Name: Stereotaxic Surgery for Excitotoxic Lesion of Specific Brain Areas in the Adult Rat (Video) | JoVE Journal
Uploaded: 2012-07-19T13:00:00.000+00:00
Duration: 12 min 45 s
Description: Learn stereotaxic surgery for targeted brain lesioning in adult rats using excitotoxin infusion. This method enables precise ablation of specific neural regions for neuroscience research.

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5 minutes

This work was supported y a CIRM predoctoral fellowship (EDK), NARSAD Young Investigator Award (DK) and the NIMH BRAINS award (R01MH087495) (DK).

Anesthesia and analgesia: Thirty minutes prior to anesthesia, inject the rat with 0.05 mg/kg subcutaneous buprenorphine for analgesia. Initiate anesthesia with 30-40 mg/kg intraperitoneal sodium pentobarbital. At this point, also inject atropine to prevent respiratory failure (0.4 mg/kg, subcutaneous) and meloxicam as further analgesic (2 mg/kg subcutaneous). If after 5 minutes, the rat is still mobile or responsive to toe pinch, give subsequent doses of sodium pentobarbital at 5 mg/kg (intraperitoneal) until the rat is unresponsive to pain. Before performing the first incision, inject lidocaine (5 mg/kg, intradermal) at the incision site for local anesthesia. Six to eight hours after initial injection, inject the rat with 0.05 mg/kg subcutaneous buprenorphine for analgesia. Buprenorphine can be injected every 6-8 hours thereafter if needed, though this is usually not necessary.
It is important to note that other forms of anesthesia can interfere with excitotoxic lesions. For example, although ketamine is a commonly used form of anesthesia in rodents, it can interfere with lesions induced with NMDA because it is an NMDA receptor antagonist. It is important to select a method of inducing anesthesia that does not reduce lesion size. If gas anesthesia is desired, most stereotaxic devices including those described here can accommodate gas mask adaptors.
Note: Materials are described further in the Table of specific reagents and equipment below.
1. Preparation of the Pump and Stereotax
<ol>
	<li>Fill a 10 μl Hamilton syringe with sterile water and mount it into the caddy of a 6 syringe programmable pump. Secure the end of the syringe plunger into the clamped holder on the pump.</li>
	<li>Prefill gas-sterilized PE20 tubing with sterile water using a needle and 1 ml syringe. Slide the open end of the tubing on to the Hamilton syringe, being careful to avoid creating any air bubbles in the tube.</li>
	<li>Clamp the 30ga, flat-cut, 1 inch infusion needle end of the tubing on to the arm of a stereotaxic device using a barrel-style electrode manipulator. Insert the infusion needle between the clamp and the grooved barrel. Secure the clamp over a piece of the tubing containing metal needle within it to avoid flattening the tubing and restricting fluid flow.</li>
	<li>Adjust the pump rate to &gt; 5 μl/min and turn on the pump. A bead of water should appear at the tip of the infusion needle. Check the length of the tubing and its joints to make sure there are no leaks.</li>
	<li>Wipe away the water bead with a sterile swab and set the pump to withdraw. Withdraw a 2 μl air bubble. The top of the bubble should be visible in the tubing.</li>
	<li>Immerse the infusion needle tip in a 20 mg/ml solution of NMDA in sterile 0.1M PBS and withdraw 4-5 μl. An air bubble should be visible between the sterile water and the NMDA solution in the tip. It is important to note that NMDA is a potent neurotoxin and caution should be used in the preparation and handling of this solution. Gloves and eye protection are advised at all times.</li>
</ol>
2. Mount the Rat in the Stereotaxic Device
<ol>
	<li>Figure 1 depicts the basic parts of the stereotaxic device. After shaving the rat's head from the line of the eyes to the ears, load the rat's front teeth over the bite bar, with the rat's body resting on a heating pad set on medium. The tongue should hang below the bite bar, with the lower jaw. If available, a temperature feedback heating pad would be preferable. Monitoring rectal temperature would also be ideal.</li>
	<li>Holding the rat's neck with thumb and forefinger, move the right horizontal ear canal onto the right ear bar. The bar should feel like it is resting on a solid place, not like it sinks far into the head. Holding the right side of the head steady, slide the left ear bar into the rat's left horizontal ear canal. It should feel like it is resting on a solid place. The pinnae should look symmetrical and should appear to lie flat on the ear bars. Upward flaring of pinnae indicates improper placement of the ear bars. The head should not wobble in response to pressure at the neck.</li>
	<li>Screw down the nose bar. Be gentle. The bones in the nose are delicate and the nose bar does not need to be tight. Check that the rat is still anesthetized before proceeding.</li>
</ol>
3. Preparing the Rat for Surgery
<ol>
	<li>Using a sterile swab, gently place a small amount of veterinary ophthalmic lubricating ointment into each of the rat's eyes to protect them from drying and debris.</li>
	<li>Concentrically from the planned incision site to the edge of the shaved region, wipe the rat's scalp with 2% chlorhexidine solution followed 70% ethanol three times. Chlorhexidine scrub may also be used if more effective disinfectant is desired. Finish by wiping with 10% povidone iodine solution. The prepared scalp is now sterile and should only be touched with sterile materials.</li>
	<li>Don sterile gloves for further steps because you will next deal with the prepared surgical site.</li>
