Protocol for behavioral and neural recording during stimulation of the macaque monkey nucleus basalis

Summary We present an experimental protocol to record neuronal activity during intermittent stimulation of nucleus basalis (NB), as macaque monkeys perform cognitive tasks. This protocol includes implantation of electrodes and generator devices to deliver electrical stimulation to NB using multiple approaches in monkeys. Direct stimulation of NB avoids peripheral cholinergic side effects, optimizes timing, and activates non-cholinergic projection neurons. We describe electrode preparation, surgery, and implantation for direct evaluation of how stimulation affects monkeys’ behavior and neuronal activity. For complete details on the use and execution of this profile, please refer to Qi et al. (2021).


SUMMARY
We present an experimental protocol to record neuronal activity during intermittent stimulation of nucleus basalis (NB), as macaque monkeys perform cognitive tasks. This protocol includes implantation of electrodes and generator devices to deliver electrical stimulation to NB using multiple approaches in monkeys. Direct stimulation of NB avoids peripheral cholinergic side effects, optimizes timing, and activates non-cholinergic projection neurons. We describe electrode preparation, surgery, and implantation for direct evaluation of how stimulation affects monkeys' behavior and neuronal activity. For complete details on the use and execution of this profile, please refer to Qi et al. (2021).

BEFORE YOU BEGIN
The methods that follow describe methods of implanting Deep Brain Stimulation (DBS) electrodes targeting the Nucleus Basalis (NB) of Meynert in macaque monkeys and recording behavior and/or neural activity during NB intermittent stimulation. Procedures described below followed guidelines by the U.S. Public Health Service Policy on Humane Care and Use of Laboratory Animals and the National Research Council's Guide for the Care and Use of Laboratory Animals and were reviewed and approved by the Wake Forest University Institutional Animal Care and Use Committee (IACUC), the Vanderbilt University IACUC and the Augusta University IACUC. Animals were maintained in the AAALAC accredited facilities managed by the Animal Resources Program of the Wake Forest University School of Medicine; in the Vanderbilt University facilities maintained by the Division of Animal Care; and the Augusta University animal facilities. Animals were group-or pair-housed in cages whenever compatible peers were available, in facilities housing several individuals of the same species. Methods for behavioral and neural acquisition (which require specialized apparatus, including a microelectrode drive, recording system, and dedicated software) are not described here explicitly; we refer to published studies for more detail. The protocol that follows presents the methods that are necessary to deliver DBS targeting the NB in order to make behavioral and neural data acquisition possible. 75

STEP-BY-STEP METHOD DETAILS
Creating the stimulation electrodes (see Figure 1 for this stage) Timing: 1.5 h (for a batch of electrodes) 1. Collect materials. A typical electrode uses 31 mm of polyimide tubing, 30 mm of hypodermic tubing, and 55 mm of 75 mm platinum/iridium Teflon insulated wire ( Figure 1A). 2. Use iris scissors to carefully remove 1 mm of insulation from the stimulation end to achieve impedances under 1000 Ohms ( Figure 1B). For higher impedances, less insulation may be removed. 3. Use a Dremel cut-off wheel to cut and file down the desired amount of hypodermic tubing.
Probe the hypodermic tubing with 26G Precisionglide syringe needle to make sure the holes of both ends of the tube have not been blocked by the cutting process. Use file to smooth down both ends of the tubing. Ream the entire section of tubing using tungsten wire.
4. Thread the materials. Locate the stimulation end of the wire (straight) and thread the other end through the metal hypodermic and then the polyimide sheath. The metal hypodermic is threaded through the polyimide sheath ( Figure 1C). 5. Pull the wire so less than 1.5 mm protrudes from the polyimide on the stimulating end, with up to 25 mm on the tail end. 6. Glue the wire into the tubing on both ends with cyanoacrylate. The Krazy Glue brand seems to wick into the tubing most effectively. Not all cyanoacrylates are equal in this attribute (Figure 1D). 7. Free insulation from the tail end by mechanical stripping. In some applications, a butane torch may be used, but if the wire is to be welded, mechanical stripping is preferred because it avoids leaving a residue that may interfere with welding. 8. Measure the impedance after removing the insulation. The ideal value is 1000 Ohms at 1 kHz but range of 100-3000 U will be acceptable. Typically the reference path electrode is placed in a saline solution, and a 1 kHz signal is pushed from electrode to reference using a current of 1 microamp, and the peak-to-peak voltage measurement is the impedance in megaOhms. 9. Store electrodes in a conventional microelectrode storage box and sterilize them using a low heat method, plasma or gas (McEvoy and Rowan, 2019). The electrode should be steam durable, but the electrode boxes are typically not. 10. The length of insulation removed will dictate the impedance. Preventing irreversible reactions at the metal/brain interface will require charge densities stay within acceptable limits, and delivery of 0.5 mA through a 1 mm length of platinum or iridium with a diameter of 75 mm falls well within these limits (Shannon, 1992). If the impedance measurement is too high, above 10 kOhms, or too low, under 100 Ohms, the tip must be inspected and/or replaced.

