The authors would like to thank the staff from the GreD, the Universit&#233; Clermont Auvergne, and the Centre International de Chirurgie Endoscopique (all in Clermont-Ferrand, France).

The study protocol was approved by the Animal Ethics Committee of the French Minist&#232;re de l&#8217;Education Nationale, de l&#8217;Enseignement Sup&#233;rieur et de la Recherche (approval number 01505.03) before being registered at preclinicaltrials.eu (Pre-clinical registry identifier PCTE0000129). All procedures were performed in the Centre International de Chirurgie Endoscopique, Universit&#233; Clermont Auvergne, Clermont-Ferrand, France, in accordance with the Animal Research: Reporting In Vivo Experiments (ARRIVE) guidelines36.
1. Animal preparation and anesthesia
<ol>
	<li>Piglet mode
	<ol>
		<li>Ensure that the experimental protocol is consistent with guidelines for animal experiments, including the 3R principles (replacement, reduction, and refinement) and national/international regulations.</li>
		<li>Obtain approvals from the ethics committee for care and use of experimental animals at the relevant institution before starting the protocol.</li>
		<li>Use a male white Landrace piglet (2&#8211;4 months old; weighing 10&#8211;15 kg).</li>
		<li>Place the piglet in the supine position after premedication using intramuscular azaperone (described in 1.2.2).</li>
	</ol>
	</li>
	<li>Anesthesia induction
	<ol>
		<li>Restrict animals from having food for overnight while allowing free access to water.</li>
		<li>Administer anxiolytic premedication to the piglet using intramuscular azaperone (2 mg.kg-1) behind the ear.</li>
		<li>Apply a finger pressure on the soft tissues of the auricular base of the piglet to identify the medial and lateral auricular vein.</li>
		<li>Insert a peripheral intravenous 22 G catheter in the medial or lateral auricular vein of the piglet. Follow with the catheter at a shallow angle of 45&#730; through the skin and advance until blood appears through the catheter.</li>
		<li>Induce general anesthesia with intravenous propofol (3 mg.kg-1) and sufentanil (0.3 &#181;g.kg-1)37. Check the depth of the anesthesia by lack of response to pedal reflex.</li>
	</ol>
	</li>
	<li>Tracheal intubation38,39
	<ol>
		<li>Prepare the laryngoscope using a size 4 straight Miller laryngoscope blade.</li>
		<li>Pass the laryngoscope into the pharyngeal cavity and depress the tongue with the laryngoscope blade, making the epiglottis visible.</li>
		<li>Visualize the larynx opening of the piglet prior to orotracheal intubation.</li>
		<li>Insert a 6 mm internal diameter cuffed endotracheal tube.</li>
		<li>Inflate the endotracheal tube cuff to reach a cuff pressure around 20&#8211;30 cmH2O.</li>
		<li>Fix the endotracheal tube to the piglet&#8217;s nose with micropore surgical tape.</li>
		<li>Connect to the ventilator and initiate mechanical ventilation following the settings described in section 3.</li>
	</ol>
	</li>
	<li>Sedation maintenance
	<ol>
		<li>Maintain anesthesia with continuous intravenous infusion of propofol (5 mg.kg-1.h-1) before acid-induced lung injury. The infusion of propofol will be stopped when halogenated agents are started.</li>
		<li>Add a continuous intravenous infusion of remifentanil (10&#8211;20 &#956;g.kg&#8722;1.h&#8722;1 = 0.15&#8211;0.33 &#956;g.kg&#8722;1.min&#8722;1) for pain management.</li>
		<li>Add continuous intravenous infusion of cisatracurium (0.2 mg.kg-1.h-1) for a neuromuscular blockade.</li>
		<li>Keep the body temperature of the piglet at approximately 38 &#176;C using warm blankets.</li>
		<li>Monitor electrocardiogram activity, the peripheral oxygen saturation (SpO2), and arterial pressure continuously using an external monitor.</li>
	</ol>
	</li>
	<li>Surgery
	<ol>
		<li>Insert central venous access using a surgical exposure of the right internal jugular vein and the Seldinger method to insert a 3-lumen catheter (7 French, 16 cm).
		<ol>
			<li>Make a cutaneous midline incision on the ventral aspect of the neck, 2 cm lateral from the trachea. Use surgical forceps to dissect the tissues.</li>
			<li>Localize the internal jugular vein (approximately 1&#8211;2 cm deep, lateral to the internal carotid artery) and, using the needle (18 G, 6.35 cm), make a puncture with a craniocaudal direction orientation.</li>
			<li>With the hand, insert the &#8220;J&#8221; guidewire (0.81 mm diameter, 60 cm) through the needle. Gently remove the needle and quickly insert a venous catheter with three lines into the internal jugular vein along the &#8220;J&#8221; guidewire. Remove the &#8220;J&#8221; guidewire while maintaining the venous catheter in place.</li>
