eoe sub articles

EoE Sub Articles

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.&#160;&#160;
1. Preparations
<ol>
	<li>Modify a 16 G peripheral vein catheter for intubation. To modify, shorten the needle by 1 cm and bend the remaining distal 1 cm by 30&#176; toward the injection valve. Remove the catheter wings (multiple use).</li>
	<li>To make an intracranial pressure or ICP probe, cut a 20 mm piece of polythene tubing (inner diameter (ID): 0.58 mm, outer diameter (OD): 0.96 mm) and burn one end to make a circular plate, keeping an open lumen. Circumvent the polythene tubing with 1 mm of silicone tubing (ID: 1.0 mm, OD: 3.0 mm) before connecting 10 mm of silicone tubing (ID: 0.76 mm, OD: 2.4 mm) to the end of the polythene tubing.</li>
	<li>Power on the laptop and open the data acquisition software. Calibrate the blood pressure (BP) and intracerebral pressure (ICP) transducers and the Laser-Doppler according to the manufacturer&#39;s instructions.</li>
	<li>Prepare the blood gas analyzer apparatus. 
	CAUTION: Make sure there is enough isoflurane in the vaporizer.</li>
	<li>Turn on the O2 and atmospheric air flow. Set the flow of O2 at 30% and atmospheric air at 70%.</li>
	<li>Place the heating pad and set the temperature to 37 &#176;C.</li>
</ol>
2. Anesthesia
<ol>
	<li>Place the rat in the anesthesia chamber with a flow of 30% of O2 and 70% of atmospheric air. Administer 5% of isoflurane gas into the chamber. Adequate anesthesia will take around 4 min. Control the breathing carefully.</li>
	<li>When anesthetized, place the rat in supine position on a heavy plate circumvented by a rubber band. Place the front teeth of the rat below the rubber band.</li>
	<li>Draw the tongue out carefully with curved forceps. Clean the larynx with a cotton tip. Place an external light in the midline of the throat to visualize the vocal cords.</li>
	<li>Intubate during inspiration using the modified 16 G peripheral vein catheter. When correctly inserted, remove the stiletto. Connect the catheter to the ventilator. 
	NOTE: Correct placement of the tube is confirmed by chest movements in sync with respiration rate. If movements of the abdomen are seen, extubate and reintroduce the rat into the anesthesia bell. Do not repeat the procedure more than three times due to the risk of damaging the airways.</li>
	<li>When intubated, keep the animal on artificial respiration with 30% of O2 and 70% of atmospheric air. Maintain the anesthesia at 1.5%-3% of isoflurane. Adjust the isoflurane to keep the blood pressure between 80-100 mmHg.</li>
	<li>Keep the inspiratory volume of the respirator at 3 mL and the frequency at 40-45 inspirations/min. Adjust the inspiratory volume according to the blood gas analysis.</li>
	<li>Make a stitch through the inner soft tissue of the cheek with a 2-0 suture. Tie the suture around the injection tube and the injection valve of the peripheral vein catheter to fasten the catheter.</li>
	<li>Move the rat to the operating field and place it in supine position with the tail facing toward the surgeon.</li>
	<li>Apply the eye gel when needed to counter dry eyes.</li>
	<li>Perform a toe pinch to confirm an adequate depth of anesthesia. Assess and maintain anesthesia depth during surgery.</li>
</ol>
3. Tail catheter
<ol>
	<li>Disinfect the proximal 3-4 cm of the tail with 0.5% of chlorhexidine ethanol. 
	NOTE: From now on, use the surgical microscope at the surgeon&#39;s discretion.</li>
	<li>Make a 15-20 mm skin incision in the proximal end of the tail on the ventral side. Be careful not to incise the artery.</li>
	<li>Loosen the skin from the underlying connective tissue using a curved forceps.</li>
	<li>Carefully penetrate the fascia exposing the artery.</li>
	<li>Carefully release the tail artery from the underlying tissue using a curved forceps.</li>