OPEN ACCESS
Note: These methods have been previously reported briefly (McCairn and Turner, 2009;Turner and DeLong, 2000).

Stimulation electrode and implantable pulse generator (IPG) implantation
Timing: 3 h This step describes implantation of stimulation electrodes to deliver stimulation into NB using an IPG (see Figures 2 and 3 for this stage).
11. Before starting surgery, determine target coordinates for implantation of electrode into the NB. a. A preoperative MRI is optimal for determining placement. Alternatively, we have used average stereotaxic coordinates when obtaining an MRI was not practical for an animal. b. Using a vertically oriented electrode approach, NB is accessible at the following stereotaxic coordinates: 16 mm anterior to the interaural line; 9 mm lateral to the midline; 29 mm below the surface of the cortex.
Note: This position places the electrode 2 mm posterior to the rear margin of the anterior commissure where it crosses the globus pallidus in coronal view.
Note: This landmark is easily viewed on coronal MRI or histological sections and implantation coordinates may be adjusted to target it more accurately in individual cases.
Note: The vertical oriented approach in the macaque monkey minimizes disruption of cortical gray matter and is consistent with most stereotaxic frames and positioners.
12. Prepare the monkey for surgery a. Solid food is withheld for 12 h before surgery. b. Sedate monkey with intramuscular (i.m.) ketamine (5 mg/kg) and dexmedetomidine (0.2 mg/kg) Optional: the monkey may receive i.m. injections of atropine (0.04-0.06 mg/kg, im) to control secretions.
c. Shave surgery area comprising the scalp and subscapular area with hair clipper and scrub with Nolvasan. d. Intubate and secure endotracheal tube in location.
CRITICAL: make sure the tube is in place by tightening a strip of gauze around the subject's head.
e. Put the monkey prone on the surgery table.
CRITICAL: Use a heating pad to maintain the body temperature during surgery, aiming for 36 -38 C.
f. Anesthetize with isoflurane, 3% for induction, 1.5% for maintenance delivered in 2 L/min oxygen. Monitor anesthesia by means of SpO2, Heart Rate, Respiration Rate, End-title CO 2 , temperature. g. Administer ophthalmic ointment in both eyes of the animal to protect the corneas from drying and abrasion. h. Secure monkey on stereotax. 13. Disinfect shaved area: clean the surgical site with antiseptic scrub. 14. Administer lidocaine subcutaneously at the surgical site (2 cc of 2% solution). 15. Cut a flap of scalp through the galea to expose the bone. 16. Drill a hole according to coordinates obtained from MRI.
a. Implant a bone screw with a radially centered hole in it.

CRITICAL:
The bone screw and hole should both be performed in stereotaxic vertical orientation and not orthogonal to the bone angulation and should be vertically above the desired stimulation location.
Note: Bone screws were custom designed with Gray-Matter Research, and are 4.5 mm long, with a 1.85 mm diameter through hole. The threads are bone screw pitch, 1.25 mm spacing.

