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 management2,3,4,5. 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 patients6,7,8,9,10,11,12. 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.
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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
- Weigh a mouse on a digital scale, and place it into a 4 in x 4 in x 7 in plexiglass anesthesia induction box.
- Adjust the anesthesia vaporizer to 5% isoflurane, the oxygen flow meter to 30, and the nitrogen flow meter to 70 (see Table of Materials).
- 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.
- 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.
- 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.
- 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. - Connect the intubated mouse to a small animal ventilator (see Table of Materials), and reduce the isoflurane to 1.5%.
- Input the mouse's body weight into the control panel of the ventilator to determine the tidal volume and respiratory rate.
- Keep the mouse in a supine position under a heat lamp, and maintain the rectal temperature at 37 °C with a temperature controller.
- 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.
- Apply eye ointment to both eyes and administer 5 mg/kg carprofen subcutaneously before surgery.
- 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.
- 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. - Inject 50 µL of 1:10 heparinized saline into each arterial line to prevent clotting in the line.
- Turn the mouse to the prone position, and mount it on a stereotaxic head frame.
- 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.
- 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.
- 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.
- Cut a 2.5 cm midline skin incision, and use four small retractors to expose the entire skull surface for brain imaging.
- 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
- 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). - Wait until the body temperature reaches 37 °C. Adjust the oxygen meter to 100% to oxygenate the blood for 2 min.
- 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.
- 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. - Turn off the ventilator, isoflurane vaporizer, temperature controller, and nitrogen flowmeter. Adjust the oxygen to 100% in preparation for resuscitation.
3. Resuscitation procedure
- Turn the ventilator on at 8 min after CA onset.
- 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
- Place the mouse in the supine position after removing it from the stereotaxic frame, and remove the PE-10 catheters from the femoral arteries.
- 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.
- Disconnect the mouse ventilator when spontaneous respiration is restored.
- Transfer the mouse to a recovery chamber with a controlled temperature of 32 °C.
- 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.
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Representative 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 to about 32 °C at the end of CA (Figure 1).
Immediately following 8 min of CA, the ventilator was turned on, and the mouse was supplied with 100% oxygen. The blood-resuscitation mixture was infused into the circulation via the arterial catheter. Shortly after the injection of the blood-resuscitation mixture, cardiac function started to recover. After a short interval, the systemic and cerebral blood flow was restored, and ROSC was established. The success rate of ROSC is almost 100% in our lab. This model has been successfully performed in young and aged mice.
Enabled by this model, two imaging modalities were used in this study, including laser speckle contrast imaging (LSCI) and photoacoustic imaging, to monitor the cerebral blood flow and blood oxygenation at the whole-brain level during CA and resuscitation. LSCI confirmed the complete absence of blood flow in the brain during CA (Figure 3). More detailed changes in blood flow, structure, and oxygenation during the CA procedure can be obtained from the photoacoustic images (Figure 4).
Figure 1: Physiologic recording during CA and resuscitation. Cerebral blood flow (% baseline; measured by laser Doppler flowmetry), blood pressure (mmHg), heart activity (beats per minute), and body temperature (°C) changes pre-CA, during CA, and after CA. The x-axis depicts the time in minutes. Please click here to view a larger version of this figure.
Figure 2: Heart activity during CA and resuscitation. The heart rate was continuously recorded, and panels (A), (B), and (C) are representative of the heart rate pre-CA, during CA, and post-CA, respectively. The y-axis depicts the absolute voltage values (mV). Please click here to view a larger version of this figure.
Figure 3: Laser speckle contrast images during CA and resuscitation. Global cerebral blood flow was monitored. CA led to a complete loss of cerebral blood flow (B) compared to baseline (A) . Hyperperfusion was present in the brain immediately after resuscitation (C) , and this was then followed by hypoperfusion during the late phase (D). Please click here to view a larger version of this figure.