	<li>Slip three black silk threads under the vessel. Place one thread as distally as possible and tie a surgical knot tightly around the artery. Hold the loose ends of the thread with a hemostat.</li>
	<li>Tie the two remaining threads loosely around the artery.</li>
	<li>Push the proximal thread as proximally as possible. Apply a hemostat to hold the ends of the proximal thread. Pull the hemostat lightly, but enough to restrict and block the blood flow. Place the hemostat on the abdomen.</li>
	<li>Cut the tip of the catheter at a 45&#176; angle. Cut the sharp point to prevent arterial wall penetration.</li>
	<li>Using a Vannas scissor, make an artery incision 1/3 of the artery&#39;s diameter at a 30&#176; angle, 3-5 mm from the distal knot.</li>
	<li>Insert the catheter into the artery using two straight forceps. Use one forceps to hold the catheter and the other to carefully pull the artery over the catheter.</li>
	<li>Insert the catheter up the vessel to the proximal knot and loosen the knot from the hemostat. Visualize the blood flow in the catheter. Fasten the middle thread loosely to the catheter.</li>
	<li>Continue insertion to, and if possible, just beyond, the point where the artery is covered again by fascia.</li>
	<li>Control the catheter placement and possible leak by flushing with saline.</li>
	<li>Fasten the two proximal threads using surgical knots. 
	NOTE: The blood pressure measurement needs to be pulsatile; if not, the catheter is not properly placed.</li>
	<li>Fasten the catheter at the end of the incision by tying a surgical knot using the distal thread.</li>
	<li>Stitch the skin incision loosely together with two non-resorbable monofilament 4-0 suture. Be careful not to penetrate the catheter. 
	NOTE: Throughout the surgery be aware of the amplitude of pulsation. If this is low, flush the catheter with saline.</li>
	<li>Loosen the arterial catheter from the pressure transducer to allow blood flow for blood gas sampling. Place a microcapillary tube at the end of the catheter. Let the blood flow into the tube. Re-attach the catheter to the transducer after blood collection and flush the catheter.</li>
	<li>Insert the capillary tube in the blood gas analyzer. Measure the pH, pCO2, and pO2 and note them down 
	&#8203;NOTE: Depending on the blood gas and blood pressure values, change the ventilation rate. If the mean arterial pressure (MAP) is too low, try to turn down the flowrate of isoflurane. Test the reflexes to ensure proper depth of anesthesia.</li>
</ol>
4. ICP probe
<ol>
	<li>Place the rat in the stereotaxic frame. It is important to position the rat symmetrically.</li>
	<li>Place a cylindrical pillow under the stereotaxic frame to create anterior flexion of the neck.</li>
	<li>Shave the rat&#39;s scalp, neck, and the area behind the ears. Remove the superfluous hair.</li>
	<li>Disinfect the area with 0.5% of chlorhexidine ethanol.</li>
	<li>Anesthetize locally with 0.7 mL of 10 mg/5 &#181;g/mL lidocaine with adrenaline, insert the needle at the caudal end of the skull in the midline. Inject into the musculature of the neck with 0.3-0.4 mL. Inject the rest subcutaneously around and anterior to the bregma.</li>
	<li>Make a skin incision from the needle puncture ~8 mm caudally in the midline.</li>
	<li>Dissect all the muscles bluntly in layers to identify the atlantooccipital membrane (marble-colored triangle caudally to the skull in the midline).</li>
	<li>Use the Alm retractor to restrain the neck musculature. Place the pronged retractor caudally if needed.</li>
	<li>Check whether the sterile ICP probe is connected to the ICP transducer. Flush the ICP probe with saline. Ensure no air bubbles are present in the ICP probe.</li>
