This study was financially supported by the National Science Foundation of China (81471181 and 81870933) and the Opening Lab Program of Guangzhou Medical University (0506308) to Y Jiang, and by the National Science Foundation of China (81701471) and the Scientific Program of Guangzhou Municipal Health Commission (20191A011083) to Z Qiu, and by the National Science Foundation of China (81501009) to L Wu.

The protocol was reviewed and approved by the Institutional Animal Care and Use Committee of the Second Affiliated Hospital of Guangzhou Medical University. Rats were provided by the Animal Center of Southern Medical University. The experimental design is shown in Figure 1.
1. Animal and instruments
<ol>
	<li>Use 8-week-old male Sprague-Dawley rats weighing 250 &#177; 10 g.</li>
	<li>House the rats for at least 7 days before surgery under controlled environmental conditions with an ambient temperature of 25 &#176;C, relative humidity of 65%, and a 12/12-h light-dark cycle.</li>
	<li>Provide food and water with no limit.</li>
	<li>Prepare the instruments (Figure 2A-E).</li>
</ol>
2. Inject the blood in the pons
<ol>
	<li>Weigh the rats again 3 days before surgery to select rats with suitable body weight for the experiments.</li>
	<li>During the 3 days before modeling, train the rats to walk on balance beam 3 times per day to ensure normal rats could pass the balance beam without pause.</li>
	<li>Preheat the heating pad to 37 &#176;C before anesthesia.</li>
	<li>Attach a microdrill to the holder on the stereotaxic frame.</li>
	<li>Inject the rats with 50 mg/kg ketamine and 5 mg/kg xylazine intraperitoneally. Wait until there is no toe-pinch response.</li>
	<li>Transport the rat into the stereotaxic frame in a prone position. Put the ear bars above the ear canal to secure the head. Fix the skull in a horizontal position to avoid skewing of the injection (Figure 2F).</li>
	<li>Maintain anesthesia by isoflurane (97.5% oxygen and 2.5% isoflurane) through a stereotaxic nose cone with an inlet and outlet port.</li>
	<li>Use eye ointment to keep the cornea moist.</li>
	<li>Shave the hair above the skull with a micro shaver.</li>
	<li>Draw a 3 cm mid-line with a marker pen in the skull from the line of the bilateral lateral canthus to 0.5 cm behind the posterior fontanelle, as shown in Figure 2F.</li>
	<li>Apply Entoiodine surgical scrub in a circular fashion, starting at the middle of the marked mid-line and rotating outward.</li>
	<li>Place the surgical drape.</li>
	<li>Make an incision with a scalpel along the marked mid-line.</li>
	<li>Use a cotton swab to remove any potential blood.</li>
	<li>Place a piece of forceps on each side of the scalp flap to expose the skull (Figure 2G).</li>
	<li>Dip a cotton swab in 0.9% saline and gently remove the connective tissues from the skull bone to avoid the connective tissues getting caught in the microdrill.</li>
	<li>Mark the central point of the bregma as the origin point via a marker pen.</li>
	<li>Perform a craniotomy (1 mm in diameter) using a microdrill at AP-9.0 mm, Lat 0 mm (Figure 1B and Table 1). Proceed carefully because this point is very close to the venous sinus (Figure 2G).</li>
	<li>Remove the microdrill from the stereotaxic frame.</li>
	<li>Put a 100 &#181;L Hamilton syringe into the stereotaxic holder (Figure 2K). Turn on the injection pump switch, click the Rapid Inhalation button, aspirate heparin solution (12500 U diluted in 100 mL of saline) to 100 &#181;L and then drain it completely to prevent blood clotting too fast.</li>
	<li>Apply chlorhexidine and 75% alcohol surgical scrub to the whole tail from root to tip at least 3 times to disinfect the skin, soften the horniness, and dilate the tail vein to increase the success rate of injection.</li>
	<li>Attach a scalp acupuncture to a 1 mL syringe.</li>
	<li>Insert the scalp acupuncture into a lateral tail vein 3 cm from the tail tip and take 150 &#181;L of blood (Figure 2H).</li>
	<li>Remove the scalp acupuncture from the 1 mL syringe.</li>
	<li>Transfer the blood into a tube (Figure 2I). 
	NOTE: Transfer quickly to prevent blood clotting.</li>
	<li>Change the surgical gloves.</li>
	<li>Choose Withdraw mode, set the volume to 100 &#181;L, set the speed at 200 &#181;L/min, click the RUN button, aspirate 100 &#181;L of blood into the Hamilton syringe (Figure 2J).</li>
	<li>Choose Infuse mode, set the speed at 1 &#181;L/min.</li>
	<li>Advance the syringe until the tip reaches 9.2 mm below the surface of the brain (Figure 1C and Figure 2L).</li>
	<li>Click the Run button, inject the first 10 &#181;L of blood at a speed of 1 &#181;L/min (Table 1).</li>
	<li>Stop the injection and leave the syringe in position for 20 min to prevent blood from flowing into the subarachnoid space.</li>
	<li>Retract the syringe until the tip arrives at 9.0 mm below the surface of the brain.</li>
	<li>Restart injection at the same speed of 1 &#181;L/min until the residual blood has been injected completely.</li>
	<li>Leave the syringe in position for 10 min to avoid blood backflow.</li>
	<li>Remove the syringe from the brain slowly.</li>
	<li>Use bone cement to cover the craniotomy hole.</li>
	<li>When the cement dries, suture the wound with 4-0 polyamide suture filament. After sewing 3 or 4 stiches, tie 2-1-1 standard surgical knots.</li>
	<li>To prevent infection, inject the rat with penicillin (0.25 mL, 80 IU diluted in 4 mL saline) intraperitoneally.</li>
	<li>Inject the rats subcutaneously with Butorphanol tartrate (2.5 mg/kg) and repeat the injection every 2 hours for relieving postoperative pain until 24 hours after surgery.</li>
	<li>Observe the rat every 15 min until it fully recovers from anesthesia. Return it to the original cage, with a heating pad underneath the cage. Provide the rat free access to food and water until sacrifice.</li>
</ol>
3. Behavioral tests
NOTE: Perform behavioral tests on Day 1, Day 3, Day 7, and Day 14 after modeling, including the balance beam test, limb placement test, and the modified Voetsch neuroscore.
