This work was funded by the Center for Neural Communication Technology (CNCT), a P41 Resource Center funded by the National Institute of Biomedical Imaging and Bioengineering (NIBIB, P41 EB002030) and supported by the National Institutes of Health (NIH).

1. Prepare Fixative
(See table Fixative and Buffers.)
2. Prepare Perfusion Buffers
(See table Fixative and Buffers.)
3. Prepare Apparatus and Anesthesia
<ol>
 <li>Using the water bath, warm perfusion buffer to 37 &deg;C. Place outlet valve in a beaker filled with buffer. Fill a 50 ml syringe with buffer and attach to fixative tubing. Flush tubing repeatedly by expelling and withdrawing buffer.</li>
 <li>Clear the line with buffer until all air bubbles in the tubing are eliminated. It is crucial for the success of a perfusion not to have air bubbles in any of the lines.</li>
 <li>Remove the syringe and connect the 4% paraformaldehyde fixative (room temperature) container. To avoid air bubbles at the tubing end, squeeze the tube while positioning the tube into the container so that a drop of buffer protrudes from the end to contact the fluid surface within the container.</li>
 <li>Close the outlet port (needle end). Turn buffer valve (blue) to the same position as the fixative valve (white). This will allow flow from the buffer line only.</li>
 <li>Open the outlet port and repeat step A1 through A3 for filling the buffer line. After all the air bubbles are eliminated close the outlet port, remove the syringe and connect the buffer container, taking care not to introduce air bubbles into the tubing.</li>
 <li>Test system for ability to hold pressure by pumping the rubber manometer blub. There is normally some resistance due to the compression of air in the system.</li>
 <li>Setup surgery tools in order for easy of access. Fill perfusion needle with buffer to eliminate the possibility of air bubbles.</li>
 <li>Prior to surgery, a ketamine/xylazine mixture (up to 80 mg/kg body weight ketamine and 10 mg/kg body weight xylazine) is administered via intraperitoneal injection (27 gauge needle and 1 cc syringe). Additional administration of anesthetic will be performed as necessary during the course of each operation to maintain a surgical plane of anesthesia.</li>
</ol>
4. Perfusion Surgery
<ol>
 <li>Once the animal has reached a surgical plane of anesthesia, place it on the shallow tray filled with crushed ice. (Use toe pinch-response method to determine depth of anesthesia. Animal must be unresponsive before continuing).</li>
 <li>Make a 5-6 cm lateral incision through the integument and abdominal wall just beneath the rib cage. Carefully separate the liver from the diaphragm.</li>
 <li>Make a small incision in the diaphragm using the curved, blunt scissors. The position and pressure of your finger can aid in the ability to cut the diaphragm.</li>
 <li>Continue the diaphragm incision along the entire length of the rib cage to expose the pleural cavity.</li>
 <li>Place curved, blunt scissors along one side of the ribs, carefully displacing the lungs, and make a cut through the rib cage up to the collarbone. Make a similar cut on the contralateral side.</li>
 <li>Lifting the sternum away, carefully trim any tissue connecting it to the heart. Clamp the tip of the sternum with the hemostat and place the hemostat over the head. When done properly, the thymus lifts away from the heart along with the sternum, providing a clear view of the major vessels.</li>
 <li>Make a small incision to the posterior end of the left ventricle using iris scissors.</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/3564/3564step7.jpg" /> 
<a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/3564/3564step7large.jpg" target="_blank"> Click here to view larger figure</a>.
<ol start="8">
 <li>Pass a 15-gauge blunt- or olive-tipped perfusion needle through the cut ventricle into the ascending aorta. The tip should be visible through the wall of the aorta, and should not reach the aortic arch where the brachial and carotid arteries diverge.</li>
 <li>Use a hemostat to clamp the heart, this secures the needle and prevents leakage. If desired, the modified hemostat can be used to clamp the aorta around the needle tip (these hemostats remain in place until dissection begins, but are omitted from future illustrations for clarity).</li>
 <li>Finally, make an incision to the animal's right atrium using iris scissors to create as large an outlet as possible without damaging the descending aorta. At this point the animal is ready to be perfused.</li>
</ol>
5. Perfusion
<ol>
 <li>Open and attach outlet port to needle base taking care not to introduce any air bubbles.</li>
 <li>Pump up the manometer bulb to a pressure of 80 mm Hg quickly &amp; evenly. Maintain this pressure throughout the buffer infusion period. Start the timer.</li>
 <li>Adjust the needle angle. The angle of the needle is critical to the achievement of a maximum flow rate (note the flow change with angle adjustment).</li>
 <li>Switch the buffer valve (blue) once buffer is almost finished (200 ml). The fluid should be running clear. The clearing of the liver is an indicator of a good perfusion. The liver should be clear at this point. Indicate time for your records.</li>
 <li>Fixation tremors should be observed within seconds; this should be considered the true time of fixation. Indicate time for your records.</li>
 <li>The pressure can gradually be increase up to a maximum of 130 mm Hg 2 to maintain a steady flow rate.</li>
 <li>Close the outlet valve once the fixative is nearly finished. Indicate ending time for your records.</li>
 <li>The rat should be stiff at this stage.</li>
 <li>The used paraformaldehyde must be collected and stored for disposal according to the regulations of your institution.</li>
</ol>
6. Dissection 4,11
<ol>
 <li>Remove the head using a pair of scissors.</li>
 <li>Make a midline incision along the integument from the neck to the nose and expose the skull.</li>
 <li>Trim off the remaining neck muscle so that the base of the skull is exposed; remove any residual muscle using scissor or rongeurs.</li>
 <li>Place the sharp end of a pair of iris scissors into the foramen magnum on one side, carefully sliding the scissors along the inner surface of the skull.</li>
 <li>Next, make a cut extending to the distal edge of the posterior skull surface. Make an identical cut on the contralateral side. Use the rongeurs to clear away the skull around the cerebellum.</li>
 <li>Carefully slide the scissors along the inner surface of the skull as the tip travels from the dorsal distal posterior corner to the distal frontal edge of the skull, lifting up on the blade as you are cutting to prevent damage to the brain. Repeat for opposite side.</li>
 <li>Using rongeurs peel the dorsal surface of the skull away from the brain. Trim away the sides of the skull using rongeurs as well.</li>
 <li>Using a spatula, sever the olfactory bulbs and nervous connections along the ventral surface of the brain.</li>
 <li>Gently tease the brain away from the head, trimming any dura that still connects the brain to the skull using iris scissors.</li>
