electroencephalographic recording of epileptic seizures in epilepsy-induced rats

Electroencephalographic Recording of Epileptic Seizures in Epilepsy-Induced Rats

Begin by switching on the amplifier positioned outside of the Faraday cage. Open the EEG software, and start the EEG acquisition, and observe the EEG signal produced by unconnected cables.
Next, connect the animal to the tethered EEG recording system by holding the animal&#39;s head between two stretched fingers of one hand, and screwing down the connectors to the electrode pedestal using the other free hand.
Set an amplification factor on each channel of the amplifier according to the electrode signal of a single animal so the EEG signal is in scale. Then, let the animal explore the new cage for at least one hour under the direct observation of the researcher.
Adjust the cable according to the animal&#39;s commodity, and ensure that the cables do not interfere with the animal&#39;s movements and lying posture.

electroencephalographic-recording-epileptic-seizures-epilepsy-induced

Research

JoVE Encyclopedia of Experiments

Neuroscience

Neurophysiology

Stimulation and Modulation Techniques

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. Assembly of the Microdialysis Probe-electrode Device 
<ol>
	<li>Use a 3-channel two-twisted electrode (with at least a 20 mm cut length of the registering electrode and a 10 cm long grounding electrode) and couple it to a guide cannula to prepare the device. See examples of 3-channel electrodes and guide cannula for dialysis in Figure 1A-1B.</li>
	<li>Remove (Figure 1C) and insert (Figure 1D) the metal guide cannula into the dummy plastic cannula a few times prior to the usage in order to ease its removal at the moment of its switch for microdialysis probe in animal.</li>
	<li>Bend the twisted wires of registering electrode two times (Figure 1E-1F) in order to align the wires with the dummy cannula of the guide and cut the electrode tip (Figure 1G) to be 0.5 mm longer than the tip of the guide cannula (Figure 1H) using the digital caliper.</li>
	<li>Have ready the 1 mm long silicon circlet (outer diameter [O.D.] 2 mm, thickness 0.3 mm; Figure 1I) and insert the tip of the guide cannula and the tip of the twisted electrodes into the silicon circlet using the tweezers (Figure 1J). Fix it onto the foot of the guide cannula pedestal with polymer glue of rapid action or resin (Figure 1K). See the example of the completed devices in Figure 1L and Figure 2A.</li>
	<li>Sterilize the device under germicidal ultraviolet (UV) light for 4 h. Turn over the device four times so as each of its sides is exposed to the light for 1 h. 
	Note: Many home-made electrodes and microdialysis probes may be assembled in a similar way. The head of the above-described implant for rats has the following dimensions: 7 mm width x 5 mm depth x 10 mm length from the top to pedestal toe; the implant tip is about 11 mm long, 600 &#181;m in diameter and all the device weights about 330-360 mg. The device may be reused two or three times if (i) a sufficient length of the ground electrode is left on the skull during the surgery for the next use and (ii) when the animal is killed, and the device recovered together with the dental cement it is left in acetone overnight, such that the cement may be mechanically disaggregated, and the device washed and sterilized again.</li>
</ol>
2. Stereotaxic Surgery 
<ol>
	<li>Use a stereotaxic apparatus and probe clip holder (Figure 2B) for the device implantation following the contemporary standards for aseptic and pain-free surgeries.
	<ol>
		<li>Anesthetize the adult Sprague-Dawley rats with ketamine/xylazine mixture (43 mg/kg and 7 mg/kg, intraperitoneally [i.p.]) and fix it onto the stereotaxic frame. Add isoflurane anesthesia (1.4% in air; 1.2 mL/min) to initial ketamine/xylazine injection as it allows to control the depth of anesthesia in time. Shave the fur on the animal&#39;s head.</li>
	</ol>
	</li>
	<li>Swab the head skin surface by iodine-based solution followed by 70% ethanol to prepare it for aseptic surgery.
	<ol>
		<li>Implant the guide cannula-electrode device prepared in precedence (1.1 &#8211; 1.5) into the right ventral hippocampus using the following coordinates: nose bar + 5.0 mm, A &#8211; 3.4 mm, L + 4.5 mm, P + 6.5 mm to bregma. Follow standard techniques for stereotaxic surgeries.</li>