			<li>Aspirate blood through each line of the venous catheter to remove the air from the different lines and flush with 5 mL of saline solution (0.9% NaCl) to rinse the three lines.</li>
			<li>Suture the skin with a 3.0 non-absorbable suture thread following the continuous Lembert pattern and fix the catheter to the skin with a single stitch and triple knots on each lateral perforation of the central venous catheter.</li>
		</ol>
		</li>
		<li>Insert an arterial line via surgical exposure of the right femoral artery and use the Seldinger method to insert the thermodilution catheter (3&#8211;5 French, 20 cm).
		<ol>
			<li>Place the right forelimb of the piglet in extension.</li>
			<li>Make a cutaneous incision on the right groin area of the piglet. Use surgical forceps to dissect the subcutaneous and muscular tissues.</li>
			<li>Localize the right femoral artery by palpating the femoral pulse (approximately 3&#8211;4 cm deep) and, using the needle (19 G, 54 mm), make a puncture with a caudocranial direction orientation.</li>
			<li>Insert the &#8220;J&#8221; guidewire through the needle. Gently remove the needle and quickly insert an arterial catheter into the femoral artery up along the guidewire. Remove the guidewire while maintaining the catheter in place.</li>
			<li>Remove the air from the arterial catheter and flush with saline solution to rinse the line.</li>
			<li>Suture the skin with a 3.0 non-absorbable suture thread following the continuous Lembert pattern and fix the catheter to the skin with a single stitch and triple knots on each lateral perforation of the arterial catheter.</li>
			<li>Plug the catheter on an arterial line tubing to allow retrieval of serial blood samples and continuous hemodynamic monitoring (arterial pressure, cardiac index, and extravascular lung water, as indexed to body weight) with a pulse contour cardiac output monitor device.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Acid-induced acute lung injury
CAUTION: Use gloves and glasses during this step to avoid any risk of contact of the acid with the skin or the eyes)
<ol>
	<li>Make 100 mL of HCl at 0.05 M and pH 1.4.</li>
	<li>Using the anatomical landmark of the last segment of the sternum, measure the distance between the tip of the endotracheal tube and the carina of the piglet.</li>
	<li>Mark this distance with a black pen on a Ch14 suction catheter.</li>
	<li>Insert the suction catheter through the endotracheal tube up to the black landmark.</li>
	<li>Gently instill 4 mL.kg-1 (bodyweight) of acid through the suction catheter for over 3 min.</li>
	<li>Remove the suction catheter.</li>
</ol>
3. Mechanical ventilation
<ol>
	<li>Use volume-controlled ventilation on an intensive care ventilator.</li>
	<li>Use a tidal volume of 6 mL.kg-1, a positive end-expiratory pressure (PEEP) of 5 cmH2O, and an inspired oxygen fraction (FiO2) of 40%.</li>
	<li>Adjust the respiratory rate to maintain the end-tidal carbon dioxide between 35 and 45 mmHg. 
	NOTE: Based on previous studies37,40,41, lung injury is considered established when the arterial oxygen tension (PaO2)-to-FiO2 ratio decreases to 25% from the baseline, approximately 1 h after airway HCl instillation.</li>
</ol>
4. Halogenated anesthetics
NOTE: Start sedation using halogenated anesthetics (sevoflurane or isoflurane) once acid-induced lung injury is achieved. The intravenous sedation using propofol should then be interrupted.
<ol>
	<li>Filling the syringe (Figure 1A): Attach the filling adapter provided by the manufacturer to the 250 mL bottle of the halogenated agent and a 60 mL syringe to the filling adapter. Turn the bottle upside down and fill the syringe by pushing and pulling the plunger. Turn the bottle upright and remove the syringe.</li>
	<li>Scavenging (Figure 1B)
	<ol>
		<li>Place the charcoal filter, used to remove halogenated hydrocarbon anesthetic gases, close to the ventilator.</li>
		<li>Remove the protective cap from the charcoal filter.</li>
		<li>Connect the charcoal filter to the expiratory valve of the ventilator with a flex tube.</li>
	</ol>
	</li>
	<li>Use the anesthetic conserving device (device used for inhaled ICU sedation) (Figure 1C) as described below.
	<ol>
		<li>Connect the ionomer membrane dryer line to the gas sampling port of the anesthetic conserving device.</li>
		<li>Connect one side of the gas sampling line to the ionomer membrane dryer line.</li>
		<li>Connect the other side of the gas sampling line to the gas analyzer.</li>
		<li>Insert the anesthetic conserving device between the Y-piece of the respiratory circuit and the endotracheal tube.</li>
		<li>Ensure that the anesthetic conserving device has the black side up and is sloped down toward the piglet.</li>
	</ol>
	</li>
	<li>Deliver inhaled sedation through the anesthetic conserving device (Figure 2).
