Vagus Nerve Stimulation as a Tool to Induce Plasticity in Pathways Relevant for Extinction Learning

All procedures described in this protocol are carried out in accordance with the NIH Guide for the Care and Use of Laboratory Animals, and they were approved by the Institutional Animal Care and Use Committee of The University of Texas at Dallas.

1. Construction of VNS Cuffs

1. Create a drilling tool by sawing off the sharp end of a 22 ½ G needle.
2. Run the now blunt end of the 22 ½ G needle over a metal file several times to flatten it. Hold the needle at a 45º angle to the file and run it several times over the file while rotating it. This will cause the metal to become thinner and furl inward. CAUTION: Insert the tip of the scalpel into the cut end of the needle and rotate with some downward force to unfurl the metal.
3. Use magnifying binoculars for the remaining steps in section 1.
4. Using a scalpel (10 or 15 blade) cut 4 mm segments of tubing.
5. Place one 4 mm segment over a small drill bit or other similarly shaped tool. This is to hold the tubing in place while it is being manipulated.
6. Drill 4 holes into the tubing (Figure 1A). Holes should make the four points of a 2 mm by 2 mm square and should be clean with no rough edges.
   NOTE: The drilling tool needs to be resharpened (step 1.2) every 2 or 3 holes. Difficulty with this step is almost certainly due to a drilling tool that is not sharp enough.
7. With the tubing still on the drill bit, use a scalpel to cut the tubing lengthwise between the holes so that two holes end up on either side of the cut (Figure 1B).
8. Using a sewing needle and suture thread, pass suture through the holes to create the rigging for the ultimate placement of the cuff around the vagus nerve. Start with the needle inside the cuff and pass it through one of the holes, then go back through the adjacent hole from the outside (Figure 1C). Tie the thread together ~2 cm from the plastic so that the tubing and thread make a triangle. Allow ~8 cm of thread after the knot and trim. Repeat the process for the holes on the opposite side of the cut. The tubing is now ready to be wired.
9. Prepare wires for cuffs.
   1. Cut platinum iridium wire into 70 mm segments.
   2. Using the finest tip for the jewelry torch, create a sharp flame with blue center that is as refined as possible and use it to strip ~1 cm of the plastic coating from the wire. Apply the blue center of the flame to stripped end of the wire to create a small ball. Apply the blue center of the flame to several points of the stripped portion to fuse the seven strands together. The wire in these spots will appear to kink.
   3. On the opposite end of the wire, apply the blue center of the flame to the end to create a small ball. Minimize stripping the plastic on this end.
10. Wire the cuff (Figure 1D).
    NOTE: Steps in section 1.10 must be done under magnifying binoculars.
    1. Tape the cuff down using the threads so that it is oriented with the cut running horizontally. Pull the threads tight so that the cuff is pulled open and then tape down the threads.
    2. Grasp the balled end of the stripped side of the prepared wire with #5 forceps and push through the bottom right hole. Pull the forceps out of the hole, leaving the end of the wire in the middle of the cuff.
    3. Re-grasp the balled end of the stripped wire (now in the middle of the cuff) and push it through the top right hole. Pull the forceps out of the hole, leaving the wire passed completely through the cuff and loose on the top side of the cuff.
    4. Re-grasp the balled end of the stripped wire (now outside the cuff on the top side) and push it back through the top right hole from the inside. Continue doing so until the wire is securely in place: test by tugging on the opposite end of the wire.
       NOTE: During this process it is critical to differentiate the stripped/insulated portions of the wire. The wire that is ultimately positioned in the ‘trough’ of the cuff must be stripped, but everything below the bottom holes (outside the cuff on the bottom end) must be insulated. This ensures delivery of current only to the vagus nerve. 
    5. Grasp the balled end of the insulated side and push it through the bottom right hole from the inside of the cuff, looping it around once.
    6. Repeat steps 1.10.2 – 1.10.5 on the left side of the cuff so that two wires end up fixed to the tubing, one on the right side and one on the left side.
    7. Place a gold pin into the arm of the helping hand with the hole facing up. Fill the hole with flux.
    8. Solder the insulated end of the wire (now attached to the cuff) into the pin.
    9. Allow the solder to cool, and then melt it again. This ensures a good connection between the end of the wire and the inside of the pin. Apply more solder if necessary. Repeat for second wire.
    10. With the leads running to the right, mark the top of the cuff with permanent marker. Also mark the gold pin attached to the top lead.

