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

Summary

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.

Abstract

A low-cost electroencephalographic (EEG) recording system is proposed here to drive transcranial magnetic stimulation (TMS) of the mouse brain in vivo, utilizing a millimeter-sized coil. Using conventional screw electrodes combined with a custom-made, flexible, multielectrode array substrate, multi-site recording can be carried out from the mouse brain. In addition, we explain how a millimeter-sized coil is produced using low-cost equipment usually found in laboratories. Practical procedures for fabricating the flexible multielectrode array substrate and the surgical implantation technique for screw electrodes are also presented, which are necessary to produce low-noise EEG signals. Although the methodology is useful for recording from the brain of any small animal, the present report focuses on electrode implementation in an anesthetized mouse skull. Furthermore, this method can be easily extended to an awake small animal that is connected with tethered cables via a common adapter and fixed with a TMS device to the head during recording.The present version of the EEG-TMS system, which can include a maximum of 32 EEG channels (a device with 16 channels is presented as an example with fewer channels) and one TMS channel device, is described. Additionally, typical results obtained by the application of the EEG-TMS system to anesthetized mice are briefly reported.

Introduction

Transcranial magnetic stimulation (TMS) is a promising tool for human brain science, clinical application, and animal model research because of its non-/low invasiveness. During the early stage of TMS applications, measurement of the cortical effect in response to single- and paired-pulse TMS in humans and animals was restricted to the motor cortex; easily measurable output was limited to motor evoked potentials and induced myoelectric potentials involving the motor cortex^1,2. To expand the brain regions that can be measured by TMS modulation, electroencephalographic (EEG) recording was integrated with single- and paired-pulse TMS as a useful method to directly examine the excitability, connectivity, and spatiotemporal dynamics of areas throughout the whole brain^3,4,5. Thus, the simultaneous application of TMS and EEG recording (TMS-EEG) to the brain has been used to probe various superficial cortical brain areas of humans and animals to investigate intracortical neural circuits (see Tremblay et al.⁶). Moreover, TMS-EEG systems can be used to examine additional cortical spatiotemporal characteristics, including the propagation of signals to other cortical areas and the generation of oscillatory activity^7,8.

However, the mechanism of action of TMS in the brain remains speculative because of the non-invasiveness of TMS, which limits our knowledge of how the brain functions during TMS applications. Therefore, invasive translational studies in animals ranging from rodents to humans are of crucial importance to understand the mechanism of the effects of TMS on neural circuits and their activity. In particular, for combined TMS-EEG experiments in animals, a simultaneous stimulation and measurement system has not been intensively developed for small animals. Therefore, experimentalists are required to construct such a system by trial and error according to their specific experimental requirements. In addition, mouse models are useful among other in vivo animal species models because many transgenic and strain-isolated mice strains are available as biological resources. Thus, a convenient method to build a TMS-EEG-combined measurement system for mice would be desirable for many neuroscience researchers.

This study proposes a TMS-EEG-combined method that can be applied for simultaneous stimulation and recording of the mouse brain, which is the main type of transgenic animal used in research, and that can easily be constructed in typical neuroscience laboratories. First, a low-cost EEG recording system is described using conventional screw electrodes and a flexible substrate to reproducibly assign an electrode-array position in each experiment. Second, a magnetic stimulation system is constructed using a millimeter-sized coil, which can easily be custom-made in typical laboratories. Third, the TMS-EEG-combined system records neural activity in response to sound and magnetic stimulation. The method presented in this study can reveal the mechanisms that generate specific disorders in small animals, and the results obtained in the animal models can be translated to understand the corresponding human disorders.

Protocol

In the present study, all animal experiments were performed following the National Institutes of Health Guide for the Care and Use of Laboratory Animals and with approval from the Institutional Animal Care and Use Committee of Hokkaido University. C57BL/6J mice, two male and three female, 8 to 10 weeks old, were used for the present study. This is a terminal procedure. The animals were obtained from a commercial source (see Table of Materials).