	<li>Place a sterile drape so that the fenestration exposes the surgical site.</li>
</ol>
4. Creating the Surgical Window
<ol>
	<li>Using a #10 scalpel blade, make an incision (approximately 2 cm in length) along the longitudinal midline of the scalp. The incision should start posterior to the line of the eyes. With the thumb and forefinger of the non-dominant hand, hold the scalp in tension perpendicular to the direction of the incision.</li>
	<li>Pull the fascia overlying the skull surface to the edges of the surgical site using sterile cotton-tips. It will take some force to clear the skull so it can be viewed.</li>
	<li>Use four curved, sterile hemostats to create a surgical window. Clamp fascia (NOT skin) on each side of the anterior and posterior aspects of the incision and then lay the clamps alongside the animal, thereby pulling the incision open and fully exposing the skull.</li>
</ol>
5. Leveling the Skull and Creating the Pilot Hole
<ol>
	<li>Locate bregma, the anterior point where the three main plates of the skull meet, as depicted in Figure 2. Move the infusion needle so the needle tip just touches the skull at bregma. Note the dorsal-ventral coordinate. Figure 3 demonstrates how to read coordinates on a Stoelting stereotaxic device. Note, at this point, gloves are no longer sterile and should not touch the sterile surgery site directly. Note: After touching the nonsterile stereotaxic device, the surgeon is no longer sterile and should not touch the surgical site nor parts of equipment which will touch the surgical site. </li>
	<li>Locate lambda, the posterior point where the three main plates of the skull meet, as depicted in Figure 2. Raise the infusion needle slightly and move it straight back to lambda. Lower the needle until it just touches the skull at lambda. Read the dorsal-ventral coordinate. If the skull is level, it will be within 0.1 mm of the bregma reading. If it is further off than this, move the needle up (out of the way), loosen the nose bar, either raise or lower the bite bar platform, depending on which end of the head is too high, and then re-measure.</li>
	<li>Move the infusion needle over bregma and record the anterior-posterior and medial-lateral coordinates. Calculate where the needle must be moved based on the coordinates of the brain area of interest according to a brain atlas12. For the BLA, subtract 2.8 mm from the anterior-posterior coordinate and either add or subtract 5.1 mm to/from the medial-lateral coordinate depending on whether you wish to target the left or right. Move the infusion needle to the calculated position.</li>
	<li>Swing the needle out of the way, taking care to keep track of where it was located and then drill a hole approximately 2 mm in diameter in that location using a Dremel moto-tool with a sterile dental drill bit attached. Drill carefully as when the drill breaks through the skull, it will tend to pull down, possibly damaging the dura and brain tissue underneath. Ream the hole to enlarge it.</li>
	<li>Absorb any blood with a sterile cotton swab.</li>
</ol>
6. Infusion of Neurotoxin
<ol>
	<li>Swing the needle back into position. Lower the needle until it touches the surface of the brain. Record the dorsal-ventral coordinate. This is the level of dura. Subtract the appropriate value to reach the brain region of interest. For the basolateral amygdala, subtract 6.7 mm.</li>
	<li>Lower the infusion needle to the calculated level slowly. Then raise it back up, well above the skull. Use a Hamilton syringe cleaning wire to unclog the tip of the infusion needle, being careful not to touch the end of the wire that comes in contact with the infusion needle with non-sterile gloves.</li>
	<li>Run the pump at &gt;5 μl/min and make sure you see a bead of liquid come out of the infusion needle. If not, unclog with the wire and try again.</li>
	<li>Lower the infusion needle to the desired dorsal/ventral level slowly.</li>
	<li>Set the rate on the infusion pump to 0.1 μl/min and start the infusion. Infuse for 2 min.</li>
	<li>Once the infusion has started, mark the two ends of the air bubble in the tubing with a black marker. If both ends do not move, your needle is clogged. Stop the infusion, raise the needle, unclog with the wire, check for flow with a high flow rate, re-lower the needle and try again.</li>
	<li>Once the infusion is complete, stop the pump and wait 5 min for the NMDA to diffuse away from the needle tip.</li>
	<li>Raise the needle 0.5 mm, and infuse again for 1 min. Wait 2.5 min for the NMDA to diffuse away from the needle-tip. This extra infusion helps ensure all of the BLA is covered by the infusion. Whether this is necessary for other brain regions must determined experimentally.</li>
	<li>Raise the needle out of the brain slowly and swing the stereotaxic arm out of the way.</li>
</ol>
7. Closing the Incision
<ol>
	<li>Don new sterile gloves to close the incision.</li>
	<li>Wipe away any excess blood on the skull with sterile cotton-tips and remove the clamps from the fascia.</li>
	<li>Using London forceps, push the two sides of the incision together. The inner edges of the skin should meet and the outer edge of the skin should not be permitted to curl into the incision. While pressing the incision closed, apply wound clips to close the protrusion firmly. Continue to push the skin together and staple along the length of the incision. This usually takes 3-5 wound clips.</li>
