This study has been funded by the Instituto de Salud Carlos III (ISCIII) through the project&#160;&#34;PI20/01706&#34; and co-founded by the European Union. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. We would like to show out gratitude to all our colleagues from Gregorio Mara&#241;&#243;n Pediatric Intensive Care Unit and from the Gregorio Mara&#241;&#243;n Experimental Institute, as without their work this project would not have been possible.

The experiments in this protocol were approved by the Institutional Ethics Committee for Animal Research of the Gregorio Mara&#241;&#243;n University Hospital, Madrid, Spain, and the Agriculture and the Environment Council of the Madrid Autonomous Government (permit number: 12/0013). European and Spanish guidelines for ethical care and the use of experimental animals were applied throughout the study. The experiments were performed in the Department of Experimental Medicine and Surgery, Gregorio Mara&#241;&#243;n University Hospital, Madrid, Spain.
NOTE: The animal model chosen consisted of healthy 2-3-month-old (8-12 kg) minipigs (Sus scrofa domestica). Minipigs are the result of a crossing of three different breeds that make them suitable for biomedical research. The animals are near-identical lines and are provided by a specifically authorized breeding facility in Madrid (IMIDRA), which keeps the maintenance of three homozygous genetic lines in purity. Male and female animals were used interchangeably. Animals were fed a standard swine diet and observed for a minimum of 2 days to ensure good health. Food, but not water, was withdrawn the night before the procedures to reduce the risk of aspiration. A typical experiment requires approximately 6 h to complete, including 30 min for anesthesia induction and surgical preparation, 60 min for instrumentation, 30 min for recovery, 60 min for hemorrhage induction and posterior stabilization, 30 min for resuscitation, and 120 min for follow-up.
1. Anesthesia, intubation, and mechanical ventilation
<ol>
	<li>Premedicate the pig with an intramuscular injection of ketamine (10 mg/kg) and atropine (0.02 mg/kg) in the lateral region of the neck, behind the ear, or in the posterior femoral region. 
	NOTE: Anticholinergic drugs, such as atropine, are useful as swine may salivate excessively under anesthesia10.&#160;In our experience, this dose of ketamine is enough to reduce stress and induces adequate sedation and analgesia in pigs without adverse effects. However, if the animal is not properly sedated or if the distance from the housing to the operating room is long, another dose of ketamine (10 mg/kg) can be administered safely. 
	CAUTION: Gloves are necessary when handling animals.</li>
	<li>Transport the sedated animal to the operating room and place it on a surgical table provided with a warming blanket.</li>
	<li>Measure peripheral oxygen saturation (Sp02) with a sensor clipped to the pig&#39;s ear and initiate continuous three-lead electrocardiographic (EKG) monitoring.</li>
	<li>Disinfect the skin with at least 3 alternating rounds of povidone-iodine or chlorhexidine scrub and alcohol. Insert a peripheral vein catheter (22-24 G) into the ear vein. Disinfect the skin priorly with an antiseptic solution.</li>
	<li>Induce anesthesia by an intravenous injection of fentanyl (5 &#181;g/kg), propofol (4 mg/kg), and atracurium (0.5 mg/kg). Once spontaneous breathing disappears and the absence of reflexes is confirmed, place the animal in the dorsal recumbency position and immediately initiate hand-bag mask ventilation with a dog mask with fraction of inspired oxygen (Fi02) set at 100%. 
	NOTE: In order to reduce the risk of accidental awareness related to the use of neuromuscular blockers,&#160;anesthetic agents with known effectivity in swine and with doses on the higher limit should be used to ensure an adequate level of anesthesia. Additionally,&#160;continuously monitor cardiovascular signs such as heart rate, blood pressure, and body temperature, and administer neuromuscular blockers only when withdrawal reflexes are absent (pedal withdrawal, palpebral reflexes, and jaw tone) and muscle tone is relaxed.</li>
	<li>Perform endotracheal intubation. At least two operators are needed for this procedure.
	<ol>
		<li>Ensure that the basic equipment and surgical tools needed for endotracheal intubation are ready: tie gauze to open the mouth and secure the tube, veterinary laryngoscope with a straight blade between 17 and 25 cm long, a common endotracheal tube (ID 4-5), stylet, syringe with air, and adhesive tape.</li>
		<li>Pull out the tongue slightly and hold the jaw open using tie gauze placed behind the upper and lower canine teeth.</li>
		<li>Perform a laryngoscopy and, once the epiglottis is visible, use the tip of the laryngoscope to press the epiglottis upward toward the base of the tongue. 
		NOTE: If the epiglottis is stuck to the soft palate, it can be displaced dorsally with the tip of the tube. Operator 1 performs step 1.6.2 while operator 2 performs step 1.6.3.</li>
		<li>Once the vocal cords are visualized, gently advance the tube with slight rotation into the trachea. 
		NOTE: The narrowest point of the trachea is on the subglottic level. If tube insertion is difficult, try slight rotation or a smaller tube.</li>
		<li>Remove the stylet and use a 5 mL syringe to inflate the cuff.</li>
		<li>Ensure placement of the endotracheal tube by observing symmetrical chest rise, adequate oxygen saturation (95%-100%), and a proper waveform and end-tidal CO2 (EtCO2) reading. 
		CAUTION: Pigs are very susceptible to laryngospasm and edema of the larynx mucosa, and laryngeal perforation may even occur after several attempts of intubation or if sedation is inadequate10.</li>
	</ol>
	</li>
	<li>After confirming intubation, initiate mechanical ventilation using a mechanical ventilator with a respiratory rate of 20 breaths per minute, tidal volume of 8 mL/kg, FiO2 of 40%, and positive end-expiratory pressure of 4 cm H2O. Adjust ventilation to achieve a partial pressure of carbon dioxide (PaCO2) between 35 and 45 mmHg.</li>
	<li>Maintain deep anesthesia throughout the experiment via a continuous infusion of fentanyl (10 &#181;g/kg/h), propofol (10 mg/kg/h), and atracurium (2 mg/kg/h).</li>
</ol>
2. Instrumentation
<ol>
	<li>Prepare the femoral area for vessel catheterization. Use bandages to pull the legs back and disinfect the inguinal area with at least 3 alternating rounds of povidone-iodine or chlorhexidine scrub and alcohol.</li>
	<li>Assess the femoral vessels with an ultrasound and use the Doppler technique to distinguish between the artery and the vein. Depending on the size of the vein, insert a 5.5-7.5 French (F) central venous catheter with three ports in one of the femoral veins under continuous ultrasound view and using the Seldinger technique11,12.</li>
	<li>Immediately after central venous catheter placement, connect a transducer system to measure the central venous pressure.</li>
	<li>Ensure that an electrolyte with glucose infusion (20 mL/h) is connected to one of the central line ports and that a maintenance saline infusion (5 mL/h) is infused via the remaining port to prevent occlusion of the catheter.</li>
