All authors disclose no financial or any other conflicts of interests.
GRANTS:
This study was supported by a grant from the Deutsche Forschungsgemeinschaft to P. Pickerodt and W. Boemke (Pi795/2-2).

The experiments were conducted at the Department of Experimental Medicine, Charit&#233; - Universit&#228;tsmedizin, Berlin, Germany (certi&#64257;ed according to the EN DIN ISO 9001:2000), and were approved by the federal authorities for animal research in Berlin, Germany prior to the experiments. The principles of laboratory animal care, which were used in all experiments, were in accordance with the guidelines of the European and German Society of Laboratory Animal Sciences.
1. Animal Welfare and Laboratory Animals
<ol>
	<li>All experiments were conducted in fully anesthetized male pigs (German Landrace &#215; Large White) of 3-4 months of age, weighing 30-60 kg.</li>
</ol>
2. Anesthesia, Intubation, and Mechanical Ventilation
<ol>
	<li>Withhold food for 12 hr prior to anesthesia to avoid a full stomach of the pig, but allow free access to water to minimize stress.</li>
	<li>For premedication, inject a combination of azaperone (3 mg/kg), atropine (0.03 mg/kg), ketamine (25 mg/kg), and xylazine (3.5 mg/kg) into the neck muscles of the pig while the animal is still kept in its box to minimize stress.
	<ol>
		<li>Place the animal onto a stretcher and cover the eyes with a cloth for transportation once an adequate level of anesthesia is achieved.</li>
		<li>Transport the pig to the surgical theater and ensure sufficient spontaneous breathing at all times by keeping the snout unobstructed.</li>
		<li>Place the pig in the prone position and preoxygenate with a mask that fits the animal&#39;s snout using a high flow of oxygen (e.g. 10 L/min).</li>
	</ol>
	</li>
	<li>Start monitoring the peripheral oxygen saturation (SpO2) by clipping the respective sensor of the monitor onto one of the ears. Gain venous access with a clinically used peripheral vein catheter (usually 18 or 20 G) placed into one of the ear veins after a wipe down procedure with alcohol swaps.
	<ol>
		<li>Start an infusion with a balanced crystalloid solution with a 500 ml bolus followed by continuous infusion of about 4 ml/kg/hr (as deemed clinically necessary depending on the experiment) and ensure correct placement of the catheter for the subsequent infusion of anesthetics. 
		NOTE: The infusion of larger volumes of normal saline instead of a balanced crystalloid solution can result in hyperchloremic acidosis, whereas infusion of a solution containing lactate can result in increased serum lactate concentrations, and thus interfere with the interpretation of the blood gas analysis or the results of subsequent experiments.</li>
	</ol>
	</li>
	<li>After sufficient preoxygenation (preoxygenation during the complete time for peripheral vein access, measured SpO2 of 95-100%) inject propofol (about 5-10 mg/kg body weight - the exact dose depends on the effect of the premedication and differs from animal to animal) using the peripheral vein catheter.</li>
	<li>Intubate the pig in the prone position using an endotracheal tube for clinical application (7.5-8.5 ID) and a laryngoscope designed for large animals (straight blade of about 25 cm length).
	<ol>
		<li>Strap two bandages through the snout of the animal (first investigator). Pull one bandage upwards to move the head into the correct position and straighten the oropharyngeal structures, pull the other bandage down to open the snout. Pull the tongue to one side (second investigator).</li>
		<li>Press the tongue down with the blade of the laryngoscope and advance the blade towards the epiglottis. Note, in this position the epiglottis is often wedged behind the soft palate of the pigs.</li>
		<li>Mobilize the epiglottis with the tube, press it down with the blade of the laryngoscope and visualize the vocal cords for intubation.</li>
		<li>Advance the tube through the vocal cords while turning the tube upwards and block the cuff as described in detail by Theisen et al. 5. 
		NOTE: Intubation may also be possible in the supine position, depending on the training of the investigator as well as the standard procedures of a certain institution. A large tube diameter supports the surfactant washout due to faster inflow and outflow of the lavage fluid.</li>
		<li>Check the correct placement of the tube using capnography and auscultation. For this, ensure the capnogram is &#39;normally&#39; shaped and auscultate both lungs for equal breath sounds as done in the clinic. 
		NOTE: Mechanically ventilate the pig with chest compressions in case of failed or delayed intubation. This requires manual compression of the rib cage from both sides while supplying oxygen with a high flow via a tight fitting face mask.</li>
	</ol>
	</li>
	<li>Start mechanical ventilation, setting the fraction of inspired oxygen (FIO2) to 1, respirator frequency to 15-20/min, tidal volume of 8-9 ml/kg body weight, inspiration to expiration ratio (I:E) of 1:1.5, and apply a positive end expiratory pressure (PEEP) of 5 cm H2O. Adjust the settings to target an end expiratory partial pressure of carbon dioxide (PetCO2) of 35-40 mmHg and a SpO2 above 95%.
	<ol>
		<li>Maintain anesthesia with continuous i.v. infusion of thiopentone (20 mg/kg/h) and fentanyl (7 &#181;g/kg/h). 
		NOTE: The necessary dosage may vary from animal to animal. Do not leave the animal unattended. Ensure a sufficient anesthesia at all times during the experiment for animal welfare and scientific reasons.</li>
		<li>Check for absence of corneal reflexes and monitor the animal closely for stress/pain reactions during instrumentation. Instrumentation should be possible without administering a muscle relaxant if anesthesia is sufficient. Administer pancuronium bromide (0.15 mg/kg BW i.v. bolus followed by a continuous infusion of 0.15 mg/kgBW/hr) if muscle relaxation is necessary for the experiment (e.g. lung compliance measurements).</li>
	</ol>
	</li>
</ol>
3. Instrumentation Techniques
<ol>
	<li>Place the animal into the supine position and retract the legs using bandages to stretch the skin above the planned incision sites. Sterilize the operating areas using a pre-operative skin disinfectant like an alcohol/1% iodine solution. 
	NOTE: We use a wipe down procedure to sterilize the operating area but do not use complete aseptical techniques, since this is a nonsurvival model. The level of surgical asepsis depends upon the investigation following the induction of lung injury.</li>
	<li>Make a 10 cm incision on the line connecting the mandible and the sternum (left or right side possible) cutting through the skin using a scalpel for placement of the central venous line and the introducer sheath of the pulmonary artery catheter. Reassess the depth of anesthesia and increase the dosage if necessary.</li>
	<li>Separate the subcutaneous tissue and the platysma using tissue forceps and surgical scissors. Once the brachiocephalic and the sternocephalic muscles are visible perform a blunt cut down procedure separating the fascia between the muscles until the external jugular vein is visible using instruments like forceps or the fingers.</li>
	<li>Cannulate the external jugular vein with the central venous catheter and the introducer sheath by means of a modified Seldinger technique. Flush all catheters with normal saline before introducing them.
	<ol>
		<li>For this, advance the respective needle from the introducer set into the vein until venous blood (dark, not pulsating) can be aspirated. Advance the guide wire through the cannula into the vein for about 15 cm. Remove the needle and advance the introducer sheath into the vein. Remove the guide wire. Repeat the same procedure for placement of a central venous catheter if necessary for the experiment.</li>
	</ol>
	</li>
	<li>Control the correct placement of the catheters by aspiration of venous blood. Close with standard sutures. 
	NOTE: Do not dilate the vein with a vasodilator as you would in case of a percutaneous approach, because this will tear the vein (Figure 1). An ultrasound-guided approach is also possible if the investigator is trained in ultrasound-guided cannulization techniques in pigs.</li>
	<li>Identify the fold between the gracilis and sartorius muscle of the hind leg (left or right is possible) for placement of the arterial catheter. This is the fold where the pulsation of the femoral artery can be palpated.</li>
	<li>Make a 5 cm incision along the fold cutting through the skin using a scalpel.</li>
	<li>Separate the subcutaneous tissue using tissue forceps and surgical scissors. Use a blunt cut down procedure separating the fascia between the muscles to the level of the femoral artery. Note, avoid cutting the saphenous vessels by performing the cut down procedure cranial of them.</li>
	<li>Cannulate the femoral artery by means of a modified Seldinger technique as described in 3.2. A ligature can be looped around the artery and closed in case of bleeding at the site of puncture. This step should be avoided, if possible, as it compromises blood flow to the hind leg. Close with standard sutures.</li>
	<li>Connect the arterial catheter and the central venous line to the transducer system and calibrate both against the atmosphere (zero) and either 200 mmHg (arterial line) or 50 mmHg (central venous line) to start monitoring.</li>
	<li>Place all pressure transducers at the height of the right atrium (in pigs in the supine position about half the height of the thorax).</li>