1. Flexible two-dimensional array design and construction

1. Prepare the required number of miniature screw electrodes (stainless, SUS XM7; see Table of Materials) with the following structural properties for use as EEG recording and reference electrodes: nominal diameter, neck length, and head diameter of 0.6 mm, 1.5 mm, and 1.1 mm, respectively (Figure 1A).
   NOTE: In the present study, 16 miniature screw electrodes were used.
2. Prepare a blueprint of a printed circuit diagram on a flexible substrate following the steps below.
   1. Create a two-dimensional (2D) electrode pad pattern on a flexible substrate (whole size, 41.2 mm × 19.9 mm; see Table of Materials) for the screw electrodes to be read out. Design the 2D electrode arrangement. Figure 1B shows the specific arrangements used in this study and the relative coordinates from a baseline point (cross marked at the origin [0, 0]).
      NOTE: In the present study, to record neural activity in the auditory cortex within the temporal lobes, electrode placement in the lateral-to-medial (horizontal) direction was longer than that in the rostral-to-caudal (vertical) direction (Figure 1B).
   2. Ensure that for the EEG recording electrodes, each copper pad (see Table of Materials) on the flexible substrate has a ring shape with an outer diameter of 1.3 mm and an inner diameter of 0.8 mm (Figure 1C, left). Make a small hole (0.8 mm diameter) in the center for each screw electrode to pass through the substrate. For the reference electrodes, each copper pad should have a square shape with a side length of 1.4 mm; similarly, make a small hole (0.8 mm diameter) in the center for each screw to pass through the square pad on the substrate (Figure 1C, right).
   3. Next, to solder a surface-mount connector (Figure 1D, left), design readout pads (2D array) leading to the connector (Figure 1D, right). For example, use a connector with 2 × 10 pins and a 1.27 mm pitch between adjacent pins (Figure 1D, right).
   4. Wire the screw electrode pads and connector pads using both the surface and back layer with a line width of 0.03 mm and line interval of 0.03 mm (thin lines in Figure 1E).
   5. Furthermore, to connect the reference and ground channels to the amplifier, connect the electrode pads for the reference and ground electrodes to the isolated part at the outside of the flexible 2D array (two vertical rectangles indicated by "G" and "HR" at the bottom of Figure 1E). After determining the reference and ground channels, remember to solder the electrode pads to the corresponding connectors (see step 2.1).
   6. Properly design an exposed area that is not covered with a protection layer (polyimide layer). Expose the connector pads in the surface layer while exposing the screw electrode pads in both the surface and back layers. The whole electrode design, sizes, and fabricated flexible 2D array are illustrated in Figure 1E, and the image of a fabricated substrate is shown in Figure 1F.
   7. In the top electrode part (head part) of the flexible 2D array, ensure that the three-layered structure from top to bottom is composed of the following (total thickness of 49.0 µm): a top copper layer (12.0 µm thickness), a middle layer of core polyimide (25.0 µm), and a bottom copper layer (12.0 µm) (Figure 1G, top).
   8. Etch the copper layers on the top and bottom surface of the substrate, for example, using wet etching and the standard fabrication technique⁹.
   9. In the bottom square pad part (connector part) of the flexible 2D array, ensure that the six-layered structure is composed of three layers, including a top copper layer (12.0 µm thickness), a middle core polyimide layer (25.0 µm), and a bottom copper layer (12.0 µm), which are sandwiched by protective polyimide layers, including top and bottom (both 12.5 µm) layers. Attach a 2 mm polyimide board from the bottom as reinforcing material (Figure 1G, bottom).
      NOTE: To maintain flexibility, the reinforcing polyimide board is not mounted to the neck part of the flexible 2D array between the head and connector part.
   10. Similarly, in the connector part, etch the copper and protective polyimide layers on the top using wet etching and the standard fabrication technique.
       NOTE: The total weight of the fabricated, flexible, 2D array device, including the connector, is 0.84 g. After designing a layout for a flexible 2D array, the substrates from a commercial manufacturer (see Table of Materials) are sometimes recommended for convenience.

Figure 1: Component parts of the flexible two-dimensional (2D) array for electroencephalographic (EEG) recording and the fabricated device including the array. (A) The miniature screw electrode that is embedded in the mouse skull. (B) The designed electrode pads for measuring brain activity (green circles) and the reference channel (square on the bottom right). The relative coordinates of the electrode pads from a reference point (cross mark) at the origin (0, 0) are shown; the size in millimeters is illustrated in parentheses. The center coordinates of the electrode pads are symmetrical with respect to the vertical axis passing through the cross mark. (C) The electrode pads and drill holes for a recording electrode (left) and a reference electrode (right) are illustrated. (D) A surface-mount connector (2 × 10 pins) used for the flexible 2D array (left) and the pattern and size of the designed pads on the substrate (right). (E) Designed blueprint with the size of each part in millimeters. (F) Image of a fabricated substrate indicated by the blueprint in E. (G) The layer structure of the flexible 2D array (head and connector parts). The top and side views of the screw electrode pads (top) and readout pads (bottom) are illustrated. The head and the connector parts are composed of a three-layered structure (top) and a six-layered structure (bottom), respectively. Additionally, the neck part is composed of a five-layered structure; a protective polyimide layer is mounted on the top and back surface, and the reinforcing polyimide board is not mounted on the neck part. Please click here to view a larger version of this figure.