	<li>Use the same technique to cannulate the opposite femoral artery with a 4 F arterial catheter specifically designed for cardiac output monitoring. Perform a blood gas test to establish the correct position of the catheter if ultrasound confirmation is not possible. 
	NOTE: In case of significant spasm or hematoma, cross over to the contralateral femoral artery.</li>
	<li>Once the arterial catheter is inserted, connect the arterial wire of the cardiac output monitor system and the arterial transducer directly to the monitor port. Simultaneously connect the venous measuring unit of the monitor to the central venous transducer. 
	NOTE: The cardiac output monitor used in this experiment is specified in the Table of Materials. For setup, calibration, and measures, see the manufacturer instructions13.</li>
	<li>Ensure both the venous and arterial transducers are calibrated to zero.</li>
	<li>Expose the left internal carotid artery and left external jugular vein via the cut-down technique.
	<ol>
		<li>Ensure the necessary equipment and surgical tools are available: scalpel, blunt-tip surgical scissors, tissue forceps, small self-retaining tissue retractor, needle holder, surgical swaps, suture with needle, one 18 G IV cannula, one 5 F catheter sheath with an introducer, and one Seldinger guide wire.</li>
		<li>With the animal in the dorsal recumbency position, disinfect the neck skin with an antiseptic solution.</li>
		<li>Use a scalpel to make an ~10 cm left paratracheal incision, bisecting a line between the manubrium and the angle of the jaw.</li>
		<li>To expose the external jugular vein, dissect the tissue lateral to the SCM and isolate the vein from the surrounding fascia.</li>
		<li>After being isolated, use two non-absorbable silk sutures (USP-0) looped around the vein to fixate the vessel prior to the puncture.</li>
		<li>Incise the vein with a Venflon needle (18 G). Once inside the vein, retract the needle and insert the guide wire through the Venflon tube.</li>
		<li>Remove the Venflon tube and insert the sheath with the introducer (5 F) over the wire. After insertion, remove both the introducer and the wire.</li>
		<li>Immediately after insertion, rinse the sheaths with 0.9% NaCl (5 mL/h) to prevent thrombus formation.</li>
		<li>Tie up the proximal silk suture around the sheath to fix it. After that, secure the handle of the sheath to the SCM and close the skin with staples.</li>
	</ol>
	</li>
	<li>Following surgical preparation, allow the animals to stabilize for 30 min before obtaining the baseline monitoring values and blood samples.</li>
	<li>Maintain the blood temperature at 37-39 &#730;C using a thermal blanket and an overhead warmer throughout the experiment. 
	&#8203;NOTE: Temperature is measured with a thermistor located at the thermodilution arterial catheter tip.</li>
</ol>
3. Hemodynamic and perfusion monitoring
<ol>
	<li>Monitor the EKG, peripheral oxygen saturation, respiratory volumes and pressures, and Fi02.</li>
	<li>Connect a spirometer between the endotracheal tube and a multiparameter monitor to measure qualitative and quantitative EtC02. 
	NOTE: For more information about the multiparameter monitor, see the Table of Materials.</li>
	<li>Use near-infrared spectroscopy (NIRS) to monitor the brain tissue oxygenation index (bTOI) and splanchnic tissue oxygenation index (aTOI). Place the sensors on the skin of the forehead and the anterior abdominal wall (subhepatic region). 
	NOTE: Do not place the brain sensor in the midline, as it may be contaminated with the superior sagittal sinus venous blood14.</li>
	<li>Connect the blood flow probe attached to the internal carotid artery to a flow monitor to measure the carotid blood flow (CaBF).</li>
	<li>Place a laser Doppler sensor over the skin of the anterior abdominal wall for continuous measurement of the cutaneous tissular blood flow (CuTBF). 
	NOTE: For more information about the carotid and cutaneous tissular blow flow sensors, see the Table of Materials.</li>
	<li>Record the following parameters at baseline and every 30 min: blood temperature, inspiratory tidal volume, EtCO2, cardiac rhythm, heart rate (HR), systolic and diastolic blood pressures, mean arterial blood pressure (MAP), shock index (HR/systolic blood pressure)15, central venous pressure, cardiac index (CI), global end diastolic volume index (GEDVI), stroke volume index (SVI), left ventricular contractility (Dt/Dpmax), systemic vascular resistance index (SVRI), extravascular lung water index (ELWI), pressure pulse variation (PPV), peripheral hemoglobin saturation, central venous saturation (ScvO2), cerebral (bTOI) and splanchnic (aTOI) tissue oxygenation index by NIRS, CaBF, and CuTBF.</li>
	<li>To obtain CI values, infuse 5 mL boluses of 0.9% normal saline at a temperature below 8 &#176;C through the central venous catheter. Record the average of two consecutive measures.</li>
	<li>Determine the arterial and venous blood gas profiles and lactate concentration every 30 min by drawing 0.3 mL of blood from the femoral vessels. Perform standard complete blood counts, coagulation studies, and biochemistry at baseline, after hemorrhage induction, and at the end of the experiment.</li>
	<li>After each blood draw, flush the lines with 0.5 mL of 100 IU/mL heparin.</li>
</ol>
4. Hemorrhagic shock induction
<ol>
	<li>Once a steady state is achieved after instrumentation and baseline data has been gathered, induce hypovolemic shock by withdrawing 30 mL/kg of blood from the jugular vein over 30 min.</li>
	<li>Allow a 30 min period for stabilization. Do not make resuscitation efforts during this period to emulate the delay in arrival of emergency medical teams.</li>
</ol>
5. Infusion and follow-up
<ol>
	<li>After the stabilization period, infuse a bolus of a volume expander or a vasoactive agent over a 30 min period. 
	NOTE: Examples of volume expanders and vasoactive agents tested are normal saline, hypertonic albumin, angiotensin, and terlipressin. In this study, 30 mL/kg normal saline (NS) (n = 13), 15 mL/kg 5% albumin plus 3% hypertonic saline (AHS) (n = 13), or a single intravenous bolus of 15 &#181;g/kg terlipressin plus 15 mL/kg 5% albumin plus 3% hypertonic saline (TAHS) (n = 13) were used.</li>
	<li>After the infusion, follow up the animal for 120 min. Record the hemodynamic parameters and obtain blood samples every 30 min for arterial and venous blood gas profiles and lactate concentration determination. Do not make resuscitation efforts during this period.</li>
</ol>
6. End of the experiment and euthanasia
<ol>
	<li>Once the experiment is completed, use a sedative overdose (5 &#181;g/kg fentanyl and 10 mg/kg propofol) and an intravenous injection of potassium chloride (2 mEq/kg) to sacrifice all successfully resuscitated animals.</li>
	<li>Confirm the absence of circulation by asystole or pulseless electrical activity on a continuous EKG display, the absence of pulsatile flow during invasive arterial pressure monitoring, and the absence of other vital signs.</li>
	<li>If, during the follow-up period, arterial blood pressure decreases below 25 mmHg, sacrifice the animal to avoid further suffering.</li>
</ol>

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The authors of this work do not have any conflict of interest.

