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

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 200μm 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.

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.

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 (~1M neurons in a 36mm³ volume) across the entire brain at very low (~0.2 Hz) temporal resolution. EEG/MEG enables monitoring 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 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.^5,8-11. In rodents, it has recently been shown that micro-ECoG (μECoG) enables high temporal resolution (~100Hz) direct electrical monitoring of neuronal populations with columnar spatial resolution (~200um) 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^14-20, 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, 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 multi-scale 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^27-29. 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 (~200 μm)¹¹. This integrated methodology allows for simultaneous multi-scale 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, we provide detailed protocols for the use of high-density μECoG arrays in two combinations.

First, we describe μECoG combined with 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 multi-channel 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.

Protocols

Both protocols follow the same key steps (anesthesia, fixation, craniotomy, μECoG recording) but have notable differences. In the following description we merged the shared steps, while the specificities of each protocol are labeled with an annotation [Mouse] or [Rat]. These two labels correspond to the distinct protocols: μECoG recording with optogenetics (Mouse) or μECoG recording with a laminar probe (Rat).

All procedures described here were conducted in compliance with our local ethical or legal authorities (IACUC or Ethics Committees).

1. Preparation and Protocol for Mouse and Rat Procedures

1.1. Most notable differences between Mouse and Rat Protocols.

1.1.1. Setups for surgery versus electrophysiological recordings

• [Rat] The same setup is used during both surgery and electrophysiological recordings.

• [Mouse] Surgery is performed in a first setup and electrophysiological recordings in a second setup.

1.1.2. Head fixation

• [Rat] The rat protocol uses the same snout clamp for surgery and electrophysiological recording.

• [Mouse] The mouse protocol uses a snout clamp for surgery and an external metal headbar in the setup for electrophysiology to allow fixation under light isoflurane anesthesia. The headbar is implanted with dental cement on the skull.

1.1.3. Recording setup. Distinct acquisition electronics, recording software, and sensory stimulation software are used for the two species.

• The mouse protocol utilizes the SpikeGadgets system (https://spikegadgets.com) and the open-source Trodes software (https://spikegadgets.com/trodes/) for data acquisition.

• The rat protocol utilizes the recording software Synapse – Neurophysiology Suite (https://www.tdt.com/component/synapse-software/) for data acquisition.

1.1.4. Anesthesia is induced by injection (Rat) or by inhalation (Mouse).

1.1.5. Recording location

• [Mouse] Recordings are conducted in the somatosensory cortex (S1),

• [Rat] Recordings are conducted in the primary auditory cortex (A1).

• This difference in anatomical localization requires different craniotomy sites for each species.

1.2. Preparing and testing the Grid

• Soak the grid (excluding the connector board) in a diluted enzymatic detergent (50% Enzol, 50% distilled water) for at least 1 hour.

• Transfer it to a bath of pure distilled water and allow it to air-dry in a safe, clean location.

• Perform Platinum Black electrodeposition as part of the initial preparation of the μECoG device, not before every recording session. 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.

• To perform the electrodeposition, prepare a chloroplatinic acid solution (typically 1-3% 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.

• Apply a constant current density of approximately −0.5 to −2 mA/cm² for 10-30 seconds while monitoring impedance values.

• Test and record impedance of grid electrodes (e.g., with a Nano-Z).

• Check the grid over a light source and prepare the grid for use for post-surgery recording.

1.3. Reference cable soldering

• [Mouse] Solder the end of a silver wire, (10 mm long, 30 gauge) to a gold pin for connecting to the reference lead of the recording system.

2. Surgery

2.1. Preparation of Materials and General Monitoring (Animal Care and Recording)

The surgical procedures must be conducted in compliance with local ethical or legal authorities (IACUC or Ethics Committees). Medications used may vary according to the approved ethical protocol.

2.1.1. Preparation

Clean and disinfect the surgical area thoroughly with an appropriate disinfectant. Ensure all surgical instruments are sterilized, typically using an autoclave.

2.1.2. Surgical Tool Placement

Arrange surgical tools on the sterile surgical pad. Stock the surgical area with Puritan cotton-tip applicators and absorbent cotton surgical triangles. Dispose of any biohazardous waste in a dedicated disposal bag.

2.1.3. Temperature Regulation

Turn on the heating pad within the surgical and electrophysiological recording site. The heating pad’s temperature will be controlled throughout the surgery and recording.

2.1.4. Surgical Pad

Place a blue surgical pad or blanket over the temperature regulation bed. This pad should feature a soft cotton white bottom, which should face upwards.

2.1.5. Microscope Positioning:

Prepare the microscope and attached illuminator (e.g., LED ring) off to one side of the surgical area. Check it is functioning properly.

2.1.6. Surgical Drill

Prepare the surgical drill for the craniotomy procedure.

2.1.7. 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.

2.1.8. Fluid Replacement

Throughout the surgery, any fluid loss must be replaced—aiming for a replacement of at least 1.5% of the animal’s body weight (Mouse) or 1 mL per hour (Rat). Prepare isotonic solutions accordingly.

2.1.9. Animal

Bring the animal from the animal facility to the surgery room, according to approved procedures. We typically use mice aged 8 to 16 weeks either male or female, of C57Bl6 background; likewise, we use rats aged 7-weeks old, males, of the Sprague Dawley strain.