2. Adaptor construction and channel mapping

1. Perform adaptor construction following the steps below.
   1. Spread soldering flux on the 2 × 10 pin, surface-mount connector (Figure 1D, left) and the connector pads of the 2D array (Figure 1D, right) (see Table of Materials) on the flexible substrate.
   2. Solder the 2 × 10 pin, surface-mount connector to the connector pads. In particular, confirm the connection between the two pads on the lower part of the 2D array and the two connector pins used as reference and ground channels (Figure 2A).
   3. Connect each of the two pads to individual lead wires to feed baseline signals to an external point (e.g., a ground point connected to the ground channel of the measurement system; Figure 2A).
      NOTE: In this study, however, one of the circular electrode pads with screw electrodes was used as a reference electrode instead of a square electrode in the connector part.
   4. After soldering, cover the soldering points using epoxy resin (see Table of Materials) to protect the exposed points and prevent short circuits.
2. Pin the connector cable and head amplifier following the steps below.
   1. Prepare an insulation-displacement connector (IDC) with 2 × 10 pins and a 1.27 mm pitch (Figure 2B, top left) and a flat 20 pin ribbon cable (see Table of Materials) with a 0.635 mm pitch (Figure 2B, bottom left). Cut the flat ribbon cable to the required length (e.g., 40 cm).
   2. Crimp the IDC and one end of the flat ribbon cable using an IDC crimping tool (Figure 2B, top right) (see Table of Materials).
   3. Separate each line of the other end of the cable up to approximately 15 mm from the tip end using a cutter. Strip the insulation 3 mm from the tip end.
   4. Connect the crimped IDC to the flat ribbon cable and the 2 × 10 pin connector soldered to the flexible substrate (Figure 2C).
   5. Confirm the correspondence between the recording electrode and the separated line of the cable. Ensure that each line used does not produce an incorrect connection error.
   6. Solder the exposed copper wires of the individual lines corresponding to the output of each electrode to the 20 pin connector (1.25 mm pitch) of the measurement system, including the main amplifier (Figure 2B, bottom right).
   7. After soldering, confirm conduction between the screw electrode pads and the connector pins using a testing equipment (e.g., an LCR meter; see Table of Materials).
   8. Cover the soldering points using epoxy resin and shielding tape to protect them from damage and prevent contact with other signal lines.
   9. Using epoxy resin, adhere a thin stainless-steel rod (diameter: 1.1-1.2 mm; length: 100 mm) to the back side of the connector part of the 2D array on the flexible substrate.
      NOTE: This stainless-steel rod can be grasped by a micromanipulator holder during experiments (Figure 2C).
   10. Finally, confirm the mapping between the screw electrodes and signal output channels (Figure 2D).

Figure 2: Constructing the adaptor for a two-dimensional (2D) electrode array on the flexible substrate and recording channel mapping. (A) In the connector part, the reference and ground channels are connected to the bottom electrode pads with lead wires. If the reference and ground channels are determined in advance, the channels should be connected to the corresponding bottom electrode pads during the design phase. In such cases, soldering lead wires to the channels and electrode pads is unnecessary. (B) Insulation-displacement connectors (top left) are crimped to one end of the flat cable (bottom left) to link the measurement amplifier connector (top right). All lines that correspond to the channels to be used are soldered to the green connectors (bottom right). In this case, because each green connector connected to the head amplifier is assigned for an eight-channel measurement, at least two connectors are needed to record 16-channel brain activity signals.The soldered points are covered with epoxy resin and shielding tape to prevent contact with other signal lines. (C) The connector and fabricated cable are placed on the surface of the flexible 2D array substrate. The thin stainless-steel rod is attached to the back side of the flexible substrate. (D) The spatial locations of recording channels on the mouse brain surface and the channel maps for each point for the measurement system are shown. In this case, there are 16 recording channels with screw electrodes (red circles), although the total number of possible recording sites is 32. The other 16 non-recording channels are also shown as green circles on the brain surface. In the mapping plot, "G" and "R" indicate the channels designed for ground and reference electrodes, respectively. Please click here to view a larger version of this figure.

