This research was supported by the National Institute of Mental Health MH 086960-01A1 (Christa K. McIntyre).

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
<ol>
	<li>Create a drilling tool by sawing off the sharp end of a 22 &#189; G needle.</li>
	<li>Run the now blunt end of the 22 &#189; G needle over a metal file several times to flatten it. Hold the needle at a 45&#186; 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.</li>
	<li>Use magnifying binoculars for the remaining steps in section 1.</li>
	<li>Using a scalpel (10 or 15 blade) cut 4 mm segments of tubing.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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).</li>
	<li>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.</li>
	<li>Prepare wires for cuffs.
	<ol>
		<li>Cut platinum iridium wire into 70 mm segments.</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Wire the cuff (Figure 1D). 
	NOTE: Steps in section 1.10 must be done under magnifying binoculars.
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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 &#8216;trough&#8217; 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.&#160;</li>
		<li>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.</li>
		<li>Repeat steps 1.10.2 &#8211; 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.</li>
		<li>Place a gold pin into the arm of the helping hand with the hole facing up. Fill the hole with flux.</li>
		<li>Solder the insulated end of the wire (now attached to the cuff) into the pin.</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>
2. Construction of Headcap for VNS Input Site
<ol>
	<li>Cut 30 mm segments of 26 AWG copper wire. Strip a small portion on each end.</li>
	<li>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).</li>
	<li>Mark one of the wires with sharpie. During the surgery, position the implant with the marked wire rostral to the unmarked wire.</li>
</ol>
3. VNS Surgery
<ol>
	<li>Create custom glass tools for handling of vagus nerve during surgery.
	<ol>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>VNS surgical procedures
	<ol>
		<li>Gather and sterilize all tools. Prepare a sanitary, heated surgical area.</li>
		<li>Anesthetize animal with ketamine/xylazine (85 mg/kg, 5mg/kg, IP). Assess the depth of the anesthetic plane by monitoring the animal&#8217;s vocalizations and withdrawal-reflexes in response to toe and/or tail pinching.</li>
		<li>Shave the top of head and left side of animal&#8217;s neck. Protect the animal&#8217;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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Pull the incision site open with hemostats. Using cotton swabs, apply hydrogen peroxide to the exposed skull to remove any remaining tissue.</li>
		<li>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.&#160;
		<ol>
			<li>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.</li>
		</ol>
		</li>
		<li>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.</li>
		<li>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.</li>
		<li>Use magnifying binoculars for the remaining steps in section 3.2.</li>
		<li>Remove the animal from the stereotaxic instrument. Place the animal on its right side, rotating it slightly toward the ventral position.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Identify the vagus nerve. It is the largest nerve in the carotid sheath and is usually to the animal&#8217;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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Make sure the cuff is oriented with the marked &#8216;top&#8217; 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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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).</li>
	</ol>
	</li>
</ol>
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.
<ol>
	<li>House animals on a 12 hr&#160;light/dark cycle with ad libitum access to food and water. Handle animals daily during recovery from surgery.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Wipe chambers with 70% ethanol before and after each session to eliminate olfactory cues.</li>
	<li>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.
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>On the third day, test the strength of the tone/shock association.&#160;Play 4 tones with an ISI of 3, 4, or 5 min (4 min average) in the absence of footshocks and record the animals&#8217; freezing behavior during the tone presentations and during the inter-stimulus-intervals as measures of the conditioned fear response (CFR).</li>
	<li>On day 4, begin extinction training with VNS or Sham VNS.
	<ol>
		<li>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 &#181;s 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.</li>
	</ol>
	</li>
	<li>Periodically test the electrical integrity of cuff and input site using an oscilloscope.
	<ol>
		<li>Plug the animal in as normal and split the output from the stimulator to the oscilloscope.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>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).</li>
	<li>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.&#160;</li>
</ol>
5. In Vivo Recordings of Evoked Field Potentials
Note: This step is optional. Evoked field potentials (EFPs) are recorded 24 hr&#160;after tests of reinstatement (Day 5) in isoflurane-anesthetized rats mounted in a stereotaxic apparatus, following standard procedures23,24.
<ol>
	<li>Gather and sterilize all tools.</li>
	<li>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&#8217;s vocalizations and withdrawal-reflexes in response to toe and/or tail pinching. Use mineral oil or eye ointment to protect eyes.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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).</li>
	<li>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&#8211;output curve determined before collection of baseline data), delivered every 15 sec.</li>
	<li>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.</li>
	<li>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.</li>
	<li>After the end of the recording decapitate the anesthetized animal and extract the brain. Prepare tissue for histological verification of electrode placement.</li>
</ol>

The authors have no competing interests or conflicts.