OPEN ACCESS
Note: The bone screw seats well in the cranial bone and creates a minimal defect in the sealing of the cranium that can be easily sealed. It also enables later removal of the electrode should it become necessary for repositioning or MRI imaging (Figure 2A).
17. Lower a 19 ga. sharp hypodermic guide tube using the manual rotational drive on the stereotaxic micropositioner. The tip should be carefully advanced at least 5 mm below the dura mater. The guide tube is advanced in stereotaxic vertical orientation, and acts as the primary targeting vector for the electrode tip. Check its orientation carefully. Measure the length of the electrode to be inserted, and the length of its wire tail ( Figure 2B).
CRITICAL: The guide tube for this procedure is 23 mm long, which is important for its later removal. The guide tube needs to be long enough to pass the dura, and short enough that it can be later removed while visualizing the tail end of the electrode.
18. Insert the stimulation electrode manually into the guide tube, and push the electrode until the polyimide sheath is fully within the guide tube. Insert the stylus into the guide tube and advance electrode to appropriate depth using a clamped manipulator with Vernier scale or microdrive. The guide tube is then raised while the stylus depth is maintained (Figures 2C-2F).
Alternatives: Use a longer guide tube in a stylus free preparation. In that scenario, advance the electrode until its shank is flush with the rear of the guide tube. Then, remove the guide tube. Advance the electrode to target depth. This approach will also work if the tail of the electrode shank will be approximately positioned at the top of the bone screw, and will thus provide its own guidance on depth.
19. Check tail length of electrode after dropping to confirm depth ( Figure 2G). Glue electrode into bone screw head using one drop of quick curing silicone to stabilize the electrode in the bone screw. 20. Repeat for the second electrode. The tweezers are masked with electrical tape except around the tips. Not shown: the tweezers squeeze the joint between the two wires creating an electrical path from one tweezer face, through both wires, to the other tweezer face. This path creates a thermocouple between the two wires which, with appropriate current pulse, will melt the two wires together. A pulse is delivered. (B) The two wires with a weld are shown. Scale bar = 1 mm. The non-stranded wire used in these images is 0.003'' wide at its uninsulated section.

OPEN ACCESS
STAR Protocols 3, 101136, March 18, 2022 21. A surgical site for the IPG is located on the back, caudal enough to avoid touching the scapula, and lateral enough to avoid touching the spine. Make a horizontal incision perpendicular to the spine. Dissect a pocket bluntly in the subcutaneous space to fit the IPG caudal to the incision. 22. Use a tunneling kit to tunnel from the head surgical location to the back incision.
Note: Large hemostats (8'' or 12'') are useful surgical tools before moving to the tunneling tool.
a. The tunneling tool is advanced subcutaneously from scalp incision to IPG pocket. b. The straw is left in the tunnel, and the pigtail wire inserted through the straw.
Note: The pigtail wire is a Boston Scientific product, and contains an industry standard 8 lead connector on one end, and eight stranded wires on the other end.
c. Once the pigtail is inserted, the straw is removed. Figure 3.

Welding. See
a. Use a Sunstone 100 ws single pulse resistance spot welder, along with its Tweezer hand piece weld attachment. b. Set the energy to 4% with a 0.45 ms pulse width. c. Use electrical tape to mask the Tweezer except where the two faces come together. d. The electrode wire tail, and pigtail wire, both mechanically stripped of insulation, are overlaid at a 90 degree angle in between the tweezer faces so that the only current path is from one tweezer face through the joint between the wires to the other tweezer face.
Note: Typically two people are needed to hold the Tweezer and both wires. One person holds both wires at 90 degree angles, while the other positions the Tweezer.
e. Then, a pulse can be used to spot weld the wires. Wire alignment on the Tweezers is critical, and practice prior to surgery recommended. These welding settings are specific to the wire diameters and composition used in this study.
CRITICAL: The welding step must not be performed while the IPG is connected to avoid damage to the IPG.
24. Check channel number using a multimeter. The connectivity between the weld and the pigtail connector contacts can be used to determine which channel is attached to the electrode. A nonsterile multimeter and two sets of sterilized alligator clip wires, and two pairs of forceps, are used to find the connector contact matching each weld. 25. Check impedance.
a. Plug the pigtail connector into the IPG, and place IPG into a previously created subcutaneous pocket. b. Test impedance using a Boston Scientific tablet, which is a specific accessory associated with their IPGs.
Note: Impedances in the 4 kOhm range or lower are expected. Impedances above 10 kOhm indicate replacement of the electrode should occur.
Note: Often the bone screw will need to be removed to facilitate removal, if needed. Once both electrodes are welded and verified, move on.
26. Place a drop of silicone on the skull for each of the two welds. Hold the uninsulated section of the welds in the silicone to insulate them.