	<li>After placing all the wound clips, use the forceps to pinch the staples again, making sure they are secure. The more secure the clips, the less likely a rat will pull them out before the incision has healed.</li>
	<li>Swab the stapled incision with 10% povidone iodine solution and return the rat to its home cage and monitor until mobility is restored (usually 1-2 hours after anesthesia, total surgery time from induction to closure being 30-45 min). At this point, the rat may be returned to the colony room. After one week, remove staples with a staple remover tool under isoflurane anesthesia.</li>
	<li>Use a hot bead sterilizer to sterilize the tip surfaces of all instruments before use on the next rat. Instruments should be immersed in the beads for no longer than 15 seconds, or they may become too hot to handle. Instruments should be fully cooled before used on another animal.</li>
</ol>
8. Recovery
<ol>
	<li>It is possible that rats will exhibit seizure activity in the days following the lesion. Depending on the brain region damaged, other behaviors such as food consumption or grooming may be affected. It is important that animals be monitored on a daily basis to ensure that they remain healthy during recovery. Supportive care, such as liquid food, high-calorie food, or additional analgesia may be required. Mortality due to anesthesia occurs in a low percentage of rats, usually less than 5%. Mortality due to other causes (infection, bleeding) is extremely rare. Any signs of distress after recovery of anesthesia may be cause for exclusion from future studies, but are extremely rare. Figure 4 depicts the weight loss during the first few days following surgery as it should be expected. Body weight should rebound within 1 week.</li>
</ol>
9. Representative Results
One week or more after NMDA infusion, the lesion can be visualized using a cresyl-violet counterstain and brightfield microscopy. Brains should be perfused with ice cold 4% paraformaldehyde, equilibrated in 30% sucrose in 0.1M PBS and frozen. They should then be sliced at 30-60 μm, in a series of 1:6 to 1:12 and mounted on to gelatin-coated glass slides before undergoing a standard cresyl violet counterstain. The lesion should correspond to a "bald spot" or an area of decreased cresyl violet stain due to cell death following lesion. Figures 5A and B show a representative lesion of the BLA and CeA. This image was obtained using a flatbed scanner with a high resolution scan (1200 dpi). To visualize the lesion, it helps to counterstain a series of sections through much of the tissue surrounding the target area so a misplaced lesion can be located.
<img alt="Stereotaxic surgery setup diagram with labeled components for precise animal brain positioning." src="/files/ftp_upload/4079/4079fig1.jpg" data-altupdatedat="1787630408000" data-alt="Figure 1"> 
Figure 1. Stereotaxic setup. This figure shows the rat stereotaxic with relevant parts labeled.
<img alt="Static equilibrium diagram; illustrating suture intersection points: Bregma and Lambda; labeled setup." src="/files/ftp_upload/4079/4079fig2.jpg" data-altupdatedat="1787630408000" data-alt="Figure 2"> 
Figure 2. Bregma and lambda on the rat skull. This illustration depicts the locations of bregma and lambda. All coordinates are measured relative to bregma. Lambda is used to ensure that the skull is level.
<img alt="Vernier scale diagram, 10.9 mm measurement, decimal point alignment, metric ruler markings." src="/files/ftp_upload/4079/4079fig3.jpg" data-altupdatedat="1787630408000" data-alt="Figure 3"> 
Figure 3. Reading stereotaxic coordinates. This drawing depicts a standard stereotaxic reading. To obtain the digits to the left of the decimal place, use the markers on the right side. The labeled digit corresponds to the tens place (i.e. 1 = 10, 2 = 20). The hash marks between the labeled digits correspond to the single units (1-9). The zero line indicates at what integer the stereotax is set. Here, the zero line is between the 10 and 11 markers, indicating that the reading is between 10 and 11. To determine the numbers to the right of the decimal place, use the markers on the left side. The hash marks on the left are numbered 0 to 10, with each unlabeled hash representing one unit difference from those next to it. Whichever of the hash marks on the left lines up best with the hash marks on the right indicates the first decimal place of the coordinate reading. Here, the hash mark on the left corresponding to number 9 lines up best with the right-side hash marks so the decimal place is 9. The final reading is 10.9 mm.
<img alt="Weight comparison graph: surgical vs. non-surgical over 9 days; data on weight gain trends." src="/files/ftp_upload/4079/4079fig4.jpg" data-altupdatedat="1787630408000" data-alt="Figure 4"> 
Figure 4. Weight change after surgery. This graph shows the average (± SEM) weight of rats who underwent stereotaxic surgery (n = 17) or did not have surgery (n = 6). Day 0 is the day of surgery. Following surgery, the weight of rats decreased for 2-3 days after surgery and then increased around day 5-6.
<img alt="Histology, brain cross-sections; stain analysis; morphology comparison; cerebral cortex detail." src="/files/ftp_upload/4079/4079fig5.jpg" data-altupdatedat="1787630408000" data-alt="Figure 5"> 
Figure 5. Excitotoxic lesion example. A) This image shows a cresyl violet counterstain on a 30 μm coronal slice with the "bald spot" from NMDA lesion of the basolateral amygdala. The white arrows outline the lesion. It can be compared to the contralateral side of the brain slice which has no lesion. B) A similar lesion of the central nucleus amygdala.