Performing procedures on young pigs can be complex and potentially life-threatening due to certain anatomical and physiological features of these animals. To achieve consistent results and reduce the loss of animals, there are some critical steps that should be carefully considered. First, achieving an adequate level of sedation is essential to minimize the animal stress response, which, if excessive, may alter the results due to endogenic catecholamine release. It is also important to avoid delays between the intramuscular injection and intubation, as animals can develop a severe stress response with tachycardia and irreversible metabolic acidosis that may precipitate the end of the experiment. Although other groups use inhaled anesthetics with good results20,21, we prefer intravenous medications, as inhaled sedatives do not allow the measurement of respiratory gas exchange with indirect calorimetry. In our experience, a combination of propofol and fentanyl is effective and has very few adverse effects. Careful temperature management throughout the experiment is another key aspect of the protocol, as rapid changes in temperature can significantly affect the animal&#39;s hemodynamic response to shock, falsifying the results or ultimately leading to the failure of the experiment.
Another crucial part of instrumentation is intubation, given the particularities of porcine anatomy and their susceptibility to laryngospasm. Therefore, the procedure should be performed by at least one operator with previous experience, and the use of a stylet and muscle relaxation is advisable10,22. Catheterization of vessels can also be challenging due to the small size of the animals. For femoral access, a sonography-guided puncture is preferable, as the vessels are located deep, usually have small diameters, and show different courses and positions22. For cervical access, we use surgical access to allow the placement of the carotid flow probe, but the ultrasound technique is also feasible23,24.&#160;Cannulation of the external jugular vein is usually preferred due to its wider diameter, its superficial location, and lower number of surrounding structures22. Catheters should be flushed immediately after insertion with saline solutions to prevent occlusion. We do not use heparin to avoid coagulation alterations. Also, initially, we avoided the administration of glucose infusions to prevent potential distortion of the hemodynamic response by the administration of extra fluids, but we found that animals developed severe and early hypoglycemia. Finally, even with anesthesia and the less invasive techniques used nowadays, instrumentation generates a significant stress response in animals, so it is desirable to leave enough time for recovery before initiating the removal of blood. Regarding induction of the hemorrhagic shock, we recommend the removal of 30 mL/kg, as this generates a significant pathophysiological response with excellent survival rates. In our experience, infant swine do not tolerate larger amounts of blood loss, and mortality is high. Gradual withdrawal of blood is also important, as rapid removal can result in severe hemodynamic instability and early death of the animal.
Although there is a wide variety of species and experimental models available to researchers, the ideal model of animal hemorrhagic shock-simple, easily reproducible, and accurate replication of the clinical situation-still represents a challenge. Small animal models-primarily mice and rats-are used to investigate the pathophysiological mechanisms of shock. However, their small size significantly complicates the performance of surgical and sampling procedures. Larger animals, such as dogs and pigs, are more expensive and complex to handle, but their size and physiological similarities with humans make them more suitable for preclinical evaluation of the treatment strategies. However, the use of dogs in the past and still nowadays is ethically questionable. They do not offer any advantage over pigs as experimental animal models, and their intelligence and the special bilateral relationship between humans and dogs situate them in a higher position in the phylogenetic scale6,7,8.
In view of all this, adult swine have been extensively used for cardiovascular research due to their similarities with adult human physiology, size, and anatomy, which is better than most species. However, as it has been well established in the literature, there are significant differences between human adult and pediatric patients in terms of the cardiovascular system, blood volume, temperature regulation, and response to shock2,3,4. At the same time, evidence shows that these differences also apply to swine, and piglets have been found to have cardiovascular, cerebrovascular, hematological, and electrolyte profiles very similar to those in pediatric human patients9,25. Finally, beyond these anatomic and physiologic differences between adults and infants in both species, the use of infant animal models, especially minipigs, provides the opportunity to test the same devices that are used in the real clinical setting for monitoring. In many cases, the reliability of these devices has been proven to be low because of a simple adaptation of the adult algorithms, sensors, or scales. All these aspects support the importance of developing specific pediatric animal models and their relevance in terms of translational utility to the pediatric clinical setting.
Besides the type of animal, there are three basic models generally used in the study of hemorrhagic shock: controlled hemorrhage-either by volume or pressure-and uncontrolled hemorrhage. The protocol presented in this article describes a fixed-volume hemorrhage model, in which a fixed blood volume, usually calculated by the percentage of body weight, is removed over a time period set by the observer. On the contrary, in fixed-pressure hemorrhage models, animals are bled to a predetermined MAP, which is then maintained with periodic bleeding or fluid infusions during a specified period, depending on the animal species and the degree or outcome of shock. Both fixed-volume and fixed-pressure hemorrhagic shock models allow the study of shock-induced pathophysiological changes under controlled conditions, offering a clear advantage in terms of reproducibility and standardization. However, their main limitation is that they do not allow the study of the effects of different resuscitation strategies on active bleeding, where aggressive fluid resuscitation before the surgical control of hemorrhage is known to increase bleeding and decrease survival, due to inhibition of the formation of the thrombus and the rise in the mean blood pressure. Uncontrolled hemorrhage models induced by a standardized vascular trauma-crush/laceration of liver and spleen, artery injury, or amputation of an appendage-have been suggested to better reflect the clinical situation, thereby allowing a better understanding of the effects of different fluid resuscitation strategies and other interventions, such as hypothermia and hemostatic products. However, despite being clinically most relevant, these uncontrolled hemorrhage models exert some clear disadvantages in terms of standardization and reproducibility. In view of all this, it seems that the ideal model does not exist, and therefore research in this field must balance clinical relevance with experimental standardization and reliability6,7,8,9,26.
The model described in this study may offer broad potential applications in cardiovascular research, such as the investigation of endothelial dysfunction and microcirculation alterations18 during shock, as well as the validation of different hemodynamic monitoring systems. Moreover, it can also be used in other research fields, allowing the study of endocrine or immune responses after severe hemorrhage as well as the determination of side effects of different fluids and vasopressors. However, regarding the research on different resuscitation strategies, it is advisable to study their effects in uncontrolled hemorrhage models before implementing changes in the clinical setting7,26.
Besides the difficulty of extrapolating the results to real life, this model has other limitations. To begin with, there are some confounding variables related to the experimental setup, such as the use of anesthetic agents or mechanical ventilation, that may attenuate the physiologic responses during shock and complicate the interpretation of results. Besides, instrumentation stress response on the animals and temperature control may affect macro- and microcirculation through different mechanisms. Another important limitation of this model-related to the experimental necessities and availability of resources-is the limited post-traumatic observation period, which further limits the study of the long-term consequences of hemorrhagic shock. In addition, despite the physiological similarities between humans and pigs, there are some differences between species that should be considered. The coagulation system, for example, appears to be more effective in swine27,28.&#160;Also, lactate and succinate plasma levels differ between species, and pigs have basal alkalosis, which may lead to an underestimation of the effects of hemorrhage on acid-base balance29. Finally, it is also well known that the inflammatory and immune responses, as well as some vasopressor receptors, are different in swine9. Specific animal differences also need to be considered as influencing factors. Several studies have indicated gender differences in terms of susceptibility to shock, with females having a significant survival advantage over males6,9.&#160;Nevertheless, in the experiments conducted in this study, we use animals from the same age group and with a similar genetical background to minimize the potential variability inherent to species.
In conclusion, this article provides a practical and step-by-step guide to setting up a swine model of pediatric hemorrhagic shock. Compared to other existing models, this is a reliable and easy-to-follow protocol with broad applicability in biomedical research, either for the investigation of pathophysiological responses after severe hemorrhage or for the evaluation of different resuscitation strategies.