 <li>Remove the brain and place it in a vial of fixative containing fluid at least 10x the volume of the brain itself. Swirl the vial occasionally.</li>
</ol>
7. Post-fixation &amp; Storage
<ol>
 <li>Keep the brain in fixative for 24 hours at 4 &deg;C, swirling occasionally.</li>
 <li>After 24 hours, wash the brain with phosphate buffered saline by exchanging the media 3 times and swirling each time.</li>
 <li>Brains can then be stored in phosphate buffered saline or HBHS with sodium azide and kept at 4 &deg;C.</li>
</ol>
8. Representative Results
An initial indicator of the success of the perfusion is the clearing of extremities such as the nose, ears, and paws and internal organs such as the thymus gland and liver (IHC World). Gross inspection of the brain reveals the blood vessel void of blood (white to pale yellow appearance). This will be also true within tissue sections destine for staining and immunohistochemistry. The final indicator of outcome of the perfusion is the condition of the ultrastructure in the tissue.1,6,7,8
<table border="1">
 <tbody>
 <tr>
 <td>Prepare Fixative:</td>
 </tr>
 <tr>
 <td>Prepare 8% Paraformaldehyde stock</td>
 </tr>
 <tr>
 <td>
 <ol>
 <li>Add 40 g Paraformaldehyde to 500 ml dH2O. Heat the solution to 60- 65 &deg;C while stirring (Do not exceed 65 &deg;C, doing so can adversely affect the success of the immunohistochemical procedure).</li>
 <li>To clear the solution, reduce heat and add 2-3 ml of 1.0 M NaOH with a dropper.</li>
 <li>Filter and store at 4 &deg;C for up to 1 month.</li>
 </ol>
 </td>
 </tr>
 <tr>
 <td>Prepare 0.2 M Sodium Phosphate Buffer, pH 7.4</td>
 </tr>
 <tr>
 <td>
 <ol start="4">
 <li>For the sodium phosphate monobasic stock, add 27.8 g NaH2PO4* H2O to 1 L dH2O.</li>
 <li>For the sodium phosphate dibasic stock, add 28.4 g Na2HPO4 to 1 L dH2O.</li>
 <li>Add 810 ml of the monobasic stock to 190 ml of the dibasic stock.</li>
 </ol>
 </td>
 </tr>
 <tr>
 <td>Prepare 4% Paraformaldehyde Fixative</td>
 </tr>
 <tr>
 <td>
 <ol start="7">
 <li>Add equal parts 8% paraformaldehyde stock to 0.2M Sodium Phosphate Buffer</li>
 <li>Note: this fix is best prepared fresh, no more than 72 hours in advance.</li>
 </ol>
 </td>
 </tr>
 <tr>
 <td>Prepare Perfusion and Storage Buffers:</td>
 </tr>
 <tr>
 <td>Prepare Phosphate Buffered Saline, pH 7.4</td>
 </tr>
 <tr>
 <td>
 <ol start="9">
 <li>1 L dH2O</li>
 <li>9 g NaCl</li>
 <li>144 mg KH2PO4</li>
 <li>795 mg Na2HPO4</li>
 <li>Check pH</li>
 </ol>
 </td>
 </tr>
 <tr>
 <td>HEPES Buffered Hanks Solution (HBHS) with Sodium Azide pH 7.4</td>
 </tr>
 <tr>
 <td>
 <ol start="14">
 <li>990 ml dH2O</li>
 <li>7.5 g NaCl,</li>
 <li>0.3 g KCl</li>
 <li>0.06 g K H2PO4</li>
 <li>0.13 g Na2HPO4</li>
 <li>2 g Dextrose/Glucose</li>
 <li>2.4 g 10 mm Hepes</li>
 <li>0.1 g MgCl2 6 parts dH2O</li>
 <li>0.05 g MgSO4 7 parts dH2O</li>
 <li>0.165 g CaCl2 2 parts dH2O</li>
 <li>90 mg NaN3</li>
 <li>Check pH</li>
 </ol>
 </td>
 </tr>
 </tbody>
</table>
Table 1. Preparation of Fixative and Buffers.
<img alt="Figure 1" src="/files/ftp_upload/3564/3564fig1.jpg" /> 
Figure 1. Perfusion Apparatus with labeled components.
<img alt="Figure 2" src="/files/ftp_upload/3564/3564fig2.jpg" /> 
Figure 2. Preparation of the Perfusion Apparatus I. Begin with the fixative line. Flush the tubing of air bubbles and fill with buffer by rapidly expelling and withdrawing buffer into the syringe and through the line. Shut the valve on the needle end off. Place the fixative line into the fixative bottle without introducing air bubbles.
<img alt="Figure 3" src="/files/ftp_upload/3564/3564fig3.jpg" /> 
Figure 3. Preparation of the Perfusion Apparatus II. 1. Open valve on the needle end. 2. Turn buffer valve (blue) to position for flow. 3. Flush the tubing of air bubbles and fill with buffer by rapidly expelling and withdrawing buffer with the syringe. 4. Close valve on the needle end. Place tubing into the buffer bottle. Test pressure to make sure the system is sealed correctly by pumping up the manometer bulb and watching the gauge. The apparatus is now ready for the perfusion procedure.
<img alt="Figure 4" src="/files/ftp_upload/3564/3564fig4.jpg" /> 
Figure 4. Perfusion Apparatus in position for buffer delivery.
<img alt="Figure 5" src="/files/ftp_upload/3564/3564fig5.jpg" /> 
Figure 5. Perfusion Surgery I. a) Make a lateral incision through the integument and abdominal wall. b) Make an incision in the diaphragm and cut across the diaphragm exposing the heart. Make parallel cuts on either side of the ribs up to the collarbone. c) clamp the tip of the sternum with the hemostat and place the hemostat over the head.
<img alt="Figure 6" src="/files/ftp_upload/3564/3564fig6.jpg" /> 
Figure 6. Perfusion Surgery II. a) Pass the perfusion needle through the cut ventricle into the ascending aorta. b) Secure the perfusion needle use one set of hemostats to clamp the heart. A second set of a modified hemostat can also be used to clamp the aorta around the needle tip to prevents leakage. Using iris scissors make a small incision to the posterior end of the left ventricle.
<img alt="Figure 7" src="/files/ftp_upload/3564/3564fig7.jpg" /> 
Figure 7. Perfusion with Buffer Pump the manometer bulb to create pressure in the line. Open up the valve (1) and attach to the needle hub. It is critical to make sure no air bubbles are introduced. Pump up the manometer bulb to a pressure of 80 mm Hg and maintain this pressure throughout the buffer infusion period.
<img alt="Figure 8" src="/files/ftp_upload/3564/3564fig8.jpg" /> 
Figure 8. Perfusion with Fixative Once buffer is almost finished (200 ml) switch the buffer valve (1) to allow for the fixative to flow. The pressure can gradually be increased up to a maximum of 130 mm Hg.
<img alt="Figure 9" src="/files/ftp_upload/3564/3564fig9.jpg" /> 
Figure 9. Dissection I. a) remove the head using a pair of scissor. b) make an incision in the skin along the midline from the neck to the nose and expose the skull. c) trim off the remaining neck muscle so that the base of the skull is exposed. Hold the head so that the large opening in the cranium surface (foramen magnum) is accessible. Carefully insert the sharp end of a pair of iris scissors into the foramen magnum and make a cut. Repeat the same maneuver for the contralateral side. Use the rongeurs to clear away the skull around the cerebellum.
<img alt="Figure 10" src="/files/ftp_upload/3564/3564fig10.jpg" /> 
Figure 10. Dissection II. a) carefully slide the scissors along the inner surface of the skull. Lift up on the tip as you are cutting to prevent damage to the brain. Repeat the same for opposite side. Use rongeurs peel the skull away from the brain. b) illustration with the skull removed and the brain exposed. c) use a spatula to sever the olfactory bulbs and nerve connections at the most anterior location of the brain. Carefully guide the spatula tip along the underside of the brain to sever connections for ease of remove. Remove the brain and place it in a vial of fixative.

