Name: Surgical Protocol for a Large, Resealable Cranial Window Enabling Longitudinal, Multi-Modal Electrophysiology Recordings of Mouse Default Mode Network (Video) | JoVE Journal
Uploaded: 2026-05-29T16:25:00.000+00:00
Duration: 14 min 34 s
Description: Learn a two-phase surgical protocol for creating a large, resealable cranial window in mice. This method enables stable, longitudinal electrophysiological recordings from the default mode network using micro-ECoG and Neuropixels probes.

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5 minutes

This work was supported by the Research Council of Finland grants #327192 and 347358, and the Sigrid Juse&#769;lius foundation. The authors thank the members of the Castr&#233;n and Palva laboratories for their technical assistance and helpful discussions throughout this project.

EC is a co-founder, Board member and recipient of research support from Kasvu Therapeutics, Ltd, and has received a speaker fee from Janssen-Cilag. The other authors have nothing to disclose.

This report details a two-phase, dura-sparing surgical protocol that provides a reliable and highly effective method for creating a large, chronic, and resealable cranial window in mice. The significance of this technique extends beyond the size of the window. Its principal advantage is the capacity for simultaneous, multi-scale interrogation of large-scale brain networks over longitudinal timescales. By combining surface &#181;ECoG grids (providing mesoscale LFP data) with deep, high-density intracranial electrode probe...

The Default Mode Network (DMN) was first identified in human brain imaging studies as a constellation of regions consistently suppressed during externally oriented, attention-demanding tasks1. It is now considered to be a fundamental large-scale network supporting internally oriented cognitive processes, such as self-referential thought, autobiographical memory, and planning of future events2,3. In humans, the canonical DMN comprises a set of anatomically connected core regions, including the medial prefrontal cortex (mPFC), posterior cingulate cortex (PCC), inferior parietal lobule (IPL), and the hippocampal formation4. Dysfunction within this network is a key feature of several neuropsychiatric disorders. In major depressive disorder (MDD), the DMN commonly exhibits hyperactivity and altered patterns of functional connectivity, which are thought to underlie core symptoms of MDD, such as rumination and negative self-image5,6.
To study the cellular and circuit mechanisms driving DMN dysfunction, robust animal models are indispensable. A homologous DMN has been reliably identified in rodents, consisting of comparable brain regions such as the mPFC, cingulate cortex, and retrosplenial cortex (RSP), thereby validating the mouse as a translationally relevant model for mechanistic investigation7,8. Furthermore, chronic stress, a key factor in depression, has been shown to alter DMN connectivity in rodents in a manner similar to that observed in human MDD patients9.
Understanding the neuroplastic changes that occur in disease states or in response to chronic therapeutic interventions require longitudinal observation of neural activity, since a single recording session cannot capture the dynamic adaptation of neural circuits over time10. In vivo electrophysiology provides the millisecond-scale temporal resolution required to study neural oscillations and synchrony in the local field potential (LFP). LFP exhibits the summed extracellular signal reflecting synchronized synaptic activity of local neuronal populations, which are the fundamental substrates of network communication11,12. However, performing chronic, large-scale electrophysiological recordings in awake mice remains technically demanding. Conventional chronic cranial windows were developed for optical imaging in the awake mouse and remain the standard preparation for that purpose, where their typical single-hemisphere footprint and transparent glass coverslip are optimal for the intended readout13. They were not, however, designed for awake multi-site electrophysiology, so the working area and the optical interface preclude simultaneous high-density recording from the distributed bilateral nodes of a network such as the DMN14. Chronic electrode implants, in turn, enable longitudinal recordings but elicit progressive foreign-body encapsulation and glial scarring around the permanently positioned shank, reducing signal quality over time15,16.
The two-phase surgical procedure described here directly addresses these limitations. It produces a large, resealable cranial window closed between sessions by a passive polydimethylsiloxane (PDMS) membrane, silicone sealant, and a 3D-printed cap permitting repeated non-terminal insertion of &#181;ECoG grids for mesoscale LFP recordings together with multiple high-density intracranial electrode probes for microscale, laminar-resolved single-unit and LFP activity17,18. In contrast to prior chronic approaches, which either rely on permanently implanted electrode arrays that accumulate glial encapsulation of the electrodes over time or require terminal acute craniotomies that preclude longitudinal designs, the window described here preserves an intact dura and avoids permanent foreign-body contact15,16,18. Furthermore, in a validation cohort in which the cranial window was reopened three times (post-operative days 0, 21, 22) without electrophysiological recording, quantitative immunoreactivity of glial fibrillary acidic protein (GFAP) and ionized calcium-binding adapter molecule 1 (IBA1) did not differ significantly from non-surgical controls, suggesting low astrogliosis and microgliosis under the repeated-opening schedule used here. The method is particularly suited to longitudinal, multi-site recordings of distributed cortical networks, including the DMN, combined with laminar high-density intracranial electrode probe access to deep structures in awake head-fixed mice. Its main operational constraints are the microsurgical expertise required for reliable dura-sparing bone-flap removal and its incompatibility with freely moving recordings, since the high-density intracranial electrode probes require head-fixation.

