Measuring Cerebral Vasospasm after Subarachnoid Hemorrhage in Mice

Summary

The present protocol provides a detailed procedure for inducing subarachnoid hemorrhage in mice via autologous blood injection to the anterior circulation and measuring delayed cerebral vasospasm by vascular gel casting.

Abstract

Subarachnoid hemorrhage (SAH) is a devastating illness, and patients who survive are still at risk for long-term neurological deficits. Cerebral vasospasm is one of several contributing factors to morbidity and mortality after SAH. Preclinical animal models are essential resources to investigate pathophysiology and novel therapeutics. This protocol provides a two-phase method for both inducing SAH in mice and evaluating delayed cerebral vasospasm by measuring cerebral artery diameter. In the first step, an anterior circulation autologous blood injection method is used to reproduce the most common anatomical location of human non-traumatic SAH and reliably control the volume and distribution of hemorrhage. The duration of this procedure is approximately 20 min per animal. Next, on post-SAH day 5, the cerebral vasculature is fixed and casted using a gelatin-dye solution before removing the brain for imaging. Finally, the diameter of many cerebral arteries can be measured simultaneously using a variety of image analysis software platforms. These procedures are rigorous and reproducible while also offering a time- and resource-efficient method for studying the pathophysiology of SAH and cerebral vasospasm.

Introduction

The morbidity and mortality caused by subarachnoid hemorrhage (SAH) are staggering despite decades of research and a plethora of disappointing candidate therapeutics^1,2,3,4,5,6. Delayed cerebral ischemia (DCI) is a fundamental cause of poor outcomes among SAH patients, yet is an incompletely understood pathophysiologically^7,8,9. There are many contributing factors to DCI, including microcirculatory dysfunction, early brain injury, cortical spreading depolarizations, and cerebral vasospasm¹⁰. Developing novel treatment strategies will require innovative basic science and translational investigation, including preclinical animal models. Mouse models are an appealing option due to their similar cerebrovascular anatomy compared to humans and the availability of genetically-modified strains¹¹. The effectiveness of mouse models is dependent on the method of SAH induction and the selection of appropriate experimental endpoints. Critical evaluation and rigorous implementation of these variables are essential to improving SAH-related morbidity and mortality.

There are two primary variables in models of SAH induction; method of hemorrhage (endovascular puncture or transcranial blood injection) and location of hemorrhage (cisterna magna/posterior circulation or prechiasmatic cistern/anterior circulation)¹². Endovascular puncture models have several strengths, including maintaining cranial integrity compared to transcranial procedures and the substantial severity of hemorrhage, allowing for more readily detectable neurological deficits. On the other hand, endovascular puncture methods are highly variable in the volume and location of hemorrhage. This variability can skew results if the study design is not adequately powered. Additionally, the perioperative mortality rate of endovascular puncture models is approximately 40%^12,13, which can drastically increase time and cost. Overall, the transcranial blood injection model is much more consistent, allows precise control over the volume and location of hemorrhage, and is much lower in perioperative mortality^12,14,15.

Transcranial blood injection was initially developed for injection in the cisterna magna via puncture of the atlanto-occipital membrane¹⁶; however, prechiasmatic injection through a cranial burr hole has now been reported as well^14,17. The primary benefit of anterior circulation models is the similarity to human SAH, where most non-traumatic SAH/vasospasm occurs in the anterior circulation. Transcranial autologous blood injection targeting the prechiasmatic cistern is an ideal preclinical model for investigating SAH and cerebral vasospasm.

There are also critical methodologic variables to consider when measuring cerebral vasospasm. Many researchers perform hematoxylin and eosin (H&E) staining on tissue sections to measure the inner and outer diameter, vessel wall thickness, and other histological parameters. This method has some critical limitations. First, cerebral vessels are rarely in a straight line; thus, tissue sections may be oblique for the ideal orthogonal orientation of a cross-section. This issue may be compounded when attempting to compare across samples. Second, histological analysis of vessels may limit the number of vessels to be studied. The sectioning angle needed to measure one cerebral artery can be quite different from the required angle to measure a nearby vessel. A gel-dye vascular casting method has been developed to solve these challenges^18,19,20. In this method, gelatin casts the morphology of the cerebral vessels, and India ink dye allows for high contrast upon digital image analysis. This technique permits the measurement of many vessels along their natural anatomical course.

The purpose of this report is to demonstrate a beginning-to-end experimental procedure for inducing subarachnoid hemorrhage and measuring delayed cerebral vasospasm. This representative experiment compared vasospasm between sham and SAH groups. Mice were allocated to each group by block randomization of mouse cages. Carefully considering the variables in each step of the protocol ensures the results are rigorous and reproducible while maintaining cost and resource efficiency.

Protocol

The present protocol was approved and performed in compliance with the institutional policy and guidelines of the University of Florida Institutional Animal Care and Use Committee (#201910613). The representative experiment was conducted consistently with ARRIVE guidelines²¹. Female C57BL/6J mice, 12-15-week-old, were used for the experiments; however, mice of any age, strain, or sex can be used. Mice were housed in ventilated cages on a 12 h/12 h light-dark cycle with ad libitum access to food and water. An overview of the protocol is illustrated in Figure 1.