<ol>
	<li>Cinar, O., Semiz, O., Can, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+microscopic+survey+on+the+efficiency+of+well-known+routine+chemical+fixatives+on+cryosections.">A microscopic survey on the efficiency of well-known routine chemical fixatives on cryosections.</a> Acta. Histochem. 108, 487-496 (2006).</li><li>Fritz, M., Rinaldi, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Blood+pressure+measurement+with+the+tail-cuff+method+in+Wistar+and+spontaneously+hypertensive+rats:+Influence+of+adrenergic-+and+nitric+oxide-mediated+vasomotion.">Blood pressure measurement with the tail-cuff method in Wistar and spontaneously hypertensive rats: Influence of adrenergic- and nitric oxide-mediated vasomotion.</a> J. Pharmacol. Toxicol. Methods. 58, 215-221 (2008).</li><li>Ikeda, K., Nara, Y., Yamorii, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Indirect+systolic+and+mean+blood+pressure+determination+by+anew+tail+cuff+method+in+spontaneously+hypertensive+rats.">Indirect systolic and mean blood pressure determination by anew tail cuff method in spontaneously hypertensive rats.</a> Laboratory Animals. 25, 26-29 (1991).</li><li>Jacobowitz, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Removal+of+discrete+fresh+regions+of+rat+brain.">Removal of discrete fresh regions of rat brain.</a> Brain Res. 80, 111-115 (1974).</li><li>Jonkers, B. W., Sterk, J. C., Wouterlood, F. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transcardial+perfusion+fixation+of+the+CNS+by+means+of+a+compressed-air-driven+device.">Transcardial perfusion fixation of the CNS by means of a compressed-air-driven device.</a> Journal of Neurosci. Meth. 12, 141-149 (1984).</li><li>Jung-Hwa, T. a. o. -. C. h. e. n. g., Gallant, J., Brightman, P. E., Dosemeci, M. W., A, ., Reese, T. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structural+changes+at+the+synapse+after+delayed+perfusion+fixation+in+different+regions+of+the+mouse+brain.">Structural changes at the synapse after delayed perfusion fixation in different regions of the mouse brain.</a> J. Comp. Neurol. 501, 731-740 (2007).</li><li>Kasukurthi, R., Brenner, M. J., Morre, A. M., Moradzadeh, A., Wilson, Z. R., Santosa, K. B., Mackinnon, S. E., Hunter, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transcardial+perfusion+versus+immersion+fixation+for+assessment+of+peripheral+nerve+regeneration.">Transcardial perfusion versus immersion fixation for assessment of peripheral nerve regeneration.</a> J. Neurosci. Meth. 184, 303-309 (2009).</li><li>Lamberts, R., Goldsmith, P. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fixation,+fine+structure,+and+immunostaining+for+neuropeptides:+perfusion+versus+immersion+of+the+neuroendocrine+hypothalamus.">Fixation, fine structure, and immunostaining for neuropeptides: perfusion versus immersion of the neuroendocrine hypothalamus.</a> J. Histochem. Cytochem. 34, 389-398 (1986).</li><li>Ramos-Vara, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Technical+Aspects+of+Immunohistochemistry.">Technical Aspects of Immunohistochemistry.</a> Vet. Pathol. 42, 405-426 (2005).</li><li>Shi, Z. R., Itzkowitz, S. H., Kim, Y. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+comparison+of+three+immunoperoxidase+techniques+for+antigen+detection+in+colorectal+carcinoma+tissues.">A comparison of three immunoperoxidase techniques for antigen detection in colorectal carcinoma tissues.</a> J. Histochem. Cytochem. 36, 317-322 (1988).</li><li>Walker, W. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Vertebrate dissection. , (1980).</li><li>Zwienenberg, M., Gong, Q., Lee, L. L., Berman, R. F., Lyeth, B. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=J.+Neurotrauma.">J. Neurotrauma.</a> Monitoring in the Rat: Comparison of Monitoring in the Ventricle, Brain Parenchyma, and Cisterna Magna. 16, 1095-1102 (1999).</li></ol>