2.1.10. 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.

2.2. Anesthesia Induction.

2.2.1. Mouse

• Place the animal in the isoflurane induction chamber (3–5% Isoflurane in 0.5 L/min O₂).

• Once deep anesthesia is confirmed (absence of reflex in tail/toe pinch), place the animal on the surgery heating pad and headfix it (see below section 2.3.2).

• Inject subcutaneous drugs for general analgesia:

• Meloxicam: 5 mg/kg

• Buprenorphine: 0.1 mg/kg

• Mouse Head fixation

• 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.

• 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.

• Protect the animal’s eyes with a petroleum-based eye ointment or lubricant to prevent drying during surgery.

• Anesthesia is maintained throughout the procedure with continuous flow of isoflurane through the anesthesia mask.

2.2.2. Rat

• Isoflurane is only used to initially sedate the animal to ease the injection of induction anesthesia.

• 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

• The animal should reach deep anesthesia within 15–30 minutes, depending on weight and age.

2.2.3. Anesthesia monitoring. Continuously monitor the animal’s vitals (heart rate, respiratory rate) throughout the procedure. Check the respiratory rate as a particularly useful sign of early changes in anesthetic state, and adjust anesthetic level if respiratory rate changes. The paw withdrawal reflex is a critical sign of anesthetic state. Test this reflex periodically, as its total absence ensures sufficient levels of anesthesia for surgery.

2.3. Head Fixation and Monitoring of Vital Signs.

2.3.1. Animal Vitals Monitoring

• 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].

2.3.2. [Rat] Head Fixation

• 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.

• Carefully insert the points of the mounting arms into the ridge of the nose to fix the head during surgery, ensuring not to make contact with the eyes.

• 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.

• Secure both arms of the mount by tightening the screws with a hex wrench.

2.3.3. [Rat] Oxygen Setup

• 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 minute intervals throughout the procedure.

2.4. Scalp Incision.

2.4.1. Shaving and Preparation

• Shave the area from the upper snout to the back of the head, extending from one eye to the other and around the ears. This can be done efficiently by removing the bulk of the fur with scissors or an electric clipper and then applying depilatory cream.

2.4.2. Disinfection

• 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.

2.4.3. Local anesthetic Injection

• 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 minutes to allow the anesthetic to take effect.

2.4.4. Incision

• [Mouse]: Lift a point on the skin with a pair of tweezers and resect a small section of skin (approximately 1 cm in diameter) using surgical scissors.

• [Rat]: Using the scalpel, make a precise incision on the anterior side of the scalp, just above the nose, at the midline. Gently pull back the skin, creating a straight incision from between the eyes to the base of the skull. Carefully lift the scalp and cut away connective tissue, and expose the skull fully.

• Exposure of the Craniotomy Site:

  • 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.
  • Surgical clips on the skin margins can be helpful to facilitate clear exposure of the skull region where the craniotomy will be performed.

2.5. Craniotomy.

2.5.1. General Drilling Procedure

• Set surgical drill speed to a low setting 5000 rpm or 7000 rpm for experienced surgeons. Do all drilling while visualizing through the microscope.

• Hold the drill parallel to the surface of the skull and rest gently against the surface.

• With light pressure on the pedal, begin drilling in a single location. Drilling is done in short intervals (5-10 seconds) with frequent checks for changes in bone color. The bone will begin an opaque white and as the hole becomes deeper, will become more translucent, revealing a pink tint.

• 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 30G needle, gently puncture the remaining layer of bone. Clear liquid should well out of the new hole.

2.5.2. [Mouse]

• To place a reference electrode for physiological recordings, drill a burr hole in the frontal part of the hemisphere ipsilateral to the recorded area.

• 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. 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.

2.5.3. [Rat]

• Drill two holes: one in the left posterior quadrant, the other in the right anterior quadrant.

• Inject masseter muscle with a second dose of lidocaine (0.4 mL / kg at 10 mg / ml), distribute evenly where you will cut.

• Resect only the minimal set of muscles needed to expose the craniotomy area.

• 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.

• Resect the anterior muscle in a similar way until you reveal a fissure line in the skull. This line will be the anterior boundary of the craniotomy window.

• Clear muscle from around the posterior ridge with the scalpel and forceps, using a strong light source to avoid cutting into highly vascularized regions.

• Grind down the posterior ridge using the drill until it is no longer raised above the skull’s surface. This step is essential for setting down the μECoG grid to make direct contact with the cortical surface.

• Drill the dorsal edge of the window just above the ridge where the resected muscle was attached. The posterior edge will be just anterior to the drilled-down posterior ridge. The anterior edge will be placed just 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.

2.6. Craniotomy Window Drilling.

2.6.1. Drilling the Craniotomy Window (Tips)

• 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.

• [Rat]: 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.

• 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.

2.6.2. Removing the thinned skull within the craniotomy window

• When the skull has been thinned enough that extremely light pressure causes the whole window to wiggle visibly, it is time to remove the thinned skull.

• Flush the craniotomy site with a drop of saline and wait at least 1 minute. This weakens the thinned bone and helps the bone to detach from the dura. Drain excess saline with an absorbent cotton triangle or vacuum.

