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 °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="Rodent dissection diagram illustrating surgical steps, likely for organ extraction research." src="/files/ftp_upload/3564/3564step7.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
<a href="https://www.jove.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 °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 °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 °C while stirring (Do not exceed 65 °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 °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="Perfusion apparatus diagram with manometer gauge, fixative, buffer inlets, and perfusion needle setup." src="/files/ftp_upload/3564/3564fig1.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1. Perfusion Apparatus with labeled components.
<img alt="Flow cytometry setup diagram; fluidics system for cell analysis using fixative and buffer solutions." src="/files/ftp_upload/3564/3564fig2.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
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="Chromatography system diagram; showing fluid paths, fixative buffer setup; chemical separation." src="/files/ftp_upload/3564/3564fig3.jpg" data-altupdatedat="1747911167000" data-alt="Figure 3"> 
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="Histology tissue fixation setup, diagram of fluid flow through fixative, buffer containers." src="/files/ftp_upload/3564/3564fig4.jpg" data-altupdatedat="1747911167000" data-alt="Figure 4"> 
Figure 4. Perfusion Apparatus in position for buffer delivery.
<img alt="Rodent dissection diagram; surgical steps for thoracic cavity exposure; anatomical study." src="/files/ftp_upload/3564/3564fig5.jpg" data-altupdatedat="1747911167000" data-alt="Figure 5"> 
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="Rodent cardiac surgery diagram showing heart exposure and surgical incision setup." src="/files/ftp_upload/3564/3564fig6.jpg" data-altupdatedat="1747911167000" data-alt="Figure 6"> 
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="Rat perfusion method diagram showing fixative and buffer flow for tissue preservation research." src="/files/ftp_upload/3564/3564fig7.jpg" data-altupdatedat="1747911167000" data-alt="Figure 7"> 
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="Rat perfusion setup diagram; fixative-buffer process for biological preservation experiment." src="/files/ftp_upload/3564/3564fig8.jpg" data-altupdatedat="1747911167000" data-alt="Figure 8"> 
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="Rat dissection diagram showing incision lines for anatomical study and organ examination guidelines." src="/files/ftp_upload/3564/3564fig9.jpg" data-altupdatedat="1747911167000" data-alt="Figure 9"> 
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="Nasal cavity development diagram; stages a-c; embryonic growth; anatomical changes; red arrows." src="/files/ftp_upload/3564/3564fig10.jpg" data-altupdatedat="1747911167000" data-alt="Figure 10"> 
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.

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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><tbody><tr><td colspan="4">&lt;strong&gt;Anesthetic:&lt;/strong&gt;</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 colspan="4">&lt;strong&gt;Surgery:&lt;/strong&gt;</td></tr><tr><td>Needle tip, 27 GA x 1.25&quot;</td><td>Materiel Services</td><td>25251</td><td></td></tr><tr><td>Large blunt/blunt curved scissors (~14.5 cm)</td><td>Fine Science Tools</td><td>14519-14</td><td></td></tr><tr><td>Straight iris scissors</td><td>Fine Science Tools</td><td>14058-11</td><td></td></tr><tr><td>Standard tweezers</td><td>Fine Science Tools</td><td>11027-12</td><td></td></tr><tr><td>Pair of fine (Graefe) tweezers</td><td>Fine Science Tools</td><td>11050-10</td><td></td></tr><tr><td>1 large hemostat forceps - curved or straight (~19 cm)</td><td>surgicaltools.com</td><td>17.21.51</td><td></td></tr><tr><td>2 standard hemostat forceps - straight serrated (14 cm)</td><td>Fine Science Tools</td><td>13013-14</td><td></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></td></tr><tr><td>15-gauge blunt or olive-tipped needle (perfusion needle)</td><td>Fisnar</td><td>5601137</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;Perfusion&lt;/strong&gt;:</td></tr><tr><td>HyPerfusion system or equivalent</td><td></td><td></td><td></td></tr><tr><td>Phosphate buffered saline, pH 7.4</td><td></td><td></td><td></td></tr><tr><td>4% Paraformaldehyde in 0.1 M Phosphate Buffer, pH 7.4</td><td></td><td></td><td></td></tr><tr><td>Shallow glass or plastic tray, approximately 10&quot; x 10&quot;</td><td></td><td></td><td></td></tr><tr><td>Crushed ice</td><td></td><td></td><td></td></tr><tr><td>Water bath (37 &amp;deg;C)</td><td></td><td></td><td></td></tr><tr><td>Timer</td><td></td><td></td><td></td></tr><tr><td>50 ml syringe</td><td>Medline Industries</td><td>NPMJD50LZ</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;Dissection&lt;/strong&gt;:</td></tr><tr><td>Pair of standard sharp/blunt straight scissors (~12 cm)</td><td>Fine Science Tools</td><td>14054-13</td><td></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></td></tr><tr><td>Straight iris scissors (~9 cm)</td><td>Fine Science Tools</td><td>14058-11</td><td></td></tr><tr><td>Micro-spatula (double 2&quot; flat ends, one rounded, one tapered to 1/8&quot;)</td><td>Fine Science Tools</td><td>10091-12</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;Post-Fixation &amp;amp; Storage:&lt;/strong&gt;</td></tr><tr><td>50 ml glass vial</td><td></td><td></td><td></td></tr><tr><td>40 ml HEPES-Buffered Hanks Solution (HBHS) with sodium azide (90mg/l)</td><td></td><td></td><td></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

