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Construction of Local Field Potential Microelectrodes for in vivo Recordings from Multiple Brain Structures Simultaneously

Published: March 14, 2022 doi: 10.3791/63633
* These authors contributed equally


The present protocol describes the construction of custom-made microelectrode arrays to record local field potentials in vivo from multiple brain structures simultaneously.


Researchers often need to record local field potentials (LFPs) simultaneously from several brain structures. Recording from multiple desired brain regions requires different microelectrode designs, but commercially available microelectrode arrays often do not offer such flexibility. Here, the present protocol outlines the straightforward design of custom-made microelectrode arrays to record LFPs from multiple brain structures simultaneously at different depths. This work describes the construction of the bilateral cortical, striatal, ventrolateral thalamic, and nigral microelectrodes as an example. The outlined design principle offers flexibility, and the microelectrodes can be modified and customized to record LFPs from any structure by calculating stereotaxic coordinates and quickly changing the construction accordingly to target different brain regions in either freely moving or anesthetized mice. The microelectrode assembly requires standard tools and supplies. These custom microelectrode arrays allow investigators to easily design microelectrode arrays in any configuration to track neuronal activity, providing LFP recordings with millisecond resolution.


Local field potentials (LFPs) are the electric potentials recorded from the extracellular space in the brain. They are generated by ion concentration imbalances outside of neurons and represent the activity of a small, localized population of neurons, allowing to precisely monitor the activity of a specific brain region compared to the macroscale EEG recordings1. As an estimate, the LFP microelectrodes separated by 1 mm correspond to two completely different populations of neurons. While EEG signal is filtered by brain tissue, cerebrospinal fluid, skull, muscle, and skin, LFP signal is a reliable marker of local neuronal activity1.

Researchers often need to simultaneously record LFPs from several brain structures, but commercially available microelectrode arrays often do not offer such flexibility. Here, the present protocol describes fully customizable, easily constructed microelectrodes to simultaneously record LFPs from any desired brain region at different depths. Although LFPs have extensively been used to record the neuronal activity of a specific brain region2,3,4,5,6,7,8,9, the current easy customizable design allows recording LFPs from any multiple superficial or deep brain regions11,12. The protocol can also be modified to construct any desired microelectrode array by determining stereotaxic coordinates of the brain regions and assembling the array accordingly. These microelectrodes with a 10 kHz sampling rate and 60-70 kΩ resistance (2 cm length) allow us to record LFPs with millisecond precision. The data can then be amplified by a 16-channel amplifier, filtered (low pass 1 Hz, high pass 5 kHz), and digitized.

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The present work is approved by the University of Virginia Animal Care and Use Committee. C57Bl/6 mice of both sexes (7-12 weeks) were used for the experiments. The animals were maintained on a 12 h light/12 h dark cycle and had ad libitum access to food and water.

1. Microelectrode construction

  1. To construct the microelectrodes, use 50 µm (diameter) diamel-coated nickel-chromium wire (see Table of Materials). Tape one end of the wire at the back of the platform and wrap the wire three times around the nearest knob on the platform (Figure 1A,C).
    NOTE: An acrylic platform with two knobs (2 x 5 inches) was used here, but any platform can be used.
    1. Stretch the wire around the farthest second knob to make two loops between the knobs. Wrap the wire three more times around the first knob to fix the wire in place and tape the end again at the back of the platform.
      NOTE: After the wires are separated (steps 1.2-1.3.1), there must be two wires on each side (four wires in total, Figure 1B).
  2. Place the tension bars under the wires with the tape wrapped around them (sticky side up) (Figure 1C).
    NOTE: Triangular acrylic pieces were used for the tension bars, with tape wrapped around them (sticky side outside to attach the wires). The sticky side of the tape outside of the tension bars will keep the wires in place to adjust the distance between them. The tension bars must be ~2.5 cm away from the knobs, and the wires must not be loose.
  3. Using a microscope and fine forceps, make either a 3 mm or 4.5 mm gap between the wires (3 mm gap between the wires to make cortical (Ctx) - ventrolateral thalamic nucleus (VL) microelectrodes; 4.5 mm gap to make striatal (Str) - nigral (SNR) microelectrodes) (Figure 1B).
    1. If magnification is used on the microscope, ensure to calculate and adjust for the difference in magnification and the actual distance between the wires.
      NOTE: If microelectrodes are constructed for structures other than those used here, the distance between the wires needs to be adjusted to the stereotaxic distance between the structures. Figure 2B provides an example of how the wires will be organized; accordingly, the stereotaxic coordinates for other structures must be adjusted.
  4. Cut four small pieces of plastic (0.5 mm thick) ~6 mm (width) x 3 mm (height) (Figure 1C).
    NOTE: Any plastic pieces can be used as long as they are 0.5 mm thick; here, square tubing was used in which the pins were sold (Pins, see Table of Materials). If a different thickness is used, please add more or fewer pieces of plastic to fit the required stereotaxic coordinates.
  5. Apply glue (see Table of Materials) on the plastic and place them on the wires (Figure 1C). Place plastic pieces ~1.0 cm away from the middle of the wire, which is 1.0 cm away from the tension bar. Remove the excess of superglue with a cotton swab.
  6. After the superglue dries, cut the wires using fine scissors, in the order indicated in Figure 1C.
  7. Cut four 7 mm glass tubes using a commercially available kit (see Table of Materials) and insert the electrode wires into the glass tubes as indicated in Figure 2A.
    1. Insert VL and SNR electrode pairs into the glass tubes.
      NOTE: Only the wires for deep structures need to be inserted into the glass tubes to support surgical implantation. Ensure not to insert cortical electrodes into the glass tube.
  8. Put the glue at the base of the glass tubes to connect them to the plastic. Wait some time until the glue dries.
  9. Cut the glass tubes and wires using a scalpel as indicated in Table 1; ensure that the lengths of the microelectrodes are correct. If the microelectrodes target different structures, adjust the cutting distance according to the required stereotaxic coordinates.

