Multiscale Investigations of Cortical Processing by Integrating Laminar Polytrodes and Optogenetics with Micro Electrocorticography in Rodents

Summary

Here, we present two protocols for high-density micro-electrocorticography (µEcoG) recording in rats and mice, including surgical, implantation, and recording methods. µECoG recordings are performed in combination with either laminar polytrode recording in the rat auditory cortex or with optogenetic manipulation of neural activity in the mouse somatosensory cortex.

Abstract

Electrocorticography (ECoG) is a methodological bridge between basic neuroscience and understanding human brain function in health and disease. ECoG records neurophysiological signals directly from the cortical surface at millisecond temporal resolution and columnar spatial resolution over large regions of cortical tissue simultaneously, making it uniquely positioned to study both local and distributed cortical computations. Here, we describe the design of custom, high-density micro-ECoG (µECoG) devices and their use in two procedures. These grids have 128 low-impedance electrodes with 200 µm spacing fabricated on a clear polymer substrate with perforations between electrodes; these features enable simultaneous µECoG recording with laminar polytrode recordings and optogenetic manipulations. First, we present a protocol for combined epidural µECoG recording over the whisker somatosensory cortex of mice with optogenetic manipulation of specific genetically defined cortical cell types. This allows causal dissection of the distinct contributions of different neuronal populations to sensory processing while also monitoring their specific signatures in µECoG signals. Second, we present a protocol for acute experiments to record neural activity from the rat auditory cortex using µECoG grids and laminar polytrodes. This allows detailed topographic mapping of sensory-evoked neural responses across the cortical surface simultaneously with recordings from multiple neural units distributed across the cortical depth. These protocols enable experiments that characterize distributed cortical activity and may contribute to understanding and eventual interventions for diverse neurological disorders.

Introduction

Brain functions underlying sensation, cognition, and action are organized and distributed across vast spatial and temporal scales, ranging from the spikes of single neurons to the electrical fields generated by populations of neurons in a cortical column to the topographic organization of columns across brain areas (e.g., somatotopy in somatosensory cortex, tonotopy in primary auditory cortex). Understanding brain function requires sensing electrical signals across these spatial scales¹. Neuroscience currently has many widely used methods for monitoring the activity of the brain. Electrophysiologically, laminar polytrodes (such as Neuropixels) enable monitoring of a modest number (~300) of single neurons, typically within a handful of distantly spaced columns, with high (≥1 kHz) temporal resolution. Ca²⁺ imaging enables monitoring of modest to large numbers of genetically and anatomically identified single neurons within ~1-2 mm spatial extent at a lower (~10 Hz) temporal resolution². fMRI enables monitoring the metabolic state of large numbers of neurons (~1 M neurons in a 36 mm³ volume) across the entire brain at very low (~0.2 Hz) temporal resolution. EEG/MEG enables monitoring of electrical activity from the whole cortical surface/brain at modest temporal resolution (<100 Hz) and very low spatial resolution (centimeters)³. While each of these methodologies has provided fundamental, synergistic insights into brain function, methods that enable direct sensing of electrophysiological signals at high temporal resolution from precise anatomical locations across broad spatial regions of the cortex are nascent. The need for broad spatial coverage is emphasized by the fact that in the brain, neuronal function changes much more dramatically across the surface compared to the depth⁴.

Electrocorticography (ECoG) is a method in which grids of low-impedance electrodes are implanted onto the surface of the brain and allow for recording or stimulation of the cortex^1,5. ECoG is typically deployed in human neurosurgical settings as part of the clinical work-up for treating pharmacologically intractable epilepsy. However, it also provides unique insights into distributed cortical processing in humans, such as speech and sensory topographic mapping^6,7. These capabilities have motivated its use in animal models, including monkeys, rats, and mice^5,8,9,10,11. In rodents, it has recently been shown that micro-ECoG (µECoG) enables high temporal resolution (~100 Hz) direct electrical monitoring of neuronal populations with columnar spatial resolution (~200 µm) and broad spatial coverage (many millimeters). µECoG enables researchers to investigate distributed neural dynamics associated with complex sensory processing, cognitive functions, and motor behaviors in animal models^12,13. Recent advances have integrated µECoG with optogenetics and laminar polytrode recordings^{14,15,16,17,18,19,20}, allowing for multiscale investigations of cortical networks and bridging the gap between micro-scale neuronal activity and macro-scale cortical dynamics^21,22. Critically, because the µECoG signal is very similar in humans and non-human animal models, the use of µECoG makes translation of results and findings from animal models to humans much more direct²³. As such, integrative approaches are crucial for advancing our understanding of neural circuitry and hold promise for developing novel therapeutic interventions for neurological disorders^5,24,25.