	<li>Perform a small (4-5 cm) incision cutting through the skin above the bladder using a scalpel for catherization of the urinary bladder. Again, separate the subcutaneous tissue using blunt instruments.</li>
	<li>Perform a purse-string suture (1-2 cm in diameter) in the wall of the bladder once it is visualized. 
	NOTE: The sutures should not penetrate through all layers of the bladder wall since this would result in the loss of urine through the punctures.</li>
	<li>Do a minimal incision in the middle of the suture, introduce the urinary catheter, block the balloon with 10 ml of aqua dest, pull the catheter back until a light resistance is felt, and close the purse-string suture around the catheter. Close the skin using standard sutures.</li>
</ol>
4. Introduction of the Pulmonary Artery Catheter
<ol>
	<li>Inject 0.5-1 ml of air into the balloon of the pulmonary artery catheter (depending on the size of the catheter) and check for possible damage of the balloon. Deflate the balloon again.</li>
	<li>Connect the pulmonary artery catheter to the pressure transducer system and calibrate the PAC against the atmosphere (zero) and 100 mmHg (Figure 2 and 3).
	<ol>
		<li>Introduce the pulmonary artery catheter through the introducer sheath (deflated balloon) for 10 to 15 cm, depending on the sheath length.</li>
		<li>Inflate the balloon (the balloon has to have left the sheath for this) and advance the pulmonary artery catheter further while monitoring the pressure and the typical wave forms on the hemodynamic monitor.</li>
		<li>Advance the PAC while the wave forms which are typical for the right atrium, right ventricle, and pulmonary artery appear and stop advancing when the pulmonary capillary wedge pressure (PCWP) curve appears (Figure 4). Deflate the balloon. 
		NOTE: Once the balloon is deflated the PCWP-waveform must disappear and the pulmonary arterial pressure waveform must be visible. Otherwise the catheter is most likely inserted too far into a pulmonary artery resulting in permanent occlusion of the artery (auto wedge position). In this case, pull the catheter back until the pulmonary arterial pressure waveform reappears to avoid serious complications (e.g. rupture of the blood vessel) 6.</li>
		<li>Make sure the balloon is deflated whenever the catheter is pulled back to avoid serious complications. 
		NOTE: Pulmonary artery catheters are often accidentally advanced into liver veins via the inferior caval vein in pigs. Thus, pull back the catheter and start all over again, if the right ventricle is not reached after about 30 cm.</li>
	</ol>
	</li>
</ol>
5. Pulmonary Artery Thermodilution Technique and Hemodynamic Measurements
<ol>
	<li>Copy all hemodynamic values like heart rate, systolic, diastolic, and mean arterial pressure (MAP), pulmonary arterial pressures, and central venous pressure (CVP) from the hemodynamic monitor.</li>
	<li>Measure the PCWP promptly. For this, inflate the balloon of the pulmonary artery catheter and ensure that a correct PCWP curve is displayed (Figure 4). Copy the pulmonary capillary wedge pressure (PCWP) at end-expiration from the monitor. Immediately deflate the balloon afterwards (see 4.2.4). Deflate the balloon, pull the catheter back and reposition it if you are unable to see a correct PCWP curve as described in 4.2.2.
	<ol>
		<li>Connect the thermistor and an appropriate flow through housing to the central venous lumen of the pulmonary artery catheter and to the monitor for measuring cardiac output (CO). Next, connect the distal temperature port of the catheter (red cap) with the monitor.</li>
		<li>Start the monitor and select &#39;bolus CO&#39; to monitor time-temperature curves and thus measure cardiac output (CO) with the pulmonary artery thermodilution technique 7.</li>
		<li>Press &#39;Inj Vol&#39; and select the volume of cooled saline (5 ml in the experiments presented here). Return to the previous screen. Press &#39;catheter&#39; and select the size of the pulmonary artery catheter that is used. Return to the previous screen.</li>
		<li>Select &#39;start bolus&#39; and inject 5 ml of normal saline of a temperature of 4 &#176;C as quickly as possible using the flow through housing. Wait until the measurement is completed and the respective time-temperature curve appears on the monitor. Copy the CO value from the monitor.</li>
		<li>Perform 5 measurements in quick succession in a randomized order over the respiratory cycle of the ventilator as described in 5.3.4. Ignore the highest and the lowest value and use the remaining three to calculate the mean value of the cardiac output. 
		NOTE: This monitoring setup is described for an Edwards Vigilance monitor, model VGS1. The setup may differ depending on the monitor. Nevertheless, it is essential to select the correct injection volume of saline as well as size of the catheter. Some monitors require the selection of a computation constant that codes the respective amount of saline and catheter size. The constants are usually found in a leaflet inside the packaging of the catheter. Keep the saline at the same temperature throughout the experiment (&#60;5 &#176;C) to assure correct measurements. Use 5% glucose solutions instead of saline for studies involving exact measurements of electrolyte intake and homeostasis.</li>
	</ol>
	</li>
	<li>Make sure that all parameters have been recorded and that arterial and mixed venous blood samples were taken to enable calculation of the intra-pulmonary right-to-left shunt.</li>
	<li>Record all needed respiratory data promptly like peak and plateau inspiratory pressure from the respirator, or perform additional measurements like measuring transpulmonary pressure to complete the data at any given time point of the experiment.</li>
</ol>
6. Lung Lavages to Induce Lung Injury
<ol>
	<li>Ensure that the animal is ventilated with a FIO2 of 1.0 and set the PEEP to 2-4 cm H2O for the lavage procedure. Disconnect the animal from the respirator.</li>
	<li>Fill the lungs with warmed normal sterile saline (37 &#176;C, 50 ml/kg body weight). For this, prefill a funnel and connect it to the endotracheal tube with a fitting elastic tube. Raise the funnel 1 m above the animal and pour the saline into the lungs as quickly as possible. The hydrostatic pressure will allocate the saline into all pulmonary sections. 
	NOTE: Sterile normal saline is used to avoid the pulmonary wash in of pathogens and possible, septic decompensation of the animal. The use of 0.9% saline is crucial, since hypotonic fluids will result in immediate pulmonary edema, electrolyte imbalance and death of the animal. Do not reuse the saline after a lavage to maximize surfactant wash out.</li>
	<li>Stop filling when MAP falls below 50 mmHg. 
	NOTE: Only hemodynamic parameters and SpO2 can be used to monitor the animal for decompensation, since the animal is disconnected from the ventilator during lung lavages.</li>
	<li>Lower the funnel manually to ground level, drain the lavage fluid passively and reconnect the animal to the ventilator for oxygenation.</li>
	<li>Wait until the animal compensates (increase in MAP and SpO2) and repeat the lavage as soon as possible. The time frame for re-lavage should not exceed 5 min. 
	NOTE: Stabilization of the animal between two lavages in the case of hemodynamic decompensation serves to avoid the use of vasopressors to treat systemic hypotension, and to further avoid recruitment maneuvers to encounter hypoxemia. The animals are ventilated with a FIO2 of 1.0 during the lavages and the following experiments to maintain oxygenation despite reduced PaO2/FIO2 ratios. Setting the PEEP at 2-4 cm H2O during the lavages will promote rapid formation of atelectasis. But, PEEP has to be set at or above 5 cm H2O after the induction of lung injury to fulfill the Berlin definition of ARDS. During the experiment no recruitment maneuver or change in PEEP is allowed, to prevent any investigator-induced bias with regards to the severity of lung injury.</li>
	<li>Take an arterial blood gas sample after the second or third lavage depending on the hemodynamic deterioration and compromise in SpO2.</li>
	<li>Repeat lavages until the PaO2/FIO2 ratio (Horowitz index) is persistently measured below 100 mmHg for at least 60 min at FIO2 1.0 and PEEP &#8805; 5 cmH2O.</li>
	<li>Adjust the ventilator rate during the period of lavages to keep the arterial pH above 7.25 in order to prevent hemodynamic decompensation.</li>
	<li>Start the experiment/treatment based on the surfactant washout model once the PaO2/FIO2 ratio (Horowitz index) is persistently measured below 100 mmHg for 60 min. 
	NOTE: After the induction of the lung injury as described, the changes in lung function will remain stable for hours, deteriorate, or even improve depending on the ventilator settings. 
	NOTE: This animal model is based on surfactant washout and consequent formation of atelectasis. Hence, any deviation from the specified ventilator settings, which may lead to recruitment of atelectatic lung regions (increase in PIP or PEEP), will partially reverse the deleterious effect of the lavages and hinder standardization of this model.</li>
</ol>
7. End of Experiment and Euthanasia
<ol>
	<li>Ensure that all measurements are performed and data is secured before the end of the experiment.</li>
	<li>Inject 0.5 mg of fentanyl additionally to the continuous anesthesia and wait at least 5 min. Inject an overdose of thiopental (at least 1,000 mg) quickly followed by at least 60 mmol of potassium using the central line.</li>
</ol>