	<ol>
		<li>Place the specific syringe in the syringe pump.</li>
		<li>Connect the anesthetic agent line to the syringe.</li>
		<li>Prime the agent line with a bolus of 1.5 mL of the halogenated agent.</li>
		<li>Adapt the initial pump rate in mL.h-1 (initial syringe pump rate settings of isoflurane and sevoflurane are 3 and 5 mL/h, respectively) to the targeted expired sevoflurane fraction (FEsevo) or the expired isoflurane fraction (FEiso) value, as displayed on the gas analyzer.</li>
		<li>Ensure that the gas analyzer displays a FEsevo %&#8211;FEiso % or equivalent minimal alveolar concentration value greater than zero. If necessary, give an additional bolus of 0.3 mL of the halogenated agent.</li>
		<li>Adapt the syringe pump rate necessary to reach a certain concentration depending on the minute volume and the targeted concentration, with rates of 2&#8211;7 mL.h-1 and 4&#8211;10 mL.h-1 being, in general, associated with expired fractions of 0.2%&#8211;0.7% and 0.5%&#8211;1.4% for isoflurane42 and sevoflurane28,43, respectively.</li>
		<li>During the experiment, continue administration of the halogenated agents with FEsevo and FEiso targets of 0.8&#8211;1.1 and 0.5&#8211;0.8, respectively.</li>
	</ol>
	</li>
</ol>
5. Measurements
<ol>
	<li>Monitoring
	<ol>
		<li>Collect different parameters as measured by the external monitor: heart rate, blood pressure, and peripheral oxygen saturation.</li>
		<li>Record parameters as measured by the ventilator: tidal volume, respiratory rate, set PEEP, auto-PEEP (by applying an expiratory hold maneuver of 5 s on the ventilator), compliance of the respiratory system, airway resistance, inspiratory plateau pressure (by applying an inspiratory hold maneuver of 2 s on the ventilator), peak inspiratory pressure, and driving pressure.</li>
		<li>Calculate the lung functional residual capacity using the Nitrogen Wash In/Wash Out method if integrated in the ventilator.</li>
		<li>Use the thermal indicator previously inserted in the femoral artery to measure the extravascular water volume of the lungs, cardiac index, and systemic vascular resistance.</li>
	</ol>
	</li>
	<li>Undiluted pulmonary edema fluid sampling to measure the net AFC rate.
	<ol>
		<li>Insert a soft 14 Fr suction catheter into a wedged position in the distal bronchus through the endotracheal tube.</li>
		<li>Sample edema fluid into a suction trap by applying gentle suction.</li>
		<li>Centrifuge all samples at 240 x g at 4 &#176;C for 10 min in a refrigerated centrifuge.</li>
		<li>Collect the supernatants. 
		NOTE: Total protein concentration in undiluted pulmonary edema fluid is measured with a colorimetric method. Because the rate of clearance of edema fluid from the alveolar space is much faster than the rate of protein removal, the net AFC rate was calculated as Percent AFC = 100 &#215; [1 - (initial edema protein/final edema total protein)] and thereafter was reported as %/h37. Undiluted pulmonary edema fluid samples are collected from the animals at baseline and 4 h later, as previously described34,44,45,46,47,48,49.</li>
	</ol>
	</li>
	<li>Mini bronchoalveolar lavage sampling.
	<ol>
		<li>Insert a soft 14 Fr suction catheter into a wedged position in a distal bronchus through the endotracheal tube.</li>
		<li>Instill 50 mL of a 0.9% sodium chloride solution into the suction catheter.</li>
		<li>Promptly sample the fluid into a suction trap.</li>
		<li>Collect the mini bronchoalveolar lavage. 
		NOTE: Total protein concentration in mini BAL is measured with a colorimetric method and, for example, the levels of proinflammatory cytokines, such as TNF-&#945;, IL-6, IL-1&#946;, and IL-18, are measured using a multiplex immunoassay method. Samples are collected 4 h after the acid-induced lung injury.</li>
	</ol>
	</li>
	<li>Blood gas analysis
	<ol>
		<li>Collect arterial blood gases through the arterial line in a 3 mL BD Preset syringe with BD Luer-Lok tip at baseline. Immediately measure PaO2/FiO2, PaCO2, pH, serum lactate, and serum creatinine using a point-of-care blood gas analyzer.</li>
		<li>Repeat this step every hour for 4 h after acid instillation.</li>
	</ol>
	</li>
	<li>Lung sampling
	<ol>
		<li>Sacrifice the piglet with an intravenous injection of pentobarbital (150 mg.kg-1) at the end of the experiment (4 h after acid-induced lung injury).</li>
		<li>Dissect and remove the whole lungs. Fix with alcohol acetified formalin.</li>
		<li>Embed in paraffin and slice at a 10 &#956;m thickness.</li>
		<li>Stain with hematoxylin and eosin. 
		NOTE: Histological evidence of lung injury can be assessed using a standardized histology injury score50.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

