Electrophysiology of Laminar Cortical Activity in the Common Marmoset

Summary

Custom-built micro-drives enable the sub-millimeter targeting of cortical recording sites with linear silicon arrays.

Abstract

The marmoset monkey provides an ideal model for examining laminar cortical circuits due to its smooth cortical surface, which facilitates recordings with linear arrays. The marmoset has recently grown in popularity due to its similar neural functional organization to other primates and its technical advantages for recording and imaging. However, neurophysiology in this model poses some unique challenges due to the small size and lack of gyri as anatomical landmarks. Using custom-built micro-drives, researchers can manipulate linear array placement to sub-millimeter precision and reliably record at the same retinotopically targeted location across recording days. This protocol describes the step-by-step construction of the micro-drive positioning system and the neurophysiological recording technique with silicon linear electrode arrays. With precise control of electrode placement across recording sessions, researchers can easily traverse the cortex to identify areas of interest based on their retinotopic organization and the tuning properties of the recorded neurons. Further, using this laminar array electrode system, it is possible to apply a current source density analysis (CSD) to determine the recording depth of individual neurons. This protocol also demonstrates examples of laminar recordings, including spike waveforms isolated in Kilosort, which span multiple channels on the arrays.

Introduction

The common marmoset (Callithrix jacchus) has quickly grown in popularity as a model to study brain function in recent years. This growing popularity is due to the accessibility of the marmoset's smooth cortex, the similarities in neural functional organization with humans and other primates, and the small size and fast breeding rate¹. As this model organism has grown in popularity, there has been rapid development in the neurophysiological techniques suited for use in the marmoset brain. Electrophysiology methods are widely used in neuroscience to study the activity of single neurons in the cortex of both rodents and primates, resulting in unparalleled temporal resolution and location access. Due to the relative novelty of the marmoset monkey as a model of visual neuroscience, the optimization of awake-behaving electrophysiology techniques is still evolving. Previous studies have shown the establishment of robust protocols for electrophysiology in anesthetized preparations², and early awake-behaving neurophysiology studies have shown the reliability of single-channel tungsten electodes³. In recent years, researchers have established the use of silicon-based microelectrode arrays for awake-behaving neurophysiology⁴. However, the marmoset poses unique targeting challenges due to its small brain size and lack of anatomical landmarks. This protocol outlines how to construct and use a micro-drive recording system suitable for the marmoset that allows for the recording of large populations of neurons with silicon linear arrays while producing minimal tissue damage.

Working with the marmoset poses a challenge due to the smaller scale of the retinotopic maps in the visual cortex as compared to larger primates. A slight shift of the electrodes by just 1 mm can result in significant changes in the maps. Moreover, researchers often need to alter the placement of the electrodes between the recording sessions to obtain a broader range of retinotopic positions in the visual cortex. Current semi-chronic preparations do not allow for the adjustment of the electrode positioning daily or with enough precision to target specific locations at sub-millimeter scales⁵. With this in mind, the proposed micro-drive system utilizes an X-Y electrode stage that mounts a lightweight micro-drive to a recording chamber and allows for the sub-millimeter targeting of cortical sites. The moveable X-Y stage components allow for vertical and horizontal movement of the linear array in order to traverse the cortical areas systematically, which is required to identify areas of interest (via retinotopy and tuning properties). Across recording sessions, researchers can also manually adjust the X-Y stage to shift the targeted sites within the area. This is a key advantage over alternative techniques using semi-chronic recording preparations, which do not have easy electrode targeting mechanisms.

The micro-drive is a versatile tool that enables the attachment of various silicon arrays to be mounted for lowering into the cortex. In this protocol, a custom probe with two 32-channel linear arrays spaced 200 µm apart was used for the investigation of laminar circuits spanning the cortical depth. Most methods for probing the neural circuitry typically sample the electrical potentials or single units averaged across all the layers of the cerebral cortex. However, recent research has revealed intriguing findings about cortical laminar microcircuits⁶. By utilizing the micro-drive, researchers can use laminar probes and make fine adjustments to the recording depth to ensure comprehensive sampling across all the layers.

This system can be constructed with commercially available components and is easily modified for different experimental techniques or probes. The key advantages of this preparation are the ability to change the X-Y recording position with sub-millimeter precision and to control the depth of the recording within the cortex. This protocol presents step-by-step instructions for constructing the X-Y stage micro-drive and neurophysiology recording techniques.

Protocol

The experimental procedures followed the National Institutes of Health Guide for the Care and Use of Laboratory Animals. The protocols for the experimental and behavioral procedures were approved by the University of Rochester Institutional Animal Care and Use Committee.

1. Construction of the micro-drive containing the electrode for recording (Figure 1)

NOTE: Custom-built X-Y stages holding multi-channel linear silicon arrays allow for sub-millimeter targeting of the recording sites.