Life-threatening hemorrhage due to trauma, although uncommon, is the leading cause of death in pediatric patients1,2. Additional causes of hemorrhagic shock include hemorrhagic fever, gastrointestinal bleeding, liver surgery, and cardiac surgery, especially when cardiopulmonary bypass is used3.
Contrary to the adult population, there is insufficient data on the management of pediatric hemorrhagic shock, which is largely based on expert opinions or directly translated from adult practice2,4. However, the translation of management strategies from adults may not be appropriate. For example, clinical indicators validated in adults are difficult to extrapolate to pediatric patients due to the physiologic heterogeneity present across groups of different ages and the different injury patterns predominant in the pediatric population. Consequently, specific endpoints that would trigger intervention in the pediatric patient are not well defined. Moreover, there is not enough evidence on the deleterious effects that therapies currently implemented in adults may have on children2,4,5.
In view of all this, further investigation is needed to establish specific resuscitation thresholds to prompt intervention, as well as to better determine which are the most appropriate therapies for pediatric hemorrhagic shock. However, the development of quality and clinically relevant studies of life-threatening hemorrhage in children is difficult, due to the paucity of patients and the already mentioned heterogeneity in the pediatric population from the neonatal period through adolescence.
The clinical relevance of hemorrhagic shock, in addition to the difficulties in performing clinical studies on pediatric patients, emphasize the need for preclinical evaluations on animal models to study pathophysiology following hemorrhagic shock in children, as well as to compare different therapies.Several animal models have been widely used in research to study hemorrhagic shock6,7,8,9.&#160;Due to their anatomical and physiological similarities with humans, pigs are highly valued in biomedical research. Regarding the advantages of using specific infant models, evidence shows that immature swine hemodynamics, as well as respiratory, hematologic, and metabolic systems, are highly comparable to those in young humans9. This confers a unique opportunity to simulate a clinical scenario of hemorrhagic shock in children.
In this model, hemorrhage is induced by withdrawing a previously calculated blood volume. Subsequently, the pig is monitored, and different resuscitation fluids are administered.
Here, we describe a precise and highly reproducible model of hemorrhagic shock in immature swine. The model yields hemodynamic data that characterizes compensatory mechanisms activated in response to severe hemorrhage.