The authors have nothing to disclose.

Additional notes for a successful surgery: 
<ol>
<li>	Once the diaphragm is breached, precede rapidly as hypoxia and hypercapnia will initiate irreversible physiological changes that may confound subsequent analysis.</li>
<li>	When the needle is inserted and clamped in place, the residual pressure will push blood to the base of the needle (flashback). In fact, larger animals may vigorously expel blood. It is critical that blood at least reach the base of the needle to avoid introducing air bubbles that will present an obstacle to complete perfusion. If no blood is observed at the needle base, remove the needle and refill with buffer by attaching it to the outlet port of the perfusion rig. Once buffer has been run through the tip, the needle can be reinserted into the heart.</li>
<li>	Proper placement of the needle within the aorta is critical, it should not reach as far as the aortic arch.</li>
<li>	If multiple perfusions are taking place it is not necessary to setup from the beginning every time. The user needs to have the proper volume for both the buffer and fixative in each bottle (200mls animal/solution). The bottles can also be refilled if needed. The most important aspect to consider is flushing out the perfusion line that runs from the setup to the animal (output valve). After the completion of a perfusion, this line will have 4% paraformaldehyde in it. Remove the tubing from the buffer bottle and use a 50 ml syringe filled with water to flush out the fixative. Make sure the outlet valve is placed in a waste collection beaker for proper disposal of the paraformaldehyde. Repeat this three times. Refill with buffer using the same method shown in 2.3.</li>
<li>	This method for fixation is highly adaptable with the ability to change both buffer and fixative other fixatives (such as those used for EM) according to the requirements of the experiment.</li>
<li>	The system is also frequently used for the extraction of live tissue. For this process one simply uses the delivery of buffer only (the buffer bottle while still attached to entire system is placed in an ice bucket and the fixative bottle is setup as empty but sealed). This assures a cold buffer perfusion at the optimal pressure. From here the system can be adapted to the infusion of fixable dyes by either adding the dye directly to the buffer (if the quantity isn't an issue) or use a syringe with a minimal amount of buffer/dye solution in between the buffer &#x2013; fixative step. </li>
<li>	In the case of vascular casting, the pressure monitoring buffer step is crucial however due to the viscosity and solidifying nature of the casting solution either a 50ml syringe must be used for direct delivery or a disposable secondary attachment to the apparatus (tubing, valve and holding container) made to interface with the current system be used. Our laboratories' vascular casting preceded the design and manufacturing of the perfusion apparatus. Therefore we cannot state that the pressure measurements will be an accurate measurement at the delivery site. The numbers may have to be adjusted to achieve successful delivery of viscous fluids.</li>
</ol>