We present here a protocol that is used to facilitate extinction of conditioned fear during a single session of exposure to conditioned cues19 and to modulate plasticity in the pathway between the infralimbic cortex and the basolateral amygdala that may mediate extinction learning20. A crucial step for the success of this protocol is the proper delivery of VNS during extinction training. Therefore, special care should be given to the construction of the cuff electrodes and the placement of the cuff ...

Classical fear conditioning provides a widely used animal model to study the biological basis of anxiety disorders. During fear conditioning, an aversive stimulus (the unconditioned stimulus, US, e.g., a footshock) is presented in conjunction with a neutral stimulus, such as a tone and/or a context (the conditioned stimulus; CS). During fear conditioning, associations between the CS and the US are formed. Eventually the presentation of the CS alone elicits a fear response (the conditioned response; CR). In fear extinction, the CS is presented repeatedly in the absence of the US, causing the CR to gradually diminish1. Thus, extinction of conditioned fear is an active process in which fearful behavioral responses to neutral stimuli are attenuated when they no longer predict aversive outcomes. Extinction of conditioned responses requires consolidation of new memories that compete with learned associations. A hallmark of anxiety disorders is impaired extinction2-4. Thus, extinction of conditioned fear in animal models serves as an important paradigm both for inhibitory learning and as a model of behavior therapy for human anxiety disorders5,6.
Because there is close correspondence between fear and human anxiety, it is thought that these studies can provide insight into the biological nature of anxiety-related disorders such as post-traumatic stress disorder and will help to develop strategies to treat them. An important goal of preclinical research is to aid extinction learning and to induce targeted plasticity in extinction circuits to consolidate extinction learning. Vagus nerve stimulation (VNS) is a minimally invasive neuroprosthetic approach that might be used to provide tight temporal and circuit-specific modulation of brain areas and synapses engaged in an ongoing task. A series of recent studies from Michael Kilgard&#39;s group at The University of Texas at Dallas have shown that pairing VNS with discrete sensory or motor stimuli (e.g., a tone or a lever pull) is highly effective in promoting cortical plasticity to treat tinnitus7, or to overcome motor deficits following stroke8-10. In addition, non-contingent VNS that occurs within a short time-window after learning similarly promotes cortical plasticity and enhances memory consolidation in rats and in humans11-13.
Considering the role of the vagus nerve in the parasympathetic pathway, it is not surprising that it could participate in modulating memories and synaptic plasticity. Highly emotional events tend to produce stronger memories than non-emotional memories. This is likely due to the influence of stress hormones on memory consolidation. Posttraining administration of the stress hormone adrenaline enhances memory consolidation in human and non-human animals, but adrenaline does not cross the blood-brain-barrier14, 15. Therefore, stress-induced adrenaline release must impact the brain indirectly to enhance memory consolidation. Strong evidence suggests that the vagus nerve may be the link between circulating adrenaline and the brain. Miyashita and Williams16 found that systemic administration of adrenaline increased vagal nerve firing, and increased levels of norepinephrine in the amygdala17. Systemic administration of adrenaline does not enhance memory consolidation when &#946;-adrenergic receptors are blocked in the amygdala18 suggesting that the vagus nerve plays a role in the pathway that turns emotionally arousing experiences into long-term memories.
Thus, pairing VNS with training has the potential to enhance the brain changes that support memory consolidation and exposure to conditioned cues in the absence of reinforcement enhances the consolidation of extinction learning in rats19,20. Here we describe the use of VNS as a tool to promote cortical plasticity and facilitate extinction of a conditioned fear response.