1. Gather all the micro-drive parts outlined in Figure 1. Use clear acrylonitrile butadiene styrene (ABS) plastic to 3D print the custom parts. This includes the protective sleeve, the base, the X stage, and the Y stage. Designs for 3D parts are available online (https://marmolab.bcs.rochester.edu/resources.html). Additionally, gather the electrode, plastic rectangular platform, grid, micromanipulator, steel tubing, and six stainless steel screws (screw size: 00).
   1. Using a rotary tool with a diamond wheel cutting attachment, cut out a 4 mm x 3 mm x 3 mm section of the plastic grid with 1 mm x 1 mm hole spacing.
   2. Use epoxy to fix the small cut-out grid section to the top of the Y-stage. Then, fit the micro-drive over that grid with a screw. Add epoxy at its base for stability.
   3. Attach the small 3D printed rectangular platform to a 28 G steel tube with epoxy. This will be used in future steps to hold the silicon electrode.
   4. Thread a 23 G guide tube through the grid. Then, thread the steel tube attached to the plastic platform through the 23 G guide tube. Place the 28 G steel tube into one of the slots of the micro-drive.
   5. Then, assemble the 3D parts (base, X stage, Y stage) using stainless steel hex screws (size: 00, length: 1/8 in) to build the base for the drive.
      1. First, use a hand-tapping tool to tap the premade screw holes in both the base and X stage. Then, start by inserting four screws through the long slots in the X stage and securing them to the corresponding screw holes premade in the base.
      2. Once the moveable X stage is secured to the base, insert two screws through the long slot in the Y stage, and secure them to the premade screw holes in the X stage.
         NOTE: Both the X stage and Y stage should remain moveable until the screws are fully tightened.
2. Ensure the micromanipulator drive is outfitted with a plastic platform to attach the electrode connector, as well as an extended polyimide tube to hold the electrode. Attach the 64-pin connector to the platform on the micromanipulator using tape.
3. Carefully place a dab of bacitracin (sterile ointment) onto the plastic tube on the micromanipulator, and attach the electrode connected to the 64-pin connector via a flexible ribbon cable.
4. Use the micromanipulator drive to hold the silicon electrode and its connector while threading the plastic tube through a hole in the Y-stage. The electrode will be hanging beneath the stage of the probe, waiting to be attached to the plastic rectangular platform (step 7, section 1).
5. Detach the 64-pin connector from the micromanipulator drive, and use glue gel to secure the connector to the platform on the X-stage. Additional epoxy may be added to strengthen the bond of the connector once the glue is holding it in place. Leave overnight to dry.
6. Carefully detach the electrode from the plastic tube, and remove the micromanipulator drive from the assembly.
   NOTE: At this point, the drive is only held with the alligator clip of a Helping Hands, and the electrode is not secure within the drive construction.
7. Carefully move the alligator clip to flip the drive over to see the unsecured electrode. Using blunt tip forceps, place the electrode onto the plastic holder below the Y stage, and secure it with a small amount of silicone elastomer (silastic). Wait for 5 min until the adhesive has cured.
8. Use the screw control on the micro-drive to retract the electrode.
9. Place the protective sleeve over the electrode on the micro-drive. This protective sleeve is the same size as the custom-printed recording chambers and fits securely into the base of the drive assembly.
10. Using a flathead screwdriver, tighten the three side-located screws on the base component of the micro-drive to secure the protective sleeve in place. Connect the head-stage used for the recordings onto the 64-pin connector.

2. Polytrode plating of electrodes to reduce the overall impedance (Figure 2)

NOTE: For the best recording quality, it is useful to electrode-plate the silicon electrode arrays with a poly(3,4-ethylenedioxythiophene) solution (PEDOT). This method has been shown to increase the signal-to-noise ratio^7,8.

