Name: standardized hemorrhagic shock induction guided by cerebral oximetry and extended hemodynamic monitoring in pigs
Uploaded: 2019-05-21T13:00:00.000+00:00
Duration: 7 min 51 s
Description: Watch this Scientific Journal Video about Standardized Hemorrhagic Shock Induction Guided by Cerebral Oximetry and Extended Hemodynamic Monitoring in Pigs at JoVE.com

The authors want to thank Dagmar Dirvonskis for her excellent technical support.

The experiments in this protocol were approved by the State and Institutional Animal Care Committee (Landesuntersuchungsamt Rheinland-Pfalz, Koblenz, Germany; Chairperson: Dr. Silvia Eisch-Wolf; reference number: 23 177-07/G 14-1-084; 02.02.2015). The experiments were conducted in accordance with the Animal Research&#160;Reporting of In Vivo Experiments (ARRIVE) guidelines. The study was planned and conducted between November 2015 and March 2016. After extended literature research, the pig model was chosen as a well-established model for hemorrhagic shock. Seven anesthetized male pigs (Sus scrofa domestica) with a mean weight of 28 &#177; 2 kg and an age of 2-3 months were included in the protocol. The animals were cared for by a local breeder that was recommended by the State and Institutional Animal Care Committee. The animals were kept in their known environment as long as possible to minimize stress. Food, but not water was denied 6 h before the experiment was scheduled, to reduce the risk of aspiration. The representative time course is displayed in Figure 1.
1. Anesthesia, Intubation, and Mechanical Ventilation
<ol>
	<li>Sedate pigs with a combined injection of ketamine (4 mg&#183;kg-1) and azaperone (8 mg&#183;kg-1) in the neck or the gluteal muscle with a needle for intramuscular injection (1.2 mm). Ensure that the animals remain stable until the sedation sets in. 
	CAUTION: Gloves are absolutely necessary when handling animals.</li>
	<li>Transport the sedated animals to the laboratory. 
	NOTE: The animals fall deeply asleep and do not awake during normal handling, like when they are lifted into the transport cage. In this setting, the transport time was about 20 min with a special van for animal transport.</li>
	<li>Monitor the peripheral oxygen saturation (SpO2) with a sensor clipped to the pig&#39;s tail or ear directly after arrival.</li>
	<li>Disinfect the skin with colorless disinfection tincture and wait for 3 min before inserting a peripheral vein catheter (1.2 mm) into an ear vein. Then, induce anesthesia by an intravenous injection of fentanyl (4 &#181;g&#183;kg-1) and propofol (3 mg&#183;kg-1).</li>
	<li>When all reflexes are absent and spontaneous breathing expires, place the pigs in supine position on a stretcher and fix them with bandages. 
	NOTE: Adequate levels of anesthesia have to be confirmed by an experienced researcher by the absence of an eyelid reflex and other reactions to external stimuli.</li>
	<li>Immediately start noninvasive ventilation with a dog ventilation mask (size 2). Use the following ventilation parameters: inspiratory oxygen fraction (FiO2) = 1.0; respiratory rate = 14-16 min-1; peak inspiratory pressure &#60;20 cm H2O, positive end-expiratory pressure (PEEP) = 5 cm H2O.</li>
	<li>Maintain anesthesia via a continuous infusion of fentanyl (0.1-0.2 &#181;g&#183;kg-1&#183;h-1) and propofol (8-12 mg&#183;kg-1&#183;h-1) and start an infusion of balanced electrolyte solution (5 mL&#183;kg-1&#183;h-1).</li>
	<li>Facilitate endotracheal intubation by the application of a muscle relaxant (atracurium 0.5 mg&#183;kg-1).</li>
	<li>Secure the airway via intubation with a common endotracheal tube (ID 6-7) and an introducer. Use a common laryngoscope with a Macintosh blade (size 4). Two persons facilitate the procedure.
	<ol>
		<li>Person 1: Fix the tongue outside with a piece of tissue and open the snout with the other hand.</li>
		<li>Person 2: Perform a laryngoscopy.</li>
		<li>Person 2: When the epiglottis comes into view, move the laryngoscope ventrally. Lift up the epiglottis and make sure the vocal cords are visible. 
		NOTE: If the epiglottis does not move dorsally, it sticks to the soft palatine and can be mobilized by the tip of the tube. Alternatively, a blade with another size (3 or 5) or type (Miller blade) can be used.</li>
	</ol>
	</li>
	<li>Move the tube carefully through the vocal cords. 
	NOTE: The narrowest point of the trachea is not on the level of the vocal cords but subglottic. If tube insertion is not possible, try to rotate the tube or use a smaller tube.</li>
	<li>Pull the introducer out of the tube, use a 10 mL syringe to block the cuff with 10 mL of air, and control the cuff pressure with a cuff manager (30 cm H2O).</li>
	<li>Start mechanical ventilation after the tube is connected to a ventilator (PEEP = 5 cm H2O; tidal volume = 8 mL&#183;kg-1; FiO2 = 0.4; inspiration-to-expiration ratio = 1:2; respiratory rate = variable to achieve an end-tidal CO2 of &#60;6 kPa). 
	NOTE: Avoid fluctuation of the CO2 to minimize any respiratory effects on the cerebral perfusion.</li>
	<li>Make sure that tube position is correct by regular and periodic exhalation of CO2 via capnography, and check the double-sided ventilation through auscultation. 