<ol>
	<li>Rubenfeld, G. 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=Incidence+and+Outcomes+of+Acute+Lung+Injury.">Incidence and Outcomes of Acute Lung Injury.</a> N Engl J Med. 353 (16), 1685-1693 (2005).</li><li>Ballard-Croft, C., Wang, D., Sumpter, L. R., Zhou, X., Zwischenberger, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Large-animal+models+of+acute+respiratory+distress+syndrome.">Large-animal models of acute respiratory distress syndrome.</a> Ann Thorac Surg. 93 (4), 1331-1339 (2012).</li><li>Lachmann, B., Robertson, B., Vogel, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+lung+lavage+as+an+experimental+model+of+the+respiratory+distress+syndrome.">In vivo lung lavage as an experimental model of the respiratory distress syndrome.</a> Acta Anaesthesiol Scand. 24 (3), 231-236 (1980).</li><li>Donaubauer, B., 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-dose+inhalation+of+an+endothelin-A+receptor+antagonist+in+experimental+acute+lung+injury:+ET-1+plasma+concentration+and+pulmonary+inflammation.">Low-dose inhalation of an endothelin-A receptor antagonist in experimental acute lung injury: ET-1 plasma concentration and pulmonary inflammation.</a> Exp Biol Med (Maywood). 231 (6), 960-969 (2006).</li><li>Theisen, M. M., 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=Ventral+recumbency+is+crucial+for+fast+and+safe+orotracheal+intubation+in+laboratory+swine.">Ventral recumbency is crucial for fast and safe orotracheal intubation in laboratory swine.</a> Lab Anim. 43 (1), 96-101 (2009).</li><li>Kelly, C. R., Rabbani, L. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Videos+in+clinical+medicine.+Pulmonary-artery+catheterization.">Videos in clinical medicine. Pulmonary-artery catheterization.</a> N Engl J Med. 369 (25), 35 (2013).</li><li>Forrester, J. S., 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=Thermodilution+cardiac+output+determination+with+a+single+flow-directed+catheter.">Thermodilution cardiac output determination with a single flow-directed catheter.</a> Am Heart J. 83 (3), 306-311 (1972).</li><li>Deja, M., 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=The+inhaled+ET(A)+receptor+antagonist+LU-135252+acts+as+a+selective+pulmonary+vasodilator.">The inhaled ET(A) receptor antagonist LU-135252 acts as a selective pulmonary vasodilator.</a> Clin Sci (Lond). 103, 21-24 (2002).</li><li>Matute-Bello, G., Frevert, C. W., Martin, T. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+of+acute+lung+injury.">Animal models of acute lung injury.</a> Am J Physiol Lung Cell Mol Physiol. 295 (3), 379-399 (2008).</li><li>Kloot, T. E., 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=Recruitment+maneuvers+in+three+experimental+models+of+acute+lung+injury.+Effect+on+lung+volume+and+gas+exchange.">Recruitment maneuvers in three experimental models of acute lung injury. Effect on lung volume and gas exchange.</a> Am J Respir Crit Care Med. 161 (5), 1485-1494 (2000).</li></ol>