<ol>
	<li>Balance beam test11. 
	&#8203;NOTE: This is a test specifically for examining the sensorimotor function of the hindlimb.
	<ol>
		<li>Prepared the apparatus to ensure that the beam (3 cm wide &#215; 70 cm long) is 20 cm above the floor.</li>
		<li>Put a dark box at the back end of the beam with a narrow entryway.</li>
		<li>Place a white noise generator and a bright light source, which are used to motivate the rat to traverse the beam and enter the goal box, at the head end of the beam.</li>
		<li>Stop the noise and light when the animal enters the goal box. Record the latency from head end to the goal box (in seconds) and the performance of hindlimb during traversing the beam.</li>
		<li>Record the performance score as following: 0, balances with steady posture; 1, grips side of beam; 2, snuggles the beam with 1 hindlimb falling off; 3, snuggles the beam with 2 limbs falling off, or spinning around the beam for more than 60 s; 4, attempts to balance on beam &#62;40 s, but falls off; 5, attempts to balance on beam &#62;20 s, but falls off; and 6, falls off, with no attempt to balance or balancing on beam &#60;20 s.</li>
	</ol>
	</li>
	<li>Limb placement test 
	&#8203;NOTE: The limb placement test examines 3 independent stimuli of vision, touch, and proprioception with 6 parameters, to evaluate the sensorimotor function of rat. The total neurological function score ranged from 0 to 12. Before the test, rat should be habituated to handling.
	<ol>
		<li>Hold the rat by the back and place it on the table slowly from a height of 10 cm. Normally, the rat would stretch forelimbs and place them on the table.</li>
		<li>Grab the rat by the back and hold it facing the edge of the table. The reaction of forelimbs is observed. A normal rat would place the forelimbs on the table.</li>
		<li>Let the rat grasp the edge of the table. A normal rat would use the chin to prevent the nose and nose hair from touching the table edge.</li>
		<li>Put the rat on the table, push the rat with a gentle lateral pressure behind the rat&#39;s shoulder toward the edge of the table, and observe the placement of the forelimbs and hindlimbs. A normal rat would grasp the edge with its forelimbs and hindlimbs.</li>
		<li>Place the rat on the table with the face toward the edge, and gently push it from the back to the edge. A normal rat would grasp the edge with its forelimbs.</li>
		<li>Place the rat on the table with its back to the edge and push it by the back to the edge of the table. Observe the reaction of the hindlimbs. A normal rat would grasp the edge with its hindlimbs.</li>
		<li>Assess the placement of the forelimbs or hindlimbs to the table edge. Record the scores as follows: 0, no placement; 1, unfinished and/or delayed placement; 2, immediate, complete placement.</li>
	</ol>
	</li>
	<li>The modified Voetsch neuroscore8 
	NOTE: The modified Voetsch neuroscore is a vertebrobasilar scale score of sensorimotor ability, containing 14 parameters: head movement, activity, hearing, pain reflex, corneal reflex, proprioception, neck sensation, exploration, circling, axial torso sensation, 4-limb movement, forelimb movement, climbing, and beam walk. The score for each parameter ranges from 0 (complete neurological deficit) to 3 (no neurological deficit). The total neurological function score ranges from 0 to 42.
	<ol>
		<li>Place the rat on the table (50 cm long &#215; 35 cm wide) and allow it to move around for five minutes. Observe the spontaneous movement of its head: moves in all dimensions, 3; prefers one side, 2; only movement to 1 side, 1; flexed to unilateral side, 0.</li>
		<li>Place the rat on the table (50 cm long &#215; 35 cm wide) and allow it to move around for five minutes. Activity is defined as: fully responsive, 3; moderately responsive, 2; minimally responsive, 1; coma, 0.</li>
		<li>Observe the craniocaudal circling: turns bilaterally, 3; prefers 1 side, 2; only to 1 side, 1; fallen to 1 side, 0.</li>
		<li>Evaluate the forelimbs movement: equal and bilateral movement, 3; slight asymmetry, 2; great asymmetry, 1; paresis, 0.</li>
		<li>Evaluate the 4-limbs movement: equal and bilateral movement, 3; slight asymmetry, 2; great asymmetry, 1; paresis, 0.</li>
		<li>When the rat is stationary, perform an ear pinch to stimulate the auricles and test the pain reflex: moves away quickly and symmetrically from stimulus, 3; moves away slowly or asymmetrically from stimulus, 2; shows some movement in response to pain, 1; no reaction, 0.</li>
		<li>When the rat is stationary, make a noise on either side of the rat&#39;s body to test hearing: fingers rubbing, 3; snap of fingers, 2; loud clapping, 1 and no startling, 0.</li>
		<li>To check the proprioception, touch the rat&#39;s vibrissae on both sides respectively with a blunt stick and observe its response to the stimulus: react on touch, 3; diminished reaction on one side, 2; diminished on both sides, 1; absent, 0.</li>
		<li>To evaluate the torso axial sensation, place a blunt stick on either side of the rat&#39;s body and observe its response to the stimulus: brisk and symmetrical reaction to stimuli, 3; slightly diminished or asymmetrical reaction, 2; greatly diminished and asymmetrical reaction, 1 and no reaction, 0.</li>
		<li>To test the corneal reflex, hold the rat and quickly touch the cornea on both sides with a cotton swab: both eyes close quickly, 3; diminished reflex on one side, 2; diminished on both sides, 1; absent, 0.</li>
		<li>To check the sensation of the neck, hold the rat and use a blunt stick to touch the neck: reacts to touch actively, 3; slow reaction to touch, 2; greatly diminished reaction, 1; no reaction, 0.</li>
		<li>To observe exploration, use a light dark box. 
		&#8203;NOTE: Two thirds of the box should be open and lit, while the rest of it is covered and dark. A 7 cm door connects the two compartments.</li>
