We gratefully acknowledge the excellent technical assistance of Birgit Brandt. This study was supported by a grant of the German Federal Ministry of Education and Research (FKZ 13GW0240A-D).

The experiments were conducted at the Department of Experimental Medicine, Charit&#233; - University Medicine, 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 (G0229/18). The principles of laboratory animal care were used in all experiments and are in accordance with the guidelines of the European and German Society of Laboratory Animal Sciences.
1. Laboratory animals and animal welfare
<ol>
	<li>Conduct all the experiments in deeply anesthetized male pigs (German Landrace &#215; Large White) of 3-4 months of age with a body weight (bw) of 30-40 kg.</li>
</ol>
2. Anesthesia, intubation, and mechanical ventilation
<ol>
	<li>Do not provide dry food for 12 h prior to anesthesia to avoid a full stomach of the pigs. Allow free access to water and straw/hay to minimize stress.</li>
	<li>Premedicate with an intramuscular injection of a combination of azaperone (3 mg/kg bw), atropine (0.03 mg/kg bw), ketamine (25 mg/kg bw), and xylazine (3.5 mg/kg bw) into the neck musculature of the pig, while the animals are still kept in their housing facility to minimize stress. 
	&#8203;NOTE: Daily training of petting the animal&#39;s neck while feeding a few sugar cubes prior to the experiment and applying the injection while feeding sugar cubes in the trained fashion will facilitate a smooth premedication and reduce stress further.
	<ol>
		<li>Place the animal onto a stretcher and cover the eyes with a cloth for transportation once an adequate level of anesthesia is reached.</li>
		<li>Transfer the pig to the surgical theater and always ensure sufficient spontaneous breathing.</li>
		<li>Take an oxygen cylinder, fitting tubing, and mask to provide supplemental oxygen while transporting the pigs, if the housing facilities are not adjacent to the laboratory.</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>Use a peripheral vein catheter (usually 18 or 20 G) to gain venous access. Place the peripheral vein catheter into one of the ear veins after a wipe-down procedure with alcohol swaps.
	<ol>
		<li>Start an infusion with a balanced crystalloid solution and ensure the correct placement of the catheter for subsequent infusion of anesthetics.</li>
		<li>Infuse 500 mL of a balanced crystalloid solution as bolus i.v. followed by continuous infusion of 4 mL/kg/h for fluid support.</li>
		<li>Start monitoring the peripheral oxygen saturation (SpO2) by securing the SpO2-Sensor at one of the ears or the tail.</li>
	</ol>
	</li>
	<li>Induce anesthesia by injecting propofol (about 5-10 mg/kg - the exact dose depends on the effect of the premedication and differs from animal to animal) for orotracheal intubation. 
	&#8203;NOTE: Prior injection of an opioid will facilitate intubation further but requires ample experience to avoid a premature apnea of the animal. An injection of 100 &#181;g of fentanyl (fentanyl citrate, 100 &#181;g/mL) may be repeated until the spontaneous respiratory rate slows down to about 20/min before injecting propofol.</li>
	<li>Intubate the animal with a cuffed endotracheal tube (7.5 - 8.0 mm ID) and a laryngoscope designed for large animals (straight blade of about 25 cm length). 
	NOTE: Intubation is easiest in the prone position as described in detail by Theisen et al.8.
	<ol>
		<li>Verify the placement of the endotracheal tube by observing the typical waveform of CO2 during expiration on the CO2-monitor (capnograph).</li>
		<li>Use auscultation to check for equal bilateral breath sounds. 
		NOTE: The pigs can be mechanically ventilated with manual compression of the rib cage from both sides while supplying oxygen with a high flow in case of failed or delayed intubation.</li>
	</ol>
	</li>
	<li>Set the fraction of inspired oxygen (FIO2) to 1.0, respirator frequency to 15-20/min, tidal volume to 8-9 mL/kg bw, inspiration to expiration ratio (I:E) to 1:1.5, and apply a positive end-expiratory pressure (PEEP) of 5 cmH2O to start mechanical ventilation. 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>Use a continuous i.v. infusion of thiopentone (20 mg/kg/h) and fentanyl (7 &#181;g/kg/h) to maintain anesthesia. 