<ol><li>Kirby, E. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Basolateral+amygdala+regulation+of+adult+hippocampal+neurogenesis+and+fear-related+activation+of+newborn+neurons%5BTitle%5D)+AND+%22Mol.+Psychiatry%22%5BJournal%5D)">Basolateral amygdala regulation of adult hippocampal neurogenesis and fear-related activation of newborn neurons.</a> Mol. Psychiatry. , Forthcoming (2011).</li>
<li>Goosens, K. A., Maren, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Contextual+and+auditory+fear+conditioning+are+mediated+by+the+lateral%2C+basal%2C+and+central+amygdaloid+nuclei+in+rats%5BTitle%5D)+AND+%22Learn+Mem%22%5BJournal%5D)">Contextual and auditory fear conditioning are mediated by the lateral, basal, and central amygdaloid nuclei in rats.</a> Learn Mem. 8, 148-155 (2001).</li>
<li>Korz, V., Frey, J. U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Bidirectional+modulation+of+hippocampal+long-term+potentiation+under+stress+and+no-stress+conditions+in+basolateral+amygdala-lesioned+and+intact+rats%5BTitle%5D)+AND+%22J.+Neurosci%22%5BJournal%5D)">Bidirectional modulation of hippocampal long-term potentiation under stress and no-stress conditions in basolateral amygdala-lesioned and intact rats.</a> J. Neurosci. 25, 7393-7400 (2005).</li>
<li>Bhatnagar, S., Vining, C., Denski, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Regulation+of+chronic+stress-induced+changes+in+hypothalamic-pituitary-adrenal+activity+by+basolateral+amygdala%5BTitle%5D)+AND+%22Ann.+N.Y.+Acad.+Sci%22%5BJournal%5D)">Regulation of chronic stress-induced changes in hypothalamic-pituitary-adrenal activity by basolateral amygdala.</a> Ann. N.Y. Acad. Sci. 1032, 315-319 (2004).</li>
<li>Ikegaya, Y., Saito, H., Abe, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Attenuated+hippocampal+long-term+potentiation+in+basolateral+amygdala-lesioned+rats%5BTitle%5D)+AND+%22Brain+Res%22%5BJournal%5D)">Attenuated hippocampal long-term potentiation in basolateral amygdala-lesioned rats.</a> Brain Res. 656, 157-164 (1994).</li>
<li>Ikegaya, Y., Saito, H., Abe, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+basomedial+and+basolateral+amygdaloid+nuclei+contribute+to+the+induction+of+long-term+potentiation+in+the+dentate+gyrus+in+vivo%5BTitle%5D)+AND+%22Eur.+J.+Neurosci%22%5BJournal%5D)">The basomedial and basolateral amygdaloid nuclei contribute to the induction of long-term potentiation in the dentate gyrus in vivo.</a> Eur. J. Neurosci. 8, 1833-1839 (1996).</li>
<li>Carter, R. N., Pinnock, S. B., Herbert, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Does+the+amygdala+modulate+adaptation+to+repeated+stress%5BTitle%5D)+AND+%22Neurosciene%22%5BJournal%5D)">Does the amygdala modulate adaptation to repeated stress.</a> Neurosciene. 126, 9-19 (2004).</li>
<li>Roozendaal, B., McGaugh, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Basolateral+amygdala+lesions+block+the+memory-enhancing+effects+of+glucocorticoid+administration+in+the+dorsal+hippocampus+of+rats%5BTitle%5D)+AND+%22Eur.+J.+Neurosci%22%5BJournal%5D)">Basolateral amygdala lesions block the memory-enhancing effects of glucocorticoid administration in the dorsal hippocampus of rats.</a> Eur. J. Neurosci. 9, 76-83 (1997).</li>
<li>Roesler, R., Roozendaal, B., McGaugh, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Basolateral+amgdala+lesions+block+the+memory-enhancing+effect+of+8-Br-cAMP+infused+into+the+entorhinal+cortex+of+rats+after+training%5BTitle%5D)+AND+%22Eur.+J.+Neurosci%22%5BJournal%5D)">Basolateral amgdala lesions block the memory-enhancing effect of 8-Br-cAMP infused into the entorhinal cortex of rats after training.</a> Eur. J. Neurosci. 15, 905-910 (2002).</li>
<li>Roozendaal, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Glucocorticoid+effects+on+memory+consolidation+depend+on+functional+interactions+between+the+medial+prefrontal+cortex+and+basolateral+amygdala%5BTitle%5D)+AND+%22J.+Neurosci%22%5BJournal%5D)">Glucocorticoid effects on memory consolidation depend on functional interactions between the medial prefrontal cortex and basolateral amygdala.</a> J. Neurosci. 29, 14299-14308 (2009).</li>
<li>Kaufer, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Restructuring+the+neuronal+stress+response+with+anti-glucocorticoid+gene+delivery%5BTitle%5D)+AND+%22Nat.+Neurosci%22%5BJournal%5D)">Restructuring the neuronal stress response with anti-glucocorticoid gene delivery.</a> Nat. Neurosci. 7, 947-953 (2004).</li>
<li>Paxinos, G., Watson, C. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aPaxinos%2C+G+%22The+rat+brain%3A+in+stereotaxic+coordinates-the+new+coronal+set%22">The rat brain: in stereotaxic coordinates-the new coronal set</a>. , Elsevier Academic Press. Burlington, MA. (2004).</li>
<li>Geiger, B. M., Frank, L. E., Caldera-Siu, A. D., Pothos, E. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Survivable+Stereotaxic+Surgery+in+Rodents%5BTitle%5D)+AND+%22J.+Vis.+Exp%22%5BJournal%5D)">Survivable Stereotaxic Surgery in Rodents.</a> J. Vis. Exp. (20), e880(2008).</li>
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No conflicts of interest declared.