		<li>Place the rat in the light compartment for 5 min and then the dark compartment for another 5 min, allow it to move around and record the rat&#39;s activities. When 4 limbs are placed in one room, record it as an entrance. Record scores as follows: reaches the light and dark compartments actively, 3; reaches 1 compartment, 2; moves slowly only after stimulus, 1; and no movement, 0.</li>
		<li>To check the climbing ability, place the rat at the bottom of a 20 cm wide, 70 cm long plane with a 30-degree angle to the horizontal ground. Record scores as follows: able to climb to top, 3; impaired climbing, 2; stationary grip, 1; and falls immediately, 0.</li>
		<li>To examine the beam walk, put the rat on one end of a 3 cm wide, 70 cm long beam which is 20 cm above the ground, record scores as following: explores the entire beam, 3; explores part of the beam, 2; some movement and falls, 1; no movement, 0.</li>
		<li>Calculate the points.</li>
	</ol>
	</li>
</ol>
4. Hemorrhage confirmation by MRI
<ol>
	<li>Perform the MRI scan at 24 h post-surgery.</li>
	<li>Anesthetize the rat with isoflurane (5% for induction, 1%- 1.5% for maintenance).</li>
	<li>Secure the rat&#39;s head in the rat brain array coil combined with a transmit-only volume coil.</li>
	<li>Place the coil together with the rat in the MRI scanner. Secure the rat within the cradle via the tooth and ear bars.</li>
	<li>Use a closed-circuit thermal jacket to maintain the rat&#39;s body temperature at 37 &#177; 0.5 &#176;C during MRI scanning.</li>
	<li>Perform a pilot sequence to ensure correct geometry.</li>
	<li>Apply a fast-spin echo sequence to collect T2-weighted scans. Set the parameters as follows: echo time (TE), 132 ms; repetition time (TR), 2,500 ms; acquisition matrix, 148 &#215; 148; field of view, 100 mm &#215; 100 mm; 12 slices; 1.5 mm thick.</li>
	<li>Return the rat to the cage.</li>
</ol>
5. Hemorrhage confirmation by gross anatomy
NOTE: Sacrifice the rats at the designated timepoint, 24 h and 14 d after surgery (Figure 1D).
<ol>
	<li>Anesthetize the rat with 5% isoflurane until loss of consciousness. Then, euthanize it with carbon dioxide (CO2) (20-30% of the volume of the cage per minute).</li>
	<li>Confirm death using the following signs: no chest rising and falling, no palpable heartbeat, no response to toe pinch, poor mucosa color, color change, or opacity in eyes.</li>
	<li>Perform cervical dislocation.</li>
	<li>Secure the rat by taping the paws on a sterile platform. Create a midline incision from the cervix to expose the hypogastrium to thorax and liver. Make a lateral incision from the upper sternal margin along the clavicle to the far left and another lateral cut from the xiphoid along the diaphragm to the far left, to expose heart. Lift the ribcage flap and fix it to the platform with a pin.</li>
	<li>Connect the end of a needle (27 G) to a perfusion pump containing 4 &#176;C saline. Advance the needle tip into the heart along the left edge of the left ventricle to avoid entering the atrium. Turn on the perfusion pump to ensure the tip is in the left ventricle, then make a cut in the right atrium.</li>
	<li>Turn off the perfusion pump when the liquid drained out of the right atrium turns colorless and the liver turns white. This procedure needs approximately 100 mL 4 &#176;C saline.</li>
	<li>Decapitate the rat and harvest the whole brain using scissors and forceps. Remove any moisture from the brain surface with blotting paper.</li>
	<li>Keep the whole brain at -80 &#176;C for 1 min. 
	NOTE: This step could be skipped if the brain can be cut without freezing.</li>
	<li>Lay the brain into the coronal rat brain matrix with the dorsal side up.</li>
	<li>Insert a 0.21 mm thick stainless-steel blade into the hemorrhage center according to the hole in the surface of the brain.</li>
	<li>Insert other blades into the brain at an interval of 2 mm.</li>
	<li>Immerse the brain sections in 10 mL of 4% paraformaldehyde (PFA) solution for 24 h at 4 &#176;C. Rinse them with 0.01 mmol/L phosphate buffered saline (PBS).</li>
	<li>Organize the sections from rostral to caudal and image them.</li>
</ol>
6. Paraffin section and hematoxylin and eosin (HE) staining
<ol>
	<li>Fix the brain with 4% PFA solution for at least 24 h at room temperature. The volume of PFA should be 5-10 times of brain volume.</li>
	<li>Cut the brain from the hemorrhage center into two pieces via blade and coronal rat brain matrix.</li>
	<li>Make paraffin-embedded tissue blocks.</li>
	<li>Section the paraffin-embedded tissue block coronally in 40 &#181;m thickness slides on a microtome, cut 4 sections in succession, starting from hemorrhage center, and float them in a 40 &#176;C water bath.</li>
	<li>Mount the 40 &#181;m sections (8 slides in total for one brain) onto clean glass slides and air dry overnight at room temperature.</li>
	<li>Bake the slides at 56 &#176;C for 1 h.</li>
	<li>Wash for 3 min in xylene, 3 times.</li>
	<li>Dip sections 30 times in 100% ethanol, 30 more times in 100% ethanol, 30 times in 95% ethanol, and 30 more times in 95% ethanol.</li>
	<li>Rinse in tap water until clear.</li>
	<li>Immerse the sections in Hematoxylin for about 10 min.</li>
	<li>Rinse in the tap water until clear.</li>
	<li>Immerse sections in eosin stain for 30 s.</li>
	<li>Dehydrate slides with 3-4 dips 95% ethanol, 3-4 dips in 100% ethanol, 100% Ethanol for 1 minute, and 10 dips in 100% ethanol + xylene (1:1).</li>
	<li>Clean sections with xylene for 1 min, 2 times.</li>
	<li>Mount using non-aqueous mounting medium and a coverslip.</li>
	<li>Dry the sections in the air overnight at room temperature, then san them.</li>
</ol>
7. Statistics
<ol>
	<li>Use GraphPad Prism 6.0 to calculate Student&#39;s t-test or Mann Whitney U test. 
	NOTE: All data should be expressed as mean &#177; SE. Differences between two groups are determined with a two-tailed Student&#39;s t-test or Mann Whitney U test. P&#60;0.05 is defined as statistical significance.</li>
</ol>