<table><tbody><tr><td>Alcohol</td><td></td><td></td><td></td></tr><tr><td>Atropine</td><td>Fisher</td><td>A0132-5G</td><td></td></tr><tr><td>Betadine</td><td>Henry Schein</td><td>69066950</td><td></td></tr><tr><td>Hydrogen peroxide&amp;nbsp;</td><td>CVS</td><td>209478</td><td></td></tr><tr><td>Ketamine</td><td>Henry Schein&amp;nbsp;</td><td>1129300</td><td></td></tr><tr><td>Marcaine</td><td>Henry Schein</td><td>6312615</td><td></td></tr><tr><td>Mineral Oil</td><td>CVS</td><td>152355</td><td></td></tr><tr><td>Neosporin</td><td>CVS</td><td>629451</td><td></td></tr><tr><td>Oxygen</td><td>Home Depot</td><td>304179</td><td></td></tr><tr><td>Pennicillin</td><td>Fisher</td><td>PENNA-10MU</td><td></td></tr><tr><td>Propane</td><td>Home Depot</td><td>304182</td><td></td></tr><tr><td>Xylazine</td><td>Henry Schein</td><td>4019308</td><td></td></tr><tr><td>Tools</td><td></td><td></td><td></td></tr><tr><td>Jewelery Torch</td><td>Smith Equipment</td><td>23-1001D</td><td></td></tr><tr><td>Sewing Needle</td><td>Walgreens</td><td>441831</td><td></td></tr><tr><td>#5 Forceps (2)</td><td>Fine Science Tools</td><td>11254-20</td><td></td></tr><tr><td>Soldering Iron</td><td>Home Depot&amp;nbsp;</td><td>203525863</td><td></td></tr><tr><td>AmScope SM-4TX-144A 3.5X-45X Circuit Board Boom Stereo Microscope + 144 LED</td><td>AmScope</td><td>SM-4TX-144A</td><td></td></tr><tr><td>Helping Hands</td><td>A-M Systems&amp;nbsp;</td><td>726200</td><td></td></tr><tr><td>Scalpel Blade Holder</td><td>Fine Science Tools</td><td>10003-12</td><td></td></tr><tr><td>Metal File</td><td>Home Depot</td><td>6601</td><td></td></tr><tr><td>Ruler</td><td>Home Deopt</td><td>202035324</td><td></td></tr><tr><td>Curved Hemostats&amp;nbsp;</td><td>Fine Science Tools</td><td>130009-12</td><td></td></tr><tr><td>Fine Scissors</td><td>Fine Science Tools</td><td>14058-09</td><td></td></tr><tr><td>Spatula</td><td>Fine Science Tools</td><td></td><td></td></tr><tr><td>Small Screwdriver</td><td>Home Depot</td><td>646507</td><td></td></tr><tr><td>Magnetic Fixator Retraction System</td><td>Fine Science Tools</td><td>18200-04, 18200-01, 18200-05</td><td></td></tr><tr><td>Heating Pad</td><td>Walgreens</td><td>30294</td><td></td></tr><tr><td>Clippers</td><td>Walgreens</td><td>277966</td><td></td></tr><tr><td>Sharpie</td><td>Staples</td><td>125328</td><td></td></tr><tr><td>Ring Forceps</td><td>Fine Science Tools</td><td>11103-09</td><td></td></tr><tr><td>Custom Micropipette Glass Tools (J shape and Straight) - Borosilicate glass</td><td>Sutter Instrument</td><td>B150-110-10</td><td></td></tr><tr><td>Adson Forceps</td><td>Fine Science Tools</td><td>11006-12</td><td></td></tr><tr><td>Cuffs</td><td></td><td></td><td></td></tr><tr><td>Tubing</td><td>Braintree Scientific Inc</td><td>MRE-065</td><td></td></tr><tr><td>Platinum Iridium Wire</td><td>Medwire</td><td>10IR9/49T</td><td></td></tr><tr><td>Gold Pins</td><td>Mill-Max</td><td>1001-0-15-15-30-27-04-0</td><td></td></tr><tr><td>Suture Thread</td><td>Henry Schein</td><td>100-5797</td><td></td></tr><tr><td>22 G needles</td><td>Fisher&amp;nbsp;</td><td>14-815-525</td><td></td></tr><tr><td>Paper Tape</td><td>Fisher&amp;nbsp;</td><td>03-411-602</td><td></td></tr><tr><td>Solder</td><td>Home Depot</td><td>327793</td><td></td></tr><tr><td>Flux&amp;nbsp;</td><td>Home Depot</td><td>300142</td><td></td></tr><tr><td>Scalpel Blade, 10 or 15</td><td>Stoelting</td><td>52173-10</td><td></td></tr><tr><td>Silastic Laboratory Tubing .51 mm ID x .94 mm OD</td><td>Fisher&amp;nbsp;</td><td>508-002</td><td></td></tr><tr><td>Headcaps</td><td></td><td></td><td></td></tr><tr><td>Connector Pieces (male)</td><td>Omnetics Connector Corporation</td><td>A25001-004</td><td></td></tr><tr><td>Headcap pieces (female)</td><td>Omnetics Connector Corporation</td><td>A24001-004</td><td></td></tr><tr><td>Teets Dental Acrylic, Liquid and Powder</td><td>A-M Systems</td><td>525000, 526000</td><td></td></tr><tr><td>26 Gauge Solid Copper Wire</td><td>Staples</td><td>1016882 &amp;nbsp;</td><td></td></tr><tr><td>Surgery</td><td></td><td></td><td></td></tr><tr><td>Bone Screws</td><td>Stoelting+CB33:C61</td><td>51457</td><td></td></tr><tr><td>Scalpel Blades, 10 or 15</td><td>Stoelting</td><td>52173-10</td><td></td></tr><tr><td>1 ml syringes</td><td>Fisher</td><td>14-826-261</td><td></td></tr><tr><td>22 G Needles</td><td>Fisher&amp;nbsp;</td><td>14-815-525</td><td></td></tr><tr><td>27 G Needles</td><td>Fisher</td><td>14-826-48</td><td></td></tr><tr><td>2&quot; 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vagus nerve stimulation as a tool to induce plasticity in pathways relevant for extinction learning

