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

Summary

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

Abstract

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 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.

Introduction

The utilization of porcine models in translational research holds significant importance in advancing our understanding of human medicine¹. Porcine models, due to their physiological and anatomical similarities to humans, provide a valuable platform for studying complex diseases and therapeutic interventions¹. In the context of acute respiratory failure, porcine models offer a unique opportunity to investigate the pathophysiological mechanisms, evaluate treatment strategies, and assess potential interventions^1,2,3. The ability to replicate key aspects of human respiratory physiology and responses to various stimuli in porcine models allows for a comprehensive evaluation of therapeutic modalities before progressing to human trials^1,2,3. This research paradigm enables researchers to bridge the gap between preclinical investigations and clinical application, facilitating the development of novel therapies and improving patient outcomes¹. Therefore, the establishment of an efficient, effective, and reproducible acute respiratory failure porcine model serves as a crucial tool in advancing the knowledge of respiratory diseases and guiding the development of effective interventions in human medicine¹.

Respiratory distress, a critical medical condition, has witnessed limited advancements in its diagnosis and management over the past three decades⁴. The currently employed evaluation and triage metrics, which include subjective symptoms, physical examination findings, SpO₂, and respiratory rate, often exhibit limitations in detecting acute pulmonary conditions at an early stage^5,6,7. This inadequacy not only hampers efficient triage and resource allocation but also fails to provide effective, quantitative monitoring of disease progression and treatment response in patients with chronic pulmonary diseases. The emerging landscape of chronic respiratory conditions, such as long COVID, along with the burden of acute respiratory insufficiencies on hospital resources, underscores the urgent need to expand translational research and foster innovation in respiratory disease management.

The direct infusion of oleic acid into a pig's bloodstream has been recognized as a robust method to induce acute respiratory distress⁸. Oleic acid, a monounsaturated fatty acid, has demonstrated the ability to trigger significant pulmonary injury and compromise respiratory function when introduced into the pulmonary circulation⁸. Upon infusion, oleic acid provokes vasoconstriction, resulting in increased pulmonary arterial pressure and resistance, leading to impaired gas exchange and oxygenation⁹. Furthermore, oleic acid promotes the activation of inflammatory pathways, including the release of pro-inflammatory mediators and recruitment of immune cells, which contribute to the development of lung injury and respiratory distress¹⁰. All of this results in severe hypoxemia, increases in pulmonary arterial pressures, and the accumulation of extravascular lung water¹¹. Histological evaluation of the lung parenchyma has demonstrated injury that is indistinguishable from human acute respiratory distress⁹.

This article details a method involving the direct administration of oleic acid into the pulmonary artery to induce acute respiratory distress, avoiding untreatable, severe hemodynamic compromise. The described method is anticipated to be a valuable tool for future researchers exploring the underlying pathophysiological mechanisms of acute respiratory failure and assessing potential therapeutic interventions and innovations.

Protocol

The protocol received approval from the Vanderbilt University Institutional Animal Care and Use Committee (protocol M1800176-00) and strictly adhered to the National Institute of Health Guidelines for the Care and Use of Laboratory Animals. Male and female Yorkshire pigs, weighing approximately 40-45 kg, were utilized in this experiment. The animals were obtained from a commercial source (see Table of Materials). The current practice does not involve screening for any pre-existing medical conditions in the acquired swine. While it is acknowledged that this practice could potentially interfere with or mask intended results, it is considered unlikely according to the vendor, and this limitation is accepted.