<ol>
	<li>Charidimou, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Brain+hemorrhage+recurrence,+small+vessel+disease+type,+and+cerebral+microbleeds:+A+meta-analysis.">Brain hemorrhage recurrence, small vessel disease type, and cerebral microbleeds: A meta-analysis.</a> Neurology. 89 (8), 820-829 (2017).</li><li>Tao, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+brainstem+hemorrhage+model+by+autologous+blood+infusion+in+Rat:+White+Matter+Injury,+Magnetic+Resonance+Imaging,+and+Neurobehavioral+features.">A novel brainstem hemorrhage model by autologous blood infusion in Rat: White Matter Injury, Magnetic Resonance Imaging, and Neurobehavioral features.</a> Journal of Stroke and Cerebrovascular Diseases. 25 (5), 1102-1109 (2016).</li><li>Ichimura, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Surgical+treatment+for+primary+brainstem+hemorrhage+to+improve+postoperative+functional+outcomes.">Surgical treatment for primary brainstem hemorrhage to improve postoperative functional outcomes.</a> World Neurosurgery. 120, 1289-1294 (2018).</li><li>Behrouz, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Prognostic+factors+in+pontine+haemorrhage:+A+systematic+review.">Prognostic factors in pontine haemorrhage: A systematic review.</a> European Stroke Journal. 3 (2), 101-109 (2018).</li><li>Guo, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Brainstem+iron+overload+and+injury+in+a+rat+model+of+brainstem+hemorrhage.">Brainstem iron overload and injury in a rat model of brainstem hemorrhage.</a> Journal of Stroke and Cerebrovascular Diseases. 29 (8), 104956 (2020).</li><li>Chung, Y., Haines, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Experimental+brain+stem+surgery.">Experimental brain stem surgery.</a> Neurosurgery Clinics of North America. 4 (3), 405-414 (1993).</li><li>Lekic, T., Tang, J., Zhang, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+rat+model+of+pontine+hemorrhage.">A rat model of pontine hemorrhage.</a> Acta Neurochirurgica Supplement. 105, 135-137 (2008).</li><li>Lekic, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+the+hematoma+consequences,+neurobehavioral+profiles,+and+histopathology+in+a+rat+model+of+pontine+hemorrhage.">Evaluation of the hematoma consequences, neurobehavioral profiles, and histopathology in a rat model of pontine hemorrhage.</a> Journal of Neurosurgery. 118 (2), 465-477 (2013).</li><li>Shrestha, B. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rat+brainstem+hemorrhage+model:+Key+points+to+success+in+modeling.">Rat brainstem hemorrhage model: Key points to success in modeling.</a> World Neurosurgery. 117, 106-116 (2018).</li><li>Luo, M., Tang, X., Zhu, J., Qiu, Z., Jiang, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Establishment+of+acute+pontine+infarction+in+rats+by+electrical+stimulation.">Establishment of acute pontine infarction in rats by electrical stimulation.</a> Journal of Visualized Experiments. (162), (2020).</li><li>Wu, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Keep+warm+and+get+success:+The+role+of+postischemic+temperature+in+the+mouse+middle+cerebral+artery+occlusion+model.">Keep warm and get success: The role of postischemic temperature in the mouse middle cerebral artery occlusion model.</a> Brain Research Bulletin. 101, 12-17 (2014).</li></ol>