1. To make the PEDOT solution, combine 0.075 mL of 3,4-ethylenedioxythiophene (EDOT) and 1.4 g of poly(sodium 4-styrenesulfonate) (PSS) in 70 mL of distilled water. Vortex this solution briefly to ensure the complete dissolution of the components. Mix this solution the day before use for the best dissolution.
2. After launching the impedance tester software (see Table of Materials), prepare the settings with the chosen electrode preparation. Using the upper drop-down menu at the top left of the software window, select the adapter being used (N2T A32). Using the second drop-down menu, select the electrode being used (NLX-EIB-36). If the electrode is not listed in the pull-down menu, see the manual for how to define a new electrode. Ensure that the number of channels is correct.
3. Connect the probe to the impedance tester system via the connector, and lower the probe into grounded saline to obtain a baseline reading.
   1. In the impedance tester system, press the button Test Impedances on the left side, and ensure the correct settings (Settings: Test Frequency: 1,004 Hz, Cycles: 100, Pause: 0). Then, press the Test probe button on the lower right.
   2. A probe report window will appear as the impedance tester runs, storing results in a spreadsheet. Save these results for future reference for both shanks of the electrode (the mounting of the electrode into a beaker for use with the impedance tester is depicted in Figure 2D). If using a probe with multiple shanks, ensure step 3 is repeated for each shank.
4. Remove the probe from the saline by lowering the pedestal (i.e., lab jack) holding the beaker. Replace the beaker fluid with distilled water. Raise the lab jack to submerge the electrode, rinse off the remaining saline solution, and then lower the lab jack again.
5. Replace the distilled water in the beaker with PEDOT solution, and raise the lab jack until the electrode is submerged in the solution.
6. Select the DC Electroplate button on the left side of the software window, and ensure the correct settings are used (Settings: Autoplate: 0.004 µA, 350 Hz, Duration: 5, Pause: 1). Then, press the Autoplate button on the lower right. Once again, a probe report window will appear with the results of the electroplating. Save these results for future reference. If using a probe with multiple shanks, ensure this step is repeated for each shank.
7. Remove the probe from the PEDOT solution by lowering the lab jack, and rinse the probe with distilled water as performed previously (step 4). After rinsing, fill the beaker with saline solution, and raise the lab jack until the electrode is submerged.
8. Press the Test Impedances button on the left side of the software window, and ensure the correct settings are used (Settings: Test Frequency: 1,004 Hz, Cycles: 100, Pause: 0). Press the Test Probe button on the lower right. Save these results for future reference. If using a probe with multiple shanks, ensure this step is repeated for each shank.
   NOTE: Compare the post-electroplated values with the pre-electroplated values. There should be a decrease in the impedance values.
9. With continual use, the probes may show increases in the impedance values after 6 months. If there is a concern about the impedance values from the recording signal, retest the probe impedances following step 3. If the impedance values have increased from the initial values, repeat the electroplating (step 6).

3. Surgical placement of the head cap, chambers, and craniotomy (Figure 3A-C)

NOTE: In this work, at the termination of the study, the animal was anesthetized under isoflurane and received intramuscular (IM) injections of ketamine, followed by an intraperitoneal (IP) injection of euthasol. The brain was extracted following transcardial perfusion with saline followed by 10% formalin.

1. Train the marmosets (at least 1.5 years of age) to sit in a small primate chair following previously described methods^3,9,10,11 2 months prior to surgery.
2. On the day of surgery, induce the subjects with an intramuscular (i.m.) injection of ketamine (5-15 mg/kg)/dexdormitor (0.02-0.1 mg/kg) (and midazolam at 0.25 mg/kg as a muscle relaxant), and confirm proper anesthetization based on the heart rate, respiration, and toe pinch reflex. Intubate the animals for continuous isoflurane (0.5%-2% in 100% oxygen) administration throughout surgery while monitoring their vitals. Use vet ointment on the eyes during surgery to prevent dryness while under anesthesia.
3. After the surgeons have put on sterile PPE, prepare a sterile surgical field using sterile drapes to cover the surfaces and the operative field. Using aseptic techniques, have the surgeons implant an acrylic head cap with titanium posts and screws to stabilize the head using methods described in detail previously by Nummela et al.¹¹.
4. During the implant surgery, plan out the recording chamber placements over the areas of interest (e.g., visual areas middle temporal area [MT], and primary visual cortex [V1]) based on stereotaxic coordinates¹².
5. Place a grounding wire.
   1. Start by drilling a 1.1 mm burr hole into the skull near the area of the planned recording chamber. Bend a 36 G stainless steel wire about 0.075-1 mm from the tip, and slide the bent portion of the wire between the skull and the dura in the bottom of the burr hole.
   2. Initially seal the wire and hole with bone wax and then with cement and acrylic when placing the chambers. Ensure to leave a portion outside near the chamber to attach for grounding.
6. After placing a base layer of quick adhesive cement to cover the entire area of the recording site and grounding wire, adhere the recording chambers (consisting of custom 3D-printed parts) to the skull using the quick adhesive cement, and reinforce with acrylic when building the head implant.
7. Seal the chambers with a 3D-printed chamber cap that is designed to fit tightly inside the chamber (and cannot be removed by cage-mates pair-housed with the subject).
   1. Dispense a small amount of silicone elastomer (silastic) around the inner edges of the chamber using the included mixing tips, and then insert the chamber cap until flush with the outside chamber walls.
   2. The chambers are designed with access holes (1 mm diameter) on either side of the chamber to ensure the chamber cap can be removed easily. Use silastic to seal these access holes, resulting in an airtight chamber seal.
      NOTE: To remove the chamber caps, use tweezers to remove the silastic sealing the access holes first to break the airtight seal of the chamber. Then, a pin can be used to leverage the chamber cap out of the chamber.
8. Recover the animals from anesthesia, and do not leave them unattended until they have regained sufficient consciousness to maintain sternal recumbency. Treat the animals post-surgically with antibiotics (amoxicillin-clavulanic acid at 13.75 mg/kg) for 7 days and anti-inflammatories (meloxicam at 0.1 mg/kg) for 3 days. Give slow-release pain medication prior to recovery to last 72 h post-operatively, and house the animals independently until full recovery.
9. Two to three weeks after surgery, begin to train head-restraint and visual behavior. Place the marmoset in a primate chair, and allow for acclimation to the head restraint. Once comfortable, train the marmoset to perform several basic tasks, including central visual fixation¹³, and a saccade task toward a peripherally detected Gabor grating, which is used to measure their visual acuity¹¹.
10. After sufficient behavioral training, perform a craniotomy within the recording chamber. Under sterile techniques, perform a second surgery to create a craniotomy (2-3 mm in diameter) in the recording chamber over the area of interest (e.g., area MT). Seal the craniotomy with a thick application of silastic of about 1 mm or the depth of the craniotomy to protect the brain from infection and reduce granulation growth¹⁴. Ensure that the silastic seal has edges grabbing on to at least 2 mm or more of the dental cement on all sides. Replace the chamber cap as described in step 7.
11. If any bleeding occurs or the silastic seal leaks clear fluids in the days following surgery, then the chamber will need to be cleaned.
    1. If the silastic seal is intact, use sterile techniques to clean with 10% betadine, and wash with sterile saline before removing the seal. If the silastic seal is not intact, only clean it with sterile saline under sterile techniques.
    2. Once the silastic is removed, flush the dura with sterile saline several times, and dry with a cotton tip applicator. Further dry the chamber thoroughly using a cotton tip applicator before applying a new layer of silastic. Typically, the chamber stabilizes and remains dry with a tight seal after a few days to 1 week.
12. Once the craniotomy has stabilized and shows no signs of bleeding, perform a dural scrape to remove excess tissue over the dura. Then, apply a thin layer (<1 mm) of silastic (thin enough to enable passage of the electrodes for recordings), and use a 1/4 burr drill bit to wick the silastic up over the walls of the craniotomy, as shown in Figure 3C, for mechanical purchase.
    NOTE: This will also allow for easier removal for cleaning purposes. The silastic can remain in place for the duration of the study and can be punctured with linear array electrodes for recording.