Consequently, there is an emerging need for protocols that integrate high-density µECoG arrays with laminar recordings and optogenetic tools to enable comprehensive multiscale investigations of cortical processing^8,26. To address this gap, we have developed custom-designed µECoG devices featuring 128 low-impedance electrodes with 40 µm electrode diameter and 20 µm inter-electrode spacing on a flexible, transparent polymer substrate (parylene-C and polyimide) with perforations between electrodes, enabling simultaneous µECoG and laminar polytrode recordings with optogenetic manipulations^13,22. Key aspects of this experimental protocol include: (i) columnar spatial resolution and large-scale coverage of cortical activity through high-density µECoG arrays; (ii) the ability to record from multiple cortical layers using laminar polytrodes inserted through the µECoG grid; and (iii) the incorporation of optogenetic techniques to selectively activate or inhibit specific neuronal populations, thus enabling causal dissection of neural circuits^27,28,29. The high-density configuration allows for high spatial resolution recordings, effectively providing a "columnar view" of cortical activity, as previous studies have shown that µECoG signals can resolve activity at a spatial scale comparable to the diameter of the rodent cortical column (~20 µm)¹¹. This integrated methodology allows for simultaneous multiscale monitoring and manipulation of neural activity, potentially enabling causal experiments to determine the neuronal sources of µECoG signals as well as distributed cortical processing. To achieve these objectives, this manuscript provides detailed protocols for the use of high-density µECoG arrays in two combinations.

First, we describe µECoG combined with the manipulation of layer 5 (L5) pyramidal cells in the mouse primary somatosensory cortex (S1). In the mouse, the µECoG array is placed epidurally (due to the surgical intractability of durotomy in mice). An optic fiber is positioned over the grid or combined with a lens to focus the optogenetic light over a small target area of the cortical surface. The optogenetic strategy is described here for inhibition of layer 5 excitatory neurons but can be readily adapted to any population of neurons provided with the corresponding, population-specific, Cre-expressing mouse line. Second, we describe the combined use of µECoG with silicon laminar polytrodes to simultaneously record cortical surface electrical potentials (CSEPs) and single-unit spiking activity from multiple neurons across cortical layers from rat auditory cortex (A1). The array has perforations between electrodes, enabling the insertion of multichannel laminar polytrodes through the grid to record neuronal activity across different cortical layers. During the craniotomy procedure, the µECoG array is placed subdurally over the auditory cortex, and the laminar polytrode is inserted through the perforations. Neural signals from the µECoG and laminar probe are recorded simultaneously, sampled at 6 kHz and 24 kHz, respectively, using an amplifier system optically connected to a digital signal processor.

Protocol

Both protocols follow the same key steps (anesthesia, fixation, craniotomy, µECoG recording) but have notable differences. In the following description, the shared steps are merged, while the specificities of each protocol are annotated. These steps below correspond to µECoG recording with optogenetics (Mouse) or µECoG recording with a laminar probe (Rat). All procedures described here were conducted in compliance with the local ethical or legal authorities (IACUC or Ethics Committees). Medications used may vary according to the approved ethical protocol.

1. Preparation and protocol for mouse and rat procedures

1. Notable differences between mouse and rat protocols
   1. Setups for surgery versus electrophysiological recordings
      1. For a rat, use the same setup during both surgery and electrophysiological recordings.
      2. For a mouse, perform the surgery in the first setup and electrophysiological recordings in the second setup.
   2. Head fixation
      1. For a rat, use the same snout clamp for surgery and electrophysiological recording.
      2. For a mouse, use a snout clamp for surgery and an external metal headbar in the setup for electrophysiology to allow fixation under light isoflurane anesthesia. Implant the headbar with dental cement on the skull.
   3. Recording setup: Use distinct acquisition electronics, recording software, and sensory stimulation software for the two species.
      1. For the mouse protocol, utilize the SpikeGadgets system (https://spikegadgets.com) and the open-source Trodes software (https://spikegadgets.com/trodes/) for data acquisition.
      2. For the rat protocol, utilize the recording software Synapse - Neurophysiology Suite (https://www.tdt.com/component/synapse-software/) for data acquisition.
   4. Induce anesthesia by injection (Rat) or inhalation (Mouse).
   5. Recording location
      1. Conduct recordings in the somatosensory cortex (S1) for a mouse.
      2. Conduct recordings in the primary auditory cortex (A1) for a rat.
         NOTE: This difference in anatomical localization requires different craniotomy sites for each species.
2. Preparing and testing the grid
   1. Soak the grid (excluding the connector board) in a diluted enzymatic detergent (50% detergent, 50% distilled water) for at least 1 h.
   2. Transfer it to a bath of pure distilled water and allow it to air-dry in a safe, clean location.
   3. Perform platinum black electrodeposition as part of the initial preparation of the µECoG device, not before every recording session.
      NOTE: Once deposited, the Platinum Black coating forms a stable layer that remains effective for multiple recordings, though its performance should be monitored through regular impedance testing. Platinum Black electrodeposition (target range of 10-20 kΩ at 1 kHz.) decreases electrode impedance and improves signal-to-noise ratio in neural recordings.
   4. To perform the electrodeposition, prepare a chloroplatinic acid solution (typically 1-3% Chloroplatinic acid [H2PtCl6]) containing a small amount of lead acetate (around 0.005%) as a deposition modifier. Connect the µECoG electrodes to serve as the working electrode in a three-electrode electrochemical cell, with a platinum counter electrode and Ag/AgCl reference electrode.
   5. Apply a constant current density of approximately -0.5--2 mA/cm² for 10-30 s while monitoring impedance values.
   6. Test and record the impedance of grid electrodes (e.g., with a Nano-Z).
   7. Check the grid over a light source and prepare the grid for use in post-surgery recording.
3. Reference cable soldering
   1. For a mouse, solder the end of a silver wire (10 mm long, 30 G) to a gold pin for connecting to the reference lead of the recording system.