No conflicts of interest.

In the present study, we provided a protocol to generate a massive pontine hemorrhage rat model. This model can be used for the research on the pathophysiological mechanism and prognosis of massive pontine hemorrhage.
Throughout the experiment, 25 rats were used, of which only one died. The verification of MRI, gross anatomy, and the HE staining indicated that this method had a very low mortality rate and a high success rate. To establish massive pontine hemorrhage model, two problems must be ...

Intracerebral hemorrhage accounts for one-fifth of stroke patients. The prognosis of intracerebral hemorrhage depends on the speed, volume, and location of bleeding1,2. Compared to the forebrain hemorrhage, the brainstem hemorrhage has higher mortality and morbidity3. About 40% of brainstem hemorrhage occurs in the pons4. The etiology and pathophysiology of pontine hemorrhage are quite different and less studied than forebrain hemorrhage5.
There are two kinds of pontine hemorrhage animal models. One is spontaneous hemorrhage model induced by infusion of bacterial collagenase in the pons6,7,8. The biggest advantage of this model is that the bleeding is spontaneous. However, collagenase can only induce a small volume of pontine hemorrhage. Besides, collagenase might cause other injuries to the brain. The other model is induced by stereotactic injection of autologous blood into the pons9. The advantage of this model is that it is easy to master with a high success rate. Theoretically, researchers could inject any volume of blood into any location of pons. However, due to the back-leakage through the needle route, the injected volume is limited. Recently, the double-injection method has been promoted to reduce the back-leakage9. This method injects autologous blood twice with a 20-minute interval between the injections. The double-injection method is applied to induce mild (30 &#181;L) and moderate (60 &#181;L) pontine hemorrhage but not massive pontine hemorrhage. In the clinic, the majority of pontine hemorrhage patients with poor prognosis have massive hemorrhage (more than 10 mL).
In the previous study, we provided a protocol to establish a pontine ischemic stroke model in rat10. In this study, we modify the existing dual injection method, and provide a detailed protocol to induce massive pontine hemorrhage in a rat by dual injection of 100 &#181;L autologous blood at two different locations in the pons.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>100ml Saline solution</td>
			<td>Guangdong yixiang</td>
			<td>191222201 C1</td>
			<td>Preparing heparin diluent</td>
		</tr>
		<tr>
			<td>100&#956;l Microinjector</td>
			<td>Shanghai Gaoge</td>
			<td></td>
			<td>Injection of autologous blood</td>
		</tr>
		<tr>
			<td>1ml Syringe</td>
			<td>Jiangsu Zhiyu</td>
			<td>20191014</td>
			<td>Withdraw autologous blood from the tail vein</td>
		</tr>
		<tr>
			<td>75% Alcohol</td>
			<td>Shandong Lierkang</td>
			<td></td>
			<td>Disinfection of rat tail</td>
		</tr>
		<tr>
			<td>Adhesive tape</td>
			<td>Shanghai Jinzhong</td>
			<td></td>
			<td>Surgicl instruments</td>
		</tr>
		<tr>
			<td>Animal anesthesia system</td>
			<td>RWD</td>
			<td>R510-31S-6</td>
			<td>Inducing and maintaining anesthesia</td>
		</tr>
		<tr>
			<td>Balance beam</td>
			<td>Jiangsu Saiangsi</td>
			<td></td>
			<td>For neurological deficit scores</td>
		</tr>
		<tr>
			<td>Blades</td>
			<td>Shanghai Feiying</td>
			<td>74-C</td>
			<td>For gross anatomy</td>
		</tr>
		<tr>
			<td>Bone cement</td>
			<td>Shanghai Xinshiji</td>
			<td>20180306</td>
			<td>Surgicl instruments</td>
		</tr>
		<tr>
			<td>Brain tank</td>
			<td>Shenzhen LEIYEA</td>
			<td></td>
			<td>For gross anatomy</td>
		</tr>
		<tr>
			<td>Butorphanol tartrate</td>
			<td>Jiangsu Hengrui</td>
			<td></td>
			<td>For pain management</td>
		</tr>
		<tr>
			<td>Electric cranial drill</td>
			<td>Nanjing&#160; Darwin biotechnology</td>
			<td>20180090018</td>
			<td>Making a burr hole on the skull</td>
		</tr>
		<tr>
			<td>EP tube</td>
			<td>Nantong Surui</td>
			<td></td>
			<td>Transfer autologous blood</td>
		</tr>
		<tr>
			<td>Erythromycin eye cream</td>
			<td>Yunnan pharmacy</td>
			<td></td>
			<td>Eyes protection</td>
		</tr>
		<tr>
			<td>HE dye liquor</td>
			<td>Solarbio</td>
			<td>G1120</td>
			<td>For HE staining</td>
		</tr>
		<tr>
			<td>Heating&#160;pad</td>
			<td>Dangerous Jungle</td>
			<td>JR01</td>
			<td>Keeping warm</td>
		</tr>
		<tr>
			<td>Heparin sodium injection</td>
			<td>Chengdu Haitong Pharmacal Company</td>
			<td>190701</td>
			<td>Preparing heparin diluent</td>
		</tr>
		<tr>
			<td>IndoPhors</td>
			<td>Guoyao of China</td>
			<td></td>
			<td>Sterilization</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>RWD</td>
			<td>20080701</td>
			<td>Inducing and maintaining anesthesia</td>
		</tr>
		<tr>
			<td>Light dark box</td>
			<td>Jiangsu Saiangsi</td>
			<td></td>
			<td>For neurological deficit scores</td>
		</tr>
		<tr>
			<td>Micro-injection pump</td>
			<td>Baoding Leifu</td>
			<td>TFD03-01-C</td>
			<td>Injection of autologous blood</td>
		</tr>
		<tr>
			<td>MRI system</td>
			<td>Philips</td>
			<td></td>
			<td>Confirmation of infarction in vivo</td>
		</tr>
		<tr>
			<td>Needle holder</td>
			<td>Shanghai Jinzhong</td>
			<td>J32020</td>
			<td>Surgicl instruments</td>
		</tr>
		<tr>
			<td>Penicilin</td>
			<td>Guoyao of China</td>
			<td></td>
			<td>Infection Prevention</td>
		</tr>
		<tr>
			<td>Q-tips</td>
			<td>Jiangxi Songhe</td>
			<td></td>
			<td>Surgicl instruments</td>
		</tr>
		<tr>
			<td>Scalp heedle</td>
			<td>Jiangxi Hongda</td>
			<td>20200313</td>
			<td>Withdraw autologous blood from the tail vein</td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>Shanghai Kaiyuan</td>
			<td>170902</td>
			<td>Surgicl instruments</td>
		</tr>
		<tr>
			<td>Shearing scissors</td>
			<td>Shanghai Jinzhong</td>
			<td>Y00040</td>
			<td>Surgicl instruments</td>
		</tr>
		<tr>
			<td>Stereotaxic apparatus</td>
			<td>RWD</td>
			<td>900-00001-00</td>
			<td>for surgical positioning</td>
		</tr>
		<tr>
			<td>Surgical towel</td>
			<td>Xinxiang Huakangweicai</td>
			<td>20070601</td>
			<td>Surgicl instruments</td>
		</tr>
		<tr>
			<td>Suture needle</td>
			<td>Shanghai Jinzhong</td>
			<td></td>
			<td>Surgicl instruments</td>
		</tr>
		<tr>
			<td>Suture scissors</td>
			<td>Shanghai Jinzhong</td>
			<td>J25041</td>
			<td>Surgicl instruments</td>
		</tr>
		<tr>
			<td>Tissue holding forcepts</td>
			<td>Shanghai Jinzhong</td>
			<td>J31080</td>
			<td>Surgicl instruments</td>
		</tr>
	</tbody>
</table>