<table><tbody><tr><td>&amp;micro;ECoG array &amp;mdash; 512-channel custom (2 &amp;times; 256 ch)</td><td>NeuroNexus</td><td>E256-200-30-80</td><td>Packaging: IH256. Two 16&amp;times;16 sub-grids (each 3.3 mm &amp;times; 3.5 mm); 200 &amp;mu;m pitch; 80 &amp;mu;m electrode diameter. Surface-level DMN recordings.</td></tr><tr><td>3-axis left-hand micromanipulator arm</td><td>World Precision Instruments (WPI)</td><td>504926</td><td>10 &amp;mu;m resolution; digital display; used for headplate/probe positioning during surgery</td></tr><tr><td>3D-printed protective cap</td><td>In-house (Anycubic Photon Mono 4K)</td><td>N/A</td><td>Custom design; printed in Anycubic High Clear UV resin; STL file available at https://github.com/RazBalin/ProtectiveDMN_Cap</td></tr><tr><td>Acetone (laboratory grade)</td><td>Sigma-Aldrich</td><td>179973</td><td>Skull surface degreasing and periosteum removal prior to adhesive application</td></tr><tr><td>Adson Dura Hook 5 mm (Fine dura hook)</td><td>gSource</td><td>gS 25.2570</td><td>Used to detach bone plate from dura along perimeter; fashioned from fine tungsten wire or similar</td></tr><tr><td>Anycubic High Clear UV Resin</td><td>Anycubic</td><td>STMGCL-102B&amp;nbsp;</td><td>Photopolymer resin used with Anycubic Photon Mono 4K printer for 3D-printed protective caps</td></tr><tr><td>Anycubic Photon Mono 4K 3D resin printer</td><td>Anycubic</td><td>PHOTON-MONO-4K</td><td>Used to print protective window caps from Anycubic High Clear Resin</td></tr><tr><td>Anycubic UV Resin Wash &amp;amp; Cure Machine 2.0</td><td>Anycubic</td><td>285-550</td><td>Post-print washing and UV curing of 3D-printed caps</td></tr><tr><td>Articulating arm with magnetic base (&amp;times;2)</td><td>Haas Tooling</td><td>09-0535</td><td>Used to hold &amp;micro;ECoG sub-grids in place on cortical surface during recordings</td></tr><tr><td>Artificial CSF (aCSF)</td><td>In-house preparation</td><td>N/A (In-house)</td><td>aCSF recipe: 124 mM NaCl, 3 mM KCl, 26 mM NaHCO&lt;sub&gt;3&lt;/sub&gt;, 2 mM CaCl&lt;sub&gt;2&lt;/sub&gt;, 1 mM MgSO&lt;sub&gt;4&lt;/sub&gt;, 1.25 mM KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, 10 mM D-Glucose; oxygenate with 95% O&lt;sub&gt;2&lt;/sub&gt; / 5% CO&lt;sub&gt;2&lt;/sub&gt; immediately after preparation. Catalog numbers listed separately.</td></tr><tr><td>Blunt microsurgical blade (Circular scalpel)&amp;nbsp;</td><td>Iogen</td><td>374769</td><td>Used for skull surface scraping and cap removal</td></tr><tr><td>Buprenorphine (Temgesic)</td><td>Indivior</td><td>BE-112515</td><td>Opioid analgesic; 0.05 mg/kg s.c. pre-operatively</td></tr><tr><td>C57BL/6JRj mice</td><td>Janvier Labs</td><td></td><td>Male or female; minimum 10 weeks old, &gt;20 g body weight at time of surgery</td></tr><tr><td>Calcium Chloride dihydrate (CaCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Sigma-Aldrich</td><td>C-7902</td><td>2 mM; MW 147.02; 0.29 g/L. Prepared as Ca &amp;times;100 stock. Component of in-house aCSF.</td></tr><tr><td>Carprofen (Rimadyl)</td><td>Zoetis Animal Health</td><td>RXRIM-INJ&amp;nbsp;</td><td>NSAID analgesic; 5 mg/kg s.c. pre-operatively</td></tr><tr><td>Corticosterone</td><td>Sigma-Aldrich</td><td>C2505</td><td>Dissolved in drinking water for 21-day chronic stress / depression model</td></tr><tr><td>Cotton swab</td><td>&amp;nbsp; Fisherbrand</td><td>&amp;nbsp;22-363-157&amp;nbsp;</td><td>Serves as handle for 30 G dura puncture device and for degreasing the skull surface with acetone</td></tr><tr><td>Cyanoacrylate glue &amp;mdash; Loctite 401</td><td>Henkel Adhesives</td><td>Loctite 401</td><td>Thin-viscosity cyanoacrylate; used for skull surface priming, headplate and cap fixation</td></tr><tr><td>Dental drill &amp;mdash; Foredom K.1070</td><td>Foredom</td><td>K.1070</td><td>Flexible-shaft rotary drill; used for craniotomy thinning and pilot hole drilling</td></tr><tr><td>Dexamethasone</td><td>Dopharma</td><td>1DEX004</td><td>Corticosteroid anti-inflammatory; 2 mg/kg s.c. pre-operatively</td></tr><tr><td>D-Glucose</td><td>Sigma-Aldrich</td><td>G-7021</td><td>10 mM; MW 180.20; 1.80 g/L. Prepared as Glucose &amp;times;100 stock. Component of in-house aCSF.