The stereotaxic method presented here allows for excitotoxic lesion of specific brain areas via infusion of NMDA. The basic stereotaxic methods can be adapted to infuse a variety of pharmacological and biological agents in a site-specific manner. It can also be adapted to target a variety of brain areas, defined by their stereotaxic coordinates in a brain atlas12. Adaptation to other species such as mice can be made with similar equipment built for smaller animals. The present procedures are optimized for 3 mo...

<table><tbody><tr><td>N-Methyl-D-aspartic Acid 98%</td><td>Fisher Scientific</td><td>AC32919-0500</td><td></td></tr><tr><td>Dual Lab Standard Stereotaxic w/45 deg. Ear Bars</td><td>Stoelting</td><td>51653</td><td>Alternative vendor: Kopf *note of caution: assure compatibility of stereotaxic accessories if purchasing from multiple vendors</td></tr><tr><td>10&amp;mu;l SYR SPECIAL (*/*/*)</td><td>Hamilton</td><td>701SN</td><td></td></tr><tr><td>Dremel Moto-Tool</td><td>Stoelting</td><td>58600</td><td>Alternative vendor: Kopf</td></tr><tr><td>Carbide burs, handpiece HP, size 2</td><td>Schein Dental</td><td>2284578</td><td></td></tr><tr><td>St&amp;#8211;lting 6 Syringe Programmable Pump</td><td>Stoelting</td><td>53140</td><td>Alternative vendor: Kopf</td></tr><tr><td>Stainless Steel 316 Hypodermic Regular Wall Tubing 30 Gauge .0123&quot; OD x .00625&quot; ID x .003&quot; Wall (infusion needle)</td><td>Small Parts</td><td>HTXX-30R-06-05</td><td></td></tr><tr><td>Intramedic PE 20 tubing (infusion tubing)</td><td>VWR</td><td>63019-025</td><td></td></tr><tr><td>Reflex Clips, 9mm, non-sterile</td><td>Kent Scientific Corp.</td><td>INS500346</td><td>Alternative vendor: Fine Science Tools</td></tr><tr><td>Reflex Clip Applier for 9mm clips</td><td>Kent Scientific Corp.</td><td>12031-09</td><td>Alternative vendor: Fine Science Tools</td></tr><tr><td>Curved Hartman hemostat</td><td>Fine Science Tools</td><td>13003-10</td><td></td></tr><tr><td>London forceps</td><td>Fine Sceince Tools</td><td>11080-02</td><td></td></tr><tr><td>2% chlorhexidine solution</td><td>Allivet</td><td>30159</td><td>Alternative vendor: PetSolutions</td></tr><tr><td>10% povidone iodine solution</td><td>CVS</td><td>SKU #739575</td><td></td></tr><tr><td>Hot bead sterilizer</td><td>Harvard Apparatus</td><td>610183</td><td></td></tr><tr><td colspan="4">&lt;p class=&quot;jove_content&quot;&gt;&lt;strong&gt;Table 1.&lt;/strong&gt; Table of specific reagents and equipment.&lt;/p&gt;</td></tr></tbody></table>

stereotaxic surgery for excitotoxic lesion of specific brain areas in the adult rat