We gratefully acknowledge the help of Prof. Burkhard Lachmann when contemplating the intricacies of PEEP settings during surfactant washout and the excellent technical assistance of Birgit Brandt and Sabine Molling.

This article describes a step by step instruction to induce severe lung injury in pigs due to surfactant washout by repeated lung lavages. This specific method enables a reproducible and comparable deterioration in lung function and pulmonary vascular resistance. It is imperative to lavage the pigs until the PaO2/FIO2 ratio decreased below 100 mmHg and stays below 100 mmHg for one hr. Once, this is achieved the animals usually do not recover from the lung injury for at least 4 ...

The mortality of human acute respiratory distress syndrome (ARDS) remains high with values between 40 and 50% 1 despite more than 4 decades of intense research. Animal models of lung injury play a major role in investigating the complex pathomechanisms or novel therapeutical approaches to reduce mortality and limit long term disabilities.
Various models have been established to induce lung injury that simulates aspects of human ARDS in either large (e.g. pigs) or small animals (e.g. rodents). Methods differ strongly, including pulmonary arterial infusion of oleic acid, intravenous (i.v.) infusion of bacteria, and endotoxins or cecal ligation and puncture (CLP) models causing sepsis-induced ARDS. In addition, direct lung injuries due to large tidal volumes and high peak inspiratory pressures (ventilator-induced lung injury; VILI), smoke/burn injuries or lung ischemia/reperfusion (I/R) models are often used 2. One major disadvantage of CLP models, as well as models working with endotoxins, is the underlying inflammation which hinders the analysis of biotrauma caused by lung injury alone. Furthermore, it may take hours to days to result in lung injury, as is the case for VILI in large animals.
The induction of lung injury by surfactant washout with repeated lung lavages, as it was first described by Lachmann et al. in guinea pigs 3, is a time efficient method to induce lung injury with reproducible functional and mechanical compromises, as well as changes in pulmonary vascular resistance. The adaption of this model to mechanically ventilated pigs of about 30-60 kg body weight supports basic research with clinically used mechanical ventilators, catheters and monitors, while the compromises in gas exchange and hemodynamics are highly reproducible at the same time 4. In addition, induction of lung injury by lavages does not require specific equipment that is not commonly available in respiratory laboratories designed for experiments in large animals. The model presented in this article is suitable for research demanding equipment (e.g. ventilators) that is designed for use in humans, and furthermore ensures a high reproducibility in the occurring deteriorations in lung function. Standardization of this model helps to ensure comparability between experiments and reduce the number of animals needed. The potential recruitability of atelectatic lung regions with deliberate or unknown recruitment maneuvers is a severe limitation of this specific model. In the following article we give a detailed description of the lavage model for the induction of lung injury and provide representative data to characterize the stability of the compromises in lung function.

