Name: standardized model of ventricular fibrillation and advanced cardiac life support in swine
Uploaded: 2020-01-30T15:30:00.000+00:00
Duration: 5 min 36 s
Description: Watch this Scientific Journal Video about Standardized Model of Ventricular Fibrillation and Advanced Cardiac Life Support in Swine at JoVE.com

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><tbody><tr><td>1 M- Kaliumchlorid-L&amp;ouml;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&amp;ouml;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&amp;uuml;hrungsstab, Durchmesser 4.3</td><td>R&amp;uuml;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&amp;uuml;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>&amp;Ouml;ls&amp;auml;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&amp;uuml;schelit Super Safety Clear &gt;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&amp;uuml;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&amp;ouml;&amp;szlig;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 ...

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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

Myocardial Infarction and Functional Outcome Assessment in Pigs

Introduction of newly discovered cardiovascular therapeutics into first-in-man trials depends on a strictly regulated ethical and legal roadmap. One important prerequisite is a good understanding of all safety and efficacy aspects obtained in a large animal model that validly reflect the human scenario of myocardial infarction (MI). Pigs are widely used in this regard since their cardiac size, hemodynamics, and coronary anatomy are close to that of humans. Here, we present an effective protocol for using the porcine MI model using a closed-chest coronary balloon occlusion of the left anterior descending artery (LAD), followed by reperfusion. This approach is based on 90 min of myocardial ischemia, inducing large left ventricle infarction of the anterior, septal and inferoseptal walls. Furthermore, we present protocols for various measures of outcome that provide a wide range of information on the heart, such as cardiac systolic and diastolic function, hemodynamics, coronary flow velocity, microvascular resistance, and infarct size. This protocol can be easily tailored to meet study specific requirements for the validation of novel cardioregenerative biologics at different stages (i.e. directly after the acute ischemic insult, in the subacute setting or even in the chronic MI once scar formation has been completed). This model therefore provides a useful translational tool to study MI, subsequent adverse remodeling, and the potential of novel cardioregenerative agents.

This protocol describes the porcine myocardial infarction (MI) model using a 90 min closed-chest coronary balloon occlusion of the left anterior descending artery (LAD), followed by reperfusion. Furthermore, the protocol for several outcome parameters, such as cardiac function, hemodynamics, microvascular resistance, and infarct size, are also presented.

Introduction of newly discovered cardiovascular therapeutics into first-in-man trials depends on a strictly regulated ethical and legal roadmap. One ...

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 ...

Improvement of a Closed Chest Porcine Myocardial Infarction Model by Standardization of Tissue and Blood Sampling Procedures

Myocardial ischemia reperfusion (I/R) injury contributes to almost half of the necrotic area after myocardial infarction. To date there is no approved drug to prevent or reduce myocardial I/R injury. The study and understanding of the pathophysiological mechanisms of myocardial I/R injury is essential to develop successful treatments. Large animal experiments are an important step in translational methods. The porcine model of acute myocardial infarction has been established and described by ourselves and others. We aimed to further improve the value of the model by focusing in detail on the sampling techniques for use in future experiments. Furthermore, we emphasize small but important steps that can affect the quality of the final results. To mimic the clinical situation of myocardial I/R injury, a percutaneous coronary intervention (PCI) catheter was inserted into the left anterior descending coronary artery (LAD) of an anesthetized pig. &#176;&#176;&#176;This model mimics acute myocardial infarction and PCI treatment in humans with the possibility of accurately determining the area at risk as well as the necrotic- and viable ischemic tissue. Here the model was used to investigate the effect of a bicyclic peptide inhibitor of FXIIa. The model can also be modified to allow longer reperfusion times to study later effects of myocardial infarction.

Here we demonstrate a protocol to standardize sampling procedures of an established porcine model of acute myocardial infarction in order to increase its translational value in the understanding of the pathophysiology of myocardial ischemia/reperfusion injury and to test novel drug candidates.

Myocardial ischemia reperfusion (I/R) injury contributes to almost half of the necrotic area after myocardial infarction. To date there is no approved ...

Tachycardia-Induced Cardiomyopathy As a Chronic Heart Failure Model in Swine

A stable and reliable model of chronic heart failure is required for many experiments to understand hemodynamics or to test effects of new treatment methods. Here, we present such a model by tachycardia-induced cardiomyopathy, which can be produced by rapid cardiac pacing in swine.
A single pacing lead is introduced transvenously into fully anaesthetized healthy swine, to the apex of the right ventricle, and fixated. Its other end is then tunneled dorsally to the paravertebral region. There, it is connected to an in-house modified heart pacemaker unit that is then implanted in a subcutaneous pocket. 
After 4 - 8 weeks of rapid ventricular pacing at rates of 200 - 240 beats/min, physical examination revealed signs of severe heart failure - tachypnea, spontaneous sinus tachycardia, and fatigue. Echocardiography and X-ray showed dilation of all heart chambers, effusions, and severe systolic dysfunction. These findings correspond well to decompensated dilated cardiomyopathy and are also preserved after the cessation of pacing.
This model of tachycardia-induced cardiomyopathy can be used for studying the pathophysiology of progressive chronic heart failure, especially hemodynamic changes caused by new treatment modalities like mechanical circulatory supports. This methodology is easy to perform and the results are robust and reproducible.