<table border="1">
 <tbody>
 <tr>
 <td>Equipment</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments </td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Anesthetic:</td>
 </tr>
 <tr>
 <td>Ketamine/xylazine mixture (Anesthetic may vary per laboratory / institution)</td>
 <td>Ketaset</td>
 <td>NDC 0856-2013-01</td>
 <td>10 mL vial</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Surgery:</td>
 </tr>
 <tr>
 <td>Needle tip, 27 GA x 1.25"</td>
 <td>Materiel Services</td>
 <td>25251</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Large blunt/blunt curved scissors (~14.5 cm)</td>
 <td>Fine Science Tools</td>
 <td>14519-14</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Straight iris scissors</td>
 <td>Fine Science Tools</td>
 <td>14058-11</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Standard tweezers</td>
 <td>Fine Science Tools</td>
 <td>11027-12</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Pair of fine (Graefe) tweezers</td>
 <td>Fine Science Tools</td>
 <td>11050-10</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>1 large hemostat forceps - curved or straight (~19 cm)</td>
 <td>surgicaltools.com</td>
 <td>17.21.51</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>2 standard hemostat forceps - straight serrated (14 cm)</td>
 <td>Fine Science Tools</td>
 <td>13013-14</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>1 modified hemostat (with 15-gauge hole filed through the tip)</td>
 <td>Fine Science Tools</td>
 <td>13013-14</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>15-gauge blunt or olive-tipped needle (perfusion needle)</td>
 <td>Fisnar</td>
 <td>5601137</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Perfusion:</td>
 </tr>
 <tr>
 <td>HyPerfusion system or equivalent</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Phosphate buffered saline, pH 7.4</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>4% Paraformaldehyde in 0.1 M Phosphate Buffer, pH 7.4</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Shallow glass or plastic tray, approximately 10" x 10"</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Crushed ice</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Water bath (37 &deg;C)</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Timer</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>50 ml syringe</td>
 <td>Medline Industries</td>
 <td>NPMJD50LZ</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Dissection:</td>
 </tr>
 <tr>
 <td>Pair of standard sharp/blunt straight scissors (~12 cm)</td>
 <td>Fine Science Tools</td>
 <td>14054-13</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Medium curved or straight rongeurs (14-16 cm) or skull bone removal pliers</td>
 <td>Fine Science Tools</td>
 <td>16020-14</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Straight iris scissors (~9 cm)</td>
 <td>Fine Science Tools</td>
 <td>14058-11</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Micro-spatula (double 2" flat ends, one rounded, one tapered to 1/8")</td>
 <td>Fine Science Tools</td>
 <td>10091-12</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Post-Fixation &amp; Storage:</td>
 </tr>
 <tr>
 <td>50 ml glass vial</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>40 ml HEPES-Buffered Hanks Solution (HBHS) with sodium azide (90mg/l)</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>

whole animal perfusion fixation for rodents

The goal of fixation is to rapidly and uniformly preserve tissue in a life-like state. While placing tissue directly in fixative works well for small pieces of tissue, larger specimens like the intact brain pose a problem for immersion fixation because the fixative does not reach all regions of the tissue at the same rate 5,7. Often, changes in response to hypoxia begin before the tissue can be preserved 12. The advantage of directly perfusing fixative through the circulatory system is that the chemical can quickly reach every corner of the organism using the natural vascular network. In order to utilize the circulatory system most effectively, care must be taken to match physiological pressures 3. It is important to note that physiological pressures are dependent on the species used. Techniques for perfusion fixation vary depending on the tissue to be fixed and how the tissue will be processed following fixation. In this video, we describe a low-cost, rapid, controlled and uniform fixation procedure using 4% paraformaldehyde perfused via the vascular system: through the heart of the rat to obtain the best possible preservation of the brain for immunohistochemistry. The main advantage of this technique (vs. gravity-fed systems) is that the circulatory system is utilized most effectively.

Watch this Scientific Journal Video about Whole Animal Perfusion Fixation for Rodents at JoVE.com

Whole Animal Perfusion Fixation for Rodents

Here we describe a low-cost, rapid, controlled and uniform fixation procedure using 4% paraformaldehyde perfused via the vascular system: through the heart of the rat to obtain the best possible preservation of the brain.

The goal of fixation is to rapidly and uniformly preserve tissue in a life-like state. While placing tissue directly in fixative works well for small ...

whole-animal-perfusion-fixation-for-rodents

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

380.2K Views.  University of Michigan. Here we describe a low-cost, rapid, controlled and uniform fixation procedure using 4% paraformaldehyde perfused via the vascular system: through the heart of the rat to obtain the best possible preservation of the brain.

Video: Whole Animal Perfusion Fixation for Rodents

Automated Interactive Video Playback for Studies of Animal Communication

Video playback is a widely-used technique for the controlled manipulation and presentation of visual signals in animal communication. In particular, parameter-based computer animation offers the opportunity to independently manipulate any number of behavioral, morphological, or spectral characteristics in the context of realistic, moving images of animals on screen. A major limitation of conventional playback, however, is that the visual stimulus lacks the ability to interact with the live animal. Borrowing from video-game technology, we have created an automated, interactive system for video playback that controls animations in response to real-time signals from a video tracking system. We demonstrated this method by conducting mate-choice trials on female swordtail fish, Xiphophorus birchmanni. Females were given a simultaneous choice between a courting male conspecific and a courting male heterospecific (X. malinche) on opposite sides of an aquarium. The virtual male stimulus was programmed to track the horizontal position of the female, as courting males do in the wild. Mate-choice trials on wild-caught X. birchmanni females were used to validate the prototype's ability to effectively generate a realistic visual stimulus.

Video playback is a widely used technique in animal behavior. We created and evaluated a program that applies rules-based, interactive playback of 3-D computer animations in response to real-time, automated data on subject behavior.

Video playback is a widely-used technique for the controlled manipulation and presentation of visual signals in animal communication. In particular, ...