massive pontine hemorrhage by dual injection of autologous blood

We provide a protocol to establish a massive pontine hemorrhage model in a rat. Rats weighing about 250 grams were used in this study. One hundred microliters of autologous blood was taken from the tail vein and stereotaxically injected into the pons. The injection process was divided into 2 steps: First, 10 &#181;L of blood was injected into a specific location, anteroposterior position (AP) -9.0 mm; lateral (Lat) 0 mm; vertical (Vert) -9.2 mm, followed by a second injection of the residual blood located at AP -9.0 mm; Lat 0 mm; Vert -9.0 mm with a 20-minute interval. The balance beam test, limb placement test, and the modified Voestch neuroscore were used to evaluate neurological function. Magnetic Resonance Imaging (MRI) was used to assess the volume of hemorrhage in vivo. The symptoms of this model were in line with patients with massive pontine hemorrhage.

A total of 25 animals were used, 3 for control, 6 for 30 &#181;L, 6 for 60 &#181;L, and 10 for 100 &#181;L blood injections. One rat that received a 100 &#181;L injection of autologous blood (1/10) died within 24 hours after surgery.
Behavioral tests were conducted on Day 1, Day 3, Day 7 and Day 14 after surgery. The scores for the control group and blood-injection groups on different timepoints after surgery are presented in <s...

Watch this Scientific Journal Video about Massive Pontine Hemorrhage by Dual Injection of Autologous Blood at JoVE.com

Massive Pontine Hemorrhage by Dual Injection of Autologous Blood

We present a protocol to establish a massive pontine hemorrhage model in a rat via dual injection of autologous blood.

We provide a protocol to establish a massive pontine hemorrhage model in a rat. Rats weighing about 250 grams were used in this study. One hundred ...

massive-pontine-hemorrhage-by-dual-injection-of-autologous-blood

Research

JoVE Journal

Neuroscience

4.0K Views.  The Second Affiliated Hospital of Guangzhou Medical University. We present a protocol to establish a massive pontine hemorrhage model in a rat via dual injection of autologous blood.

Video: Massive Pontine Hemorrhage by Dual Injection of Autologous Blood

Dual Somatic Recordings from Gonadotropin-Releasing Hormone (GnRH) Neurons Identified by Green Fluorescent Protein (GFP) in Hypothalamic Slices

Gonadotropin-Releasing Hormone (GnRH) is a small neuropeptide that regulates pituitary release of luteinizing hormone (LH) and follicle-stimulating hormone (FSH). These gonadotropins are essential for the regulation of reproductive function. The GnRH-containing neurons are distributed diffusely throughout the hypothalamus and project to the median eminence where they release GnRH from their axon terminals into the hypophysiotropic portal system (1). In the portal capillaries, GnRH travels to the anterior pituitary gland to stimulate release of gonadotropins into systemic circulation. GnRH release is not continuous but rather occurs in episodic pulses. It is well established that the intermittent manner of GnRH release is essential for reproduction (2, 3).
Coordination of activity of multiple GnRH neurons probably underlies GnRH pulses. Total peptide content in GnRH neurons is approximately 1.0 pg/cell (4), of which 30% likely comprises the releasable pool. Levels of GnRH during a pulse (5, 6), suggest multiple GnRH neurons are probably involved in neurosecretion. Likewise, single unit activity extracted from hypothalamic multi-unit recordings during LH release indicates changes in activity of multiple neurons (7). The electrodes with recorded activity during LH pulses are associated with either GnRH somata or fibers (8). Therefore, at least some of this activity arises from GnRH neurons.
The mechanisms that result in synchronized firing in hypothalamic GnRH neurons are unknown. Elucidating the mechanisms that coordinate firing in GnRH neurons is a complex problem. First, the GnRH neurons are relatively few in number. In rodents, there are 800-2500 GnRH neurons. It is not clear that all GnRH neurons are involved in episodic GnRH release. Moreover, GnRH neurons are diffusely distributed (1). This has complicated our understanding of coordination of firing and has made many technical approaches intractable. We have optimized loose cell-attached recordings in current-clamp mode for the direct detection of action potentials and developed a recording approach that allows for simultaneous recordings from pairs of GnRH neurons.

Dual Somatic Recordings from Gonadotropin-Releasing Hormone GnRH Neurons Identified by Green Fluorescent Protein GFP in Hypothalamic Slices

Activity in neuronal systems often requires synchronous action potential discharges from neurons within a specific population. For example, pulses of gonadotropin-releasing hormone (GnRH) likely require coordinated activity between GnRH neurons. We present our methodological approach for reliably obtaining simultaneous electrophysiological recordings from the diffusely distributed GnRH neurons.

Gonadotropin-Releasing Hormone (GnRH) is a small neuropeptide that regulates pituitary release of luteinizing hormone (LH) and follicle-stimulating ...

Gene Delivery to Postnatal Rat Brain by Non-ventricular Plasmid Injection and Electroporation

Creation of transgenic animals is a standard approach in studying functions of a gene of interest in vivo. However, many knockout or transgenic animals are not viable in those cases where the modified gene is expressed or deleted in the whole organism. Moreover, a variety of compensatory mechanisms often make it difficult to interpret the results. The compensatory effects can be alleviated by either timing the gene expression or limiting the amount of transfected cells.
The method of postnatal non-ventricular microinjection and in vivo electroporation allows targeted delivery of genes, siRNA or dye molecules directly to a small region of interest in the newborn rodent brain. In contrast to conventional ventricular injection technique, this method allows transfection of non-migratory cell types. Animals transfected by means of the method described here can be used, for example, for two-photon in vivo imaging or in electrophysiological experiments on acute brain slices.