3. Animal surgery

1. Prepare the sterile surgical environment.
   1. Wear protective equipment, such as latex gloves, during the whole experimental procedure involving animals.
   2. Sterilize the stereotaxic apparatus and surgical instruments (see Table of Materials).
   3. After sterilizing the surgical instruments, wash them using sterile saline.
2. Anesthetize the animals.
   1. Measure the weight of the mouse before surgery. Administer atropine sulfate (0.04 mg/kg; see Table of Materials) via intraperitoneal injection.
   2. Anesthetize the mouse via an intraperitoneal injection of a mixture of medetomidine (0.3 mg/kg), midazolam (4.0 mg/kg), and butorphanol (5.0 mg/kg).
   3. Confirm the anesthetic depth by the level of response via pinching the toe.
      NOTE: The anesthesia will wear off after approximately 40 min. If the mouse responds to a toe pinch, administer the same dose of the anesthetic mixture via an intraperitoneal injection.
3. Pre-prepare for the electrode implantation surgery.
   1. Cut the mouse whiskers to prevent haptic sensation.
   2. Lubricate both eyes with an ophthalmic ointment to prevent drying. Close the eyelids to obstruct the sense of sight and maintain closure by adhering the upper and lower eyelids with mending tape.
   3. Shave the hair on the head of the mouse with electric clippers. Insert a thermometer into the rectum and maintain the body temperature at 37 °C using a heating pad.
   4. Administer lidocaine hydrochloride as a topical local anesthetic to the part of the mouse scalp that will be incised.
   5. Incise the mouse scalp using a scalpel or surgical scissors in a rostral-to-caudal direction (area size: 7 × 10 mm²).
   6. Pinch the scalp near the incised part with tweezers and lift. Remove the visible membrane on the skull using a scalpel or surgical scissors. Do not break the blood vessels around the eyes during the operation.
   7. Grasp the skin near both centers of the incision line of the scalp with forceps and widen the incised part to widely expose the top of the skull.
   8. Confirm the complete removal of all membranes on the surface of the skull and the tissue around lambda with surgical scissors.
   9. Wet the skull surface with physiological saline to enhance the visibility of the brain surface under the skull and locate the transverse sinus.
      ​NOTE: When implanting screw electrodes into the skull, remember not to embed them above and into the transverse sinus.

4. Electrode implantation

1. Attach the stainless-steel rod mounted to the 2D electrode array on the back side of the flexible substrate to a micromanipulator. Place the flexible substrate on the skull.
2. Adjust the location of channels (Chs) 3 and 14 (Figure 2D) on the array to fit within the inferior colliculus.
   NOTE: The inferior colliculus is located along the transverse sinus. We recommend confirming the location of the inferior colliculus using a mouse brain atlas in advance.
3. Draw small circles at the locations of Chs 3, 8, 9, and 14 (Figure 2D) on the skull with a permanent marker to use as targeting landmarks.
4. Dry the skull surface to enhance adherence to the dental cement and to electrically isolate the 2D electrode array on the flexible substrate from the mouse skull.
5. Apply dental cement (approximately 1 mm thickness; see Table of Materials) to the skull surface. After applying the dental cement, wait approximately 30 min for it to cure.
6. Align the flexible substrate according to the small circular marks on the surface of the skull.
7. Align the tip of a dental drill to each electrode pad hole on the flexible substrate. Drill carefully into the skull through each of the electrode pad holes.
8. Screw each of the miniature screw electrodes through the drilled holes in the skull using a dedicated screwdriver for miniature screws.
9. Crimp the head of the screw electrode and the electrode pad tightly. Finally, measure the conductance between each screw electrode and the connector with testing equipment (e.g., an LCR meter) to confirm electrical conductivity.