• Carefully lift the thinned skull using forceps, avoiding damage to the underlying tissue.

• 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.

• For mouse μECoG recordings, the dura is left intact.

2.7. [Mouse] Cement and headpost Implant

2.7.1. Insert and secure the reference wire

• Insert the silver wire end ~ 1 mm into the burr hole, enough to contact the surface of the brain, but not to cause bleeding.

• Cement in place during application of the first layer of dental cement.

2.7.2. Preparation of Dental Cement

• Use a cooled ceramic mixing dish to prepare the dental cement mixture. This cement thickens quickly, and requires preparation of new mixture regularly. Wipe clean the mixing dish before preparing a new mixture. Note: The cement should never be in direct contact with the brain.

2.7.3. Application of First Layer

• 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.

• Completely surround the craniotomy with cement, including lateral coverage, to provide adequate protection for the open craniotomy and μECoG grid.

2.7.4. Positioning the metal headbar

• Attach the large section of the headbar to its holder without fully tightening it.

• Position the headbar as desired, laying the thin section along the skull midline in contact with the cement surface.

2.7.5. Securing the Implant

• Cover the headbar with dental cement and connect it to the cement surface.

2.7.6. Removing the Holder

• Allow a few minutes for the dental cement to strengthen

• 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.

2.8. [Rat] Durotomy for Rat surgery. (Note- this is a challenging surgical step)

2.8.1. Lifting the Dura

• 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.

• Use a fresh 30G needle (as short as possible) to carefully tear the lifted dura.

• 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.

2.8.2. Resection of the Dura

• Continue gripping the dura with the forceps and lifting it away from the brain. Create a diagonal tear with the needle while lifting.

• Use the forceps to carefully peel the dura towards the sides of the craniotomy window, ensuring the brain surface remains undisturbed.

2.9. [Mouse] Transfer the animal in the setup for electrophysiological recording.

2.9.1. 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), which is a sedative that enables continuous anesthesia to be maintained using a lower Isoflurane concentration.

2.9.2. Place the mouse in the electrophysiological recording setup

• Make sure that the heating pad is in place and working properly.

• Head fix the animal using the headbar on the holder in the electrophysiological setup

• Bring the isoflurane mask close to fully cover the animal's snout.

2.9.3. Anesthesia Adjustment

• Gradually reduce anesthesia levels to 0.7–1% Isoflurane (in increments of 0.5% maximum every 5 minutes).

• Monitor the animal’s respiratory rate and movements. Respiratory rate should increase slightly compared to the surgical state, but the animal should not be moving.

• 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%.

2.9.4. Insert whiskers for sensory stimulation

• Attach the mouse’s vibrissae to the whisker stimulation device. In this protocol, nine vibrissae are inserted into short 10 μL pipette tips, which are connected to piezoelectric actuators that provide rapid deflections of the vibrissae.

3. Recording

3.1. Installing the Grid

3.1.1. Preliminary Steps

• Turn on the recording system and amplifier.

• Check the animal’s vital signs.

3.1.2. Procedure

• Positioning the Animal and Tools

  • The animal is placed in the recording setup, the craniotomy remains moist by regularly applying saline solution.
  • [Rat] Position the micromanipulator on the rig’s railing, located well behind the craniotomy site to avoid interference.
  • [Mouse] The micromanipulator is placed laterally to the craniotomy site, alongside the animal.
• [Mouse] Attaching and Positioning the Grid

  • Attach the μECoG grid to the headstage using the ZIF-clip connectors. The headstage's electronic board is held in place via a mechanical bar fixed to a micromanipulator.
  • Lower the μECoG grid horizontally so it aligns 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.)
  • 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.
  • Further lower the grid to contact the brain.
  • Move the grid laterally so it "glides" over the moist dura surface. Continue adjusting until the grid is centered along the mediolateral axis.
  • Use aspiration or a surgical sponge around the craniotomy's edges to remove any excess saline solution.
  • Once the preparation is slightly drier, the grid should adhere more firmly to the dura and 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.
• [Rat] Positioning the Grid

  • 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.
  • 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.
  • 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.
  • Wick away moisture carefully using an absorbent cotton triangle, without touching the grid itself. Every row of the grid should be in contact with the brain surface. Removing moisture prevents 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.
  • 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. Wires should be inserted only ~1mm, enough to contact the brain, but not to cause bleeding or trauma to the brain.

3.2. Check Positioning of the Grid

3.2.1. Monitoring Electrophysiological Activity

• Electrophysiological activity can be observed using the recording software. Under light anesthesia, brain signals are variable and can exhibit a variety of patterns. 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. Noise in the high-frequency range can be monitored using bandpass filtering with Trodes (e.g., 100-6000 Hz), and should not exceed a few tens of μV.

3.2.2. Sensory responsiveness can be assessed using noise (e.g., clapping or snapping fingers) to induce visible event-related cortical surface electrical potentials (CSEPs). Stimulation of a single whisker should evoke a clear, sharp event-related CSEP in only a few channels (Mouse).

3.2.3. Grid Position Verification:

• [Rat] It is important to confirm that the grid is correctly positioned over the auditory cortex. The first block recorded should typically be a 60-second white noise stimulus set to verify that the grid registers a proper response from the brain. Conduct white noise and tone diagnostic recordings with grid only before inserting polytrode. This will help determine whether the grid was placed correctly and there is signal response.