	NOTE: If the tube is placed incorrectly, air inflation into the stomach rapidly forms a visible bulge in the abdominal wall, even before the capnography is installed. In this case, replacement of the tube and the insertion of a gastric tube are absolutely necessary.</li>
	<li>With two persons, place a gastric tube into the stomach to avoid reflux and vomiting.
	<ol>
		<li>Person 1: Fix the tongue outside with a piece of tissue and open the snout with the other hand.</li>
		<li>Person 2: Perform a laryngoscopy of the porcine larynx.</li>
		<li>Person 2: Visualize the esophagus.</li>
		<li>Person 2: Push the gastric tube inside the esophagus with a pair of Magill forceps until gastric fluid is drained. 
		NOTE: Sometimes, visualization is not easy. In this case, move the laryngoscope dorsally to the tube and push it ventrally to open the esophagus. During the procedure, the animal body is covered with blankets to avoid hypothermia. If the animal&#39;s body temperature decreases, use a heating system to stabilize the temperature on a physiological level (see the Table of Materials). The body temperature is displayed on the screen of the PiCCO.</li>
	</ol>
	</li>
</ol>
2. Instrumentation
<ol>
	<li>Use bandages to pull back the hind legs to smoothen the folds in the femoral area for vessel catheterization.</li>
	<li>Prepare the following materials: one 5 mL syringe, one 10 mL syringe, one 50 mL syringe, one Seldinger needle, introducer sheaths (2 mm, 2.7 mm, 2.7 mm), guidewires for the sheaths, a central venous catheter with three ports (2.3 mm, 30 cm) with guidewire, and a PiCCO catheter (1.67 mm, 20 cm).</li>
	<li>Disinfect the inguinal area with colored disinfection, wait for 2 min, and wipe the disinfection off with a sterile tissue. Repeat this procedure 3x. After the third time, do not remove the disinfection.</li>
	<li>Fill all catheters with saline solutions.</li>
	<li>Apply ultrasound gel to the ultrasound probe. Cover the inguinal area with a sterile fenestrated drape and scan the right femoral vessels with ultrasound. Use the Doppler technique to distinguish between the artery and the vein18.</li>
	<li>Bright red pulsating blood confirms the aspired needle position. Disconnect the syringe and insert the guidewire into the right femoral artery.</li>
	<li>Visualize the longitudinal axis of the right femoral vein and insert the Seldinger needle under permanent aspiration with the 5 mL syringe.</li>
	<li>Aspirate dark red nonpulsating venous blood.</li>
	<li>Visualize the right femoral artery axially and switch to a longitudinal view of the artery by rotating the probe 90&#176;.</li>
	<li>Puncture the right femoral artery under ultrasound visualization with the Seldinger needle under permanent aspiration with the 5 mL syringe. 
	NOTE: The ultrasound-guided Seldinger technique is associated with significantly lower blood loss, tissue trauma, and time consumption than other methods of vascular access19,20.
	<ol>
		<li>If the correct position of the needle in the different vessels cannot be established for certain, take the blood probes and analyze the blood gas content with a blood gas analyzer (see the Table of Materials). A high oxygen&#160;level is a good sign of arterial blood, and a low oxygen level is a sign of venous blood.</li>
	</ol>
	</li>
	<li>Insert the guidewire for the central venous catheter into the right femoral vein after disconnecting the syringe and retracting the Seldinger needle.</li>
	<li>Visualize both right vessels with ultrasound to control the correct wire position.</li>
	<li>Push the arterial introducer sheath (2 mm) over the guidewire into the right artery and secure the position with blood aspiration.</li>
	<li>Use the Seldinger technique to position the central venous line into the right femoral vein. Aspirate all ports and flush them with saline solution.</li>
	<li>Perform the same procedure on the left inguinal side to insert the other introducer sheaths in the Seldinger technique into the left femoral artery (2.7 mm) and femoral vein (2.7 mm).</li>
	<li>Connect the right arterial introducer sheath and the central venous catheter with two transducer systems for the measurement of invasive hemodynamics, and position both transducers on the heart level to get appropriate values.</li>
	<li>Switch the three-way-stopcocks of both transducers open to the atmosphere to calibrate the systems to 0 as is prescribed in the operation instructions. 
	NOTE: It is absolutely necessary to avoid any air bubbles and bloodstains in the systems to generate plausible values.</li>
	<li>Switch all infusions for maintaining anesthesia from the peripheral vein to the central venous line.</li>
	<li>Take baseline values (hemodynamics, spirometry, NIRS (see section 4) and PiCCO&#160;(see section 3) after 15 min of recovery.</li>
	<li>Initiate hemorrhagic shock (see section 5).</li>
</ol>
3. PiCCO Measurement
NOTE: For the PiCCO equipment, see the Table of Materials.
<ol>
	<li>Insert the PiCCO catheter into the right arterial introducer sheath. 