Here, we present a protocol to produce tachycardia-induced cardiomyopathy in swine. This model represents a potent way to study the hemodynamics of progressive chronic heart failure and the effects of applied treatment.

A stable and reliable model of chronic heart failure is required for many experiments to understand hemodynamics or to test effects of new treatment ...

Complete and Partial Aortic Occlusion for the Treatment of Hemorrhagic Shock in Swine

Hemorrhage remains the leading cause of preventable deaths in trauma. Endovascular management of non-compressible torso hemorrhage has been at the forefront of trauma care in recent years. Since complete aortic occlusion presents serious concerns, the concept of partial aortic occlusion has gained a growing attention. Here, we present a large animal model of hemorrhagic shock to investigate the effects of a novel partial aortic balloon occlusion catheter and compare it with a catheter that works on the principles of complete aortic occlusion. Swine are anesthetized and instrumented in order to conduct controlled fixed-volume hemorrhage, and hemodynamic and physiological parameters are monitored. Following hemorrhage, aortic balloon occlusion catheters are inserted and inflated in the supraceliac aorta for 60 min, during which the animals receive whole-blood resuscitation as 20% of the total blood volume (TBV). Following balloon deflation, the animals are monitored in a critical care setting for 4 h, during which they receive fluid resuscitation and vasopressors as needed. The partial aortic balloon occlusion demonstrated improved distal mean arterial pressures (MAPs) during the balloon inflation, decreased markers of ischemia, and decreased fluid resuscitation and vasopressor use. As swine physiology and homeostatic responses following hemorrhage have been well-documented and are like those in humans, a swine hemorrhagic shock model can be used to test various treatment strategies. In addition to treating hemorrhage, aortic balloon occlusion catheters have become popular for their role in cardiac arrest, cardiac and vascular surgery, and other high-risk elective surgical procedures.

Here, we present a protocol demonstrating a hemorrhagic shock model in swine that uses aortic occlusion as a bridge to definitive care in trauma. This model has application in testing a wide range of surgical and pharmacological therapeutic strategies.

Hemorrhage remains the leading cause of preventable deaths in trauma. Endovascular management of non-compressible torso hemorrhage has been at the ...

Measuring Cardiac Output in a Swine Model

Swine are frequently used in medical research given their similar cardiac physiology to that of humans. Measuring cardiac parameters such as stroke volume and cardiac output are essential in this type of research. Contrast ventriculography, thermodilution, and pressure-volume loop (PV-loop) catheters can be used to accurately obtain cardiac performance data depending on which resources and expertise are available. For this study,five Yorkshire swine were anesthetized and intubated. Central venous and arterial access was obtained to place the necessary measurement instruments.A temperature probe was placed in the aortic root. A cold saline bolus was delivered to the right atrium and temperature deflection curve was recorded. Integration of the area under the curve allowed for the calculation of the current cardiac output.A pigtail catheter was percutaneously placed in the left ventricle and 30 mL of iodinated contrast was power injected over 2 seconds. Digital subtraction angiography images were uploaded to volumetric analysis software to calculate the stroke volume and cardiac output. A pressure volume-loop catheter was placed into the left ventricle (LV) and provided continuous pressure and volume data of the LV, which allowed the calculation of both stroke volume and cardiac output.All three methods demonstrated good correlation with each other. The PV-loop catheter and thermodilution exhibited the best correlation with a 3% error and a Pearson coefficient of 0.99, with 95% CI=0.97 to 1.1, (p=0.002). The PV-loop catheter against ventriculography also showed good correlation with a 6% error and a Pearson coefficient of 0.95, 95% CI=0.96 to 1.1 (p=0.01). Finally, thermodilution against ventriculography had a 2% error with r=0.95, 95% CI=0.93 to 1.11, (p=0.01). In conclusion, we state that the PV-loop catheter, contrast ventriculography, and thermodilution each offer certain advantages depending on the researcher's requirements. Each method is reliable and accurate for measuring various cardiac parameters in swine such as the stroke volume and cardiac output.

Thermodilution, pressure-volume loop catheters, and contrast ventriculography are reliable and accurate methods for determining cardiac physiology such as stroke volume and cardiac output in a laboratory setting in swine.

Swine are frequently used in medical research given their similar cardiac physiology to that of humans. Measuring cardiac parameters such as stroke volume ...

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 ...