• [Mouse] To verify grid positioning, perform a quick mapping session with 20–30 whisker deflections spaced by 350 ms. Activity in the LFP band can be recorded using Trodes and analyzed offline with a custom MATLAB code to visualize the spatial extent of whisker-evoked activity.

3.2.4. Repositioning

• If the grid requires adjustment, moisten the cortical surface with drops of saline over the grid.

• Leave the saline for 30 seconds to a minute before attempting to lift the grid.

• Carefully and slowly lift the grid

• Reposition it using the steps described in Section 3.1.

3.3. [Rat] Laminar Polytrodes

3.3.1. Polytrode Setup

• First, connect the ZIF clip 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.

3.3.2. Polytrode Insertion

• 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 minutes, allowing the brain to adjust to the polytrode's presence.

• After 15 minutes, check whether the last electrodes have entered the cortical surface. If not, lower the polytrode slightly more and wait an additional 10 minutes before proceeding.

3.4. [Mouse] Positioning the Optogenetic Light Source

3.4.1. Use either a fine adjustment screw system in three dimensions, mounted on an articulated arm, or a micromanipulator to mount the optical fiber holder.

3.4.2. To guid 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.

3.4.3. Focus and fine-tune the fiber’s position using either a micromanipulator or fine adjustment screws.

3.5. Recording

3.5.1. Preparation

• Unplug all unnecessary lights, extension cords, and surge protectors in the surgical rig to reduce electrical interference. Turn off overhead lights in the rig.

• Close the door to the isolated recording space and the door to the surgery room before commencing the experiment.

3.5.2. Starting Acquisition

• [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.

• [Mouse] Start the Recording Session in Trodes

3.5.3. Hydration

• Inject the rat or mouse subcutaneously with respectively 1 mL or 0.1 mL of saline every 1–2 hours during recording to prevent dehydration. Wait 5–10 minutes after administering saline before running a new recording block (Rat).

3.5.4. Stimulus Sets

• [Rat] Once the recording site is confirmed, proceed with recording the required stimulus sets. An example set might include

  • White Noise (60 seconds)
  • Tone Diagnostic (5 minutes)
  • Pure Tone (23 minutes)
  • Dynamic Moving Ripple
  • Tone 150 (15 minutes)
  • TIMIT (38 minutes)

• [Rat] White Noise and Tone Diagnostics should be re-presented any time the grid is repositioned.

• [Mouse] Tactile Stimuli

  • 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.
• [Mouse] Optogenetic stimuli

  • In some trials, apply a square pulse of optogenetic light over the entire trial duration (5s). The required light level can be determined 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)

3.6. Cleanup

3.6.1. Lifting and Cleaning the Grid

• Once recording has finished, close the recording software and replug light sources in the rig.

• If the brain is dry, apply a small drop of saline onto the brain surface using a syringe. Leave the saline for 30 seconds to 1 minute before attempting to lift the grid.

• Working under the microscope, gently lift the grid from the brain surface using micromanipulators.

• If additional force is needed while lifting the grid, carbon-tipped forceps can be used (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.

• Once the grid has been fully removed, detach it from the gripping fork and clean it.

  • Soak the grid (excluding the connector board) in a diluted enzymatic detergent (50% Enzol, 50% distilled water) for at least 1 hour. Afterward, transfer it to a second bath of pure distilled water and allow it to air-dry in a safe, clean location.
  • If areas of the grid have deposited blood or tissue, use a cotton triangle soaked in enzymatic solution to gently wipe it clean.

• Once dry, return the grid to its box.

3.6.2. Euthanizing the Animal

• Remove the animal from the head fixation device and close the oxygen tank. Turn off the heating pad system.

• Euthanize the animal using approved methods.

3.6.3. Cleaning Equipment

• 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.

• Alternatively, use a powder detergent (Contrex AP) with water by scrubbing the instruments with a brush in the sink.

• Once tools are clean and rinsed, wipe them down with alcohol wipes and return them to their storage space.

3.6.4. Sanitizing the Workspace

• Dispose of all used needles and blades in the sharps container.

• Dispose of contaminated cotton swabs, triangles, and alcohol wipes in the biohazard bag.

• Wipe down all work surfaces in the rig room with alcohol and clean all instruments before closing the workspace.

Representative results

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

Figure 1: Recording μECoG signals over the mouse whisker somatosensory cortex.

A. Components of the setup. See text.

B. Top view of the surgical setup before μECoG recording, showing placement of the head restraint and craniotomy site.

C. Detailed view of the μECoG grid and optic fiber positioned on the exposed cortex.

D. Top: Light source is located outside of the recording box. Light from a 473 nm LED is collected in the optical fiber through a lens tandem. Bottom: close-up of the cortical grid over the mouse cortex in an example recording.

E. Representative μECoG traces from all 16x8 channels showing trial averaged responses to whisker D2 stimulation

F. Raw voltage trace from the single channel indicated in (E) across single deflections of multiple whiskers. The channel exhibited the strongest response during stimulation of the whisker D2.

G. Average evoked response across trials in the single channel (E,F), which exhibited a sharp voltage deflection following whisker stimulation.

Figure 3: Light delivery to focal points of the cortex.