1. Animal preparation

1. Anesthesia and ventilation
   1. Anesthetize the pig with intramuscular injection of ketamine (2.2 mg/kg) / xylazine (2.2 mg/kg) / telazol (4.4 mg/kg) and position the animal in a supine (lying down) posture on the operating table.
   2. Maintain general anesthesia by initiating inhalational anesthetic, 1% isoflurane.
   3. Expose the vocal cords through the mouth using a laryngoscope and intubate¹² with a 6.5 mm endotracheal tube (see Table of Materials). Inflate the tube cuff with 3-5 mL of air using a syringe without a needle attached.
      NOTE: Immediately perform carbon dioxide (CO₂) capnography post-tracheal cannulation to ensure proper tube placement, and measure CO₂ with ventilation indicating appropriate ventilation.
   4. Use an 18 G to 24 G intravenous (IV) catheter placed in the central or marginal ear vein on the posterior side of the auricle to administer intraoperative (as needed) and euthanasia drugs to the pig.  Secure the IV catheter with 1-inch adhesive tape.
   5. Set mechanical ventilation to volume-controlled ventilation settings with a tidal volume of 8 mL/kg.
      NOTE: Anesthesia monitoring is conducted throughout the experiment. Vital signs, response to stimulus, presence/absence of movement, jaw tone laxity, changes in heart rate, end-tidal CO₂, and respiratory rate variation are monitored by an independent animal lab technician. Adjustments to the inhaled anesthetic dose are made based on these assessments. Analgesic administration of buprenorphine via bolus is performed. Adjust the respiratory rate on the mechanical ventilator to maintain an end-tidal CO₂ of 35-40 mmHg throughout the experiment.
2. Cannulation and monitoring
   1. Disinfect the entire anterior neck with a 2% chlorhexidine scrub solution, followed by a spray of 5% providone-iodine solution.
   2. Surgically expose both the right and left internal and external jugular (IJ and EJ) veins and carotid arteries (CA) with a vertical incision, approximately 7-8 cm, immediately lateral to the trachea on either side using a No. 23 blade to the sternum¹³.
      NOTE: A surgical cut-down approach is chosen for vascular access due to the challenges of a percutaneous, ultrasound-guided Seldinger technique¹⁴ in swine. The tough skin and vascular size make a cut-down approach more feasible. Cervical vessels are preferred for bilateral pulmonary artery catheters (PAC), though femoral access is an option. The choice which jugular vein (IJ or EJ) is at the discretion of procedurelist. Which ever has the larger diameter can be cannulated and used for heart catheterization.
   3. Dissect the strap muscles and tract as needed using Kelly tissue scissors and Lahey retractors¹³ (see Table of Materials).
   4. After exposure, place two 8.5 Fr cannulas and two PACs using the Seldinger technique¹⁴.
      NOTE: Right jugular vein catheter and PAC are dedicated for volume administration and hemodynamic monitoring. Left jugular vein catheter and corresponding PAC are used for oleic acid administration. Dual-PAC placement is confirmed using fluoroscopy.
   5. Using the Seldinger technique, place an arterial line in the right CA for invasive blood pressure monitoring throughout the experiment.
   6. Attach all desired monitoring equipments. Monitor heart rate (HR) with telemetry leads. Monitor systolic blood pressure (SBP), diastolic blood pressure (DBP), and mean arterial pressure (MAP) by connecting a pressure transducer to the CA catheter. Monitor mean pulmonary artery pressure (MPAP) and central venous pressure (CVP) using an independent pressure transducer/amplifier system set-up.
      ​NOTE: Calculate pulse pressure as the difference between SBP and DBP, and pulse pressure variability (PPV) by calculating the difference between peak pulse pressure at inspiration and expiration during the respiratory cycle. Perform thermodilution cardiac output (CO) using device-specific volume temperature/volume calibration. To obtain a pulmonary capillary wedge pressure (PCWP), inflate the PAC balloon with 1.5 mL of air, advance the catheter until visualization of both "v" and "a" waves, representing restricted right-to-left blood flow, and record the PCWP as the pressure value of the "a" wave at end expiration¹⁶.
   7. To monitor urine output, place a Foley catheter (see Table of Materials) in the pig's urethra. For male pigs, surgical suprapubic catheterization is required¹⁷.
   8. Administer crystalloids (PlasmaLyte, see Table of Materials) at a rate of 100 mL over 10 min to achieve a PCWP of 8-12 mmHg (euvolemia) before initiating oleic acid. Check PCWP every 100 mL until 10 mmHg is achieved.

2. Oleic acid infusion

1. Oleic acid preparation
   1. Prepare the oleic acid solution (16%) by combining 16 mL of oleic acid with 84 mL of normal saline.
      NOTE: Handle oleic acid with care, ensuring personnel wear protective gloves, safety goggles, and masks to prevent direct skin contact, inhalation, or eye exposure. Frequent agitation is necessary to prevent separation, and dimethyl sulfoxide may be added if separation occurs.
   2. Prime the oleic acid solution into an IV fluid line and connect it to the distal port of the left jugular pulmonary artery catheter (see Table of Materials).
2. Oleic acid initiation
   1. Start the oleic acid infusion at a rate of 0.2 mL/kg/h, marking the official start time^18,19,20(Figure 1).
      NOTE: Confirm hemodynamic monitoring and catheter placement before starting the oleic acid infusion. Techniques such as fluoroscopy or transthoracic echocardiography can ensure proper placement^21,22.
   2. Immediately after starting oleic acid, set ventilator settings to mimic room air conditions (FiO₂ = 21%, PEEP = 0 cm H₂O).
3. Hemodynamic and respiratory monitoring during oleic acid infusion
   1. Continuously monitor heart rate (HR), fraction of oxygen-saturated hemoglobin (SpO₂), respiratory rate (RR), end-tidal carbon dioxide (ETCO₂), central venous pressure (CVP), systolic blood pressure (SBP), diastolic blood pressure (DBP), mean arterial pressure (MAP), pulse pressure variability (PPV), and mean pulmonary artery pressure (MPAP).
      1. Measure partial arterial oxygen pressure (PaO₂), pH, lactate, base excess, cardiac output (CO), and pulmonary capillary wedge pressure (PCWP) every 30 min over the first 60-90 min and then every 15 min thereafter until sacrifice (Figure 1). Record peak airway pressure and plateau pressure at the same time PaO₂ is measured.
         NOTE: Typical starting values for continuously monitored variables are 95%-100%, 15-20 breaths per minute, 25-35 mmHg, 70-80 mmHg, 40-50 mmHg, 55-65 mmHg, 1%-4%, and 10-20 mmHg, respectively. Starting values for intermittent variables are 10 mmHg, 7.4, 0-2 mg/dL, -2 +2 mEq/L, >5 L/min, and 8-10 mmHg, respectively.
   2. Consider the experiment complete once arterial PaO₂/fraction of inspired O₂ (P/F) is less than <100²³.
      ​NOTE: At this point, animal can be euthanized (see below), and pulmonary pathology samples can be obtained, if needed (Figure 2).