Isolating Nasal Olfactory Stem Cells from Rodents or Humans

The olfactory mucosa, located in the nasal cavity, is in charge of detecting odours. It is also the only nervous tissue that is exposed to the external environment and easily accessible in every living individual. As a result, this tissue is unique for anyone aiming to identify molecular anomalies in the pathological brain or isolate adult stem cells for cell therapy. 
Molecular abnormalities in brain diseases are often studied using nervous tissue samples collected post-mortem. However, this material has numerous limitations. In contrast, the olfactory mucosa is readily accessible and can be biopsied safely without any loss of sense of smell1. Accordingly, the olfactory mucosa provides an "open window" in the adult human through which one can study developmental (e.g. autism, schizophrenia)2-4 or neurodegenerative (e.g. Parkinson, Alzheimer) diseases4,5. Olfactory mucosa can be used for either comparative molecular studies4,6 or in vitro experiments on neurogenesis3,7.
The olfactory epithelium is also a nervous tissue that produces new neurons every day to replace those that are damaged by pollution, bacterial of viral infections. This permanent neurogenesis is sustained by progenitors but also stem cells residing within both compartments of the mucosa, namely the neuroepithelium and the underlying lamina propria8-10. We recently developed a method to purify the adult stem cells located in the lamina propria and, after having demonstrated that they are closely related to bone marrow mesenchymal stem cells (BM-MSC), we named them olfactory ecto-mesenchymal stem cells (OE-MSC)11.
Interestingly, when compared to BM-MSCs, OE-MSCs display a high proliferation rate, an elevated clonogenicity and an inclination to differentiate into neural cells. We took advantage of these characteristics to perform studies dedicated to unveil new candidate genes in schizophrenia and Parkinson's disease4. We and others have also shown that OE-MSCs are promising candidates for cell therapy, after a spinal cord trauma12,13, a cochlear damage14 or in an animal models of Parkinson's disease15 or amnesia16.
In this study, we present methods to biopsy olfactory mucosa in rats and humans. After collection, the lamina propria is enzymatically separated from the epithelium and stem cells are purified using an enzymatic or a non-enzymatic method. Purified olfactory stem cells can then be either grown in large numbers and banked in liquid nitrogen or induced to form spheres or differentiated into neural cells. These stem cells can also be used for comparative omics (genomic, transcriptomic, epigenomic, proteomic) studies.

We describe here a method for biopsying olfactory mucosa from rat and human nasal cavities. These biopsies can be used for either identifying molecular anomalies in brain diseases or isolating multipotent adult stem cells that can be utilized for cell transplantation in animal models of brain trauma/disease.

The olfactory mucosa, located in the nasal cavity, is in charge of detecting odours. It is also the only nervous tissue that is exposed to the external ...

Rodent Stereotaxic Surgery and Animal Welfare Outcome Improvements for Behavioral Neuroscience

Stereotaxic surgery for the implantation of cannulae into specific brain regions has for many decades been a very successful experimental technique to investigate the effects of locally manipulated neurotransmitter and signaling pathways in awake, behaving animals. Moreover, the stereotaxic implantation of electrodes for electrophysiological stimulation and recording studies has been instrumental to our current understanding of neuroplasticity and brain networks in behaving animals. Ever-increasing knowledge about optimizing surgical techniques in rodents1-4, public awareness concerning animal welfare issues and stringent legislation (e.g., the 2010 European Union Directive on the use of laboratory animals5) prompted us to refine these surgical procedures, particularly with respect to implementing new procedures for oxygen supplementation and the continuous monitoring of blood oxygenation and heart rate levels during the surgery as well as introducing a standardized protocol for post-surgical care. Our observations indicate that these modifications resulted in an increased survival rate and an improvement in the general condition of the animals after surgery (e.g. less weight loss and a more active animal). This video presentation will show the general procedures involved in this type of stereotaxic surgery with special attention to our several modifications. We will illustrate these surgical procedures in rats, but it is also possible to perform this type of surgery in mice or other small laboratory animals by using special adaptors for the stereotaxic apparatus6.

Stereotaxic surgery on rodents allows for targeted administration of drugs or electrical stimulation and recordings in awake, behaving animals. In this video presentation we will demonstrate recent procedural refinements to this long-standing procedure that successfully improved survival rate and reduced post-surgical weight loss.

Stereotaxic surgery for the implantation of cannulae into specific brain regions has for many decades been a very successful experimental technique to ...

Large-scale Recording of Neurons by Movable Silicon Probes in Behaving Rodents

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of neurons and circuits requires methods with the spatial selectivity and temporal resolution appropriate for mechanistic analysis of neural ensembles in the behaving animal, i.e. recording of representatively large samples of isolated single neurons. Ensemble monitoring of neuronal activity has progressed remarkably in the past decade in both small and large-brained animals, including human subjects1-11. Multiple-site recording with silicon-based devices are particularly effective because of their scalability, small volume and geometric design.
Here, we describe methods for recording multiple single neurons and local field potential in behaving rodents, using commercially available micro-machined silicon probes with custom-made accessory components. There are two basic options for interfacing silicon probes to preamplifiers: printed circuit boards and flexible cables. Probe supplying companies (<a href="http://www.neuronexustech.com/" >http://www.neuronexustech.com/</a>; <a href="http://www.sbmicrosystems.com/">http://www.sbmicrosystems.com/</a>; <a href="http://www.acreo.se/">http://www.acreo.se/</a>) usually provide the bonding service and deliver probes bonded to printed circuit boards or flexible cables. Here, we describe the implantation of a 4-shank, 32-site probe attached to flexible polyimide cable, and mounted on a movable microdrive. Each step of the probe preparation, microdrive construction and surgery is illustrated so that the end user can easily replicate the process.

