The authors want to thank Dagmar Dirvonskis for excellent technical support.

The experiments in this protocol were approved by the State and Institutional Animal Care Committee (Landesuntersuchungsamt Rheinland-Pfalz, Koblenz, Germany; Chairperson: Dr. Silvia Eisch-Wolf; approval no. G16-1-042). The experiments were conducted in accordance with the ARRIVE guidelines. Seven anesthetized male pigs (sus scrofa domestica) with a mean weight of 30 &#177; 2 kg and 12-16 weeks in age were included in the protocol.
1. Anesthesia, intubation, and mechanical ventilation13,14
<ol>
	<li>Maintain animals in their normal environment as long as possible to minimize stress. Withhold food 6 h before the scheduled experiment to reduce the risk of aspiration, but do not refuse water access.</li>
	<li>Sedate pigs with a combined injection of ketamine (4 mg/kg) and azaperone (8 mg/kg) in the neck or gluteal muscle with a needle (20 G) for intramuscular injection. Leave the animals undisturbed in their stables until sedation sets in (15-20 min). 
	CAUTION: Gloves are absolutely necessary when handling animals.</li>
	<li>Transport the sedated animals to the laboratory. Transport time should not exceed effective sedation time (here, 30-60 min).</li>
	<li>Monitor the peripheral oxygen saturation (SpO2) with a sensor clipped to the tail or ear.</li>
	<li>Disinfect the skin with an alcoholic disinfectant before insertion of a peripheral vein catheter (20 G) into an ear vein. Spray the area, wipe 1x, spray again, and let the disinfectant dry.</li>
	<li>Administer analgesia via intravenous injection of fentanyl (4 &#181;g/kg). Induce anesthesia with intravenous injection of propofol (3 mg/kg)</li>
	<li>Place the pig in supine position on a stretcher with a vacuum mattress and fix it with bandages. Apply muscle relaxant via intravenous injection of atracurium (0.5 mg/kg)</li>
	<li>Directly start noninvasive ventilation with a dog ventilation mask (size 2). Ventilation parameters are as follows: FiO2 (inspiratory oxygen fraction) = 100%, respiratory rate = 18-20 breaths/min, peak inspiratory pressure = &#60;20 cmH20, PEEP (positive end-expiratory pressure) = 5 cmH20.</li>
	<li>Maintain anesthesia via continuous infusion of fentanyl (0.1-0.2 mg kg-1 h-1) and propofol (8-12 mg kg-1 h-1). Start a continuous infusion of balanced electrolyte solution (5 mL kg-1 h-1).</li>
	<li>Secure the airway via intubation with a common endotracheal tube (ID 6-7) and an introducer. Use a common laryngoscope with a Macintosh blade (size 4). Two people are necessary for this step.
	<ol>
		<li>Ensure that one person fixes the tongue outside with a piece of tissue and opens the snout with the other hand.
		<ol>
			<li>Ensure that the second person performs a laryngoscopy of the porcine larynx. When the epiglottis comes into view, move the laryngoscope ventrally. The epiglottis should be lifted up and the vocal cords will be visible. 
			NOTE: If the epiglottis does not move ventrally, it will stick to the soft palatine and can be mobilized by the tip of the tube.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Move the tube carefully through the vocal cords. 
	NOTE: The narrowest point of the trachea is not on the level of the vocal cords but is subglottic. If tube insertion is not possible, try to rotate the tube clockwise or use a smaller tube.</li>
	<li>Pull the introducer out of the tube. Use a 10 mL syringe to block cuff with 10 mL of air. Control the cuff pressure with a cuff manager (30 cmH2O).</li>
	<li>Start mechanical ventilation after tube connection with a ventilator (PEEP = 5 cmH2O, tidal volume = 8 mL/kg, FiO2 = 0.4, I:E [inspiration to expiration ratio] = 1:2, respiratory rate = variable to achieve an end-tidal CO2 of &#60;6 kPa, usually 20-30/min). Make sure that tube position is correct by regular and periodic exhalation of carbon dioxide via capnography.</li>
	<li>Check double-sided ventilation via auscultation. 
	NOTE: In case of incorrect placement of the tube, an air-filled stomach rapidly forms a clearly visible bulge through the abdominal wall. In this case, immediate replacement of the tube and insertion of a gastric tube is necessary. If intubation is not successful, switch back to mask ventilation and try a smaller tube or better positioning of the snout.</li>
	<li>Place gastric tube into the stomach to avoid reflux and vomiting with two people.
	<ol>
		<li>Fix the tongue outside with a piece of tissue and open the snout with the other hand.
		<ol>
			<li>Ensure that a second person performs a laryngoscopy of the porcine larynx then visualizes the esophagus. Push the gastric tube inside the esophagus with a Magill&#39;s forceps until gastric fluid is drained. 
			NOTE: Visualization may be difficult. In this case, lift the tube with the laryngoscope ventrally to open the esophagus.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Instrumentation
<ol>
	<li>Use bandages to pull back the hindlegs to smooth the folds in the femoral area for vessel catherization.</li>
	<li>Prepare the following materials: syringes (5 mL, 10 mL, and 50 mL), Seldinger needle, introducer sheaths (6 Fr, 8 Fr, 8 Fr), guidewires for the sheaths, central venous catheter with three ports (7 Fr, 30 cm) with guidewire, cardiac output monitor (Table of Materials), and a catheter (5 Fr, 20 cm).</li>
	<li>Disinfect the inguinal area (see step 1.6). Repeat this process 2x.</li>
	<li>Fill all catheters with saline solution. Apply ultrasound gel on the ultrasound probe. Cover the inguinal area with a sterile fenestrated drape.</li>
	<li>Scan the right femoral vessels with ultrasound and use doppler technique to identify the artery and vein15. Visualize the right femoral artery axially. Switch to a longitudinal view of the arteria by rotating the probe 90&#176;.</li>
	<li>Puncture the right femoral artery under ultrasound visualization with the Seldinger needle under permanent aspiration with the 5 mL syringe. 
	NOTE: In our opinion, the ultrasound guided Seldinger&#39;s technique is associated with significantly less blood loss and tissue trauma than other methods of vascular access.</li>
	<li>Confirm the desired needle position by observing bright red pulsating blood. Disconnect the syringe and quickly insert the guidewire into the right femoral artery.</li>
	<li>Visualize the longitudinal axis of the right femoral vein. Insert the Seldinger needle under permanent aspiration with the 5 mL syringe. Aspirate any dark red non-pulsating venous blood. 
	NOTE: If the correct position of the needle in the different vessels cannot be visually confirmed, take blood samples and analyze the blood gas content. A high oxygen level is a good sign for arterial blood, while low oxygen saturation indicates intravenous position.</li>
	<li>Insert the guidewire for the central venous catheter into the right femoral vein after disconnecting the syringe. Retract the Seldinger needle.</li>
	<li>Visualize both right vessels using ultrasound to control the correct wire position. Push the arterial introducer sheath (6 Fr) over the guidewire into the right artery and secure the position with blood aspiration. 
	NOTE: Placing the sheath through the skin can be difficult. It can be helpful to perform a small incision along the wire to facilitate better placement.</li>
	<li>Use Seldinger&#39;s technique to position the central venous line into the right femoral vein. Aspirate all ports and flush them with saline solution.</li>
	<li>Perform the same procedure on the left inguinal side to insert the other introducer sheaths in Seldinger&#39;s technique into the left femoral artery (8 Fr) and femoral vein (8 Fr).</li>