This article describes a reproducible experimental model of ARDS induced by the intratracheal instillation of HCl in piglets to investigate the lung-protective effects of halogenated volatiles, such as sevoflurane or isoflurane, delivered using an anesthetic conserving device.
The primary goal of this study was to develop an experimental model of ARDS in which volatile agents could be delivered by an anesthetic conserving device, such as those used in ICU patients. Although some effects of hal...

Acute respiratory distress syndrome (ARDS) is a common cause of hypoxemic respiratory failure and death in critically ill patients1. It is characterized by both diffuse alveolar epithelial and endothelial injuries, leading to increased permeability and pulmonary edema, altered alveolar fluid clearance (AFC), and worsened respiratory distress2. The resorption of alveolar edema and recovery from ARDS require epithelial fluid transport through the alveoli to remain intact, which suggests that a therapy improving AFC could be useful3,4. Although lung-protective ventilation and a restrictive strategy for intravenous fluid therapy have proven beneficial in improving outcomes2,5, they are still associated with high mortality and morbidity6. Therefore, there is an urgent need to develop effective therapies for the syndrome and to better understand the precise mechanisms through which such therapies might work.
Halogenated anesthetics, such as isoflurane or sevoflurane, have been widely used for general anesthesia in the operating room. Sevoflurane is associated with decreased inflammation in the lungs of patients undergoing thoracic surgery and with a decrease in postoperative pulmonary complications, such as ARDS7. Similar results have been found in a meta-analysis of patients after cardiac surgery8. Halogenated volatiles also have a bronchodilatory effect9,10 and perhaps some properties that protect several organs, such as the heart8,11 and the kidneys12,13,14. Recently, there has been growing interest in the clinical use of inhaled anesthetics as sedatives in the intensive care unit (ICU). Both animal and human studies support the protective effects of pretreatment with halogenated agents before prolonged ischemia of the liver15, the brain16, or the heart11. Halogenated agents also have potential pharmacokinetic and pharmacodynamic advantages over other intravenous agents for the sedation of critically ill patients, including a rapid onset of action and fast offset due to little accumulation in tissues. Inhaled halogenated agents decrease intubation times in comparison with intravenous sedation in patients undergoing cardiac surgery17. Several studies support the safety and efficacy of halogenated agents in the sedation of ICU patients18,19,20. In experimental models of ARDS, inhaled sevoflurane improves gas exchange21,22, reduces alveolar edema21,22, and attenuates both pulmonary and systemic inflammation23. Isoflurane also ameliorates lung repair after injury by maintaining the integrity of the alveolar-capillary barrier, possibly by modulating the expression of a key tight junction protein24,25,26. In addition, mouse macrophages that were cultured and treated with isoflurane had better phagocytic effects on neutrophils than macrophages that were not treated with isoflurane27.
However, the precise biological pathways and mechanisms accounting for the lung-protective properties of volatile anesthetics remain largely unknown to date, requiring further investigation18. Additional studies are also warranted to investigate the precise effects of sevoflurane on lung injury and to verify whether experimental evidence can be translated to patients. The first randomized control trial from our team found that the administration of inhaled sevoflurane in patients with ARDS was associated with oxygenation improvement and decreased levels of both pro-inflammatory cytokines and lung epithelial injury markers, as assessed by plasma and alveolar soluble receptors for advanced glycation end products (sRAGE)28. As sRAGE is now considered as a marker of alveolar type 1 cell injury and a key mediator of alveolar inflammation, these results could suggest some beneficial effects of sevoflurane on the lung alveolar epithelial injury21,29,30.
The use of halogenated agents for inhaled ICU sedation has long required operating room anesthesia ventilators and gas vaporizers to be deployed in the ICU. Since then, anesthetic reflectors suitable for the use with modern critical care ventilators have been developed for specific use in the ICU31. These devices feature modified heat and moisture exchanging filters inserted between the Y-piece of the respiratory circuit and the endotracheal tube. They allow the administration of halogenated agents, with isoflurane and sevoflurane being the most frequently used, and they consist of a porous polypropylene evaporator rod, into which a liquid agent, delivered by a specific syringe pump, is released. The halogenated agent is absorbed during expiration by a reflecting medium contained in the device and it is released during the next inspiration, allowing recirculation of approximately 90% of the expired halogenated agent31,32. Recently, a miniaturized version of the device was developed with an instrumental dead space of 50 mL, making it even more suitable for use during ultra-protective ventilation in ARDS patients, with tidal volumes that could be as low as 200 mL31. Such a miniaturized device has never been studied in an experimental piglet model of ARDS.
Because previous research supports the promising roles of halogenated volatiles in lung alveolar inflammation and injury during ARDS, we designed an experimental animal model to achieve a translational understanding of the mechanisms of the effects of halogenated agents on lung injury and repair33,34,35. In this study, we developed a model of hydrochloric acid (HCl)-induced ARDS in piglets in whom inhaled sedation can be delivered using the miniaturized version of the anesthetic conserving device, an ICU-type device. This large animal model of ARDS could be used to further our understanding of the potential lung-protective effects of inhaled halogenated agents.