2. Construction of Headcap for VNS Input Site

1. Cut 30 mm segments of 26 AWG copper wire. Strip a small portion on each end.
2. Cut the narrow end off loose gold pins and solder the stripped end of the wire to the cut end of the gold pin. Create two wire/pin compounds for each desired input site. Place a connector into the helping hands and solder the wire end of a wire/pin compound to the each of the two fluxed teeth of the connector (Figure 2G).
3. Mark one of the wires with sharpie. During the surgery, position the implant with the marked wire rostral to the unmarked wire.

3. VNS Surgery

1. Create custom glass tools for handling of vagus nerve during surgery.
   1. Use borosilicate glass to pull a micropipette so that it has a long tapered tip. If no pipette puller is available, break the glass to create a longer edge.
   2. Hold the non-tapered end of the pipette with a thick cloth (to prevent burns) and press the tapered/broken end into a smooth fire-resistant surface while applying the blue flame from the jewelry torch to the tapered end. The glass will bend as it is pressed into the surface. Apply flame until a hook or J shape forms.
2. VNS surgical procedures
   1. Gather and sterilize all tools. Prepare a sanitary, heated surgical area.
   2. Anesthetize animal with ketamine/xylazine (85 mg/kg, 5mg/kg, IP). Assess the depth of the anesthetic plane by monitoring the animal’s vocalizations and withdrawal-reflexes in response to toe and/or tail pinching.
   3. Shave the top of head and left side of animal’s neck. Protect the animal’s eyes with mineral oil or eye ointment. Apply iodine cleansing solution with gauze and then alcohol with gauze to the shaved areas. Repeat once.
   4. Inject 0.05 ml marcaine subcutaneously on the top of head and allow the bolus to disperse while placing the animal into the stereotaxic instrument.
   5. Use a scalpel to make an incision in the skin on the skull to expose both lambda and bregma. Prepare a path for the cuff by using blunt forceps to tunnel subcutaneously from the incision site down the left side in front of the ear to the left side of the neck.
   6. Pull the incision site open with hemostats. Using cotton swabs, apply hydrogen peroxide to the exposed skull to remove any remaining tissue.
   7. Using a scalpel, drill two shallow starter holes in the skull to place anchor screws. They should be placed far enough apart to allow room for the implant, but not too close to the surrounding tissue. Avoid placing screws directly on the midline. 
      1. With forceps and screwdriver, drive a bone screw into both holes. The screws should be tight in the holes, with the caps 2 – 3 mm above the surface of the skull to allow room for acrylic to fill in under and around the screws.
   8. Fill the space under, around, and between the screws with a small amount of acrylic, avoiding the surrounding tissue. Then place a larger amount of acrylic in the middle of the skull between the two screws.
   9. Grab the implant so the marked wire is oriented rostral to the unmarked wire and quickly place the implant in the acrylic, taking care not to get acrylic into the gold pins, the clasp area, or the input site on top. Once positioned allow to set ~5 min until dry. This can also be done using the arm of the stereotaxic for support. Fill in any cracks or gaps between the implant and skull with a low viscosity blend of acrylic. Allow to dry.
   10. Use magnifying binoculars for the remaining steps in section 3.2.
   11. Remove the animal from the stereotaxic instrument. Place the animal on its right side, rotating it slightly toward the ventral position.
   12. Make a small incision approximately over the left jugular vein. The jaw bone and clavicle should be roughly equidistant to the incision site. Widen the incision using blunt dissection until the muscle layer is reached. The sternomastoid, sternohyoid, and omohyoid muscles should be visible. Use muscle retractors to keep the site open.
   13. Continue blunt dissecting along the natural furrows between the muscles. Look for the pulsing of the carotid artery. Heading through the muscle toward the pulsing will reveal the carotid artery. Pull the muscles back with the muscle retractor. The sheath containing the carotid artery also contains the vagus nerve. Carefully blunt dissect the sheath with the scissors.
   14. Identify the vagus nerve. It is the largest nerve in the carotid sheath and is usually to the animal’s left side of the artery but can be found on any side. Switch to the custom glass tools and separate the vagus nerve from the carotid artery. The nerve should be free of any other tissue for at least 5 mm.
   15. From the incision on the head, using the threads on the side of the cuff opposite the leads, pull the cuff through the previously made subcutaneous tunnel with forceps or small hemostats. Push it through the tissue into the incision site on the neck.
   16. Gently lift the nerve up using the glass tools and push the threads on the side of the cuff opposite the leads under the nerve. Pull the threads all the way through, taking care not to rub against the nerve. The cuff should be immediately adjacent to the nerve.
   17. Make sure the cuff is oriented with the marked ‘top’ side superior. Drop the nerve into the middle of the cuff. The nerve should now lie across both wires in the trough of the cuff (Figure 2B). Tie the threads together to close the cuff.
   18. On the headcap, plug the pins attached to the cuff into the gold pins on the stimulation input site. Plug the marked pin into the gold pin attached to the most anterior tooth on the simulation input site.
   19. To verify that the cuff properly stimulates the vagus nerve, perform a cessation of breathing test by, connecting the stimulator to the stimulation input site on the headcap and running stimulation (0.2 mA, 60 Hz, up to 10 sec). Breathing should briefly stop and heart rate should drop, verifying cuff function.
   20. Secure the pins on the stimulation input site with acrylic. Cover the wires and verify that exposed pins and wires do not lead to short circuits. Use acrylic to smooth over any bumps or to fill in any gaps. Allow to dry.
   21. Suture closed both incision sites. Inject 0.05 ml marcaine subcutaneously near the neck incision site. Apply antibiotic ointment to incision sites. Optionally, leave a male connector in place on the stimulation input site to prevent damage or obstruction during the healing process.
   22. Treat with antibiotic and follow standard post-operative care including appropriate analgesia. Return animals to animal housing facility after they regain mobility. Allow 5 days for recovery. To ensure maximal longevity and function of the headcap, house animals singly for the remainder of the experiment.
       NOTE: Sham-VNS rats undergo the same surgery, however the circuit is designed to short at the level of the headcap (i.e., a headcap is implanted and the vagus nerve is separated from the carotid artery, but no electrode cuff is placed around the nerve).