<table><tbody><tr><td>Evita Infinity V500</td><td>Dr&amp;auml;ger</td><td></td><td>intensive care ventilator</td></tr><tr><td>Vigilance I&amp;nbsp;</td><td>Edwards</td><td></td><td>monitor</td></tr><tr><td>Vasofix Braun&amp;uuml;le 20G</td><td>B Braun</td><td>4268113B</td><td>peripheral vein catheter</td></tr><tr><td>Mallinckrodt Tracheal Tube Cuffed</td><td>Covidien</td><td>107-80</td><td>&amp;nbsp;8.0 mm ID</td></tr><tr><td>MultiCath3</td><td>Vygon</td><td>157,300</td><td>3 lumen central venous catheter, 20 cm length</td></tr><tr><td>Leader Cath Set</td><td>Vygon</td><td>115,805</td><td>arterial catheter</td></tr><tr><td>Percutaneus Sheath Introducer Set</td><td>Arrow</td><td>SI-09600</td><td>introducer sheath for pulmonary artery catheter of 4-6 Fr., 10 cm length</td></tr><tr><td>Swan-Ganz True Size Thermodilution Catheter</td><td>Edwards</td><td>132F5</td><td>pulmonary artery catheter, 75 cm length</td></tr><tr><td>Flow through chamber thermistor</td><td>Baxter</td><td>93-505</td><td>for measuring cardiac output</td></tr><tr><td>urinary catheter</td><td></td><td></td><td>no specific model requiered</td></tr></tbody></table>

lavage-induced surfactant depletion in pigs as a model of the acute respiratory distress syndrome (ards)

Various animal models of lung injury exist to study the complex pathomechanisms of human acute respiratory distress syndrome (ARDS) and evaluate future therapies. Severe lung injury with a reproducible deterioration of pulmonary gas exchange and hemodynamics can be induced in anesthetized pigs using repeated lung lavages with warmed 0.9% saline (50 ml/kg body weight). Including standard respiratory and hemodynamic monitoring with clinically applied devices in this model allows the evaluation of novel therapeutic strategies (drugs, modern ventilators, extracorporeal membrane oxygenators, ECMO), and bridges the gap between bench and bedside. Furthermore, induction of lung injury with lung lavages does not require the injection of pathogens/endotoxins that impact on measurements of pro- and anti-inflammatory cytokines. A disadvantage of the model is the high recruitability of atelectatic lung tissue. Standardization of the model helps to avoid pitfalls, to ensure comparability between experiments, and to reduce the number of animals needed.

PaO2/FIO2-ratio decreases during lung lavages, but the exact impact of a single lavage is difficult to predict. We start to take arterial blood gas samples from the third lavage onwards to detect a decrease in PaO2/FIO2-ratio below 100 mmHg. Once a decrease in PaO2/FIO2-ratio below 100 mmHg is achieved, we require this ratio to remain below 100 mmHg for one h...

Watch this Scientific Journal Video about Lavage-induced Surfactant Depletion in Pigs As a Model of the Acute Respiratory Distress Syndrome ARDS at JoVE.com

Lavage-induced Surfactant Depletion in Pigs As a Model of the Acute Respiratory Distress Syndrome ARDS

Repeated pulmonary lavages in anesthetized pigs induce lung injury resembling major aspects of human acute respiratory distress syndrome (ARDS). For this purpose the lungs are repeatedly lavaged with 0.9% saline at 37 &#176;C. The goal of the protocol is a reproducible mitigation of gas exchange and hemodynamics for research in ARDS.

Lavage-induced Surfactant Depletion in Pigs As a Model of the Acute Respiratory Distress Syndrome (ARDS)

Various animal models of lung injury exist to study the complex pathomechanisms of human acute respiratory distress syndrome (ARDS) and evaluate future ...

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11.2K Views.  Charité - Universitätsmedizin, Berlin.  Herhaalde pulmonale lavages in verdoofde varkens induceren longschade lijkt op belangrijke aspecten van het menselijk acute respiratory distress syndrome (ARDS). Hiertoe worden de longen herhaaldelijk gespoeld met 0,9% zoutoplossing bij 37 ° C. Het doel van het protocol is een reproduceerbare vermindering van de gasuitwisseling en hemodynamica voor onderzoek in ARDS. 

Video: Lavage-induced Surfactant Depletion in Pigs As a Model of the Acute Respiratory Distress Syndrome ARDS

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 ...