Figure 1
Figure 1: Schematic of the microelectrode construction. (A) Set up of wires on the platform with tension bars below the wires. (B) The gap between the wires. (C) Four pieces of plastic are glued to the wires. Please click here to view a larger version of this figure.

Ctx Str VL SNR
AP (Anterior/Posterior) 2.2 1.2 -1.3 -3.3
ML (Medial/Lateral) 1.8 1.5 1 1.5
DV (Dorsal/Ventral) 0.5 3.5 4 4.75
Electrode length 4 4.75 5.25 6

Table 1: Stereotaxic implantation coordinates and dimensions of the microelectrodes.

2. Microelectrode array assembly

  1. Use the glue to attach plastics in the desired order of target regions. An example for cortical, thalamic, striatal, and nigral electrodes is shown in Figure 2B,C.
    1. Place Ctx-VL electrode pair face down (side with electrode wires has to face down) and connect two 6 mm x 3 mm empty pieces of plastic on top with glue.
    2. On top of the three pieces of plastic, place the second Ctx-VL electrode pair with the electrodes facing upwards (use a microscope and ensure that the VL electrodes are aligned).
      NOTE: Alignment of bilateral electrodes (here, the alignment of left and right VL electrodes) is essential to target desired bilateral structures appropriately.
    3. Use the glue to attach the SNR electrodes on top with the SNR electrodes 2.0 mm away from the VL electrodes and ~5.0 mm away from the cortical electrodes (the SNR electrode wires have to face upwards).
    4. Repeat step 2.1.3. for the other side (the SNR electrode wires have to face outside of the microelectrode array).
  2. Apply epoxy resin around the plastic to bind the electrodes together. Avoid putting epoxy resin on the electrodes.
  3. Take a thick wire and make a loop on one end. Dip the loop in the epoxy solution and place it on the plastic, ensuring the thick wire is lying flat (Figure 2D) so that for the next steps, this wire could be used as a handle. Wait until the electrodes are fully dry.
  4. Cut the wires to 2 cm, as shown in Figure 2E.

Figure 2
Figure 2: Microelectrode construction and dimensions. (A) Four pairs of electrodes formed after the wires were cut with scissors, as indicated in Figure 1C (2 pairs of Ctx-VL electrodes and 2 pairs of Str-SNR electrodes). Insert deep structure electrodes (VL and SNR) into the glass tubes and glue their bases to plastic (red dots). (B) Top view: The electrode pairs from (A) are glued in a stack to create the microelectrode core. Red lines indicate glue lines. (C) Front side view of (B). (D) The thick wire was attached to the microelectrodes. (E) The wires are grouped as indicated, and the isolated ends are scraped off and cut into 2 cm. Please click here to view a larger version of this figure.