5. Small coil design and construction

1. Design a donut-shaped disk (see Supplementary Coding File 1) with a hole in the center (inner diameter: 2 mm; outer diameter: 7 mm; thickness: 1 mm) using computer-aided design (CAD) software (see Table of Materials).
2. Using a 3D printer, print two disks (Figure 3A, left) made of non-heat-resistant material (e.g., poly-lactic acid filament); non-heat-resistant material is not always necessary (see below).
3. Trim a permalloy-45 rod (diameter: 2 mm; see Table of Materials) to form a short shaft (length: 60 mm).
4. Insert the shaft into each hole of the two 3D-printed disks (Figure 3A, right). Place one disk at the end of the shaft and the other 11 mm from the end, resulting in a 10 mm distance between the two disks. Adhere the disks with instant glue (see Table of Materials).
5. Attach the end of the shaft without a disk to an impact driver (Figure 3B). Attach a small magnet to the permalloy-45 shaft. Place a hall-effect sensor near the magnet 5 mm from the shaft. Connect the hall-effect sensor to a data acquisition (DAQ; see Table of Materials) system.
6. To count the number of turns, prepare a computer program (see Table of Materials) that analyzes output signals from the hall-effect sensor through the DAQ system.
7. Connect a thin copper wire (diameter: 0.16 mm) to the shaft and adhere to the top end of the wire with instant glue.
8. Using the impact driver, wind the copper wire for 1,000 turns between the two disks. Although the rotation speed is arbitrary, approximately 5 rotations per second is typically used. Then, adhere to the wound wire with instant glue.
9. Detach the two disks from the shaft. If the disks are adhered strongly to the shaft, melt the disks using a heat gun.
10. Cover the coil with epoxy resin to insulate and secure the surface. Then, cut off the unwound shaft portion as excess.
11. Ensure that the obtained coil has height of 10 mm and a diameter of 6 mm (Figure 3B, left). For coil manipulation, either construct a coil holder (Figure 3C, right) or adhere a stainless-steel rod to the coil (not shown here).
12. Measure the resistance and inductance of the coil using an LCR meter (see Table of Materials). For example, the coil used here had a direct current (DC) resistance of 18.3 Ω and an inductance of 7.9 mH at 1 kHz alternating current (AC) input. The AC properties (resistance and inductance) are shown in Figure 3D.
13. Use a function generator to apply a bipolar square wave to the coil. The typical amplitude of the input voltage is 20 V through a bipolar power supply with a 10x gain, following a 2 V generator output. The resulting waveform is a bipolar square wave with an approximate amplitude of 20 V (i.e., a peak-to-peak voltage of 40 V) (Figure 3E).
14. Measure the magnetic flux density using the hall-effect sensor and the DAQ system. In this case, for example, the magnetic flux density (B) of the coil was 113.6 ±2.5 mT (mean ± SEM) when the coil bottom was in contact with the hall-effect sensor (Figure 3F).

Figure 3: Small coil for magnetic stimulation. (A) Three-dimensional (3D)-printed disk (left). Two identical disks are adhered to the permalloy-45 shaft; one is at the end of the shaft, and the other is 10 mm away (right). (B) Setup for winding the coil. The 60 mm shaft with the two disks is attached to an impact driver. A hall-effect sensor is placed near the small magnet attached to the shaft. The copper wire is wound between the two disks. (C) Constructed coil. The coil is 10 mm in height, 6 mm in diameter, and has 1,000 turns of copper wire. The right side of the figure shows the coil manipulated by a 3D-printed coil holder. (D) AC properties of the coil recorded by an LCR meter: (top) resistance versus frequency of sinusoidal input; (bottom) inductance versus input frequency. A typical coil has a resistance and inductance of 21.6 and 7.9 mH, respectively, at 1 kHz of AC input. (E) Biphasic rectangular waveform used as the coil input recorded by an oscilloscope. (F) Relationship between magnetic flux density and the distance between a constructed coil and the hall-effect sensor. The magnetic flux density was recorded by five different hall-effect sensors, once for each sensor. The average of five measurements is plotted, and error bars represent the standard errors of the mean. Please click here to view a larger version of this figure.

6. Signal recording system and procedure

1. Connect the flexible 2D array to the recording system (see Table of Materials) with the flat ribbon cable.
2. Attach the stainless-steel rod mounted on the coil to a micromanipulator (see Table of Materials).
3. Place the coil above bregma and adjust the position in the caudal direction to locate the focal point above the inferior colliculus. The focal point of the emitting electric field is the midline of the wound area on the bottom surface of the coil (i.e., 1 mm from the edge to the center).
4. Prepare a stimulation system consisting of a bipolar power supply and a function generator (see Table of Materials) and connect the coil to the system.
5. Connect a cable between the input terminal of the function generator and the output terminal of the DAQ system to apply trigger signals to the function generator from the DAQ system. Prepare an appropriate computer program for trigger signals to initiate stimuli. Additionally, connect the DAQ system to the recording system to save the stimulation times as timestamps.
6. Start the acquisition process for the recording system.
   NOTE: If the recording system is picking up noise, find the source of the noise and reduce it.
7. Test the magnetic stimulation by triggering the stimulation system.
   NOTE: If the noise produced by the magnetic stimulation saturates the measurement range, adjust the range properly. Additionally, confirm that the recording system saves the stimulation timestamps properly.
8. Start recording the response data and begin stimulation sessions. Stop the recording when each stimulation session is complete. Save all recorded data for subsequent analysis.
   ​NOTE: To perform all experimental conditions with five different magnetic intensities, for example, the total time required for all sessions was about 75 min. The endpoint was usually determined after all the recording sessions were over. However, when the animals showed clinical signs including coughing, labored breathing, and gasping, the experimental session was immediately terminated. For euthanasia, decapitation was performed using sharp, clean scissors while animals were under anesthesia.