2. Surgery

1. Preparation of materials and general monitoring (Animal care and recording)
   1. Preparation: Clean and disinfect the surgical area thoroughly with an appropriate disinfectant. Ensure all surgical instruments are sterilized, typically using an autoclave.
   2. Surgical tool placement: Arrange surgical tools on the sterile surgical pad. Stock the surgical area with cotton-tip applicators and absorbent cotton surgical triangles. Dispose of any biohazardous waste in a dedicated disposal bag.
   3. Temperature regulation: Turn on the heating pad within the surgical and electrophysiological recording site. Control the heating pad's temperature throughout the surgery and recording.
   4. Surgical pad: Place a blue surgical pad or blanket over the temperature-regulation bed.
      NOTE: This pad should feature a soft cotton white bottom, which should face upwards.
   5. Microscope positioning: Prepare the microscope and the attached illuminator (e.g., LED ring) off to one side of the surgical area. Check that it is functioning properly.
   6. Surgical drill: Prepare the surgical drill for the craniotomy procedure.
   7. Oxygen supply: Set the oxygen tank flow rate to 1.0 L/min (Rat) or 0.5 L/min (Mouse) and place the oxygen mask near the regulation pad. The animal will need continuous oxygen following anesthesia.
   8. Fluid replacement: Throughout the surgery, replace any fluid loss-aiming for a replacement of at least 1.5% of the animal's body weight (Mouse) or 1 mL per h (Rat). Prepare isotonic solutions accordingly.
   9. Animal: Bring the animal from the animal facility to the surgery room according to approved procedures. Use mice aged 8 to 16 weeks, either male or female, of C57Bl6 background; likewise, use rats aged 7 weeks old, males, of the Sprague Dawley strain.
   10. Preparation of medications: Weigh the animal using a scale with 0.1 g precision. Prepare the adequate drug quantities for the surgery, using pre-diluted solutions if necessary.
2. Anesthesia induction
   1. Anesthesia induction for a mouse
      1. Place the animal in the isoflurane induction chamber (3-5% Isoflurane in 0.5 L/min O₂).
      2. Once deep anesthesia is confirmed (absence of reflex in tail/toe pinch), place the animal on the surgery heating pad and headfix it.
      3. Inject subcutaneous drugs for general analgesia: Meloxicam: 5 mg/kg and Buprenorphine: 0.1 mg/kg.
      4. Headfix the mouse following steps 2.2.1.5-2.2.1.6.
      5. Place the snout in the anesthesia mask and the head loosely in the head mount. To place the mouse in the bite bar, first, ensure that the tongue is below the rod and not between the rod and the roof of the mouth. Use forceps to move the tongue if necessary.
      6. Insert the animal's incisors into the hole on the rod of the bite bar. Secure the mouse anesthesia mask (1.5-2% Isoflurane in 0.5 L/min O₂) by gently tightening the fixation screw. Head stabilization during surgery is ensured solely by the bite bar.
      7. Protect the animal's eyes with a petroleum-based eye ointment or lubricant to prevent drying during surgery.
      8. Maintain anesthesia throughout the procedure with a continuous flow of isoflurane through the mask.
   2. Anesthesia induction for a rat
      1. Use isoflurane initially to sedate the animal to ease the injection of induction anesthesia.
      2. Administer the drugs for anesthesia and analgesia:
         Meloxicam: Dosage of 5 mg/kg, concentration 10 mg/mL, 0.4 mL/kg
         Ketamine: Dosage of 90 mg/kg, concentration 100 mg/mL, 0.9 mL/kg
         ​Xylazine: Dosage of 10 mg/kg, concentration 100 mg/mL, 0.1 mL/kg
      3. Allow the animal to reach deep anesthesia within 15-30 min, depending on weight and age.
   3. Anesthesia monitoring
      1. Continuously monitor the animal's vitals (respiratory rate) throughout the procedure. Check the respiratory rate as a particularly useful sign of early changes in the anesthetic state, and adjust the anesthetic level if the respiratory rate changes.
      2. The paw withdrawal reflex is a critical sign of the anesthetic state. Test this reflex periodically, as its total absence ensures sufficient levels of anesthesia for surgery.
3. Head fixation and monitoring of vital signs
   1. Animal vitals monitoring
      1. Check and record the animal's vital signs on an experimental sheet. If the animal's reflexes (e.g., paw withdrawal) are not fully extinguished, administer an additional half-dose of supplementary ketamine (Rat) or increase the isoflurane concentration by increments of 0.5% (Mouse).
   2. Head fixation for a rat
      1. Once the rat is fully anesthetized (no paw or tail reflexes), insert the animal's incisors into the hole on the rod of the head mount.
      2. Carefully insert the points of the mounting arms into the ridge of the nose to fix the head during surgery, ensuring that it does not make contact with the eyes.
      3. Adjust the angle of the arms until the roof of the animal's mouth is firmly pressed against the rod. Ensure the skull remains immobile under pressure.
      4. Secure both arms of the mount by tightening the screws with a hex wrench.
   3. Oxygen setup for a rat
      1. Secure the plastic tubing from the oxygen tank over the animal's muzzle and nose, fastening it with surgical tape. Avoid creases in the tubing that may obstruct airflow. Set the oxygen tank to a flow rate of 1 L/min.
         NOTE: Animal vitals, including heart rate and respiratory rate, should be checked at 15-30 min intervals throughout the procedure.