</td></tr><tr><td>Drill bit Steel Bur round 1/007W</td><td>Hager &amp;amp; Meisinger</td><td>18089</td><td>Used for the reference site and the cranial window</td></tr><tr><td>Ethanol 80% (v/v) - A12t Dilutus</td><td>Berner</td><td>13221132</td><td>Pre-operative soaking of surgical instruments and surgical laminar hood for sterilization</td></tr><tr><td>Gas Evacuation Apparatus</td><td>RWD Life Science</td><td>R546W</td><td>Scavenges waste anesthetic gases from surgical area</td></tr><tr><td>Gold-plated reference socket</td><td>DigiKey</td><td>310-13-102-41-001000</td><td>Implanted over left cerebellum; serves as ECoG ground/reference</td></tr><tr><td>GRANT Ultrasonic Cleaner &amp;mdash; Model 17002A</td><td>GRANT Instruments</td><td>17002A</td><td>EAN: 4064343191654. Hot-water ultrasonic bath; used with TICKOPUR TR 3 for instrument cleaning</td></tr><tr><td>Heating pad &amp;mdash; Supertech TMP-5B</td><td>Supertech</td><td>TMP-5B</td><td>Maintains body temperature at 37 &amp;deg;C throughout anesthesia</td></tr><tr><td>Isoflurane</td><td>Piramal Healthcare</td><td>66794-017-25</td><td>Inhalation anesthetic; induction 4%, maintenance 1.5&amp;ndash;2.5%</td></tr><tr><td>Kimtech Science, Precision Wipes (Non-fibrous tissue)</td><td>KIMBERLY CLARK</td><td>7552</td><td>Used with pointy tip to dry residual ACSF and assist PDMS membrane adhesion to dura</td></tr><tr><td>Kwik-Cast Sterile Elastomer Sealant</td><td>World Precision Instruments (WPI)</td><td>KWIK-CAST-s</td><td>Model: KWIK-CAST-S. Biocompatible silicone; used to create removable seal around PDMS membrane.&amp;nbsp;</td></tr><tr><td>LED UV Curing Pen 405 nm</td><td>CureUV</td><td>514424-405</td><td>Used to cure UV-activated dental cement (Total C-Ram) during implantation</td></tr><tr><td>Lidocaine-epinephrine solution</td><td>Orion Pharma</td><td>05 10 31</td><td>Local anesthetic; s.c. injection under scalp skin</td></tr><tr><td>Magnesium Sulfate heptahydrate (MgSO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;7H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Sigma-Aldrich</td><td>M-7774</td><td>1 mM; MW 246.48; 0.25 g/L. Prepared as Mg &amp;times;100 stock. Component of in-house aCSF.</td></tr><tr><td>Metal headplate &amp;mdash; Neurotar Model 13</td><td>Neurotar</td><td>Model 13</td><td>Titanium headplate; round opening 11 mm diameter; affixed to skull with cyanoacrylate and dental cement</td></tr><tr><td>NeuroNexus headstage adapter )</td><td>NeuroNexus</td><td>Custom IH256 64&amp;times;4 Molex&amp;nbsp;</td><td>Custom adapter board; sandwiches each &amp;micro;ECoG sub-grid between two Intan RHD 128-ch headstages</td></tr><tr><td>Neuropixels 1.0 probe with metal cap (&amp;times;2)</td><td>IMEC</td><td>PRB_1_4_0480_1_C</td><td>960 recording sites; 10 mm shank. One probe targets hippocampal-thalamic-visual column, one targets mPFC column</td></tr><tr><td>NI PXIe-1083 chassis</td><td>National Instruments</td><td>PXIe-1083</td><td>9-slot PXI Express chassis housing Neuropixels module and NI DAQ cards</td></tr><tr><td>NI PXIe-3839 multifunction I/O module</td><td>National Instruments</td><td>PXIe-3839</td><td>Installed in NI PXIe-1083 chassis; used for synchronization and auxiliary I/O</td></tr><tr><td>NI PXIe-6363 multifunction DAQ</td><td>National Instruments</td><td>PXIe-6363</td><td>Installed in NI PXIe-1083 chassis; analog and digital I/O for synchronization</td></tr><tr><td>Nova Microcinerator -&amp;nbsp; Infrared Sterilizer</td><td>Argos Tech</td><td>S3022</td><td>Used for quick sterilization of any instrument that may touch exposed brain tissues</td></tr><tr><td>Olympus SZ51 stereo binocular microscope</td><td>Olympus</td><td>SZ51</td><td>Surgical visualization; used throughout craniotomy and implantation steps</td></tr><tr><td>Open Ephys Acquisition Board (One Box, 1st generation)</td><td>Open Ephys</td><td>OEPS-9029</td><td>Receives &amp;micro;ECoG data streams via SPI; open-source hardware. See open-ephys.org</td></tr><tr><td>Open Ephys GUI</td><td>Open Ephys</td><td>N/A (open-source)</td><td>Free, open-source electrophysiology acquisition software. Available at: open-ephys.org</td></tr><tr><td>Potassium Chloride (KCl)</td><td>Sigma-Aldrich</td><td>P-5405</td><td>3 mM; MW 74.55; 0.22 g/L. Component of in-house aCSF.</td></tr><tr><td>Potassium dihydrogen Phosphate (KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>P-5655</td><td>1.25 mM; MW 136.09; 0.17 g/L. Component of in-house aCSF (Klebs &amp;times;10 stock).