Name: halogenated agent delivery in porcine model of acute respiratory distress syndrome via an intensive care unit type device
Uploaded: 2020-09-24T13:30:00.000+00:00
Duration: 9 min 36 s
Description: 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

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 halogenated agents have been previously studied in animal models, the strength of our model is that it is a clinically relevant, translational model to further our understanding of such effects. Another advantage of this model is that substantial lung injury can be induced in animals larger than mice with low mortality over time. Indeed, an important consideration in the choice of an animal model of ARDS should be the experimental question to be addressed51. In a mouse model, experimental techniques to induce lung injuries, such as intravenous oleic acid52, lavage-induced surfactant depletion40, and high-stretch mechanical ventilation53, can induce intensive injury over a timescale from hours to days, but they do not allow the investigation of lung injury repair/resolution. Furthermore, some challenges of the animal model (e.g., extremely large tidal volumes in certain models of ventilator-induced lung injury) may be extreme, such that they are unrepresentative of the range of conditions present in humans with ARDS. Conversely, such models as intratracheal endotoxin54 may allow the investigation of certain aspects of the resolution of inflammation and fibrotic processes that can occur following clinical ARDS, but they do not produce the substantial hypoxemia that is a prerequisite for a diagnosis of the syndrome51. To characterize their specific effects better, therapeutics should likely be tested in multiple models, as none sufficiently reproduces the heterogeneity of ARDS55.
Our model has some inherent limitations. First, because we euthanized the animals 4 h after experimental ARDS onset, we only collected parameters during the early phase of ARDS. More elaborate facilities, such as &#8220;animal ICUS,&#8221; are needed to investigate later phases of ARDS in piglets. Second, during the current experiments, we only assessed the degree of lung injury using an index of arterial oxygenation, such as PaO2/FiO2. However, most features of experimental ARDS were present when we previously reported use of this acid-induced ARDS model37. To improve our model, it could be interesting to add non-invasive measures of the degree of lung injury, determined, for example, using electrical impedance tomography or lung ultrasound56. Third, we only report the use of a &#8220;one-hit model&#8221; to induce lung injury in piglets, while models in which more than one inciting stimulus for lung injury is induced are likely more reflective of the pathological human situation, in which a single inciting stimulus is rarely present (&#8220;two-hit hypothesis&#8221;)51. From this perspective, ventilator-induced lung injury could, for example, be added to our model to produce an additional hit, and this model can be combined with other injurious &#8220;hits&#8221; if needed to investigate more complex clinical scenarios involving multiple features of the ARDS pathophysiology, such as lung endothelial injury, alveolar macrophage activation, and the effects of cell-free hemoglobin57, among others58. Fourth, we did not test other halogenated agents such as desflurane. Indeed, we only used isoflurane and sevoflurane for inhaled sedation in piglets because they are most frequently used in clinical ICU practice, at least in Europe and in some other areas of the world.
The porcine model has contributed to significant advances in experimental research over the last decades, and it has become an increasingly important translational bridge between traditional small laboratory animal models and human medicine. A major advantage of using experimental models in large animals is to allow investigations that involve the ventilation of animals over time. Nevertheless, such models can be extremely expensive, and they may sometimes require the availability of an animal ICU. In addition, the limited availability of some molecular reagents in pigs is an important limitation. Studies in smaller animals, such as mice, rats, or rabbits, have been highly useful in studying individual pathways, but the generalizability of the results to humans appears limited59. Larger animal studies can provide focused evaluations of key physiological and molecular pathways and can be used to test new therapies in humans, such as sedation with halogenated agents. Furthermore, the size of the animals supports the use of clinically used catheters, endotracheal tubes, ventilators, and monitors that are not fully available for use in smaller mammals. Indeed, the important advantages of using experimental models with large animals include the ability to take multiple longitudinal blood samples and perform blood gas analyses over time. In addition, invasive hemodynamic monitoring could be used as transpulmonary thermodilution with a pulse contour cardiac output monitor device, allowing the study of the degree of alveolar edema by measuring extravascular lung water, a highly relevant parameter in the alteration of the alveolar&#8211;capillary barrier during ARDS51. Nevertheless, caution is necessary when data aside from size are interpreted from animal models, because important anatomical, physiological, and immunological differences exist among animal species. Animal models could have anatomical differences that will impact research and the translation to humans. Indeed, many animals, such as mice or rabbits, have an incomplete mediastinum and thin visceral membranes, prohibiting, for example, the use of the contralateral pleura as a control. However, larger animals (e.g., sheep or pigs) have one pleural cavity around each lung and a thick visceral pleura resembling that of humans60.