<table>
	<tbody>
		<tr>
			<td>Tracheal intubation</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Endotracheal tube 6-mm</td>
			<td>Covidien</td>
			<td>18860</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Animal preparation</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Central venous catheter 3-lumens catheter (7 French - 16 cm)</td>
			<td>Arrow</td>
			<td>CV-12703</td>
			<td></td>
		</tr>
		<tr>
			<td>Pulse contour cardiac output monitor PiCCO catheter (3-5 French - 20 cm)</td>
			<td>Getinge Pulsion Medical System</td>
			<td>catheter</td>
			<td></td>
		</tr>
		<tr>
			<td>Warm blankets WarmTouch5300</td>
			<td>MedTronic</td>
			<td>5300</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Monitoring</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>External monitor IntelliVue MP40</td>
			<td>Phillips</td>
			<td>MNT 142</td>
			<td></td>
		</tr>
		<tr>
			<td>Point-of-care blood gas analyzer Epoc&#174; Blood Analysis System</td>
			<td>Siemens</td>
			<td>20093</td>
			<td></td>
		</tr>
		<tr>
			<td>Pulse contour cardiac output monitor PiCCO Device PulsioFlex Monitor</td>
			<td>Getinge Pulsion Medical System</td>
			<td>PulsioFlex</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Mechanical ventilation</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ventilator Engstr&#246;m Carestation</td>
			<td>General Electrics</td>
			<td>Engstr&#246;m</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Halogenated anesthetics</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Anaconda Syringe</td>
			<td>SedanaMedical</td>
			<td>26022</td>
			<td></td>
		</tr>
		<tr>
			<td>Anesthetic conserving device AnaConDa-S</td>
			<td>SedanaMedical</td>
			<td>26050</td>
			<td></td>
		</tr>
		<tr>
			<td>Charcoal filter FlurAbsorb</td>
			<td>SedanaMedical</td>
			<td>26096</td>
			<td></td>
		</tr>
		<tr>
			<td>Filling Adaptaters</td>
			<td>SedanaMedical</td>
			<td>26042</td>
			<td></td>
		</tr>
		<tr>
			<td>Ionomer membrane dryer line Nafion</td>
			<td>SedanaMedical</td>
			<td>26053</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Products</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Propofol</td>
			<td>Mylan</td>
			<td>66617123</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Virbac</td>
			<td>QN01AB06</td>
			<td></td>
		</tr>
		<tr>
			<td>Pentobarbital</td>
			<td>PanPharma</td>
			<td>68942457</td>
			<td></td>
		</tr>
		<tr>
			<td>Sevoflurane</td>
			<td>Abbvie</td>
			<td>N01AB08</td>
			<td></td>
		</tr>
		<tr>
			<td>Sufentanil</td>
			<td>Mylan</td>
			<td>62404996</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

