Massive Pontine Hemorrhage by Dual Injection of Autologous Blood

Summary

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

Abstract

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 µL 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.

Introduction

Intracerebral hemorrhage accounts for one-fifth of stroke patients. The prognosis of intracerebral hemorrhage depends on the speed, volume, and location of bleeding^1,2. 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 pons^6,7,8. 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 µL) and moderate (60 µL) 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 µL autologous blood at two different locations in the pons.

Protocol

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

1. Use 8-week-old male Sprague-Dawley rats weighing 250 ± 10 g.
2. House the rats for at least 7 days before surgery under controlled environmental conditions with an ambient temperature of 25 °C, relative humidity of 65%, and a 12/12-h light-dark cycle.
3. Provide food and water with no limit.
4. Prepare the instruments (Figure 2A-E).

2. Inject the blood in the pons

1. Weigh the rats again 3 days before surgery to select rats with suitable body weight for the experiments.
2. 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.
3. Preheat the heating pad to 37 °C before anesthesia.
4. Attach a microdrill to the holder on the stereotaxic frame.
5. Inject the rats with 50 mg/kg ketamine and 5 mg/kg xylazine intraperitoneally. Wait until there is no toe-pinch response.
6. 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).
7. Maintain anesthesia by isoflurane (97.5% oxygen and 2.5% isoflurane) through a stereotaxic nose cone with an inlet and outlet port.
8. Use eye ointment to keep the cornea moist.
9. Shave the hair above the skull with a micro shaver.
10. 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.
11. Apply Entoiodine surgical scrub in a circular fashion, starting at the middle of the marked mid-line and rotating outward.
12. Place the surgical drape.
13. Make an incision with a scalpel along the marked mid-line.
14. Use a cotton swab to remove any potential blood.
15. Place a piece of forceps on each side of the scalp flap to expose the skull (Figure 2G).
16. 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.
17. Mark the central point of the bregma as the origin point via a marker pen.
18. 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).
19. Remove the microdrill from the stereotaxic frame.
20. Put a 100 µL 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 µL and then drain it completely to prevent blood clotting too fast.
21. 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.
22. Attach a scalp acupuncture to a 1 mL syringe.
23. Insert the scalp acupuncture into a lateral tail vein 3 cm from the tail tip and take 150 µL of blood (Figure 2H).
24. Remove the scalp acupuncture from the 1 mL syringe.
25. Transfer the blood into a tube (Figure 2I).
    NOTE: Transfer quickly to prevent blood clotting.
26. Change the surgical gloves.
27. Choose Withdraw mode, set the volume to 100 µL, set the speed at 200 µL/min, click the RUN button, aspirate 100 µL of blood into the Hamilton syringe (Figure 2J).
28. Choose Infuse mode, set the speed at 1 µL/min.
29. Advance the syringe until the tip reaches 9.2 mm below the surface of the brain (Figure 1C and Figure 2L).
30. Click the Run button, inject the first 10 µL of blood at a speed of 1 µL/min (Table 1).
31. Stop the injection and leave the syringe in position for 20 min to prevent blood from flowing into the subarachnoid space.
32. Retract the syringe until the tip arrives at 9.0 mm below the surface of the brain.
33. Restart injection at the same speed of 1 µL/min until the residual blood has been injected completely.
34. Leave the syringe in position for 10 min to avoid blood backflow.
35. Remove the syringe from the brain slowly.
36. Use bone cement to cover the craniotomy hole.
37. 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.
38. To prevent infection, inject the rat with penicillin (0.25 mL, 80 IU diluted in 4 mL saline) intraperitoneally.
39. 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.
40. 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.

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.