1. SAH procedure

1. Prepare the mice following the steps below.
   1. Anesthetize the mice via intraperitoneal injection of ketamine (100 mg/kg) and xylazine (15 mg/kg). Ensure appropriate level of anesthesia by performing a toe pinch.
      NOTE: This procedure can also work with inhaled anesthetics, provided the stereotaxic frame is adapted with a gas delivery-compatible nose cone.
   2. Shave the scalp, including hair between the eyes and the scalp. Then clean the shaved skin with swabs of 10% povidone-iodine and sterile saline (a total of at least three swabs to be used). Shave the tail's ventral base and clean in the same manner, as this is the incision site for blood collection.
   3. After cleaning, fix the mice into the stereotaxic frame (see Table of Materials) using the nose cone and ear supports. Maintain the body temperature at 37 °C using an adjustable heating pad integrated into the stereotaxic frame.
2. Incise the scalp and drill the burr hole.
   1. Once fixed in the stereotaxic frame, create a longitudinal incision in the scalp along the midsagittal plane of the skull. Begin the incision rostrally from bregma (meeting point of frontal bones and parietal bones) and extend past the anterior edges of the frontal bones in the midline; at least 0.75 cm is needed for sufficient exposure.
   2. Create a burr hole using a variable speed rotary drill with a 1.5 mm drill bit (see Table of Materials). Make the burr hole 5 mm rostral to bregma and 0.5 mm right of midline, avoiding puncture of the superior sagittal sinus. Quickly stop any resultant bleeding by pressing manually using a sterile cotton-tipped applicator.
3. Perform autologous blood collection and injection.
   1. First, collect the autologous blood from the ventral tail artery. Make a 2-3 mm transverse incision along the base of the tail, and allow the blood to drop onto a paraffin wax paper. After collecting 5 drops (~150-200 µL) of blood, stop the bleeding with sterile gauze and manual pressure.
      NOTE: The nonpolar wax paper will slow blood clotting to help the loading of blood in the syringe.
   2. Load 60 µL of autologous blood into a microlitre syringe with a 26 G needle (see Table of Materials) and secure in the manipulator arm of the stereotaxic frame. When performing a saline injection control procedure, load 60 µL of the sterile saline into the syringe during this step. Then, pass the needle through the burr hole until it comes in contact with the skull base (~7.2 mm, but ultimately judge visually by the skull movement).
      NOTE: The manipulator's arm is angled to 30° such that the needle is in the longitudinal plane and pointing rostrally.
   3. Leave the needle in place for 20 s to allow for tissue accommodation, and then inject the blood (or saline control) at a rate of 10 µL/min.
      NOTE: In this experiment, the injection volume was 50 µL, leaving 10 µL in the syringe to prevent any associated air bubbles from being injected in the subarachnoid space.
   4. Leave the needle in place for an additional 3 min to permit coagulation of the injected blood.
      NOTE: Perform a sham procedure by collecting and inserting the needle as described, but without blood injection or by injecting an equivalent volume of saline.
   5. Remove the needle at a 10 mm/min rate, and fill the burr hole with bone wax (see Table of Materials). Then approximate the skin flaps using 5-0 sutures.
   6. Post-operatively, administer each mouse with a bolus of pre-warmed saline equal to the volume collected minus the volume injected in addition to the analgesics and other agents as outlined in institutionally approved animal protocols.
      ​NOTE: The mice used for the representative experiment received bupivacaine (1.5 mg/kg) once post-surgery, carprofen (5 mg/kg) once per day for 48 h post-surgery, and buprenorphine (0.1 mg/kg) every 8 h for 48 h post-surgery. Bodyweight, body condition score, sensorimotor function, hydration status, and general appearance of distress were used to monitor the clinical status of animals throughout the protocol.