	<li>Incise the atlantooccipital membrane using a 23 G needle. Make a hole to coax the ICP probe through the membrane.</li>
	<li>Coax the probe through the atlantooccipital membrane gently. Pull the probe lightly and ensure that it shows a pulsating curve ranging between 0-5 mmHg. If not, remove the probe, check the connection to the transducer, and confirm the flow through the lumen.</li>
	<li>Apply two drops of the tissue glue. Move the 1 mm silicone tubing forward to the membrane and apply additional glue to minimize the risk of ICP-probe displacement.</li>
	<li>Remove the retractor(s).</li>
	<li>Make one horizontal mattress suture to the cephalic end of the incision and one simple interrupted suture to the caudal end using a non-resorbable monofilament 4-0 suture.</li>
</ol>
5. Placement of the needle and the Laser-Doppler probe
<ol>
	<li>Make an incision in the midline just anterior to the eyes, 15 mm caudally.</li>
	<li>Remove the connective tissue and the muscles with forceps. Use the end of a sterile cotton swab as a rougine making it possible to identify the bregma and the coronal sutures.</li>
	<li>Place the Alm retractor.</li>
	<li>Place a 25 G spinal needle in the stereotaxic frame. Place the needle exactly on the bregma and note the position. 
	NOTE: Place the midline joint of the stereotaxic frame at 30&#176; toward the animal in the vertical plane.</li>
	<li>Remove the needle from the bregma, move the frame 65 mm anteriorly, and then replace the needle in the midline to mark the site of drilling.</li>
	<li>Drill until the dura mater is identified below the bone. Gently remove the bone fragments using straight forceps and fill the cavity with bone wax.</li>
	<li>Drill another hole 3-4 mm lateral to the right of the bregma and just anterior to the coronal suture for the Laser-Doppler. It is not necessary to drill all the way through the bone. Be careful not to penetrate the dura mater.</li>
	<li>Look for the vessels where the laser-doppler can measure the blood flow. Place the laser-doppler and check the values. A minimum value of 100 FU is required. Remove the microscope (artificial light).</li>
	<li>If the values are still acceptable, add one drop of glue to fix the probe.</li>
	<li>Recheck to confirm whether the value is above 80 FU. If the value is below 80 FU, remove and reposition the probe to reach a value above 80 FU. 
	&#8203;NOTE: The value, FU, is an arbitrary unit showing cerebral blood flow (CBF).</li>
</ol>
6. Induction of subarachnoid hemorrhage (SAH)
<ol>
	<li>Insert the needle gently through the skull in the midline between the hemispheres until resistance of the base of the skull is felt. Retract the needle by 1 mm to ensure correct placement just anteriorly to the optic chiasm.</li>
	<li>Turn the needle 90&#176; clockwise so that the needle tip points to the right to ensure the most homogenous result when injecting the blood. Remove the stiletto (Figure 1).</li>
	<li>Equilibrate for 15 min and adjust the level of anesthesia to obtain a mean arterial blood pressure in the range 80-100 mmHg.</li>
	<li>Perform a blood gas analysis. Adjust the level of anesthesia accordingly.</li>
	<li>Withdraw 500 &#181;L of blood from the tail catheter using a 1 mL syringe with a blunt 23 G needle.</li>
	<li>Fill the dead space of spinal needle chamber with blood to avoid injection of air. Remove the 23 G needle from the blood-filled syringe and confirm that the syringe contains 300 &#181;L of blood.</li>
	<li>Connect the syringe to the spinal needle. Grasp firmly and inject the blood manually to surpass MAP.</li>
	<li>Observe a steep rise in ICP and a steep fall in CBF on the laptop. 