Stereotaxic Surgery for Excitotoxic Lesion of Specific Brain Areas in the Adult Rat

Many behavioral functions in mammals, including rodents and humans, are mediated principally by discrete brain regions. A common method for discerning the function of various brain regions for behavior or other experimental outcomes is to implement a localized ablation of function. In humans, patient populations with localized brain lesions are often studied for deficits, in hopes of revealing the underlying function of the damaged area. In rodents, one can experimentally induce lesions of specific brain regions.
Lesion can be accomplished in several ways. Electrolytic lesions can cause localized damage but will damage a variety of cell types as well as traversing fibers from other brain regions that happen to be near the lesion site. Inducible genetic techniques using cell-type specific promoters may also enable site-specific targeting. These techniques are complex and not always practical depending on the target brain area. Excitotoxic lesion using stereotaxic surgery, by contrast, is one of the most reliable and practical methods of lesioning excitatory neurons without damaging local glial cells or traversing fibers.
Here, we present a protocol for stereotaxic infusion of the excitotoxin, N-methyl-D-aspartate (NMDA), into the basolateral amygdala complex. Using anatomical indications, we apply stereotaxic coordinates to determine the location of our target brain region and lower an injection needle in place just above the target. We then infuse our excitotoxin into the brain, resulting in excitotoxic death of nearby neurons. While our experimental subject of choice is a rat, the same methods can be applied to other mammals, with the appropriate adjustments in equipment and coordinates.
This method can be used on a variety of brain regions, including the basolateral amygdala1-6, other amygdala nuclei6, 7, hippocampus8, entorhinal cortex9 and prefrontal cortex10. It can also be used to infuse biological compounds such as viral vectors1, 11. The basic stereotaxic technique could also be adapted for implantation of more permanent osmotic pumps, allowing more prolonged exposure to a compound of interest.

Targeted ablation of specific brain region(s) by infusion of an excitotoxin using stereotaxic coordinates is described. This technique could also be adapted for infusion of other chemicals into the rat brain.

Many behavioral functions in mammals, including rodents and humans, are mediated principally by discrete brain regions. A common method for discerning the ...