4. Neurophysiology recording setup (Figure 3D-F)

NOTE: The animal handling steps will vary depending on the lab and experiment. The following steps are specific to the placement of the micro-drive and the penetration of the dura for the recordings.

1. Remove the cap from the recording chamber (described in section 3, step 7.2). Place the cap in 91% isopropyl alcohol to soak during the recording. Make sure the edges of the chamber are clear from silastic. If silastic is left on the edges of the chamber, the micro-drive will not sit correctly.
2. The base of the micro-drive has three screws for tightening it to the chamber on the animal. Normally, those screws are tightened in place around a protective sleeve that is the same size as the chamber to keep the silicon probe inside protected from being hit.
   1. First, use the screwdriver to loosen the three screws holding the protective sleeve over the silicon probe, and carefully remove the sleeve. Attach the micro-drive to the recording chamber, being careful not to hit the silicon electrodes on the walls of the chamber (Figure 3D, E).
   2. Additionally, ensure the electrodes are in a fully retracted position before placing the drive onto the chamber, such that they sit well above the cortex on initial placement.
3. Once placed, use the screwdriver to tighten down each screw on the side of the micro-drive (Figure 3F). Pick an order to tighten the screws, and use this every time to maintain stability in the location. Researchers may also want to count the number of turns when initially loosening the screws to tighten them by a similar amount. Check if the probe is loose, and tighten it as needed, but be careful not to strip the screw threads in the plastic by over-tightening.
4. Once the micro-drive is secured to the chamber, place all the grounding wires needed.
   NOTE: Leave the head stage inserted into the connector overnight, and only insert the serial peripheral interface (SPI) cable into the head stage for recordings because it is difficult to insert the head stage into the connector while it is on the animal.
5. Carefully plug the SPI cable into the head stage (already connected to the probe). Ensure not to bend the wire. Amplify and digitize all the neurophysiology data from the head stage at 30 kHz using the Open-Ephys GUI (https://github.com/open-ephs/plugin-GUI). High-pass-filter the wideband signal from the head stage at 0.1 Hz. Correct for the phase shifts from this filtering, as described previously¹⁵. For linear arrays, preprocess the resulting traces by common-average referencing.
6. Enter the channel map for the particular electrode array being used in the recording system so the channels plotted during the recording are ordered by depth in the high-pass-filtered recording signal view. This view will be utilized when advancing and retracting the electrode.