Extinction describes the process of attenuating behavioral responses to neutral stimuli when they no longer provide the reinforcement that has been maintaining the behavior. There is close correspondence between fear and human anxiety, and therefore studies of extinction learning might provide insight into the biological nature of anxiety-related disorders such as post-traumatic stress disorder, and they might help to develop strategies to treat them. Preclinical research aims to aid extinction learning and to induce targeted plasticity in extinction circuits to consolidate the newly formed memory. Vagus nerve stimulation (VNS) is a powerful approach that provides tight temporal and circuit-specific release of neurotransmitters, resulting in modulation of neuronal networks engaged in an ongoing task. VNS enhances memory consolidation in both rats and humans, and pairing VNS with exposure to conditioned cues enhances the consolidation of extinction learning in rats. Here, we provide a detailed protocol for the preparation of custom-made parts and the surgical procedures required for VNS in rats. Using this protocol we show how VNS can facilitate the extinction of conditioned fear responses in an auditory fear conditioning task. In addition, we provide evidence that VNS modulates synaptic plasticity in the pathway between the infralimbic (IL) medial prefrontal cortex and the basolateral complex of the amygdala (BLA), which is involved in the expression and modulation of extinction memory.

This section illustrates examples of results that can be obtained by using VNS in combination with extinction learning to reduce the expression of the conditioned fear response in rats. For Days 1 and 2 (Auditory Fear Conditioning), rats were trained on an auditory fear conditioning task in which footshocks were paired with a tone. On Day 3 (Pre Treatment Test), tones were presented in the absence of footshocks to measure freezing levels and infer conditioned fear response acquisition. On Day 4 (Treatment) rats received ...

Watch this Scientific Journal Video about Vagus Nerve Stimulation as a Tool to Induce Plasticity in Pathways Relevant for Extinction Learning at JoVE.com

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

Vagus nerve stimulation (VNS) has emerged as a tool to induce targeted synaptic plasticity in the forebrain to modify a range of behaviors. This protocol describes how to implement VNS to facilitate the consolidation of fear extinction memory.

Extinction describes the process of attenuating behavioral responses to neutral stimuli when they no longer provide the reinforcement that has been ...

vagus-nerve-stimulation-as-tool-to-induce-plasticity-pathways

Universidad de Cartagena UDEC

Research

JoVE Journal

Behavior

23.2K Views.  The University of Texas at Dallas. Vagus nerve stimulation (VNS) has emerged as a tool to induce targeted synaptic plasticity in the forebrain to modify a range of behaviors. This protocol describes how to implement VNS to facilitate the consolidation of fear extinction memory.

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

Long-term Potentiation of Perforant Pathway-dentate Gyrus Synapse in Freely Behaving Mice

Studies of long-term potentiation of synaptic efficacy, an activity-dependent synaptic phenomenon having properties that make it attractive as a potential cellular mechanism underlying learning and information storage, have long been used to elucidate the physiology of various neuronal circuits in the hippocampus, amygdala, and other limbic and cortical structures. With this in mind, transgenic mouse models of neurological diseases represent useful platforms to conduct long-term&#160;potentiation (LTP) studies to develop a greater understanding of the role of genes in normal and abnormal synaptic communication in neuronal networks involved in learning, emotion and information processing. This article describes methodologies for reliably inducing LTP in the freely behaving mouse. These methodologies can be used in studies of transgenic and knockout freely&#160;behaving mouse models of neurodegenerative diseases.