We describe methods for large-scale recording of multiple single units and local field potential in behaving rodents with silicon probes. Drive fabrication, probe attachment to the drive and probe implantation processes are illustrated in sufficient details for easy replication.

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of ...

A Low Cost Setup for Behavioral Audiometry in Rodents

In auditory animal research it is crucial to have precise information about basic hearing parameters of the animal subjects that are involved in the experiments. Such parameters may be physiological response characteristics of the auditory pathway, e.g. via brainstem audiometry (BERA). But these methods allow only indirect and uncertain extrapolations about the auditory percept that corresponds to these physiological parameters. To assess the perceptual level of hearing, behavioral methods have to be used. A potential problem with the use of behavioral methods for the description of perception in animal models is the fact that most of these methods involve some kind of learning paradigm before the subjects can be behaviorally tested, e.g. animals may have to learn to press a lever in response to a sound. As these learning paradigms change perception itself 1,2 they consequently will influence any result about perception obtained with these methods and therefore have to be interpreted with caution. Exceptions are paradigms that make use of reflex responses, because here no learning paradigms have to be carried out prior to perceptual testing. One such reflex response is the acoustic startle response (ASR) that can highly reproducibly be elicited with unexpected loud sounds in na&iuml;ve animals. This ASR in turn can be influenced by preceding sounds depending on the perceptibility of this preceding stimulus: Sounds well above hearing threshold will completely inhibit the amplitude of the ASR; sounds close to threshold will only slightly inhibit the ASR. This phenomenon is called pre-pulse inhibition (PPI) 3,4, and the amount of PPI on the ASR gradually depends on the perceptibility of the pre-pulse. PPI of the ASR is therefore well suited to determine behavioral audiograms in na&iuml;ve, non-trained animals, to determine hearing impairments or even to detect possible subjective tinnitus percepts in these animals. In this paper we demonstrate the use of this method in a rodent model (cf. also ref. 5), the Mongolian gerbil (Meriones unguiculatus), which is a well know model species for startle response research within the normal human hearing range (e.g. 6).

A fast and inexpensive method for the behavioral determination of hearing parameters like hearing thresholds, hearing impairments or phantom perceptions (subjective tinnitus) is described. It uses pre-pulse inhibition of the acoustic startle response and can be easily implemented in a personal computer using a programmable AD/DA-converter and a piezo sensor.

In auditory animal research it is crucial to have precise information about basic hearing parameters of the animal subjects that are involved in the ...

Recording and Analysis of Circadian Rhythms in Running-wheel Activity in Rodents

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, including rats, hamsters, and mice, are active during the night and relatively inactive during the day. Many other behavioral and physiological measures also exhibit daily rhythms, but in rodents, running-wheel activity serves as a particularly reliable and convenient measure of the output of the master circadian clock, the suprachiasmatic nucleus (SCN) of the hypothalamus. In general, through a process called entrainment, the daily pattern of running-wheel activity will naturally align with the environmental light-dark cycle (LD cycle; e.g. 12 hr-light:12 hr-dark). However circadian rhythms are endogenously generated patterns in behavior that exhibit a ~24 hr period, and persist in constant darkness. Thus, in the absence of an LD cycle, the recording and analysis of running-wheel activity can be used to determine the subjective time-of-day. Because these rhythms are directed by the circadian clock the subjective time-of-day is referred to as the circadian time (CT). In contrast, when an LD cycle is present, the time-of-day that is determined by the environmental LD cycle is called the zeitgeber time (ZT).
Although circadian rhythms in running-wheel activity are typically linked to the SCN clock6-8, circadian oscillators in many other regions of the brain and body9-14 could also be involved in the regulation of daily activity rhythms. For instance, daily rhythms in food-anticipatory activity do not require the SCN15,16 and instead, are correlated with changes in the activity of extra-SCN oscillators17-20. Thus, running-wheel activity recordings can provide important behavioral information not only about the output of the master SCN clock, but also on the activity of extra-SCN oscillators. Below we describe the equipment and methods used to record, analyze and display circadian locomotor activity rhythms in laboratory rodents.

Circadian rhythms in voluntary wheel-running activity in mammals are tightly coupled to the molecular oscillations of a master clock in the brain. As such, these daily rhythms in behavior can be used to study the influence of genetic, pharmacological, and environmental factors on the functioning of this circadian clock.

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, ...

Deep Brain Stimulation with Simultaneous fMRI in Rodents

In order to visualize the global and downstream neuronal responses to deep brain stimulation (DBS) at various targets, we have developed a protocol for using blood oxygen level dependent (BOLD) functional magnetic resonance imaging (fMRI) to image rodents with simultaneous DBS. DBS fMRI presents a number of technical challenges, including accuracy of electrode implantation, MR artifacts created by the electrode, choice of anesthesia and paralytic to minimize any neuronal effects while simultaneously eliminating animal motion, and maintenance of physiological parameters, deviation from which can confound the BOLD signal.&#160;Our laboratory has developed a set of procedures that are capable of overcoming most of these possible issues. For electrical stimulation, a homemade tungsten bipolar microelectrode is used, inserted stereotactically at the stimulation site in the anesthetized subject. In preparation for imaging, rodents are fixed on a plastic headpiece and transferred to the magnet bore. For sedation and paralysis during scanning, a cocktail of dexmedetomidine and pancuronium is continuously infused, along with a minimal dose of isoflurane; this preparation minimizes the BOLD ceiling effect of volatile anesthetics. In this example experiment, stimulation of the subthalamic nucleus (STN) produces BOLD responses which are observed primarily in ipsilateral cortical regions, centered in motor cortex. Simultaneous DBS and fMRI allows the unambiguous modulation of neural circuits dependent on stimulation location and stimulation parameters, and permits observation of neuronal modulations free of regional bias. This technique may be used to explore the downstream effects of modulating neural circuitry at nearly any brain region, with implications for both experimental and clinical DBS.