Excitotoxic Stimulation of Brain Microslices as an In vitro Model of Stroke

Examining molecular mechanisms involved in neuropathological conditions, such as ischemic stroke, can be difficult when using whole animal systems. As such, primary or &#39;neuronal-like&#39; cell culture systems are commonly utilized. While&#160;these systems are relatively easy to work with, and are useful model systems in which various functional outcomes (such as cell death) can be readily quantified, the examined outcomes and pathways in cultured immature neurons (such as excitotoxicity-mediated cell death pathways) are not necessarily the same as those observed in mature brain, or in intact tissue. Therefore, there is the need to develop models in which cellular mechanisms in mature neural tissue can be examined. We have developed an in vitro technique that can be used to investigate a variety of molecular pathways in intact nervous tissue. The technique described herein utilizes rat cortical tissue, but this technique can be adapted to use tissue from a variety of species (such as mouse, rabbit, guinea pig, and chicken) or brain regions (for example, hippocampus, striatum, etc.). Additionally, a variety of stimulations/treatments can be used (for example, excitotoxic, administration of inhibitors, etc.). In conclusion, the brain slice model described herein can be used to examine a variety of molecular mechanisms involved in excitotoxicity-mediated brain injury.

Excitotoxic Stimulation of Brain Microslices as an In vitro Model of Stroke

We have developed a brain slice model which can be used to examine molecular mechanisms involved in excitotoxicity-mediated brain injury.&#160;This technique generates viable mature brain tissue and reduces animal numbers required for experimentation, whilst keeping the neuronal circuitry, cellular interactions, and postsynaptic compartments partly intact.

Examining molecular mechanisms involved in neuropathological conditions, such as ischemic stroke, can be difficult when using whole animal systems. As ...

Sublingual Immunotherapy as an Alternative to Induce Protection Against Acute Respiratory Infections

Sublingual route has been widely used to deliver small molecules into the bloodstream and to modulate the immune response at different sites. It has been shown to effectively induce humoral and cellular responses at systemic and mucosal sites, namely the lungs and urogenital tract. Sublingual vaccination can promote protection against infections at the lower and upper respiratory tract; it can also promote tolerance to allergens and ameliorate asthma symptoms. Modulation of lung&#8217;s immune response by sublingual immunotherapy (SLIT) is safer than direct administration of formulations by intranasal route because it does not require delivery of potentially harmful molecules directly into the airways. In contrast to intranasal delivery, side effects involving brain toxicity or facial paralysis are not promoted by SLIT. The immune mechanisms underlying SLIT remain elusive and its use for the treatment of acute lung infections has not yet been explored. Thus, development of appropriate animal models of SLIT is needed to further explore its potential advantages. 
This work shows how to perform sublingual administration of therapeutic agents in mice to evaluate their ability to protect against acute pneumococcal pneumonia. Technical aspects of mouse handling during sublingual inoculation, precise identification of sublingual mucosa, draining lymph nodes and isolation of tissues, bronchoalveolar lavage and lungs are illustrated. Protocols for single cell suspension preparation for FACS analysis are described in detail. Other downstream applications for the analysis of the immune response are discussed. Technical aspects of the preparation of Streptococcus pneumoniae inoculum and intranasal challenge of mice are also explained.
SLIT is a simple technique that allows screening of candidate molecules to modulate lungs&#8217; immune response. Parameters affecting the success of SLIT are related to molecular size, susceptibility to degradation and stability of highly concentrated formulations.

The present work illustrates the convenience of using sublingual immunotherapy to boost the innate immune response in the lungs and confer protection against acute pneumococcal pneumonia in mouse.

Sublingual route has been widely used to deliver small molecules into the bloodstream and to modulate the immune response at different sites. It has been ...

Primary Outcome Assessment in a Pig Model of Acute Myocardial Infarction

Mortality after acute myocardial infarction remains substantial and is associated with significant morbidity, like heart failure. Novel therapeutics are therefore required to confine cardiac damage, promote survival and reduce the disease burden of heart failure. Large animal experiments are an essential part in the translational process from experimental to clinical therapies. To optimize clinical translation, robust and representative outcome measures are mandatory. The present manuscript aims to address this need by describing the assessment of three clinically relevant outcome modalities in a pig acute myocardial infarction (AMI) model: infarct size in relation to area at risk (IS/AAR) staining, 3-dimensional transesophageal echocardiography (TEE) and admittance-based pressure-volume (PV) loops. Infarct size is the main determinant driving the transition from AMI to heart failure and can be quantified by IS/AAR staining. Echocardiography is a reliable and robust tool in the assessment of global and regional cardiac function in clinical cardiology. Here, a method for three-dimensional transesophageal echocardiography (3D-TEE) in pigs is provided. Extensive insight into cardiac performance can be obtained by admittance-based pressure-volume (PV) loops, including intrinsic parameters of myocardial function that are pre- and afterload independent. Combined with a clinically feasible experimental study protocol, these outcome measures provide researchers with essential information to determine whether novel therapeutic strategies could yield promising targets for future testing in clinical studies.

Reliable and accurate outcome assessment is the key for translation of preclinical therapies into clinical treatment. The current paper describes how to assess three clinically relevant primary outcome parameters of cardiac performance and damage in a pig acute myocardial infarction model.

Mortality after acute myocardial infarction remains substantial and is associated with significant morbidity, like heart failure. Novel therapeutics are ...

