Mar 30, 2026
In Vivo Electrophysiology Protocol
- Egoa Ugarte Perez1,
- Silvia De Santis1
- 1Instituto de Neurociencias (Consejo Superior de Investigaciones Científicas - Universidad Miguel Hernández), San Juan de Alicante 03550, Spain
Protocol Citation: Egoa Ugarte Perez, Silvia De Santis 2026. In Vivo Electrophysiology Protocol . protocols.io https://dx.doi.org/10.17504/protocols.io.4r3l2dprjg1y/v1
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: March 13, 2026
Last Modified: March 30, 2026
Protocol Integer ID: 313219
Abstract
This protocol describes the procedure for performing in vivo electrophysiological stimulation and multichannel recordings in anesthetized rats.
Guidelines
This protocol was carried out using Wistar rats.
It is important to control the anesthesia according to body weight and to closely monitor the first 10 minutes after injection, as respiratory arrest may occur. This issue was observed particularly in aged rats, which may be more sensitive to urethane anesthesia.
Materials
-Cotton swabs
-Vaseline
-Viscotears
-Medical tape
-Craniotomy drill bit with the appropiate diameter required for the experiment
-Surgical drill to performing the craniotomy
-Fine surgical forceps
-Coarse surgical forceps
-Scalpel
-Surgical scissors with curved tip
-Four hemostatic forceps
-Pasteur pipette
-30G and 25G needles and syringes
-Electric heating pad
-High-intensity surgical lamp (preferably two) to provide adequate illumination of the surgical field
-High-resolution camera with optical magnification for visualization and documentation of the surgical field
-Fine-tip marker
-Small vessel cauterizer kit
-Three stereotaxic posts
Troubleshooting
Safety warnings
Improper grounding can introduce electrical noise into the recording system. Ensure that all grounding cables are correctly connected before starting the experiment.
Ethics statement
All experimental procedures were approved by the Animal Care and Use Committee of the Instituto de Neurociencias de Alicante (Spain) and were conducted in compliance with Spanish (Law 32/2007) and European regulations (EU Directive 86/609, EU Decree 2001-486, and EU Recommendation 2007/526/EC).
Before start
Prepare materials:
-Cotton swabs
-Vaseline
-Viscotears
-Medical tape
-Craniotomy drill bit with the appropiate diameter required for the experiment
-Surgical drill to performing the craniotomy
-Fine surgical forceps
-Coarse surgical forceps
-Scalpel
-Surgical scissors with curved tip
-Four hemostatic forceps
-Pasteur pipette
-30G and 25G needles and syringes
-Electric heating pad
-High-intensity surgical lamp (preferably two) to provide adequate illumination of the surgical field
-High-resolution camera with optical magnification for visualization and documentation of the surgical field
-Fine-tip marker
-Small vessel cauterizer kit
-Three stereotaxic posts
Anesthesia
Weigh the animal to determine the appropriate anesthetic dose. Prepare a solution of urethane diluted in PBS and administer it intraperitoneally (1.4 g/kg).
After the injection, place the animal on top of a heating pad to maintain body temperature during the induction of anesthesia.
Continuously observe the animal while anesthesia develops. Check for ear and paw reflexes to assess the level of anesthesia.
After 1 hour, reassess the reflexes. If reflexes are still present, administer an additional 1/5 of the initial urethane dose intraperitoneally.
Second supplemental dose (if necessary): Continue monitoring the animal. After another hour, if reflexes are still present, administer 1/10 of the original dose.
Repeat monitoring until surgical anesthesia is reached.
Surgery
Carefully transfer the animal to the surgical setup. Place the animal on an electric heating pad to maintain body temperature and insert a rectal probe to continuously monitor temperature at 36–37 °C.
Fix the head in the stereotaxic frame using the ear bars. Ensure that the head is stable and properly aligned.
Shave the hair from the cranial area before performing the incision.