</td></tr><tr><td>Povidone-iodine solution (BETADINE)</td><td>Takeda</td><td>45 70 28</td><td>Topical disinfectant for surgical field preparation</td></tr><tr><td>PXIe acquisition module</td><td>Neuropixels</td><td>PXIE_1000</td><td>PXI-format module for Neuropixels 1.0 data acquisition; installed in NI PXIe-1083 chassis</td></tr><tr><td>Reference socket connector</td><td>DigiKey</td><td>342-10-102-00-591000</td><td>Corresponding mating connector for reference socket above; confirm part number with DigiKey</td></tr><tr><td>RHD 128-channel headstage (&amp;times;4)</td><td>Intan Technologies</td><td>RHD 128</td><td>Two headstages per &amp;micro;ECoG sub-grid; confirm model (e.g., RHD2132 or RHD2164)</td></tr><tr><td>RHD SPI interface cable, 0.9 m</td><td>Intan Technologies / Open Ephys</td><td>C3203</td><td>Standard SPI cable connecting RHD headstages to Open Ephys acquisition box</td></tr><tr><td>RWD Air Pump</td><td>RWD Life Science</td><td>R510-29</td><td>Provides airflow through anesthesia circuit</td></tr><tr><td>RWD Animal Anesthesia Vaporizer</td><td>RWD Life Science</td><td>R5835</td><td>Precision isoflurane vaporizer for induction and maintenance of anesthesia</td></tr><tr><td>Scalpel Blade #11</td><td>Swann Morton</td><td>SM0103</td><td>Used for engraving crisscross pattern on skull surface to enhance adhesive grip</td></tr><tr><td>Small animals hair trimmer - GT416</td><td>Aesculap Schermaschinen GmbH</td><td>D-98528H</td><td>Used to trim the animal scalp before surgery</td></tr><tr><td>Sodium Bicarbonate (NaHCO&lt;sub&gt;3&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>S-5761</td><td>26 mM; MW 84.01; 2.18 g/L. Component of in-house aCSF.</td></tr><tr><td>Sodium Chloride (NaCl)</td><td>Sigma-Aldrich</td><td>S-5886</td><td>124 mM; MW 58.44; 7.25 g/L. Component of in-house aCSF.</td></tr><tr><td>SOL-M (Sterile hypodermic needle 30 G)</td><td>Sol-Millennium Europe</td><td>113005</td><td>Bent 90&amp;deg; at tip and affixed to cotton swab to create dura puncture device</td></tr><tr><td>Standard manipulator arm, 3-axis LEFT (Stoelting)</td><td>Stoelting Co.</td><td>51604</td><td>Used for Neuropixels probe insertion during recording sessions</td></tr><tr><td>Standard manipulator arm, 3-axis RIGHT (Stoelting)</td><td>Stoelting Co.</td><td>51606</td><td>Used for Neuropixels probe insertion during recording sessions</td></tr><tr><td>Stereotactic frame &amp;mdash; RWD F67001</td><td>RWD Life Science</td><td>F67001</td><td>Standard stereotaxic frame; used for head stabilization and coordinate-referenced implant placement</td></tr><tr><td>Surgical Marker skin pen with ruler (Violet Ink)</td><td>Fannin</td><td>SM0372</td><td>Biocompatible gentian-violet based surgical skin marker used to outline the planned craniotomy and implant footprint on bone. Resistant to cyanoacrylate priming (Loctite 401) and remains visible through skull degreaseing and headplae fixation, allowing reliable transfer of stereotactic coordinates onto the bone surface.</td></tr><tr><td>SYLGARD 184 Silicone Elastomer Kit (PDMS)</td><td>Dow (Germany)</td><td>1673921</td><td>Polydimethylsiloxane; cast into thin (&amp;lt;1 mm) membranes; placed on dural surface to protect brain</td></tr><tr><td>Tergazyme enzymatic detergent</td><td>Alconox</td><td>1304-1</td><td>Protease-based cleaner used at 1% (w/v) in lukewarm RO water for post-session electrode cleaning (45 min soak, then triple rinse in purified water).</td></tr><tr><td>Total C-Ram Automix UV Dental Cement (White)</td><td>Itena</td><td>R301950</td><td>Model: TTCRAM-BLC. UV-curable dental resin; used for headplate reinforcement and socket fixation</td></tr><tr><td>Ultrasonic Cleaning Solution (TICKOPUR TR 3)</td><td>&amp;nbsp;Dr. H. Stamm GmbH</td><td>TR-3-913&amp;nbsp;</td><td>Citric-acid-based concentrate; used in ultrasonic bath for post-surgical instrument cleaning</td></tr><tr><td>Viscotears Eye Gel 2 mg/g</td><td>Alcon</td><td>503849</td><td>Carbomer-based lubricating eye ointment; applied to corneas during anesthesia to prevent drying</td></tr><tr><td>Zeiss Cold Light Source KL1500 LCD</td><td>Carl Zeiss</td><td>KL1500 LCD</td><td>Fiber-optic cold illumination for surgical field; prevents thermal damage</td></tr></tbody></table>