		&#8203;NOTE: The necessary dosage may vary from animal to animal and between experimental settings. It is essential to maintain a sufficient depth of anesthesia during the course of the experiment for animal welfare and scientific reasons.</li>
		<li>Monitor the animal closely for stress/pain reactions (such as an increase in heart rate, blood pressure, or respiratory rate) during instrumentation. 
		&#8203;NOTE: Instrumentation should be possible without administering a muscle relaxant if the depth of anesthesia is sufficient.</li>
		<li>Administer a muscle relaxant, e.g., pancuronium bromide (0.15 mg/kg bw i.v. bolus, followed by a continuous infusion of 0.15 mg/kg bw/h or repeated bolus injections), if muscle relaxation is necessary for the experiment (e.g., before a surfactant depletion, before injurious ventilaion&#160;lung compliance measurements).</li>
	</ol>
	</li>
	<li>Instrumentation techniques
	<ol>
		<li>Turn the animal into the supine position.</li>
		<li>Secure the endotracheal tube and i.v. line while turning the animal.</li>
		<li>Retract the legs using bandages to stretch the skin above the planned incision sites.</li>
		<li>Sterilize the operating areas with an appropriate skin disinfectant such as an alcohol and iodine 1% solution.</li>
	</ol>
	</li>
	<li>Cannulate the external jugular vein with a central venous catheter and, in addition, introduce the introducer sheath of the pulmonary arterial catheter (PAC) into the same vein.
	<ol>
		<li>Perform a 10 cm skin incision on the line connecting the mandible and the sternum (left or right side possible).</li>
		<li>Always reassess the depth of anesthesia and adjust the dosage, if necessary.</li>
		<li>Separate the subcutaneous tissue and the platysma with tissue forceps and surgical scissors until the brachiocephalic and the sternocephalic muscles are visible.</li>
		<li>Continue with a blunt cut down procedure to separate the fascia between the muscles until the external jugular vein is visible.</li>
		<li>Use the Seldinger technique9 to cannulate the external jugular vein with the central venous catheter and the introducer sheath for later insertion of the PAC. 
		NOTE: Do not dilate the vein with a dilator as it is done in case of a percutaneous approach. This would tear the vein. Close with standard sutures. The sizes of the sheath depend on the size of the chosen PAC. A 6F introducer sheath (10 cm length) and a 5F PAC of 75 cm length in pigs of 30-40 kg body weight are&#160;typically used.</li>
	</ol>
	</li>
	<li>Cannulate the femoral artery for invasive blood pressure monitoring.
	<ol>
		<li>Identify the fold between the gracilis and sartorius muscle of the hind leg (left or right is possible) to place an arterial line. 
		NOTE: The pulsation of the femoral artery should be easily palpable.</li>
		<li>Cannulate the artery percutaneously with the Seldinger technique9.</li>
		<li>Use a direct approach if the artery is not easily palpated.
		<ol>
			<li>Cut through the skin with a 5 cm long incision and separate the subcutaneous tissue with tissue forceps and surgical scissors.</li>
			<li>Use a blunt cut down procedure separating the fascia between the muscles to the level of the femoral artery. 
			&#8203;NOTE :Do not injure the saphenous vessels by performing the cut down procedure cranial of them.</li>
			<li>Loop a ligature around the femoral artery so that the vessel can be closed in case of bleeding at the site of puncture. Avoid this step whenever possible, as it compromises blood flow to the hind leg.</li>
			<li>Cannulate the artery with the Seldinger technique9.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Calibrate the transducers against the atmosphere (zero) and either 200 mmHg (arterial line) or 50 mmHg (central venous line) and connect them to the arterial catheter and the central venous line to start monitoring.
	<ol>
		<li>Place the pressure transducers about half the height of the thorax at the estimated position of the right atrium.</li>
	</ol>
	</li>
	<li>Perform a small (4-5 cm) incision cutting through the skin above the bladder for catherization of the urinary bladder.
	<ol>
		<li>Separate the subcutaneous tissue using blunt instruments.</li>
		<li>Place a purse-string suture (1-2 cm in diameter) in the wall of the bladder. 
