Electroencephalographic Recording of Epileptic Seizures in Epilepsy-Induced Rats

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

1. 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.
2. 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.
3. 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.
4. 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.
5. 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 µm 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.

2. Stereotaxic Surgery

1. Use a stereotaxic apparatus and probe clip holder (Figure 2B) for the device implantation following the contemporary standards for aseptic and pain-free surgeries.
   1. 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's head.
2. Swab the head skin surface by iodine-based solution followed by 70% ethanol to prepare it for aseptic surgery.
   1. Implant the guide cannula-electrode device prepared in precedence (1.1 – 1.5) into the right ventral hippocampus using the following coordinates: nose bar + 5.0 mm, A – 3.4 mm, L + 4.5 mm, P + 6.5 mm to bregma. Follow standard techniques for stereotaxic surgeries.
   2. 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.
3. 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.
   1. 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.
4. 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.
5. 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.
6. 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.

3. Temporal Lobe Epilepsy Induction by Pilocarpine and Assignment of Animals to Experimental Groups

1. 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.
2. 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.
   1. 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).
3. 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.
4. 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.
   1. 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).
5. 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.

4. Epileptic Behavior Monitoring and Analysis

1. Long-term monitoring of epileptic behavior
   1. 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.
   2. Continue the 24 h video monitoring until day 5, using a digital video surveillance system.
   3. Beginning at day 5, connect the rats in their home rectangular cages to tethered EEG recording system and continue the 24 h video monitoring.
   4. 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.
   5. Connect the animal to cables holding an animal'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.
   6. 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.

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 few times to ease its release once implanted into the animal's head. The second step is to bend two times the twisted registering electrode to be aligned to dialysis guide cannula (E, F). (G) The electrode tip should be cut to be 0.5 mm longer than the tip of the metal guide cannula. (H) Check for the precision of the cut using the digital caliper. Subsequently, about 1 mm long silicon circlet should be used to fix the alignment of electrode to guide cannula foot (I). (J) The photograph showing how to ring the electrode and guide cannula shaft. The final step is to put a drop of resin or glue onto the guide cannula pedestal fixing the silicon circlet to it (K). (L) Assembled device ready to be sterilized.

Figure 2. Photographs of different types of devices for microdialysis-EEG in rats used (A) and the photograph of the probe clip holder (B) used to implant these devices. (A) The guide cannula (in green) is replaced by a microdialysis cannula typically 24 h before the experiment. The electrical connector of the device (first left was used for the recordings described in this manuscript) permits the attachment of wires that conduct electrical signal to amplifier and data collection equipment. The device is surgically attached to the skull of anesthetized rats and recordings may be obtained later without causing pain or discomfort in freely behaving rats.

Figure 3. Experimental design. The week before status epilepticus (SE) induction, the rats are implanted with the device. SE is induced by pilocarpine and animals (if not dialyzed and killed at 24 h after SE; the rats from acute group) are video monitored for 5 days (blue line), then video-EEG monitored to assess the seizure frequency and duration in their home cages (green line). For the microdialysis experiment, the epileptic and respective non-epileptic control rats are transferred to another EEG set up equipped with cylinder cages in 24 h before the microdialysis session and still video-EEG monitored (light green line). The vertical red lines represent the dialysis sessions at different timepoints of epileptic disease development. The horizontal red lines represent the different groups of epileptic animals (and respective non-epileptic animals), where the arrow indicates the last day of the microdialysis and the day of animal's death.