	<li>Connect the right arterial introducer sheath and the central venous catheter with two transducer systems for measurement of invasive hemodynamics. Position both transducers at heart level.</li>
	<li>Switch the tree-way stopcocks of both transducers open to the atmosphere to calibrate the system to zero. 
	NOTE: It is necessary to avoid any air bubbles and bloodstains in the system to generate plausible values.</li>
	<li>Switch all infusions for maintaining anesthesia from the peripheral vein to a central venous line. Take baseline values (hemodynamics, spirometrics, and other output from the cardiac monitor; see section 3) after a 15 min recovery.</li>
	<li>Initiate ventricular fibrillation (see section 4).</li>
</ol>
3. Pulse contour cardiac output
<ol>
	<li>Insert transpulmonal thermodilution catheter into the right arterial introducer sheath. 
	NOTE: In clinical medicine, thermodilution catheters are directly placed by Seldinger&#39;s technique. However, placement via an introducer sheath is also feasible. In the proposed protocol, sheaths are placed as a standardized vascular access for maximum flexibility in instrumentation throughout different experiments.</li>
	<li>Connect the catheter with the arterial wire of the cardiac monitor system. Switch the arterial transducer directly with the cardiac monitor port and recalibrate as described in step 2.14. Connect the venous measuring unit of the cardiac monitor system with the left venous introducer sheath. 
	NOTE: It is necessary to connect the venous and arterial probes as far apart as possible; otherwise, the measurement will be disturbed, because the application of cold water into the venous system will affect the arterial measurement. More details regarding PiCCO2 have been provided previously16.</li>
	<li>Turn on the cardiac monitor system. Confirm that a new patient is being measured. Enter the size and weight.</li>
	<li>Switch the category to adults. Enter the protocol name and ID. Click on Exit.</li>
	<li>Set the injection volume to 10 mL. 
	NOTE: The volume of chosen injection solution can be changed in the software. A higher volume makes the measured values more valid. A small volume was chosen for this experiment to avoid any hemodilution effects.</li>
	<li>Enter the central venous pressure.</li>
	<li>Open the three-way stopcock to the atmosphere.</li>
	<li>Click on Zero for system calibration. Click on Exit.</li>
	<li>Calibrate the continuous cardiac output measurement.
	<ol>
		<li>Click on TD (thermodilution). Prepare a physiological saline solution with a temperature of 4 &#176;C in a 10 mL syringe. Click on Start.</li>
		<li>Inject 10 mL of cold saline solution quickly and steadily into the venous measuring unit. Wait until the measurement is completed and the system requests a repetition.</li>
		<li>Repeat the previous step until three measurements are completed. The system will calculate the mean of all parameters. Click on Exit. 
		NOTE: Measurements will start immediately after calibration has been completed. Although cardiac output measurements during CPR are not performed regularly, plausible results have been able to be affirmed after adequate calibration17,18.</li>
	</ol>
	</li>
</ol>
4. Ventricular fibrillation and mechanical resuscitation
<ol>
	<li>Place defibrillator patch electrodes in anterior-posterior position on the torso. The posterior electrode should be positioned on the central left hemithorax. 
	NOTE: Use a razor to remove excess hair and dirt to facilitate optimal conduction.</li>
	<li>Connect the electrodes to a defibrillator and establish an ECG.</li>
	<li>Immobilize the pig inside the vacuum mattress. Deflate the mattress to prevent unwanted movement during CPR. Control fixation of the limbs.</li>
	<li>Place chest compression device (here, LUCAS-2) around the chest and under the vacuum mattress according to the manufacturer&#39;s recommendations. Adjust the pressure pad to the lower third of the sternum in median position.</li>
	<li>Turn on chest compression device (&#34;power&#34; button) and lower the pressure pad to skin level. Set the compression frequency to 100/min, if not otherwise defined in the protocol. Press the Pause button to prepare the compression device for chest compressions.</li>
	<li>Insert a fibrillation/pacing catheter into the left femoral vein through the i.v. sheath.</li>
	<li>Inflate the catheter cuff with 1-2 mL of air. Slowly push the inflated cuff further until it is placed next to the right atrium (usually about a 50 cm distance).</li>
	<li>Connect catheter electrodes to an adequate oscilloscope/function generator. Adjust fibrillation parameters to the desired values (here, a 13.8 V current with frequencies between 50-200 Hz).</li>
	<li>Turn on generator and monitor ECG changes. Move the catheter slowly forward until arrhythmias can be detected in the ECG. 
	CAUTION: Prevent the separate electrodes at the end of the catheter from touching human skin or each other to prevent short circuits and possibly life-threatening situations.</li>
	<li>Carefully vary the catheter position until ventricular fibrillation can be detected. 
	NOTE: It can be difficult to induce fibrillation right away. If a position is reached at which ECG effects can be seen, changing the frequency or repeatedly turning the generator on and off can sometimes be helpful.</li>
	<li>Once ventricular fibrillation is confirmed, turn off the generator, deflate the balloon, and remove the fibrillation catheter. Maintain fibrillation with or without ventilation for as long as required.</li>
	<li>Start mechanical chest compressions by pressing the Play button on the compression device. To interrupt chest compressions, press the Pause button on the compression device.</li>
	<li>Analyze ECG patterns. If ventricular fibrillation persists, prepare defibrillation.
	<ol>
		<li>Enter Manual mode in the defibrillator menu. Adjust the energy to 200 J biphasic.</li>
		<li>Press the Load button. Wait until acoustic signal turns on to indicate a prepared shock value. Initiate electric shock. 
		CAUTION: Only experienced users should handle defibrillators and fibrillation catheters. No shocks should be initiated if there is any indication for faulty or worn materials. The initiation of an electric shock must always be announced clearly audible to every person in the room, and the person launching the defibrillation is responsible for ensuring that nobody is touching the animal or stretcher prior to releasing the shock. 
		NOTE: Here, guideline-based resuscitation protocol was used (i.e., 2 min of chest compressions, ECG assessment, shock, 2 min of chest compressions, adrenaline administration, etc.). For more information, consult with the guidelines4.</li>
	</ol>
	</li>
	<li>In the case of return of spontaneous circulation (ROSC), stop chest compressions, continue ventilation, and apply monitoring as extensively and for as long as needed. 
	NOTE: Anesthetic drug administration may or may not be interrupted during CPR, depending on the protocol. If sedation is discontinued, infusion should be restarted upon confirmed ROSC.</li>
	<li>A goal-directed approach for the guidance of fluid and catecholamine administration as well as standardized respiratory and ventilation settings are recommended to prevent cardiorespiratory deterioration in the ROSC phase leading to experimental failure.</li>
</ol>
5. End of experiment and euthanasia (in the case of ROSC)
<ol>
	<li>Inject 0.5 mg of fentanyl into the central venous line. Wait 5 min. Inject 200 mg of propofol into the central venous line.</li>
	<li>Euthanize the animal with a 40 mmol potassium chloride injection.</li>
	<li>Perform organ removal/fixation or analyses as needed.</li>
</ol>