Open Tracheostomy Gastric Acid Aspiration Murine Model of Acute Lung Injury Results in Maximal Acute Nonlethal Lung Injury

Acid pneumonitis is a major cause of sterile acute lung injury (ALI) in humans. Acid pneumonitis spans the clinical spectrum from asymptomatic to acute respiratory distress syndrome (ARDS), characterized by neutrophilic alveolitis, and injury to both alveolar epithelium and vascular endothelium. Clinically, ARDS is defined by acute onset of hypoxemia, bilateral patchy pulmonary infiltrates and non-cardiogenic pulmonary edema. Human studies have provided us with valuable information about the physiological and inflammatory changes in the lung caused by ARDS, which has led to various hypotheses about the underling mechanisms. Unfortunately, difficulties determining the etiology of ARDS, as well as a wide range of pathophysiology have resulted in a lack of critical information that could be useful in developing therapeutic strategies.
Translational animal models are valuable when their pathogenesis and pathophysiology accurately reproduce a concept proven in both in vitro and clinical settings. Although large animal models (e.g., sheep) share characteristics of the anatomy of human trachea-bronchial tree, murine models provide a host of other advantages including: low cost; short reproductive cycle lending itself to greater data acquisition; a well understood immunologic system; and a well characterized genome leading to the availability of a variety of gene deletion and transgenic strains. A robust model of low pH induced ARDS requires a murine ALI that targets mainly the alveolar epithelium, secondarily the vascular endothelium, as well as the small airways leading to the alveoli. Furthermore, a reproducible injury with wide differences between different injurious and non-injurious insults is important.
The murine gastric acid aspiration model presented here using hydrochloric acid employs an open tracheostomy and recreates a pathogenic scenario that reproduces the low pH pneumonitis injury in humans. Additionally, this model can be used to examine interaction of a low pH insult with other pulmonary injurious entities (e.g., food particles, pathogenic bacteria).

This protocol induces acute lung injury in a mouse that has close fidelity to the pathogenesis of acid pneumonitis observed in humans. We generate a maximal acute nonlethal low pH lung injury and account for differences in rodent-human anatomic respiratory structure using an open tracheostomy coupled with circumferential pressure release.

Acid pneumonitis is a major cause of sterile acute lung injury (ALI) in humans. Acid pneumonitis spans the clinical spectrum from asymptomatic to acute ...

Oleic Acid-Injection in Pigs As a Model for Acute Respiratory Distress Syndrome

The acute respiratory distress syndrome is a relevant intensive care disease with an incidence ranging between 2.2% and 19% of intensive care unit patients. Despite treatment advances over the last decades, ARDS patients still suffer mortality rates between 35 and 40%. There is still a need for further research to improve the outcome of patients suffering from ARDS. One problem is that no single animal model can mimic the complex pathomechanism of the acute respiratory distress syndrome, but several models exist to study different parts of it. Oleic acid injection (OAI)-induced lung injury is a well-established model for studying ventilation strategies, lung mechanics and ventilation/perfusion distribution in animals. OAI leads to severely impaired gas exchange, deterioration of lung mechanics and disruption of the alveolo-capillary barrier. The disadvantage of this model is the controversial mechanistic relevance of this model and the necessity for central venous access, which is challenging especially in smaller animal models. In summary, OAI-induced lung injury leads to reproducible results in small and large animals and hence represents a well-suited model for studying ARDS. Nevertheless, further research is necessary to find a model that mimics all parts of ARDS and lacks the problems associated with the different models existing today.

In this article, we present a protocol to induce acute lung injury in pigs by central-venous injection of oleic acid. This is an established animal model for studying the acute respiratory distress syndrome (ARDS).

The acute respiratory distress syndrome is a relevant intensive care disease with an incidence ranging between 2.2% and 19% of intensive care unit ...

Bronchoalveolar Lavage and Oleic Acid-Injection in Pigs as a Double-Hit Model for Acute Respiratory Distress Syndrome (ARDS)

The treatment of ARDS continues to pose major challenges for intensive care physicians in the 21st century with mortality rates still reaching up to 50% in severe cases. Further research efforts are needed to better understand the complex pathophysiology of this disease. There are different well-established animal models to induce acute lung injury but none has been able to adequately mimic the complex pathomechanisms of ARDS. The most crucial factor for the development of this condition is the damage to the alveolar capillary unit. The combination of two well-established lung injury models allow us to mimic in more detail the underlying pathomechanism. Bronchoalveolar lavage (BAL) leads to surfactant depletion as well as alveolar collapse. The repeated instillation of fluid volumes causes subsequent hypoxemia. Surfactant depletion is a key factor of ARDS in humans. BAL is often combined with other lung injury approaches, but not with a second hit followed by oleic acid injection (OAI) yet. Oleic acid injection leads to severely impaired gas exchange, a deterioration of lung mechanics and disruption of the alveolo-capillary barrier. The OAI mimics most of the expected effects of ARDS consisting of extended inflammation of lung tissue with an increase of alveolar leakage and gas exchange impairment. A disadvantage of the combination of different models is the difficulty to determine the influence to the lung injury caused by BAL alone, OAI alone or both together. The model presented in this report represents the combination of BAL and OAI as a new double-hit lung injury model. This new model is easy to implement and an alternative to study different therapeutic approaches in ARDS in the future.

Bronchoalveolar Lavage and Oleic Acid-Injection in Pigs as a Double-Hit Model for Acute Respiratory Distress Syndrome ARDS

Different, complex animal models exist to study the pathophysiology of acute respiratory distress syndrome (ARDS). Bronchoalveolar lavage and oleic acid injection induced lung injury is suitable as a new double-hit animal model for studying the acute respiratory distress syndrome.

The treatment of ARDS continues to pose major challenges for intensive care physicians in the 21st century with mortality rates still reaching up to 50% ...