Research

JoVE Journal

Neuroscience

381.4K 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

2-Vessel Occlusion/Hypotension: A Rat Model of Global Brain Ischemia

Cardiac arrest followed by resuscitation often results in dramatic brain damage caused by ischemia and subsequent reperfusion of the brain. Global brain ischemia produces damage to specific brain regions shown to be highly sensitive to ischemia 1. Hippocampal neurons have higher sensitivity to ischemic insults compared to other cell populations, and specifically, the CA1 region of the hippocampus is particularly vulnerable to ischemia/reperfusion 2.
The design of therapeutic interventions, or study of mechanisms involved in cerebral damage, requires a model that produces damage similar to the clinical condition and in a reproducible manner. Bilateral carotid vessel occlusion with hypotension (2VOH) is a model that produces reversible forebrain ischemia, emulating the cerebral events that can occur during cardiac arrest and resuscitation. We describe a model modified from Smith et al. (1984) 2, as first presented in its current form in Sanderson, et al. (2008) 3, which produces reproducible injury to selectively vulnerable brain regions 3-6. The reliability of this model is dictated by precise control of systemic blood pressure during applied hypotension, the duration of ischemia, close temperature control, a specific anesthesia regimen, and diligent post-operative care. An 8-minute ischemic insult produces cell death of CA1 hippocampal neurons that progresses over the course of 6 to 24 hr of reperfusion, while less vulnerable brain regions are spared. This progressive cell death is easily quantified after 7-14 days of reperfusion, as a near complete loss of CA1 neurons is evident at this time.
In addition to this brain injury model, we present a method for CA1 damage quantification using a simple, yet thorough, methodology. Importantly, quantification can be accomplished using a simple camera-mounted microscope, and a free ImageJ (NIH) software plugin, obviating the need for cost-prohibitive stereology software programs and a motorized microscopic stage for damage assessment.

Bilateral carotid occlusion coupled with systemic hypotension produces global brain ischemia in the rat, resulting in damage to the hippocampus with reproducible severity. Animal subjects are impaired with predictable patterns of brain damage, they recover expediently, and mortality rates are comparatively low.

Cardiac arrest followed by resuscitation often results in dramatic brain damage caused by ischemia and subsequent reperfusion of the brain. Global brain ...

Medicine

Intact Histological Characterization of Brain-implanted Microdevices and Surrounding Tissue

Research into the design and utilization of brain-implanted microdevices, such as microelectrode arrays, aims to produce clinically relevant devices that interface chronically with surrounding brain tissue. Tissue surrounding these implants is thought to react to the presence of the devices over time, which includes the formation of an insulating "glial scar" around the devices. However, histological analysis of these tissue changes is typically performed after explanting the device, in a process that can disrupt the morphology of the tissue of interest.
Here we demonstrate a protocol in which cortical-implanted devices are collected intact in surrounding rodent brain tissue. We describe how, once perfused with fixative, brains are removed and sliced in such a way as to avoid explanting devices. We outline fluorescent antibody labeling and optical clearing methods useful for producing an informative, yet thick tissue section. Finally, we demonstrate the mounting and imaging of these tissue sections in order to investigate the biological interface around brain-implanted devices.

Here we present a histological method for capturing, labeling, optically clearing, and imaging the intact brain tissue interface around chronically implanted microdevices in rodent brain tissue. Results from the techniques comprising this method are useful for understanding the impact of various penetrating brain-implants on their surrounding tissue.

Research  into the design and utilization of brain-implanted microdevices, such as  microelectrode arrays, aims to produce clinically relevant devices ...