	NOTE: In clinical medicine, PiCCO catheters are directly placed by the Seldinger technique. However, placement via an introducer sheath is feasible as well.</li>
	<li>Connect the catheter with the arterial wire of the PiCCO system and the arterial transducer directly with the PiCCO port. Then, recalibrate as described in step 2.17.</li>
	<li>Connect the venous measuring unit of the PiCCO&#160;system with the left venous introducer sheath. 
	NOTE: It is necessary to connect the venous and arterial probes at some distance from each other. Otherwise, the measurement will be disturbed, because the application of cold saline solution into the venous system will influence the arterial measurement. For more details on PiCCO, see&#160;Mayer and Suttner21.</li>
	<li>Turn on the PiCCO system and confirm that a new patient is measured.</li>
	<li>Enter the animal&#39;s size and weight and switch the category to adults.</li>
	<li>Enter the protocol name and ID and enter Exit.</li>
	<li>Set the injection volume to 10 mL. 
	NOTE: The volume of the chosen injection solution can be varied. A higher volume makes the measured values more valid. Chose a small volume to avoid any hemodilution effects through repetitive application.</li>
	<li>Enter the central venous pressure.</li>
	<li>Open the three-way stopcock to the atmosphere, click on Zero for system calibration, and click on Exit.</li>
	<li>Calibrate the continuous cardiac output measurement as described next and click on TD (Thermodilution). Prepare physiological saline solution with a temperature of 4 &#176;C in a 10 mL syringe and click on Start.</li>
	<li>Inject 10 mL of the cold saline solution quickly and steadily into the venous measuring unit and wait until the measurement is completed and the system requests a repetition.</li>
	<li>Repeat this procedure until three measurements are completed.</li>
	<li>Let the system calculate the mean of all parameters and click on Exit.</li>
	<li>After complete calibration, immediately start the measurement. To monitor shock induction, focus on the PiCCO-derived parameter cardiac index.</li>
</ol>
4. Cerebral Regional Oxygenation Saturation
NOTE: For the equipment to monitor cerebral regional oxygenation, see the Table of Materials.
<ol>
	<li>Shave the forehead of the pig with a disposable razor and water and stick two self-adherent sensors (see the Table of Materials) for NIRS to the forehead of the pig.</li>
	<li>Connect the preamplifier to the monitor and connect the sensor cable connectors color-coded to the preamplifier.</li>
	<li>Close the preamp locking mechanism and attach the sensors to the sensor cables. 
	NOTE: In order to record real-time data, a USB flash drive has to be connected to the NIRS monitor.</li>
	<li>Switch on the monitor, click on New Patient, enter the study name, and click on Done.</li>
	<li>Check the incoming signal. When the signal is stable, click on Baseline Menu and click on Set Baselines. If the baseline has already been entered, confirm the new baseline by clicking Yes and click on Event Mark.</li>
	<li>Chose the event with the arrow buttons on the keyboard and with Next Event; select the event 3 Induction and press Select Event. 
	NOTE: If further information is necessary, consult the operation manual of the NIRS system22.</li>
</ol>
5. Hemorrhagic Shock Induction
<ol>
	<li>Connect left introducer sheath with a tree-way stopcock. Connect one port of the three-way stopcock with a 50 mL syringe and one with an empty infusion bottle. 
	NOTE: Alternatively, the withdrawn blood may be collected in citrated bags for later autotransfusion. This is a major advantage of controlled blood withdrawal.</li>
	<li>Measure and document the exact hemodynamic parameters and calculate 40% of the cardiac index and mean arterial pressure as hemodynamic targets. Set the event 93 Blood Loss in the NIRS system as described in step 4.6. 
	NOTE: Hemorrhagic shock is achieved if the cardiac index and mean arterial pressure drops below 40% of the baseline value. A considerable cerebral regional oxygenation saturation (crSO2) decline of 20% is preferable to depict microcirculatory impairment. The average blood loss to achieve this lies within a range of 25-35 mL&#183;kg-1.</li>
	<li>Aspirate 50 mL of blood into the syringe and switch the three-way stopcock. Push the blood into the empty bottle.</li>
	<li>Note the removed blood volume.</li>
	<li>Monitor the arterial blood pressure, the cardiac index, and the crSO2 closely. Repeat the blood withdrawal until the target blood pressure and cardiac index are achieved (after 20-30 min).</li>
	<li>Set the event 97 Hypotension in the NIRS device as described in step 4.6. 
	NOTE: Do not withdraw the blood too quickly, because this bears the risk of immediate cardio-circulatory failure. After finishing the shock induction procedure, the animals can be used for various therapeutic interventions.</li>
</ol>
6. End of the Experiment and Euthanasia
<ol>
	<li>Inject 0.5 mg of fentanyl into the central venous line and wait for 5 min.</li>
	<li>Inject 200 mg of propofol into the central venous line and euthanize the animal with 40 mmol potassium chloride.</li>
</ol>