4. Scalp incision
   1. Shaving and preparation
      1. Shave the area from the upper snout to the back of the head, extending from one eye to the other and around the ears. Remove the bulk of the fur with scissors or an electric clipper, and then apply depilatory cream.
   2. Disinfection
      1. Disinfect the area using a cotton swab soaked in Betadine, then rinse with a cotton swab soaked in 70% ethanol. Repeat this process three times and finish with a final Betadine application to ensure the area is sterile.
   3. Local anesthetic Injection
      1. Inject the local anesthetic Lidocaine (1%, 0.1 mL for mouse/0.4 mL per kg for rat) subcutaneously into the midline of the animal's scalp. Gently massage the scalp to spread the lidocaine, and wait for 5 min to allow the anesthetic to take effect.
   4. Incision
      1. For a mouse, lift a point on the skin with tweezers and resect a small section of skin (approximately 1 cm in diameter) using surgical scissors.
      2. For a rat, make a precise incision on the anterior side of the scalp, just above the nose, at the midline using the scalpel. Gently pull back the skin, creating a straight incision from between the eyes to the base of the skull. Carefully lift the scalp, cut away connective tissue, and expose the skull fully.
      3. Expose the craniotomy site following steps 2.4.4.4-2.4.4.5.
      4. With a scraper, clear away connective tissue and periosteum on the top of the skull. Flush saline and use aspiration or a surgical sponge to clean the site.
      5. Use surgical clips on the skin margins to facilitate clear exposure of the skull region where the craniotomy will be performed.
5. Craniotomy
   1. General drilling procedure
      1. Set surgical drill speed to a low setting of 5000 rpm or 7000 rpm for experienced surgeons. Do all drilling while visualizing through the microscope.
      2. Hold the drill parallel to the surface of the skull and rest gently against the surface.
      3. With light pressure on the pedal, begin drilling in a single location. Perform drilling in short intervals (5-10 s) with frequent checks for changes in bone color.
         NOTE: The bone will begin an opaque white, and as the hole becomes deeper, it will become more translucent, revealing a pink tint.
      4. When the drilling has gotten close to the brain, slow down and look for signs of moisture seeping into the hole. When the hole is dark pink and has a slight shine, stop drilling. Using a short 30 G needle, gently puncture the remaining layer of bone. Clear liquid should well out of the new hole.
   2. Drilling procedure for a mouse
      1. To place a reference electrode for physiological recordings, drill a burr hole in the frontal part of the hemisphere ipsilateral to the recorded area.
      2. Define the contour of the craniotomy by drilling a shallow trench on its perimeter. In the medio-lateral axis, start from the lateral bone ridge as a reference, and trace a 4 mm window.
      3. In the antero-posterior axis, drill a 3 mm window starting ~ 1mm anterior of the posterior bone ridge. The final open craniotomy size is approximately a 4 x 3 mm window.
   3. Drilling procedure for a rat
      1. Drill two holes: one in the left posterior quadrant, the other in the right anterior quadrant.
      2. Inject masseter muscle with a second dose of lidocaine (0.4 mL/kg at 10 mg/mL), and distribute evenly where the cut is to be made.
      3. Resect only the minimal set of muscles needed to expose the craniotomy area.
      4. Using a fresh #10 scalpel blade, create a transverse dorsal-ventral cut in the bundle of muscle above the animal's jaw (right side). Hold the posterior edge of the cut with gripping forceps and peel away from the skull while cutting along the bony ridge of the cheekbone. In this way, the muscle can be detached from the bone with minimal bleeding.
      5. Resect the anterior muscle in a similar way until a fissure line in the skull is revealed. This line will be the anterior boundary of the craniotomy window.
      6. Clear muscle from around the posterior ridge with the scalpel and forceps, using a strong light source to avoid cutting into highly vascularized regions.
      7. Grind down the posterior ridge using the drill until it is no longer raised above the skull's surface.
         NOTE: This step is essential for setting down the µECoG grid to make direct contact with the cortical surface.
      8. Drill the dorsal edge of the window just above the ridge where the resected muscle was attached. Place the posterior edge anterior to the drilled-down posterior ridge. Place the anterior edge posterior to the fissure line extending down near the eye socket.
         NOTE: When clearing the anterior muscle, care must be taken to avoid the eye.
6. Craniotomy window drilling
   1. Drilling the craniotomy window (Tips)
      1. When drilling, ensure that the drill bit is kept parallel to the skull surface. Apply as little force as possible, using the drill like a brush by allowing the drill to make light contact with the skull while using short, repetitive motions along the intended craniotomy line.
         NOTE: In rats, the posterior edge of the window has the thickest bone. If drilling too far back, the bone may exhibit a flaky, "crunchy" quality that complicates gauging drilling progress. If placed incorrectly, this bone area may reveal veiny red coloration that gives a false impression of proximity to the brain.