Inject approximately 0.3 ml of lidocaine subcutaneously into the shaved scalp using a 30G needle to provide local anesthesia before making the incision.
Protect the eyes. Apply Viscotears to both eyes to prevent corneal drying and protect them from light exposure during the experiment.
Provide supplemental oxygen. Place an oxygen mask over the animal’s nose and mouth to deliver oxygen at approximately 0.8 L/min throughout the experiment.
Make a midline scalp incision.
Using a scalpel, make a straight incision along the midline of the skull from anterior to posterior.
Remove blood from the surgical area using cotton swabs moistened with PBS.
Continue cleaning until the area is as blood-free and dry as possible.
Use a small vessel cauterizer to stop bleeding in areas with excessive blood and to keep the surgical field clean. Important: Use cauterization only in terminal (non-recovery) experiments, as it can cause irreversible tissue damage.
Use four hemostatic forceps to hold back the tissue and keep the surgical field as open as possible, exposing the maximum area of the skull for the procedure.
Attach a stereotaxic post with its holder and a 22G needle, and use a fine-tip marker to mark the coordinates of the ROIs on the exposed skull, according to the anteroposterior (AP) and mediolateral (ML) coordinates relative to Bregma.
Perform the craniotomy using a manual or electric drill.
Exercise caution to avoid damaging the underlying brain tissue, especially in regions close to the venous sinus.
Once the small piece of skull is removed, place a cotton swab moistened with PBS over the exposed area to keep it continuously wet.
Carefully pierce and lift the dura mater, the outermost protective layer of the brain, using a 30G needle with a slightly bent tip, taking extreme caution to avoid damaging the underlying brain tissue. This step allows for subsequent insertion of the electrode into the brain without breaking it, as the electrode is especially fragile.
Electrophysiological recordings
Set up the stimulation and recording system.
Begin by arranging the three stereotaxic posts:
- One post with its holder will be used to secure the stimulation electrode.
- The other two posts, each equipped with a pre-amplifier, will be used to hold and connect the Neuronexus recording electrodes.
Ensure that all towers are stable and properly positioned to allow precise placement of the electrodes and minimal movement during the experiment.
Place all recording and stimulation electrodes in their respective holders, and connect them to the main amplifier and stimulator, respectively.
Lower the electrodes carefully.
Adjust the camera focus and zoom to clearly visualize electrode insertion.
Slowly and carefully lower the recording and stimulation electrodes while respecting the anteroposterior (AP), mediolateral (ML), and dorsoventral (DV) coordinates. Just before inserting the recording electrodes into the brain, connect the ground wire by attaching it to the animal tissue to ensure proper electrical grounding. Preferably, insert the stimulation electrode first, followed by the recording electrodes, with both the stimulator and amplifier turned off.
Maintain a layer of PBS over the entire surface of the skull to ensure proper electrical conduction throughout the experiment.
Turn on the Multichannel Systems stimulator and recording amplifier, and launch the MC_Rack and MC_Stimulus software to proceed with stimulation and recording according to the experimental protocol. Record for the required duration.
After completing the recording, carefully remove the electrodes and disassemble the surgical setup. Gently rinse the electrodes with distilled water, avoiding direct contact of the water stream with the electrode tip, as this may damage it.
As urethane anesthesia is toxic and non-recoverable, the animal is subsequently subjected to terminal perfusion.
Protocol references
- Pérez-Cervera L, De Santis S, Marcos E, Ghorbanzad-Ghaziany Z, Trouvé-Carpena A, Selim MK, Pérez-Ramírez Ú, Pfarr S, Bach P, Halli P, Kiefer F, Moratal D, Kirsch P, Sommer WH, Canals S. Alcohol-induced damage to the fimbria/fornix reduces hippocampal-prefrontal cortex connection during early abstinence. Acta Neuropathol Commun. 2023 Jun 21;11(1):101. doi: 10.1186/s40478-023-01597-8. PMID: 37344865; PMCID: PMC10286362.