For this experiment, 25 piglets were anesthetized and divided in two groups: 12 piglets in the untreated group (SHAM group) and 13 piglets in the acid-injured group (HCl group). No piglet died before the end of the experiment. A two-way repeated-measures analysis of variance (RM-ANOVA) indicated a significant time by group interaction (P &#60; 10&#8722;4) with a detrimental effect of HCl-induced ARDS on PaO2/FiO2, compared to sham animals without ARDS (Figure 3</stron...

Watch this Scientific Journal Video about Halogenated Agent Delivery in Porcine Model of Acute Respiratory Distress Syndrome via an Intensive Care Unit Type Device at JoVE.com

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

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 ...

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2.6K Views.  CHU Clermont-Ferrand. 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.

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

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 ...

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 ...

A Protocol to Set Up Needle-Free Connector with Positive Displacement on Central Venous Catheter in Intensive Care Unit

Needle-free connectors were initially designed and promoted to avoid blood exposure for healthcare workers. Some recent data suggest that the latest generation of connectors (with positive displacement) may be of interest for reducing central venous line infections. We have been using needle-free connectors for several years in our intensive care unit and here we present a protocol for installing these connectors on central venous catheters. After insertion of the catheter and control of the permeability of the lines, the connectors must be purged with 0.9% NaCl before being connected. The connectors replace all disposable caps used on infusion stopcocks and manifolds. All the connectors are changed every 7 days as recommended by the manufacturer (except when there is macroscopic contamination, which requires an immediate change of the connector). Before each injection, the connector must be disinfected for at least 3 seconds with 70% isopropyl alcohol. The connectors must not be disconnected (unless changed), as the injection is done through the device. Setting up the connectors slightly increases the total time required to place the catheter and there is no formal evidence that these connectors reduce the incidence of infectious or thrombotic complications. However, these devices simplify the management of central venous lines and prevent the catheter circuit from &#34;opening&#34; once it has been sterilely installed.

We present a protocol to show the installation of a needle-free connector with positive displacement on a central venous catheter.

Needle-free connectors were initially designed and promoted to avoid blood exposure for healthcare workers. Some recent data suggest that the latest ...

Inducing Acute Lung Injury in Mice by Direct Intratracheal Lipopolysaccharide Instillation

Airway administration of lipopolysaccharide (LPS) is a common way to study pulmonary inflammation and acute lung injury (ALI) in small animal models. Various approaches have been described, such as the inhalation of aerosolized LPS as well as nasal or intratracheal instillation. The presented protocol describes a detailed step-by-step procedure to induce ALI in mice by direct intratracheal LPS instillation and perform FACS analysis of blood samples, bronchoalveolar lavage (BAL) fluid, and lung tissue. After intraperitoneal sedation, the trachea is exposed and LPS is administered via a 22 G venous catheter. A robust and reproducible inflammatory reaction with leukocyte invasion, upregulation of proinflammatory cytokines, and disruption of the alveolo-capillary barrier is induced within hours to days, depending on the LPS dosage used. Collection of blood samples, BAL fluid, and lung harvesting, as well as the processing for FACS analysis, are described in detail in the protocol. Although the use of the sterile LPS is not suitable to study pharmacologic interventions in infectious diseases, the described approach offers minimal invasiveness, simple handling, and good reproducibility to answer mechanistic immunological questions. Furthermore, dose titration as well as the use of alternative LPS preparations or mouse strains allow modulation of the clinical effects, which can exhibit different degrees of ALI severity or early vs. late onset of disease symptoms.

Presented here is a step-by-step procedure to induce acute lung injury in mice by direct intratracheal lipopolysaccharide instillation and to perform FACS analysis of blood samples, bronchoalveolar lavage fluid, and lung tissue. Minimal invasiveness, simple handling, good reproducibility, and titration of disease severity are advantages of this approach.