<table><tbody><tr><td>ADA swabs</td><td>Albino Dias de Andrade, S.A.</td><td>300575750400&amp;nbsp;&amp;nbsp;&amp;nbsp;</td><td>Non-woven swabs</td></tr><tr><td>Alaris SE</td><td>Carefusion</td><td>N/A</td><td>Volumetric infusion pump</td></tr><tr><td>Atracurium</td><td>Aspen Pharma Trading Limited. Dublin, Ireland</td><td>N/A</td><td>Muscle relaxant</td></tr><tr><td>Atropine 1 mg/mL</td><td>B. Braun</td><td>481377/1013</td><td></td></tr><tr><td>Barrier adhesive aperture drape</td><td>M&amp;ouml;lnlycke</td><td>63621</td><td></td></tr><tr><td>BD emerald syringe 5 mL, 10 mL, 20 mL&amp;nbsp;</td><td>Becton Dickinson S.A</td><td>https://www.bd.com/en-eu/offerings/capabilities/syringes-and-needles/injection-syringes/bd-emerald-3-piece-syringe</td><td>various options available</td></tr><tr><td>BLF21A laser doppler monitor</td><td>Transonic Systems Inc.</td><td>BLF21A</td><td>Skin blood flow monitor</td></tr><tr><td>&amp;nbsp;BlueSensor NF ECG electrodes</td><td>Ambu&amp;nbsp;</td><td>NF-50-A/12</td><td></td></tr><tr><td>Check-Flo performer introducer set 5Fr</td><td>Cook Medical</td><td>G12018</td><td>Vascular Sheath</td></tr><tr><td>Datex ohmeda S5</td><td>GE Healthcare Finland Oy, Helsinki, Finland</td><td>M1162897</td><td>Hemodynamic monitor</td></tr><tr><td>Fentanyl 0.05 mg/mL</td><td>Kern Pharma</td><td>N/A</td><td>Anesthesia</td></tr><tr><td>GE Vivid S5</td><td>GE Healthcare&amp;nbsp;</td><td>S series</td><td>Ultrasound machine</td></tr><tr><td>Introcan Safety 18 G, 22 G, 24 G</td><td>B. Braun</td><td>Introcan series</td><td>Safety intravenous catheter</td></tr><tr><td>INVOS cerebral/somatic oximetry adult sensors</td><td>Medtronic PLC, USA</td><td>https://www.medtronic.com/covidien/en-us/products/cerebral-somatic-oximetry/invos-cerebral-somatic-oximetry-adult-sensors.html</td><td></td></tr><tr><td>INVOS OXIMETER cerebral/somatic</td><td>Somanetics</td><td>08-10566</td><td>Regional oxygenation monitor</td></tr><tr><td>Ketamin 50 mg/mL</td><td>Pfizer, S.L.</td><td>47034</td><td>Sedation</td></tr><tr><td>Leon plus</td><td>Heinen + L&amp;ouml;wenstein</td><td>N/A</td><td>Ventilator</td></tr><tr><td>Life scope VS</td><td>Nihon Kohden</td><td>N/A</td><td>Bedside monitor</td></tr><tr><td>Miller laryngoscope blade 12&amp;Prime;</td><td>Jorgensen Labs, USA</td><td>J0449F</td><td>Laryngoscope</td></tr><tr><td>Multi-lumen central venous catheterization set 7 French, 3 lumen, 30 cm</td><td>Arrow</td><td>CS-14703</td><td>Central venous catheter</td></tr><tr><td>Nellcor WarmTouch 5300A</td><td>Covidien</td><td></td><td>Thermal blancket</td></tr><tr><td>Nitrile gloves</td><td>Medihands</td><td>KS-ST RT021</td><td>Single use gloves</td></tr><tr><td>Pediatric SomaSensor INVOS cerebral/somatic</td><td>Covidien</td><td>https://www.medtronic.com/covidien/en-us/products/cerebral-somatic-oximetry.html</td><td>Disposable regional oxygen saturation sensor</td></tr><tr><td>PICCO monitoring kit</td><td>Pulsion Medical Systems</td><td>PV8215</td><td></td></tr><tr><td>PICCO thermodilution catheter 5F/20 cm</td><td>Pulsion Medical Systems</td><td>N/A</td><td></td></tr><tr><td>Propofol Lipoven 10 mg/mL</td><td>Fresenius Kabi, Spain</td><td>N/A</td><td>Anesthesia</td></tr><tr><td>Pulse contour cardiac output (PiCCO2)</td><td>Pulsion Medical Systems</td><td>N/A</td><td>Hemodynamic monitor</td></tr><tr><td>R&amp;uuml;sch flexislip</td><td>Teleflex Medical</td><td>503700</td><td>Endotracheal tube stylet</td></tr><tr><td>Softa swabs</td><td>B. Braun</td><td>19579</td><td>Alcohol pads</td></tr><tr><td>Surgical silk sutures USP 0</td><td>Arag&amp;oacute;, Barcelona, Spain.&amp;nbsp;</td><td>6245</td><td></td></tr><tr><td>TruWave pressure monitoring set</td><td>Edwards</td><td>T001767A</td><td>Pressure monitoring set</td></tr><tr><td>Ultrasound transmission gel</td><td>Ultragel Hungary 2000 Kft.&amp;nbsp;</td><td>UC260</td><td></td></tr></tbody></table>

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.

The presented model has been successfully used in several experiments to study macrocirculatory and microcirculatory changes following hemorrhagic shock and subsequent resuscitation, comparing different fluids and vasoactive drugs16,17,18,19.
Considering the response to shock, this model has consistently shown that a controlled hemorrhage produces marked changes in hemodynamic parameters, as well as in cerebral and tissue perfusion.
Following volume withdrawal, significant tachycardia and a decrease in MAP, CI, SVI, blood volume parameters (GEDVI and ITBI), and carotid arterial blood flow, along with an increase in systemic vascular resistance index, are detected (Figure 1 and Figure 2).
Regarding systemic perfusion parameters, lactate increases significantly, whereas ScvO2, CuTBF, and bTO decreases (Figure 3). Variations in central venous pressure, Dt/Dpmax, and ELWare not usually registered.
As for laboratory parameters, hemoglobin content and hematocrit do not decrease until after fluids have been administered. Albumin concentration decreases, and troponin levels increase significantly after controlled hemorrhage. Other parameters, including core temperature, PaO2, PaCO2, arterial oxygen saturation, EtCO2, electrolytes, and kidney and liver function parameters, usually remain stable.
Besides its utility in analyzing the cardiovascular and biochemical responses to shock, this model has been shown to successfullydiscriminate between different resuscitation fluids.
In previous studies, we have aimed to determine if, in an infant animal model of hemorrhagic shock, the use of a lower volume infusion of hypertonic fluids-alone or combined with different vasopressors- would improve global hemodynamic and perfusion parameters when compared to normal saline.
As reported previously, we have consistently observed that the infusion of hypertonic fluids produces a similar response to the infusion of twice the volume of isotonic fluid16,17,18.
More specifically, the use of albumin plus hypertonic saline produced a greater and longer volume expansion than normal saline or hypertonic saline alone, with significant differences in HR, SVI, and PPV, and the absence of a progressive fall after volume expansion in blood pressure and GEDVI, as observed in the other groups (Figure 1 and Figure 2). Furthermore, we have also observed a greater improvement of perfusion parameters with hypertonic albumin, represented as a greater increase of bTOI and CaBF, and a greater decrease of lactate levels than the other groups in comparison with the beginning of the fluid expansion (Figure 3). We believe that this difference might be secondary to the capacity of albumin to increase the blood volume and remain for a longer period of time within the intravascular compartment than normal saline.Interestingly, we have seen that the addition of a single bolus of terlipressin at the beginning of fluid resuscitation yielded similar results to those observed in the hypertonic albumin group, without any extra benefits in terms of hemodynamic or perfusion parameters17,18.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64815/64815fig1v2.jpg" /> 
Figure 1: Hemodynamic parameters. (A) Evolution of heart rate, (B) mean arterial pressure, (C) cardiac index at baseline (t0&#39;),&#160;and (D) systemic vascular resistance index at baseline (t0&#39;). Throughout the course of the experiment: end of controlled bleeding (Shock30&#39;); the beginning of infusion, 30 min after the end of controlled bleeding (Res0&#39;); end of infusion (Res30&#39;); follow-up 30 min after the end of infusion (Obs30&#39;); follow-up 60 min after the end of infusion (Obs60&#39;); follow-up 90 min after the end of infusion (Obs90&#39;). (*) Significant difference (p &#60; 0.05) from the baseline, same group. (&#8225;) p &#60; 0.05 from hemorrhage, same group. (#) p &#60; 0.05 from group NS. Abbreviations: NS = normal saline; AHS = hypertonic saline albumin; TAHS = terlipressin plus hypertonic saline albumin. Data are presented as mean and standard deviation. This figure is adapted with permission from Urbano et al.17. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64815/64815fig1v2large.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/64815/64815fig2v2.jpg" /> 
Figure 2: Blood volume parameters. (A) Evolution of stroke volume index, (B) pulse pressure variation, and (C) global end diastolic volume index at baseline (t0&#39;). Throughout the course of the experiment: end of controlled bleeding (Shock30&#39;); beginning of infusion, 30 min after the end of controlled bleeding (Res0&#39;); end of infusion (Res30&#39;); follow-up 30 min after the end of infusion (Obs30&#39;); follow-up 60 min after the end of infusion (Obs60&#39;); follow-up 90 min after the end of infusion (Obs90&#39;). (*) Significant difference (p &#60; 0.05) from baseline, same group. (&#8225;) p &#60; 0.05 from hemorrhage, same group. (#) p &#60; 0.05 from group NS. Abbreviations: NS = normal saline; AHS = hypertonic saline albumin; TAHS = terlipressin plus hypertonic saline albumin. Data are presented as mean and standard deviation. This figure is adapted with permission from Urbano et al.17. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64815/64815fig2v2large.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/64815/64815fig3v2.jpg" /> 
Figure 3: Systemic perfusion parameters. (A) Evolution of arterial blood lactate, (B) central venous blood oxygen saturation, and (C) brain tissue oxygenation index at baseline (t0&#39;). Throughout the course of the experiment: end of controlled bleeding (Shock30&#39;); beginning of infusion, 30 min after the end of controlled bleeding (Res0&#39;); end of infusion (Res30&#39;); follow-up 30 min after the end of infusion (Obs30&#39;); follow-up 60 min after the end of infusion (Obs60&#39;); follow-up 90 min after the end of infusion (Obs90&#39;). (*) Significant difference (p &#60; 0.05) from baseline, same group. (&#8225;) p &#60; 0.05 from hemorrhage, same group. (#) p &#60; 0.05 from group NS. Data are presented as mean and standard deviation. This figure is adapted with permission from Urbano et al.17. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64815/64815fig3v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Description of a Swine Infant Model of Volume-Controlled Hemorrhagic Shock at JoVE.com