2. Vascular casting

1. Prepare the mice and the perfusion agents.
   1. Prepare solutions of normal saline, 4% paraformaldehyde (PFA) in PBS, and 5% gelatin dissolved in PBS with 5% India ink dye (see Table of Materials).
      NOTE: Processing of each animal require approximately 3 mL of saline, 10 mL of 4% PFA solution, and 1 mL of gelatin-India ink solution.
   2. Anesthetize the mice by intraperitoneal injection of ketamine (100 mg/kg) and xylazine (15 mg/kg) and then place them on absorbent paper towels. Confirm the appropriate level of anesthesia with a toe pinch.
2. Perform cardiac perfusion.
   1. Make a 3 cm transverse incision in the abdomen immediately inferior to the diaphragm. Make two more incisions, each 2-3 cm in length, in the rib cage along the midclavicular lines, permitting exposure of the heart and lungs by reflecting the front of the ribcage superiorly.
   2. Using a 23 G needle, puncture the right atrium and then place the needle in the left ventricle via puncturing the apex of the heart. Secure the needle with locking forceps or hold it in place manually.
   3. Perfuse¹⁸ the left ventricle serially with saline, PFA, and gelatin-India ink solution (for the required volume of each solution, see step 2.1.1). Maintain the infusion rate at ~15 mL/min to mimic cardiac output²². After cardiac perfusion, place the carcasses into a 4 °C refrigerator overnight to allow for gelatin hardening.
3. Measure the cerebral artery diameter.
   1. The next day, make a 2 cm longitudinal incision along the midsagittal plane from the foramen magnum through the frontal bones and reflect the frontal and parietal bones laterally. Then remove the brain by carefully dissecting the cranial nerves and cutting the internal carotid artery. Ensure not to disturb or damage the casted cerebral arteries.
   2. Rinse the isolated brain gently with saline to remove any debris. Also, dissect any remaining cranial nerve segments that obscure the cerebral vessels.
   3. Next, place the brain into a brain matrix and place it under a dissecting microscope with an attached digital camera for imaging (see Table of Materials). Adjust the lighting and camera exposure settings to make all cerebral vessels of interest visible without glare or other obstruction.
   4. Take a calibration image of a ruler or other precise measurement standard to calibrate the image analysis software (see Table of Materials).
   5. Finally, measure the diameter of the vessels of interest at various points using software capable of linear measurement.
      NOTE: Observers blinded to the treatment group are used to avoid potential bias. Data can be expressed as relative measurements (e.g., percent of sham group), or they can be calibrated to report the diameters in microns.

Results

One of the primary advantages of this SAH model is the low mortality. Perioperative mortality is less than 10% for the SAH procedure, and perioperative deaths from the sham procedure are very rare; mortality rates during our development of this protocol were 6.7% and 0%, respectively (Table 1). Perioperative deaths in this SAH model, similar to other methods of SAH induction, are typically due to brain herniation secondary to increased intracranial pressure¹².

Discussion

This protocol is a two-phase procedure for inducing SAH in mice and measuring cerebral vasospasm. This procedure is rigorous and reproducible while also maintaining time and resource efficiency. There are several critical steps in this procedure that must be adhered to closely. First, the coordinates of blood injections are of vital importance. Even a slight error in the location of the burr hole can lead to excessive bleeding from the dural venous sinuses and/or unintended injury to the cerebral vessels when the needle ...

Materials

Name	Company	Catalog Number	Comments
1 qt plastic bags	Ziploc
1.5 mm drill bit	Dremel	106
1 mL insulin syringes	ThermoFisher Scientific	BD 329461
23 G Luer-lock needle	Beckton-Dickinson (BD)	305145
4% paraformaldehyde	Elabscience	E-IR-R113
5% Povidone-Iodine Solution	PBS Animal Health	11205	Chlorhexadine may be used per institutional guidelines or investigator preference
5-0 monofilament suture	Ethicon	698H
Absorbent bench covering	ThermoFisher Scientific	22-131-401
Adjustable heating pad	Thermotech	S766D
Bone wax	Braintree Scientific	DYNJBW 26
Camera microscope mount adapter	AmScope	CA-NIK-SLR
Digital camera	Nikon		Any high resolution digital camera will be sufficient; specific model up to investigator preference
Dissecting microscope	Leica	M60
Ear tags	Kent Scientific	INS1005-5LS
Electric hair trimmer	Kent Scientific	CL7300-Kit
Forceps	Roboz Surgical	RS-5136
Gelatin	ThermoFisher Scientific	S25335
Hamilton syringe with 28 G needle	Hamilton Company	80630
Handheld rotary tool	Dremel	Model #4000
Image analysis software	ImagePro	10.0.04	Other software platforms (e.g. ImageJ, Orbit) may be sufficient
India ink dye	ThermoFisher Scientific	NC9903975
Ketamine	Patterson Veterinary	07-893-6763
Luer-lock syringes (3, 5, & 10 mL)	Beckton-Dickinson (BD)	309657, 309646, 309604
Mice	Charles River Laboratory	C57BL/6	Genetically-modfied strains may be used per study design
Mouse brain frame matrix	Harvard Apparatus	51386
Needle driver	Roboz Surgical	RS-7860
Opthalmic ointment	Dechra	17033-211-38
Paraffin wax paper	Stellar Scientific	HS234526A	Parafilm or equivalent
Phosphate-buffered saline	ThermoFisher Scientific	50488886
Ruler or other measurement standard	Harvard Apparatus	51386
Scalpel	Roboz Surgical	65-9843
Scissors	Roboz Surgical	RS-5882
Stereotaxic frame with syringe holder	Harvard Apparatus	75-1822
Sterile cotton-tipped applicators	Uline	S-18991
Sterile gauze	ThermoFisher Scientific	19-090-729
Sterile saline	Patterson Veterinary	07-869-6657
Sterile surgical gloves	AD Surgical	A5CS-LTX65
Sterile surgical gown	AD Surgical	GWN-D02-CS
Surgical microscope	World Precision Instruments	PSMB5N
Xylazine	Patterson Veterinary	07-893-8424