		<li>Ensure that it does not cover the anchoring screws. When mounting it onto the stereotaxic apparatus, grasp the device for the guide cannula head as this may be easily aligned to the probe holder.</li>
	</ol>
	</li>
	<li>Anchor the device to the skull with at least four stainless screws screwing them into the skull bone (1 screw into the left and 1 screw into the right frontal bone plates, 1 screw into the left parietal and 1 screw into the interparietal bone plates). Add a drop of tissue glue to further fix each screw to the skull bone.
	<ol>
		<li>Cover half of screw threads with methacrylic cement. Promote the binding of the cement by making shallow grooves in the bone to increase the adherence.</li>
	</ol>
	</li>
	<li>Once the tip of the device is positioned into the brain tissue, twist the wire of the ground electrode around 3 anchoring screws. Cover all mounted screws and the device with dental cement.</li>
	<li>Monitor the animals during the surgery and for about 1 h thereafter until they are upright and moving around the cage. Keep them on a warming pad to avoid hypothermia. Allow the rats to recover for at least 7 days after the device implantation.</li>
	<li>Monitor the animals at least once daily for 3 days after the surgery for signs of pain or distress. Give the animals with the antibiotic cream (gentamycin 0.1%) close to the incised site to prevent the infection and an analgesic (tramadol 5 mg/kg, i.p.) for 3 days to prevent the postsurgical pain.</li>
</ol>
3. Temporal Lobe Epilepsy Induction by Pilocarpine and Assignment of Animals to Experimental Groups 
<ol>
	<li>After a week of post-surgical recovery, assign the animals randomly to groups: (i) control animals receiving vehicle and (ii) epileptic animals that will receive pilocarpine. Use a proportionally higher number of animals for epileptic group since not all of the pilocarpine administered rats will develop the disease.</li>
	<li>Inject a dose of methylscopolamine (1 mg/kg, subcutaneously [s.c.]) and 30 min after, a single injection of pilocarpine (350 mg/kg, i.p.) to induce the status epilepticus (SE). Inject methylscopolamine and the vehicle (saline) to the control rats. Use 1 mL syringe with 25G needle for all i.p. administrations.
	<ol>
		<li>Check visually the animals to start to have behavioral seizures (moving vibrissa within 5 min, nodding head, cloning the limbs) and within 25 min to clone continuously all the body (SE).</li>
	</ol>
	</li>
	<li>Arrest the SE 2 h after the onset to have a mortality of about 25% and a mean latent period of approximately 10 days by administration of diazepam (20 mg/kg, i.p.). Observe and record any seizure behavior beginning immediately after the pilocarpine injection and continue for at least 6 h thereafter.</li>
	<li>Give the animals saline (1 mL, i.p.) using 1 mL syringe with 25G needle and sucrose solution (1 mL, p.o.) using 1 mL syringe and flexible feeding 17G needle for 2-3 days after SE to promote the recovery of body weight loss.
	<ol>
		<li>Exclude the animals that do not achieve the initial body weight within the first week after pilocarpine SE from the study (except for the acute group killed 24 h after SE, where the body weight follow up is not possible).</li>
	</ol>
	</li>
	<li>Assign post-SE animals randomly to different experimental groups (Figure 3): acute phase (where the microdialysis takes place 24 h after SE), latency (7-9 days after SE), first spontaneous seizure (approximately 11 days after SE), and chronic period (starts about 22-24 days after SE, i.e. about 10 days after the first seizure). Monitor the animals for the occurrence of spontaneous seizures. 
	NOTE: Use the following inclusion/exclusion criteria for further experiments in epileptic rats: development of convulsive SE within 1 h after pilocarpine administration; weight gain in the first week after SE and the correct positioning of the microdialysis probe and electrode.</li>
</ol>
4. Epileptic Behavior Monitoring and Analysis 
<ol>
	<li>Long-term monitoring of epileptic behavior
	<ol>
		<li>Approximately 6 h after pilocarpine administration (i.e., at the end of direct observation by the researchers), place the animals into the clean home cages and start the 24 h video monitoring.</li>