The NIRS device was provided unconditionally by Medtronic PLC, USA, for experimental research purposes. Alexander Ziebart, Andreas Garcia-Bardon, and Erik K. Hartmann received instructor honoraria for physician training courses from Medtronic PLC. None of the authors report financial or other conflicts of interest.

The protocol describes one method of inducing hemorrhagic shock via controlled arterial bleeding in pigs that is guided by systemic hemodynamics, as well as by cerebral microcirculatory impairment. Shock conditions were achieved by a calculated blood withdrawal of 25-35 mL kg-1 and confirmed by the mentioned composite of surrogate parameters indicating considerable cardio-circulatory failure. If untreated, this procedure was lethal within 2 h in 66% of the animals, which underlines the severity and reproducibi...

Massive blood loss is among the main causes of injury-related deaths1,2,3. The loss of circulatory fluid and oxygen carriers leads to hemodynamic failure and severe oxygen undersupply and can cause irreversible organ failure and death. The severity level of shock is influenced by additional factors like hypothermia, coagulopathy, and acidosis4. Particularly the brain, but also the kidneys lack compensation capacity due to high oxygen demand and the incapability of adequate anaerobic energy generation5,6. For therapeutic purposes, fast and immediate action is pivotal. In clinical practice, fluid resuscitation with a balanced electrolyte solution is the first option for treatment, followed by the administration of red blood cell concentrates and fresh frozen plasma. Thrombocyte concentrates, catecholamines, and the optimization of coagulation and the acid-base status support the therapy to regain normal physiological conditions after sustained trauma. This concept focuses on the restoration of hemodynamics and macrocirculation. Several studies, however, show that microcirculatory perfusion does not recover simultaneously with the macrocirculation. Especially, cerebral perfusion remains impaired and further oxygen undersupply may occur7,8.
The use of animal models allows scientists to establish novel or experimental strategies. The comparable anatomy, homology, and physiology of pigs and humans enable conclusions on specific pathological factors. Both species have a similar metabolic system and response to pharmacologic treatments. This is a great advantage in comparison to small animal&#160;models where differences in blood volume, hemodynamics, and overall physiology make it almost impossible to mimic a clinical scenario9. Furthermore, authorized medical equipment and consumables can be easily used in porcine models. Additionally, it is easily possible to obtain pigs from commercial suppliers, which allows a high diversity of genetics and phenotypes and is cost reducing10. The model of blood withdrawal via vessel cannulation is quite common11,12,13,14,15.
In this study, we extend the concept of hemorrhagic shock induction via arterial blood withdrawal with an exact titration of hemodynamic failure and cerebral oxygenation impairment. Hemorrhagic shock is achieved if the cardiac index and mean arterial pressure drops below 40% of the baseline value, which has been shown to cause considerable deterioration of the cerebral regional oxygenation saturation8. Pulse contour cardiac output (PiCCO) measurement is used for continuous hemodynamic monitoring. First, the system has to be calibrated by transpulmonary thermodilution, which enables the calculation of the cardiac index of the extravascular lung water content and the global end-diastolic volume. Subsequently, the continuous cardiac index is calculated by pulse contour analysis and also provides dynamic preload parameters like pulse pressure and stroke volume variation.
This technique is well established in clinical and experimental settings.Near-infrared spectroscopy (NIRS) is a clinically and experimentally established method to monitor changes in cerebral oxygen supply in real-time. Self-adherent sensors are attached to the left and right forehead and calculate the cerebral oxygenation non-invasively in the cerebral frontal cortex. Two wavelengths of infrared light (700 and 900 nm) are emitted and detected by the sensors after being reflected from the cortex tissue. To assess the cerebral oxygen content, contributions of arterial and venous blood are calculated in 1:3 relations and updated in 5 s intervals. The sensitivity in depth of 1-4 cm is exponential decreasing and influenced by the penetrated tissue (e.g., skin and bone), although the skull is translucent to infrared light. The technique facilitates quick therapeutic actions to prevent patients from adverse outcomes like delirium or hypoxic cerebral injury and serves as the target parameter in case of impaired cardiac output16,17. The combination of both techniques during experimental shock enables an exact titration of macrocirculation, as well as cerebral microcirculatory impairment, to study this life-threatening event.