This protocol describes a non-viral method of delivery of genetic constructs to a certain area of living rodent brain. The method consists of plasmid preparation, micropipette fabrication, neonatal rat pup surgery, microinjection of the construct, and in vivo electroporation.

Creation of transgenic animals is a standard approach in studying functions of a gene of interest in vivo. However, many knockout or transgenic animals ...

Autologous Blood Injection to Model Spontaneous Intracerebral Hemorrhage in Mice

Investigation of the pathophysiology of injury after intracerebral
hemorrhage (ICH) requires a reproducible animal model. While ICH
accounts for 10-15% of all strokes, there remains no specific
effective therapy. The autologous blood injection model in mice
involves the stereotaxic injection of arterial blood into the basal
ganglia mimicking a spontaneous hypertensive hemorrhage in man. The
response to hemorrhage can then be studied in vivo and the
neurobehavioral deficits quantified, allowing for description of the
ensuing pathology and the testing of potential therapeutic agents. The
procedure described in this protocol uses the double injection
technique to minimize risk of blood reflux up the needle track, no
anticoagulants in the pumping system, and eliminates all dead space
and expandable tubing in the system.

The autologous blood injection model of intracerebral hemorrhage in mice described in this protocol uses the double injection technique to minimize risk of blood reflux up the needle track, no anticoagulants in the pumping system, and eliminates all dead space and expandable tubing in the system.

Investigation of the pathophysiology of injury after intracerebral
hemorrhage (ICH) requires a reproducible animal model.  While ICH
accounts for 10-15% ...

Experimental Demyelination and Remyelination of Murine Spinal Cord by Focal Injection of Lysolecithin

Multiple sclerosis is an inflammatory demyelinating disease of the central nervous system characterized by plaque formation containing lost oligodendrocytes, myelin, axons, and neurons. Remyelination is an endogenous repair mechanism whereby new myelin is produced subsequent to proliferation, recruitment, and differentiation of oligodendrocyte precursor cells into myelin-forming oligodendrocytes, and is necessary to protect axons from further damage. Currently, all therapeutics for the treatment of multiple sclerosis target the aberrant immune component of the disease, which reduce inflammatory relapses but do not prevent progression to irreversible neurological decline. It is therefore imperative that remyelination-promoting strategies be developed which may delay disease progression and perhaps reverse neurological symptoms. Several animal models of demyelination exist, including experimental autoimmune encephalomyelitis and curprizone; however, there are limitations in their use for studying remyelination. A more robust approach is the focal injection of toxins into the central nervous system, including the detergent lysolecithin into the spinal cord white matter of rodents. In this protocol, we demonstrate that the surgical procedure involved in injecting lysolecithin into the ventral white matter of mice is fast, cost-effective, and requires no additional materials than those commercially available. This procedure is important not only for studying the normal events involved in the remyelination process, but also as a pre-clinical tool for screening candidate remyelination-promoting therapeutics.

Demyelinating diseases can be modeled in animals by focal application of lysolecithin into the CNS. A single injection of lysolecithin into mouse spinal cord produces a lesion that spontaneously repairs over time. The goal is to study factors involved in de- and remyelination, and to test agents for enhancing repair.

Multiple sclerosis is an inflammatory demyelinating disease of the central nervous system characterized by plaque formation containing lost ...

Rat Model of Widespread Cerebral Cortical Demyelination Induced by an Intracerebral Injection of Pro-Inflammatory Cytokines

Multiple sclerosis (MS) is the most common immune-mediated disease of the central nervous system (CNS) and progressively leads to physical disability and death, caused by white matter lesions in the spinal cord and cerebellum, as well as by demyelination in grey matter. Whilst conventional models of experimental allergic encephalomyelitis are suitable for the investigation of the cell-mediated inflammation in the spinal and cerebellar white matter, they fail to address grey matter pathologies. Here, we present the experimental protocol for a novel rat model of cortical demyelination allowing the investigation of the pathological and molecular mechanisms leading to cortical lesions. The demyelination is induced by an immunization with low-dose myelin oligodendrocyte glycoprotein (MOG) in an incomplete Freund&#39;s adjuvant followed by a catheter-mediated intracerebral delivery of pro-inflammatory cytokines. The catheter, moreover, enables multiple rounds of demyelination without causing injection-induced trauma, as well as the intracerebral delivery of potential therapeutic drugs undergoing a preclinical investigation. The method is also ethically favorable as animal pain and distress or disability are controlled and relatively minimal. The expected timeframe for the implementation of the entire protocol is around 8 - 10 weeks.

The protocol presented here allows the reproduction of a widespread grey matter demyelination of both cortical hemispheres in adult male Dark Agouti rats. The method comprises of intracerebral implantation of a catheter, subclinical immunization against myelin oligodendrocyte glycoprotein, and intracerebral injection of a pro-inflammatory cytokine mixture through the implanted catheter.

Multiple sclerosis (MS) is the most common immune-mediated disease of the central nervous system (CNS) and progressively leads to physical disability and ...

A Volumetric Method for Quantification of Cerebral Vasospasm in a Murine Model of Subarachnoid Hemorrhage

Subarachnoid hemorrhage (SAH) is a subtype of hemorrhagic stroke. Cerebral vasospasm that occurs in the aftermath of the bleeding is an important factor determining patient outcome and is therefore frequently taken as a study endpoint. However, in small animal studies on SAH, quantification of cerebral vasospasm is a major challenge. Here, an ex vivo method is presented that allows quantification of volumes of entire vessel segments, which can be used as an objective measure to quantify cerebral vasospasm. In a first step, endovascular casting of the cerebral vasculature is performed using a radiopaque casting agent. Then, cross-sectional imaging data are acquired by micro computed tomography. The final step involves 3-dimensional reconstruction of the virtual vascular tree, followed by an algorithm to calculate center lines and volumes of the selected vessel segments. The method resulted in a highly accurate virtual reconstruction of the cerebrovascular tree shown by a diameter-based comparison of anatomical samples with their virtual reconstructions. Compared with vessel diameters alone, the vessel volumes highlight the differences between vasospastic and non-vasospastic vessels shown in a series of SAH and sham-operated mice.