A. Light from the 1 mm diameter optical fiber is focused on the brain surface. This simple adjustment allows for more precise targeting of light on the brain surface and within the tissue.

B. The spatial extent of the light delivered to the brain is measured from the opto-electric artifact

C-D. as in A-B with a 200 μm optical fiber which allows it to directly target an area of 250 μm diameter, roughly the size of a cortical column.

Figure 4: Recording μECoG and laminar polytrode signals over the rat primary auditory cortex (A1).

A. Components of the μECoG and laminar polytrode setup. See text.

B. Top view of the surgical setup before μECoG and laminar polytrode recording, showing placement of the head restraint and craniotomy site.

C. Detailed view of the μECoG grid and laminar polytrode on the exposed auditory cortex.

D. A close-up view of the μECoG grid placement on the cortex, including size of individual electrodes (40 μm) and between electrodes (200 μm).

E. Average z-scored wavelet decomposition of a single-channel response to a single frequency-attenuation pair. The vertical red lines indicate the peak frequency response period shown in F.

F. Red- Average peak frequency response across 20 presentations of a single frequency-attenuation pair. Gray- Standard error. The frequency axis is on a log-scale. Canonical neural frequency bands are indicated across the top.

G. Photomicrograph of an 8x16 μECoG grid on the surface of rat primary auditory cortex (A1). A 32-channel laminar polytrode has been inserted in the center μECoG window.

H. Schematic of 3D multiscale recording of cortical activity.

I. Top: Spectrogram of 50ms pure tone pips. Middle: In red, the average cortical surface electrical potential of 4 μECoG electrodes (gray). Bottom: 32-channel laminar polytrode voltage traces are arranged by cortical depth

Figure 6. μECoG signals are similarly tuned to neural unit spike recordings across the depth and reveal tonotopic organization of the auditory cortex.

A. Frequency-Response Amplitude plots (FRAs), which display the high-gamma response (color bar) as a function of the frequency (x-axis) and amplitude (y-axis) of the auditory stimulus. (top) FRAs from 2x16 subset of μECoG electrodes on the auditory cortex; (bottom) FRAs from 1x16 subset of laminar polytrode electrodes inserted through perforations in the μECoG array, indicated by gray arrows.

B. High-resolution tonotopic organization of multiple auditory cortical fields derived from high-gamma activity. Each pixel is color coded according to the best frequency of that electrode. The 8x16 μECoG array displayed here covers multiple auditory cortical fields (A1, PAF, and VAF) and the approximate boundaries are demarcated (black lines).

In Figure 1A-D we provide images of various components of the system used for recording and for optogenetics manipulation in mouse S1 cortex. Labeled elements include: 1- nine independent whisker stimulator in a 3x3 array, 2- nose cone for isoflurane anesthesia, 3- head-post holder/counterpart, 4- μECoG back-end PCB, connected to the headstage PCB with a double ZIF connector; 5- μECoG grid of electrodes at the tip of an 8 mm flexible cable; 6- reference wire; 7- ground wire; 8- headpost implant; 9- reference gold pin (insulated from the headpost with dental cement); 10- craniotomy on the left hemisphere, over the S1 whisker barrel cortex; 11- 1 mm diameter optogenetic fiber (held by a micromanipulator, not shown); 12- a star-shaped LED driven by an LED driver (not shown); 13- light collection through a tandem of aspheric lenses. When stimulating a single whisker, we observe a rapid deflection of the surface potential at a small number of electrodes (Fig. 1E). This cluster of electrodes represents the local signal that peaks in the cortical column of the stimulated whisker³⁰. Looking at a single electrode, we observe the strongest response for stimulation of its preferred whisker, and weaker or no response to stimulation of more distant whiskers (Fig. 1F). In this example, the onset of the deflection occurs around 10 ms after whisker vibration, with an average amplitude of 1 mV (Fig. 1G).

In Figure 2 we present example recordings during optogenetic inhibition. The light can reach the cortical tissue through the transparent substrate of the grid (Fig. 2A). However, when using a large diameter fiber, or any spatially broad source of light, photons also hit the electrodes, producing an opto-electric artifact (Fig. 2B). In this protocol of inhibition, we used a 5 second long square pulse of light. The resulting opto-electric artifacts are only present at onset and offset of the light. In an animal without opsin, whisker stimulation presented during the light on do not evoke a different response from the whisker stimulation presented in trials with the light off (Fig. 2C). In contrast, light stimulation of the inhibitory opsin in a subpopulation of excitatory neurons leads to a decrease in the amplitude of the sensory evoked response (Fig. 2D)

Figure 2: Optogenetic suppression in wS1 during μECoG recording in mice.

A. Schematic of a representative optogenetic experiment. An optical fiber is directly pointed at the brain, allowing transient inhibition of the neuronal population expressing an inhibitory opsin.

B. Voltage trace recorded in the channel at the center of optogenetic stimulation, averaged across trials. The black arrow indicates opto-electric artifacts at onset and offset of the 473 nm light. Note the response to whisker deflection (random whisker identity) in the middle of the trial. The artifact is transient and does not affect recording of sensory evoked activity after a delay (here 500 ms).