Transgenic and knockout mouse models of neurological diseases are useful for studying the role of genes in normal and abnormal neurophysiology. This article describes methodologies which can be used to study long-term potentiation, a cellular mechanism which may underlie learning and memory, in transgenic and knockout freely&#160;behaving mouse models of neuropathology.

Studies of long-term potentiation of synaptic efficacy, an activity-dependent synaptic phenomenon having properties that make it attractive as a potential ...

A Method for Remotely Silencing Neural Activity in Rodents During Discrete Phases of Learning

This protocol describes how to temporarily and remotely silence neuronal activity in discrete brain regions while animals are engaged in learning and memory tasks. The approach combines pharmacogenetics (Designer-Receptors-Exclusively-Activated-by-Designer-Drugs) with a behavioral paradigm (sensory preconditioning) that is designed to distinguish between different forms of learning. Specifically, viral-mediated delivery is used to express a genetically modified inhibitory G-protein coupled receptor (the Designer Receptor) into a discrete brain region in the rodent. Three weeks later, when designer receptor expression levels are high, a pharmacological agent (the Designer Drug) is administered systemically 30 min prior to a specific behavioral session. The drug has affinity for the designer receptor and thus results in inhibition of neurons that express the designer receptor, but is otherwise biologically inert. The brain region remains silenced for 2-5 hr (depending on the dose and route of administration). Upon completion of the behavioral paradigm, brain tissue is assessed for correct placement and receptor expression. This approach is particularly useful for determining the contribution of individual brain regions to specific components of behavior and can be used across any number of behavioral paradigms.

This protocol describes how to temporarily and remotely silence neuronal activity in discrete brain regions while rats are engaged in learning and memory tasks. The approach combines pharmacogenetics (Designer-Receptors-Exclusively-Activated-by-Designer-Drugs) with a behavioral paradigm (sensory preconditioning) that is designed to distinguish between different components of learning.

This protocol describes how to temporarily and remotely silence neuronal activity in discrete brain regions while animals are engaged in learning and ...

Extinction Training During the Reconsolidation Window Prevents Recovery of Fear

Fear is maladaptive when it persists long after circumstances have become safe. It is therefore crucial to develop an approach that persistently prevents the return of fear. Pavlovian fear-conditioning paradigms are commonly employed to create a controlled, novel fear association in the laboratory. After pairing an innocuous stimulus (conditioned stimulus, CS) with an aversive outcome (unconditioned stimulus, US) we can elicit a fear response (conditioned response, or CR) by presenting just the stimulus alone1,2 . Once fear is acquired, it can be diminished using extinction training, whereby the conditioned stimulus is repeatedly presented without the aversive outcome until fear is no longer expressed3. This inhibitory learning creates a new, safe representation for the CS, which competes for expression with the original fear memory4. Although extinction is effective at inhibiting fear, it is not permanent. Fear can spontaneously recover with the passage of time. Exposure to stress or returning to the context of initial learning can also cause fear to resurface3,4.
Our protocol addresses the transient nature of extinction by targeting the reconsolidation window to modify emotional memory in a more permanent manner. Ample evidence suggests that reactivating a consolidated memory returns it to a labile state, during which the memory is again susceptible to interference5-9. This window of opportunity appears to open shortly after reactivation and close approximately 6hrs later5,11,16, although this may vary depending on the strength and age of the memory15. By allowing new information to incorporate into the original memory trace, this memory may be updated as it reconsolidates10,11. Studies involving non-human animals have successfully blocked the expression of fear memory by introducing pharmacological manipulations within the reconsolidation window, however, most agents used are either toxic to humans or show equivocal effects when used in human studies12-14. Our protocol addresses these challenges by offering an effective, yet non-invasive, behavioral manipulation that is safe for humans.
By prompting fear memory retrieval prior to extinction, we essentially trigger the reconsolidation process, allowing new safety information (i.e., extinction) to be incorporated while the fear memory is still susceptible to interference. A recent study employing this behavioral manipulation in rats has successfully blocked fear memory using these temporal parameters11. Additional studies in humans have demonstrated that introducing new information after the retrieval of previously consolidated motor16, episodic17, or declarative18 memories leads to interference with the original memory trace14. We outline below a novel protocol used to block fear recovery in humans.