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K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+inhaled+tumor+necrosis+factor-alpha-derived+TIP+peptide+improves+the+pulmonary+function+in+experimental+lung+injury.">An inhaled tumor necrosis factor-alpha-derived TIP peptide improves the pulmonary function in experimental lung injury.</a> Acta Anaesthesiol Scand. 57 (3), 334-341 (2013).</li><li>Ziebart, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Low+tidal+volume+pressure+support+versus+controlled+ventilation+in+early+experimental+sepsis+in+pigs.">Low tidal volume pressure support versus controlled ventilation in early experimental sepsis in pigs.</a> Respiratory Research. 15, 101 (2014).</li><li>Tan, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Duration+of+cardiac+arrest+requires+different+ventilation+volumes+during+cardiopulmonary+resuscitation+in+a+pig+model.">Duration of cardiac arrest requires different ventilation volumes during cardiopulmonary resuscitation in a pig model.</a> Journal of Clinical Monitoring and Computing. , (2019).</li><li>Kill, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanical+ventilation+during+cardiopulmonary+resuscitation+with+intermittent+positive-pressure+ventilation,+bilevel+ventilation,+or+chest+compression+synchronized+ventilation+in+a+pig+model.">Mechanical ventilation during cardiopulmonary resuscitation with intermittent positive-pressure ventilation, bilevel ventilation, or chest compression synchronized ventilation in a pig model.</a> Critical Care Medicine. 42 (2), e89-e95 (2014).</li><li>Speer, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanical+Ventilation+During+Resuscitation:+How+Manual+Chest+Compressions+Affect+a+Ventilator's+Function.">Mechanical Ventilation During Resuscitation: How Manual Chest Compressions Affect a Ventilator's Function.</a> Advances in Therapy. 34 (10), 2333-2344 (2017).</li><li>Kill, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chest+Compression+Synchronized+Ventilation+versus+Intermitted+Positive+Pressure+Ventilation+during+Cardiopulmonary+Resuscitation+in+a+Pig+Model.">Chest Compression Synchronized Ventilation versus Intermitted Positive Pressure Ventilation during Cardiopulmonary Resuscitation in a Pig Model.</a> PLoS ONE. 10 (5), e0127759 (2015).</li><li>Newell, C., Grier, S., Soar, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Airway+and+ventilation+management+during+cardiopulmonary+resuscitation+and+after+successful+resuscitation.">Airway and ventilation management during cardiopulmonary resuscitation and after successful resuscitation.</a> Critical Care. 22 (1), 190 (2018).</li></ol>

The LUCAS-2 device was provided unconditionally by Stryker/Physio-Control, Redmond, WA, USA for experimental research purposes. No authors report any conflicts of interest.

Some major technical issues regarding anesthesia in a porcine model have previously been described by our group13,14. These include the strict avoidance of stress and unnecessary pain for the animals, possible anatomical problems during airway management, and specific personnel requirements19.
Additionally, the benefits of ultrasound-guided catheterization was highlighted previously and remains the preferable ap...