3. Venous waveform analysis and ventilator management procedure

1. Respiratory non-Invasive Venous waveform Analysis
    - Respiratory Index (RIVA-RI)
   NOTE: Our research team employs this model to investigate changes in venous waveforms during respiratory distress. Peripheral venous waveforms are noninvasively captured at the upper arm of a pig using a piezoelectric sensor (see Table of Materials). Signal processing and amplification are required for analysis of these low-amplitude waveforms. Fourier transformation is then applied to present the data in the frequency domain, revealing a low-amplitude waveform at approximately 0.2 Hz (termed "f_R0") corresponding to respiration. This hypothesis suggests that this wave results from retrograde propagation of negative intrathoracic pressure during inspiration from the right atrium/vena cava throughout the venous system. The weighted contributions of the amplitudes of the respiratory signal (f_R0) can be ratiometrically normalized to compare data on a common scale and improve performance, and to the amplitude of the frequency of the pulse rate (f₀) to produce a RIVA-RI⁷.
   1. Place the piezoelectrode on the anterior upper extremity venous plexus immediately proximal to the elbow.
      NOTE: Ensure recording and uploading abilities with the piezoelectrode device. Examples of previous recording prototypes can be found in referenced literature^{7,24,25,26,27}.
   2. Start recording venous waveforms with LabChart software (see Table of Materials) whenever venous waveforms are desired during experimentation.
2. Euthanasia
   1. Confirm the maintenance of isoflurane at 1%.
   2. Induce cardiac arrest by IV injection of sodium pentobarbital (125 mg/kg) (following institutionally approved protocols).
   3. Confirm the lack of vitals post-injection to verify demise.

Results

Early single pig, pilot data demonstrates an increase in RIVA-RI prior to changes in other respiratory monitoring measures (RR and SpO₂), in line with changes in PaO₂ (Figure 3). The drop in PaO₂ is the "positive" result this model intends to achieve. Preliminary data also shows that RIVA-RI increases and the PaO₂ decreases with disease progression starting at the 30-min mark (Figure 3; red arrow). PaO

Discussion

The key element in this protocol is to closely monitor the hemodynamic condition of the pig during the administration of oleic acid to induce respiratory distress¹⁵. It is of the utmost importance for researchers to take the necessary time to appropriately position the hemodynamic monitoring devices. One specific drawback of this model is the potential hemodynamic instability that may arise as a result of inflammation and injury to the respiratory vasculature^8,