OPEN ACCESS
27. Flush head and IPG surgical sites with 3% hydrogen peroxide diluted 50:50 in 0.9% NaCl irrigation solution. Insert IPG into the subcutaneous pocket. Coil extra pigtail wire under IPG. Silk sutures may be used to secure the IPG. Subcutaneous silk sutures should be used to secure the pigtail at the cranial location to avoid pigtail creep. 28. Suture both sites closed using interrupted 3-0 Vicryl for the galea (scalp) or cutaneous (back) layer, and a subcuticular running 4-0 Monocryl for skin, and apply Dermabond to the incisions.
The relative positions of pigtail wire, electrodes, and IPG are shown in Figure 4.
29. Treat the monkey post-operatively with opioid analgesics (e.g., buprenorphine) for up to 72 h, followed by nonsteroidal anti-inflammatory drugs (e.g., ketoprofen) for up to 1 week.
Note: Suspected infections around the wound margin may (rarely) require antibiotic treatment. Suspected brain edema may require treatment with steroid drugs (e.g. dexamethasone).

Stimulation electrode implantation -Stimulation through external connector
Timing: 3 h In some preparations it is desirable to have the connector to the electrode externalized through an acrylic cap that will also be used to anchor recording chambers. This alternate approach describes implantation of stimulation electrodes to deliver stimulation into NB using an external stimulator, connected to the electrodes through a removable connector, in dental cement or acrylic.
Repeat steps 1-21 of electrode fabrication and stimulation electrode implantation, above.
30. Once both electrodes are implanted, and wires are loose, the wires are attached to the connector via soldering. A bare piece of platinum wire is wrapped around a separate bone screw to act as the ground. 31. The wires and connector are encased in a bone-cement head cap, adhering to the screws. 32. Implantation of additional screws may be needed for added stability. Alternatively, this procedure may be performed in monkeys who already have a headpost/recording cylinder implanted and covered by bone-cement. The bone cement that has been added to stabilize the electrodes then attaches to the rest of the implant. 33. A cap may be added to protect the connector; all parts of the electrode are encased in acrylic at the end of the procedure.

Combined stimulation -Neural recording
Timing: 2 h Monkeys that have already had cranial implants that allow neurophysiological recordings prior to the stimulation electrode implantation (e.g., headpost and cranial well that allows insertion and removal of micro-electrodes through a craniotomy into the brain) may undergo simultaneous deep brain stimulation and neural recording with a data acquisition system such as the APM recording system. The first step in this experiment is to load the electrodes into the micro-electrode microdrive and prepare and test the neural recording system (see Figure 5 for this stage).
34. Head fix monkey and put recording stage on and connect to preamplifier 35. Test impedance of stimulation electrode by connecting external connector to an impedance meter. The use of implanted electrodes includes the possibility of failure, and regular impedance checks are necessary to ensure connectivity (see also problem 1 in Troubleshooting section). 36. Connect stimulator to stimulation electrode (e.g., isolated pulse stimulator Model 2100, A-M Systems, Sequim WA).
Note: Connect a 10 mF capacitor in series to the stimulating electrode to prevent net charge accumulation.
37. Advance the micro-electrodes into the brain (e.g., prefrontal cortex) through the cranial well and stop when neural signals have been isolated. 38. Start behavioral task, implemented in the MATLAB environment e.g., as described in (Meyer and Constantinidis, 2005) or equivalent. Behavioral program will trigger stimulation.
CRITICAL: During the time interval of electrical stimulation, electrical artifacts are visible in the neurophysiological data acquisition. These make recording of neurophysiological signals anywhere in the brain impossible, for a brief period of time (<1 s). It is imperative therefore that the behavioral task and data acquisition be structured so that artifacts do not interfere with recordings during task execution. At the same time, presence of artifacts