An In Vivo Blood-brain Barrier Permeability Assay in Mice Using Fluorescently Labeled Tracers

Blood-brain barrier (BBB) is a specialized barrier that protects the brain microenvironment from toxins and pathogens in the circulation and maintains brain homeostasis. The principal sites of the barrier are endothelial cells of the brain capillaries whose barrier function results from tight intercellular junctions and efflux transporters expressed on the plasma membrane. This function is regulated by pericytes and astrocytes that together form the neurovascular unit (NVU). Several neurological diseases such as stroke, Alzheimer&#39;s disease (AD), brain tumors are associated with an impaired BBB function. Assessment of the BBB permeability is therefore crucial in evaluating the severity of the neurological disease and the success of the treatment strategies employed.
We present here a simple yet robust permeability assay that have been successfully applied to several mouse models both, genetic and experimental. The method is highly quantitative and objective in comparison to the tracer fluorescence analysis by microscopy that is commonly applied. In this method, mice are injected intraperitoneally with a mix of aqueous inert fluorescent tracers followed by anesthetizing the mice. Cardiac perfusion of the animals is performed prior to harvesting brain, kidneys or other organs. Organs are homogenized and centrifuged followed by fluorescence measurement from the supernatant. Blood drawn from the cardiac puncture just before perfusion serves for normalization purpose to the vascular compartment. The tissue fluorescence is normalized to the wet weight and serum fluorescence to obtain a quantitative tracer permeability index. For additional confirmation, the contralateral hemi-brain preserved for immunohistochemistry can be utilized for tracer fluorescence visualization purposes.

An In Vivo Blood-brain Barrier Permeability Assay in Mice Using Fluorescently Labeled Tracers

Here we present a mouse brain vascular permeability assay using intraperitoneal injection of fluorescent tracers followed by perfusion that is applicable to animal models of blood-brain barrier dysfunction. One hemi-brain is used for assessing permeability quantitatively and the other for tracer visualization/immunostaining. The procedure takes 5 - 6 h for 10 mice.

Blood-brain barrier (BBB) is a specialized barrier that protects the brain microenvironment from toxins and pathogens in the circulation and maintains ...

Novel Passive Clearing Methods for the Rapid Production of Optical Transparency in Whole CNS Tissue

Since the development of CLARITY, a bioelectrochemical clearing technique that allows for three-dimensional phenotype mapping within transparent tissues, a multitude of novel clearing methodologies including CUBIC (clear, unobstructed brain imaging cocktails and computational analysis), SWITCH (system-wide control of interaction time and kinetics of chemicals), MAP (magnified analysis of the proteome), and PACT (passive clarity technique), have been established to further expand the existing toolkit for the microscopic analysis of biological tissues. The present study aims to improve upon and optimize the original PACT procedure for an array of intact rodent tissues, including the whole central nervous system (CNS), kidneys, spleen, and whole mouse embryos. Termed psPACT (process-separate PACT) and mPACT (modified PACT), these novel techniques provide highly efficacious means of mapping cell circuitry and visualizing subcellular structures in intact normal and pathological tissues. In the following protocol, we provide a detailed, step-by-step outline on how to achieve maximal tissue clearance with minimal invasion of their structural integrity via psPACT and mPACT.

Here, we present two novel methodologies, psPACT and mPACT, for achieving maximal optical transparency and subsequent microscopic analysis of tissue vasculature in the intact rodent whole CNS.

Since the development of CLARITY, a bioelectrochemical clearing technique that allows for three-dimensional phenotype mapping within transparent tissues, ...

A Micro-CT-based Method for Characterizing Lesions and Locating Electrodes in Small Animal Brains

Lesion and electrode location verification are traditionally done via histological examination of stained brain slices, a time-consuming procedure that requires manual estimation. Here, we describe a simple, straightforward method for quantifying lesions and locating electrodes in the brain that is less laborious and yields more detailed results. Whole brains are stained with osmium tetroxide, embedded in resin, and imaged with a micro-CT scanner. The scans result in 3D digital volumes of the brains with resolutions and virtual section thicknesses dependent on the sample size (12&#8211;15 and 5&#8211;6 &#181;m&#160;per voxel for rat and zebra finch brains, respectively). Surface and deep lesions can be characterized, and single tetrodes, tetrode arrays, electrolytic lesions, and silicon probes can also be localized. Free and proprietary software allows experimenters to examine the sample volume from any plane and segment the volume manually or automatically. Because this method generates whole brain volume, lesions and electrodes can be quantified to a much higher degree than in current methods, which will help standardize comparisons within and across studies.