C. Average spectrogram of an example channel in light-off trials, light-on trials, and the difference between the two, in an animal where no opsin was expressed. Note the opto-electric effect induces a broadband, transient artifact and the whisker evoked response is unaffected by the light.

D. Example trial-averaged spectrogram across light-off trials, light-on trials, and the difference between the two, in a Rbp4-Cre mouse³² in which st-GtACR2.0 was expressed in excitatory neurons of layer 5. Note the suppression of whisker evoked response in the spectrogram.

Figure 2C,D and present examples of optogenetic suppression in the time-frequency domain. To analyze the μECoG data, we first apply a common average referencing (CAR) to remove signals that are undifferentiated across electrodes (e.g., breathing) followed by a Morse wavelet transform which reveals active frequency bands over time. Neural activity in the frequency domain typically exhibits an approximate power law of 1/f^2-3. To reveal the sensory evoked signal more uniformly across the frequency domain, we apply Z-scoring separately for each frequency band. The Z-scoring is based on the statistics of the signal during baseline epochs. Here, we use a time window that precedes the stimulus, −3000 to −1000 ms prior to the trial start. This process yields the stimulus evoked Z-score for each frequency band (Fig. 2C,D).

In Figure 3, we present easy-to-implement variations in the light delivery system. Using a smaller optical fiber, or a simple lens, it is possible to target a specific area of the cortex¹⁵. The light coming out of the fiber is rapidly diverging, and reaches cortical tissue beyond the targeted area. By incorporating an aspheric lens (f =16 mm) at the optical fiber output, it is possible to focus the light to a smaller surface area (Fig. 3A,C), down to roughly the diameter of a single cortical column (Fig. 3C). Light is also less diverging within the cortex. Ideally using a laser or a powerful source of light with a tandem set of lenses, it is possible to target a single cortical column with a collimated beam of light. However, it is important to consider that light will scatter within the tissue, which may partially illuminate neighboring columns. The light artifact measured in the μECoG data will reveal where light has been delivered over the cortical surface, see comparison of Fig. 3B versus Fig. 3D. The light artifact is measured as the peak power in a high frequency range (65 - 500 Hz) 5 ms after light onset.

Rat μECoG and Laminar Polytrode recordings

In Figure 4A-D we provide images of various components of the system used for rat primary auditory cortex recordings. Specifically, labeled elements include: (1) μECoG micromanipulator, (2) rodent stereotaxic rig, (3) μECoG cable with ZIF-clip connectors, (4) laminar polytrode micromanipulator, (5) rat nose cone and head stabilization apparatus, (6) electrostatic speaker for stimulus presentation, (7) laminar polytrode cable with ZIF-clip adapter, (8) μECoG ZIF-clip attached by a two-pronged fork, (9) 32-channel polytrode device, (10) electrostatic speaker, (11) alligator clips to hold surgical site open, (12) grounding wires, one inserted into the cortex through the burr holes in the left posterior quadrant or right anterior quadrant and another is grounded to tissue, (13) close-up view of μECoG grid, (14) close-up view of inserted polytrode. Figure 4E displays the average neural spectrogram derived from the recorded electrical potentials of a single μECoG electrode in response to 50 ms tone pips of the same frequency and attenuation (N = 20 trials). Across frequencies, the evoked response exhibits a sharp peak between 25 ms and 30 ms (indicated by red solid lines) after the stimulus onset (left gray dotted line). The apparent response preceding the stimulus onset (ostensibly acausal) is due to the large bandwidth at lower frequencies of the constant-Q transform, which smooths (acausal, but no phase offset) rather than filters (causal, but induces a phase offset) the signal. Therefore, the time of the peak response remains accurate. At the time of the peak evoked response, we observed that the z-scored cortical surface electrical potential (CSEP) was multimodal across frequencies. Specifically, it exhibited three primary non-harmonic peaks: the first and largest in the 40–180 Hz gamma/high-gamma (γ/Hγ) range, the second in the 200–450 Hz ultrahigh-gamma (uHγ) range, and a final peak above 500 Hz associated with multi-unit activity (MUA) (Fig. 4F)¹¹. This stimulus-evoked multimodal structure is robust across all tuned electrodes. Here, we focus on Hγ because of its prevalence in human electrocorticography (ECoG) recordings³¹. In Figure 4G, we display a μECoG array that was placed subdurally alongside a silicon laminar polytrode to simultaneously measure cortical surface electrical potentials (CSEPs) and spiking activity across cortical laminae. The custom-fabricated μECoG array consisted of 8×16 electrodes with a 200 μm pitch and 40 μm contact diameter, and we utilized a 32-channel laminar polytrode configured as 2×16 channels with a 46 μm pitch and 15 μm contact diameter. The μECoG array was sufficiently large to cover the entire rat primary auditory cortex (A1), and its small 40 μm diameter electrodes enabled the measurement of local CSEPs suitable for deriving auditory tuning. Spatial resolution of the μECoG high-gamma (Hγ) signal is ~200 μm, comparable to the radius of a rat cortical column. Thus, μECoG provides a "columnar view" of cortical activity. Perforations in the μECoG grid allowed the laminar polytrode to pass between surface contacts, enabling direct recording of neuronal activity across cortical laminae (Fig. 4H). Example voltage traces recorded from μECoG and laminar polytrode electrodes are shown in Figure 4I.