5. Performing the neurophysiology recording experiment (Figure 4)

NOTE: Here, the method for lowing the electrode arrays into the cortex is described; this method has been optimized to avoid excessive dimpling of the underlying tissue. An increase in noise in the electrophysiological recordings provides a good indication of dimpling prior to penetrating the silastic and entering the brain. Once in the brain, if there is too much dimpling, the researcher may notice units shifting on the probe even without the manipulation of the drive (units gradually moving across channels in depth), or alternatively, the researcher may notice suppression of the neural activity, particularly at superficial sites on the probe. In those conditions, the drive is retracted to relieve dimpling and facilitate better recordings.

1. Using the screw control on the micro-drive, lower the silicon probe, making one turn (250 µm) every 1-2 s until units are observed at the array tip.
   NOTE: A minor increase in noise during the penetration of the silastic placed over the dura may be observed. The shanks will dimple the silastic before popping through, which can increase the noise on the probe. A fast descent until the first sign of units aids in the penetration of the silastic layer.
2. Once the first neurons are observed, retract one turn of the micro-drive (250 µm) to reduce the speed of entry as the electrode begins to pop through the silastic and dura into the cortical tissue. If the electrode appears to continue advancing into the tissue over time without micromanipulation, retract an additional turn to relieve pressure from the probe pushing on the silastic and tissue beneath it.
   NOTE: It should be possible to track the location of neurons if the correct channel map for the linear array has been loaded (the channels will appear ordered by their depth relative to the cortex).
3. Continue driving the array into the cortex very slowly, advancing by approximately four to six turns (1-1.5 mm) over a 20-30 min duration until neurons are evenly distributed across the length of the probe.
4. Slowly retract the array by one to two turns (0.25-0.5 mm) to reduce the pressure on the tissue before beginning the recordings.
   NOTE: Neurons do not typically shift channel location on the linear array during this retraction, suggesting that it primarily acts to reduce pressure on the tissue. Without retraction, a continual array shift may be seen during the recording as the tissue relaxes, thus disrupting the recording stability (Figure 4C).
5. After retracting, wait 20 min before starting the recordings. This will limit the amount of movement seen in the recording. Use this time to prepare for the recording (i.e., calibrate the eye tracking and prepare the stimulus presentation systems).
6. Select Record on the recording software.
7. When the experiment is completed, select the recording button again. This will create the full recording file in the selected destination. Ensure that the file has been saved correctly before continuing.
8. Slowly begin retracting the array out of the cortex at a similar speed as before (four to six turns over a 20-30 min duration). Visualize the probe retracting by watching the neurons shift along the probe. After retracting by the number of turns since first seeing neurons and when no more neurons are observed on the probe, continue retracting more quickly until returning to the fully retracted starting position.
9. To remove the probe from the recording chamber, perform steps 1-3 in section 4 in reverse order (i.e., first perform step 3, followed by step 2, and then step 1).
10. Once the probe has been removed from the recording chamber, soak it in contact lens solution for 20 min to remove any tissue or blood from the electrode. Afterward, place the probe in alcohol for 1 min to remove the contact lens solution, and let the electrode dry.
11. Use Kilosort2 to spike-sort the array data after the intial filtering from the recording system (as described earlier in section 4, step 5).
    1. Manually label the outputs from the spike-sorting algorithms using the "phy" GUI (https://github/kwikteam/phy). Classify the units with tiny or physiologically implausible waveforms as noise, and exclude them.
    2. If using laminar probes, view the spike waveforms across each channel to ensure single units are labeled on the channel with the peak waveform (Figure 5A). Only include units that have clear clusters in the PCA space, less than 1% inter-spike interval violations, and bi-phasic spike waveforms (which should be localized to adjacent channels on the linear array). An example single unit is shown in Figure 5B, which displays clear clusters in the PCA space (top right) and stability over time (bottom right).

Representative Results

This protocol describes how to build an X-Y electrode stage (Figure 1) that allows for the sub-millimeter targeting of sites and maintains reliable positioning across separate recording sessions. The reliability of the X-Y positioning is illustrated in Figure 6, which demonstrates that two recording sessions conducted a week apart showed a 70.8% overlap in their mean RF locations (Figure 6A). Furthermore, minor adjustments to the micro-drive positioning can support precise movement in the retinotopic space. Small X and Y stage movements up to 0.2 mm can be made by referencing the predrilled holes in each stage, which are 2 mm apart. In a series of recording sessions that transversed the MT/MTC border using minor positioning adjustments (Figure 6B), precise movements in retinotopic space could be seen (Figure 6C). The recording sites for the MT and MTC aligned with electrophysiology maps previously depicted in Solomon and Rosa². Similarly, with this protocol, the Z-positioning can be tracked by the number of turns to the preferred electrode depth. While the Z-positioning of the drive to place electrodes in cortical depth must be adjusted daily and includes some variation, it is possible to obtain estimates of depth in early visual areas offline by using current source density analysis (CSD), as described previously¹⁶.