Materials

Name	Company	Catalog Number	Comments
1% Isoflurane	Primal, Boston, MA, USA	26675-46-7	https://www.sigmaaldrich.com/US/en/product/aldrich/792632?gclid=Cj0KCQjw9fqnBhDSARIsAHl cQYS_W-q6tS2s6LQw2Qn7Roa3T GIpTLPf52351vrhgp44foEcRozPqt YaAtvfEALw_wcB
Arterial Catheter	Merit Medical, South Jordan, UT, USA	MAK401	MAK Mini Access Kit 4F
Blood Pressure Amp	AD Instruments, Colorado Springs, CO, USA	FE117	https://www.adinstruments.com/products/bp-blood-pressure-amp
Central Venous Catheter	Arrow International, Cleveland, OH, USA	AK-09800	8.5 Fr. x 4" (10 cm) Arrow-Flex
Disposable Pressure Transducers	AD Instruments, Colorado Springs, CO, USA	MLT0670	https://www.adinstruments.com/products/disposable-bp-transducers
Edwards Lifesciences Triple Stage Venous Cannulas	Edwards Life Sciences, Irvine, CA	TF293702	https://www.graylinemedical.com/products/edwards-lifesciences-triple-stage-venous-cannulas-venous-dual-stage-cannula-tf293702?variant=31851942576185&gad=1& gclid=Cj0KCQiAr8eqBhD3ARIsAIe -buNdmkzavUBaIx-1be7boWn2kW hbUR6QCjaobB08uuK9qJW66JvY TM4aAufGEALw_wcB
Kelly Scissors	MPM Medical Supply, Freehold, NJ 07728	104-5516	https://www.mpmmedicalsupply.com/products/kelly-scissors
Kendall 930 FoamElectrodes	Covidien, Mansfield, MA, USA	22935	https://www.cardinalhealth.com/en/product-solutions/medical/patient-monitoring/electrocardiography/monitoring-ecg-electrodes/radiolucent-electrodes/kendall-930-series-radiolucent-foam-electrodes.html
Ketamine Hydrochloride 100 mg/mL, Injectable Solution, 10 mL	Patterson Veterinary, Loveland, CO 80538	07-894-8462	https://www.pattersonvet.com/ProductItem/078948462?omni=ketamine
LabChart 8 software	AD Instruments, Colorado Springs, CO, USA	N/A	https://www.adinstruments.com/products/labchart
Lahey Retractor	BOSS Instruments LTD, Gordonsville, VA 22942	18-1210	https://bossinstruments.com/product/7-3-4-lahey-thyroid-retractor-6mmx28mm/
Oleic Acid	Sigma-Aldrich, Merck, Darmstadt, Germany	O1008	https://www.sigmaaldrich.com/US/en/product/sial/o1008?gclid=CjwKCAjwzJmlBhBBEiwAEJy Lu2047wRpXqF_Z2BegUyhgZJ _WygsWfErhgrGCIyMp8PxwNH sTZ8qARoCl1QQAvD_BwE&gcl src=aw.ds
Peripheral IV Catheter Angiocath 18-24 G 1.16 inch	McKesson, Irving, TX, USA	329830	https://mms.mckesson.com/product/329830/Becton-Dickinson-381144
Piezoelectrode	MuRata Manuractoring Co, Ltd., Nagaokakyo, Kyoto, Japan	7BB-12-9	https://www.murata.com/en-us/products/productdetail?partno=7BB-12-9
PlasmaLyte	Baxter International, Deerfield, IL, USA	2B2544X	https://www.ciamedical.com/baxter-2b2544x-each-solution-plasma-lyte-a-inj-ph-7-4-1000ml
Pulmonary Artery Catheter	Edwards Life Sciences, Irvine, CA	131F7	Swan Ganz 7F x 110cm
Standard Endotracheal Tubes	Teleflex, Morrisville, NC 27560	5-10313	https://www.teleflex.com/usa/en/product-areas/anesthesia/airway-management/endotracheal-tubes/standard-tubes/index.html
SurgiVet Clearview Foley Catheter, 8 Fr, 55 cm Silicone	Penn Veterinary Supply, Inc, West Rendering, PN 13971	SVCFC1030	https://www.pennvet.com/customer/portal/catalog/home?urile=wcm:path%3APennVet+Catalog/Super+Sku+Catalog/SS0672/Surgivet+Clearview+Silicone+Foley+Catheters
Telazol (Tiletamine HCl and Zolazepam HCl), Injectable Solution, 5 mL	Patterson Veterinary, Loveland, CO 80538	07-801-4969	https://www.pattersonvet.com/ProductItem/078014969?omni=telazol
Welch Allyn E-MacIntosh Standard Laryngoscope Blade	MFIMedical, San Diego, CA 92131	WLA-69242	https://mfimedical.com/products/welch-allyn-e-macintosh-standard-laryngoscope-blade?variant=12965771870285&currency =USD&utm_medium=product_sync &utm_source=google&utm_content =sag_organic&utm_campaign=sag _organic&gclid=Cj0KCQiAr8eqBhD 3ARIsAIe-buMhpgM96qRXkCUKA 6Mhmdat0p93JbecCGTaLStexhV pkUVa9VkWUzgaAr-iEALw_wcB
Xylazine HCl 100 mg/mL, Injectable Solution, 50 mL	Patterson Veterinary, Loveland, CO 80538	07-894-5244	https://www.pattersonvet.com/ProductItem/078945244
Yorkshire Pigs	Oak Hill Genetics, Ewing, IL, USA	138274	Female/Male Swine- Yorkshire/Landrace 81-100lbs