Cardiac arrest and cardiopulmonary resuscitation (CPR) are regularly encountered medical emergencies in hospital wards as well as preclinical emergency provider scenarios1,2. While there have been extensive efforts to characterize the optimal treatment for this situation3,4,5,6, international guidelines and expert recommendations (e.g., ERC and ILCOR) usually rely on low-grade evidence due to the lack of prospective randomized trials3,4,5,7,8,9. This is in part due to obvious ethical reservations regarding randomized resuscitation protocols in human trials10. However, this may also point towards a lack of strict protocol adherence when confronted with a life-threatening and stressful situation11,12. The protocol presented in this report aims to provide a standardized resuscitation model in a realistic clinical setting, which generates valuable, prospective data while being as valid and accurate as possible without the need for human subjects. It adheres to common resuscitation guidelines, can be easily applied, and enables researches to examine and characterize various aspects and interventions in a critical but controlled setting. This will lead to 1) a better understanding of the pathological mechanisms underlying cardiac arrest and ventricular fibrillation and 2) higher quality evidence in order to optimize treatment options and increase survival rates.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1 M- Kaliumchlorid-L&#246;sung 7,46% 20ml</td>
			<td>Fresenius, Kabi Deutschland GmbH</td>
			<td></td>
			<td>potassium chloride</td>
		</tr>
		<tr>
			<td>Arterenol 1mg/ml 25 ml</td>
			<td>Sanofi- Aventis, Seutschland GmbH</td>
			<td></td>
			<td>norepinephrine</td>
		</tr>
		<tr>
			<td>Atracurium Hikma 50mg/5ml</td>
			<td>Hikma Pharma GmbH, Martinsried</td>
			<td></td>
			<td>atracurium</td>
		</tr>
		<tr>
			<td>BD Discardit II Spritze 2,5,10,20 ml</td>
			<td>Becton Dickinson S.A. Carretera Mequinenza Fraga, Spain</td>
			<td></td>
			<td>syringe</td>
		</tr>
		<tr>
			<td>BD Luer Connecta</td>
			<td>Becton Dickinson Infusion Therapy AB Helsingborg, Schweden</td>
			<td></td>
			<td>3-way-stopcock</td>
		</tr>
		<tr>
			<td>BD Microlance 3 20 G</td>
			<td>Becton Dickinson S.A. Carretera Mequinenza Fraga, Spain</td>
			<td></td>
			<td>canula</td>
		</tr>
		<tr>
			<td>CorPatch Easy Electrodes</td>
			<td>CorPuls, Kaufering, Germany</td>
			<td></td>
			<td>defibrillator electrodes</td>
		</tr>
		<tr>
			<td>Corpuls 3</td>
			<td>Corpuls, Kaufering, Germany</td>
			<td></td>
			<td>defibrillator</td>
		</tr>
		<tr>
			<td>Datex Ohmeda S5</td>
			<td>GE Healthcare Finland Oy, Helsinki, Finland</td>
			<td></td>
			<td>hemodynamic monitor</td>
		</tr>
		<tr>
			<td>Engstr&#246;m Carestation</td>
			<td>GE Heathcare, Madison USA</td>
			<td></td>
			<td>ventilator</td>
		</tr>
		<tr>
			<td>Fentanyl-Janssen 0,05mg/ml</td>
			<td>Janssen-Cilag GmbH, Neuss</td>
			<td></td>
			<td>fentanyl</td>
		</tr>
		<tr>
			<td>F&#252;hrungsstab, Durchmesser 4.3</td>
			<td>R&#252;sch</td>
			<td></td>
			<td>endotracheal tube introducer</td>
		</tr>
		<tr>
			<td>Incetomat-line 150 cm</td>
			<td>Fresenius, Kabi Deutschland GmbH</td>
			<td></td>
			<td>perfusorline</td>
		</tr>
		<tr>
			<td>Ketamin-Hameln 50mg/ml</td>
			<td>Hameln Pharmaceuticals GmbH</td>
			<td></td>
			<td>ketamine</td>
		</tr>
		<tr>
			<td>laryngoscope</td>
			<td>R&#252;sch</td>
			<td></td>
			<td>laryngoscope</td>
		</tr>
		<tr>
			<td>logicath 7 Fr 3-lumen 30cm lang</td>
			<td>Smith- Medical Deutschland GmbH</td>
			<td></td>
			<td>central venous catheter</td>
		</tr>
		<tr>
			<td>LUCAS-2</td>
			<td>Physio-Control/Stryker, Redmond, WA, USA</td>
			<td></td>
			<td>chest compression device</td>
		</tr>
		<tr>
			<td>Masimo Radical 7</td>
			<td>Masimo Corporation Irvine, Ca 92618 USA</td>
			<td></td>
			<td>periphereal oxygen saturation</td>
		</tr>
		<tr>
			<td>Neofox Oxygen sensor 300 micron fiber</td>
			<td>Ocean optics Largo, FL USA</td>
			<td></td>
			<td>ultrafast pO2-measurements</td>
		</tr>
		<tr>
			<td>&#214;ls&#228;ure reinst Ph. Eur NF C18H34O2 M0282,47g/mol Dichte 0,9</td>
			<td>Applichem GmbH Darmstadt, Deutschland</td>
			<td></td>
			<td>oleic acid</td>
		</tr>
		<tr>
			<td>Original Perfusor syringe 50ml Luer Lock</td>
			<td>B.Braun Melsungen AG, Germany</td>
			<td></td>
			<td>perfusorsyringe</td>
		</tr>
		<tr>
			<td>Osypka pace, 110 cm</td>
			<td>Osypka Medical GmbH, Rheinfelden-Herten, Germany</td>
			<td></td>
			<td>Pacing/fibrillation catheter</td>
		</tr>
		<tr>
			<td>PA-Katheter Swan Ganz 7,5 Fr 110cm</td>
			<td>Edwards Lifesciences LLC, Irvine CA, USA</td>
			<td></td>
			<td>PAC</td>
		</tr>
		<tr>
			<td>Percutaneous sheath introducer set 8,5 und 9 Fr, 10 cm with integral haemostasis valve/sideport</td>
			<td>Arrow international inc. Reading, PA, USA</td>
			<td></td>
			<td>introducer sheath</td>
		</tr>
		<tr>
			<td>Perfusor FM Braun</td>
			<td>B.Braun Melsungen AG, Germany</td>
			<td></td>
			<td>syringe pump</td>
		</tr>
		<tr>
			<td>Propofol 2% 20mg/ml (50ml flasks)</td>
			<td>Fresenius, Kabi Deutschland GmbH</td>
			<td></td>
			<td>propofol</td>
		</tr>
		<tr>
			<td>Radifocus Introducer II, 5-8 Fr</td>
			<td>Terumo Corporation Tokio, Japan</td>
			<td></td>
			<td>introducer sheath</td>
		</tr>
		<tr>
			<td>R&#252;schelit Super Safety Clear &#62;ID 6/ 6,5 /7,0 mm</td>
			<td>Teleflex Medical Sdn. Bhd, Malaysia</td>
			<td></td>
			<td>endotracheal tube</td>
		</tr>
		<tr>
			<td>Seldinger Nadel mit Fixierfl&#252;gel</td>
			<td>Smith- Medical Deutschland GmbH</td>
			<td></td>
			<td>seldinger canula</td>
		</tr>
		<tr>
			<td>Sonosite Micromaxx Ultrasoundsystem</td>
			<td>Sonosite Bothell, WA, USA</td>
			<td></td>
			<td>ultrasound</td>
		</tr>
		<tr>
			<td>Stainless Macintosh Gr&#246;&#223;e 4</td>
			<td>Welsch Allyn69604</td>
			<td></td>
			<td>blade for laryngoscope</td>
		</tr>
		<tr>
			<td>Stresnil 40mg/ml</td>
			<td>Lilly Deutschland GmbH, Abteilung Elanco Animal Health</td>
			<td></td>
			<td>azaperone</td>
		</tr>
		<tr>
			<td>Vasofix Safety 22G-16G</td>
			<td>B.Braun Melsungen AG, Germany</td>
			<td></td>
			<td>venous catheter</td>
		</tr>
		<tr>
			<td>Voltcraft Model 8202</td>
			<td>Voltcraft, Hirschau, Germany</td>
			<td></td>
			<td>oscilloscope/function generator</td>
		</tr>
	</tbody>
</table>