4. Auditory Fear Conditioning

NOTE: This fear conditioning protocol is more intensive than most²¹ because the goal of these experiments is to enhance extinction. With mild fear conditioning that is easily extinguished, a floor effect can obscure this enhancement.

1. House animals on a 12 hr light/dark cycle with ad libitum access to food and water. Handle animals daily during recovery from surgery.
2. Set up the conditioning and testing apparatus, consisting of an operant box housed in a sound-attenuated chamber (Figure 2C). The operant box has clear plastic walls, 20 x 20 x 20 cm, and has a stainless steel grid floor which is connected to a foot shock generator. Use a white house-light to illuminate the chamber for video recording. Use a 9 kHz, 85 dB SPL tone as the conditioned stimulus.
3. Record behavior using a digital camera located inside the chamber, above the operant box. View and monitor the session on a computer located outside the behavior room. Save videos for later analysis.
4. Wipe chambers with 70% ethanol before and after each session to eliminate olfactory cues.
5. Fear-condition the rats for 2 days (Figure 3A). Confirm that the rats are not innately afraid of the tone by presenting 5 tones (9 khz, 85 db, 30 sec) on the first day. Ensure that the freezing levels are negligible.
   1. Follow the initial tone presentations with 8 tone-footshock (1 sec, 0.5 mA) pairings on each of 2 consecutive days. Repeat the tone-footshock pairings again on the second day. Vary the inter-stimulus-interval (ISI) between 2 and 4 min, averaging 3 min for every trial. Randomize the point at which the shock occurs during the tone.
6. On the third day, test the strength of the tone/shock association. Play 4 tones with an ISI of 3, 4, or 5 min (4 min average) in the absence of footshocks and record the animals’ freezing behavior during the tone presentations and during the inter-stimulus-intervals as measures of the conditioned fear response (CFR).
7. On day 4, begin extinction training with VNS or Sham VNS.
   1. Plug the rats into the stimulator by inserting the male connectors from the stimulator into the stimulation input site. Place animals into the chamber (Figure 2A, 2C). Set the stimulator to 0.4 mA, 500 µs pulse width at 30 Hz. Set stimulation to a total duration of 30.15 sec, starting 150 msec before the onset of the tone. Play animals 4 tones (as in step 4.6) and pair each tone presentation with VNS or Sham VNS.
8. Periodically test the electrical integrity of cuff and input site using an oscilloscope.
   1. Plug the animal in as normal and split the output from the stimulator to the oscilloscope.
   2. Set the range on the oscilloscope as -20 V to +20 V and run stimulation. The waveform of the 30 Hz stimulation should be visible on the oscilloscope. Stimulations exceeding 10 V in size indicate high impedance and an improperly functioning cuff or connection at the stimulation input site.
9. To test the effect of VNS on extinction training run a second CFR test (as in step 4.6) on day 5. Record the time spent freezing during tone presentations and compare to baseline freezing recorded during the first CFR test (step 4.6).
10. Analyze the videos using an independent observer who is blind to the treatment conditions. Measure time spent freezing during tone presentations using a stopwatch. Freezing is defined as complete immobility, during which the rat exhibits rapid respiration, lowered head, and spread paws²². Analysis of freezing behavior can be split into two phases: during tone presentation and during the inter-stimulus interval. 

