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

doi: 10.3791/60707 Published: January 30, 2020


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 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.


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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.

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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 ± 2 kg and 12-16 weeks in age were included in the protocol.

1. Anesthesia, intubation, and mechanical ventilation13,14

  1. 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.
  2. 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.
  3. Transport the sedated animals to the laboratory. Transport time should not exceed effective sedation time (here, 30-60 min).
  4. Monitor the peripheral oxygen saturation (SpO2) with a sensor clipped to the tail or ear.
  5. 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.
  6. Administer analgesia via intravenous injection of fentanyl (4 µg/kg). Induce anesthesia with intravenous injection of propofol (3 mg/kg)
  7. 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)
  8. 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 = <20 cmH20, PEEP (positive end-expiratory pressure) = 5 cmH20.
  9. 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).
  10. 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.
    1. Ensure that one person fixes the tongue outside with a piece of tissue and opens the snout with the other hand.
      1. 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.
  11. 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.
  12. 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).
  13. 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 <6 kPa, usually 20-30/min). Make sure that tube position is correct by regular and periodic exhalation of carbon dioxide via capnography.
  14. 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.
  15. Place gastric tube into the stomach to avoid reflux and vomiting with two people.
    1. Fix the tongue outside with a piece of tissue and open the snout with the other hand.
      1. 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'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.

2. Instrumentation

  1. Use bandages to pull back the hindlegs to smooth the folds in the femoral area for vessel catherization.
  2. 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).
  3. Disinfect the inguinal area (see step 1.6). Repeat this process 2x.
  4. Fill all catheters with saline solution. Apply ultrasound gel on the ultrasound probe. Cover the inguinal area with a sterile fenestrated drape.
  5. 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°.
  6. 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's technique is associated with significantly less blood loss and tissue trauma than other methods of vascular access.
  7. Confirm the desired needle position by observing bright red pulsating blood. Disconnect the syringe and quickly insert the guidewire into the right femoral artery.
  8. 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.
  9. Insert the guidewire for the central venous catheter into the right femoral vein after disconnecting the syringe. Retract the Seldinger needle.
  10. 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.
  11. Use Seldinger's technique to position the central venous line into the right femoral vein. Aspirate all ports and flush them with saline solution.
  12. Perform the same procedure on the left inguinal side to insert the other introducer sheaths in Seldinger's technique into the left femoral artery (8 Fr) and femoral vein (8 Fr).
  13. 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.
  14. 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.
  15. 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.
  16. Initiate ventricular fibrillation (see section 4).

3. Pulse contour cardiac output

  1. Insert transpulmonal thermodilution catheter into the right arterial introducer sheath.
    NOTE: In clinical medicine, thermodilution catheters are directly placed by Seldinger'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.
  2. 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.
  3. Turn on the cardiac monitor system. Confirm that a new patient is being measured. Enter the size and weight.
  4. Switch the category to adults. Enter the protocol name and ID. Click on Exit.
  5. 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.
  6. Enter the central venous pressure.
  7. Open the three-way stopcock to the atmosphere.
  8. Click on Zero for system calibration. Click on Exit.
  9. Calibrate the continuous cardiac output measurement.
    1. Click on TD (thermodilution). Prepare a physiological saline solution with a temperature of 4 °C in a 10 mL syringe. Click on Start.
    2. 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.
    3. 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.

4. Ventricular fibrillation and mechanical resuscitation

  1. 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.
  2. Connect the electrodes to a defibrillator and establish an ECG.
  3. Immobilize the pig inside the vacuum mattress. Deflate the mattress to prevent unwanted movement during CPR. Control fixation of the limbs.
  4. Place chest compression device (here, LUCAS-2) around the chest and under the vacuum mattress according to the manufacturer's recommendations. Adjust the pressure pad to the lower third of the sternum in median position.
  5. Turn on chest compression device ("power" 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.
  6. Insert a fibrillation/pacing catheter into the left femoral vein through the i.v. sheath.
  7. 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).
  8. 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).
  9. 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.
  10. 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.
  11. 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.
  12. 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.
  13. Analyze ECG patterns. If ventricular fibrillation persists, prepare defibrillation.
    1. Enter Manual mode in the defibrillator menu. Adjust the energy to 200 J biphasic.
    2. 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.
  14. 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.
  15. 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.

5. End of experiment and euthanasia (in the case of ROSC)

  1. Inject 0.5 mg of fentanyl into the central venous line. Wait 5 min. Inject 200 mg of propofol into the central venous line.
  2. Euthanize the animal with a 40 mmol potassium chloride injection.
  3. Perform organ removal/fixation or analyses as needed.

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Representative Results

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Cardiac arrest was induced in seven pigs. Return of spontaneous circulation following CPR was achieved in four Pigs (57%) with a mean of 3 ± 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 performed and all values including temperature should be normalized.

If placed in the right position, the pacing catheter should start to influence heart rhythm. This can result in extrasystoles, tachycardia and all forms of ventricular and supraventricular arrhythmias. Cardiac arrest can be assumed if 1) the ECG reading shows ventricular fibrillation and 2) no cardiac output or pressure variations are measured by the arterial line (Figure 1). If this state persists with the generator turned off, fibrillation is likely not to spontaneously subside anymore17.

Once chest compressions are started, sufficient cardiac output generation is indicated by a mean arterial pressure of 30-50 mmHg. (Figure 1) If adhering to resuscitation guidelines, the administration of adrenaline (1 mg) should result in a substantial rise in blood pressure within 1 min.