In Figure 5, we display the recording of μECoG signals simultaneously with spiking activity using a laminar silicon polytrode (Camb64). The μECoG signal, recorded from a surface electrode, is shown as a raw voltage trace (Fig. 5A) and its wavelet transform, highlighting frequency decomposition over time (Fig. 5B). Mulli-unit activity was recorded using the polytrode, as illustrated by a representative raw voltage trace from a single channel (Fig. 5C), where individual action potentials (spikes) were detected using a simple thresholding method (voltage crossing a −120 μV threshold). The extracted spike waveforms from this channel are well-defined for the majority of spikes, although they could originate from multiple sources of neurons (Fig. 5D, inset). Across multiple channels in the same penetration, the average spike waveforms further confirm the consistency of well defined spiking activity recordings (Fig. 5E). Thus, these recording methods could support single unit recording with spike sorting analysis, and analysis of quality metrics such as the interspike interval or firing rates (Fig. 5F). In summary, the quality of the laminar probe recordings is good, with clear waveform morphology on single channels. These results illustrate that this method enables simultaneous acquisition of μECoG and spiking activity.

Figure 5. Simultaneous recordings of μECoG and columnar spiking activity.

A. Example raw voltage trace from a single μECoG channel.

B. Wavelet transform of the same μECoG channel, showing frequency decomposition over time

C. Example raw voltage trace from a single polytrode channel (acute 64-channels silicon probe camb64), showing action potentials (spikes) detected using a simple voltage threshold.

D. Extracted spike waveforms from the polytrode channel in (C) aligned and color coded based on the distribution of voltage amplitude. 7 out of 142 spike waveforms are not shown, as they were considered outliers based on their distance from the main cluster in spike width and amplitude (see inset)

E. Average multi-unit spike waveforms extracted from 9 channels in the same penetration.

F. Interspike interval (ISI) histograms for different polytrode channels (e.g., Channels 21, 27, 63, and 61), with corresponding firing rates (FR) indicated in Hz. ISI violations (< 5ms) represented less than 5% of spikes in all units.

With the capability to perform simultaneous recordings using the custom-designed μECoG array and laminar polytrode, we investigated how μECoG signals compare to neural unit recordings across cortical depth in terms of auditory tuning. Figure 6A presents Frequency-Response Amplitude (FRA) plots, which depict the high-gamma (Hγ) response as a function of auditory stimulus frequency and amplitude. The top panel shows FRAs from a 2×16 subset of μECoG electrodes positioned on the auditory cortex, while the bottom panel displays FRAs from a 1×16 subset of laminar polytrode electrodes inserted through the μECoG array's perforations (indicated by gray arrows). Remarkably, the FRAs obtained from the μECoG electrodes closely resemble those from the laminar polytrode recordings, suggesting that μECoG signals are similarly tuned to neural unit activity across cortical layers.

Leveraging the high spatial resolution of the μECoG array, we generated a high-resolution tonotopic map of multiple auditory cortical fields based on the Hγ activity¹¹. In Figure 6B each electrode's best frequency is color-coded, revealing the tonotopic organization across the cortical surface. The 8×16 μECoG array covered several auditory cortical fields—including the primary auditory cortex (A1), posterior auditory field (PAF), and ventral auditory field (VAF)—with approximate boundaries demarcated by black lines. This detailed mapping underscores the μECoG array's capability to provide a "columnar view" of cortical activity, capturing functional organization with a spatial resolution of less than 200 μm, comparable to the dimensions of a rat cortical column. These findings demonstrate that μECoG recordings not only reflect the tuning properties observed in neural unit recordings across cortical depth but also enable high-resolution mapping of functional cortical organization.

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 representations in brain organization³³. The addition of laminar recording allows for the examination of cortical dynamics across multiple spatial and temporal scales. The inclusion of optogenetics enables causal manipulation to determine the relationships between specific neural populations and their contributions to cortical surface-evoked potentials (CSEPs) and cortical processing³⁴.

Optogenetics allows for the selective modulation of specific neuronal populations, enabling the investigation of their causal roles in generating cortical activity patterns and participating in computation whose signatures can be sensed with μECoG. For example, our studies demonstrate that targeting a specific cell type can alter cortical surface electrical potentials (CSEPs). This approach could be used to dissect the neural cell type basis of μECoG signals, allowing us to identify which neuronal cell types contribute to characteristic sensory-evoked activities, such as those observed in the high-gamma band. Other large-scale phenomena, such as cortical rhythms in different frequency bands²¹, or spatially organized activity, including traveling waves^35,36, could similarly be investigated. Additionally, a range of genetically modified mouse lines and opsins are readily available, providing opportunities to explore specific circuit mechanisms. Optogenetic techniques could be employed to examine the functional effects of horizontal connections between columns³⁷, which play a crucial role in various sensory computations, such as surround suppression³⁸ or perceptual binding³⁹. In summary, the ability to manipulate neuronal activity through optogenetics enables testing the links between specific neuronal populations and the properties of CSEPs or between population and specific cortical computations measurable with μECoG. This approach could effectively dissect the relationships between local neural structures and global cortical activity.