The goal of this article is to present a method that allows a 3-dimensional reconstruction of the cerebrovascular tree in mice after micro computed tomography and determination of volumes of entire vessel segments that can be used to quantify cerebral vasospasm in murine models of subarachnoid hemorrhage.

Subarachnoid hemorrhage (SAH) is a subtype of hemorrhagic stroke. Cerebral vasospasm that occurs in the aftermath of the bleeding is an important factor ...

Minimally Invasive Endoscopic Intracerebral Hemorrhage Evacuation

Intracerebral hemorrhage (ICH) is a subtype of stroke with high mortality and poor functional outcomes, largely because there are no evidence-based treatment options for this devastating disease process. In the past decade, a number of minimally invasive surgeries have emerged to address this issue, one of which is endoscopic evacuation. Stereotactic ICH Underwater Blood Aspiration (SCUBA) is a novel endoscopic evacuation technique performed in a fluid-filled cavity using an aspiration system to provide an additional degree of freedom during the procedure. The SCUBA procedure utilizes a suction device, endoscope, and sheath and is divided into two phases. The first phase involves maximal aspiration and minimal irrigation to decrease clot burden. The second phase involves increasing irrigation for visibility, decreasing aspiration strength for targeted aspiration without disturbing the cavity wall, and cauterizing any bleeding vessels. Using the endoscope and aspiration wand, this technique aims to maximize hematoma evacuation while minimizing collateral damage to the surrounding brain. Advantages of the SCUBA technique include the use of a low-profile endoscopic sheath minimizing brain disruption and improved visualization with a fluid-filled cavity rather than an air-filled one.

This paper details the surgical protocol for minimally invasive endoscopic intracerebral hemorrhage evacuation using the SCUBA technique.

Intracerebral hemorrhage (ICH) is a subtype of stroke with high mortality and poor functional outcomes, largely because there are no evidence-based ...

Sub-acute Cerebral Microhemorrhages Induced by Lipopolysaccharide Injection in Rats

Cerebral microhemorrhages (CMHs) are common in aged patients and are correlated to various neuropsychiatric disorders. The etiology of CMHs is complex, and neuroinflammation is often observed as a co-occurrence. Here, we describe a sub-acute CMHs rat model induced by lipopolysaccharide (LPS) injection, as well as a method for detecting CMHs. Systemic LPS injection is relatively simple, economical, and cost-effective. One major advantage of LPS injection is its stability to induce inflammation. CMHs caused by LPS injection could be detected by gross observation, hematoxylin and eosin (HE) staining, Perl&#39;s Prussian staining, Evans blue (EB) double-labeling, and magnetic resonance imaging-susceptibility weighted imaging (MRI-SWI) technology. Finally, other methods of developing CMHs animal models, including their advantages and/or disadvantages, are also discussed in this report.

We present a protocol to induce and detect CMHs caused by LPS injection in Sprague-Dawley rats, which may be utilized in future research investigations on the pathogenesis of CMHs.

Cerebral microhemorrhages (CMHs) are common in aged patients and are correlated to various neuropsychiatric disorders. The etiology of CMHs is complex, ...

In Vivo Direct Reprogramming of Resident Glial Cells into Interneurons by Intracerebral Injection of Viral Vectors

Converting resident glia in the brain into functional and subtype-specific neurons in vivo provides a step forward towards the development of alternative cell replacement therapies while also creating tools to study cell fate in situ. To date, it has been possible to obtain neurons via in vivo reprogramming, but the precise phenotype of these neurons or how they mature has not been analyzed in detail. In this protocol, we describe a more efficient conversion and cell-specific identification of the in vivo reprogrammed neurons, using an AAV-based viral vector system. We also provide a protocol for functional assessment of the reprogrammed cells&#8217; neuronal maturation. By injecting flip-excision (FLEX) vectors, containing the reprogramming and synapsin-driven reporter genes&#160;to specific cell types in the brain that serve as the target for cell reprogramming. This technique allows for the easy identification of newly reprogrammed neurons. Results show that the obtained reprogrammed neurons functionally mature over time, receive synaptic contacts and show electrophysiological properties of different types of interneurons. Using the transcription factors Ascl1, Lmx1a and Nurr1, the majority of the reprogrammed cells have properties of fast-spiking, parvalbumin-containing interneurons.

This protocol aims at generating directly reprogrammed interneurons in vivo, using an AAV-based viral system in the brain and a FLEX synapsin-driven GFP reporter, which allows for cell identification and further analysis in vivo.

Converting resident glia in the brain into functional and subtype-specific neurons in vivo provides a step forward towards the development of alternative ...

Establishment of Acute Pontine Infarction in Rats by Electrical Stimulation

Pontine infarction is the most common stroke subtype in the posterior circulation, while there lacks a rodent model mimicking pontine infarction. Provided here is a protocol for successfully establishing a rat model of acute pontine infarction. Rats weighing about 250 g are used, and a probe with an insulated sheath is injected into the pons using a stereotaxic apparatus. A lesion is produced by the electrical stimulation with a single pulse. The Longa score, Berderson score, and beam balance test are used to assess neurological deficits. Additionally, the adhesive-removal somatosensory test is used to determine sensorimotor function, and the limb placement test is used to evaluate proprioception. MRI scans are then used to assess the infarction in vivo, and TTC staining is used to confirm the infarction in vitro. Here, a successful infarction is identified that is located in the anterolateral basis of the rostral pons. In conclusion, a new method is described to establish an acute pontine infarction rat model.

Presented here is a protocol for establishing acute pontine infarction in a rat model via electrical stimulation with a single pulse.

Pontine infarction is the most common stroke subtype in the posterior circulation, while there lacks a rodent model mimicking pontine infarction. Provided ...