Figure 4: Photoacoustic images during CA and resuscitation. Local vascular changes were accessed using photoacoustic imaging. The arteries and branches were not perfused with blood during CA (B) compared to baseline (A) . All the arteries and branches were perfused immediately after resuscitation, including even some tiny bridges between branches (C, arrows). However, these bridges disappeared late (D) due to hypoperfusion. The bar shows the sO2 level. Please click here to view a larger version of this figure.
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Discussion
In experimental CA studies, asphyxia, potassium chloride injections, or electrical current-derived ventricular fibrillation have been used to induce CA16,17,18,19,20,21,22,23. Normally, CPR is required for resuscitation in these CA models, especially in mice. We have formulated a resuscitation mixture that enables spontaneous resuscitation after asphyxia CA in mice. Eliminating the CPR step opens more opportunities for monitoring brain physiology during CA and resuscitation using current imaging modalities.
This resuscitation cocktail stock solution includes sodium bicarbonate, heparin, oxygenated arterial blood, and epinephrine. It is well known that CA induces both metabolic and respiratory acidosis. Sodium bicarbonate is expected to normalize the pH in the blood. Heparin is an anticoagulant and is used to prevent harmful clot formation during reperfusion. Oxygenated blood and epinephrine are the most critical components for resuscitation in this model. Although the exact mechanisms that underpin this spontaneous resuscitation are still unknown, it is speculated that when an adequate amount of oxygenated blood reaches the coronary arteries, thus delivering oxygen and epinephrine, the restoration of the myocardial contractility and the generation of cardiac output can be achieved without chest compressions. In this process, the infusion pressure, which is only achievable in the non-collapsed and thicker-walled arterial vasculature, is critical, since this facilitates the delivery of oxygenated blood to the heart. In support of this notion, we found that infusing the same mixture via the femoral vein did not result in the restoration of cardiac function, and resuscitation could not be achieved. Therefore, this resuscitation cocktail must be administered through the arterial line to achieve the restoration of cardiac function without chest compressions.
The dosage of epinephrine used in the current model is similar to what is used in standard CA experiments. Each milliliter of resuscitation cocktail stock solution contains 400 µg of epinephrine. The syringe is prepared with 26 µL of resuscitation cocktail stock solution, and arterial blood is withdrawn to 200 µL in the syringe. As the 1 mL plastic syringe has a 60 µL dead space in the front end, the blood remaining in the syringe following resuscitation is 60 µL, which includes 6 µL of resuscitation cocktail stock solution. Thus, the final injected resuscitation cocktail stock solution is 20 µL in each mouse, representing a dose of 8 µg of epinephrine in this procedure. In this protocol, the amount of resuscitation solution is not adjusted according to the body weight, similar to in clinical settings. We have not experienced any resuscitation issues in mice with a body weight of 20-32 g.
Of note, this resuscitation protocol was used successfully only in this asphyxia CA model. In our pilot study, this protocol failed to resuscitate mice after KCl-induced CA. Thus, the model described here is specifically useful for studying the brain physiology of asphyxia CA.
In summary, since this model does not require chest compressions during resuscitation, 1) the mouse can be kept in the prone position, and 2) the head can be mounted in a stereotaxic head frame, allowing imaging and electrophysiologic measurements without any movement during the entire recording phase. This perfectly fits the requirements for imaging/monitoring brain physiology during CA and resuscitation. This model has been successfully used in experiments that aim to dynamically track cerebral blood flow, vascular responses, and brain tissue oxygen in CA mice, and these experiments have generated invaluable data on vascular changes and responses to drug administration in CA.
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Disclosures
The authors have no conflicts of interest.
Acknowledgments
The authors thank Kathy Gage for her editorial support. This study was supported by funds from the Department of Anesthesiology (Duke University Medical Center), American Heart Association grant (18CSA34080277), and National Institutes of Health (NIH) grants (NS099590, HL157354, NS117973, and NS127163).
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 |
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