1. Balance beam test¹¹.
   ​NOTE: This is a test specifically for examining the sensorimotor function of the hindlimb.
   1. Prepared the apparatus to ensure that the beam (3 cm wide × 70 cm long) is 20 cm above the floor.
   2. Put a dark box at the back end of the beam with a narrow entryway.
   3. 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.
   4. 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.
   5. 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 >40 s, but falls off; 5, attempts to balance on beam >20 s, but falls off; and 6, falls off, with no attempt to balance or balancing on beam <20 s.
2. Limb placement test
   ​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.
   1. 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.
   2. 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.
   3. 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.
   4. Put the rat on the table, push the rat with a gentle lateral pressure behind the rat'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.
   5. 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.
   6. 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.
   7. 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.
3. 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.
   1. Place the rat on the table (50 cm long × 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.
   2. Place the rat on the table (50 cm long × 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.
   3. Observe the craniocaudal circling: turns bilaterally, 3; prefers 1 side, 2; only to 1 side, 1; fallen to 1 side, 0.
   4. Evaluate the forelimbs movement: equal and bilateral movement, 3; slight asymmetry, 2; great asymmetry, 1; paresis, 0.
   5. Evaluate the 4-limbs movement: equal and bilateral movement, 3; slight asymmetry, 2; great asymmetry, 1; paresis, 0.
   6. 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.
   7. When the rat is stationary, make a noise on either side of the rat's body to test hearing: fingers rubbing, 3; snap of fingers, 2; loud clapping, 1 and no startling, 0.
   8. To check the proprioception, touch the rat'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.
   9. To evaluate the torso axial sensation, place a blunt stick on either side of the rat'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.
   10. 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.
   11. 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.
   12. To observe exploration, use a light dark box.
       ​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.
   13. 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'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.
   14. 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.
   15. 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.
   16. Calculate the points.

4. Hemorrhage confirmation by MRI

1. Perform the MRI scan at 24 h post-surgery.
2. Anesthetize the rat with isoflurane (5% for induction, 1%- 1.5% for maintenance).
3. Secure the rat's head in the rat brain array coil combined with a transmit-only volume coil.
4. Place the coil together with the rat in the MRI scanner. Secure the rat within the cradle via the tooth and ear bars.
5. Use a closed-circuit thermal jacket to maintain the rat's body temperature at 37 ± 0.5 °C during MRI scanning.
6. Perform a pilot sequence to ensure correct geometry.
7. 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 × 148; field of view, 100 mm × 100 mm; 12 slices; 1.5 mm thick.
8. Return the rat to the cage.

5. Hemorrhage confirmation by gross anatomy

NOTE: Sacrifice the rats at the designated timepoint, 24 h and 14 d after surgery (Figure 1D).

1. 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).
2. 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.
3. Perform cervical dislocation.
4. 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.
5. Connect the end of a needle (27 G) to a perfusion pump containing 4 °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.
6. 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 °C saline.
7. Decapitate the rat and harvest the whole brain using scissors and forceps. Remove any moisture from the brain surface with blotting paper.
8. Keep the whole brain at -80 °C for 1 min.
   NOTE: This step could be skipped if the brain can be cut without freezing.
9. Lay the brain into the coronal rat brain matrix with the dorsal side up.
10. Insert a 0.21 mm thick stainless-steel blade into the hemorrhage center according to the hole in the surface of the brain.
11. Insert other blades into the brain at an interval of 2 mm.
12. Immerse the brain sections in 10 mL of 4% paraformaldehyde (PFA) solution for 24 h at 4 °C. Rinse them with 0.01 mmol/L phosphate buffered saline (PBS).
13. Organize the sections from rostral to caudal and image them.

6. Paraffin section and hematoxylin and eosin (HE) staining

1. 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.
2. Cut the brain from the hemorrhage center into two pieces via blade and coronal rat brain matrix.
3. Make paraffin-embedded tissue blocks.
4. Section the paraffin-embedded tissue block coronally in 40 µm thickness slides on a microtome, cut 4 sections in succession, starting from hemorrhage center, and float them in a 40 °C water bath.
5. Mount the 40 µm sections (8 slides in total for one brain) onto clean glass slides and air dry overnight at room temperature.
6. Bake the slides at 56 °C for 1 h.
7. Wash for 3 min in xylene, 3 times.
8. 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.
9. Rinse in tap water until clear.
10. Immerse the sections in Hematoxylin for about 10 min.
11. Rinse in the tap water until clear.
12. Immerse sections in eosin stain for 30 s.
13. 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).
14. Clean sections with xylene for 1 min, 2 times.
15. Mount using non-aqueous mounting medium and a coverslip.
16. Dry the sections in the air overnight at room temperature, then san them.

7. Statistics

1. Use GraphPad Prism 6.0 to calculate Student's t-test or Mann Whitney U test.
   NOTE: All data should be expressed as mean ± SE. Differences between two groups are determined with a two-tailed Student's t-test or Mann Whitney U test. P<0.05 is defined as statistical significance.

Results

A total of 25 animals were used, 3 for control, 6 for 30 µL, 6 for 60 µL, and 10 for 100 µL blood injections. One rat that received a 100 µL 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 Table 2

Discussion

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 ...

Materials

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