7. Data analysis

1. Filter the wideband (raw) signal using a low-pass filter with a cut-off frequency of 200 Hz.
2. Collect filtered waveforms during a time window around each stimulation timestamp. Average the waveforms to obtain the event-related potential (ERP) waveforms (Figure 4 and Figure 5).

Representative Results

Sample EEG data recorded in anesthetized C57BL/6J mice with the flexible substrate combined with the screw electrodes are presented below.

As a typical example, the average EEG waveforms generated in response to sound stimulation (8 kHz tone-burst, 80 dB sound pressure level [SPL]) are shown for 60 trials with identical stimuli (Figure 4A). A schematic of recording channel mapping is also presented in the middle of Figure 4A. The responses from Chs 5, 7, 10, and 12 are recorded from areas near the auditory cortex in both temporal lobes. In the individual EEG waveforms of the channels located around the auditory areas (the inferior colliculus and auditory cortex), the responses excluding the stimulation artifacts were first negative-going immediately after sound stimulation onset (e.g., Chs 3 and 10); the peak amplitudes were 45.6 ± 4.0 µV and25.6 ± 1.5 µV, respectively. The responses were subsequently positive-going to some extent over the baseline (Figure 4B,C) and oscillating while dampening. In contrast, responses from other channels were nearly independent of the stimulation onset, although some channel waveforms showed similar responses.

Figure 4: Sound event-related potential (ERP) waveforms at 16 sites in the mouse brain. (A) In response to sound (8 kHz tone-burst, 80 dB SPL) stimulation applied to an anesthetized mouse, 16-channel ERP waveforms are illustrated. The schematic of a mouse brain is shown in the center, and the 16 recording sites (red circles) on the mouse brain surface are indicated by channel numbers. In this case, 16 recording channels are used; the other 16 non-recording channels are shown as green circles. (B) Expanded views of ERP waveforms for Ch 3. (C) Expanded views of ERP waveforms for Ch 10. Please click here to view a larger version of this figure.

Similarly, the average waveforms of EEG recordings in response to short magnetic stimulation (V_in = 60 V_pp) of the area near the right inferior colliculus are shown for 60 trials with identical stimuli in Figure 5A. A schematic of recording channel mapping is also presented in the middle of Figure 5A. Because the stimulation coil was located near the area of Ch 14, the stimulation artifact was largest at that channel. However, relatively large stimulation artifacts were observed for most channels immediately after stimulation onset, indicating that the magnetic stimulation influenced all recording sites. Because the responses from Chs 5, 7, 10, and 12 were recorded from areas near the auditory cortex in both temporal lobes, the individual EEG waveforms excluding the stimulation artifacts were first negative-going and then positive-going to some extent, depending on the channel positions (Figure 5A-C). Near the auditory areas, response time courses induced by magnetic stimulation were different from those induced by sound stimulation. For Chs 3 and 10, for example, the responses were negative-going immediately after sound stimulation onset, although the peak amplitudes were 58.8 ± 4.0 µV and 28.2 ± 2.0 µV, respectively. Furthermore, with increasing magnetic stimulation intensities, the peak amplitudes of driven responses for Ch 10 were increased (Figure 5D), suggesting that the magnetic stimulation affected evoked neural responses.

Figure 5: Transcranial magnetic stimulation (TMS)-driven event-related potential (ERP) waveforms at 16 sites in the mouse brain. (A) The 16-channel ERP waveforms in response to TMS (V_in = 60 V_pp) applied to an anesthetized mouse are illustrated. A schematic of a mouse brain is shown in the center, and the 16 recording sites (red circles) on the mouse brain surface are indicated by the channel numbers. (B) Expanded views of ERP waveforms for Ch 3. (C) Expanded views of ERP waveforms for Ch 10. (D) Summary for the amplitudes of Ch. 10 ERPs evoked by different magnetic intensities (input voltage). For statistical analysis, an ANOVA for multiple comparisons followed by a post-hoc Tukey-Kramer test is used. * and *** represent p < 0.05 and p < 0.001, respectively. The trial number for a session is 60 times for each condition of individual animals. The statistics are calculated for the samples obtained from two animals. Please click here to view a larger version of this figure.

This method can also be easily extended to an awake small animal that is connected with tethered cables via a common adapter and fixed with a TMS device to the head during recording (Supplementary Figure 1 and Supplementary Figure 2).

Supplementary Figure 1: Fixture of the stimulation coil attached to a mouse skull. (A) For an awake mouse, a stimulation coil fixed with the fixture attached to the mouse skull is shown. (B) Event-related potentials (ERPs) of the awake mouse were recorded in an acryl box, where the mouse could move inside the box. Please click here to download this File.