	NOTE: CBF should be 50% or lower compared to the baseline score for at least 5 min for the surgery to be successful, see Figure 2. Sham rats do not undergo the steps 6.1-6.7, thereby omitting the introduction of the spinal needle into the cerebrum, minimizing possible spontaneous hemorrhage, and iatrogenic brain damage.</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>16 G peripheral vein catheter</td>
			<td>BD Venflon</td>
			<td>393229</td>
			<td>Needle shortened, distal 1 cm curved. Wings removed</td>
		</tr>
		<tr>
			<td>Anesthesia bell/ chamber</td>
			<td>Unknown</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Blood gas analyzer</td>
			<td>Radiometer</td>
			<td>ABL80</td>
			<td></td>
		</tr>
		<tr>
			<td>Blood pressure (BP) monitor</td>
			<td>Adinstruments</td>
			<td>ML117</td>
			<td>Connects to Powerlab</td>
		</tr>
		<tr>
			<td>Curved forceps, 12 cm x 3</td>
			<td>F.S.T</td>
			<td>11001-12</td>
			<td>For anesthesia</td>
		</tr>
		<tr>
			<td>Cylindrical pillow, 28 cm x 4 cm</td>
			<td>Homemade</td>
			<td></td>
			<td>Made from surgical towels</td>
		</tr>
		<tr>
			<td>Data acquisition hardware</td>
			<td>Adinstruments</td>
			<td>ML870 Powerlab</td>
			<td></td>
		</tr>
		<tr>
			<td>Data acquistion software</td>
			<td>Adinstruments</td>
			<td>LabChart 6.0</td>
			<td></td>
		</tr>
		<tr>
			<td>Drill</td>
			<td>KMD</td>
			<td>1189</td>
			<td></td>
		</tr>
		<tr>
			<td>Drill controller</td>
			<td>Silfradent</td>
			<td>300 IN</td>
			<td></td>
		</tr>
		<tr>
			<td>Flexible light</td>
			<td>Schott</td>
			<td>KL200</td>
			<td></td>
		</tr>
		<tr>
			<td>Heating pad</td>
			<td>Minco</td>
			<td>1135</td>
			<td></td>
		</tr>
		<tr>
			<td>Hypodermic needle, 20 G</td>
			<td>KD Medical</td>
			<td>301300</td>
			<td>Connects to stereotaxic frame</td>
		</tr>
		<tr>
			<td>ICP monitor</td>
			<td>Adinstruments</td>
			<td>ML117</td>
			<td>Connects to Powerlab</td>
		</tr>
		<tr>
			<td>Isoflurane vaporizer</td>
			<td>Ohmeda</td>
			<td>TEC3</td>
			<td></td>
		</tr>
		<tr>
			<td>Laptop</td>
			<td>Lenovo</td>
			<td>T410</td>
			<td></td>
		</tr>
		<tr>
			<td>Laser doppler monitor</td>
			<td>Adinstruments</td>
			<td>ML191</td>
			<td></td>
		</tr>
		<tr>
			<td>Laser doppler probe</td>
			<td>Oxford Optronics</td>
			<td>MSF100XP</td>
			<td>Connects to laser doppler monitor</td>
		</tr>
		<tr>
			<td>Needle holder, 13 cm</td>
			<td>F.S.T</td>
			<td>12001-13</td>
			<td>For anesthesia</td>
		</tr>
		<tr>
			<td>Precision syringe, 0.025 mL</td>
			<td>Hamilton</td>
			<td>547407</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereotaxic frame</td>
			<td>Kopf Instruments</td>
			<td>M900</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical microscope</td>
			<td>Carl Zeiss</td>
			<td>F170</td>
			<td></td>
		</tr>
		<tr>
			<td>Suture needle</td>
			<td>Allgaier</td>
			<td>1245</td>
			<td>For anesthesia</td>
		</tr>
		<tr>
			<td>Temperaure controller</td>
			<td>CWE,INC.</td>
			<td>TC-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Transducer x 2</td>
			<td>Adinstruments</td>
			<td>MLT0699</td>
			<td>Connects to BP and ICP monitor</td>
		</tr>
		<tr>
			<td>Ventilator</td>
			<td>Ugo Basile</td>
			<td>7025</td>
			<td></td>
		</tr>
		<tr>
			<td>Veterinary clipper</td>
			<td>Aesculap</td>
			<td>GT421</td>
			<td></td>
		</tr>
		<tr>
			<td>3-pronged Blair retractor, 13.5 cm</td>
			<td>Agnthos</td>
			<td>17022--13</td>
			<td></td>
		</tr>
		<tr>
			<td>Blunt Alm retractor</td>
			<td>F.S.T</td>
			<td>17008-07</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved forceps, 12 cm x 2</td>
			<td>F.S.T</td>
			<td>11001-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle holder, 13 cm</td>
			<td>F.S.T</td>
			<td>12001-13</td>
			<td></td>
		</tr>
		<tr>
			<td>Straight Dumont forceps, 11 cm</td>
			<td>F.S.T</td>
			<td>11252-00</td>
			<td></td>
		</tr>
		<tr>
			<td>Straight Halsted-Mosquito hemostat x 2</td>