standardized model of ventricular fibrillation and advanced cardiac life support in swine

Cardiopulmonary resuscitation after cardiac arrest, independent of its origin, is a regularly encountered medical emergency in hospitals as well as preclinical settings. Prospective randomized trials in human subjects are difficult to design and ethically ambiguous, which results in a lack of evidence-based therapies. The model presented in this report represents one of the most common causes of cardiac arrests, ventricular fibrillation, in a standardized setting in a large animal model. This allows for reproducible observations and various therapeutic interventions under clinically accurate conditions, hence facilitating the generation of better evidence and eventually the potential for improved medical treatment.

Cardiac arrest was induced in seven pigs. Return of spontaneous circulation following CPR was achieved in four Pigs (57%) with a mean of 3 &#177; 1 biphasic defibrillations. Healthy and adequately anesthetized pigs should stay in supine position without shivering and signs of agitation throughout the entire experiment. Mean arterial blood pressures should not drop below 50 mmHg before initiation of fibrillation18. For optimal results, blood gas analyses can be perf...

Watch this Scientific Journal Video about Standardized Model of Ventricular Fibrillation and Advanced Cardiac Life Support in Swine at JoVE.com

Standardized Model of Ventricular Fibrillation and Advanced Cardiac Life Support in Swine

Cardiopulmonary resuscitation and defibrillation are the only effective therapeutic options during cardiac arrest caused by ventricular fibrillation. This model presents a standardized regimen to induce, assess, and treat this physiological state in a porcine model, thus providing a clinical approach with various opportunities for data collection and analysis.

Cardiopulmonary resuscitation after cardiac arrest, independent of its origin, is a regularly encountered medical emergency in hospitals as well as ...

standardized-model-ventricular-fibrillation-advanced-cardiac-life

Research

JoVE Journal

Medicine

7.6K Views.  University Medical Center of the Johannes Gutenberg-University. Cardiopulmonary resuscitation and defibrillation are the only effective therapeutic options during cardiac arrest caused by ventricular fibrillation. This model presents a standardized regimen to induce, assess, and treat this physiological state in a porcine model, thus providing a clinical approach with various opportunities for data collection and analysis.

Video: Standardized Model of Ventricular Fibrillation and Advanced Cardiac Life Support in Swine