Airway administration of lipopolysaccharide (LPS) is a common way to study pulmonary inflammation and acute lung injury (ALI) in small animal models. ...

Assessment of Dependence in Activities of Daily Living Among Older Patients in an Acute Care Unit

During an acute medical problem, older people may lose functional independence. ADL scales are used to assess this loss of independence. The simplest and most convenient ADL scale is the Katz Index, which measures six ADL: bathing, dressing, toileting, transferring, continence, and feeding. A lower ADL score indicates greater loss of functional independence. The ADL score prior to the acute medical problem (baseline) is estimated by questioning the patient or the caregivers, and this score is then compared with that on hospital admission. The ADL score should be monitored from hospital admission until discharge to allow early detection of changes in functional independence. Identifying any loss of functional independence before and during hospitalization provides information to caregivers regarding the risk of short-term mortality risk and complications, and the prognosis after hospitalization.

During acute medical problems, older people may lose independence in activities of daily living (ADL). Assessment of baseline ADL and ADL on admission can guide personalized treatment plans aimed at preventing nosocomial dependence and ensuring better functional outcomes.

During an acute medical problem, older people may lose functional independence. ADL scales are used to assess this loss of independence. The simplest and ...

A Reproducible Intensive Care Unit-Oriented Endotoxin Model in Rats

Sepsis and septic shock remain the leading cause of death in intensive care units. Despite significant improvements in sepsis management, mortality still ranges between 20 and 30%. Novel treatment approaches in order to reduce sepsis-related multiorgan failure and death are urgently needed. Robust animal models allow for one or multiple treatment approaches as well as for testing their effect on physiological and molecular parameters. In this article, a simple animal model is presented.
First, general anesthesia is induced in animals either with the use of volatile or by intraperitoneal anesthesia. After placement of an intravenous catheter (tail vein), tracheostomy, and insertion of an intraarterial catheter (tail artery), mechanical ventilation is started. Baseline values of mean arterial blood pressure, arterial blood oxygen saturation, and heart rate are recorded.
The injection of lipopolysaccharides (1 milligram/kilogram body weight) dissolved in phosphate-buffered saline induces a strong and reproducible inflammatory response via the toll-like receptor 4. Fluid corrections as well as the application of norepinephrine are performed based on well-established protocols.
The animal model presented in this article is easy to learn and strongly oriented towards clinical sepsis treatment in an intensive care unit with sedation, mechanical ventilation, continuous blood pressure monitoring, and repetitive blood sampling. Also, the model is reliable, allowing for reproducible data with a limited number of animals in accordance with the 3R (reduce, replace, refine) principles of animal research. While animal experiments in sepsis research cannot easily replaced, repetitive measurements allow for a reduction of animals and keeping septic animals anesthetized diminishes suffering.

Here, we present a reproducible intensive care unit-oriented endotoxin model in rats.

Sepsis and septic shock remain the leading cause of death in intensive care units. Despite significant improvements in sepsis management, mortality still ...

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 ...

Porcine Liver Transplantation Without Veno-Venous Bypass As an Extended Criteria Donor Model

Liver transplantation is regarded as the gold standard for the treatment of a variety of fatal hepatic diseases. However, unsolved issues of chronic graft failure, ongoing organ donor shortages, and the increased use of marginal grafts call for the improvement of current concepts, such as the implementation of organ machine perfusion. In order to evaluate new methods of graft reconditioning and modulation, translational models are required. With respect to anatomical and physiological similarities to humans and recent progress in the field of xenotransplantation, pigs have become the main large animal species used in transplantation models. After the initial introduction of a porcine orthotopic liver transplant model by Garnier et al. in 1965, several modifications have been published over the past 60 years.
Due to specifies-specific anatomical traits, a veno-venous bypass during the anhepatic phase is regarded as a necessity to reduce intestinal congestion and ischemia resulting in hemodynamic instability and perioperative mortality. However, the implementation of a bypass increases the technical and logistical complexity of the procedure. Furthermore, associated complications such as air embolism, hemorrhage, and the need for a simultaneous splenectomy have been reported previously.
In this protocol, we describe a model of porcine orthotopic liver transplantation without the use of a veno-venous bypass. The engraftment of donor livers after static cold storage of 20 h - simulating extended criteria donor conditions - demonstrates that this simplified approach can be performed without significant hemodynamic alterations or intraoperative mortality and with regular uptake of liver function (as defined by bile production and liver-specific CYP1A2 metabolism). The success of this approach is ensured by an optimized surgical technique and a sophisticated anesthesiologic volume and vasopressor management.
This model should be of special interest for workgroups focusing on the immediate postoperative course, ischemia-reperfusion injury, associated immunological mechanisms, and the reconditioning of extended criteria donor organs.