surgical protocol for a large, resealable cranial window enabling longitudinal, multi-modal electrophysiology recordings of mouse default mode network

Surgical Protocol for a Large, Resealable Cranial Window Enabling Longitudinal, Multi-Modal Electrophysiology Recordings of Mouse Default Mode Network

The default mode network (DMN) is a central, large-scale brain network implicated in a range of cognitive functions and neuropsychiatric disorders, such as major depressive disorder (MDD). Studying the DMN&#39;s complex dynamics in animal models provides invaluable insights into its function in both healthy and pathological states. However, performing stable, long-term, large-scale electrophysiological recordings from the multiple deep and distributed nodes of the DMN in awake, behaving mice remains a significant challenge.
Here, a novel two-phase surgical protocol is presented to create a large (4 mm &#215; 7.6 mm), durable, and resealable cranial window in mice. The procedure is designed to preserve the integrity of the dura mater, which is paramount for long-term brain health and recording stability.
This window enables simultaneous, repeated, longitudinal recordings from over 1,000 electrodes by combining surface-level micro-electrocorticography (&#181;ECoG) with two high-density intracranial electrode probes, providing unprecedented access to the DMN. This technique provides a robust platform for multi-modal, multi-scale interrogation of network-wide electrophysiological dynamics over several weeks, opening new avenues for investigating the neuroplastic changes underlying the pathophysiology of brain disorders and for evaluating the chronic effects of novel therapeutics.

A successful surgery results in a clear, transparent window over the cortex, with visible vasculature and minimal signs of inflammation or infection. This clarity can be maintained for over 21 days (Figure 1), enabling long-term longitudinal studies. The robustness of the implant is demonstrated by its ability to remain stable even throughout a 21-day chronic corticosterone administration paradigm, a common model for inducing depression-like states in rodents10,<...

This protocol describes a two-phase surgical method to create a large, resealable, dura-sparing cranial window in mice. This technique enables chronic, multi-modal electrophysiological recordings from distributed, deep-brain networks, such as the Default Mode Network, over several weeks.

The default mode network (DMN) is a central, large-scale brain network implicated in a range of cognitive functions and neuropsychiatric disorders, such ...