The administration of volatile anesthetics to ICU patients has been increasingly studied in the last decade, mostly because of the development of dedicated devices based either on reflection or on a circle system. Such devices can be inserted into any mechanical ventilation circuit to administer the two agents most frequently used in the ICU setting, sevoflurane and isoflurane61. Due to their hypnotic, bronchodilator, and anticonvulsant properties, halogenated agents have been used for a long time in the ICU to manage patients with refractory status asthmaticus, status epilepticus, and complex sedation scenarios with high sedation requirements, such as burns, chronic pain, high-risk surgeries, or a history of drug abuse. Although recent international guidelines do not recommend using volatile agents for procedural pain management62, halogenated anesthetics are increasingly popular in Europe, and they are considered a feasible option for sedation in the 2015 German Guidelines63, especially if short wake-up times are needed. Potential therapeutic end-organ-protective properties via the cytoprotective and anti-inflammatory mechanisms64 of volatile anesthetics have attracted the attention of researchers and physicians. In fact, their emerging use in ICUs has paved the way to the study of their potential benefits in patients with ARDS. ARDS represents the ultimate and most severe form of lung organ dysfunction, as well as a major challenge for patients, their families, healthcare providers from various disciplines, and healthcare systems when caring for critically ill patients, especially under some exceptional circumstances, such as during the current COVID-19 pandemic65,66,67. Beyond the current efforts to find specific antiviral therapies, improving supportive care and treatment options for patients with COVID-19-related ARDS is, therefore, of major importance65,68,69. From this perspective, the rationale supporting inhaled sedation with sevoflurane or isoflurane as a way to improve lung epithelial permeability, to decrease the inflammatory response, and, potentially, to improve patient outcomes is strong. In addition, several non-human models have shown that a volatile anesthetic agent, such as inhaled sevoflurane, improves gas exchange21,70,71, reduces alveolar edema22, and decreases levels of pro-inflammatory cytokines72,73. These effects could be explained by restored lung epithelial function and by the immunomodulatory effects of the halogenated agent. In a previous pilot randomized control trial, the use of a halogenated agent, such as sevoflurane, to sedate ARDS patients in the ICU improved oxygenation and decreased levels of a marker of lung epithelial injury and of some pro-inflammatory cytokines (interleukin [IL]-1&#946;, IL-6, and IL-8 and tumor necrosis factor-&#945;) compared to intravenous sedation28. These results reinforce the protective effect of sevoflurane on inflammation and on reduced epithelial injury or improved AFC, as assessed by plasma sRAGE34.
Understanding the biological mechanisms and pathophysiological pathways involved in acute lung injury and its resolution under inhaled sedation with halogenated agents requires the use of experimental and preclinical models. Although in vitro studies represent an important step in describing these mechanisms74, in vivo experiments are fundamental before results can be extrapolated to the clinical setting. Furthermore, in this large animal model, halogenated agents could be administered using the same anesthetic conserving device as in humans. In fact, devices based either on reflection or on a circle system, which are both available for patients in some countries, do not have specific equivalents available for small animals, such as for mice, rats, or rabbits. Consequently, when researchers want to administer halogenated agents to animals, they must choose between either pre- or post-exposure to halogenated agents, usually via anesthesia chamber induction over a more or less long time with no specific mechanical ventilation during this period75. This piglet model allows the specific reproduction of the same treatment conditions as in ICU patients with ARDS, that is, the administration of halogenated agents, such as sevoflurane, in addition to delivering lung-protective mechanical ventilation with low tidal volumes and PEEP. Interestingly, our model reported the use of the recent, miniaturized version of the anesthetic conserving device to administer sevoflurane for the first time in piglets, thereby allowing smaller tidal volumes and further instrumental dead space to be set compared to the previous version of the device. Furthermore, in addition to administering halogenated volatiles, this model of acid-induced ARDS could be useful in studying specific pathways, such as those involved in lung epithelial injury and its repair37.
In conclusion, this experimental model of ARDS in piglets has significant advantages compared to the existing ones. These include rapid onset (within 1 h in general), good reproducibility and stability over time, a low mortality rate, and, more importantly, the use of a clinically relevant device to deliver inhaled ICU sedation, thus allowing novel translational approaches to the study of the effects of halogenated agents in ARDS.