Supplementary Figure 2: Waveforms of sound-driven and transcranial magnetic stimulation (TMS)-driven ERPs at 16 sites from the brain of an awake mouse. (A) In response to sound stimulation (8 kHz tone-burst, 80 dB SPL) applied to an awake mouse in an acryl case (Supplementary Figure 1B), 16-channel ERP waveforms are illustrated. The schematic of a mouse brain is shown in the center, and the 16 recording sites (red circles) on the mouse brain surface are indicatedby channel numbers. In this case, 16 recording channels are used; the other 16 non-recordingchannels are shown as green circles. (B) Similarly, 16-channel ERP waveforms in response to TMS (V_in = 60 Vpp) applied to the same awake mouse are illustrated. A schematic of a mouse brain is shown in the center, and the 16 recording sites (red circles) on the mouse brain surface are indicated by the channel numbers. The stimulation coil is located near the area of Ch 14. Please click here to download this File.

Supplementary Coding File 1: CAD data file for the donut-shaped disk required for the coil construction. Please click here to download this File.

Discussion

This study addresses a multi-site EEG recording system combined with a magnetic stimulation system designed for small animals, including mice. The constructed system is low-cost and easily constructed in physiological laboratories, and can extend their existing measurement setups. The surgical procedure necessary to obtain data from the mouse recording system is profoundly simple if such laboratories have previous experience with standard electrophysiological experiments.

One advantage of using this approach is the good reproducibility of electrode placement on an individual animal's head and scalp. The flexible substrate used to assign screw electrodes to brain target sites is easily replicated using standard microfabrication techniques, and the same substrates are also convenient for determining the recording sites from the scalp of each animal. In addition, the shape of the electrode array can be easily modified to optimize various experimental needs; customized electrode arrangements can be optimally created for specific experimental purposes. If the method stated in the protocol is followed, screw electrodes, connectors, cables, and surgical procedures can be easily modified and extended to a measurement system with a larger number of recording sites. A second advantage of this recording system is its low cost when laboratories are equipped with a multichannel amplifier. The present recording system can obtain neural signals from 32 input channels and up to four separate cables. Therefore, an extended 32-channel recording system would require extra cables, screw electrodes, and modified flexible substrates, and this expanded system would have a very low cost.

However, one drawback of this methodology is the precise control of the depth of screw electrodes during implantation. However, this drawback is always present for typical screw EEG electrodes, and the precise depth of the screws premortem relative to the cortical surface is unknown. Furthermore, in this system, another critical point for the recording quality of EEG signals and reducing the noise level is appropriate electrode contact with the epidural layer. We always confirm appropriate electrode contact of all screw electrodes through impedance measurement. Typically, an impedance of 5-10 kΩ at 1 kHz suggests appropriate epidural placement, and the impedance values should be confirmed before neural signal measurement.

Additionally, in the current protocol, dental cement is applied to the skull before electrode implantation. The appropriate amount of dental cement can affect the success of EEG signal recordings. That is, a thin layer of dental cement on the skull does not support the implanted electrodes or fix the position of the electrodes, whereas a thicker layer prevents proper positioning of the electrode(s) to contact the dura matter. To determine the appropriate thickness of the layer, we measured the thickness of the dental cement using a digital caliper after successful EEG recordings. The average thickness of an appropriate cement layer was 0.7 mm, suggesting that the dental cement layer could be replaced with a "skull cap" with a 0.7 mm thickness and small holes for screw electrodes.

Magnetic stimulation is a useful tool in human and animal studies for minimally invasive or noninvasive neurostimulation of the brain. Rapidly changing the currents in a coil creates a magnetic field around the coil and causes hyperpolarization or depolarization of neuron membranes when the currents pass through animal and human skulls. For animal models, action potential responses are directly kindled by the suprathreshold magnitude of the electric field change, whereas subthreshold changes in neural membranes are produced to attune the network activity of neural populations¹⁰. This coil is simulated to produce an electric field that is more than 10 V/m, up a depth of 1.8 mm from the surface of the brain (2.4 mm from the skull), corresponding to cortical layer 5/6 or deeper regions in a typical (e.g., C57BL/6J) mouse¹⁰. These millimeter-sized coils are capable of inducing suprathreshold neural activity and can even generate a more localized electric field on the surface of the brain compared to those induced by previously reported coils¹¹. Although added effects consisting of several factors, including perceived sound, skull vibration, and the thermal effect, cannot be completely excluded, these individual effects had a little influence on neural activity. Moreover, as a magnetic core, we use permalloy, whose magnetic properties usually depend on the conditions of the annealing process, including the cooling rate, annealing temperature, and holding time¹². However, its annealing conditions could not be controlled as it was a commercial permalloy.