A Swine Model of Neonatal Asphyxia

Annually more than 1 million neonates die worldwide as related to asphyxia. Asphyxiated neonates commonly have multi-organ failure including hypotension, perfusion deficit, hypoxic-ischemic encephalopathy, pulmonary hypertension, vasculopathic enterocolitis, renal failure and thrombo-embolic complications. Animal models are developed to help us understand the patho-physiology and pharmacology of neonatal asphyxia. In comparison to rodents and newborn lambs, the newborn piglet has been proven to be a valuable model. The newborn piglet has several advantages including similar development as that of 36-38 weeks human fetus with comparable body systems, large body size (&tilde;1.5-2 kg at birth) that allows the instrumentation and monitoring of the animal and controls the confounding variables of hypoxia and hemodynamic derangements.
We here describe an experimental protocol to simulate neonatal asphyxia and allow us to examine the systemic and regional hemodynamic changes during the asphyxiating and reoxygenation process as well as the respective effects of interventions. Further, the model has the advantage of studying multi-organ failure or dysfunction simultaneously and the interaction with various body systems. The experimental model is a non-survival procedure that involves the surgical instrumentation of newborn piglets (1-3 day-old and 1.5-2.5 kg weight, mixed breed) to allow the establishment of mechanical ventilation, vascular (arterial and central venous) access and the placement of catheters and flow probes (Transonic Inc.) for the continuously monitoring of intra-vascular pressure and blood flow across different arteries including main pulmonary, common carotid, superior mesenteric and left renal arteries. Using these surgically instrumented piglets, after stabilization for 30-60 minutes as defined by Z&lt;10% variation in hemodynamic parameters and normal blood gases, we commence an experimental protocol of severe hypoxemia which is induced via normocapnic alveolar hypoxia. The piglet is ventilated with 10-15% oxygen by increasing the inhaled concentration of nitrogen gas for 2h, aiming for arterial oxygen saturations of 30-40%. This degree of hypoxemia will produce clinical asphyxia with severe metabolic acidosis, systemic hypotension and cardiogenic shock with hypoperfusion to vital organs. The hypoxia is followed by reoxygenation with 100% oxygen for 0.5h and then 21% oxygen for 3.5h. Pharmacologic interventions can be introduced in due course and their effects investigated in a blinded, block-randomized fashion.

Large animal models have good translational values in the examination of physiology and pharmacology of neonatal asphyxia. Using newborn piglets, we develop an experimental protocol to simulate neonatal asphyxia which has advantages of studying the systemic and regional hemodynamics, oxygen transport with biochemical and pathologic pathways and correlations.

Annually more than 1 million neonates die worldwide as related to asphyxia. Asphyxiated neonates commonly have multi-organ failure including hypotension, ...

Assessment of Right Ventricular Structure and Function in Mouse Model of Pulmonary Artery Constriction by Transthoracic Echocardiography

Emerging clinical data support the notion that RV dysfunction is critical to the pathogenesis of cardiovascular disease and heart failure1-3. Moreover, the RV is significantly affected in pulmonary diseases such as pulmonary artery hypertension (PAH). In addition, the RV is remarkably sensitive to cardiac pathologies, including left ventricular (LV) dysfunction, valvular disease or RV infarction4. To understand the role of RV in the pathogenesis of cardiac diseases, a reliable and noninvasive method to access the RV structurally and functionally is essential.
A noninvasive trans-thoracic echocardiography (TTE) based methodology was established and validated for monitoring dynamic changes in RV structure and function in adult mice. To impose RV stress, we employed a surgical model of pulmonary artery constriction (PAC) and measured the RV response over a 7-day period using a high-frequency ultrasound microimaging system. Sham operated mice were used as controls. Images were acquired in lightly anesthetized mice at baseline (before surgery), day 0 (immediately post-surgery), day 3, and day 7 (post-surgery). Data was analyzed offline using software.
Several acoustic windows (B, M, and Color Doppler modes), which can be consistently obtained in mice, allowed for reliable and reproducible measurement of RV structure (including RV wall thickness, end-diastolic&#160;and end-systolic dimensions), and function (fractional area change, fractional shortening, PA peak velocity, and peak pressure gradient) in normal mice and following PAC.
Using this method, the pressure-gradient resulting from PAC was accurately measured in real-time using Color Doppler mode and was comparable to direct pressure measurements performed with a Millar high-fidelity microtip catheter. Taken together, these data demonstrate that RV measurements obtained from various complimentary views using echocardiography are reliable, reproducible and can provide insights regarding RV structure and function. This method will enable a better understanding of the role of RV cardiac dysfunction.

Right ventricle (RV) dysfunction is critical to the pathogenesis of cardiovascular disease, yet limited methodologies are available for its evaluation. Recent advances in ultrasound imaging provide a noninvasive and accurate option for longitudinal RV study. Herein, we detail a step-by-step echocardiographic method using a murine model of RV pressure overload.

Emerging clinical data support the notion that RV dysfunction is critical to the pathogenesis of cardiovascular disease and heart failure1-3. Moreover, ...

A Rat Model of Ventricular Fibrillation and Resuscitation by Conventional Closed-chest Technique

A rat model of electrically-induced ventricular fibrillation followed by cardiac resuscitation using a closed chest technique that incorporates the basic components of cardiopulmonary resuscitation in humans is herein described. The model was developed in 1988 and has been used in approximately 70 peer-reviewed publications examining a myriad of resuscitation aspects including its physiology and pathophysiology, determinants of resuscitability, pharmacologic interventions, and even the effects of cell therapies. The model featured in this presentation includes: (1) vascular catheterization to measure aortic and right atrial pressures, to measure cardiac output by thermodilution, and to electrically induce ventricular fibrillation; and (2) tracheal intubation for positive pressure ventilation with oxygen enriched gas and assessment of the end-tidal CO2. A typical sequence of intervention entails: (1) electrical induction of ventricular fibrillation, (2) chest compression using a mechanical piston device concomitantly with positive pressure ventilation delivering oxygen-enriched gas, (3) electrical shocks to terminate ventricular fibrillation and reestablish cardiac activity, (4) assessment of post-resuscitation hemodynamic and metabolic function, and (5) assessment of survival and recovery of organ function. A robust inventory of measurements is available that includes &#8211; but is not limited to &#8211; hemodynamic, metabolic, and tissue measurements. The model has been highly effective in developing new resuscitation concepts and examining novel therapeutic interventions before their testing in larger and translationally more relevant animal models of cardiac arrest and resuscitation.

This article describes a rat model of electrically-induced ventricular fibrillation and resuscitation by chest compression, ventilation, and delivery of electrical shocks that simulates an episode of sudden cardiac arrest and conventional cardiopulmonary resuscitation. The model enables gathering insights on the pathophysiology of cardiac arrest and exploration of new resuscitation strategies.