      2. Drill each side of the craniotomy window until the bone appears pale pink with a thin white fissure or crack running along its length. Apply light pressure; the bone should produce a distinct "wiggle" when fully drilled. If the crack appears disjointed, continue drilling lightly until achieving a continuous line.
   2. Removing the thinned skull within the craniotomy window
      1. When the skull has been thinned enough that extremely light pressure causes the whole window to wiggle visibly, remove the thinned skull.
      2. Flush the craniotomy site with a drop of saline and wait at least 1 min. This weakens the thinned bone and helps the bone to detach from the dura. Drain excess saline with an absorbent cotton triangle or vacuum.
      3. Carefully lift the thinned skull using forceps, avoiding damage to the underlying tissue.
      4. Use a hemostatic sponge to keep the brain moist.
      5. Tightly grip the window on the dorsal and ventral sides with toothed forceps and pull directly away from the skull. If there is any difficulty pulling the window out of the site, stop and resume light drilling until the bone is sufficiently weakened.
      6. For mouse µECoG recordings, leave the dura intact.
7. Cement and headpost implant for a mouse
   1. Insert and secure the reference wire.
      1. Insert the silver wire end ~1 mm into the burr hole, enough to contact the surface of the brain but not to cause bleeding.
      2. Apply the dental cement in place while applying the first layer.
   2. Preparation of dental cement
      1. Use a cooled ceramic mixing dish to prepare the dental cement mixture. This cement thickens quickly and requires the regular preparation of a new mixture. Wipe clean the mixing dish before preparing a new mixture.
         NOTE: The cement should never be in direct contact with the brain.
   3. Application of first layer
      1. Apply the first layer of cement around the craniotomy and across the entire skull using micro applicators. This layer acts as electrical insulation between the skull and the metal headbar.
      2. Completely surround the craniotomy with cement, including lateral coverage, to provide adequate protection for theopen craniotomy and µECoG grid.
   4. Positioning the metal headbar
      1. Attach the large section ofthe headbar to its holder without fully tightening it.
      2. Position the headbar as desired, laying the thin section along the skull midline in contact with the cement surface.
   5. Securing the implant
      1. Cover the headbar with dental cement and connect it to the cement surface.
   6. Removing the holder
      1. Allow a few minutes for the dental cement to strengthen.
      2. Once the headbar is fully secured, remove it by first removing the screw from the holder. Then, retract the holder backward, ensuring no force is applied to the headbar.
8. Durotomy for rat surgery
   NOTE: This is a challenging surgical step.
   1. Lifting the dura
      1. Using No. 5 forceps, held as parallel to the brain's surface as possible, lift a small portion of the dura away from the brain.
      2. Use a fresh 30 G needle (as short as possible) to carefully tear the lifted dura.
         NOTE: The dura mater is a thin, transparent layer of tissue that lies directly on top of the brain. It is removed for µECoG recordings in rats. It is critical to perform the durotomy without disturbing the vasculature on the brain's surface. Recommended methods for performing the durotomy include using forceps and a syringe needle to puncture the dura before pulling it back or using a duratome tool anchored near the skull to retract the dura carefully.
   2. Resection of the dura
      1. Continue gripping the dura with the forceps and lifting it away from the brain. Create a diagonal tear with the needle while lifting.
      2. Use the forceps to carefully peel the dura towards the sides of the craniotomy window, ensuring the brain surface remains undisturbed.
9. Transferring the mouse to the setup for electrophysiological recording
   1. Remove the animal from the surgery setup by gently lifting the snout and incisors from the incisor bar and then pulling back the animal. Inject Chlorprothixene (1 mg/kg, intraperitoneal [IP]), a sedative that enables continuous anesthesia to be maintained using a lower Isoflurane concentration.
   2. Place the mouse in the electrophysiological recording setup.
      1. Make sure that the heating pad is in place and working properly.
      2. Head fix the animal using the headbar on the holder in the electrophysiological setup.
      3. Bring the isoflurane mask close to fully cover the animal's snout.
   3. Anesthesia adjustment
      1. Gradually reduce anesthesia levels to 0.7-1% Isoflurane (in increments of 0.5% maximum every 5 min).
      2. Monitor the animal's respiratory rate and movements.
         NOTE: The respiratory rate should increase slightly compared to the surgical state, but the animal should not be moving.
      3. If the animal is moving, immediately increase the isoflurane concentration to 2% before slowly returning it to a lower level in increments of 0.5%.
   4. Inserting whiskers for sensory stimulation
      1. Attach the mouse's vibrissae to the whisker stimulation device. In this protocol, insert nine vibrissae into short 10 µL pipette tips, which are connected to piezoelectric actuators that provide rapid deflections of the vibrissae.

