Mouse Cardiac Arrest Model for Brain Imaging and Brain Physiology Monitoring During Ischemia and Resuscitation

Summary

This protocol demonstrates a unique mouse model of asphyxia cardiac arrest that does not require chest compression for resuscitation. This model is useful for monitoring and imaging the dynamics of brain physiology during cardiac arrest and resuscitation.

Abstract

Most cardiac arrest (CA) survivors experience varying degrees of neurologic deficits. To understand the mechanisms that underpin CA-induced brain injury and, subsequently, develop effective treatments, experimental CA research is essential. To this end, a few mouse CA models have been established. In most of these models, the mice are placed in the supine position in order to perform chest compression for cardiopulmonary resuscitation (CPR). However, this resuscitation procedure makes the real-time imaging/monitoring of brain physiology during CA and resuscitation challenging. To obtain such critical knowledge, the present protocol presents a mouse asphyxia CA model that does not require the chest compression CPR step. This model allows for the study of dynamic changes in blood flow, vascular structure, electrical potentials, and brain tissue oxygen from the pre-CA baseline to early post-CA reperfusion. Importantly, this model applies to aged mice. Thus, this mouse CA model is expected to be a critical tool for deciphering the impact of CA on brain physiology.

Introduction

Cardiac arrest (CA) remains a global public health crisis¹. More than 356,000 out-of-hospital and 290,000 in-hospital CA cases are reported annually in the US alone, and most CA victims are over 60 years old. Notably, post-CA neurologic impairments are common among survivors, and these represent a major challenge for CA management^2,3,4,5. To understand post-CA brain pathologic changes and their effects on neurologic outcomes, various neurophysiologic monitoring and brain tissue monitoring techniques have been applied in patients^{6,7,8,9,10,11,12}. Using near-infrared spectroscopy, real-time brain monitoring has also been performed in CA rats to predict neurologic outcomes¹³.

However, in murine CA models, such an imaging approach has been complicated by the need for chest compressions to restore spontaneous circulation, which always entails substantial physical motion and, thus, hinders delicate imaging procedures. Moreover, CA models are normally performed with mice in a supine position, whereas the mice must be turned to the prone position for many brain imaging modalities. Thus, a mouse model with minimal body movement during the surgery is required in many cases in order to perform real-time imaging/monitoring of the brain during the whole CA procedure, spanning from pre-CA to post-resuscitation.

Previously, Zhang et al. reported a mouse CA model that could be useful for brain imaging¹⁴. In their model, CA was induced by bolus injections of vecuronium and esmolol followed by the cessation of mechanical ventilation. They showed that after 5 min of CA, resuscitation could be achieved by infusing a resuscitation mixture. Notably, however, circulatory arrest in their model occurred only about 10 s after the esmolol injection. Thus, this model does not recapitulate the progression of asphyxia-induced CA in patients, including hypercapnia and tissue hypoxia during the prearrest period.

The overall goal of the current surgical procedure is to model clinical asphyxia CA in mice followed by resuscitation without chest compressions. This CA model, therefore, allows the use of complex imaging techniques to study brain physiology in mice¹⁵.

Protocol

All the procedures described here were conducted in accordance with the National Institutes of Health (NIH) guidelines for the care and use of animals in research, and the protocol was approved by the Duke Institute of Animal Care and Use Committee (IACUC). C57BL/6 male and female mice aged 8-10 weeks old were used for the present study.

1. Surgical preparation

1. Weigh a mouse on a digital scale, and place it into a 4 in x 4 in x 7 in plexiglass anesthesia induction box.
2. Adjust the anesthesia vaporizer to 5% isoflurane, the oxygen flow meter to 30, and the nitrogen flow meter to 70 (see Table of Materials).
3. Take the animal out of the induction box, and lay it in a supine position on the surgical bench when its respiratory rate has decreased to 30-40 breaths per minute.
4. Pull out the tongue with blunt forceps, and hold it using the non-dominant hand. Use the dominant hand to insert a laryngoscope (see Table of Materials) into the mouse's mouth and visualize the vocal cord.
5. Use the non-dominant hand to insert a guide wire and 20 G intravenous catheter into the mouth. Gently insert the guide wire into the trachea.
6. Push the catheter into the trachea until the wing part of the catheter is even with the nose tip.
   NOTE: Do not intubate a mouse that is not fully anesthetized since this may injure the trachea and cause airway bleeding.
7. Connect the intubated mouse to a small animal ventilator (see Table of Materials), and reduce the isoflurane to 1.5%.
8. Input the mouse's body weight into the control panel of the ventilator to determine the tidal volume and respiratory rate.
9. Keep the mouse in a supine position under a heat lamp, and maintain the rectal temperature at 37 °C with a temperature controller.
10. Shave the inguinal areas, disinfect the surgical area at least three times with iodine and alcohol (see Table of Materials), and cover the area with a sterile surgical drape.
11. Apply eye ointment to both eyes and administer 5 mg/kg carprofen subcutaneously before surgery.
12. Open the sterile instrument package for surgery. Make a 1 cm skin incision with surgical scissors to access the femoral arteries on both sides. Dissect and ligate the distal femoral artery with a single strand of 4-0 silk suture (see Table of Materials), and apply one drop of lidocaine.
13. Apply an aneurysm clip at the proximal femoral artery and make a small cut on the artery distal to the clip. Insert a polyethylene 10 (PE-10, see Table of Materials) catheter into the left and the right femoral arteries.
    NOTE: The left arterial line is used for blood pressure monitoring, while the right one is used for blood withdrawal and resuscitation mixture infusion.
14. Inject 50 µL of 1:10 heparinized saline into each arterial line to prevent clotting in the line.
15. Turn the mouse to the prone position, and mount it on a stereotaxic head frame.
16. Connect three needle electrodes (red, green, and black) to the left arm, left leg, and right arm for electrocardiogram (ECG, see Table of Materials) monitoring.
17. Glue a flexible plastic fiber probe onto the intact temporal skull through a 0.5 cm skin incision for cerebral blood flow monitoring. This step is optional.
18. Shave the top of the head, disinfect the surgical area at least three times with iodine and alcohol, and cover the area with a sterile surgical drapel.
19. Cut a 2.5 cm midline skin incision, and use four small retractors to expose the entire skull surface for brain imaging.
20. Place a monitoring imager (e.g., a laser speckle contrast imager, see Table of Materials) above the head.
    NOTE: A few drops of saline can be added to the skull surface to facilitate laser speckle contrast imaging.