		NOTE: The sutures should not penetrate through all layers of the bladder wall, which would result in the loss of urine through the punctures.</li>
		<li>Perform a small incision in the middle of the suture and introduce the urinary catheter.</li>
		<li>Immediately, block the balloon with 10 mL of distilled water and pull the catheter toward the bladder wall until a light resistance is felt.</li>
		<li>Close the purse-string suture around the catheter. Close the skin using standard sutures.</li>
	</ol>
	</li>
</ol>
3. Introduction of the pulmonary artery catheter (PAC)
<ol>
	<li>Check the patency of the balloon of the PAC with 0.5-1 mL of air depending on the size of the catheter and deflate the balloon again.</li>
	<li>Connect the PAC to the pressure transducer system and calibrate the transducer against the atmosphere (zero) and 100 mmHg.</li>
	<li>Introduce the PAC through the introducer sheath with a deflated balloon for 10-15 cm (depending on the sheath length).
	<ol>
		<li>Inflate the balloon after it has left the sheath and advance the PAC further while monitoring the pressure and the typical wave forms on the pressure monitor.</li>
		<li>Push the PAC forward while the waveforms typical of the right atrium, right ventricle, and the pulmonary artery appear and stop advancing the PAC when the pulmonary capillary wedge pressure (PCWP) waveform is seen.</li>
		<li>Record the PCWP at end-expiration and deflate the balloon (see Figure 1 for the respective curves). 
		NOTE: After deflation of the balloon, the PCWP-waveform must disappear, and the pulmonary arterial pressure waveform must be visible. If the pulmonary arterial pressure waveform cannot be seen, the catheter is most likely inserted too far into a pulmonary artery and has reached an auto-wedge position. This results in a permanent occlusion of a pulmonary vessel and must be corrected by pulling the catheter back until the pulmonary arterial pressure waveform reappears thereby avoiding complications, e.g., rupture of the pulmonary vessel10. The PAC catheters are often accidentally advanced into liver veins via the inferior caval vein in pigs. Thus, if the right ventricle pressure signal is not reached after about 30 - 50 cm, pull the catheter back and start all over again.</li>
	</ol>
	</li>
</ol>
4. Pulmonary artery thermodilution technique for hemodynamic measurements
<ol>
	<li>Measure cardiac output (CO) with the thermodilution technique11.
	<ol>
		<li>Connect the thermistor and a flow through housing to the respective lumen of the PAC.</li>
		<li>Next, connect the hemodynamic monitor with the distal temperature port of the PAC (red cap).</li>
		<li>Adjust the hemodynamic monitor to the necessary mode compensating for catheter size, catheter length, injected volume, and temperature of the injected saline solution.</li>
		<li>Inject the appropriate volume of 0.9% saline as quickly as possible (usually 5 or 10 mL of 0.9% saline with a temperature of 4 &#176;C).</li>
		<li>Wait until the measurement is completed.</li>
	</ol>
	</li>
	<li>Randomize five measurements in quick succession over the respiratory cycle of the ventilator.
	<ol>
		<li>Delete the highest and the lowest values and use the remaining three values to calculate the mean.</li>
		<li>Note this mean value as the cardiac output.</li>
		<li>Measure the PCWP afterwards by inflating the catheter balloon, and deflate it after the measurement.</li>
		<li>Use the mean arterial pressure (MAP), pulmonary arterial pressure (PAP), central venous pressure (CVP), PCWP, and the CO for all further hemodynamic calculations. 
		&#8203;NOTE: The volume of saline as well as the temperature have to be entered into the monitor before the measurements. The normal saline has to be kept at the same temperature (usually &#60;5 &#176;C) for correct measurements. The size and length of the catheter have to be entered as well. Some monitors require the entry of a correction factor.</li>
		<li>For studies involving exact measurements of electrolyte balance, use 5% glucose solution instead of 0.9% saline.</li>
	</ol>
	</li>
	<li>Ensure to record all the parameters. Take simultaneous arterial and mixed venous blood samples shortly before or after CO measurements to enable calculation of the intra-pulmonary right-to-left shunt.
	<ol>