<table><tbody><tr><td>3-way-stopcock blue</td><td>Becton Dickinson Infusion Therapy AB Helsingborg, Sweden</td><td>394602</td><td>Drug administration</td></tr><tr><td>3-way-stopcock red</td><td>Becton Dickinson Infusion Therapy AB Helsingborg, Sweden</td><td>394605</td><td>Drug administration/Shock induction</td></tr><tr><td>Atracurium</td><td>Hikma Pharma GmbH , Martinsried</td><td>AM03AC04*</td><td>Anesthesia</td></tr><tr><td>Canula 20 G</td><td>Becton Dickinson S.A. Carretera Mequinenza Fraga, Spain</td><td>301300</td><td>Vascular access</td></tr><tr><td>Datex Ohmeda S5</td><td>GE Healthcare Finland Oy, Helsinki, Finland</td><td>-</td><td>Hemodynamic monitor</td></tr><tr><td>Desinfection&amp;nbsp;</td><td>Sch&amp;uuml;lke &amp;amp; Mayr GmbH, Germany</td><td>104802</td><td>Desinfection&amp;nbsp;</td></tr><tr><td>Heidelberger Verl&amp;auml;ngerung 75CM</td><td>Fresenius Kabi Deutschland GmbH</td><td>2873112 &amp;nbsp;</td><td>Drug administration/Shock induction</td></tr><tr><td>INVOS 5100C Cerebral</td><td>Medtronic PLC, USA</td><td>-</td><td>Monitore for cerebral regional oxygenation&amp;nbsp;</td></tr><tr><td>INVOS Cerebral/Somatic Oximetry Adult Sensors</td><td>Medtronic PLC, USA</td><td>20884521211152</td><td>Monitoring of the cerebral regional oxygenation&amp;nbsp;</td></tr><tr><td>Endotracheal tube</td><td>Teleflex Medical Sdn. Bhd, Malaysia</td><td>112482</td><td>Intubation</td></tr><tr><td>Endotracheal tube introducer</td><td>&amp;nbsp; Wirutec GmbH, Sulzbach, Germany</td><td>5033062</td><td>Intubation</td></tr><tr><td>Engstr&amp;ouml;m Carestation</td><td>GE Heathcare, Madison USA</td><td>-</td><td>Ventilator</td></tr><tr><td>Fentanyl</td><td>Janssen-Cilag GmbH, Neuss</td><td>AA0014*</td><td>Anesthesia</td></tr><tr><td>Gloves</td><td>Paul Hartmann, Heidenheim, Germany</td><td>9422131</td><td>Self-protection</td></tr><tr><td>Incetomat-line 150 cm</td><td>Fresenius, Kabi GmbH, Bad Homburg, Germany</td><td>9004112</td><td>Drug administration</td></tr><tr><td>Ketamine</td><td>Hameln Pharmaceuticals GmbH, Zofingen, Schweiz</td><td>AN01AX03*</td><td>Sedation</td></tr><tr><td>Laryngoscope</td><td>Teleflex Medical Sdn. Bhd, Malaysia</td><td>671067-000020</td><td>Intubation</td></tr><tr><td>Logical pressure monitoring system</td><td>Smith- Medical GmbH,&amp;nbsp; Minneapolis, USA</td><td>MX9606</td><td>Hemodynamic monitor</td></tr><tr><td>Logicath 7 Fr 3-lumen 30 cm</td><td>Smith- Medical GmbH,&amp;nbsp; Minneapolis, USA</td><td>MXA233x30x70-E</td><td>Vascular access/Drug administration</td></tr><tr><td>Masimo Radical 7</td><td>Masimo Corporation, Irvine, USA</td><td>-</td><td>Hemodynamic