2. Induction of cardiac arrest

1. Fill a 1 mL plastic syringe with 26 µL of the resuscitation cocktail stock solution.
   NOTE: Each milliliter of this solution contains 400 µL of 1 mg/mL epinephrine, 500 µL of 8.4% sodium bicarbonate, 50 µL of 1,000 U/mL heparin, and 50 µL of 0.9% sodium chloride (see Table of Materials).
2. Wait until the body temperature reaches 37 °C. Adjust the oxygen meter to 100% to oxygenate the blood for 2 min.
3. Withdraw the oxygenated arterial blood up to 200 µL via the right femoral artery into the prepared plastic syringe containing 26 µL of resuscitation cocktail stock solution.
4. Switch off the oxygen, and increase the nitrogen to 100% to induce anoxia.
   NOTE: After approximately 45 s, the heart will fail to function, and the heart rate will rapidly decrease, indicating the onset of CA. After about 2 min of oxygen deprivation, the ECG monitoring will indicate an asystole, and there will be no measurable systemic blood pressure and negligible cerebral blood flow.
5. Turn off the ventilator, isoflurane vaporizer, temperature controller, and nitrogen flowmeter. Adjust the oxygen to 100% in preparation for resuscitation.

3. Resuscitation procedure

1. Turn the ventilator on at 8 min after CA onset.
2. Immediately start to infuse the withdrawn oxygenated blood mixed with the resuscitation cocktail into the blood circulation via the right femoral artery in 1 min.
   NOTE: The infusion leads to a gradual increase in the heart rate and the restoration of blood perfusion; eventually, the return of spontaneous circulation (ROSC) is achieved.

4. Post-CA recovery

1. Place the mouse in the supine position after removing it from the stereotaxic frame, and remove the PE-10 catheters from the femoral arteries.
2. Apply 0.25% bupivacaine to the skin incision, and suture the skin incisions using a 6-0 nylon suture (see Table of Materials). Apply antibiotic ointment to the surface of the skin incision.
3. Disconnect the mouse ventilator when spontaneous respiration is restored.
4. Transfer the mouse to a recovery chamber with a controlled temperature of 32 °C.
5. After 2 h of recovery, extubate the mouse, and return to the home cage. Inject 0.5 mL of normal saline subcutaneously to prevent dehydration.

Results

To induce CA, the mouse was anesthetized with 1.5% isoflurane and ventilated with 100% nitrogen. This condition led to severe bradycardia in 45 s (Figure 1). Following 2 min of anoxia, the heart rate dramatically reduced (Figure 2), the blood pressure decreased below 20 mmHg, and the cerebral blood flow ceased completely (Figure 1). As the isoflurane was turned off, the body temperature was no longer managed and slowly dropped ...

Discussion

In experimental CA studies, asphyxia, potassium chloride injections, or electrical current-derived ventricular fibrillation have been used to induce CA^{16,17,18,19,20,21,22,23}. Normally, CPR is required for resuscitation in these CA models, especially in mic...

Materials

Name	Company	Catalog Number	Comments
Adrenalin	Par Pharmaceutical	NDC 42023-159-01
Alcohol swabs	BD	326895
Animal Bio Amp	ADInstruments	FE232
BP transducer	ADInstruments	MLT0699
Bridge Amp	ADInstruments	FE117
Heparin sodium injection, USP	Fresenius Kabi	NDC 63323-540-05
Isoflurane	Covetrus	NDC 11695-6777-2
Laser Doppler perfusion monitor	Moor Instruments	moorVMS-LDF1
Laser speckle imaging system	RWD	RFLSI III
Lubricant eye ointment	Bausch + Lomb	339081
Micro clip	Roboz	RS-5431
Mouse rectal probe	Physitemp	RET-3
Needle electrode	ADInstruments	MLA1213	29 Ga, 1.5 mm socket
Nitrogen	Airgas	UN1066
Optic plastic fibre	Moor Instruments	POF500
Otoscope	Welchallyn	728	2.5 mm Speculum
Oxygen	Airgas	UN1072
PE-10 tubing	BD	427401	Polyethylene tubing
Povidone-iodine	CVS	955338
PowerLab 8/35	ADInstruments
Rimadyl (carprofen)	Zoetis	6100701	Injectable 50 mg/ml
Small animal ventilator	Kent Scientific	RoVent Jr.
Temperature controller	Physitemp	TCAT-2DF
Triple antibioric & pain relief	CVS	NDC 59770-823-56
Vaporizer	RWD	R583S
0.25% bupivacaine	Hospira	NDC 0409-1159-18
0.9% sodium chroride	ICU Medical	NDC 0990-7983-03
1 mL plastic syringe	BD	309659
4-0 silk suture	Look	SP116	Black braided silk
6-0 nylon suture	Ethilon	1698G
8.4% sodium bicarbonate Inj., USP	Hospira	NDC 0409-6625-02
20 G IV catheter	BD	381534	20GA 1.6 IN
30 G PrecisionGlide needle	BD	305106	30 G X 1/2