A rat model of electrically-induced ventricular fibrillation followed by cardiac resuscitation using a closed chest technique that incorporates the basic ...

A Chronic Cardiac Ischemia Model in Swine Using an Ameroid Constrictor

Cardiovascular disease remains the number one cause of mortality in the United States. There are numerous approaches to treating these diseases, but regardless of the approach, an in vivo model is needed to test each treatment. The pig is one of the most used large animal models for cardiovascular disease. Its heart is very similar in anatomy and function to that of a human. The ameroid placement technique creates an ischemic area of the heart, which has many useful applications in studying myocardial infarction. This model has been used for surgical research, pharmaceutical studies, imaging techniques, and cell therapies.
There are several ways of inducing an ischemic area in the heart. Each has its advantages and disadvantages, but the placement of an ameroid constrictor remains the most widely used technique. The main advantages to using the ameroid are its prevalence in existing research, its availability in various sizes to accommodate the anatomy and size of the vessel to be constricted, the surgery is a relatively simple procedure, and the post-operative monitoring is minimal, since there are no external devices to maintain. This paper provides a detailed overview of the proper technique for the placement of the ameroid constrictor.

The purpose of this protocol is to demonstrate the placement of a delayed constricting device (an ameroid constrictor) around a coronary artery in a swine model. This device creates an ischemic area of the heart that is useful for studying new diagnostic imaging techniques and new methods of treatment.

Cardiovascular disease remains the number one cause of mortality in the United States. There are numerous approaches to treating these diseases, but ...

Surgical Swine Model of Chronic Cardiac Ischemia Treated by Off-Pump Coronary Artery Bypass Graft Surgery

Chronic cardiac ischemia that impairs cardiac function, but does not result in infarct, is termed hibernating myocardium (HM). A large clinical subset of coronary artery disease (CAD) patients have HM, which in addition to causing impaired function, puts them at higher risk for arrhythmia and future cardiac events. The standard treatment for this condition is revascularization, but this has been shown to be an imperfect therapy. The majority of pre-clinical cardiac research focuses on infarct models of cardiac ischemia, leaving this subset of chronic ischemia patients largely underserved. To address this gap in research, we have developed a well-characterized and highly reproducible model of hibernating myocardium in swine, as swine are ideal translational models for human heart disease. In addition to creating this unique disease model, we have optimized a clinically relevant treatment model of coronary artery bypass surgery in swine. This allows us to accurately study the effects of bypass surgery on heart disease, as well as investigate additional or alternate therapies. This model surgically induces single vessel stenosis by implanting a constrictor on the left anterior descending (LAD) artery in a young pig. As the pig grows, the constrictor creates a gradual stenosis, resulting in chronic ischemia with impaired regional function, but preserving tissue viability. Following the establishment of the hibernating myocardium phenotype, we perform off-pump coronary artery bypass graft surgery to revascularize the ischemic region, mimicking the gold-standard treatment for patients in the clinic.

This protocol presents a surgical large animal model of chronic, single vessel ischemia that results in regional abnormalities but does not create infarct, known as hibernating myocardium. Following establishment of chronic ischemia, animals are treated with off-pump LIMA-LAD coronary artery bypass graft surgery to revascularize the ischemic tissue.

Chronic cardiac ischemia that impairs cardiac function, but does not result in infarct, is termed hibernating myocardium (HM). A large clinical subset of ...

Standardized Histomorphometric Evaluation of Osteoarthritis in a Surgical Mouse Model

One of the most prevalent joint disorders in the United States, osteoarthritis (OA) is characterized by progressive degeneration of articular cartilage, primarily in the hip and knee joints, which results in significant impacts on patient mobility and quality of life. To date, there are no existing curative therapies for OA able to slow down or inhibit cartilage degeneration. Presently, there is an extensive body of ongoing research to understand OA pathology and discover novel therapeutic approaches or agents that can efficiently slow down, stop, or even reverse OA. Thus, it is crucial to have a quantitative and reproducible approach to accurately evaluate OA-associated pathological changes in the joint cartilage, synovium, and subchondral bone. Currently, OA severity and progression are primarily assessed using the Osteoarthritis Research Society International (OARSI) or Mankin scoring systems. In spite of the importance of these scoring systems, they are semiquantitative and can be influenced by user subjectivity. More importantly, they fail to accurately evaluate subtle, yet important, changes in the cartilage during the early disease states or early treatment phases. The protocol we describe here uses a computerized and semiautomated histomorphometric software system to establish a standardized, rigorous, and reproducible quantitative methodology for the evaluation of joint changes in OA. This protocol presents a powerful addition to the existing systems and allows for more efficient detection of pathological changes in the joint.

The current protocol establishes a rigorous and reproducible method for quantification of morphological joint changes that accompany osteoarthritis. Application of this protocol can be valuable in monitoring disease progression and evaluating therapeutic interventions in osteoarthritis.

One of the most prevalent joint disorders in the United States, osteoarthritis (OA) is characterized by progressive degeneration of articular cartilage, ...

Advanced Cardiac Rhythm Management by Applying Optogenetic Multi-Site Photostimulation in Murine Hearts

Ventricular tachyarrhythmias are a major cause of mortality and morbidity worldwide. Electrical defibrillation using high-energy electric shocks is currently the only treatment for life-threatening ventricular fibrillation. However, defibrillation may have side-effects, including intolerable pain, tissue damage, and worsening of prognosis, indicating a significant medical need for the development of more gentle cardiac rhythm management strategies. Besides energy-reducing electrical approaches, cardiac optogenetics was introduced as a powerful tool to influence cardiac activity using light-sensitive membrane ion channels and light pulses. In the present study, a robust and valid method for successful photostimulation of Langendorff perfused intact murine hearts will be described based on multi-site pacing applying a 3 x 3 array of micro light-emitting diodes (micro-LED). Simultaneous optical mapping of epicardial membrane voltage waves allows the investigation of the effects of region-specific stimulation and evaluates the newly induced cardiac activity directly on-site. The obtained results show that the efficacy of defibrillation is strongly dependent on the parameters chosen for photostimulation during a cardiac arrhythmia. It will be demonstrated that the illuminated area of the heart plays a crucial role for termination success as well as how the targeted control of cardiac activity during illumination for modifying arrhythmia patterns can be achieved. In summary, this technique provides a possibility to optimize the on-site mechanism manipulation on the way to real-time feedback control of cardiac rhythm and, regarding the region specificity, new approaches in reducing the potential harm to the cardiac system compared to the usage of non-specific electrical shock applications.