The Default Mode Network or the DMN is a crucial, large-scale network, implicated in a range of cognitive functions and neuropsychiatric disorders such as depression. Studying the DMN's complex dynamics in animal models provides invaluable insights into its function in both healthy and pathological states. However, a significant challenge has been performing stable, long-term, and large-scale electrophysiological recordings from the multiple deep and distributed nodes, the DMN, in awake, behaving mice.Here, we present a novel two-phase surgical protocol developed in the lab of Eero Castren. This technique creates a large, durable, and resealable cranial window enabling repeated longitudinal recordings from over 1, 000 electrode channels simultaneously. This is achieved by combining surface-level micro-electrocorticography, micro-ECoG, with two Neuropixels probes, allowing unprecedented access to the Default Mode Network.This video will provide a step-by-step guide to this robust surgical procedure from animal preparation and headplate implantation to the creation of the chronic cranial window, ensuring high-quality, long-term data acquisition for network neuroscience research. All procedures shown were approved by the National Experiment Board in Finland and comply with the European directive for the protection of animals used for scientific purposes. Phase 1, Animal Preparation and Headplate Implantation.To perform this procedure, begin by preparing all necessary surgical materials and equipment. Anesthetize the mouse with 4%isoflurane and maintain anesthesia at 1.5 to 2.5%with an oxygen flow rate of 0.5 liters per minute. Shave the fur from the top of the head and place the animal on a controlled heating pad equipped with an internal temperature sensor calibrated to hold body temperature at 37 degrees Celsius throughout the procedure.Apply carbomer eye ointment to prevent corneal drying. Secure the mouse in the stereotactic frame so that the skull lies flat and administer preoperative analgesics and anti-inflammatory agents via subcutaneous injections, Carprofen, Buprenorphine, and Dexamethasone. disinfect the shaved area with povidone-iodine solution and inject lidocaine-epinephrine solution as a local anesthetic under the scalp skin.Make a small transverse incision at the earline and enlarge the incision progressively to fully expose the top of the skull surface. Carefully clean the exposed skull with acetone until all visible periosteum has been removed to ensure strong adhesion of the implant. Use a blunt, circular scalpel blade to remove any residual periosteal and connective tissue fragments that the acetone wash did not fully resolve.With the stereotactic device, mark a 4 millimeters by 7.6 millimeters rectangular area on the skull relative to bregma. The rectangle should extend two millimeters laterally from bregma on both sides, three millimeters rostrally, and 4.6 millimeters caudally. Next, mark the two intracranial probe insertion sites on the right hemisphere.Mark the rostral probe site at 1.66 millimeters anterior and 1.95 millimeters lateral from bregma, and the caudal probe site at 2.2 millimeters posterior and 1.9 millimeters lateral from bregma. Next, engrave a crisscross pattern over all exposed bone surfaces outside the designated window area using a number 11 surgical blade, and produce a shallow but defined grooves approximately 0.2 to 0.4 millimeters deep to form a uniform grid of approximately one by one millimeter squares. Create a fine glue applicator by fitting a sterile 30G needle to the tip of a cotton swab and bend the needle into a V-shape.Dispense cyanoacrylate glue into a disposable plastic weighing boat and dip the applicator into the glue reservoir. Deposit glue over every exposed and engraved bone surface, applying no more than a single deposit per site so that the coverage is thorough, but never excessive. Allow the glue to dry for seven minutes before proceeding.For implanting the reference socket, select the drilling site over the left cerebellum position to avoid visible superficial vessels. Drill the pilot hole in five to ten second bursts using a round steel burr at 20, 000 to 25, 000 rounds per minute until the hole becomes more translucent, revealing a light pink tint. Insert the gold-plated reference socket into the pilot hole, secure it with cyanoacrylate glue, and reinforce the junction with UV-curable dental cement.Cure the dental cement with a LED curing pen light for one minute. Next, place a small quantity of UV-curable dental cement on the apex of the exposed rostral bone and lay the headplate under the skull so that one edge rests on the dental cement scaffolding, cured around the reference socket, and with the opposing edge resting on the fresh rostral cement dab. Make sure the headplate is centered and leveled.Reinforce the structure by applying dental cement around the base of the head plate and circling the engraved bone and the reference socket until a continuous, sealed enclosure is formed around the perimeter of the future cranial window. Lastly, cured the dental cement with the LED curing pen light for one minute. Once the cement has fully cured, allow the mouse to wake up from the anesthesia.The first phase is now complete. Allow the mouse to recover for at least 48 hours before proceeding to the next phase. Phase 2, Chronic Cranial Window Creation.The second phase requires the same tools and medications as the first phase except for the local anesthetic. Additionally, this phase requires a sterile, thin PDMS membrane, cold, artificial cerebrospinal fluid kept on ice, an elastomer silicone sealant, and a custom 3D printed protective cap. At least 48 hours after the first surgery, re-anesthetize the mouse with isoflurane, place it on the heating pad on the stereotactic frame, and administer the preoperative drugs as in the first phase apart from the lidocaine-epinephrine local anesthetic.Begin thinning the bone with the dental drill along the rectangle outline marked in Phase 1, starting at 25, 000 rounds per minute with the drilling device held in a 90 degree angle relative to the bone surface, and perform one or two slightly deeper passes along the rectangle edges to establish an initial cutting groove. Drill shallow grooves into the bone and dental cement adjacent to the cranial window at the two probe insertion coordinates. These grooves serve as the persistent visual landmarks that remain visible after the bone flap is removed and are used to reposition the intracranial probes reproducibly in subsequent electrophysiological sessions.Apply ice-cold, sterile artificial cerebrospinal fluid regularly throughout the drilling to prevent thermal damage to the underlying cortex and to minimize bleeding. Reduce the drill speed progressively to 20, 000 rounds per minute as the bone thins. Continue drilling along the rectangle perimeter until the bone within the outline is approximately 90%thinned.Do not drill completely through the skull. Switch to the fine curved dura hook. Insert the tip under the edge of the thin bone with extreme caution, and slide the hook along the window perimeter, carefully detaching the bone flap from the surrounding skull and underlying tissue.Lift the successfully