OPEN ACCESS
in the neurophysiological recording when delivering electrical stimulation provides confirmation that the stimulation circuit is closed and the current application is effective.
39. The recommendation is to construct a recording task consisting of 45 s of trials, with 15 s pauses interleaved. During the 15 s pauses, biphasic stimulation can be delivered to one or both hemispheres at intensities up to 0.5 mA, with 100 microsecond phase length, biphasic charge balanced pulses. Good effects of stimulation have been documented at 80 pulses per second during that 15 s period (Liu et al., 2017;Qi et al., 2021). 40. Retract the recording electrodes first from the recording chamber; when task is done then power off and disconnect the stimulator. 41. Clean cranial well with sterile saline. Remove guide tube and electrodes, clean with alcohol followed by distilled water and sterilize with UV irradiation.
Combined stimulation/IPG -Behavioral recording (see Figure 6 for this stage) Timing: 1 h Monkeys that have not had cranial implants that allow neurophysiological recordings, or when obtaining behavioral measures alone is adequate, NB stimulation can be performed around behavioral recording. An example is shown in Figure 6.
42. Test impedance of stimulation electrode. Connections between the implanted electrodes and the IPG may fail, and regular impedance checks are necessary to ensure connectivity. The IPG system allows impedance testing remotely. 43. Create a program of stimulation. Intermittent stimulation comprises biphasic pulses delivered to one or both hemispheres at intensities up to 0.5 mA, with 100 ms phase duration. Positive effects of stimulation have been documented at 80 pulses per second during that 15 s period. The activating function is proportional to the square root of the current (McIntyre et al., 2004;Murasugi et al., 1993;Stoney et al., 1968), and currents of 200-500 microamps per pulse have been used in the macaque 44. Prepare and test behavioral control system. 45. Apply stimulation for a period of 1 h, starting stimulation at the same time the monkey engages with a behavioral task. 46. Daily sessions of stimulation during the task may alternate with sessions beginning after the 1 h of stimulation.

EXPECTED OUTCOMES
Absolute verification of electrode placement requires histological verification that the tip is positioned in the floor of the globus pallidus, with anterior-posterior position determined by where the anterior commissure crosses the globus pallidus (Figure 7). Electrodes may be imaged using CT postoperatively to validate placement, but final placement should be verified histologically.
We have demonstrated that these stimulation protocols improve working memory performance (Liu et al., 2017;Qi et al., 2021), and sustained attention performance (Liu et al., 2018), generate long-term neuroplasticity based effects (Liu et al., 2017), and desynchronize the EEG (Qi et al., 2021). During recording, for a successful stimulation we will expect to see artifacts from recording

OPEN ACCESS
electrodes during each stimulation pulse. Animals show no observable reactions to this form of brain stimulation, and in particular no consistent pupil dilation or contraction occurs.

LIMITATIONS
Practically, regular verification of electrode impedance and visualization of stimulation artifacts when recording are the primary methods used to ensure the circuits are intact. Reliable positioning of the animal into the stereotax using aural and orbital ridge landmarks is critical to success, and can be practiced with plastic model skulls to verify reproducibility. This process is the largest potential source of error in positioning. Enabling the tail end of the electrode to be visible in acrylic cap preparations allows verification of depth, and behavioral impairment via high frequency stimulation (Liu et al., 2017) is robust and useful to improve confidence in electrode positioning. Better biomarkers of appropriate electrode placement prior to histological verification are an area for improvement.

TROUBLESHOOTING Problem 1
When the electrode is coupled to an acrylic cap, wiring may break; for this reason impedance checks are performed manually each day to verify an intact circuit (step 35).

Potential solution
We always manually check the impedance of an actual 10 kOhm resistor before checking the implanted animal to ensure the impedance checking procedure is competent. If the circuit is broken, the most likely point of failure is in the wire where it exits the electrode tubing. We have increased the wire diameter from 50 to 75 mm from the same supplier and this problem has become rarer.

Problem 2
Stimulation parameters may not be implemented correctly (step 39).

Potential solution
The stimulation pulses should be placed on an oscilloscope during sessions to verify the voltage is negative at the electrode first (cathodal biphasic pulses).

Problem 3
It may not be possible to synchronize stimulation perfectly with events of the behavioral task (step 46).

Potential solution
We have reconstructed behavioral performance relative to time of stimulation onset in multiple experiments in each of our labs, and found performance is remarkably insensitive to the timing of the trial relative to stimulation during intermittent stimulation.

RESOURCE AVAILABILITY
Lead contact Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, David T. Blake, dblake@augusta.edu.

Materials availability
No new mouse lines or materials were generated for this protocol. Boston Scientific pigtail wires and IPGs were provided by Boston Scientific through an agreement modeled on the BRAIN Initiative Public-Private Partnership templates.

Data and code availability
No new data or code was generated for this protocol.