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

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 ...

description-swine-infant-model-volume-controlled-hemorrhagic

Nanyang Technological University

Research

JoVE Journal

Medicine

913 Views.  Gregorio Marañón University General Hospital. Hemorragische shock is de belangrijkste doodsoorzaak bij kinderen in de ontwikkelde wereld. Deze studie biedt de mogelijkheid om de respons op verschillende therapieën voor hemorragische shock te vergelijken. Dit is een eenvoudig en zeer reproduceerbaar model dat het mogelijk maakt om mechanismen te activeren als reactie op ernstige bloedingen bij een onvolwassen proefpersoon.In dit protocol is de vertaling van de resultaten hoog, en met hulp om de keuze van vloeistoffen en rust voor ...

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

Fixed Volume or Fixed Pressure: A Murine Model of Hemorrhagic Shock

It is common knowledge that severe blood loss and traumatic injury can lead to a cascade of detrimental signaling events often resulting in mortality. 1, 2, 3, 4, 5 These signaling events can also lead to sepsis and/or multiple organ dysfunction (MOD). 6, 7, 8, 9 It is critical then to investigate the causes of suppressed immune function and detrimental signaling cascades in order to develop more effective ways to help patients who suffer from traumatic injuries. 10 This fixed pressure Hemorrhagic Shock (HS) procedure, although technically challenging, is an excellent resource for investigation of these pathophysiologic conditions. 11, 12, 13 Advances in the assessment of biological systems, i.e. Systems Biology have enabled the scientific community to further understand complex physiologic networks and cellular communication patterns. 14 Hemorrhagic Shock has proven to be a vital tool for unveiling these cellular communication patterns as they relate to immune function. 15, 16, 17, 18 This procedure can be mastered! This procedure can also be used as either a fixed volume or fixed pressure approach. We adapted this technique in the murine model to enhance research in innate and adaptive immune function. 19, 20, 21 Due to their small size HS in mice presents unique challenges. However due to the many available mouse strains, this species represents an unparalleled resource for the study of the biologic responses. The HS model is an important model for studying cellular communication patterns and the responses of systems such as hormonal and inflammatory mediator systems, and danger signals, i.e. DAMP and PAMP upregulation as it elicits distinct responses that differ from other forms of shock. 22, 23, 24, 25 The development of transgenic murine strains and the induction of biologic agents to inhibit specific signaling have presented valuable opportunities to further elucidate our understanding of the up and down regulation of signal transduction after severe blood loss, i.e. HS and trauma 26, 27, 28, 29, 30. 
There are numerous resuscitation methods (R) in association with HS and trauma. 31, 32, 33, 34 A fixed volume resuscitation method of solely lactated ringer solution (LR), equal to three times the shed blood volume, is used in this model to study endogenous mechanisms such as remote organ injury and systemic inflammation. 35, 36, 38 This method of resuscitation is proven to be effective in evaluating the effects of HS and trauma 38, 39.

The Hemorrhagic Shock model has been a reliable and reproducible resource facilitating the identification and understanding of signaling cascades associated with inflammation and end-organ damage after trauma. This article provides a step-by-step description of surgical and mechanical aspects associated with the Hemorrhagic Shock experimental procedure in mice.

It is common knowledge that severe blood loss and traumatic injury can lead to a cascade of detrimental signaling events often resulting in mortality. 1, ...