detached bone flap carefully with the dura hook in a posterior to anterior direction to approximately 35 degree above the skull surface. Grasp the lifted edge with the forceps and sway gently left and right until the plate releases completely. Gently clear the exposed area of coagulated blood with repeated washes of ice-cold ACSF.Place a single sterile precast PDMS membrane directly under the dural surface once bleeding has subsided. When correctly sized, it will cover the window precisely and adhere passively to the dura, holding it flat against the brain. Seal the craniotomy by applying silicone sealant.Fill every crevice between the dental cement enclosure and inner rim of the head plate, fully surrounding and overlapping the edges of the BDMS membrane. Affix a custom 3D printed protective cap onto the headplate with a small quantity of cyanoacrylate glue to protect the window between recording sessions. The mouse is now ready for recovery and should be transferred to its home cage for individual housing to prevent the damage to the implant.A successful surgery results in a clear, transparent window over the cortex, with visible vasculature and minimal signs of inflammation or infection. This clarity can be maintained for over 21 days, enabling long-term longitudinal studies. Raw electrophysiological signals recorded through the chronic window retained high quality across the longitudinal timeline.Here, representative broadband micro-ECoG traces sampled from a posterior retrosplenial grid and simultaneous intracranial probe local field potential traces sampled from a superficial cortical channel are shown side by side for the first recording day and 21 days after. Day 21 recordings exhibit comparable signal amplitude, spectral content, and absence of motion and noise artifacts relative to Day 0 recordings from the same animal, confirming that neither the chronic presence of the PDMS membrane and silicone seal nor the repeated dural punctures introduced detectable degradation of either the surface or the intracranial probe signal quality in superficial cortical layers where degradation would be expected to emerge first. The primary validation of this technique is the acquisition of stable, high-quality multimodal electrophysiological data over time.The resealable window allows for the repeated insertion of probes to record from the same neuronal populations across weeks. Across the alpha-band, phase locking value matrices computed from channels along the caudal high-density intracranial electrode probe show a stable functional architecture between baseline and the Day 21 post-treatment session. This shows reproducible reinsertion to the same cortical location rather than formal tracing of the same individual neurons.Although minor intercession translation of the shank precludes a same unit claim, the aggregate laminar and regional structure of the alpha-band interactions is preserved, supporting that the chronic window permits longitudinal sampling of the same functional circuit. To directly verify that the chronic window supports reproducible laminar targeting of the same deep structures across sessions, current source density or CSD maps were computed from the probes'LFP channels. Stimulus-evoked CSD profiles show that the expected laminar sink source signatures, including the prelimbic cortex and anterior cingulate area are preserved between sessions.The laminar power profile overlays between the two sessions are highly similar across sessions with the Pearson correlation being 0.81. The differences observed in the secondary motor cortex are likely due to movement differences between recordings. Because CSD plotting carries anatomical information, it can provide a within-session non-terminal readout of which brain regions each probe is currently sampling independent of postmortem tissue stainings.Reproducibility of surface micro-ECoG placement across sessions is quantified independently of the intracranial CSD readout. Band-limited spatial power maps computed from the micro-ECoG grid on Day 0 and Day 21 are superimposed, and the pixel-wise correlation of the two maps is computed separately for the rostral and caudal sub-grids. Sub-grid profile correlations remain high and the spatial barcodes of the two sessions visibly co-localized the same cortical power hotspots on both the rostral and the caudal halves of the array.A sinusoidal alignment queue visible through the translucent grid together with the engraved bone grooves at the probe insertion coordinates therefore supports millimeter-scale reproducibility of micro-ECoG placement over the 21 day longitudinal interval. To further validate the technique, immunohistochemical stainings were performed for GFAP and IBA1 in post-mortem brain slices approximately four weeks after the craniotomy surgery. GFAP are expressed by astrocytes, which are upregulated in reactive gliosis such as brain injury or inflammation.IBA1 on the other hand is expressed by microglia, which are more active during neuroinflammatory processes. The validation cohort consisted of animals in which the full two-phase surgery was performed and the cranial window was subsequently reopened three times on postoperative day 0, 21 and 22. However, no micro-ECoG placement nor intracranial probe insertions were performed in this cohort.The window was resealed between sessions as during an electrophysiological recording. This cohort therefore isolates the inflammatory contribution of the chronic window and repeated dural exposure from any probe-induced tissue damage. No significant differences were observed in either of the markers between the no-surgery control group and the vehicle group that underwent surgery.These representative micrographs show no qualitative evidence of reactive gliosis or microglial activation in the region directly underlying the window. However, a slight increase of microglial and astrocyte activation was observed adjacent to the probe trajectory and in the hemisphere of the probe insertion respectively, which could likely be as a result of too-high insertion speed of the intracranial probe. Successful surgery results in a clear, transparent window over the cortex with minimal inflammation.After the chronic treatment period, some brain swelling may occur. This can be managed with careful monitoring and if necessary, administration of additional analgesics. For recordings, simply remove the cap and sealant, place the micro-ECoG grids and Neuropixels probes and begin data acquisition from the awake, behaving animal.In summary, this two-phase surgical protocol provides a reliable and highly effective method for creating a large, chronic cranial window in mice. The procedure's key advantages are its minimal damage to the dura and the large exposure it provides, which is critical for the simultaneous placement of both surface micro-ECoG grids and multiple deep brain Neuropixels probes. This method enables unprecedented longitudinal investigation of network-wide electrophysiological dynamics in the DMN and other large-scale circuits.It opens the door to deeper investigations into neural underpinnings of complex behaviors and the pathophysiology of brain disorders, ultimately aiding in the development of novel therapeutics.