ROSC is confirmed by a dramatic increase in expiratory carbon dioxide measurements (usually increasing from 10-20 mmHg during arrest to 45 mmHg and above), organized heart rhythm in the ECG, and respective cardiac output as shown by arterial measurement. Hypercapnia and a decreased Horovitz index (PaO2/FiO2) are commonly observed after ROSC. Reestablishment of controlled mechanical ventilation leads to recompensation and stable respiratory conditions (Figure 2). A ROSC rate of 50%-70% can be expected depending on the time between cardiac arrest and the start of chest compressions.

Figure 1
Figure 1: Typical hemodynamic values. (A) Heart rate monitoring during trial (depicted as mean values with standard deviation [SD] error bars). Heart rate drops to zero at cardiac arrest (CA) and is standardized during CPR according to the specifications of the chest compression device (here, 100 bpm). Tachycardia is regularly seen after achieving ROSC, initially as a result of adrenaline administration and metabolic acidosis compensation. Values usually normalize over a period of 1-2 h. (B) Mean intra-arterial blood pressure values. At cardiac arrest (CA), pressure does not drop below 10-20 mmHg but loses all signs of effective output. During CPR, especially before vasopressor effects are registered, adequate chest compressions are indicated by pressure values between 30-50 mmHg. Post-ROSC, norepinephrine might be necessary to cover low blood pressure intervals during metabolic recompensation. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Oxygenation and decarboxylation parameters during and after resuscitation. (A) Arterial partial pressure values of carbon dioxide (PaCO2) during and after CPR (depicted as mean values with standard deviation error bars). Under guideline-based ventilation, no significant differences should be detected. An increase in CO2 levels directly after ROSC is to be expected but should normalize within 1 h. (B) Typical values of Horovitz index (arterial partial pressure of oxygen [PaO2]/inspiratory oxygen fraction [FiO2]; depicted as mean values with SD error bars). During CPR, oxygenation is often highly impaired but usually fully recovers post-ROSC during the first 2 h. Please click here to view a larger version of this figure.

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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 approach to prevent vascular damages during instrumentation. However, only professionally trained users should work with this technique to yield its advantages20. For this experimental model, it must be stressed that handling electrical frequency generators as well as defibrillators should only be handled by specifically trained personnel or under their direct supervision. Failure to provide adequate expertise while conducting such trials may result in serious injury and can be life-threatening.

Correct positioning of the pacing catheter and initiation of ventricular fibrillation may prove difficult and can require reinsertion of the catheter or frequency variation. When repositioning or removing the catheter, the balloon should be deflated first to prevent internal injuries as well as damage to the catheter itself. If frequency variations are used, the catheter should be placed near the myocardium in order to detect ECG changes, then frequency should slowly be changed according to the manufacturer's instructions. Importantly, the chest compression device has to be positioned correctly and the pig has to be properly immobilized (as shown in the video). Repositioning during CPR can be necessary but often leads to insufficient resuscitation. Even though thoracic anatomy and bone structure differs compared to humans, our studies showed sufficient perfusion generation and ROSC rates with a compression device placed on the lower third of the sternum in median position.

Porcine models have been successfully used in critical care studies for decades17,21,22,23. Similar anatomic and physiologic properties comparable to humans allow for reasonably accurate deductions regarding patient reactions to certain stimuli or clinical situations. The presented resuscitation model has been used and modified in various trials18,24,25,26. It provides an experimental setting that enables the evaluation of guideline effectiveness, since (in contrast to resuscitation models in rodents) equal chest compression intervals, blood pressure thresholds, blood gas values, and defibrillation energies can be used for human comparisons as recommended by ILCOR and ERC, respectively. This facilitates internationally comparable and comprehensible study designs, thus generating a higher quality of evidence overall. The model additionally allows for adequate assessment of drug effects not only qualitatively, but also in a dose-dependent fashion.

Assuming guideline-based resuscitation with intervals of 2 min between defibrillations, pigs usually achieve ROSC within the first four shocks or within 8-10 min27. A ROSC rate of 50%-70% can be expected depending on the time between cardiac arrest and the start of chest compressions. If acceptable ROSC rates or adequate blood pressure values cannot be achieved, it is possible to add vasopressine (0.5 IU/kgBW) to the therapy regimen during CPR. During and directly after CPR, pulmonary gas exchange is heavily impaired. This is largely dependent on the ventilation mode used during chest compressions and can have long-term effects on end organ damage and inflammation18,25,28. Additionally, metabolic acidosis and stunned myocardium can lead to persistent hypotension, especially in the first 1 h following ROSC. This can be treated by fluid administration (20-30 mL/kgBW) and continuous norepinephrine infusion. Excessive acidosis can also be treated with 8.4% sodium bicarbonate solution with a maximum of 4 mL/kgBW.

This experimental protocol provides a standardized setting for resuscitation research in which the aspects of hemodynamic effects of specific drug treatments, influence of ventilation modes on ROSC rates, end-organ damage, and post-resuscitation reactions can be analyzed and evaluated under various circumstances. This will help further scientific insight into the pathophysiologic mechanisms underlying ventricular fibrillation and may lead to more effective treatment options.

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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.


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


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



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Standardized Model of Ventricular Fibrillation and Advanced Cardiac Life Support in Swine
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Ruemmler, R., Ziebart, A., Garcia-Bardon, A., Kamuf, J., Hartmann, E. K. Standardized Model of Ventricular Fibrillation and Advanced Cardiac Life Support in Swine. J. Vis. Exp. (155), e60707, doi:10.3791/60707 (2020).More

Ruemmler, R., Ziebart, A., Garcia-Bardon, A., Kamuf, J., Hartmann, E. K. Standardized Model of Ventricular Fibrillation and Advanced Cardiac Life Support in Swine. J. Vis. Exp. (155), e60707, doi:10.3791/60707 (2020).

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