l’épithélium pigmentaire rétinien de souris

The retinal pigment epithelium (RPE) is a specialized monolayer of cells strategically located between the retina and the choriocapillaris that maintain ...

Recherche

JoVE Journal

Chimie

Simultaneous Recording of Electroretinography and Visual Evoked Potentials in Anesthetized Rats

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

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

Enregistrement simultané de Électrorétinographie et potentiels évoqués visuels chez des rats anesthésiés

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

Neurosciences

Stimulus-specific Cortical Visual Evoked Potential Morphological Patterns

This paper presents a methodology for the recording and analysis of cortical visual evoked potentials (CVEPs) in response to various visual stimuli using 128-channel high-density electroencephalography (EEG). The specific aim of the described stimuli and analyses is to examine whether it is feasible to replicate previously reported CVEP morphological patterns elicited by an apparent motion stimulus, designed to simultaneously stimulate both ventral and dorsal central visual networks, using object and motion stimuli designed to separately stimulate ventral and dorsal visual cortical networks.&#160; Four visual paradigms are presented: 1. Randomized visual objects with consistent temporal presentation. 2. Randomized visual objects with inconsistent temporal presentation (or jitter).&#160; 3. Visual motion via a radial field of coherent central dot motion without jitter.&#160; 4. Visual motion via a radial field of coherent central dot motion with jitter.&#160; These four paradigms are presented in a pseudo-randomized order for each participant.&#160; Jitter is introduced in order to view how possible anticipatory-related effects may affect the morphology of the object-onset and motion-onset CVEP response.&#160; EEG data analyses are described in detail, including steps of data exportation from and importation to signal processing platforms, bad channel identification&#160;and removal, artifact rejection, averaging, and categorization of average CVEP morphological pattern type based upon latency ranges of component peaks. Representative data show that the methodological approach is indeed sensitive in eliciting differential object-onset and motion-onset CVEP morphological patterns and may, therefore, be useful in addressing the larger research aim. Given the high temporal resolution of EEG and the possible application of high-density EEG in source localization analyses, this protocol is ideal for the investigation of distinct CVEP morphological patterns and the underlying neural mechanisms which generate these differential responses.&#160; &#160; &#160;&#160;

In this paper, we present a protocol to investigate differential cortical visual evoked potential morphological patterns through stimulation of ventral and dorsal networks using high-density EEG. Visual object and motion stimulus paradigms, with and without temporal jitter, are described. Visual evoked potential morphological analyses are also outlined.&#160;&#160;

Modèles morphologiques potentiels d'evoked Visual

This paper presents a methodology for the recording and analysis of cortical visual evoked potentials (CVEPs) in response to various visual stimuli using ...

Recording of Electroretinogram and Visual Evoked Potential in a Rat

Transcription

Recording of Electroretinogram and Visual Evoked Potential in a Rat