		<li>Continue the 24 h video monitoring until day 5, using a digital video surveillance system.</li>
		<li>Beginning at day 5, connect the rats in their home rectangular cages to tethered EEG recording system and continue the 24 h video monitoring.</li>
		<li>Set the parameters on the amplifier positioned outside of the Faraday cage (set amplification factor on each channel according to the specificity of the EEG signal of each single animal) and start the EEG acquisition observing the EEG signal produced by unconnected cables. Use a sampling rate of 200 Hz and low pass filter set to 0.5 Hz.</li>
		<li>Connect the animal to cables holding an animal&#39;s head between two stretched fingers of one hand and screwing down the connectors to the electrode pedestal using the other free hand. Start the acquisition. 
		CAUTION: Ensure that the signal is free of artifacts. Common artifacts are spikes greatly exceeding the scale.</li>
		<li>A day before the microdialysis experiment, transfer the animals into the tethered EEG system equipped with plexiglass cylinders for microdialysis. Disconnect the animals from the EEG tethering system in home cage screwing up the connectors from the electrode without restrain the animal. Place the animal into the high plexiglass cylinders.</li>
	</ol>
	</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>3-channel two-twisted electrode</td>
			<td>Invivo1, Plastic One, Roanoke, Virginia, USA</td>
			<td>MS333/3-B/SPC</td>
			<td>Material</td>
		</tr>
		<tr>
			<td>guide cannula</td>
			<td>Agn Tho&#39;s, Lindig&#246;, Sweden</td>
			<td>MAB 4.15.IC</td>
			<td>Material</td>
		</tr>
		<tr>
			<td>Resin KK2 Plastik</td>
			<td>Elettra Sport, Lecco, Italy</td>
			<td>KK2</td>
			<td>Material</td>
		</tr>
		<tr>
			<td>Super Attack gel Loctite</td>
			<td>Henkel Italia Srl, Milano, Italy&#160;</td>
			<td>2047420_71941</td>
			<td>Material</td>
		</tr>
		<tr>
			<td>Imalgene-Ketamine</td>
			<td>Merial, Toulouse, France</td>
			<td>221300288 (AIC)</td>
			<td>Solution</td>
		</tr>
		<tr>
			<td>Xylazine</td>
			<td>Sigma, Milano, Italy</td>
			<td>X1251</td>
			<td>Material</td>
		</tr>
		<tr>
			<td>Isoflurane-Vet</td>
			<td>Merial, Toulouse, France</td>
			<td>103120022 (AIC)</td>
			<td>Solution</td>
		</tr>
		<tr>
			<td>Altadol 50 mg/ ml - tramadol</td>
			<td>Formevet, Milano, Italy</td>
			<td>103703017 (AIC)</td>
			<td>Solution</td>
		</tr>
		<tr>
			<td>Gentalyn 0.1% crm - gentamycine</td>
			<td>MSD Italia, Roma, Italy</td>
			<td>20891077 (AIC)</td>
			<td>Material</td>
		</tr>
		<tr>
			<td>simplex rapid dental cement</td>
			<td>Kemdent, Associated Dental Products Ltd, Swindon, United Kingdom</td>
			<td>ACR811</td>
			<td>Material</td>
		</tr>
		<tr>
			<td>GlasIonomer CX-Plus Cement</td>
			<td>Shofu, Kyoto, Japan</td>
			<td>PN1167</td>
			<td>Material</td>
		</tr>
		<tr>
			<td>probe clip holder</td>
			<td>Agn Tho&#39;s, Lindig&#246;, Sweden</td>
			<td>p/n 100 5001</td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>Histoacryl&#174; Blue Topical Skin Adhesive</td>
			<td>TissueSeal, Ann Arbor, Michigan, USA</td>
			<td>TS1050044FP</td>
			<td>Material</td>
		</tr>
		<tr>
			<td>Valium 10 mg/2 ml - diazepam</td>
			<td>Roche, Monza, Italy</td>
			<td>019995063 (AIC)</td>
			<td>Material</td>
		</tr>
		<tr>
			<td>1 mL syringe with 25G needle</td>
			<td>Vetrotecnica, Padova, Italy</td>
			<td>11.3500.05</td>
			<td>Material</td>
		</tr>
		<tr>
			<td>rat flexible feeding needle 17G</td>
			<td>Agn Tho&#39;s, Lindig&#246; Sweden</td>
			<td>7206</td>
			<td>Material</td>
		</tr>
		<tr>
			<td>Grass Technology apparatus</td>
			<td>Grass Technologies, Natus Neurology Incorporated, Pleasanton, California, USA</td>
			<td>M665G08</td>
			<td>Equipment (AS40 amplifier, head box, interconnecting cables, telefactor model RPSA S40)</td>
		</tr>
		<tr>
			<td>modular data acquisition and analysis system MP150</td>
			<td>Biopac, Goleta, California, USA</td>
			<td>MP150WSW</td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>digital video surveillance system</td>
			<td>AverMedia Technologies, Fremont, California, USA</td>
			<td>V4.7.0041FD</td>
			<td>Equipment</td>
		</tr>
	</tbody>
</table>