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></td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Tracheal intubation&lt;/strong&gt;</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>&lt;strong&gt;Animal preparation&lt;/strong&gt;</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>&lt;strong&gt;Monitoring&lt;/strong&gt;</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&amp;reg; 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>&lt;strong&gt;Mechanical ventilation&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Ventilator Engstr&amp;ouml;m Carestation</td><td>General Electrics</td><td>Engstr&amp;ouml;m</td><td></td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Halogenated anesthetics&lt;/strong&gt;</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>&lt;strong&gt;Products&lt;/strong&gt;</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>Cisatracurium</td><td>Mylan</td><td>69252651</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). A significant between-group difference was noted in the undiluted pulmonary edema fluid levels of the total protein measured after 4 h of mechanical ventilation (P &#60; 10&#8722;4). HCl-induced ARDS was associated with increased BAL protein when compared to the sham animals (Figure 4). A two-way RM-ANOVA indicated a significant time by group interaction (P &#60; 10&#8722;4) with HCl-induced ARDS being associated with increased extravascular lung water, compared to sham animals without ARDS (Figure 5A). Cardiac output and systemic vascular resistance values are reported in Figure 5B and Figure 5C, respectively. Inspiratory and expiratory fractions of sevoflurane measured in all animals are reported in Figure 6, and macroscopic evidence of histologic lung injury is shown in Figure 7.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61644/61644fig01.jpg" /> 
Figure 1: Illustration of the set up needed to administer sedation with halogenated volatile agents using the anesthetic conserving device.&#160;(A) Filling the specific syringe with the bottle adapter. (B) Scavenging the halogenated agents using the scavenging charcoal filter. (C) Assembling both the syringe pump and the gas analyzer with the anesthetic conserving device to use along with the ventilator. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61644/61644fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61644/61644fig02.jpg" /> 
Figure 2: Schematic representation of the connection of the anesthetic conserving device to the respiratory circuit of the ventilator.&#160;This includes the integration of the module to measure lung functional residual capacity. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61644/61644fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61644/61644fig03.jpg" /> 
Figure 3: Measurement of the alteration in arterial oxygenation.&#160;(A) Evolution of arterial oxygen tension (PaO2) to inspired oxygen fraction (FiO2) ratio in untreated piglets (SHAM group, N = 12) and acid-injured piglets (HCl group, N = 13) during a 4 h period. (B) Evolution of the delta of PaO2/FiO2 at a specific time point and of PaO2/FiO2 at H0 in untreated piglets (SHAM group, N = 12) and acid-injured piglets (HCl group, N = 13). Values are expressed in mmHg and represented as means, with error bars representing standard errors of the means. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61644/61644fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61644/61644fig04.jpg" /> 
Figure 4: Measurements of the alteration in alveolar-capillary membrane permeability.&#160;Mini bronchoalveolar levels (BAL) of total protein at 4 h in untreated piglets (SHAM group, N = 12) and acid-injured piglets (HCl group, N = 13). Values are expressed in g.L-1 and represented as means, with error bars representing standard errors of the means. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61644/61644fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61644/61644fig05.jpg" /> 
Figure 5: Measurements provided by transpulmonary thermodilution.&#160;(A) Lung edema, as assessed by extravascular lung water. (B) Cardiac output. (C) Systemic vascular resistance. Transpulmonary thermodilution was performed in untreated piglets (SHAM group, N = 12) and acid-injured piglets (HCl group, N = 13) using a pulse contour cardiac output monitor. Values are expressed in mL.kg-1, L.min-1, dynes.s.cm-5, respectively, and are reported as means, with error bars representing standard errors of the means. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61644/61644fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61644/61644fig06.jpg" /> 
Figure 6: Measurements of the expired fractions of halogenated agents, sevoflurane and isoflurane.&#160;(A) Expired (FEsevoflurane) and inspired (FIsevoflurane) sevoflurane fractions during the 4-h study period. (B) Expired (FEisoflurane) and inspired (FIisoflurane) isoflurane fractions during the 4-h study period. Values are expressed in % and represented as means, with error bars representing standard errors of the means. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61644/61644fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/61644/61644fig07.jpg" /> 
Figure 7: Macroscopic evaluation of the whole lung after the 4 h study period.&#160;(A) Whole lung of an untreated piglet (SHAM group). (B) Whole lung of an acid-injured piglet (HCl group). Macroscopic lung injury, with visible hemorrhage and congestion, is noticeable in the red parts of the lung (white arrows). Histologic evaluation of the lung after the 4 h study period. (C) Histological slice of the lung of an untreated piglet (SHAM group). (D) Histological slice of the lung of an acid-injured piglet (HCl group). Histologic evidence of lung injury was a greater cellularity consisting mainly of neutrophils (black arrowheads), with more areas of atelectasis and increased alveolar disruption, hyaline membranes, protein debris, hemorrhage (white arrow), and the thickening of the alveolar wall (black arrows). Scale bars equal 100 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61644/61644fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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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2.7K Views.  CHU Clermont-Ferrand. We beschrijven een model van zoutzuur-geïnduceerd acute respiratory distress syndrome (ARDS) bij biggen die sedatie krijgen met gehalogeneerde middelen, isofluraan en sevofluraan, via een apparaat dat wordt gebruikt voor geïnhaleerde intensive care-sedatie. Dit model kan worden gebruikt om de biologische mechanismen van gehalogeneerde middelen op longletsel en -herstel te onderzoeken.