3. Microelectrode connection to the headset

  1. Group the wires as indicated in Figure 2E and scrape away 1 mm of the isolated ends with a scalpel.
  2. Bend the cortical electrodes as shown in Figure 3A. Separate the wires as shown in Figure 3B. Using fine forceps, make a loop at the end of each wire (Figure 3B).
  3. Hold a 10-pin headset with a hemostat (see Table of Materials) and use the wooden end of a cotton swab to apply minimal amounts of flux on the pins (Figure 3C). Ensure not to put flux outside the pins to prevent short-circuit among the pins.
  4. Using the wooden end of a cotton swab, apply flux to the wire loops.
  5. Solder the wire loops to the 10-pin headset as shown in Figure 3C. After soldering, dry the headset to prevent short-circuit among the pins.
  6. Take a thin wire (0.005-0.008 inch) for the reference and ground wires and strip off the plastic from one end. Make a loop on the other end of the wire.
  7. Solder the stripped side of the reference and ground wires to their respective pins (Figure 3A,C).
  8. Holding on the thick wire (Figure 2D), apply cranioplasty cement around the microelectrodes, especially where the wires connect to the pins. Avoid touching the actual electrode ends with the cement.
  9. After the cement dries, put epoxy resin at the base of the glass tubes, striatal microelectrode wires, and the whole electrode. Avoid touching the actual electrode ends with epoxy resin. Wait until the electrodes are fully dry.
  10. The electrodes are ready. Drill holes in the mouse's skull (as per the required stereotaxic coordinates) using a dental drill and implant the headset as shown in Figure 3D by lowering the headset with microelectrodes facing the skull and the appropriate holes. The headset could be attached to the stereotaxic arm for support during the implantation.

Figure 3
Figure 3: Microelectrode implantation. (A) The cortical electrodes are bent as indicated. (B) The wires are separated to make loops at the ends. (C) The flux (at the red dots) and looped wires are soldered to the 10-pin headset, ensuring that each wire goes to its appropriate pin. (D) The headset is implanted to record LFPs. Please click here to view a larger version of this figure.

4. Marking electrode location after recordings

  1. At the end of the LFP recordings, confirm the correct position of the electrodes in the target region by applying a current at the electrode tips to make a lesion and waiting for 30 min before perfusing the mouse.
    NOTE: Lesion settings to confirm the location of the electrode tips: Single burst, 40 µA, 0.75 ms monophasic square wave pulse, 50 Hz, 30 s.
  2. Anesthetize the mice with isoflurane (until the mouse fell asleep) and perfuse10 transcardially with 4% paraformaldehyde (PFA) in 0.1 M sodium phosphate buffer. Section the brains (40 µm thick) on a cryostat (see Table of Materials) and stain with DAPI (0.02% in PBS). Confirm the correct location of the electrodes by the presence of the electrode tip lesions as shown in Figure 4B,C.
    NOTE: The percentage of the isoflurane needs to be applied following the individual institution's guidelines.

5. Measuring the electrode resistance

  1. Measure the resistance of the electrodes and check the short-circuit among the electrodes using a multi-range ohmmeter (see Table of Materials). Set the resistance scale at R x 10,000, indicating that a unit deflection in the pointer corresponds to 1 kΩ resistance. 2 cm long electrodes need to have 60-70 kΩ resistance.
  2. Customize the headset by taking ten individual pins (see Table of Materials). Solder each pin with a thin, multi-stranded copper wire in a cable.
    1. Press-fit the soldered pins with their corresponding cable tails into the 10-pin double-row mating socket (matching sockets). Press-fit the open-ended pins of the mating socket into the LFP headset. In this way, each LFP electrode has a designated cable wire in the assembly.
  3. Dip the tip of each LFP electrode (the resistance of which is to be measured) into 0.9% NaCl saline solution (NaCl concentration in the blood). Connect the cable wire end that corresponds to the LFP electrode to the positive terminal of the ohmmeter.
    1. Use a low resistance ( ̴100 Ω) wire with one side in the saline water and the other side as the open end. Connect the open end of the low-resistance wire to the ground of the ohmmeter pointer.
      NOTE: This arrangement completes the circuit and prompts a deflection of the ohmmeter pointer.
  4. Ensure that there is no electrical connection between any two electrodes on the headset. Check the electrical insulation of pairwise LFP electrodes (ground: one cable corresponds to one electrode; positive: another cable corresponds to another electrode). Discard the electrodes if any deflection is observed in this case.

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Representative Results

In this work, the LFP microelectrodes were used to map the seizure spread through the basal ganglia11. Simultaneous LFP recordings were performed from the right premotor cortex (where the seizure focus was) and the left VL, striatum, and SNR (Figure 4). Seizure start was identified as deflection of the voltage trace at least twice the baseline (Figure 4A, red arrow). The power spectrum plot11 shows frequency distributions for the recorded LFPs (Figure 4A). Seizure onset latencies (red bars) could be compared between each structure with millisecond precision (Figure 4A). A current pulse was applied at the end of the recordings to mark and confirm the location of the electrode tips, forming a lesion (Figure 4B,C).

Figure 4
Figure 4: Representative LFP recordings. (A) A seizure was recorded from the right premotor cortex and left VL, striatum, and SNR using LFP microelectrodes with the corresponding power spectrums. The red arrow indicates seizure onset. The red horizontal bars indicate seizure onset delay in each structure. The brain schematic shows the position of the microelectrodes (red dots). (B,C) The structures were lesioned after the recordings to mark the location of the microelectrode tips in the VL and SNR. Please click here to view a larger version of this figure.