			<td>F.S.T</td>
			<td>13008-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Straight Iris scissor, 9 cm</td>
			<td>F.S.T</td>
			<td>14090-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Straight Vannas scissor, 10.5 cm</td>
			<td>F.S.T</td>
			<td>15018-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Absorpable swabs</td>
			<td>Kettenbach</td>
			<td>31603</td>
			<td></td>
		</tr>
		<tr>
			<td>Black silk thread, 4-0, 5 x 15 cm</td>
			<td>V&#246;mel</td>
			<td>14757</td>
			<td></td>
		</tr>
		<tr>
			<td>Bone wax</td>
			<td>Aesculap</td>
			<td>1029754</td>
			<td></td>
		</tr>
		<tr>
			<td>Carbomer eye gel 2 mg/g</td>
			<td>Paranova</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton swab</td>
			<td>Heinz Herenz</td>
			<td>WA-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton tipped applicator x 4</td>
			<td>Selefa</td>
			<td>120788</td>
			<td></td>
		</tr>
		<tr>
			<td>Hypodermic needle, 23 G x2</td>
			<td>KD Medical</td>
			<td>900284</td>
			<td>Connects to stopcock. Remove distal end</td>
		</tr>
		<tr>
			<td>Hypodermic needle, 23 G x3</td>
			<td>KD Medical</td>
			<td>900284</td>
			<td>Remove distal end. 2 connects to stopcock, 1 to syringe</td>
		</tr>
		<tr>
			<td>ICP probe:</td>
			<td>Homemade</td>
			<td></td>
			<td>Made of the following:</td>
		</tr>
		<tr>
			<td>Polythene tubing, 20 mm</td>
			<td>Smiths medical</td>
			<td>800/100/200</td>
			<td>Inner diameter (ID): 0.58 mm, Outer diameter (OD): 0.96 mm.</td>
		</tr>
		<tr>
			<td>Silicone tubing, 10 mm</td>
			<td>Fisher</td>
			<td>15202710</td>
			<td>ID: 0.76 mm, OD: 2.4 mm.</td>
		</tr>
		<tr>
			<td>Silicone tubing, 2 mm</td>
			<td>Fisher</td>
			<td>11716513</td>
			<td>ID: 1.0 mm, OD: 3.0 mm.</td>
		</tr>
		<tr>
			<td>Micro hematocrit tubes</td>
			<td>Brand</td>
			<td>7493 11</td>
			<td></td>
		</tr>
		<tr>
			<td>OP-towel, 45 cm x75 cm</td>
			<td>M&#246;lnlycke</td>
			<td>800430</td>
			<td></td>
		</tr>
		<tr>
			<td>PinPort adapter, 22 G</td>
			<td>Instech</td>
			<td>PNP3F22</td>
			<td></td>
		</tr>
		<tr>
			<td>PinPort injector</td>
			<td>Instech</td>
			<td>PNP3M</td>
			<td></td>
		</tr>
		<tr>
			<td>Polythene tubing, 2 x 20 cm</td>
			<td>Smiths medical</td>
			<td>800/100/200</td>
			<td>Connects to syringe. ID: 0.58 mm, OD: 0.96 mm.</td>
		</tr>
		<tr>
			<td>Rubberband</td>
			<td>Unknown</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel, 10 blade</td>
			<td>Kiato</td>
			<td>23110</td>
			<td></td>
		</tr>
		<tr>
			<td>Spinalneedle, 25 G x 3.5&#39;&#39;</td>
			<td>Braun</td>
			<td>5405905-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Stopcock system, Discofix x 2</td>
			<td>Braun</td>
			<td>16494C</td>
			<td>Connects to transducer</td>
		</tr>
		<tr>
			<td>Suture, 4-0, monofil, non-resorbable x 3</td>
			<td>Ethicon</td>
			<td>EH7145H</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe, 1 mL</td>
			<td>BD Plastipak</td>
			<td>1710023</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe, luer-lock, 10 mL x 4</td>
			<td>BD Plastipak</td>
			<td>305959</td>
			<td>Connects to transducer</td>
		</tr>
		<tr>
			<td>Tissue adhesive glue</td>
			<td>3M</td>
			<td>1469SB</td>
			<td></td>
		</tr>
		<tr>
			<td>0.5% Chlorhexidine spirit</td>
			<td>Faaborg Pharma</td>
			<td>210918</td>
			<td></td>
		</tr>
		<tr>
			<td>Carprofen 50 mg/mL</td>
			<td>ScanVet</td>
			<td>43715</td>
			<td>Diluted 1:10</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Baxter</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Isotonic saline</td>
			<td>Amgros</td>
			<td>16404</td>
			<td></td>
		</tr>
		<tr>
			<td>Lidocaine-Adrenaline 10 mg/5 &#181;g/mL</td>
			<td>Amgros</td>
			<td>16318</td>
			<td></td>
		</tr>
	</tbody>
</table>