		<li>Record all needed respiratory settings and measurements to complete the data set, e.g.&#160;peak, plateau and end-expiratory pressure. 
		&#8203;NOTE: The induction of anesthesia, intubation, and full instrumentation may require 1.5 h depending on the experience and number of the investigators.</li>
	</ol>
	</li>
</ol>
5. Surfactant depletion
<ol>
	<li>Ventilate the animal with a FIO2 of 1.0.
	<ol>
		<li>Disconnect the animal from the ventilator.</li>
	</ol>
	</li>
	<li>Fill the lungs with prewarmed 0.9% saline (37 &#176;C, 35 mL/kg) with a funnel connected to the endotracheal tube.
	<ol>
		<li>For this, raise the funnel about 1 m above the animal. 
		&#8203;NOTE: The hydrostatic pressure will allocate the saline into all pulmonary sections.</li>
		<li>Immediately stop filling when the MAP decreases below &#60;50 mmHg.</li>
	</ol>
	</li>
	<li>Lower the funnel to ground level to drain the lavage fluid. Reconnect the animal to the ventilator for oxygenation.</li>
	<li>Wait until the animal recuperates and repeat the lavage a soon as possible, if required. 
	&#8203;NOTE: The necessity for a further lavage is defined by the PaO2/FIO2 ratio.
	<ol>
		<li>Take an arterial blood gas sample after 5 min following each lavage.</li>
		<li>Repeat lavages until the PaO2/FIO2 ratio (Horowitz index) decreases below 100 mmHg for at least 5 min at FIO2 1.0 and PEEP &#62; 5 cmH2O. 
		NOTE: The respiratory rate must be adjusted during the period of lavages to keep the arterial pH above 7.25 in order to prevent hemodynamic decompensation.</li>
	</ol>
	</li>
	<li>Be aware that this animal model is based on a combination of surfactant depletion and VILI. 
	NOTE: The lavages will be stopped after the PaO2/FIO2 ratio remains below 100 for 5 min NOT after 60 min as published previously for a model of surfactant washout without VILI5.
	<ol>
		<li>Commence with high Tv/low PEEP ventilation after the targeted PaO2/FIO2 has been reached. 
		&#8203;NOTE: Otherwise, an overly aggressive surfactant depletion combined with VILI will result in multiple organ failure and compromise the experiment. The duration of surfactant depletion varies between animals, since a defined PaO2/FIO2 is targeted. It may take 45 min to 1.5 h.</li>
	</ol>
	</li>
</ol>
6. Injurious ventilation with high tidal volume/low PEEP (high Tv/low PEEP)
<ol>
	<li>Keep an FIO2 of 1.0.</li>
	<li>Set the ventilator on a volume guaranteed, pressure-controlled ventilation mode.</li>
	<li>Increase the alarm threshold for peak inspiratory pressure to 60 mbar. 
	NOTE: The ventilator should apply an inspiratory pressure up to 60 mbar, but not higher.</li>
	<li>Lower the respiratory rate to 12/min and set the inspiration to expiration (I:E) ratio to 1:1.5 (resulting in an inspiration time of 2 s and expiration time of 3 s).</li>
	<li>Increase the tidal volume slowly up to 17 mL/kg bw over at least 2 min.
	<ol>
		<li>Do not increase the tidal volume further if an inspiratory pressure of 60 mbar is reached. 
		NOTE: The limited inspiratory pressure can result in tidal volume below 17 mL/kg body weight depending on the lung injury after surfactant washout. A sudden increase in tidal volume may result in barotrauma or hemodynamic decompensation. Therefore, it is of utmost importance to increase tidal volumes slowly over several minutes.</li>
	</ol>
	</li>
	<li>Reduce the PEEP to 2 mbar.</li>
	<li>Ventilate the animal for up to 2 h (see Figure 2 for the ventilator settings and the flow curve). 
	NOTE: Ventilation with high tidal volumes will result in good oxygenation of the animal, but the cyclic near-complete inflation and deflation results in structural injury of the lungs. Structural damage cannot be reversed with recruitment maneuvers, prone positioning, high PEEP, etc. The resulting injury has to be tolerated throughout the investigation. A shorter high Tv/low PEEP ventilation time may be required depending on the following experiment and duration of the investigation.</li>
</ol>
7. End of experiment and euthanasia
<ol>
	<li>Ensure that all measurements of the experimental protocol, which will follow the induction of lung injury, are performed.</li>
	<li>Inject fentanyl (at least 0.5 mg) additionally to the continuous anesthesia and wait at least 5 minutes. Inject thiopental (at least 1000 mg) quickly followed by at least 60 mmol of potassium using the central line.</li>
</ol>