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Historically, microelectrode arrays have been extensively used to record neuronal activity from a specific brain region of interest2,3,4,5,6,7,8,9,13. However, our easy microelectrode design allows recording from multiple structures simultaneously11,12. Here, the construction of the cortical, thalamic, striatal, and nigral microelectrodes are described as an example. Investigators can modify the microelectrode design to fit any desired structure by calculating the necessary stereotaxic distances and adjusting the construction accordingly.

For example, we have previously modified the design of these microelectrode arrays to record LFPs in the lamellar and septotemporal direction in the hippocampus12. A 50 µm spacing wire separated adjacent electrodes as four microelectrodes recorded along the hippocampal lamina to prevent cross-contamination of the signal. Although those were not single-unit recordings, each electrode represented a small group of neurons as indicated by the variability of a spike waveform as a function of distance from the cell body.

During the construction, insertion of the thalamic and nigral microelectrode wires into glass tubes was necessary to provide stability during implantation surgery to target those deep structures. There were eight bilateral microelectrodes, four of which had glass tubes (2 VL and 2 SNR), which were a limit before elevating intracranial pressure and increasing mortality. Generally, glass tubes are needed when the desired insertion depth is at least 2 mm.

Also, 0.5 mm thick plastic was needed, limiting the minimum distance separation between the electrodes to 0.5 mm, but other plastics could be used. In the present case, plastics were placed along the major axis of the headset. Plastics can also be placed across the headset axis where several electrodes have identical anterior-posterior (AP) but different medial-lateral (ML) coordinates. This method offers a wide range of possible configurations for specific brain regions.

The number of pins on a headset limits the number of microelectrodes. A headset containing 12 pins covers the anterior-posterior extent of an adult mouse head completely. Each pin should be isolated from the other pins during soldering. An ohmmeter and 0.9% saline water were needed to test the electrical isolation for each pair of electrode terminals. The 12-pin headset limits the recording to 10 regions (2 are reserved for the ground and reference).

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The authors have nothing to disclose.


This work was supported by the National Institute of Health (RO1 NS120945, R37NS119012 to JK) and the UVA Brain Institute.


Name Company Catalog Number Comments
Amplifier 16-Channel A-M Systems Model 3600 Amplifier
Cranioplasty cement Coltene Perm Reeline/Repair Resin Type II Class I Shade - Clear Cement to hold microelectrodes
Cryostat Microtome Precisionary CF-6100 To slice brain
Diamel-coatednickel-chromium wire Johnson Matthey Inc. 50 µm Microelectrode wire
Dremel Dremel 300 Series To drill holes in mouse skull
Epoxy CEC Corp C-POXY 5 Fast setting adhesive
Hemostat Any To hold the headset
Forceps Any To hold microelectrodes
Light microscope Nikon SMZ-10 To see alignment
Ohmmeter Any To measurre resistance
Pins (Headers and matching Sockets) Mill-Max Interconnects, 833 series, 2 mm grid gull wing surface mount headers and sockets To attach microelectrodes to
Polymicro Tubing Kit Neuralynx ID 100 ± 04 µm, OD 164 ± 06 µm, coating thickness 12 µm Glass tubes
Pulse Stimulator A-M Systems Model 2100 To mark the microelectrode location at the end of the recordings
Scissors Any To cut microelectrodes
Superglue Gorilla Adhesive
Thick wire 0.008 in. – 0.011 in. A-M Systems 791900 Tick wire to hold the microelectrode array
Thin wire 0.005 in. - 0.008 in. A-M Systems 791400 Thin wire for reference and ground



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Local Field Potential Microelectrodes In Vivo Recordings Multiple Brain Structures Simultaneous Recordings Easy Microelectrode Design Flexibility Modify Construction Dianal-coated Nickel Chromium Wire Platform Tension Bars Microscope Forceps Cortical Microelectrodes Ventral-lateral Microelectrodes Thalamic Nucleus Microelectrodes Striatal-nidral Microelectrodes Plastic Pieces
Construction of Local Field Potential Microelectrodes for <em>in vivo</em> Recordings from Multiple Brain Structures Simultaneously
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Cite this Article

Brodovskaya, A., Shiono, S.,More

Brodovskaya, A., Shiono, S., Batabyal, T., Williamson, J., Kapur, J. Construction of Local Field Potential Microelectrodes for in vivo Recordings from Multiple Brain Structures Simultaneously. J. Vis. Exp. (181), e63633, doi:10.3791/63633 (2022).

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