3. Recording

1. Installing the grid
   1. Preliminary steps
      1. Turn on the recording system and amplifier.
      2. Check the animal's vital signs.
   2. Procedure
      1. For positioning the animal and tools, follow steps 3.1.2.2-3.1.2.4.
      2. Place the animal in the recording setup, and ensure the craniotomy remains moist by regularly applying a saline solution.
      3. For a rat, position the micromanipulator on the rig's railing, located well behind the craniotomy site, to avoid interference.
      4. For a mouse, place the micromanipulator laterally to the craniotomy site alongside the animal.
      5. To attach and position the grid on a mouse, follow steps 3.1.2.6-3.1.2.12.
      6. Attach the µECoG grid to the headstage using the ZIF-clip connectors (headstage connector). Hold the headstage's electronic board in place via a mechanical bar fixed to a micromanipulator.
      7. Lower the µECoG grid horizontally to align flat over the craniotomy along the anteroposterior axis.
         NOTE: Along the lateral-medial axis, the grid's edge should be near the craniotomy's medial border.
      8. Once the grid is positioned near the brain but not in contact with it, attach the grid's reference wire to the implanted silver wire-gold pin. If required, attach the ground wire to the animal (e.g., to an uncovered muscle) to reduce electrical noise.
      9. Further, lower the grid to contact the brain.
      10. Move the grid laterally to "glide" over the moist dura surface. Continue adjusting until the grid is centered along the mediolateral axis.
      11. Use aspiration or a surgical sponge around the craniotomy's edges to remove any excess saline solution.
      12. Once the preparation is slightly drier, ensure the grid adheres more firmly to the dura and does not slide over its surface. When drier, apply a lateral to medial movement to the flexible grid, ensuring contact with the most lateral electrodes. The grid's flexible cable will naturally bend to match the brain's contour.
      13. To position the grid on a rat, follow steps 3.1.2.14-3.1.2.18.
      14. Secure the stem of the holding fork in the micromanipulator, ensuring that the connector board of the grid will hover over against the posterior side of the craniotomy window when lowered.
      15. Adjust the position of the micromanipulator on the railing so that the grid is roughly above the craniotomy site. Lower the grid until it hovers close above the brain surface. Moisten the brain surface with a small drop of saline.
      16. Conduct these steps using the microscope. Using the dials of the micromanipulator, adjust the grid position until it lies flat against the brain surface in the center of the craniotomy.
      17. Wick away moisture carefully using an absorbent cotton triangle without touching the grid itself. Ensure every row of the grid is in contact with the brain surface.
          NOTE: Removing moisture prevents the passive spread of the electrical signal through the fluid between the cortical surface and the grid, which spatially diffuses the signal sensed at the electrode.
      18. Using number 2 or number 5 forceps, insert the grid grounding wire into the same burr hole or insert the reference wire into a burr hole and the ground wire into nearby muscle tissue.
          NOTE: Wires should be inserted only ~1 mm, enough to contact the brain but not to cause bleeding or trauma to the brain.
2. Checking the positioning of the grid
   1. Monitoring electrophysiological activity
      1. Observe the electrophysiological activity using the recording software. Under light anesthesia, brain signals are variable and can exhibit a variety of patterns.
      2. Proper connection of the grid, reference, and ground wires should yield a high signal-to-noise ratio, with signal amplitudes in the range of mV. Monitor noise in the high-frequency range using bandpass filtering with Trodes (e.g., 100-6000 Hz) and ensure it does not exceed a few tens of microvolts (µV).
   2. Assess the sensory responsiveness using noise (e.g., clapping or snapping fingers) to induce visible event-related cortical surface electrical potentials (CSEPs).
      NOTE: Stimulation of a single whisker should evoke a clear, sharp event-related CSEP in only a few channels (Mouse).
   3. Grid position verification
      1. For a rat, confirm that the grid is correctly positioned over the auditory cortex. The first block recorded should typically be a 60-s white noise stimulus set to verify that the grid registers a proper response from the brain. Conduct white noise and tone diagnostic recordings with the grid only before inserting polytrode to help determine whether the grid was placed correctly and if there is a signal response.
      2. For a mouse, to verify grid positioning, perform a quick mapping session with 20-30 whisker deflections spaced by 350 ms. Record the activity in the local field potential (LFP) band using Trodes and analyze it offline with a custom MATLAB code to visualize the spatial extent of whisker-evoked activity.
   4. Repositioning
      1. If the grid requires adjustment, moisten the cortical surface with drops of saline over the grid.
      2. Leave the saline for 30 s to 1 min before attempting to lift the grid.
      3. Carefully and slowly lift the grid.
      4. Reposition it using the steps described in step 3.1.
3. Laminar polytrodes for a rat
   1. Polytrode setup
      1. First, connect the headstage adapter to the back side of the polytrode. Clip the connector to the third set of channels onto the board of the adapter. Ensure the black mark on the clip faces the right side of the business end of the polytrode.