<ol>
	<li>Bellani, G., 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=Epidemiology,+patterns+of+care,+and+mortality+for+patients+with+acute+respiratory+distress+syndrome+in+intensive+care+units+in+50+countries.">Epidemiology, patterns of care, and mortality for patients with acute respiratory distress syndrome in intensive care units in 50 countries.</a> JAMA. 315 (8), 788-800 (2016).</li><li>Ashbaugh, D. G., Bigelow, D. B., Petty, T. L., Levine, B. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acute+respiratory+distress+in+adults.">Acute respiratory distress in adults.</a> Lancet. 2 (7511), 319-323 (1967).</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> The Annals of Thoracic Surgery. 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 Anaesthesiologica Scandinavica. 24 (3), 231-236 (1980).</li><li>Russ, 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=Lavage-induced+surfactant+depletion+in+pigs+as+a+model+of+the+acute+respiratory+distress+syndrome+(ARDS).">Lavage-induced surfactant depletion in pigs as a model of the acute respiratory distress syndrome (ARDS).</a> Journal of Visualized Experiments: JoVE. (115), e53610 (2016).</li><li>Pomprapa, 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=Artificial+intelligence+for+closed-loop+ventilation+therapy+with+hemodynamic+control+using+the+open+lung+concept.">Artificial intelligence for closed-loop ventilation therapy with hemodynamic control using the open lung concept.</a> International Journal of Intelligent Computing and Cybernetics. 8 (1), 50-68 (2015).</li><li>Yoshida, 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=Continuous+negative+abdominal+pressure+reduces+ventilator-induced+lung+Injury+in+a+porcine+model.">Continuous negative abdominal pressure reduces ventilator-induced lung Injury in a porcine model.</a> Anesthesiology. 129 (1), 163-172 (2018).</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> Laboratory Animals. 43 (1), 96-101 (2009).</li><li>Seldinger, S. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Catheter+replacement+of+the+needle+in+percutaneous+arteriography:+A+new+technique.">Catheter replacement of the needle in percutaneous arteriography: A new technique.</a> Acta Radiologica. 39 (5), 368-376 (1953).</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> The New England Journal of Medicine. 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> American Heart Journal. 83 (3), 306-311 (1972).</li><li>Dos Santos Rocha, 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=Physiologically+variable+ventilation+reduces+regional+lung+inflammation+in+a+pediatric+model+of+acute+respiratory+distress+syndrome.">Physiologically variable ventilation reduces regional lung inflammation in a pediatric model of acute respiratory distress syndrome.</a> Respiratory Research. 21 (1), 288 (2020).</li></ol>