Using this X-Y stage technology, researchers can perform neurophysiological recording with linear arrays while the subjects perform a variety of foraging tasks for visual receptive field mapping and simple saccade tasks¹⁷. Electrode placement techniques are shown in Figure 4, which highlights the importance of retraction to alleviate pressure on the tissue and prevent long-term cortical damage. Using two shank linear arrays allows for sampling a large number of neurons across cortical depths during each recording. In this work, the spike-sorted array data obtained using Kilosort showed that the principle component features of a selection of spikes in a cluster were distinct from background spikes and that the recordings were stable over time (Figure 5).

Figure 1: Construction of the electrode micro-drive. The construction of an electrode micro-drive is possible after obtaining all the necessary components outlaid in section 1, step 1. Each successive step (section 1, steps 2-10) shows a before and an after image to aid in the visualization of the actions required. This construction is explained fully in protocol section 1. Please click here to view a larger version of this figure.

Figure 2: Constructed micro-drive and electroplating setup. Overview image of the completed micro-drive from the (A) top and (B) bottom. (C) The micro-drive is attached to a protective sleeve (same dimensions as the chambers on the animal's implant, with an extended length for protection) and then attached to the animal's chamber by tightening three side screws. (D) Configuration of the micro-drive on the impedance tester for electroplating. Please click here to view a larger version of this figure.

Figure 3: Schematic of the head cap and chamber placement (A) Drawing representation of an animal with a head cap, head post, and MT chamber. (B) Image of an MT chamber 6 months after craniotomy and sealed with a thin layer of silastic. (C) Schematic of the chamber and micro-drive preparation. The illustration shows the chamber preparation and the thin layer of silastic for use in the recordings. The micro-drive is secured to the recording chamber by tension from the side screws. (D) One should move parallel to the chamber to attach the micro-drive to the recording chamber. Coming from an angle will likely result in hitting the side of the chamber or the cement sides of the craniotomy. (E) The micro-drive secured on an example head cap. (F) Depiction of the three screws on the micro-drive to be tightened for secure placement. Note: One screw sits behind the connector and is usually tightened from above. The screws are tightened in a repeatable order to ensure drive location reliability. Please click here to view a larger version of this figure.

Figure 4: Neurophysiology recordings (A) Depiction of the electrode before and after it touches the dura (top). High-pass-filtered recording signal from a single probe in a neural recording before and after the probe touches the dura (bottom). (B) Depiction of an electrode causing dimpling after initially popping through the silastic (top). High-pass-filtered recording signal from one probe in a neural recording after the probe has popped through the silastic (bottom). Channels with neurons are labeled as filled. (C) Depiction of the electrode in the cortex before and after retraction (top). High-pass-filtered recording signal from one probe in a neural recording before and after a 500 µm retraction; the units have shifted one channel (bottom). Please click here to view a larger version of this figure.

Figure 5: Post-processing of neurophysiology data. (A) Depiction of a laminar probe (left) and a selection of spike waveforms viewed across each channel (right). (B) Example Kilosort output. (Left) Example waveform across several channels with a peak waveform on channel 46. (Top right) The principle component features of a selection of spikes in the selected cluster (blue) against a set of background spikes (uniformly spaced in time across the entire recording, grey). Spiking clusters are easily discriminated from background spikes. (Bottom right) The amplitude of a selection of spikes belonging to the selected cluster over time. The recordings show stability over time for this example neuron. Please click here to view a larger version of this figure.

Figure 6: Reliability of the X-Y stage positioning (A) Mean receptive field (RF) contours for two recording sessions 1 week apart with the same micro-drive coordinates, showing 70.8% overlap. (B) An MT/MTC map showing the successive targeting locations across recording sessions. The solid lines and numbers indicate eccentricity from the fovea in degrees. The dashed lines indicate the horizontal meridian (HM). The bolded lines indicate the vertical meridian (VM). (C) The corresponding RF locations for each shank outlined with a circle marking the center location show that minor adjustments to the micro-drive positioning can support precise movement in the retinotopic space. Panel B has been adapted with permission from Solomon and Rosa². Please click here to view a larger version of this figure.

Discussion

Several methods (e.g., chronic, semi-chronic, acute) are currently available for performing neurophysiology experiments in non-human primates. The common marmoset poses unique challenges for neurophysiology experiments due to its small size and lack of gyri as anatomical landmarks. This requires researchers to use neurophysiological landmarks such as the retinotopy and tuning properties of areas of interest to identify the recording targets. Therefore, when initially mapping out a cortical area, daily adjustments to the electrode positioning may be needed. Current preparations for marmoset neurophysiology often use semi-chronic probe positioning, which does not allow for easy-access positioning mechanisms⁵. This protocol demonstrates a lightweight micro-drive for stable linear array recordings that enables flexible positioning with precise sub-millimeter movements across sessions to map out the retinotopy within a chamber.