This article describes a straightforward method to prepare small animal brains for micro-CT imaging, in which lesions can be quantified and electrodes located with high precision in the context of the whole brain.

Lesion and electrode location verification are traditionally done via histological examination of stained brain slices, a time-consuming procedure that ...

AAV Deployment of Enhancer-Based Expression Constructs In Vivo in Mouse Brain

Enhancers are binding platforms for a diverse array of transcription factors that drive specific expression patterns of tissue- and cell-type-specific genes. Multiple means of assessing non-coding DNA and various chromatin states have proven useful in predicting the presence of enhancer sequences in the genome, but validating the activity of these sequences and finding the organs and developmental stages they are active in is a labor-intensive process. Recent advances in adeno-associated virus (AAV) vectors have enabled the widespread delivery of transgenes to mouse tissues, enabling in vivo enhancer testing without necessitating a transgenic animal. This protocol shows how a reporter construct that expresses EGFP under the control of a minimal promoter, which does not drive significant expression on its own, can be used to study the activity patterns of candidate enhancer sequences in the mouse brain. An AAV-packaged reporter construct is delivered to the mouse brain and incubated for 1-4 weeks, after which the animal is sacrificed, and brain sections are observed under a microscope. EGFP appears in cells in which the tested enhancer is sufficient to initiate gene expression, pinpointing the location and developmental stage in which the enhancer is active in the brain. Standard cloning methods, low-cost AAV packaging, and expanding AAV serotypes and methods for in vivo delivery and standard imaging readout make this an accessible approach for the study of how gene expression is regulated in the brain.

AAV Deployment of Enhancer-Based Expression Constructs In Vivo in Mouse Brain

This protocol describes a novel rAAV-based transient enhancer-reporter assay. This assay can be used to induce enhancer-driven expression in vivo in the mouse brain.

Enhancers are binding platforms for a diverse array of transcription factors that drive specific expression patterns of tissue- and cell-type-specific ...

Establishing In Situ Closed Circuit Perfusion of Lower Abdominal Organs and Hind Limbs in Mice

Ex vivo perfusion is an important physiological tool to study the function of isolated organs (e.g. liver, kidneys). At the same time, due to the small size of mouse organs, ex vivo perfusion of bone, bladder, skin, prostate, and reproductive organs is challenging or not feasible. Here, we report for the first time an in situ lower body perfusion circuit in mice that includes the above tissues, but bypasses the main clearance organs (kidney, liver, and spleen). The circuit is established by cannulating the abdominal aorta and inferior vena cava above the iliac artery and vein and cauterizing peripheral blood vessels. Perfusion is performed via a peristaltic pump with perfusate flow maintained for up to 2 h. In situ staining with fluorescent lectin and Hoechst solution confirmed that the microvasculature was successfully perfused. This mouse model can be a very useful tool for studying pathological processes as well as mechanisms of drug delivery, migration/metastasis of circulating tumor cells into/from the tumor, and interactions of immune system with perfused organs and tissues.

A protocol is described for in situ perfusion of the mouse lower body, including the bladder, the prostate, sex organs, bone, muscle and foot skin.

Ex vivo perfusion is an important physiological tool to study the function of isolated organs (e.g. liver, kidneys). At the same time, due to the small ...

Free-floating Immunostaining of Mouse Brains

Immunohistochemical staining of mouse brains is a routine technique commonly used in neuroscience to investigate central mechanisms underlying the regulation of energy metabolism and other neurobiological processes. However, the quality, reliability, and reproducibility of brain histology results may vary among laboratories. For each staining experiment, it is necessary to optimize the key procedures based on differences in species, tissues, targeted proteins, and the working conditions of the reagents. This paper demonstrates a reliable workflow in detail, including intra-aortic perfusion, brain sectioning, free-floating immunostaining, tissue mounting, and imaging, which can be followed easily by researchers in this field.
Also discussed are how to modify these procedures to satisfy the individual needs of researchers. To illustrate the reliability and efficiency of this protocol, perineuronal nets were stained with biotin-labeled Wisteria florbunda agglutinin (WFA) and arginine vasopressin (AVP) with an anti-AVP antibody in the mouse brain. Finally, the critical details for the entire procedure have been addressed, and the advantages of this protocol compared to those of other protocols. Taken together, this paper presents an optimized protocol for free-floating immunostaining of mouse brain tissue. Following this protocol makes this process easier for both junior and senior scientists to improve the quality, reliability, and reproducibility of immunostaining studies.