All authors disclose no financial or any other conflict of interests.

This article describes the induction of experimental ARDS in pigs combining surfactant depletion by repeated lung lavages and ventilation with high tidal volumes, low PEEP, and complete inflation/deflation of the lungs. This combination causes a reproducible and comparable deterioration in gas exchange and the resulting hemodynamic compromise but limits the recruitability of the lungs. Thus, this model mimics clinical ARDS with low recruitability and allows the investigation of new ventilation regimes.
<p class="jove...

The mortality of the acute respiratory distress syndrome (ARDS) remains high with values above 40%1 despite intensive research since its first description by Ashbough and Petty in 19672. Naturally, the investigation of novel therapeutic approaches is limited in the clinic due to ethical concerns and the lack of standardization of the underlying pathologies, ambient conditions, and co-medications, whereas animal models enable systematic research under standardized conditions.
Thus, experimental ARDS has been induced in either large animals (e.g., pigs) or small animals (e.g., rodents) using various methods such as pulmo-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 caused by burns and smoke inhalation or lung ischemia/reperfusion (I/R) are used3. One frequently used model of direct lung injury is surfactant depletion with lung lavages as first described by Lachmann et al. in guinea pigs4.
Surfactant depletion is a highly reproducible method that results rapidly in compromises in gas exchange and hemodynamics5. A major advantage is the possibility to apply surfactant depletion in large species which enable support research with clinically used mechanical ventilators, catheters, and monitors. However, a major disadvantage of the surfactant depletion model is the instant recruitment of atelectatic lung areas whenever high airway pressures or recruiting maneuvers, such as prone positioning, are applied. Thus, the model is not suitable to investigate, e.g., automated ventilation with high PEEP levels for prolonged times6. Yoshida et al. described a combination of surfactant depletion and ventilation with high inspiratory airway pressures to induce&#160;experimental ARDS7, but their model requires an elaborate maintenance of partial pressure of oxygen (PaO2) in a predefined corridor via repeated blood gas sampling and adjustment of the driving pressure according to a sliding table of inspiratory pressure and PEEP.
Overall, a model with an overly aggressive injurious ventilation or a laborious, repeated adjustment of the ventilation regime can result in structural damage of the lungs, which is too severe and results in subsequent multiple organ failure. Thus, this article provides a detailed description of an easily feasible model of surfactant depletion plus injurious ventilation with high Tv/low PEEP for induction of experimental ARDS, which supports research with clinically used ventilation parameters for prolonged periods.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Evita Infinity V500</td>
			<td>Dr&#228;ger</td>
			<td></td>
			<td>intensive care ventilator</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>Leader Cath Set</td>
			<td>Vygon</td>
			<td>1,15,805</td>
			<td>arterial catheter</td>
		</tr>
		<tr>
			<td>Mallinckrodt Tracheal Tube Cuffed</td>
			<td>Covidien</td>
			<td>107-80</td>
			<td>&#160;8.0 mm ID</td>
		</tr>
		<tr>
			<td>MultiCath3</td>
			<td>Vygon</td>
			<td>1,57,300</td>
			<td>3 lumen central venous catheter, 20 cm length</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>urinary catheter</td>
			<td></td>
			<td></td>
			<td>no specific model requiered</td>
		</tr>
		<tr>
			<td>Vasofix Braun&#252;le 20G</td>
			<td>B Braun</td>
			<td>4268113B</td>
			<td>peripheral vein catheter</td>
		</tr>
		<tr>
			<td>Vigilance I&#160;</td>
			<td>Edwards</td>
			<td></td>
			<td>monitor</td>
		</tr>
	</tbody>
</table>

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.

The PaO2/FIO2-ratio decreased during surfactant washout in all animals (Figure 3). The resulting hypoxemia, hypercapnia, and atelectasis caused an increase in pulmonary artery pressure. The details of the lung lavages are already described elsewhere6.
The surfactant depletion was repeated until the PaO2/FIO2 ratio remained below 100 mmHg despite mechanic...

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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.

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 ...

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JoVE Journal

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3.3K Views.  Charité – Universitätsmedizin Berlin, Corporate Member of Freie Universität Berlin, Humboldt Universität zu Berlin and Berlin Institute of Health Department of Anesthesiology and Intensive Care Medicine,  Campus Charité Mitte and Campus Virchow-Klinikum. A combination of surfactant washout using 0.9% saline (35 mL/kg body weight, 37 °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.

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

Pressure Controlled Ventilation to Induce Acute Lung Injury in Mice

Murine models are extensively used to investigate acute injuries of different organs systems (1-34). Acute lung injury (ALI), which occurs with prolonged mechanical ventilation, contributes to morbidity and mortality of critical illness, and studies on novel genetic or pharmacological targets are areas of intense investigation (1-3, 5, 8, 26, 30, 33-36). ALI is defined by the acute onset of the disease, which leads to non-cardiac pulmonary edema and subsequent impairment of pulmonary gas exchange (36). We have developed a murine model of ALI by using a pressure-controlled ventilation to induce ventilator-induced lung injury (2). For this purpose, C57BL/6 mice are anesthetized and a tracheotomy is performed followed by induction of ALI via mechanical ventilation. Mice are ventilated in a pressure-controlled setting with an inspiratory peak pressure of 45 mbar over 1 - 3 hours. As outcome parameters, pulmonary edema (wet-to-dry ratio), bronchoalveolar fluid albumin content, bronchoalveolar fluid and pulmonary tissue myeloperoxidase content and pulmonary gas exchange are assessed (2). Using this technique we could show that it sufficiently induces acute lung inflammation and can distinguish between different treatment groups or genotypes (1-3, 5). Therefore this technique may be helpful for researchers who pursue molecular mechanisms involved in ALI using a genetic approach in mice with gene-targeted deletion.