A Swine Model of Neonatal Asphyxia

Annually more than 1 million neonates die worldwide as related to asphyxia. Asphyxiated neonates commonly have multi-organ failure including hypotension, perfusion deficit, hypoxic-ischemic encephalopathy, pulmonary hypertension, vasculopathic enterocolitis, renal failure and thrombo-embolic complications. Animal models are developed to help us understand the patho-physiology and pharmacology of neonatal asphyxia. In comparison to rodents and newborn lambs, the newborn piglet has been proven to be a valuable model. The newborn piglet has several advantages including similar development as that of 36-38 weeks human fetus with comparable body systems, large body size (&tilde;1.5-2 kg at birth) that allows the instrumentation and monitoring of the animal and controls the confounding variables of hypoxia and hemodynamic derangements.
We here describe an experimental protocol to simulate neonatal asphyxia and allow us to examine the systemic and regional hemodynamic changes during the asphyxiating and reoxygenation process as well as the respective effects of interventions. Further, the model has the advantage of studying multi-organ failure or dysfunction simultaneously and the interaction with various body systems. The experimental model is a non-survival procedure that involves the surgical instrumentation of newborn piglets (1-3 day-old and 1.5-2.5 kg weight, mixed breed) to allow the establishment of mechanical ventilation, vascular (arterial and central venous) access and the placement of catheters and flow probes (Transonic Inc.) for the continuously monitoring of intra-vascular pressure and blood flow across different arteries including main pulmonary, common carotid, superior mesenteric and left renal arteries. Using these surgically instrumented piglets, after stabilization for 30-60 minutes as defined by Z&lt;10% variation in hemodynamic parameters and normal blood gases, we commence an experimental protocol of severe hypoxemia which is induced via normocapnic alveolar hypoxia. The piglet is ventilated with 10-15% oxygen by increasing the inhaled concentration of nitrogen gas for 2h, aiming for arterial oxygen saturations of 30-40%. This degree of hypoxemia will produce clinical asphyxia with severe metabolic acidosis, systemic hypotension and cardiogenic shock with hypoperfusion to vital organs. The hypoxia is followed by reoxygenation with 100% oxygen for 0.5h and then 21% oxygen for 3.5h. Pharmacologic interventions can be introduced in due course and their effects investigated in a blinded, block-randomized fashion.

Large animal models have good translational values in the examination of physiology and pharmacology of neonatal asphyxia. Using newborn piglets, we develop an experimental protocol to simulate neonatal asphyxia which has advantages of studying the systemic and regional hemodynamics, oxygen transport with biochemical and pathologic pathways and correlations.

Annually more than 1 million neonates die worldwide as related to asphyxia. Asphyxiated neonates commonly have multi-organ failure including hypotension, ...

Uncontrolled Hemorrhagic Shock Modeled via Liver Laceration in Mice with Real Time Hemodynamic Monitoring

Uncontrolled hemorrhage is an important cause of preventable deaths among trauma patients. We have developed a murine model of uncontrolled hemorrhage via a liver laceration that results in consistent blood loss, hemodynamic alterations, and survival.
Mice undergo a standardized resection of the left-middle lobe of the liver. They are allowed to bleed without mechanical intervention. Hemostatic agents can be administered as pre-treatment or rescue therapy depending on the interest of the investigator. During the time of hemorrhage, real-time hemodynamic monitoring via a left femoral arterial line is performed. Mice are then sacrificed, blood loss is quantified, blood is collected for further analysis, and organs are harvested for analysis of injury. Experimental design is described to allow for simultaneous testing of multiple animals.
Liver hemorrhage as a model of uncontrolled hemorrhage exists in the literature, primarily in rat and porcine models. Some of these models utilize hemodynamic monitoring or quantify blood loss but lack consistency. The present model incorporates quantification of blood loss, real-time hemodynamic monitoring in a murine model that offers the advantage of using transgenic lines and a high-throughput mechanism to further investigate the pathophysiologic mechanisms in uncontrolled hemorrhage.

Uncontrolled hemorrhage, an important cause of mortality among trauma patients, can be modeled using a standard liver laceration in a murine model. This model results in consistent blood loss, survival, and allows for testing hemostatic agents. This article provides the step-by-step process to perform this valuable model.

Uncontrolled hemorrhage is an important cause of preventable deaths among trauma patients. We have developed a murine model of uncontrolled hemorrhage via ...

A Chronic Cardiac Ischemia Model in Swine Using an Ameroid Constrictor

Cardiovascular disease remains the number one cause of mortality in the United States. There are numerous approaches to treating these diseases, but regardless of the approach, an in vivo model is needed to test each treatment. The pig is one of the most used large animal models for cardiovascular disease. Its heart is very similar in anatomy and function to that of a human. The ameroid placement technique creates an ischemic area of the heart, which has many useful applications in studying myocardial infarction. This model has been used for surgical research, pharmaceutical studies, imaging techniques, and cell therapies.
There are several ways of inducing an ischemic area in the heart. Each has its advantages and disadvantages, but the placement of an ameroid constrictor remains the most widely used technique. The main advantages to using the ameroid are its prevalence in existing research, its availability in various sizes to accommodate the anatomy and size of the vessel to be constricted, the surgery is a relatively simple procedure, and the post-operative monitoring is minimal, since there are no external devices to maintain. This paper provides a detailed overview of the proper technique for the placement of the ameroid constrictor.

The purpose of this protocol is to demonstrate the placement of a delayed constricting device (an ameroid constrictor) around a coronary artery in a swine model. This device creates an ischemic area of the heart that is useful for studying new diagnostic imaging techniques and new methods of treatment.

Cardiovascular disease remains the number one cause of mortality in the United States. There are numerous approaches to treating these diseases, but ...

Tachycardia-Induced Cardiomyopathy As a Chronic Heart Failure Model in Swine

A stable and reliable model of chronic heart failure is required for many experiments to understand hemodynamics or to test effects of new treatment methods. Here, we present such a model by tachycardia-induced cardiomyopathy, which can be produced by rapid cardiac pacing in swine.
A single pacing lead is introduced transvenously into fully anaesthetized healthy swine, to the apex of the right ventricle, and fixated. Its other end is then tunneled dorsally to the paravertebral region. There, it is connected to an in-house modified heart pacemaker unit that is then implanted in a subcutaneous pocket. 
After 4 - 8 weeks of rapid ventricular pacing at rates of 200 - 240 beats/min, physical examination revealed signs of severe heart failure - tachypnea, spontaneous sinus tachycardia, and fatigue. Echocardiography and X-ray showed dilation of all heart chambers, effusions, and severe systolic dysfunction. These findings correspond well to decompensated dilated cardiomyopathy and are also preserved after the cessation of pacing.
This model of tachycardia-induced cardiomyopathy can be used for studying the pathophysiology of progressive chronic heart failure, especially hemodynamic changes caused by new treatment modalities like mechanical circulatory supports. This methodology is easy to perform and the results are robust and reproducible.

Here, we present a protocol to produce tachycardia-induced cardiomyopathy in swine. This model represents a potent way to study the hemodynamics of progressive chronic heart failure and the effects of applied treatment.

A stable and reliable model of chronic heart failure is required for many experiments to understand hemodynamics or to test effects of new treatment ...