Conditioned fear can be diminished through an inhibitory process called extinction, but can resurface under conditions such as the passage of time or exposure to stress. Our protocol presents a novel way of preventing fear recovery by introducing extinction during the reconsolidation window (the re-storage phase of a reactivated memory).

Fear is maladaptive when it persists long after circumstances have become safe. It is therefore crucial to develop an approach that persistently prevents ...

Neuroscience

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated proteins and subsequent manipulation of neural activity by light. Channelrhodopsins (ChRs) are light-gated cation-channels and when fused to a fluorescent protein their expression allows for visualization and concurrent activation of specific cell types and their axonal projections in defined areas of the brain. Via stereotactic injection of viral vectors, ChR fusion proteins can be constitutively or conditionally expressed in specific cells of a defined brain region, and their axonal projections can subsequently be studied anatomically and functionally via ex vivo optogenetic activation in brain slices. This is of particular importance when aiming to understand synaptic properties of connections that could not be addressed with conventional electrical stimulation approaches, or in identifying novel afferent and efferent connectivity that was previously poorly understood. Here, a few examples illustrate how this technique can be applied to investigate these questions to elucidating fear-related circuits in the amygdala. The amygdala is a key region for acquisition and expression of fear, and storage of fear and emotional memories. Many lines of evidence suggest that the medial prefrontal cortex (mPFC) participates in different aspects of fear acquisition and extinction, but its precise connectivity with the amygdala is just starting to be understood. First, it is shown how ex vivo optogenetic activation can be used to study aspects of synaptic communication between mPFC afferents and target cells in the basolateral amygdala (BLA). Furthermore, it is illustrated how this ex vivo optogenetic approach can be applied to assess novel connectivity patterns using a group of GABAergic neurons in the amygdala, the paracapsular intercalated cell cluster (mpITC), as an example.

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are widely used to manipulate neural activity and assess the consequences for brain function. Here, a technique is outlined that&#160;upon in vivo expression of the optical activator Channelrhodopsin, allows for ex vivo analysis of synaptic properties of specific long range and local neural connections in fear-related circuits.

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated ...

Assessment of Long-term Depression Induction in Adult Cerebellar Slices

Synaptic plasticity provides a mechanism for learning and memory. For cerebellar motor learning, long-term depression (LTD) of synaptic transmissions from parallel fibers (PF) to Purkinje cells (PC) is considered the basis for motor learning, and deficiencies of both LTD and motor learning are observed in various gene-manipulated animals. Common motor learning sets, such as adaptation of the optokinetic reflex (OKR), the vestibular-ocular reflex (VOR), and rotarod test were used for evaluation of motor learning ability. However, results obtained from the GluA2-carboxy terminus modified knock-in mice demonstrated normal adaptation of the VOR and the OKR, despite lacking PF-LTD. In that report, induction of LTD was only attempted using one type of stimulation protocol at room temperature. Thus, conditions to induce cerebellar LTD were explored in the same knock-in mutants using various protocols at near physiological temperature. Finally, we found stimulation protocols, by which LTD could be induced in these gene-manipulated mice. In this study, a set of protocols are proposed to evaluate LTD-induction, which will more accurately allow examination of the causal relationship between LTD and motor learning. In conclusion, experimental conditions are crucial when evaluating LTD in gene-manipulated mice.

In some gene-manipulated animals, using a single protocol may fail to induce LTD in cerebellar Purkinje cells, and there may be a discrepancy between LTD and motor learning. Multiple protocols are necessary to assess LTD-induction in gene-manipulated animals. Standard protocols are shown.

Synaptic plasticity provides a mechanism for learning and memory. For cerebellar motor learning, long-term depression (LTD) of synaptic transmissions from ...