establishing a subarachnoid hemorrhage rat model induced by autologous blood injection

Source: B&#246;mers, J. et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/62567">Pre-Chiasmatic, Single Injection of Autologous Blood to Induce Experimental Subarachnoid Hemorrhage in a Rat Model.</a> J. Vis. Exp. (2021)
This video demonstrates the procedure for establishing a subarachnoid hemorrhage rat model. An anesthetized rat is implanted with probes to measure intracranial pressure and cerebral blood flow. The target site is identified, and the needle is inserted at a pre-chiasmatic position. Autologous blood is then injected into the subarachnoid space, increasing intracranial pressure and decreasing cerebral blood flow, establishing the model.

<img alt="Figure 1" src="/files/ftp_upload/23119/23119_Figure1.jpg" /> 
Figure 1: Overview of the setup before induction of SAH. From the top of the picture, note that the 1) injection needle, 2) laser-Doppler probe, and 3) the ICP probe are all in place.
<img alt="Figure 2" src="/files/ftp_upload/23119/23119_Figure2.jpg" /> 
Figure 2: Sample trace following intrathecal injection. The red graph shows the blood pressure in mmHg. The blue graph shows the ICP in mmHg and the green graph shows the CBF in the arbitrary unit FU. The spike in ICP is the result of blood injection. Notice that this results in a drop in the CBF &#62; 50% of baseline for more than 5 min. The ICP rise furthermore results in a small rise in blood pressure which normalizes within seconds.

Establishing a Subarachnoid Hemorrhage Rat Model Induced by Autologous Blood Injection

Source: B&#246;mers, J. et al. Pre-Chiasmatic, Single Injection of Autologous Blood to Induce Experimental Subarachnoid Hemorrhage in a Rat Model. J. Vis. ...

Take an anesthetized rat with its skull exposed and a probe implanted to measure intracranial pressure.
Using a reference point on the skull, mark the target site and drill a hole.
Remove the bone fragments and fill the hole with bone wax.
Drill another hole. Locate a blood vessel and secure a probe to measure cerebral blood flow.
Take a needle and insert it into the brain until the skull is felt.
Retract slightly to ensure its placement in front of the optic chiasm, where major blood vessels are located.
Fill the needle chamber with fresh blood drawn from the rat&#39;s tail and connect the syringe containing blood.
Inject the blood into the fluid-filled subarachnoid space between the outer and inner brain membranes, which increases intracranial pressure, causing blood vessel constriction.
This leads to a decreased cerebral blood flow, establishing a subarachnoid hemorrhage model.

establishing-subarachnoid-hemorrhage-rat-model-induced-autologous

Nanyang Technological University

Research

JoVE Encyclopedia of Experiments

Neuroscience

Neuropathology

Cerebrovascular Diseases

48 vistas. Source: Bömers, J. et al. Pre-Chiasmatic, Single Injection of Autologous Blood to Induce Experimental Subarachnoid Hemorrhage in a Rat Model. J. Vis. Exp. (2021) This video demonstrates the procedure for establishing a subarachnoid hemorrhage rat model. An anesthetized rat is implanted with probes to measure intracranial pressure and cerebral blood flow. The target site is identified, and the needle is inserted at a pre-chiasmatic position. Autologous blood is then injected into the subara...