monitor</td></tr><tr><td>Mask for ventilating dogs</td><td>Henry Schein, Melville, USA</td><td>730-246</td><td>Ventilation</td></tr><tr><td>Original Perfusor syringe 50 mL Luer Lock</td><td>B.Braun Melsungen AG, Melsungen, Germany</td><td>8728810F</td><td>Drug administration</td></tr><tr><td>PICCO Thermodilution. F5/20CM EW&amp;nbsp;</td><td>MAQUET Cardiovascular GmbH, Rastatt, Germany</td><td>PV2015L20-A &amp;nbsp;</td><td>Hemodynamic monitor</td></tr><tr><td>Percutaneous sheath introducer set 8,5 und 9 Fr, 10 cm with integral haemostasis valve/sideport</td><td>Arrow international inc., Reading, USA</td><td>AK-07903</td><td>Vascular access/Shock induction</td></tr><tr><td>Perfusor FM Braun</td><td>B.Braun Melsungen AG, Melsungen, Germany</td><td>8713820</td><td>Drug administration</td></tr><tr><td>Potassium chloride</td><td>Fresenius, Kabi GmbH, Bad Homburg, Germany</td><td>6178549</td><td>Euthanasia</td></tr><tr><td>Propofol 2%</td><td>Fresenius, Kabi GmbH, Bad Homburg, Germany</td><td>&amp;nbsp; AN01AX10*</td><td>Anesthesia</td></tr><tr><td>&amp;nbsp;Pulse Contour Cardiac Output (PiCCO&lt;sub&gt;2&lt;/sub&gt;)&amp;nbsp;</td><td>Pulsion Medical Systems, Feldkirchen, Germany</td><td>-</td><td>Hemodynamic monitor</td></tr><tr><td>Sonosite Micromaxx Ultrasoundsystem</td><td>Fujifilm, Sonosite Bothell, Bothell, USA&amp;nbsp;</td><td>-</td><td>Vascular access</td></tr><tr><td>Stainless Macintosh Size 4</td><td>Teleflex Medical Sdn. Bhd, Perak,&amp;nbsp; Malaysia</td><td>670000</td><td>Intubation</td></tr><tr><td>Sterofundin</td><td>B.Braun Melsungen AG, Melsungen, Germany</td><td>AB05BB01*</td><td>balanced electrolyte infusion</td></tr><tr><td>Stresnil 40 mg/mL&amp;nbsp;&amp;nbsp;</td><td>Lilly Germany GmbH, Wiesbaden, Germany</td><td>QN05AD90</td><td>Sedation</td></tr><tr><td>Syringe 10 mL</td><td>Becton Dickinson S.A. Carretera Mequinenza Fraga, Spain</td><td>309110</td><td>Drug administration</td></tr><tr><td>Syringe 2 mL</td><td>Becton Dickinson S.A. Carretera Mequinenza Fraga, Spain</td><td>300928</td><td>Drug administration</td></tr><tr><td>Syringe 20 mL</td><td>Becton Dickinson S.A. Carretera Mequinenza Fraga, Spain</td><td>300296</td><td>Drug administration</td></tr><tr><td>Syringe 5 mL</td><td>Becton Dickinson S.A. Carretera Mequinenza Fraga, Spain</td><td>309050</td><td>Drug administration</td></tr><tr><td>Venous catheter 22 G</td><td>B.Braun Melsungen AG, Melsungen, Germany</td><td>4269110S-01</td><td>Vascular access</td></tr><tr><td>*ATC:&amp;nbsp; Anatomical Therapeutic Chemical / Defined Daily Dose Classification&amp;nbsp;</td><td></td><td></td><td></td></tr></tbody></table>