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Research

JoVE Journal

Neuroscience

We describe methods for large-scale recording of multiple single units and local field potential in behaving rodents with silicon probes. Drive fabrication, probe attachment to the drive and probe implantation processes are illustrated in sufficient details for easy replication.

Large-scale Recording of Neurons by Movable Silicon Probes in Behaving Rodents

A procedure is described for manipulating the activity of cerebral cortical pyramidal neurons optogenetically while the electroencephalogram, electromyogram, and cerebral lactate concentration are monitored. Experimental recordings are performed on cable-tethered mice while they undergo spontaneous sleep/wake cycles. Optogenetic equipment is assembled in our laboratory; recording equipment is commercially available.

Simultaneous Electroencephalography, Real-time Measurement of Lactate Concentration and Optogenetic Manipulation of Neuronal Activity in the Rodent Cerebral Cortex

This method creates a tangible, familiar environment for the mouse to navigate and explore during microscopic imaging or single-cell electrophysiological recordings, which require firm fixation of the animal’s head.

Flat-floored Air-lifted Platform: A New Method for Combining Behavior with Microscopy or Electrophysiology on Awake Freely Moving Rodents

This protocol presents a method for creating a large unilateral craniotomy over the temporal and parietal regions of the mouse cerebral cortex. This is especially useful for real time imaging over an expansive area of a cortical hemisphere.

A Large Lateral Craniotomy Procedure for Mesoscale Wide-field Optical Imaging of Brain Activity

Presented here is a protocol for the implantation of a chronic cranial window for the longitudinal imaging of brain cells in awake, head-restrained mice.

Implantation of a Cranial Window for Repeated In Vivo Imaging in Awake Mice

Here, we present the fabrication method of an optrode system with optical fibers for light delivery and an electrode array for neural recording. In vivo experiments with transgenic mice expressing channelrhodopsin-2 show the feasibility of the system for simultaneous optogenetic stimulation and electrophysiological recording.

Optrode Array for Simultaneous Optogenetic Modulation and Electrical Neural Recording

A low-cost electroencephalographic recording system combined with a millimeter-sized coil is proposed to drive transcranial magnetic stimulation of the mouse brain in vivo. Using conventional screw electrodes with a custom-made, flexible, multielectrode array substrate, multi-site recording can be carried out from the mouse brain in response to transcranial magnetic stimulation.

Low-Cost Electroencephalographic Recording System Combined with a Millimeter-Sized Coil to Transcranially Stimulate the Mouse Brain In Vivo

We have developed a simplified and cost-effective approach for electrode fabrication and conducted recordings of signals across multiple regions in freely moving mice. Utilizing optogenetics, alongside multi-region electrophysiology and calcium signal recording, enabled the revelation of neuronal activities across regions in the seizure kindling model.

An Integrated Method for Crafting Flexible and Convenient Electrophysiological Optrodes for Multi-Region In Vivo Recording

This protocol describes a method to obtain in vivo, high-density single-neuron recordings from the brainstem of head-fixed mice. This approach is deployed to measure the action potential firing of neurons in the ventrolateral periaqueductal gray - a brainstem region inactive during Rapid Eye Movement (REM) sleep - before and during general anesthesia.

Acute Single-Unit Multi-Electrode Recordings from the Brainstem of Head-Fixed Mice

This study presents a scalable, reliable, and reproducible method for repeated chronic cortical recordings in a porcine model. The method has applications in various fields of neuroscience, including pain research and neurological disease diagnostics.

Chronic Cranial Window Technique for Repeated Cortical Recordings During Anesthesia in Pigs

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.

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

Learn a two-phase surgical protocol for creating a large, resealable cranial window in mice. This method enables stable, longitudinal electrophysiological recordings from the default mode network using micro-ECoG and Neuropixels probes.

Surgical Protocol for a Large, Resealable Cranial Window Enabling Longitudinal, Multi-Modal Electrophysiology Recordings of Mouse Default Mode Network (Video) | JoVE Journal

Surgical Protocol for a Large, Resealable Cranial Window Enabling Longitudinal, Multi-Modal Electrophysiology Recordings of Mouse Default Mode Network

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