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 ...

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

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

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

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

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

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 ...

Endotracheal Intubation Using a Flexible Intubation Endoscope as a Standardized Model for Safe Airway Management in Swine

Endotracheal intubation is often a basic requirement for translational research in porcine models for various interventions that require a secured airway or high ventilation pressures. Endotracheal intubation is a challenging skill,&#160;requiring a minimum number of successful endotracheal intubations to achieve a high success rate under optimal conditions, which is often unachievable for non-anaesthesiology researchers. Due to the specific porcine airway anatomy, a difficult airway can usually be assumed. The impossibility of establishing a secure airway can result in injury, adverse events, or death of the laboratory animal. Using a prospective, randomized, controlled evaluation approach, it has been shown that fiberoptic-assisted endotracheal intubation takes longer but has a higher first-pass success rate than conventional intubation without causing clinically relevant drops in oxygen saturation. This model presents a standardized regimen for endoscopically guided endotracheal intubation, providing a secured airway, especially for researchers who are inexperienced in the technique of endotracheal intubation via direct laryngoscopy. This procedure is expected to minimize animal suffering and unnecessary animal losses.

The use of pigs in research has increased in recent years. Nevertheless, pigs are characterized by difficult airway anatomy. By demonstrating how to perform endoscopically guided endotracheal intubation, the present protocol aims to further increase laboratory animals' safety to avoid animal suffering and unnecessary death.

Endotracheal intubation is often a basic requirement for translational research in porcine models for various interventions that require a secured airway ...

Delivery of Cardioactive Therapeutics in a Porcine Myocardial Infarction Model

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

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

Myocardial infarction is one of the leading causes of death and disability worldwide, and there is an urgent need for novel cardioprotective or ...

Description of a Swine Infant Model of Volume-Controlled Hemorrhagic Shock

Hemorrhagic shock is a leading cause of morbidity and mortality in pediatric patients. Interpretation of the clinical indicators validated in adults to guide resuscitation and comparison between different therapies is difficult in children due to the inherent heterogeneity of this population. As a result, compared to adults, appropriate management of pediatric hemorrhagic shock is still not well established. In addition, the scarcity of pediatric patients with hemorrhagic shock precludes the development of clinically relevant studies. For this reason, an experimental pediatric animal model is necessary to study the effects of hemorrhage in children as well as their response to different therapies. We present an infant animal model of volume-controlled hemorrhagic shock in anesthetized young pigs. Hemorrhage is induced by withdrawing a previously calculated blood volume, and the pig is subsequently monitored and resuscitated with different therapies. Here, we describe a precise and highly reproducible model of hemorrhagic shock in immature swine. The model yields hemodynamic data that characterizes compensatory mechanisms that are activated in response to severe hemorrhage.