This protocol describes an efficient and reproducible approach for mouse brain histological studies, including perfusion, brain sectioning, free-floating immunostaining, tissue mounting, and imaging.

Immunohistochemical staining of mouse brains is a routine technique commonly used in neuroscience to investigate central mechanisms underlying the ...

A Gravity-Fed Transcardial Perfusion Method for Histologic Analysis of the Mouse Central Nervous System

The histologic analysis of brain and spinal cord specimens isolated from mice is common practice for the assessment of pathology in this model system. To maintain the morphology of these delicate tissues, it is routine to administer a chemical fixative such as paraformaldehyde via cannulation of the heart in anesthetized animals (transcardial perfusion). Transcardial perfusion of the mouse heart has traditionally relied on the use of peristaltic pumps or air pressure to deliver both the saline and fixative solutions necessary for this process. As an easily accessible alternative to these methods, this work demonstrates the use of a gravity-fed method of perfusate delivery that uses materials available in most hardware stores. 
To validate this new perfusion method, this work demonstrates all the subsequent steps necessary for the sensitive detection of phosphorylated &#945;-synuclein in both the brain and spinal cord. Included in these steps are the dissection of the fixed brain and spinal cord tissues, rapid freezing/embedding and cryosectioning of the tissues, and immunofluorescent staining. As this method results in whole-body delivery of the fixative, it may also be used to prepare other non-neuronal tissues for histologic analysis.

A convenient gravity-fed perfusion method for histological analysis of the mouse central nervous system is presented. Immunofluorescent detection of phosphorylated &#945;-synuclein is demonstrated in a mouse model of Parkinson's disease. This work also comprehensively describes the transcardial perfusion, dissection, tissue freezing/embedding, and frozen sectioning steps.

The histologic analysis of brain and spinal cord specimens isolated from mice is common practice for the assessment of pathology in this model system. To ...

Protocol for Dendritic Spine Analysis Using Neurolucida 360

Dendritic Spine Quantification Using an Automatic Three-Dimensional Neuron Reconstruction Software

Synaptic connections allow for the exchange and processing of information between neurons. The post-synaptic site of excitatory synapses is often formed on dendritic spines. Dendritic spines are structures of great interest in research centered around synaptic plasticity, neurodevelopment, and neurological and psychiatric disorders. Dendritic spines undergo structural modifications during their lifespan, with properties such as total spine number, dendritic spine size, and morphologically defined subtype altering in response to different processes. Delineating the molecular mechanisms regulating these structural alterations of dendritic spines relies on morphological measurement. This mandates accurate and reproducible dendritic spine analysis to provide experimental evidence. The present study outlines a detailed protocol for dendritic spine quantification and classification using Neurolucida 360 (automatic three-dimensional neuron reconstruction software). This protocol allows for the determination of key dendritic spine properties such as total spine density, spine head volume, and classification into spine subtypes thus enabling effective analysis of dendritic spine structural phenotypes.

Author Spotlight: Optimizing Dendritic Spine Analysis for Balanced Manual and Automated Assessment in the Hippocampus CA1 Apical Dendrites

Dendritic spines are post-synaptic compartments of most excitatory synapses. Alterations to dendritic spine morphology occur during neurodevelopment, aging, learning, and many neurological and psychiatric disorders, underscoring the importance of reliable dendritic spine analysis. This protocol describes quantifying dendritic spine morphology accurately and reproducibly using automatic three-dimensional neuron reconstruction software.

Synaptic connections allow for the exchange and processing of information between neurons. The post-synaptic site of excitatory synapses is often formed ...

Whole Animal Perfusion Fixation for Rodents

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Przeglądaj więcej filmów

2-Vessel Occlusion/Hypotension: A Rat Model of Global Brain Ischemia

Intact Histological Characterization of Brain-implanted Microdevices and Surrounding Tissue

An In Vivo Blood-brain Barrier Permeability Assay in Mice Using Fluorescently Labeled Tracers

Novel Passive Clearing Methods for the Rapid Production of Optical Transparency in Whole CNS Tissue

A Micro-CT-based Method for Characterizing Lesions and Locating Electrodes in Small Animal Brains

AAV Deployment of Enhancer-Based Expression Constructs In Vivo in Mouse Brain

Establishing In Situ Closed Circuit Perfusion of Lower Abdominal Organs and Hind Limbs in Mice

Free-floating Immunostaining of Mouse Brains

A Gravity-Fed Transcardial Perfusion Method for Histologic Analysis of the Mouse Central Nervous System

Dendritic Spine Quantification Using an Automatic Three-Dimensional Neuron Reconstruction Software