Myocardial Infarction by Percutaneous Embolization Coil Deployment in a Swine Model

Myocardial infarction (MI) is the leading cause of mortality worldwide. Despite the use of evidence-based treatments, including coronary revascularization and cardiovascular drugs, a significant proportion of patients develop pathological left-ventricular remodeling and progressive heart failure following MI. Therefore, new therapeutic options, such as cellular and gene therapies, among others, have been developed to repair and regenerate injured myocardium. In this context, animal models of MI are crucial in exploring the safety and efficacy of these experimental therapies before clinical translation. Large animal models such as swine are preferred over smaller ones due to the high similarity of swine and human hearts in terms of coronary artery anatomy, cardiac kinetics, and the post-MI healing process. Here, we aimed to describe an MI model in pig by permanent coil deployment. Briefly, it comprises a percutaneous selective coronary artery cannulation through retrograde femoral access. Following coronary angiography, the coil is deployed at the target branch under fluoroscopic guidance. Finally, complete occlusion is confirmed by repeated coronary angiography. This approach is feasible, highly reproducible, and emulates the pathogenesis of human non-revascularized MI, avoiding the traditional open-chest surgery and the subsequent postoperative inflammation. Depending on the time of follow-up, the technique is suitable for acute, sub-acute, or chronic MI models.

Myocardial infarction (MI) animal models that emulate the natural process of the disease in humans are crucial to understanding pathophysiological mechanisms and testing the safety and efficacy of new emergent therapies. Here, we describe an MI swine model created by deploying a percutaneous embolization coil.

Myocardial infarction (MI) is the leading cause of mortality worldwide. Despite the use of evidence-based treatments, including coronary revascularization ...

Swine Models of Aneurysmal Diseases for Training and Research

Large animal models, specifically swine, are widely used to research cardiovascular diseases and therapies, as well as for training purposes. This paper describes two different aneurysmal swine models that may help researchers to study new therapies for aneurysmal diseases. These aneurysmal models are created by surgically adding a pouch of tissue to carotid arteries in swine. When the model is used for research, the pouch must be autologous; for training purposes, a synthetic pouch suffices.
First, the right external jugular vein (EJV) and right common carotid artery (CCA) must be surgically exposed. The EJV is ligated and a vein pouch fashioned from a short segment. This pouch is then sutured to an elliptical arteriotomy performed in the CCA. Animals must be kept heparinized during model creation, and local vasodilators may be used to decrease vasospasms. Once the suture is completed, correct blood flow should be inspected, checking for bleeding from the suture line and vessel patency. Finally, the surgical incision is closed by layers and an angiography performed to image the aneurysmal model.
A simplification of this aneurysmal carotid model that decreases invasiveness and surgical time is the use of a synthetic, rather than venous, pouch. For this purpose, a pouch is tailored in advance with a segment of a polytetrafluoroethylene (PTFE) prosthesis, one end of which is sutured close using polypropylene vascular suture and sterilized prior to surgery. This "sac" is then attached to an arteriotomy performed in the CCA as described. 
Although these models do not reproduce many of the physiopathological events related to aneurysm formation, they are hemodynamically similar to the situation found in the clinical setting. Therefore, they can be used for research or training purposes, allowing physicians to learn and practice different endovascular techniques in animal models that are close to the human system.

Description of two different aneurysmal swine models for neuroradiology training courses and research studies. This study provides evidence of the feasibility of these aneurysm porcine model creations and the reproducible methods that are close to the clinical setting.

Large animal models, specifically swine, are widely used to research cardiovascular diseases and therapies, as well as for training purposes. This paper ...

Delivery of Cardioactive Therapeutics in a Porcine Myocardial Infarction Model

Myocardial infarction is one of the leading causes of death and disability worldwide, and there is an urgent need for novel cardioprotective or regenerative strategies. An essential component of drug development is determining how a novel therapeutic is to be administered. Physiologically relevant large animal models are of critical importance in assessing the feasibility and efficacy of various therapeutic delivery strategies. Due to their similarities to humans in cardiovascular physiology, coronary vascular anatomy, and heart weight to body weight ratio, swine is one of the preferred species in the preclinical evaluation of new therapies for myocardial infarction. The present protocol describes three methods of administering cardioactive therapeutic agents in a porcine model. After percutaneously induced myocardial infarction, female landrace swine received treatment with novel agents through either: (1) thoracotomy and transepicardial injection, (2) catheter-based transendocardial injection, or (3) intravenous infusion via jugular vein osmotic minipump. The procedures employed for each technique are reproducible, resulting in reliable cardioactive drug delivery. These models can be easily adapted to suit individual study designs, and each of these delivery techniques can be used to investigate a variety of possible interventions. Therefore, these methods are a useful tool for translational scientists pursuing novel biological approaches in cardiac repair following myocardial infarction.

The present protocol describes three methods of administering cardioactive therapeutic agents in a porcine model. Female landrace swine received treatment through either: (1) thoracotomy and transepicardial injection, (2) catheter-based transendocardial injection, or (3) intravenous infusion via jugular vein osmotic minipump.

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

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