Vagus Nerve Stimulation As an Adjunctive Neurostimulation Tool in Treatment-resistant Depression

Vagus nerve stimulation (VNS) is an approved neurostimulation therapy. The purpose of the method is to treat patients with therapy-resistant depression (TRD). VNS exhibits antidepressive and stabilizing effects. This method is particularly useful as a long-term treatment, in which up to two-thirds of patients respond. The vagus nerve stimulator is positioned on the left vagus nerve during a surgical procedure and is activated telemetrically by a wand connected to a handheld computerized device. The treating physician can perform various adjustments of the vagus nerve stimulator during in-office visits (e.g., by modifying stimulation intensity or stimulation frequency) to achieve maximum therapeutic effects with low side effects. Set-up of the device usually takes several months. Typical side effects include wound infection, temporary salivation, coughing, paralysis of the vocal cords, bradycardia, or even asystole. The patient can stop the VNS by placing a magnet over the generator. The current protocol describes delivery of the specific stimulation tool and methods for adjusting the tuning parameters to achieve the best remission rates in patients with TRD.

Vagus nerve stimulation (VNS) has been shown to be effective as an adjunct treatment for treatment-resistant depression (TRD). VNS leads to antidepressive and antisuicidal effects and quality of life improvements. This protocol offers a step-by-step guide to managing and adjusting a vagus nerve stimulator for efficient treatment of TRD.

Vagus nerve stimulation (VNS) is an approved neurostimulation therapy. The purpose of the method is to treat patients with therapy-resistant depression ...

Medicine

Preparation of Peripheral Nerve Stimulation Electrodes for Chronic Implantation in Rats

Peripheral nerve cuff electrodes have long been used in the neurosciences and related fields for stimulation of, for example, vagus or sciatic nerves. Several recent studies have demonstrated the effectiveness of chronic VNS in enhancing central nervous system plasticity to improve motor rehabilitation, extinction learning, and sensory discrimination. Construction of chronically implantable devices for use in such studies is challenging due to rats&#8217; small size, and typical protocols require extensive training of personnel and time-consuming microfabrication methods. Alternatively, commercially available implantable cuff electrodes can be purchased at a significantly higher cost. In this protocol, we present a simple, low-cost method for construction of small, chronically implantable peripheral nerve cuff electrodes for use in rats. We validate the short and long-term reliability of our cuff electrodes by demonstrating that VNS in ketamine/xylazine anesthetized rats produces decreases in breathing rate consistent with activation of the Hering-Breuer reflex, both at the time of implantation and up to 10 weeks after device implantation. We further demonstrate the suitability of the cuff electrodes for use in chronic stimulation studies by pairing VNS with skilled lever press performance to induce motor cortical map plasticity.

Existing approaches for constructing chronically implantable peripheral nerve cuff electrodes for use in small rodents often require specialized equipment and/or highly trained personnel. In this protocol we demonstrate a simple, low-cost approach for fabricating chronically implantable cuff electrodes, and demonstrate their effectiveness for vagus nerve stimulation (VNS) in rats.

Peripheral nerve cuff electrodes have long been used in the neurosciences and related fields for stimulation of, for example, vagus or sciatic nerves. ...

Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex

Studying the physiological properties of specific synapses in the brain, and how they undergo plastic changes, is a key challenge in modern neuroscience. Traditional in vitro electrophysiological techniques use electrical stimulation to evoke synaptic transmission. A major drawback of this method is its nonspecific nature; all axons in the region of the stimulating electrode will be activated, making it difficult to attribute an effect to a particular afferent connection. This issue can be overcome by replacing electrical stimulation with optogenetic-based stimulation. We describe a method for combining optogenetics with in vitro patch-clamp recordings. This is a powerful tool for the study of both basal synaptic transmission and synaptic plasticity of precise anatomically defined synaptic connections and is applicable to almost any pathway in the brain. Here, we describe the preparation and handling of a viral vector encoding channelrhodopsin protein for surgical injection into a pre-synaptic region of interest (medial prefrontal cortex) in the rodent brain and making of acute slices of downstream target regions (lateral entorhinal cortex). A detailed procedure for combining patch-clamp recordings with synaptic activation by light stimulation to study short- and long-term synaptic plasticity is also presented. We discuss examples of experiments that achieve pathway- and cell-specificity by combining optogenetics and Cre-dependent cell labeling. Finally, histological confirmation of the pre-synaptic region of interest is described along with biocytin labeling of the post-synaptic cell, to allow further identification of the precise location and cell type.

Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex

Here we present a protocol describing viral transduction of discrete brain regions with optogenetic constructs to permit synapse-specific electrophysiological characterization in acute rodent brain slices.

Studying the physiological properties of specific synapses in the brain, and how they undergo plastic changes, is a key challenge in modern neuroscience. ...