The overall aim of this procedure is to lesion a specific area of the rat brain using stereotaxic infusion of an excitor toxin. This is accomplished by first properly mounting a rat in the stereotaxic device. The second step is to locate the infusion position based on stereotaxic coordinates.Next, the excitotoxin is infused and allowed to diffuse away from the needle tip. The final step is to remove the needle and close the incision. Ultimately, immunohistochemical counters staining of fixed tissue slices are used to show effective and correctly located lesion.This method can help answer key questions in neuroscience field, such as functions of discrete brain areas. Visual demonstration of this method is critical because to properly mount the head so that it is level and secure requires that you know what proper position looks like. To set up for the experiment, first fill a 10 microliter Hamilton syringe with sterile water.Then mount the filled syringe into the caddy of a six syringe programmable pump, and secure the end of the syringe plunger into the clamped holder on the pump. Insert a previously prepared 30 gauge flat cut one inch infusion needle into approximately two feet of gas sterilized PE 20 tubing. Next, use a 25 gauge needle and one milliliter syringe to prefill the tubing with sterile water.Slide the open end of the tubing onto the Hamilton syringe. Be careful to avoid creating air bubbles in the tube. Now clamp the infusion needle end of the tubing onto the arm of a stereotaxic device using a barrel style electrode manipulator.Then insert the infusion needle between the clamp and the grooved barrel, ensuring that the clamp is secured over a piece of tubing containing the needle within it to avoid flattening the tubing. Next, adjust the pump to a rate greater than five microliters per minute and turn on the pump. Once the pump is running, a bead of water should appear at the tip of the infusion needle.Use this opportunity to check the joints and tubing to ensure that there are no leaks. Use a sterile swab to wipe away the water bead. Then set the pump to withdraw and suck up a two microliter air bubble.The top of the bubble should be visible in the tubing. Now immerse the tip of the infusion needle in a 20 milligram per milliliter solution of NMDA in sterile 0.1 molar PBS and withdraw four to five microliters of the liquid. Ensure that the air bubble separates the sterile water and the NMDA solution.Once a suitable volume of NMDA solution has been drawn up, the receptacle containing the stock solution can be removed and then the setup is ready for surgery. After mounting the rat in the stereotaxic frame, each ear bar should feel like it is resting on a solid place, not like its sinks far into the head. The penny should look symmetrical and should appear to lie flat on the ear bars.Upward flaring of penny indicates improper placement of the ear bars. The head should not wobble in response to pressure at the neck. Use a sterile swab to apply ophthalmic ointment to the eyes.Now apply 2%chlorhexidine solution to the scalp while moving in. Concentric circles from the planned incision site towards the edge of the shaved region. Apply 70%ethanol in the same fashion.Repeat this process three times followed by one application of 10%Povidone iodine solution as the scalp is now sterile, it should only be touched with sterile materials. At this point, don sterile surgical gloves and places sterile drape over the animal so that the fenestration exposes the surgical site. The animal is now ready for surgery after using a scalpel fitted with a number 10 blade to make a two centimeter longitudinal incision along the midline of the scalp beginning posterior to the line of the eyes.Use sterile cotton tips to pull the fascia overlying the skull surface to the edges of the surgical site. Use four curved sterile hemostats to clamp the fascia on each side of the anterior and posterior aspects of the incision. To create a surgical window, lay the other end of the hemostats alongside the animal.To pull the incision open and fully expose the skull. First, locate bgma and lambda as depicted in this diagram. Then move the needle so that the tip just touches the scarlet bgma.Note the dorsal ventral coordinate according to the venia scale. It is a good idea to familiarize oneself with reading the venia scale before beginning the procedure. A guide is provided at the end of this video.After touching the non-sterile stereotaxic device, the surgeon is no longer sterile and should be careful not to touch the sterile surgical site or the parts of any tools that will touch the surgical site. Next, slightly raise the needle and move the needle tip straight back to lambda again. Lower the needle tip and note the dorsal ventral coordinate.If this coordinate is within 0.1 millimeters of that recorded for broma, then the skull is considered level. If not, then the bite bar will need to be adjusted and the measurements repeated. Once the head is level in the dorsal ventral plane, return the needle torema and record the anterior, posterior and media lateral coordinates.Calculate the target positioning of the needle based on the brain atlas coordinates of the structure of interest and move the needle to the calculated position. Mark the position of the needle and then swing the needle out of the way. Use a Dremel moto tool with a sterile dental drill bit attached to carefully drill a hole of approximately two millimeters at the site of the needle position without damaging the dura or tissue underneath.Ream the hole to enlarge it and when finished, absorb any blood with a cotton swab. After drilling the hole, swing the needle back into position. Lower the needle into the hole until the tip touches the dura.Record this dorsal ventral coordinate and subtract the appropriate valley required to reach the brain region of interest. Slowly lower the needle to the calculated level. Then raise the needle back up to a level way above the skull.Use a Hamilton syringe cleaning wire to unclog the tip of the infusion needle. Now run the pump at the same rate as before. A bead of liquid should form at the end of the infusion needle.If not, repeat the unclogging procedure and then try again. After confirming that the needle is free of blockages, slowly lower the needle back to the desired dorsal ventral level. Mark the two ends of the air bubble with black marker.Set the infusion pump to 0.1 microliter per minute and start the infusion. The movement of the bubble indicates that the needle is not clogged and that the infusion is working. Well allow the neurotoxin to infuse into the brain for two minutes.Then once the infusion is complete, stop the pump and wait five minutes. For the NMDH diffuse away from the needle tip. Depending on the brain region, it may be necessary to raise the needle a small amount and infuse again for a shorter time period.This is done to ensure that the target structure is covered by the infusion. Once the infusion of neurotoxin is complete, raise the needle outta the brain slowly and swing the stereotaxic arm out of the way. After doning a fresh pair of sterile gloves, use sterile cotton tips to wipe away any excess blood on the skull.Then remove the clamps from the facia. Then use London forceps to push the two sides of the incision together, ensuring that the inner edges of the skin meet. Do not allow the skin to curl in towards the incision.Keep pressing the incision closed and apply wound clips along the length of the incision. This usually takes three to five wound clips. After placing the wound clips, use forceps to pinch and further secure the staples.Finally, swap the stapled incision with 10%povidone iodine solution. Return the rat to the home cage. Monitor the recovery of the animal and administer analgesic according to institutional guidelines.This graph shows the average weight of 17 rats that underwent stereotypes, surgery, and six rats that did not have surgery. Day zero is the day of the surgical procedure. Following surgery, the weight of rats decreased for two to three days and then increased around day five to six.This image shows a crestal violet counter stain on a 30 micron coronal slice with the bald spot from an MDA lesion of the basal lateral amygdala. The white arrows outline the lesion. It can be compared to the contralateral side of the brain slice, which has no lesion.This image shows a similar lesion of the central nucleus amygdala. This drawing depicts a standard stereotaxic reading to obtain the digits to the left of the decimal place. Use the markers on the right side.The labeled digit corresponds to the tens place IE one equals 10 and two equals 20. The hash marks between the labeled digits correspond to the single units one to nine. The zero line indicates at what integer the stereotypes is set.Here the zero line is between the 10 and 11 markers indicating that the reading is between 10 and 11. To determine the numbers to the right of the decimal place, use the markers on the left side. The hash marks on the left are numbered zero to 10 with each unlabeled hash mark representing one unit difference from those next to it, whichever of the hash marks on the left lines up best with the hash marks on the right indicates the first decimal place of the coordinate reading here.The hash mark on the left, corresponding to number nine, lines up best with the right side hash marks, so the decimal place is nine. The final reading is 10.9 millimeters. Once mastered, this technique should take 30 minutes if performed properly.Following this procedure. Viral vectors or sir a could be infused site specifically into the brain to answer questions like how changing gene expression in certain brain regions alter brain function.