The current approach, which involves acute daily electrode penetrations for precise electrode targeting and movement across retinotopic maps, differs from chronic electrode array implants in marmosets⁵. However, acute preparations intrinsically have drawbacks, such as an increased experimental preparation time and an increased risk of infection, as well as an increased risk of cortical tissue damage due to repeated penetrations. While the use of silastic has been optimized in this preparation to limit the risk of infection, applying the silastic does require some troubleshooting to ensure it is done correctly and to obtain a dry seal. This preparation also demands care to avoid electrode damage during the drive mounting and requires care in tightening the drive screws to ensure reliable targeting across sessions.

Long-term cortical damage can be avoided in this preparation when applied correctly with a secure drive, slow penetration of the tissue, and the avoidance of major blood vessels. Additionally, the use of silastic in the craniotomy to prevent infections has been optimized in this technique. The brain is covered with a thin layer of silastic after the craniotomy is performed to prevent infection and keep the interior of the chamber dry¹⁴. When the dura is clear from blood and granulation tissue (which is possible with a tight silastic seal), this enables the visualization of the major blood vessels in the underlying tissue and for targeting the silicon probes to avoid hitting the blood vessels during recordings. This technique requires some troubleshooting during learning, since it is critical that the silastic seal is thin (<1 mm) in the center to allow electrode penetration without excessive dimpling while also being thick at the edges to provide mechanical purchase. These methods will be valuable in investigating the circuitry role of the layers across the marmoset cortex and in preventing tissue damage during multiple penetrations.

Using custom-printed plastic components along with commercially available supplies, this micro-drive design is easy to use once fabricated and can be personalized for different recording probes. While this method has been shown to be reliable, the initial fabrication can be difficult and costly to learn. However, the components of the probe are relatively economical to purchase, and, once constructed, the probes have been shown to last up to 1 year with continual use.

One of the key advantages of this preparation is the precise and repeatable targeting of areas of interest due to the X-Y stage. With this technology, researchers are able to map the retinotopic locations within the recording chambers and record from different areas across days using a coordinate system. To achieve this, it is critical to ensure the correct placement of the probe daily and to tighten all the exterior screws in a repeatable manner. Changes to the exterior screw tightness can lead to small shifts in the electrode placement across days.

Acute recordings offer the benefit of repeated independent measurements and can result in a higher yield of single-unit data. However, semi-chronic or chronic recordings may also offer advantages. In small animals such as mice, chronic recordings can be made with high-density silicon probes to obtain stable, long-term population recordings over multiple days¹⁸. Eliminating the repeated insertions of electrodes into the brain for each recording session can shorten the overall experimental preparation time and reduce the risk of infection or cortical tissue damage. Therefore, when evaluating the use of chronic or acute recording preparations, it is essential to consider the specific experimental requirements.

The proposed micro-drive design allows the flexibility of different recording techniques and array usage, meaning it could be easily adopted into existing experimental setups and pipelines. Furthermore, the flexibility of the array mounting on the micro-drive may lead to possible future applications, such as controlling laser stimulation for optogenetic purposes. This method may enable future studies to improve on targeting for laser stimulation in the marmoset.

