Electrophysiology of Laminar Cortical Activity in the Common Marmoset

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).