Source: Soukupov&#225;, M., et al. <a href="https://app.jove.com/v/58455">Microdialysis of Excitatory Amino Acids During EEG Recordings in Freely Moving Rats.</a> J. Vis. Exp. (2018)
This video demonstrates electroencephalographic (EEG) recording of epileptic seizures in epilepsy-induced rats. A rat with electrodes implanted in the ventral hippocampus is connected to the EEG recording system and placed in a plexiglass chamber. The rat is allowed to move freely while the brain activity is recorded to monitor the occurrence of seizures.

<img alt="Figure 1" src="/files/ftp_upload/22861/22861_Figure1.jpg" /> 
Figure 1. Step-by-step preparation of the device to be implanted. (A) 3-channel electrode with 10 cm long grounding electrode in its protective sleeve on the left and guide cannula for microdialysis on the right needed to assemble the device. (B). The bare electrode and guide cannula in detail. The first step is to remove (C) and insert (D) the metal guide cannula from and into its plastic dummy ...

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

Take an epilepsy-induced rat with an electrode implanted in the skull. The tip of the electrode is inserted into the ventral hippocampus region.
Connect an electroencephalography, or EEG, recording system to the implanted electrode.
Transfer the rat to a plexiglass cylinder to record brain activity while the animal moves freely.
Epilepsy induction results in an increased release of excitatory neurotransmitters in the synapse of neurons while the release of inhibitory neurotransmitters decreases, disrupting the equilibrium between excitatory and inhibitory signals.
The increased excitatory neurotransmitters trigger a continuous influx of positive ions, making the neurons hyperexcitable. This causes a continuous generation of electrical signals.
A burst of synchronous electrical activity by neighboring neurons at a high frequency is termed an epileptic seizure.
Using the implanted electrode, record the abnormal burst of brain activity to identify the occurrence of seizures in the epilepsy-induced rat.

36 次观看. 癫痫诱发大鼠癫痫发作的脑电图记录

视频: 癫痫诱发大鼠癫痫发作的脑电图记录 - 实验

Simultaneous Video-EEG-ECG Monitoring to Identify Neurocardiac Dysfunction in Mouse Models of Epilepsy

In epilepsy, seizures can evoke cardiac rhythm disturbances such as heart rate changes, conduction blocks, asystoles, and arrhythmias, which can potentially increase risk of sudden unexpected death in epilepsy (SUDEP). Electroencephalography (EEG) and electrocardiography (ECG) are widely used clinical diagnostic tools to monitor for abnormal brain and cardiac rhythms in patients. Here, a technique to simultaneously record video, EEG, and ECG in mice to measure behavior, brain, and cardiac activities, respectively, is described. The technique described herein utilizes a tethered (i.e., wired) recording configuration in which the implanted electrode on the head of the mouse is hard-wired to the recording equipment. Compared to wireless telemetry recording systems, the tethered arrangement possesses several technical advantages such as a greater possible number of channels for recording EEG or other biopotentials; lower electrode costs; and greater frequency bandwidth (i.e., sampling rate) of recordings. The basics of this technique can also be easily modified to accommodate recording other biosignals, such as electromyography (EMG) or plethysmography for assessment of muscle and respiratory activity, respectively. In addition to describing how to perform the EEG-ECG recordings, we also detail methods to quantify the resulting data for seizures, EEG spectral power, cardiac function, and heart rate variability, which we demonstrate in an example experiment using a mouse with epilepsy due to Kcna1 gene deletion. Video-EEG-ECG monitoring in mouse models of epilepsy or other neurological disease provides a powerful tool to identify dysfunction at the level of the brain, heart, or brain-heart interactions.