In this protocol, a model of porcine orthotopic liver transplantation after static cold storage of donor organs for 20 h without the use of a veno-venous bypass during engraftment is described. The approach uses a simplified surgical technique with minimization of the anhepatic phase and sophisticated volume and vasopressor management.

Liver transplantation is regarded as the gold standard for the treatment of a variety of fatal hepatic diseases. However, unsolved issues of chronic graft ...

Hemodynamic Precision in the Neonatal Intensive Care Unit using Targeted Neonatal Echocardiography

Targeted neonatal echocardiography (TnECHO) refers to the use of comprehensive echocardiographic evaluation and physiologic data to obtain accurate, reliable, and real-time information on developmental hemodynamics in sick newborns. The comprehensive assessment is based on a multiparametric approach that overcomes the reliability issues of individual measurements, allows for earlier recognition of cardiovascular compromise and promotes enhanced diagnostic precision and timely management. TnECHO-driven research has led to an enhanced understanding of the mechanisms of illness and the development of predictive models to identify at-risk populations. This information may then be used to formulate a diagnostic impression and provide individualized guidance for the selection of cardiovascular therapies. TnECHO is based on the expert consultative model in which a neonatologist, with advanced training in neonatal hemodynamics, performs comprehensive and standardized TnECHO assessments. The distinction from point of care ultrasonography (POCUS), which provides limited and brief one-time assessments, is important. Neonatal hemodynamics training is a 1-year structured program designed to optimize image acquisition, measurement analysis, and hemodynamic knowledge (physiology, pharmacotherapy) to support cardiovascular decision-making. Neonatologists with hemodynamic expertise are trained to recognize deviations from normal anatomy and appropriately refer cases of possible structural abnormalities. We provide an outline of neonatal hemodynamics training, the standardized TnECHO imaging protocol, and an example of representative echo findings in a hemodynamically significant patent ductus arteriosus.

Presented here is a protocol to perform comprehensive neonatal echocardiography by trained neonatologists in the neonatal intensive care unit. The trained individuals provide longitudinal assessments of heart function, systemic and pulmonary hemodynamics in a consultative role. The manuscript also describes the requirements to become a fully trained neonatal hemodynamics specialist.

Targeted neonatal echocardiography (TnECHO) refers to the use of comprehensive echocardiographic evaluation and physiologic data to obtain accurate, ...

Enhancing Pulmonary Infection Treatment Using M-ROSE

Microbiological Rapid On-Site Evaluation for Pulmonary Infectious Diseases

The prompt initiation of empirical anti-infective therapy is crucial in patients presenting with unexplained pulmonary infection. Although imaging acquisition is relatively straightforward in clinical practice, its lack of specificity often necessitates additional time-consuming tests such as sputum culture, bronchoalveolar-lavage fluid culture, or genetic sequencing to identify the underlying etiology of the disease accurately. Moreover, the limited efficacy of empirical anti-infective treatment may contribute to antibiotic misuse. Recent advancements in interpreting microbial background on rapid on-site evaluation (ROSE) slides have enabled clinicians to promptly obtain samples through bronchoscopy (e.g., alveolar lavage, mucosal brushing, tissue clamp), facilitating bedside staining and interpretation that provides essential microbial background information. Consequently, this establishes a foundation for developing targeted anti-infection treatment and individualized drug therapy plans. With a better understanding of which pathogens are causing infections in real-time, physicians can avoid unnecessary broad-spectrum antibiotics contributing to antibiotic resistance. Establishing a rapid and standardized M-ROSE workflow within respiratory medicine departments or intensive care units will greatly assist physicians in formulating accurate treatment strategies for patients, which holds significant clinical implications.

Author Spotlight: Advancing Pathogen Detection and Disease Assessment in Real-Time Using M-ROSE

The protocol here demonstrates a fast and standardized microbiological rapid on-site evaluation (M-ROSE) workflow, including three steps: slide making, staining, and interpretation. This protocol will help physicians make rapid clinical decisions.