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.

After starting the shock induction, a short time of compensation can be registered. With ongoing blood removal, the aforementioned cardio-circulatory decompensation, as monitored by a significant decrease of crSO2, the cardiac index, the intrathoracic blood volume index, and the global end-diastolic volume index (Figure 2, Figure 3, and Figure 4), occurs. Furthermore, significant tachycard...

Watch this Scientific Journal Video about Standardized Hemorrhagic Shock Induction Guided by Cerebral Oximetry and Extended Hemodynamic Monitoring in Pigs at JoVE.com

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

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 ...

standardized-hemorrhagic-shock-induction-guided-cerebral-oximetry

Research

JoVE Journal

Medicine

7.3K Views.  University Medical Center of the Johannes Gutenberg-University. 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. 

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

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, ...

A Piglet Model of Neonatal Hypoxic-Ischemic Encephalopathy

Birth asphyxia, which causes hypoxic-ischemic encephalopathy (HIE), accounts for 0.66 million deaths worldwide each year, about a quarter of the world&#8217;s 2.9 million neonatal deaths. Animal models of HIE have contributed to the understanding of the pathophysiology in HIE, and have highlighted the dynamic process that occur in brain injury due to perinatal asphyxia. Thus, animal studies have suggested a time-window for post-insult treatment strategies. Hypothermia has been tested as a treatment for HIE in pdiglet models and subsequently proven effective in clinical trials. Variations of the model have been applied in the study of adjunctive neuroprotective methods and piglet studies of xenon and melatonin have led to clinical phase I and II trials1,2. The piglet HIE model is further used for neonatal resuscitation- and hemodynamic studies as well as in investigations of cerebral hypoxia on a cellular level. However, it is a technically challenging model and variations in the protocol may result in either too mild or too severe brain injury. In this article, we demonstrate the technical procedures necessary for establishing a stable piglet model of neonatal HIE. First, the newborn piglet (&#60; 24 hr old, median weight 1500 g) is anesthetized, intubated, and monitored in a setup comparable to that found in a neonatal intensive care unit. Global hypoxia-ischemia is induced by lowering the inspiratory oxygen fraction to achieve global hypoxia, ischemia through hypotension and a flat trace amplitude integrated EEG (aEEG) indicative of cerebral hypoxia. Survival is promoted by adjusting oxygenation according to the aEEG response and blood pressure. Brain injury is quantified by histopathology and magnetic resonance imaging after 72 hr.

Hypoxic-ischemic encephalopathy following perinatal asphyxia can be studied using animal models. We demonstrate the procedures necessary for establishing a piglet model of neonatal hypoxic-ischemic encephalopathy.

Birth asphyxia, which causes hypoxic-ischemic encephalopathy (HIE), accounts for 0.66 million deaths worldwide each year, about a quarter of the ...

Invasive Hemodynamic Monitoring of Aortic and Pulmonary Artery Hemodynamics in a Large Animal Model of ARDS

One of the leading causes of morbidity and mortality in patients with heart failure is right ventricular (RV) dysfunction, especially if it is due to pulmonary hypertension. For a better understanding and treatment of this disease, precise hemodynamic monitoring of left and right ventricular parameters is important. For this reason, it is essential to establish experimental pig models of cardiac hemodynamics and measurements for research purpose. 
This article shows the induction of ARDS by using oleic acid (OA) and consequent right ventricular dysfunction, as well as the instrumentation of the pigs and the data acquisition process that is needed to assess hemodynamic parameters. To achieve right ventricular dysfunction, we used oleic acid (OA) to cause ARDS and accompanied this with pulmonary artery hypertension (PAH). With this model of PAH and consecutive right ventricular dysfunction, many hemodynamic parameters can be measured, and right ventricular volume load can be detected. 
All vital parameters, including respiratory rate (RR), heart rate (HR) and body temperature were recorded throughout the whole experiment. Hemodynamic parameters including femoral artery pressure (FAP), aortic pressure (AP), right ventricular pressure (peak systolic, end systolic and end diastolic right ventricular pressure), central venous pressure (CVP), pulmonary artery pressure (PAP) and left arterial pressure (LAP) were measured as well as perfusion parameters including ascending aortic flow (AAF) and pulmonary artery flow (PAF). Hemodynamic measurements were performed using transcardiopulmonary thermodilution to provide cardiac output (CO). Furthermore, the PiCCO2 system (Pulse Contour Cardiac Output System 2) was used to receive parameters such as stroke volume variance (SVV), pulse pressure variance (PPV), as well as extravascular lung water (EVLW) and global end-diastolic volume (GEDV). Our monitoring procedure is suitable for detecting right ventricular dysfunction and monitoring hemodynamic findings before and after volume administration.

We present a protocol of creating right ventricular dysfunction in a pig model by inducing ARDS. We demonstrate invasive monitoring of left and right ventricular cardiac output using flow probes around the aorta and the pulmonary artery, as well as blood pressure measurements in the aorta and pulmonary artery.

One of the leading causes of morbidity and mortality in patients with heart failure is right ventricular (RV) dysfunction, especially if it is due to ...