5. In Vivo Recordings of Evoked Field Potentials

Note: This step is optional. Evoked field potentials (EFPs) are recorded 24 hr after tests of reinstatement (Day 5) in isoflurane-anesthetized rats mounted in a stereotaxic apparatus, following standard procedures^23,24.

1. Gather and sterilize all tools.
2. Induce anesthesia with isoflurane (5% in 100% oxygen, flowrate 1l/min) in a clear plastic chamber. Assess the depth of the anesthetic plane by monitoring the animal’s vocalizations and withdrawal-reflexes in response to toe and/or tail pinching. Use mineral oil or eye ointment to protect eyes.
3. Inject 0.05 ml marcaine subcutaneously on the top of head and allow the bolus to disperse. Use a scalpel and hemostats to remove the headcap from the skull. Use as little force as possible to avoid obscuring bregma.
4. Place the animal in the stereotaxic instrument. Use a scalpel to widen the incision to expose both lambda and bregma. Maintain anesthetic plane with isoflurane (3% in 100% oxygen, flowrate 1 L/min) via a nosecone.
5. Drill holes into the skull above the infralimbic prefrontal cortex (IL) and the basolateral amygdala (BLA). Lower a glass microelectrodes (2M KCl; 1-2 MOhms resistance) into the BLA (D/V: 7.2, A/P: 2.7, M/L: 4.9 from bregma) and a stimulation electrode into the IL region of the medial prefrontal cortex (D/V: 4.6, A/P: 3.0, M/L: 0.7 from bregma) (Figure 4A).
6. Stimulate the IL to evoke EFPs in the BLA. Data shown in Figure 4 was acquired with the following settings: A stimulation pulse of 0.3 msec duration, using a stimulation intensity that corresponded to 40% of the minimum current intensity that evoked a maximum field response (based on an input–output curve determined before collection of baseline data), delivered every 15 sec.
7. Collect baseline data for a minimum of 10 – 15 min before inducing synaptic plasticity.
   NOTE: The protocol used to evoke synaptic plasticity will vary with the requirements of the experiments and must be carefully selected by each experimenter. Data in Figure 4C shows changes in the EFP following 3 bursts of 100 pulses at 50 Hz (2 sec), with 20 sec inter-burst intervals at the minimum current intensity that evoked the maximum field response.
8. Measure the amplitude of the EFP as the difference between the mean of a 5 msec window before the stimulation artifact and the mean of a 5 msec window around 20 – 25 msec after the stimulation artifact, corresponding to the negative peak of the field potential. Normalize data to baseline and set the average of a 10 min baseline as 100%. Use the averaged EFP amplitudes of another 10 min period after plasticity induction (e.g., 40 – 50 min post induction) to assess long-term changes in EFP amplitude.
9. After the end of the recording decapitate the anesthetized animal and extract the brain. Prepare tissue for histological verification of electrode placement.