Normothermic Cardiac Arrest and Cardiopulmonary Resuscitation: A Mouse Model of Ischemia-Reperfusion Injury

The overall aim of this procedure is to model Norm Themic global Ischemia reperfusion to assess the effect of interventions on this important and common disease state. First, an anesthetized instrumented mouse is prepared for cardiac arrest and resuscitation or ca CPR. Next cardiac arrest is induced.

Then the mouse is resuscitated from cardiac arrest using chest compressions and epinephrine. Ultimately, functional assays such as blood urea, nitrogen, serum creatinine, Alan Transferases, aspartate to minor transferases and histology can be performed to assess the substantial organ damage. The presence of early biomarkers such as neutrophil gelatin associated lipo callin can be measured as well.

Finally, outcome of the resuscitation is evaluated here, demonstrated as a 24 hour trans cardio, perfusion and renal harvest. The main advantage of this model over our other methods, such as larger animal models of cardiac arrest and cardiopulmonary resuscitation, is that laboratory mice are low cost, ubiquitous and available in many transgenic strains. To begin, lubricate the eyes of an anesthetized mouse and place the animal in a supine position on a heating pad.

Then immobilize the animal's extremities using tape, placing the hind pores in a neutral position, but securing the four pores as near to the chest wall as possible to allow full chest wall excursion during chest compressions. Next lubricate and place a rectal temperature probe. Intubate the trachea using a 22 gauge Teflon catheter and an angled introducer.

Endotracheal positioning of the tube may be confirmed using either positive or negative pressure. Using positive pressure, a small volume of air is forced into the tube. If the tube is trache rather than esophageal placed, the chest rises symmetrically using negative pressure.

A small amount of fluid is placed in clear tubing, which is attached to the endotracheal tube. Spontaneous respiratory effort by the mouse moves the fluid within the tubing. Secure the endotracheal catheter with a loop of wire to the incisor, maintaining a slight tension on the incisor to keep the head immobilized during chest compressions.

Mechanically ventilate the mouse with the rodent ventilator set to 140 microliters 150 breaths per minute. Using sterile technique and an operating microscope, place a pre flushed PE 10 catheter in the jugular vein. Secure the PE 10 catheter into the skin closure with cyanoacrylate surgical adhesive.

Place three subcutaneous EKG electrodes, one near each axi, and one in the left lower quadrant of the abdomen. Ensure that all the wires are secure to the operating surface. Minimize the signal crossings and minimize the insulators within the signal path.

Once connected, optimize the EKG signal on the monitor. Ensure that the mouse is norm themic. Administer 40 microliters of room temperature, 0.5 molar potassium chloride intravenously, and observe the isoelectric tracing on the EKG.

Start the arrest timer. Next, disconnect the ventilator and discontinue the anesthetic vapor. Turn off the heating pad and any other equipment that produces electronic noise and may interfere with the EKG monitoring.

Place an insulating blanket over the mouse. Record the temperature every minute during cardiac arrest. If necessary, use a heating lamp to bring the core temperature up to the normal themic range.

After seven minutes and 30 seconds of cardiac arrest, reconnect the ventilator and increase the rate to 180 breaths per minute. Performing chest compressions to return spontaneous circulation is the trickiest part of this model. The mouse is quite small, so positioning and pressure are critical.

Too much pressure will result in lung and hepatic injury and reduce survival. Too little pressure will reduce the probability of return of spontaneous circulation. The chest should be compressed from one third to one half of the ANA row.

Posterior distance and full recoil should be allowed between compressions. Failure to achieve survival in this model is almost always due to suboptimal CPR At eight minutes. Initiate chest compressions at 300 beats per minute.

Chest compression should be delivered with the index finger. Five millimeters above the xiphoid process and slightly to the left of the midline infuse. 0.5 milliliters of epinephrine diluted to 15 micrograms per milliliter.

In the first 30 seconds of the CPR, carefully observe the EKG for the return of spontaneous circulation or ROSC, frequent premature ventricular contractions and changes in EKG axis are observed in the first two minutes. ROSC and almost always resolve into steady sinus tachycardia. At two minutes, record the total time of resuscitation and epinephrine dose.

Record the temperature every minute for 10 minutes. After R-O-S-C-E-K-G leads can be removed When spontaneous respiration begins, usually within 12 to 50 minutes after ROSC, remove the jugular catheter and use direct pressure to obtain hemostasis, extubate the trachea when the spontaneous respiratory rate is greater than 60 per minute. Finally, place the mouse in a recovery cage on a temperature controlled surface set to 37 degrees Celsius for the first two hours post procedure or longer if needed for complete recovery from anesthesia, the cage may them be moved to standard postoperative housing conditions 24 hours after C-A-C-P-R anesthetize the mouse and perform trans cardio perfusion first with saline and then with formalin Following fixation, a laparotomy is performed to check for the adequacy of renal fixation.

Adequately perfused and fixed kidneys are well blanched. Cardiac arrest induces instant loss of perfusion. Pressure represented here as mean arterial pressure or map.

This loss of perfusion pressure results in near complete cessation of regional renal cortical blood flow or RR CBF throughout the period of cardiac arrest in the shaded area. As seen here, resuscitation with chest compressions and epinephrine returns map to normal and RR CBF steadily rises in the post resuscitation period. In this figure, it can be seen how 24 hours post procedure blood urea nitrogen or BUN serum creatinine and the extent of tubular cell death are all significantly elevated in animals having undergone ca CPR as compared with animals treated with a sham procedure ca CPR induces a pan organismal ischemic insult here, evidence by massive elevation of liver function enzymes, Alan Transferases or a LT and aspartate to minoro transferase or a ST in ca, CPR mice as compared with sham treated animals.

Here, a western blot performed using a polyclonal antibody to neutrophil gelatinous associated lipo callin or end gal. A sensitive indicator of renal ischemic injury is shown. Urine samples were obtained immediately prior to pre and 24 hours after C-A-C-P-R in four animals represented as A, B, C and D.This figure shows that N gal is massively upregulated in mouse urine after C-A-C-P-R.

This figure shows a hemat, toin, and eoin stain of a short axis hilar section of renal tissue 24 hours after ca CPR patchy, but clear damage to the medullary and cortico medullary tubules with tubular plugging can be seen a fluoro jade B stain of the same region in the same animal. 24 hours after C-A-C-P-R can be seen in this figure, the fluoro jade B stains the necrotic cells bright green, showing patchy cortico medullary tubulin necrosis. These findings are substantially similar to renal biopsy findings from humans who develop acute kidney injury or a KI and unlike those produced by other animal models of a KI, While attempting this procedure, it's important to remember to carefully position the mouse prior to a rest to minimize EKG signal noise, and to deliver chest compressions with optimal pressure.