Materials

Name	Company	Catalog Number	Comments
1/4 Hp burr drill bit	McMaster & Carr	Cat# 43035A32	Carbide Bur with 1/4" Shank Diameter, Rounded Cylinder Head, trade Number SC-1, single Cut(https://www.mcmaster.com/products/bur-bits/burs-7/?s=1%2F4%22+bur+bits)
1x1mm Crist Grid	Crist Instruments	1 mm x 1 mm Grid	https://www.cristinstrument.com/products/implant-intro/grids
91% isopropyl alcohol	Medline	N/A	https://www.medline.com/product/Medline-Isopropyl-Rubbing-Alcohol/Bulk-Alcohol/Z05-PF03807?question=91%25%20isopropyl%20alcohol
Acquisition Board	Open-Ephys	N/A	https://open-ephys.org/acquisition-system/eux9baf6a5s8tid06hk1mw5aafjdz1
Bacitracin Ointment	Medline: Cosette Pharmaceuticals Inc	N/A	https://www.medline.com/product/Bacitracin-Ointment/Antibiotics/Z05-PF86957?question=bacitr
Blunt straight Forceps	Medline	N/A	https://www.medline.com/category/Central-Sterile/Surgical-Instruments/Forceps/Z05-CA16_02_20/products
Bone wax	Medline	ETHW31G	https://www.medline.com/product/Ethicon-Bone-Wax/Bone-Wax/Z05-PF61528?question=bonewax
C&B Metabond Quick Adhesive Cement System	Parkell, Inc.	SKU: S380	https://www.parkell.com/C-B-Metabond-Quick-Adhesive-Cement-System
Clavamox	MWI Animal Health	N/A
Contact lens solution	Bausch and lomb		Various sources available
Custom Printed 3D printed parts	ProtoLab		https://marmolab.bcs.rochester.edu/resources.html
DB25-G2 25 Pin Male Plug Port Signal Connector	Various Sources	DB25-G2 25	DB25-G2 25 Pin Male Plug Port Signal 2 Row Terminal Breakout Board Screw Nut Connector
diamond saw attachment for dremel	Dremel	545 Diamond Wheel	https://www.dremel.com/us/en/p/545-26150545ab
Digitizing Head-stages	Intan	RHD 32channel (Part #C3314)	https://intantech.com/RHD_headstages.html?tabSelect=RHD32ch&yPos=120.80 000305175781
EDOT	Sigma Aldrich	Product # 483028	https://www.sigmaaldrich.com/US/en/product/aldrich/483028
Helping Hands	Harbor Freight	N/A	https://www.harborfreight.com/helping-hands-60501.html
Hook Electrical Clips	Various Sources	N/A	Hook test Cable wires
Interface Cables (RHD 3-ft (0.9 m) ultra thin SPI cable)	Intan	Part #C3213	https://intantech.com/RHD_SPI_cables.html
Lab jack	Various Sources	N/A	https://www.amazon.com/Stainless-Steel-Scissor-Stand-Platform/dp/B07T8FM85H/ref=asc_df_B07T8FM85H/?tag=&linkCode=df0&hvadid=366343 827267&hvpos=&hvnetw=g&hvrand =2036619536500717246&hvpone =&hvptwo=&hvqmt=&hvdev=c&hv dvcmdl=&hvlocint=&hvlocphy=900 5674&hvtargid=pla-795933567991& ref=&adgrpid=71496544770&th=1
Meloxicam	MWI Animal Health	N/A
Micro-drive	Crist Instrument	3-NRMD	https://www.cristinstrument.com/products/microdrives/miniature-microdrive-3-nrmd
Multi-channel linear silicon arrays with 64 channel connector	NeuroNexus	A1x32-5mm-25-177	https://www.neuronexus.com/products/electrode-arrays/up-to-10-mm-depth/
NanoZ Omentics Adapter- 32 Channel	NeuraLynx	ADPT-NZ-N2T-32	https://neuralynx.com/hardware/adpt-nz-n2t-32
NanoZ System	Plexon	NanoZ Impedance Tester	https://plexon.com/products/nanoz-impedance-tester/
Narishige Micromanipulator	Narishige	Stereotaxic Micromanipulator	https://usa.narishige-group.com/
Open-Ephys GUI	Open-Ephys		https://open-ephys.org/
Polyimide Tubing (OD(in): 0.021 / ID(in) 0.018 )	Various Sources (Chamfr)	Chamfr Cat#HPC01895	https://chamfr.com/sellers/teleflex-medical-oem-llc/
Primate Chair	Custom made by University of Rochester Machine Shop	Designs online	https://marmolab.bcs.rochester.edu/resources.html
Poly(sodium 4-styrenesulfonate) (PSS)	Sigma Aldrich	Product # 243051	https://www.sigmaaldrich.com/US/en/product/aldrich/243051
RHD USB Interface board	Intan	RHD2000 Evaluation Board Version 1.0	https://intantech.com/RHD_USB_interface_board.html
Silastic gel	World Precision Instruments	# KWIK-SIL	Low Toxicity Silicone Adhesive ((https://www.wpiinc.com/kwik-sil-low-toxicity-silicone-adhesive)
Slow release buprenorphine	Compounding Pharmacy
Stainless steel wire 36 gauge	McMaster & Carr	Cat# 6517K11	Round Bend-and-Stay Multipurpose 304 Stainless Steel Wire, Matte Finish, 1-Foot Long, 0.008" Diameter
Stanley 6-Piece Precision Screwdriver Set	Stanley	1.4mm flathead screwdriver	https://www.amazon.com/Stanley-Tools-6-Piece-Precision-Screwdriver/dp/B076621ZGC/ref=sr_1_3?crid=237VSK5FNFP9N&keywords= stanley+66-052&qid=1672764369&sprefix= stanley+66-052%2Caps%2C90&sr=8-3
Steel Screws	McMaster & Carr	type 00 stainless steel hex screws and 1/8” in length	https://www.mcmaster.com/
Steel Tube	McMaster & Carr	28 gauge stainless steel tubing	https://www.mcmaster.com/tubing/multipurpose-304-stainless-steel-6/id~0-055/
Superglue	Loctite	SuperGlue Gel Control	https://www.loctiteproducts.com/en/products/fix/super-glue/loctite_super_gluegelcontrol.html