   2. Polytrode Insertion
      1. Insert the polytrode into the brain until the very last (topmost) electrodes are visible above the cortical surface. A slow descent (down to 1 µm/s) improves the signal quality. Wait for 15 min, allowing the brain to adjust to the polytrode's presence.
      2. After 15 min, check whether the last electrodes have entered the cortical surface. If not, lower the polytrode slightly more and wait an additional 10 min before proceeding.
4. Positioning the optogenetic light source on a mouse
   1. Use either a fine adjustment screw system in three dimensions, mounted on an articulated arm, or a micromanipulator to mount the optical fiber holder.
   2. To guide the light source and aid in positioning the fiber, turn on the optogenetic light at a low intensity. Use the articulated arm to coarsely position the optogenetic light toward the target area.
   3. Focus and fine-tune the fiber's position using either a micromanipulator or fine adjustment screws.
5. Recording the signals
   1. Preparation
      1. Unplug all unnecessary lights, extension cords, and surge protectors in the surgical rig to reduce electrical interference. Turn off the overhead lights in the rig.
      2. Close the door to the isolated recording space and the door to the surgery room before commencing the experiment.
   2. Starting acquisition
      1. For a rat, start Synapse on the recording platform/computer and confirm that acquisition is functional by previewing and checking for signals. Elicit large, sharp voltage transients in the µECoG signal by presenting stimuli near the animal, i.e., clapping.
      2. For a mouse, start the Recording Session in Trodes.
   3. Hydration
      1. Inject the rat or mouse subcutaneously with respectively 1 mL or 0.1 mL of saline every 1-2 h during recording to prevent dehydration. For a rat, wait 5-10 min after administering saline before running a new recording block.
   4. Stimulus sets
      1. For a rat, once the recording site is confirmed, proceed with recording the required stimulus sets. An example set might include
         White Noise (60 s)
         Tone Diagnostic (5 min)
         Pure Tone (23 min)
         Dynamic Moving Ripple
         Tone 150 (15 min)
         ​TIMIT (38 min)
      2. For a rat, re-present white noise and tone diagnostics any time the grid is repositioned.
      3. Tactile stimuli for a mouse: Provide tactile stimuli in a trial structure, with each trial containing a train of random whisker deflections every 350 ms. In the provided example, each trial includes 14 deflections presented over 4500 ms.
      4. Optogenetic stimuli for a mouse: In some trials, apply a square pulse of optogenetic light over the entire trial duration (5 s). Determine the required light level based on the opsin used and on the depth of tissue to be reached using estimates of light penetrance (https://web.stanford.edu/group/dlab/cgi-bin/graph/chart.php)
6. Cleanup
   1. Lifting and cleaning the grid
      1. Once recording has finished, close the recording software and replug light sources in the rig.
      2. If the brain is dry, apply a small drop of saline onto the brain surface using a syringe. Leave the saline for 30 s to 1 min before attempting to lift the grid.
      3. Working under the microscope, gently lift the grid from the brain surface using micromanipulators.
      4. If additional force is needed while lifting the grid, use carbon-tipped forceps (closed) to gently lift the grid from the brain. Ensure the movement of the micromanipulator is slightly anterior to gently peel the grid away from the brain surface.
      5. Once the grid has been fully removed, detach it from the gripping fork and clean it following steps 3.6.1.6-3.6.1.7.
      6. Soak the grid (excluding the connector board) in a diluted enzymatic detergent (50% Enzol, 50% distilled water) for at least 1 h. Afterward, transfer it to a second bath of pure distilled water and allow it to air-dry in a safe, clean location.
      7. If areas of the grid have deposited blood or tissue, use a cotton triangle soaked in enzymatic solution to gently wipe it clean.
      8. Once dry, return the grid to its box.
   2. Euthanizing the animal
      1. For a mouse, remove the animal from head-fixation and place it in the euthanasia chamber. Add a gauze with 5 mL isoflurane and wait 60 s after cessation of respiration. Verify lack of withdrawal reflex and decapitate using sharp scissors.
      2. For a rat, inject 0.2 mL of pentobarbital IP. Wait for 60 s after cessation of respiration, lie the animal on its back, and use a #11 blade to perform a double thoracotomy.
   3. Cleaning the equipment
      1. Take all surgical tools to the lab sink and lay them on a surgical towel. Spray the tools with 10% bleach solution and wash thoroughly in the sink. For dirtier tools, allow them to soak in a bleach solution before washing.
      2. Alternatively, use a powder detergent (e.g., Contrex AP) with water by scrubbing the instruments with a brush in the sink.
      3. Once tools are clean and rinsed, wipe them down with alcohol wipes and return them totheir storage space.
   4. Sanitizing the workspace
      1. Dispose of all used needles and blades in the sharps container.
      2. Dispose of contaminated cotton swabs, triangles, and alcohol wipes in the biohazard bag.
      3. Wipe all work surfaces in the rig room with alcohol and clean all instruments before closing the workspace.