A murine model for ventilator induced lung injury is an important tool to study an acute lung injury in vivo. Here, we report an easy applicable in situ model for acute lung injury using high-pressure mechanical ventilation to induce acute failure of the lung.

Murine models are extensively used to investigate acute injuries of different organs systems (1-34). Acute lung injury (ALI), which occurs with prolonged ...

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.

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.

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

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 ...

Repeated Measurement of Respiratory Muscle Activity and Ventilation in Mouse Models of Neuromuscular Disease

Accessory respiratory muscles help to maintain ventilation when diaphragm function is impaired. The following protocol describes a method for repeated measurements over weeks or months of accessory respiratory muscle activity while simultaneously measuring ventilation in a non-anesthetized, freely behaving mouse. The technique includes the surgical implantation of a radio transmitter and the insertion of electrode leads into the scalene and trapezius muscles to measure the electromyogram activity of these inspiratory muscles. Ventilation is measured by whole-body plethysmography, and animal movement is assessed by video and is synchronized with electromyogram activity. Measurements of muscle activity and ventilation in a mouse model of amyotrophic lateral sclerosis are presented to show how this tool can be used to investigate how respiratory muscle activity changes over time and to assess the impact of muscle activity on ventilation. The described methods can easily be adapted to measure the activity of other muscles or to assess accessory respiratory muscle activity in additional mouse models of disease or injury.

This paper introduces a method for repeated measurements of ventilation and respiratory muscle activity in a freely behaving amyotrophic lateral sclerosis (ALS) mouse model throughout disease progression with whole-body plethysmography and electromyography via an implanted telemetry device.

Accessory respiratory muscles help to maintain ventilation when diaphragm function is impaired. The following protocol describes a method for repeated ...

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 ...

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 ...

Acupoint Application Combined with Acupoint Massage for Treating Constipation in a Patient with Chronic Obstructive Pulmonary Disease

Chronic obstructive pulmonary disease (COPD) is a common and frequent disease in elderly patients, with a tendency for progressive exacerbation. Constipation that reduces the quality of life and triggers the risk of diseases is a common concomitant symptom in patients with COPD. Currently, western medical treatment does not achieve the desired results for patients. A high recurrence rate accompanies it, whereas traditional Chinese medicine (TCM) has a long history and rich experience in treating chronic diseases. Both acupoint application and acupoint massage are characteristic therapies of TCM, with minor side effects, high safety, simple operation, and outstanding advantages. They are effective in treating constipation for patients with COPD and are considered an ideal alternative therapy for patients with chronic constipation. The purpose of this article is to introduce the method of acupoint application combined with acupoint massage for the treatment of constipation in patients with COPD, including the selection of points, items, treatment time, and operation procedure.

Here we demonstrate acupoint application combined with acupoint massage for treating constipation in a patient with chronic obstructive pulmonary disease (COPD).

Chronic obstructive pulmonary disease (COPD) is a common and frequent disease in elderly patients, with a tendency for progressive exacerbation. ...

Acupuncture Stimulation of Ears to Alleviate Sleeping Disorders in COPD Patients

Auricular Acupuncture as a Traditional Chinese Medicine Therapy for Chronic Obstructive Pulmonary Disease Combined with Sleep Disorders

Chronic obstructive pulmonary disease (COPD) is a clinical syndrome characterized by persistent and irreversible airflow limitation and chronic respiratory symptoms. It has a wide spectrum of complications, and sleep disorders, as part of it, are common in severe cases, especially in elderly patients. Long-term lack of sleep may lead to the aggravation of the original disease, reducing patients' quality of life. Benzodiazepines are mainly used for symptomatic treatment of COPD combined with sleep disorders. However, such drugs have the side effect of respiratory central inhibition and could probably aggravate hypoxia symptoms. Auricular acupuncture is a special method of treating physical and psychosomatic dysfunctions by stimulating specific points in the ear. This article explains the specific methods of clinical operation of auricular acupuncture in detail, including assessment of patient eligibility, medical devices used, acupuncture points, course of treatment, post-treatment care, responses to emergencies, etc. The Pittsburgh sleep quality index (PSQI) and chronic obstructive pulmonary disease assessment scale (CAT) were used as the observational index of this method. So far, clinical reports have proved that auricular acupuncture has a definite curative effect in the treatment of COPD combined with sleep disorders, and its advantages of simple operation, few adverse reactions are worthy of further study and promotion, which provide a reference for the clinical treatment of such diseases.