Video: Establishing a Subarachnoid Hemorrhage Rat Model Induced by Autologous Blood Injection - Concepto

Intracranial Pressure Monitoring In Nontraumatic Intraventricular Hemorrhage Rodent Model

Survivors of intraventricular hemorrhage are often left with significant long-term memory impairment; thus, research utilizing intraventricular hemorrhage animal models is essential. In this study, we sought out ways to measure intracranial pressure, mean arterial pressure, and cerebral perfusion pressure during nontraumatic intraventricular hemorrhage in rats. The experimental design included three Sprague Dawley groups: sham, standard 200 &#181;l intraventricular hemorrhage, and vehicle control groups. By introducing an intraparenchymal fiberoptic pressure sensor, precise intracranial pressure measurements were obtained in all groups. Cerebral perfusion pressures were calculated with the knowledge of intracranial pressure and mean arterial pressure values. As expected, the intraventricular hemorrhage and vehicle control groups both experienced a rise in the intracranial pressure and subsequent decline in cerebral perfusion pressure during intraventricular injection of autologous blood and artificial cerebrospinal fluid, respectively. The addition of an intraparenchymal fiberoptic pressure sensor is beneficial in monitoring precise intracranial pressure changes.

Monitoring intracranial pressure in rodent&#160;models of nontraumatic intraventricular hemorrhage is not common in the current literature. Herein, we demonstrate a technique for measuring intracranial pressure, mean arterial pressure, and cerebral perfusion pressure during intraventricular hemorrhage in a rat&#160;animal model.

Monitorización de la presión intracraneal en un modelo de roedor con hemorragia intraventricular no traumática

Survivors of intraventricular hemorrhage are often left with significant long-term memory impairment; thus, research utilizing intraventricular hemorrhage ...

Investigación

JoVE Journal

Neurociencias

Visualización de la contractilidad del pericito cerebral post-hemorragia subaracnoidea

Imaging Vital and Non-vital Brain Pericytes in Brain Slices following Subarachnoid Hemorrhage

Pericytes are crucial mural cells situated within cerebral microcirculation, pivotal in actively modulating cerebral blood flow via contractility adjustments. Conventionally, their contractility is gauged by observing morphological shifts and nearby capillary diameter changes under specific circumstances. Yet, post-tissue fixation, evaluating vitality and ensuing pericyte contractility of imaged brain pericytes becomes compromised. Similarly, genetically labeling brain pericytes falls short in distinguishing between viable and non-viable pericytes, particularly in neurologic conditions like subarachnoid hemorrhage (SAH), where our preliminary investigation validates brain pericyte demise. A reliable protocol has been devised to surmount these constraints, enabling simultaneous fluorescent tagging of both functional and non-functional brain pericytes in brain sections. This labeling method allows high-resolution confocal microscope visualization, concurrently marking the brain slice microvasculature. This innovative protocol offers a means to appraise brain pericyte contractility, its impact on capillary diameter, and pericyte structure. Investigating brain pericyte contractility within the SAH context yields insightful comprehension of its effects on cerebral microcirculation.

Autor destacado: Imágenes de pericitos post-hemorragia subaracnoidea en roedores 

The preliminary inquiry confirms that subarachnoid hemorrhage (SAH) causes brain pericyte demise. Evaluating pericyte contractility post-SAH requires differentiation between viable and non-viable brain pericytes. Hence, a procedure has been developed to label viable and non-viable brain pericytes concurrently in brain sections, facilitating observation using a high-resolution confocal microscope.

Establishing a Subarachnoid Hemorrhage Rat Model Induced by Autologous Blood Injection

Transcripción

Establishing a Subarachnoid Hemorrhage Rat Model Induced by Autologous Blood Injection

Monitorización de la presión intracraneal en un modelo de roedor con hemorragia intraventricular no traumática

Obtención de imágenes de pericitos cerebrales vitales y no vitales en cortes de cerebro después de una hemorragia subaracnoidea

Obtención de imágenes de pericitos cerebrales vitales y no vitales en cortes de cerebro después de una hemorragia subaracnoidea

Obtención de imágenes de pericitos cerebrales vitales y no vitales en cortes de cerebro después de una hemorragia subaracnoidea