Laminar polytrode recordings enable sampling of single neuron activity from multiple neurons within small cortical volumes, i.e., within an individual cortical column. These recordings are crucial because individual neurons can independently encode distinct information, employing selective coding of stimuli—such as the "Jennifer Aniston neurons" described by Quiroga et al.⁴⁰— or providing complementary representations in a higher-dimensional space, as seen in mixed selectivity⁴¹. Traditionally, electrophysiologists studied neural activity in the context of simple, parametrically designed stimuli and behaviors (e.g., single whisker deflection, or pure tones, as deployed here). The representation of such stimuli tends to be fairly spatially localized (e.g., individual columns). However, many ethologically relevant stimuli and behaviors are more complex, and as such typical patterns of neural activity during such paradigms frequently extend beyond single columns – even all across the brain⁴². In these instances, μECoG offers a comprehensive readout that captures high-temporal resolution, columnar resolved activity across multiple columns simultaneously. In summary, our protocol effectively bridges the gap between local processing within individual cortical columns and the more extensive dynamics that occur across multiple columns in an entire cortical area and across areas.

As general guidance and troubleshooting, we propose a few recommendations. Conventional acute in vivo electrophysiological protocols typically advise keeping the brain wet during exposure as a means of prolonging health of the exposed brain. While this is likely useful in some circumstances, our experience suggests the opposite is true for μECoG recordings in rodents. Indeed, we found that the recording quality was qualitatively better when the μECoG arrays were placed on a slightly dry cortical surface. We believe this arises because having saline, a highly conductive ionic solution positioned between the cortical surface and the recording electrodes homogenizes electrical signals generated by the brain. Effectively, saline is ‘shorting’ the electrodes together. Because μECoG grids monitor the continuous electrical field generated by populations of neurons, it is important to ensure that electrophysiological systems are well-assembled and denoised. During recordings, the connection with the reference electrode is critical. Without improvement during the recording, the pre-processing of the data can incorporate a notch filter at 50 or 60 Hz to remove line noise. However, this will dramatically alter the signal, and so must be accounted for in subsequent analysis. Considering optogenetic light stimulation, the opto-electrical artifact must be minimized or at least accounted for in the experimental design (e.g., by including a delay following onset or offset of light). Ideally, light is delivered in the area between the electrodes using a small optical fiber diameter, or focusing or collimating the light. If this artifact cannot be completely avoided (but see studies proposing different probe designs^15,20,43 including clear electrodes), it can be reduced by using the least amount of light necessary. New generation opsins require less light for effectiveness^44,45. We suggest calibrating the optogenetic light power prior to the experiment using laminar probes. The artifact shape can also be modified and reduced by avoiding sharp transitions in the light stimulus (e.g., using a light ramp instead of a square pulse). In any case, control conditions using animals that do not express opsins are advisable to differentiate genuine changes in neural activity from artifact-related signals. Finally, the electrodeposition process creates a rough, high surface area coating that enhances charge transfer between electrode and tissue while maintaining mechanical stability during recordings, reducing electrode impedance by 1-2 orders of magnitude compared to bare platinum and enabling better neural signal detection⁴⁶.

Acute μECoG offers flexibility and reduced complexity in experimental setups, allowing for detailed mapping of neural activity over the course of tens of minutes of recordings. As a mesoscale method, it enables an inter-areal but not whole-brain tracking of cortical activity, though the exact sources of the signals remain uncertain^1,47. In the future, multimodal studies should provide a better picture of the signal’s origin. Acute μECoG is limited in capturing long-term neural dynamics and may be influenced by transient factors such as surgical recovery or anesthesia^48,49. In contrast, chronic μECoG enables prolonged observation of neural activity, providing insights into processes like learning, plasticity, and disease progression^10,13. Chronic μECoG also presents challenges such as electrode stability, potential signal degradation, and risks associated with long-term implantation, including tissue scarring or infection^50,51. These challenges tend to be less severe when compared to penetrating electrodes and are supposedly further reduced with epidural implantation of the μECoG in mice (at the cost of lower signal quality)⁵². It is possible to reuse the same μECoG grid across many sessions in the same or different animals, removing the grid at the end of a session and replacing it in the next session. In this sub-acute configuration, the brain must be kept moist at all times, and protected with a glass coverslip between sessions. We have observed that μECoG grids are fairly durable; an individual grid can be reused ~20 times with proper handling and cleaning, making them cost effective recording devices. Grids can be designed with diverse numbers and geometries of electrode layouts. Our results suggest that reducing electrode pitch to < 200μm results in only negligible improvements of functional resolution, due to the granularity of neural representations in the underlying cortex.

Table of materials

Name of Material/Equipment	Company	Catalog Number	Comments/Description
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%)
A fine adjustment screw system in 3 dimension	Narishige	U-3C	for precise positioning of the optical fiber end
A micromanipulator	Narishige (Stereotaxic Rig)	SR-6R + SR-10R-HT components	Used to manipulate ECoG and rodent with fine movements
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.
An 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 Gm 3gm/Bt	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 / Steel Round Bur (5.5mm / 7.5mm)	LZQ Tools	Dental Bar Drill Bit Stainless Steel Bur	For rat procedures
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
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
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
No. 2 forceps	Fine Science Tools	91117-10	For surgical procedures
No. 55 forceps	Fine Science Tools	1129551	For surgical procedures
Ophthalmic 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 2m 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 30G 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