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Research

JoVE Journal

Neuroscience

The monitoring of extracellular neurotransmitter levels in distinct brain regions of  freely moving animals offers insights on the link between neurotransmitter release and behavior. In vivo microdialysis coupled with electrochemical detection provides excellent anatomical and chemical resolution; and information on how basal neurotransmission is altered by pharmacological or physiological manipulations.

Survivable Stereotaxic Surgery in Rodents 

Source: Batra, V., et al. A General Method for Evaluating Deep Brain Stimulation Effects on Intravenous Methamphetamine Self-Administration. J. Vis. Exp. (2016).This video demonstrates a rat model of drug addiction in which the animal self-administers intravenous methamphetamine by pressing a lever, leading to compulsive drug-seeking behavior driven by dopamine release in the nucleus accumbens. To counteract this, the rat undergoes deep brain stimulation (DBS) via an implanted bipolar...

Deep Brain Stimulation as a Treatment for Drug Addiction in a Rat Model

This video demonstrates electroencephalographic (EEG) recording of epileptic seizures in epilepsy-induced rats. A rat with electrodes implanted in the ventral hippocampus is connected to the EEG recording system and placed in a plexiglass chamber. The rat is allowed to move freely while the brain activity is recorded to monitor the occurrence of seizures.

Electroencephalographic Recording of Epileptic Seizures in Epilepsy-Induced Rats

This video demonstrates a stereotactically-guided surgical procedure for ablating the auditory cortex in a rat. The steps include identifying anatomical landmarks in an anesthetized rat secured in a stereotactic frame, creating a window on the skull above the auditory cortex, aspirating the cortical layers, and providing post-operative care to assess the impact of the ablation on sound processing.

A Stereotactically Guided Ablation Technique for Auditory Cortex Removal in Rats

The mechanism underlying the therapeutic effects of Deep Brain Stimulation (DBS) surgery needs investigation. The methods presented in this manuscript describe an experimental approach to examine the cellular events triggered by DBS by analyzing the gene expression profile of candidate genes that can facilitate neurogenesis post DBS surgery.

Analysis of Gene Expression Changes in the Rat Hippocampus After Deep Brain Stimulation of the Anterior Thalamic Nucleus

A method for implanting electrodes into the subthalamic nucleus (STN) of rats is described. Better localization of the STN was achieved by using a microrecording system. Furthermore, a stimulation set-up is presented that is characterized by long-lasting connections between the head of the animal and the stimulator.

Microelectrode Guided Implantation of Electrodes into the Subthalamic Nucleus of Rats for Long-term Deep Brain Stimulation

We describe a method for the stereotactically-guided location, exposure, and ablation of the auditory cortex in rats. The localization of the ablation is assessed using a coordinate map postmortem.

Stereotactically-guided Ablation of the Rat Auditory Cortex, and Localization of the Lesion in the Brain

We describe a protocol of stereotaxic surgery with a homemade head-fixed device for microinjecting reagents into the striatum of neonatal mouse brains. This technique allows genetic manipulation in neuronal cells of specific regions of neonatal mouse brains.

Stereotaxic Surgery for Genetic Manipulation in Striatal Cells of Neonatal Mouse Brains

This protocol describes the construction of a low-cost microinjection system, its stereotaxic implantation into deep-brain structures and the procedure for timed microinjections of tetrodotoxin in awake and unrestrained rats. The goal is to reveal the participation of hypothalamic structures in the regulation of ovulation by inhibiting their neural activity.

Unraveling the Role of Discrete Areas of the Rat Brain in the Regulation of Ovulation through Reversible Inactivation by Tetrodotoxin Microinjections

We describe a preclinical experimental method to evaluate metabolic neuromodulation induced by acute deep brain stimulation with in vivo FDG-PET. This manuscript includes all experimental steps, from stereotaxic surgery to the application of the stimulation treatment and the acquisition, processing, and analysis of PET images.

In vivo Positron Emission Tomography to Reveal Activity Patterns Induced by Deep Brain Stimulation in Rats

Learn stereotaxic surgery for targeted brain lesioning in adult rats using excitotoxin infusion. This method enables precise ablation of specific neural regions for neuroscience research.

Stereotaxic Surgery for Excitotoxic Lesion of Specific Brain Areas in the Adult Rat

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