Complete and Partial Aortic Occlusion for the Treatment of Hemorrhagic Shock in Swine

Hemorrhage remains the leading cause of preventable deaths in trauma. Endovascular management of non-compressible torso hemorrhage has been at the forefront of trauma care in recent years. Since complete aortic occlusion presents serious concerns, the concept of partial aortic occlusion has gained a growing attention. Here, we present a large animal model of hemorrhagic shock to investigate the effects of a novel partial aortic balloon occlusion catheter and compare it with a catheter that works on the principles of complete aortic occlusion. Swine are anesthetized and instrumented in order to conduct controlled fixed-volume hemorrhage, and hemodynamic and physiological parameters are monitored. Following hemorrhage, aortic balloon occlusion catheters are inserted and inflated in the supraceliac aorta for 60 min, during which the animals receive whole-blood resuscitation as 20% of the total blood volume (TBV). Following balloon deflation, the animals are monitored in a critical care setting for 4 h, during which they receive fluid resuscitation and vasopressors as needed. The partial aortic balloon occlusion demonstrated improved distal mean arterial pressures (MAPs) during the balloon inflation, decreased markers of ischemia, and decreased fluid resuscitation and vasopressor use. As swine physiology and homeostatic responses following hemorrhage have been well-documented and are like those in humans, a swine hemorrhagic shock model can be used to test various treatment strategies. In addition to treating hemorrhage, aortic balloon occlusion catheters have become popular for their role in cardiac arrest, cardiac and vascular surgery, and other high-risk elective surgical procedures.

Here, we present a protocol demonstrating a hemorrhagic shock model in swine that uses aortic occlusion as a bridge to definitive care in trauma. This model has application in testing a wide range of surgical and pharmacological therapeutic strategies.

Hemorrhage remains the leading cause of preventable deaths in trauma. Endovascular management of non-compressible torso hemorrhage has been at the ...

Standardized Hemorrhagic Shock Induction Guided by Cerebral Oximetry and Extended Hemodynamic Monitoring in Pigs

Hemorrhagic shock ranks among the main reasons for severe injury-related death. The loss of circulatory volume and oxygen carriers can lead to an insufficient oxygen supply and irreversible organ failure. The brain exerts only limited compensation capacities and is particularly at high risk of severe hypoxic damage.This article demonstrates the reproducible induction of life-threatening hemorrhagic shock in a porcine model by means of calculated blood withdrawal. We titrate shock induction guided by near-infrared spectroscopy&#160;and extended hemodynamic monitoring to display systemic circulatory failure, as well as cerebral microcirculatory oxygen depletion. In comparison to similar models that primarily focus on predefined removal volumes for shock induction, this approach highlights a titration by means of the resulting failure of macro- and microcirculation.

Hemorrhagic shock is a severe complication in seriously injured patients, which leads to life-threatening oxygen undersupply. We present a standardized method to induce hemorrhagic shock via blood withdrawal in pigs that is guided by hemodynamics and microcirculatory cerebral oxygenation.

Hemorrhagic shock ranks among the main reasons for severe injury-related death. The loss of circulatory volume and oxygen carriers can lead to an ...

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, ...

Vascular Cannulation with Hemorrhagic Shock Induction in Rats

Developing a Clinically Relevant Hemorrhagic Shock Model in Rats

Over the recent decades, the development of animal models allowed us to better understand various pathologies and identify new treatments. Hemorrhagic shock, i.e., organ failure due to rapid loss of a large volume of blood, is associated with a highly complex pathophysiology involving several pathways. Numerous existing animal models of hemorrhagic shock strive to replicate what happens in humans, but these models have limits in terms of clinical relevance, reproducibility, or standardization. The aim of this study was to refine these models to develop a new model of hemorrhagic shock. Briefly, hemorrhagic shock was induced in male Wistar Han rats (11-13 weeks old) by a controlled exsanguination responsible for a drop in the mean arterial pressure. The next phase of 75 min was to maintain a low mean arterial blood pressure, between 32 mmHg and 38 mmHg, to trigger the pathophysiological pathways of hemorrhagic shock. The final phase of the protocol mimicked patient care with an administration of intravenous fluids, Ringer Lactate solution, to elevate the blood pressure. Lactate and behavioral scores were assessed 16 h after the protocol started, while hemodynamics parameters and plasmatic markers were evaluated 24 h after injury. Twenty-four hours post-hemorrhagic shock induction, the mean arterial and diastolic blood pressure were decreased in the hemorrhagic shock group (p&#160;&#60; 0.05). Heart rate and systolic blood pressure remained unchanged. All organ damage markers were increased with the hemorrhagic shock (p &#60; 0.05). The lactatemia and behavioral scores were increased compared to the sham group (p &#60; 0.05). In conclusion, we demonstrated that the protocol described here is a relevant model of hemorrhagic shock that can be used in subsequent studies, particularly to evaluate the therapeutic potential of new molecules.

Author Spotlight: Developing Innovative Therapeutic Strategies for Hemorrhagic Shock Research

Hemorrhagic shock kills 1.9 million people worldwide every year. Small animals are frequently used as hemorrhagic shock models but are associated with issues of standardization, reproducibility, and clinical significance, thus limiting their relevance. This article describes developing a new clinically relevant hemorrhagic shock model in rats.

Over the recent decades, the development of animal models allowed us to better understand various pathologies and identify new treatments. Hemorrhagic ...

Swine Axillary Artery Injury Model for Testing Pre-Hospital Hemostatic Products

A New Hybrid Quantitative Evaluation Model for Axillary Junctional Hemorrhage in Swine

In this study, we developed and validated a hybrid quantitative model for simulating upper extremity junctional hemorrhage in swine, aiming to advance the development of pre-hospital hemostatic products. Utilizing 12 healthy 8-month-old male Yorkshire swine, we demonstrated the feasibility of a swine axillary artery injury model for evaluating hemostatic efficacy. Animals were divided into three groups to undergo volume-controlled hemorrhage (VCH), mimicking Class I-III hemorrhagic shock by withdrawing blood at different rates. Subsequent external compression was applied using a novel device consisting of a mechanical arm and an inflatable hemostatic balloon, achieving controlled pressure to enhance clot formation. Hemodynamic parameters, including heart rate and blood pressure, were continuously monitored, highlighting the impact of controlled hemorrhage and external compression on physiological responses. The findings suggest that the combination of VCH with targeted external compression effectively simulates clinical scenarios of axillary artery injury, providing a valuable model for testing hemostatic interventions in a controlled, standardized manner. This study underscores the potential of the model in facilitating the development and evaluation of new hemostatic agents and devices for managing junctional hemorrhages.

This study presents a hybrid quantitative model for axillary junctional hemorrhage in swine, enhancing pre-hospital hemostatic intervention evaluation.

In this study, we developed and validated a hybrid quantitative model for simulating upper extremity junctional hemorrhage in swine, aiming to advance the ...

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

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