Noninvasive and Invasive Renal Hypoxia Monitoring in a Porcine Model of Hemorrhagic Shock

Up to 50% of patients with trauma develop acute kidney injury (AKI), in part due to poor renal perfusion after severe blood loss. AKI is currently diagnosed based on a change in serum creatinine concentration from baseline or prolonged periods of decreased urine output. Unfortunately, baseline serum creatinine concentration data is unavailable in most patients with trauma, and current estimation methods are inaccurate. In addition, serum creatinine concentration may not change until 24-48 h after the injury. Lastly, oliguria must persist for a minimum of 6 h to diagnose AKI, making it impractical for early diagnosis. AKI diagnostic approaches available today are not useful for predicting risk during the resuscitation of patients with trauma. Studies suggest that urinary partial pressure of oxygen (PuO2) may be useful for assessing renal hypoxia. A monitor that connects the urinary catheter and the urine collection bag was developed to measure PuO2 noninvasively. The device incorporates an optical oxygen sensor that estimates PuO2 based on luminescence quenching principles. In addition, the device measures urinary flow and temperature, the latter to adjust for confounding effects of temperature changes. Urinary flow is measured to compensate for the effects of oxygen ingress during periods of low urine flow. This article describes a porcine model of hemorrhagic shock to study the relationship between noninvasive PuO2, renal hypoxia, and AKI development. A key element of the model is the ultrasound-guided surgical placement in the renal medulla of an oxygen probe, which is based on an unsheathed optical microfiber. PuO2 will also be measured in the bladder and compared to the kidney and noninvasive PuO2 measurements. This model can be used to test PuO2 as an early marker of AKI and assess PuO2 as a resuscitative endpoint after hemorrhage that is indicative of end-organ rather than systemic oxygenation.

Presented here is a protocol to measure renal oxygenation in the medulla and noninvasive urine oxygen partial pressure in a hemorrhagic shock porcine model to establish urine oxygen partial pressure as an early indicator of acute kidney injury (AKI) and a novel resuscitative endpoint.

Up to 50% of patients with trauma develop acute kidney injury (AKI), in part due to poor renal perfusion after severe blood loss. AKI is currently ...

Bioengineering

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 ...

Real-Time Assessment of Spinal Cord Microperfusion in a Porcine Model of Ischemia/Reperfusion

Spinal cord injury is a devastating complication of aortic repair. Despite developments for the prevention and treatment of spinal cord injury, its incidence is still considerably high and therefore, influences patient outcome. Microcirculation plays a key role in tissue perfusion and oxygen supply and is often dissociated from macrohemodynamics. Thus, direct evaluation of spinal cord microcirculation is essential for the development of microcirculation-targeted therapies and the evaluation of existing approaches in regard to spinal cord microcirculation. However, most of the methods do not provide real-time assessment of spinal cord microcirculation. The aim of this study is to describe a standardized protocol for real-time spinal cord microcirculatory evaluation using laser-Doppler needle probes directly inserted in the spinal cord. We used a porcine model of ischemia/reperfusion to induce deterioration of the spinal cord microcirculation. In addition, a fluorescent microsphere injection technique was used. Initially, animals were anesthetized and mechanically ventilated. Thereafter, laser-Doppler needle probe insertion was performed, followed by the placement of cerebrospinal fluid drainage. A median sternotomy was performed for exposure of the descending aorta to perform aortic cross-clamping. Ischemia/reperfusion was induced by supra-celiac aortic cross-clamping for a total of 48 min, followed by reperfusion and hemodynamic stabilization. Laser-Doppler Flux was performed in parallel with macrohemodynamic evaluation. In addition, automated cerebrospinal fluid drainage was used to maintain a stable cerebrospinal pressure. After completion of the protocol, animals were sacrificed, and the spinal cord was harvested for histopathological and microsphere analysis. The protocol reveals the feasibility of spinal cord microperfusion measurements using laser-Doppler probes and shows a marked decrease during ischemia as well as recovery after reperfusion. Results showed comparable behavior to fluorescent microsphere evaluation. In conclusion, this new protocol might provide a useful large animal model for future studies using real-time spinal cord microperfusion assessment in ischemia/reperfusion conditions.

Spinal cord microcirculation plays a pivotal role in spinal cord injury. Most methods do not allow real-time assessment of spinal cord microcirculation, which is essential for the development of microcirculation-targeted therapies. Here, we propose a protocol using Laser-Doppler-Flow Needle probes in a large animal model of ischemia/reperfusion.

Spinal cord injury is a devastating complication of aortic repair. Despite developments for the prevention and treatment of spinal cord injury, its ...

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 ...

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 ...