Using Optogenetics to Reverse Neuroplasticity and Inhibit Cocaine Seeking in Rats

This protocol demonstrates the steps needed to use optogenetic tools to reverse cocaine-induced plasticity at thalamo-amygdala circuits to reduce subsequent cocaine seeking behaviors in the rat. In our research, we had found that when rats self-administer intravenous cocaine paired with an audiovisual cue, synapses formed at inputs from the medial geniculate nucleus of the thalamus (MGN) onto principal neurons of the lateral amygdala (LA) become stronger as the cue-cocaine association is learned. We hypothesized that reversal of the cocaine-induced plasticity at these synapses would reduce cue-motivated cocaine seeking behavior. In order to accomplish this type of neuromodulation in vivo, we wanted to induce synaptic long-term depression (LTD), which decreases the strength of MGN-LA synapses. To this end, we used optogenetics, which allows neuromodulation of brain circuits using light. The excitatory opsin oChiEF was expressed on presynaptic MGN terminals in the LA by infusing an AAV containing oChiEF into the MGN. Optical fibers were then implanted in the LA and 473 nm laser light was pulsed at a frequency of 1 Hz for 15 minutes to induce LTD and reverse cocaine induced plasticity. This manipulation produces a long-lasting reduction in the ability of cues associated with cocaine to induce drug seeking actions.

The methods described here outline a procedure used to optogenetically reverse cocaine-induced plasticity in a behaviorally-relevant circuit in rats. Sustained low-frequency optical stimulation of thalamo-amygdala synapses induces long-term depression (LTD). In vivo optogenetically-induced LTD in cocaine-experienced rats resulted in the subsequent attenuation of cue-motivated drug seeking.

This protocol demonstrates the steps needed to use optogenetic tools to reverse cocaine-induced plasticity at thalamo-amygdala circuits to reduce ...

Abdominal VNS for Treating Inflammatory Conditions in Rat Models

Implantation Surgery for Abdominal Vagus Nerve Stimulation and Recording Studies in Awake Rats

Abdominal vagus nerve stimulation (VNS) can be applied to the subdiaphragmatic branch of the vagus nerve of rats. Due to its anatomical location, it does not have any respiratory and cardiac off-target effects commonly associated with cervical VNS. The lack of respiratory and cardiac off-target effects means that the intensity of stimulation does not need to be lowered to reduce side effects commonly experienced during cervical VNS. Few recent studies demonstrate the anti-inflammatory effects of abdominal VNS in rat models of inflammatory bowel disease, rheumatoid arthritis, and glycemia reduction in a rat model of type 2 diabetes. Rat is a great model to explore the potential of this technology because of the well-established anatomy of the vagus nerve, the large size of the nerve that allows easy handling, and the availability of many disease models. Here, we describe the methods for cleaning and sterilizing the abdominal VNS electrode array and surgical protocol in rats. We also describe the technology required for confirmation of suprathreshold stimulation by recording evoked compound action potentials. Abdominal VNS has the potential to offer selective, effective treatment for a variety of conditions, including inflammatory diseases, and the application is expected to expand similarly to cervical VNS.

Author Spotlight: Exploring Abdominal VNS for Inflammatory Conditions

The present protocol describes the surgical technique for implanting an electrode array onto the abdominal vagus nerve in rats, along with methods for chronic electrophysiology testing and stimulation using the implanted device.

Abdominal vagus nerve stimulation (VNS) can be applied to the subdiaphragmatic branch of the vagus nerve of rats. Due to its anatomical location, it does ...

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

Summary

Explore More Videos

Chapters in this video

Long-term Potentiation of Perforant Pathway-dentate Gyrus Synapse in Freely Behaving Mice

A Method for Remotely Silencing Neural Activity in Rodents During Discrete Phases of Learning

Extinction Training During the Reconsolidation Window Prevents Recovery of Fear

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Assessment of Long-term Depression Induction in Adult Cerebellar Slices

Vagus Nerve Stimulation As an Adjunctive Neurostimulation Tool in Treatment-resistant Depression

Preparation of Peripheral Nerve Stimulation Electrodes for Chronic Implantation in Rats

Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex

Using Optogenetics to Reverse Neuroplasticity and Inhibit Cocaine Seeking in Rats

Implantation Surgery for Abdominal Vagus Nerve Stimulation and Recording Studies in Awake Rats