Halogenated Agent Delivery in Porcine Model of Acute Respiratory Distress Syndrome via an Intensive Care Unit Type Device

Acute respiratory distress syndrome (ARDS) is a common cause of hypoxemic respiratory failure and death in critically ill patients, and there is an urgent need to find effective therapies. Preclinical studies have shown that inhaled halogenated agents may have beneficial effects in animal models of ARDS. The development of new devices to administer halogenated agents using modern intensive care unit (ICU) ventilators has significantly simplified the dispensing of halogenated agents to ICU patients. Because previous experimental and clinical research suggested potential benefits of halogenated volatiles, such as sevoflurane or isoflurane, for lung alveolar epithelial injury and inflammation, two pathophysiologic landmarks of diffuse alveolar damage during ARDS, we designed an animal model to understand the mechanisms of the effects of halogenated agents on lung injury and repair. After general anesthesia, tracheal intubation, and the initiation of mechanical ventilation, ARDS was induced in piglets via the intratracheal instillation of hydrochloric acid. Then, the piglets were sedated with inhaled sevoflurane or isoflurane using an ICU-type device, and the animals were ventilated with lung-protective mechanical ventilation during a 4 h period. During the study period, blood and alveolar samples were collected to evaluate arterial oxygenation, the permeability of the alveolar-capillary membrane, alveolar fluid clearance, and lung inflammation. Mechanical ventilation parameters were also collected throughout the experiment. Although this model induced a marked decrease in arterial oxygenation with altered alveolar-capillary permeability, it is reproducible and is characterized by a rapid onset, good stability over time, and no fatal complications.
We have developed a piglet model of acid aspiration that reproduces most of the physiological, biological, and pathological features of clinical ARDS, and it will be helpful to further our understanding of the potential lung-protective effects of halogenated agents delivered through devices used for inhaled ICU sedation.

We describe a model of hydrochloric acid-induced acute respiratory distress syndrome (ARDS) in piglets receiving sedation with halogenated agents, isoflurane and sevoflurane, through a device used for inhaled intensive care sedation. This model can be used to investigate the biological mechanisms of halogenated agents on lung injury and repair.

Acute respiratory distress syndrome (ARDS) is a common cause of hypoxemic respiratory failure and death in critically ill patients, and there is an urgent ...

Surfactant Depletion Combined with Injurious Ventilation Results in a Reproducible Model of the Acute Respiratory Distress Syndrome (ARDS)

Various animal models exist to study the complex pathomechanisms of the acute respiratory distress syndrome (ARDS). These models include pulmo-arterial infusion of oleic acid, infusion of endotoxins or bacteria, cecal ligation and puncture, various pneumonia models, lung ischemia/reperfusion models and, of course, surfactant depletion models, among others. Surfactant depletion produces a rapid, reproducible deterioration of pulmonary gas exchange and hemodynamics and can be induced in anesthetized pigs using repeated lung lavages with 0.9% saline (35 mL/kg body weight, 37 &#176;C). The surfactant depletion model supports investigations with standard respiratory and hemodynamic monitoring with clinically applied devices. But the model suffers from a relatively high recruitability and ventilation with high airway pressures can immediately reduce the severity of the injury by reopening atelectatic lung areas. Thus, this model is not suitable for investigations of ventilator regimes that use high airway pressures. A combination of surfactant depletion and injurious ventilation with high tidal volume/low positive end-expiratory pressure (high Tv/low PEEP) to cause ventilator induced lung injury (VILI) will reduce the recruitability of the resulting lung injury. The advantages of a timely induction and the possibility to perform experimental research in a setting comparable to an intensive care unit are preserved.

Surfactant Depletion Combined with Injurious Ventilation Results in a Reproducible Model of the Acute Respiratory Distress Syndrome ARDS

A combination of surfactant washout using 0.9% saline (35 mL/kg body weight, 37 &#176;C) and high tidal volume ventilation with low PEEP to cause moderate ventilator induced lung injury (VILI) results in experimental acute respiratory distress syndrome (ARDS). This method provides a model of lung injury with low/limited recruitability to study the effect of various ventilation strategies for extended periods.

Various animal models exist to study the complex pathomechanisms of the acute respiratory distress syndrome (ARDS). These models include pulmo-arterial ...

Oleic Acid&#45;Induced Acute Respiratory Decompensation in Pigs

A Porcine Model of Acute Respiratory Failure with a Continuous Infusion of Oleic Acid

This protocol outlines an acute respiratory distress model utilizing centrally administered oleic acid infusion in Yorkshire pigs. Prior to experimentation, each pig underwent general anesthesia, endotracheal intubation, and mechanical ventilation, and was equipped with bilateral&#160;jugular vein central vascular access catheters. Oleic acid was administered through a dedicated pulmonary artery catheter at a rate of 0.2 mL/kg/h. The infusion lasted for 60-120 min, inducing respiratory distress. Throughout the experiment, various parameters including heart rate, respiratory rate, arterial blood pressure, central venous pressure, pulmonary artery pressure, pulmonary capillary wedge pressure, end-tidal carbon dioxide, peak airway pressures, and plateau pressures were monitored. Around the 60 min mark, decreases in partial arterial oxygen pressure (PaO2) and fraction of oxygen-saturated hemoglobin (SpO2) were observed. Periodic hemodynamic instability, accompanied by acute increases in pulmonary artery pressures, occurred during the infusion. Post-infusion, histological analysis of the lung parenchyma revealed changes indicative of parenchymal damage and acute disease processes, confirming the effectiveness of the model in simulating acute respiratory decompensation.

Author Spotlight: Exploring Venous Waveforms for Non&#45;Invasive Respiratory Monitoring in Pigs

Infusing oleic acid continuously into the pulmonary artery of an anesthetized adult pig induces acute respiratory failure, enabling controlled experimentation during acute respiratory decompensation.

Lavage-induced Surfactant Depletion in Pigs As a Model of the Acute Respiratory Distress Syndrome (ARDS)

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