This work reports a method for controlling the cardiac rhythm of intact murine hearts of transgenic channelrhodopsin-2 (ChR2) mice using local photostimulation with a micro-LED array and simultaneous optical mapping of epicardial membrane potential.

Ventricular tachyarrhythmias are a major cause of mortality and morbidity worldwide. Electrical defibrillation using high-energy electric shocks is ...

Sterile Pericarditis in Aachener Minipigs As a Model for Atrial Myopathy and Atrial Fibrillation

Atrial fibrillation (AF) is the most common arrhythmia caused by structural remodeling of the atria, also called atrial myopathy. Current therapies only target the electrical abnormalities and not the underlying atrial myopathy. For the development of novel therapies, a reproducible large animal model of atrial myopathy is necessary. This paper presents a model of sterile pericarditis-induced atrial myopathy in Aachener minipigs. Sterile pericarditis was induced by spraying sterile talcum and leaving a layer of sterile gauze over the atrial epicardial surface. This led to inflammation and fibrosis, two crucial components of the pathophysiology of atrial myopathy, making the atria susceptible to the induction of AF. Two pacemaker electrodes were positioned epicardially on each atrium and connected to two pacemakers from different manufacturers. This strategy allowed for repeated non-invasive atrial programmed stimulation to determine the inducibility of AF at specified time points after surgery. Different protocols to test AF inducibility were used. The advantages of this model are its clinical relevance, with AF inducibility and the rapid induction of inflammation and fibrosis-both present in atrial myopathy-and its reproducibility. The model will be useful in the development of novel therapies targeting atrial myopathy and AF.

We describe a sterile pericarditis model in minipigs to study atrial myopathy and atrial fibrillation (AF). We present surgical and anesthetic techniques, strategies for vascular access, and a protocol to study the inducibility of AF.

Atrial fibrillation (AF) is the most common arrhythmia caused by structural remodeling of the atria, also called atrial myopathy. Current therapies only ...

Endotracheal Intubation Using a Flexible Intubation Endoscope as a Standardized Model for Safe Airway Management in Swine

Endotracheal intubation is often a basic requirement for translational research in porcine models for various interventions that require a secured airway or high ventilation pressures. Endotracheal intubation is a challenging skill,&#160;requiring a minimum number of successful endotracheal intubations to achieve a high success rate under optimal conditions, which is often unachievable for non-anaesthesiology researchers. Due to the specific porcine airway anatomy, a difficult airway can usually be assumed. The impossibility of establishing a secure airway can result in injury, adverse events, or death of the laboratory animal. Using a prospective, randomized, controlled evaluation approach, it has been shown that fiberoptic-assisted endotracheal intubation takes longer but has a higher first-pass success rate than conventional intubation without causing clinically relevant drops in oxygen saturation. This model presents a standardized regimen for endoscopically guided endotracheal intubation, providing a secured airway, especially for researchers who are inexperienced in the technique of endotracheal intubation via direct laryngoscopy. This procedure is expected to minimize animal suffering and unnecessary animal losses.

The use of pigs in research has increased in recent years. Nevertheless, pigs are characterized by difficult airway anatomy. By demonstrating how to perform endoscopically guided endotracheal intubation, the present protocol aims to further increase laboratory animals' safety to avoid animal suffering and unnecessary death.

Endotracheal intubation is often a basic requirement for translational research in porcine models for various interventions that require a secured airway ...

A Model of Long-Term Ventricular Fibrillation in Isolated Rat Hearts

Ventricular fibrillation (VF) is a fatal arrhythmia with a high incidence in cardiac patients, but VF arrest under perfusion is a neglected method of intraoperative arrest in the field of cardiac surgery. With recent advances in cardiac surgery, the demand for prolonged VF studies under perfusion has increased. However, the field lacks simple, reliable, and reproducible animal models of chronic ventricular fibrillation. This protocol induces long-term VF through alternating current (AC) electrical stimulation of the epicardium. Different conditions were used to induce VF, including continuous stimulation with a low or high voltage to induce long-term VF and stimulation for 5 min with a low or high voltage to induce spontaneous long-term VF. The success rates of the different conditions, as well as the rates of myocardial injury and recovery of cardiac function, were compared. The results showed that continuous low-voltage stimulation induced long-term VF and that 5 min of low-voltage stimulation induced spontaneous long-term VF with mild myocardial injury and a high rate of recovery of cardiac function. However, the low-voltage, continuously stimulated long-term VF model had a higher success rate. High-voltage stimulation provided a higher rate of VF induction but showed a low defibrillation success rate, poor recovery of cardiac function, and severe myocardial injury. On the basis of these results, continuous low-voltage epicardial AC stimulation is recommended for its high success rate, stability, reliability, reproducibility, low impact on cardiac function, and mild myocardial injury.

This protocol presents a model of long-term ventricular fibrillation in rat hearts induced by continuous stimulation with low-voltage alternating current. This model has a high success rate, is stable, reliable, and reproducible, has a low impact on cardiac function, and causes only mild myocardial injury.

Ventricular fibrillation (VF) is a fatal arrhythmia with a high incidence in cardiac patients, but VF arrest under perfusion is a neglected method of ...