Recently, combined measurement systems consisting of multi-site EEG recording and TMS have been used in medical studies, and their clinical applications have been increasing^4,6. Our proposed approach will improve small animal models (particularly mouse models) of human neurophysiology, which can provide a much easier translation of experimental rodent model results to human clinical counterparts by offering animal models that better parallel human systems. Finally, using multi-site recording techniques in genetically modified mice, combined magnetic and pharmacological interventions in animals with sensory hearing loss could help to reveal the mechanisms generating specific auditory disorders and tinnitus, which are our future research targets.

Materials

Name	Company	Catalog Number	Comments
3D printer	Zhejiang Flashforge 3D Technology Co., Ltd	FFD-101	The printer used for 3D-printing the donut-shaped disks
ATROPINE SULFATE  0.5 mg	NIPRO ES PHARMA CO., LTD.	-	Atropine sulfate
Bipolar amplifier	NF Corp.	KIT61380	For amplifying waveforms for coil input
Butorphanol	Meiji Seika Pharma Co., Ltd., Tokyo, Japan	-	For anathesis of animals
Commercial manufacturer of flexible 2D array	p-ban.com Corp.	-	URL: https://www.p-ban.com/
Computer prograom to analyze output signals	Natinal Instruments	NI-DAQ and  NI-DAQmx Python	To analyze output signals from the hall-effect sensor
Connector	Harwin Inc.	G125-FV12005L0P	For connector to conect to the measuring system
Copper pad	p-ban.com Corp.	copper	Copper pad on each substrate
Copper wire	Kyowa Harmonet Ltd.	P644432	The windings of the coil
DAQ board	National Instruments Corp.	USB-6343	For measuring the magnitic flux density of the coil
Dental cement	SHOFU INC.	Quick Resin	Self-Curing Orthodontic Resin
ECoG electrode	NeuroNexus Inc.	HC32	For reference to design of the flexible 2D array
Epoxy resin	Konishi Co. Ltd.	#16123	For coil construction
Ethyl Carbamate	FUJIFILM Wako Pure Chemical Corp.	050-05821	For urethan anesthesia
Flat ribbon cable	Oki Electric Cable Co., Ltd.	FLEX-B2(20)-7/0.1 20028 5m	For cable to connect between surface-mount connector and measuring sysytem
flexible substrate	p-ban.com Corp.	polyimide	Baseplate of flexible substrate
Function generator	NF Corp.	WF1947	For generating waveforms for coil input
Hall-effect sensor	Honeywell International Inc.	SS94A2D	For measuring the magnitic flux density of the coil
IDC crimping tool	Pro'sKit Industries Co.	6PK-214	To crimp the IDC and one end of the flat ribbon cable; Flat cable connector crimping tool
Instant glue	Konishi Co. Ltd.	#04612	For coil construction
Insulation-displacement connector (IDC )	Uxcell Japan	B07GDDG3XG	2 × 10 pins and a 1.27 mm pitch
LCR meter	NF Corp.	ZM2376	For measuring the AC properties of the coil
Manipulator	NARISHIGE Group.	SM-15L	For manipulating the coil
Medetomidine	Kobayashi Kako, Fukui, Japan	-	For anathesis of animals
Midazolam	Astellas Pharma, Tokyo, Japan	-	For anathesis of animals
Miniature screw	KOFUSEIBYO Co., Ltd.	S0.6*1.5	For EEG-senseing and reference electrode
Mouse	Japan SLC, Inc.	C57BL/6J (C57BL/6JJmsSlc)	Experimental animal
Permalloy-45 rod	The Nilaco Corp.	780544	The core of the coil
Recording system	Plexon Inc.	OmniPlex	For EEG data acquisition
Stainless wire	Wakisangyo Co., Ltd.	HW-136	For grasp by manipulator
Stereotaxic apparatus	NARISHIGE Group.	SR-5M-HT	To fix a mouse head
Surface-mount connector	Useconn Electronics Ltd.	PH127-2x10MG	For connector to mount on the flexible 2D array
Testing equipment (LCR meter)	NF Corp.	ZM2372	Contact check and impedance measurements
White PLA filament	Zhejiang Flashforge 3D Technology Co., Ltd	PLA-F13	The material used for 3D-printing the donut-shaped disks
Xylocaine Jelly 2%	Sandoz Pharma Co., Ltd.	-	lidocaine hydrochloride