Here, we present a protocol to record brain and heart bio signals in mice using simultaneous video, electroencephalography (EEG), and electrocardiography (ECG). We also describe methods to analyze the resulting EEG-ECG recordings for seizures, EEG spectral power, cardiac function, and heart rate variability.

同步视频脑电图-心电图监测对小鼠癫痫模型 Neurocardiac 功能障碍的鉴别

In epilepsy, seizures can evoke cardiac rhythm disturbances such as heart rate changes, conduction blocks, asystoles, and arrhythmias, which can ...

科研

JoVE Journal

遗传学

Recording EEG in Freely Moving Neonatal Rats Using a Novel Method

EEG is a useful method to detect electrical activity in the brain. Moreover, it is a widely used diagnostic tool for various neurological conditions, such as epilepsy and neurodegenerative disorders. However, it is technically difficult to obtain EEG recordings in neonates as it requires specialized handling and great care. Here, we present a novel method to record EEG in neonatal rat pups (P8-P15). We designed a simple and reliable electrode using computer pin loci; it can be easily implanted into the skull of a rat pup to record high-quality EEG signals in the normal and epileptic brain. Pups were given an intraperitoneal (i.p.) injection of the neurotoxin kainic acid (KA) to induce epileptic seizures. The surgical implantation performed in this procedure is less expensive than other EEG procedures for neonates. This method allows one to record high-quality and stable EEG signals for more than 1 week. Furthermore, this procedure can also be applied to adult rats and mice to study epilepsy or other neurological disorders.

Here, we introduce a novel technique designed to record electroencephalography (EEG) in freely moving neonatal epileptic pups and describe its procedures, features, and applications. This method allows one to record EEG for more than 1 week.

使用新颖的方法在自由移动的新生大鼠中记录脑电图

EEG is a useful method to detect electrical activity in the brain. Moreover, it is a widely used diagnostic tool for various neurological conditions, such ...

神经科学

A Behavioral Screen for Heat-Induced Seizures in Mouse Models of Epilepsy

Transgenic mouse models have proved to be powerful tools in studying various aspects of human neurological disorders, including epilepsy. The SCN1A-associated genetic epilepsies comprise a wide spectrum of seizure disorders with incomplete penetrance and clinical variability. SCN1A mutations can result in a large variety of seizure phenotype ranging from simple, self-limited fever-associated febrile seizures (FS), moderate-level genetic epilepsy with febrile seizures plus (GEFS+) to more severe Dravet Syndrome (DS). Although FS are commonly seen in children below 6-7 years of age who do not have genetic epilepsy, FS in GEFS+ patients continue to occur into adulthood. Traditionally, experimental FS have been induced in mice by exposing the animal to a stream of dry air or heating lamps, and the rate of change in body temperature is often not well controlled. Here, we describe a custom-built heating chamber, with a plexiglass front, that is fitted with a digital temperature controller and a heater-equipped electric fan, which can send heated forced air into the test arena in a temperature-controlled manner. The body temperature of a mouse placed in the chamber, monitored through a rectal probe, can be increased to 40-42 &#176;C in a reproducible manner by increasing the temperature inside the chamber. Continual visual monitoring of the animals during the heating period demonstrates induction of heat-induced seizures in mice carrying an FS mutation at a body temperature that does not elicit behavioral seizures in wild-type litter mates. Animals can be easily removed from the chamber and placed on a cooling pad to rapidly return body temperature to normal. This method provides for a simple, rapid, and reproducible screening protocol for the occurrence of heat-induced seizures in epilepsy mouse models.

The goal of the method is to screen for hyperthermia or heat-induced seizures in mouse models. The protocol describes the use of a custom-built chamber with continuous monitoring of the body temperature to determine whether elevated body temperature leads to seizures.

Electroencephalographic Recording of Epileptic Seizures in Epilepsy-Induced Rats

成績單

Electroencephalographic Recording of Epileptic Seizures in Epilepsy-Induced Rats