This protocol describes a standard method for simultaneous functional magnetic resonance imaging and deep brain stimulation in the rodent. The combined use of these experimental tools allows for the exploration of global downstream activity in response to electrical stimulation at virtually any brain target.

In order to visualize the global and downstream neuronal responses to deep brain stimulation (DBS) at various targets, we have developed a protocol for ...

Acrylic Resin Molding Based Head Fixation Technique in Rodents

Head fixation is a technique of immobilizing animal's head by attaching a head-post on the skull for rigid clamping. Traditional head fixation requires surgical attachment of metallic frames on the skull. The attached frames are then clamped to a stationary platform resulting in immobilization of the head. However, metallic frames for head fixation have been technically difficult to design and implement in general laboratory environment. In this study, we provide a novel head fixation method. Using a custom-made head fixation bar, head mounter is constructed during implantation surgery. After the application of acrylic resin for affixing implants such as electrodes and cannula on the skull, additional resins applied on top of that to build a mold matching to the port of the fixation bar. The molded head mounter serves as a guide rails, investigators conveniently fixate the animal&#8217;s head by inserting the head mounter into the port of the fixation bar. This method could be easily applicable if implantation surgery using dental acrylics is necessary and might be useful for laboratories that cannot easily fabricate CNC machined metal head-posts.

Traditional head fixation techniques have used metallic frames attached on the skull as an interface for head fixation apparatus. This protocol demonstrates a frameless, acrylic resin molding based head fixation technique for behavioral tasks in rodents.

Head fixation is a technique of immobilizing animal's head by attaching a head-post on the skull for rigid clamping. Traditional head fixation requires ...

A Novel Strategy Combining Array-CGH, Whole-exome Sequencing and In Utero Electroporation in Rodents to Identify Causative Genes for Brain Malformations

Birth defects that involve the cerebral cortex &#8212; also known as malformations of cortical development (MCD) &#8212; are important causes of intellectual disability and account for 20-40% of drug-resistant epilepsy in childhood. High-resolution brain imaging has facilitated in vivo identification of a large group of MCD phenotypes. Despite the advances in brain imaging, genomic analysis and generation of animal models, a straightforward workflow to systematically prioritize candidate genes and to test functional effects of putative mutations is missing. To overcome this problem, an experimental strategy enabling the identification of novel causative genes for MCD was developed and validated. This strategy is based on identifying candidate genomic regions or genes via array-CGH or whole-exome sequencing and characterizing the effects of their inactivation or of overexpression of specific mutations in developing rodent brains via in utero electroporation. This approach led to the identification of the C6orf70 gene, encoding for a putative vesicular protein, to the pathogenesis of periventricular nodular heterotopia, a MCD caused by defective neuronal migration.

A Novel Strategy Combining Array-CGH, Whole-exome Sequencing and In Utero Electroporation in Rodents to Identify Causative Genes for Brain Malformations

Periventricular nodular heterotopia (PNH) is the most common form of malformation of cortical development (MCD) in adulthood but its genetic basis remains unknown in most sporadic cases. We have recently developed a strategy to identify novel candidate genes for MCDs and to directly confirm their causative role in vivo.

Birth defects that involve the cerebral cortex &#8212; also known as malformations of cortical development (MCD) &#8212; are important causes of ...

Transcranial Electrical Brain Stimulation in Alert Rodents

Transcranial electrical brain stimulation can modulate cortical excitability and plasticity in humans and rodents. The most common form of stimulation in humans is transcranial direct current stimulation (tDCS). Less frequently, transcranial alternating current stimulation (tACS) or transcranial random noise stimulation (tRNS), a specific form of tACS using an electrical current applied randomly within a pre-defined frequency range, is used. The increase of noninvasive electrical brain stimulation research in humans, both for experimental and clinical purposes, has yielded an increased need for basic, mechanistic, safety studies in animals. This article describes a model for transcranial electrical brain stimulation (tES) through the intact skull targeting the motor system in alert rodents. The protocol provides step-by-step instructions for the surgical set-up of a permanent epicranial electrode socket combined with an implanted counter electrode on the chest. By placing a stimulation electrode into the epicranial socket, different electrical stimulation types, comparable to tDCS, tACS, and tRNS in humans, can be delivered. Moreover, the practical steps for tES in alert rodents are introduced. The applied current density, stimulation duration, and stimulation type may be chosen depending on the experimental needs. The caveats, advantages, and disadvantages of this set-up are discussed, as well as safety and tolerability aspects.

This protocol describes a surgical set-up for a permanent epicranial electrode socket and an implanted chest electrode in rodents. By placing a second electrode into the socket, different types of transcranial electrical brain stimulation can be delivered to the motor system in alert animals through the intact skull.

Transcranial electrical brain stimulation can modulate cortical excitability and plasticity in humans and rodents. The most common form of stimulation in ...