Author Spotlight: Utilizing Traditional Chinese Acupuncture of the Ear to Improve Sleep Disorders 

This protocol introduces an effective method to improve the sleep condition of patients with chronic obstructive pulmonary disease and alleviate cough symptoms by using auricular acupuncture techniques to percutaneously stimulate specific acupoints in the patients' ears.

Chronic obstructive pulmonary disease (COPD) is a clinical syndrome characterized by persistent and irreversible airflow limitation and chronic ...

Establishment of the KOA Model in Rabbits Using the Modified Videman Method

Application of Acupotomy in a Knee Osteoarthritis Model in Rabbit

Knee osteoarthritis (KOA) is one of the most frequently encountered diseases in the orthopedic department, which seriously reduces the quality of life of people with KOA. Among several pathogenic factors, the biomechanical imbalance of the knee joint is one of the main causes of KOA. Acupotomology believes that restoring the mechanical balance of the knee joint is the key to treating KOA. Clinical studies have shown that acupotomy can effectively reduce pain and improve knee mobility by reducing adhesion, contracture of soft tissues, and stress concentration points in muscles and tendons around the knee joint. 
In this protocol, we used the modified Videman method to establish a KOA model by immobilizing the left hindlimb in a straight position. We have outlined the method of operation and the precautions related to acupotomy in detail and evaluated the efficacy of acupotomy in conjunction with the theory of "Modulating Muscles and Tendons to Treat Bone Disorders" through the detection of the mechanical properties of quadriceps femoris and tendon, as well as cartilage mechanics and morphology. The results show that acupotomy has a protective effect on cartilage by adjusting the mechanical properties of the soft tissues around the knee joint, improving the cartilage stress environment, and delaying cartilage degeneration.

Author Spotlight: Investigating the Mechanism of Action of Acupotomy in Treating Knee Osteoarthritis

In this protocol, a knee osteoarthritis model was prepared using the modified Videman method, and the operation procedures and precautions of acupotomy are detailed. The effectiveness of acupotomy has been demonstrated by testing the mechanical properties of quadriceps femoris and tendon and the mechanical and morphological properties of cartilage.

Knee osteoarthritis (KOA) is one of the most frequently encountered diseases in the orthopedic department, which seriously reduces the quality of life of ...

A Reproducible Cartilage Impact Model to Generate Post-Traumatic Osteoarthritis in the Rabbit

Post-traumatic osteoarthritis (PTOA) is responsible for 12% of all osteoarthritis cases in the United States. PTOA can be initiated by a single traumatic event, such as a high-impact load acting on articular cartilage, or by joint instability, as occurs with anterior cruciate ligament rupture. There are no effective therapeutics to prevent PTOA currently. Developing a reliable animal model of PTOA is necessary to better understand the mechanisms by which cartilage damage proceeds and to investigate novel treatment strategies to alleviate or prevent the progression of PTOA. This protocol describes an open, drop tower-based rabbit femoral condyle impact model to induce cartilage damage. This model delivered peak loads of 579.1 &plusmn; 71.1 N, and peak stresses of 81.9 &plusmn; 10.1 MPa with a time-to-peak load of 2.4 &plusmn; 0.5 ms. Articular cartilage from impacted medial femoral condyles (MFCs) had higher rates of apoptotic cells (p = 0.0058) and possessed higher Osteoarthritis Research Society International (OARSI) scores of 3.38 &plusmn; 1.43 compared to the non-impacted contralateral MFCs (0.56 &plusmn; 0.42), and other cartilage surfaces of the impacted knee (p &lt; 0.0001). No differences in OARSI scores were detected among the non-impacted articular surfaces (p &gt; 0.05).

The open medial femoral condyle impact model in rabbits is reliable for studying post-traumatic osteoarthritis (PTOA) and novel therapeutic strategies to mitigate PTOA progression. This protocol generates an isolated cartilage defect of the posterior medial femoral condyle in rabbits using a carriage-based drop tower with an impactor head.

Post-traumatic osteoarthritis (PTOA) is responsible for 12% of all osteoarthritis cases in the United States. PTOA can be initiated by a single traumatic ...