Results

We have described the protocols for recording electrocorticographic signals combined with optogenetic methods and laminar recordings. Here, typical signals obtained from the somatosensory cortex of the mouse (Figure 1, Figure 2, and Figure 3) and within the auditory cortex of rats in response to sensory stimulation (Figure 4, Figure 5, and Figure 6

Discussion

The protocols described here enable integrating high-density micro-electrocorticography (µECoG) arrays with laminar probes and optogenetic techniques. The ease of use of this protocol in rodent models makes it a powerful tool for the investigation of cortical dynamics, and the number of subjects can be easily increased. The high-density µECoG grid allows for efficient, spatially precise mapping of cortical topography across multiple areas in mice and rats, leveraging the critical role of topographical represent...

Materials

Name	Company	Catalog Number	Comments
1 disposable #11 blade	Swann Morton	303	For surgical procedures
2 disposable #10 blades	Swann Morton	3901	For surgical procedures
30 mm cage bars	Thorlabs	ER	cage components
30 mm cage plate	Thorlabs	CP33T	holding the lenses
70% ethanol	Decon Labs	V1016	Cleaning / Disinfectant (diluted to 70%)
Amalgambond PLUS Adjustable Precision Applicator Brush Teal 200/Bx	Henry Schein	1869563	precision applicator for the cement
Amalgambond PLUS Catalyst 0.7 mL Syringe Ea	Henry Schein	1861119	cement component
Amplifier (Tucker-Davis Technologies)	Tucker-Davis Technologies	PZ5M-512	Used for auditory stimulus and recording software.
Articulated arm	Noga	DG60103	for holding the fine adjustment screw system
Aspheric lenses for light collection (and one for focusing the light)	Thorlabs	ACL25416U-B	for collecting LED light
Auditory equipment	Tucker-Davis Technologies, Sony, Cortera	RP2.1 Enhanced Real-Time Processor/HB7 Headphone Drive	Used for auditory stimulus and recording software.
Buprenorphine	Sterile Products LLC	#42023017905	General analgesia
C&B Metabond Base Cement Ea	Henry Schein	1864477	cement component
C&B Metabond L-Powder Cement Clear 3 g	Henry Schein	1861068	cement component
Chlorprothixene hydrochloride (mouse)	Sigma Aldrich	Cat. No. C1671	For sedation, must be prepared the same day and kept at 4
Custom-designed 128-channel micro-electrocorticography (μECoG) grids	Neuronexus	E128-200-8-40-HZ64	For neurophysiology recordings. Placed onto the cortex.
Dengofoam gelatin sponges	Dengen dental	600034 (SKU)	can be used dry or wet, saturated with sterile sodium chloride solution
Drill bit, size 5 to 9 (Mouse)	Fine Science Tools	19007-XX	XX is the size of the drill bit e.g. 05 or 09. For mouse procedures
Drill bitSteel Round Bur (5.5 mm/7.5 mm)	LZQ Tools	Dental Bar Drill Bit Stainless Steel Bur	For rat procedures
Dumont No. 5 forceps	Fine Science Tools	11251-10	For surgical procedures
Dumont tweezers #5 bent 45°	World precision instruments	14101	for removing craniotomy window
DVD Player (Sony)	Sony	CDP-C345	System used to accept and play back stimulus sets
Electrostatic Speaker	Sony	XS-162ES	Used for auditory stimulus and recording software. Located within the rig, plays sound to the sedated rodent
Enzymatic detergent (Enzol)	Advanced sterilization products	2252	Cleaning/Disinfectant
EverEdge 2.0 Scaler Sickle Double End H6/H7 #9	Henry Schein	6011862	for scrubing the skull
Fine adjustment screw system in 3 dimension	Narishige	U-3C	for precise positioning of the optical fiber end
Gold pin	Harwin Inc	G125-1020005	Used for contact reference in mouse Soldered to the silver wire
Gripping forceps	Fine Science Tools	00632-11	For surgical procedures
Isoflurane	Covetrus	11695067772	require a vaporizer
Ketamine (Hydrochloride Injection) (Rat)	Dechra	17033-101-10	Anesthesia/Analgesic
LED	New Energy	LED XLAMP XPE2 BLUE STARBOARD	Blue LED light source
LED driver	Thorlabs	LEDD1B	LED driver
Lidocaine	Covetrus	VINB-0024-6800	to be diluted to 1% in saline
Meloxicam	Covetrus	6451603845	Anti-inflammatory used for general analgesia
Micromanipulator	Narishige (Stereotaxic Rig)	SR-6R + SR-10R-HT components	Used to manipulate ECoG and rodent with fine movements
No. 2 forceps	Fine Science Tools	91117-10	For surgical procedures
No. 55 forceps	Fine Science Tools	1129551	For surgical procedures
Ophtalmic lubricant (Artificial tears)	Akorn	17478-062-35	Used to protect eyes from dessication during surgical procedures
Optical fiber 200µm Core diameter	Thorlabs	M133L02	FC/PC connector 2 m long
Pentobarbital (Rat)	Covetrus / Dechra	VINV-C0II-0008	Anesthesia/Analgesic
Platinum Black	Sigma	205915-250MG	For neurophysiology recordings (Used for electroplating the contacts on the μECoG grids).
Povidone Iodine 10%	Betadine	https://betadine.com/medical-professionals/betadine-solution/	no catalog number ( not retail )
Powder detergent (Contrex AP)	Decon Labs	5204	Cleaning / Disinfectant
Pre-cut tape for oxygen tube	ULINE (Various Providers)	S-14726	Used to attach oxygen tube to the nose-cone of the rodent stereotaxic rig
Scalpel handle # 3	World precision instruments	500236-G	for blades # 10, #11 and #15
Scraper	Fine Science Tools	1007516	For surgical procedures
Short 30 G needles	ExelInt	26437	For surgical procedures and injections
Silver Wire	Warner Instruments	63-1319	For neurophysiology recordings (Used for grounding and as a reference electrode).
Sterilized saline (0.9% sodium chloride for injection)	Hospira	00409-7101-67 (NDC)	For dilution of injectable, and replacement of body fluids
Stoelting Hopkins Bulldog	Fine Science Tools	10-000-481	For surgical procedures
Surface disinfectant (Coverage Plus NDP Disinfectant)	Steris life science	638708	Cleaning/Disinfectant
TDT ZIF-clip connectors for acquisition.	Tucker-Davis Technologies	ZIF-Clip Analog Headstages	Connects ECoG with outside acquisition equipement
Two-pronged holding fork	Tucker-Davis Technologies	Z-ROD128	Used to connect the TDT-clips with the micromanipulator
Xylazine (Rat)	Covetrus	1XYL006	Anesthesia/Analgesic