This article aims to provide researchers with a detailed and accessible guide to set up an infant porcine model of hemorrhagic shock.

Hemorrhagic shock is a leading cause of morbidity and mortality in pediatric patients. Interpretation of the clinical indicators validated in adults to ...

Dosage-Dependent Induction of Endotoxemia in Pigs via LPS Infusion

Lipopolysaccharide Infusion as a Porcine Endotoxemic Shock Model

Sepsis and septic shock are frequently encountered in patients treated in intensive care units (ICUs) and are among the leading causes of death in these patients. It is caused by a dysregulated immune response to an infection. Even with optimized treatment, mortality rates remain high, which makes further insights into the pathophysiology and new treatment options necessary. Lipopolysaccharide (LPS) is a component of the cell membrane of gram-negative bacteria, which are often responsible for infections causing sepsis and septic shock. 
The severity and high mortality of sepsis and septic shock make standardized experimental studies in humans impossible. Thus, an animal model is needed for further studies. The pig is especially well suited for this purpose as it closely resembles humans in anatomy, physiology, and size. 
This protocol provides an experimental model for endotoxemic shock in pigs by LPS infusion. We were able to reliably induce changes frequently observed in septic shock patients, including hemodynamic instability, respiratory failure, and acidosis. This will allow researchers to gain valuable insight into this highly relevant condition and evaluate new therapeutic approaches in an experimental setting.

Author Spotlight: Induction of Experimental Endotoxemic Shock in Pigs for Studying Hemodynamic and Respiratory Failure

We provide a protocol for an experimental endotoxemic shock model in pigs by infusion of lipopolysaccharide.

Sepsis and septic shock are frequently encountered in patients treated in intensive care units (ICUs) and are among the leading causes of death in these ...

Investigating Physiological Responses to Acute Volume Overload in Porcine Models

Adult and Pediatric Porcine Model of Acute Volume Overload

This protocol describes an acute volume overload porcine model for adult Yorkshire pigs and piglets. Both swine ages undergo general anesthesia, endotracheal intubation, and mechanical ventilation. A central venous catheter and an arterial catheter are placed via surgical cutdown in the external jugular vein and carotid artery, respectively. A pulmonary artery catheter is placed through an introducer sheath of the central venous catheter. PlasmaLyte crystalloid solution is then administered at a rate of 100 mL/min in adult pigs and at 20 mL/kg boluses over 10 min in piglets. Hypervolemia is achieved either at 15% decrease in cardiac output or at 5 L in adult pigs and at 500 mL in piglets. Hemodynamic data, such as heart rate, respiratory rate, end-tidal carbon dioxide, fraction of oxygen-saturated hemoglobin, arterial blood pressure, central venous pressure, pulmonary artery pressure, pulmonary capillary wedge pressure, partial arterial oxygen pressure, lactate, pH, base excess, and pulmonary artery fraction of oxygen-saturated hemoglobin, are monitored during experimentation. Preliminary data observed with this model has demonstrated statistically significant changes and strong linear regressions between central hemodynamic parameters and acute volume overload in adult pigs. Only pulmonary capillary wedge pressure demonstrated both a linear regression and a statistical significance to acute volume overload in piglets. These models can aid scientists in the discovery of age-appropriate therapeutic and monitoring strategies to understand and prevent acute volume overload.

Author Spotlight: Exploring Venous Waveforms in Porcine Models to Tackle Volume Overload in Medicine

The protocol here shows how continuous administration of crystalloids into the central veins of a euvolemic pig/piglet allows for the appropriate investigation of the physiological effects of acute volume overload.

This protocol describes an acute volume overload porcine model for adult Yorkshire pigs and piglets. Both swine ages undergo general anesthesia, ...

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

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

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

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

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

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

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