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

Summary

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.

Abstract

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 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.

Introduction

Massive blood loss is among the main causes of injury-related deaths^1,2,3. 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 generation^5,6. 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 occur^7,8.

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 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 common^{11,12,13,14,15}.

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 output^16,17. 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.

Protocol

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 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 ± 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

1. Sedate pigs with a combined injection of ketamine (4 mg·kg^-1) and azaperone (8 mg·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.
2. 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.
3. Monitor the peripheral oxygen saturation (SpO₂) with a sensor clipped to the pig's tail or ear directly after arrival.
4. 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 µg·kg^-1) and propofol (3 mg·kg^-1).
5. 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.
6. 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 <20 cm H₂O, positive end-expiratory pressure (PEEP) = 5 cm H₂O.
7. Maintain anesthesia via a continuous infusion of fentanyl (0.1-0.2 µg·kg^-1·h^-1) and propofol (8-12 mg·kg^-1·h^-1) and start an infusion of balanced electrolyte solution (5 mL·kg^-1·h^-1).
8. Facilitate endotracheal intubation by the application of a muscle relaxant (atracurium 0.5 mg·kg^-1).
9. 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.
   1. Person 1: Fix the tongue outside with a piece of tissue and open the snout with the other hand.
   2. Person 2: Perform a laryngoscopy.
   3. 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.
10. 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.
11. 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).
12. Start mechanical ventilation after the tube is connected to a ventilator (PEEP = 5 cm H₂O; tidal volume = 8 mL·kg^-1; FiO₂ = 0.4; inspiration-to-expiration ratio = 1:2; respiratory rate = variable to achieve an end-tidal CO₂ of <6 kPa).
    NOTE: Avoid fluctuation of the CO₂ to minimize any respiratory effects on the cerebral perfusion.
13. 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.
14. With two persons, place a gastric tube into the stomach to avoid reflux and vomiting.
    1. Person 1: Fix the tongue outside with a piece of tissue and open the snout with the other hand.
    2. Person 2: Perform a laryngoscopy of the porcine larynx.
    3. Person 2: Visualize the esophagus.
    4. 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'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.

2. Instrumentation

1. Use bandages to pull back the hind legs to smoothen the folds in the femoral area for vessel catheterization.
2. 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).
3. 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.
4. Fill all catheters with saline solutions.
5. 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.
6. Bright red pulsating blood confirms the aspired needle position. Disconnect the syringe and insert the guidewire into the right femoral artery.
7. Visualize the longitudinal axis of the right femoral vein and insert the Seldinger needle under permanent aspiration with the 5 mL syringe.
8. Aspirate dark red nonpulsating venous blood.
9. Visualize the right femoral artery axially and switch to a longitudinal view of the artery by rotating the probe 90°.
10. 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 access^19,20.
    1. 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 level is a good sign of arterial blood, and a low oxygen level is a sign of venous blood.
11. Insert the guidewire for the central venous catheter into the right femoral vein after disconnecting the syringe and retracting the Seldinger needle.
12. Visualize both right vessels with ultrasound to control the correct wire position.
13. Push the arterial introducer sheath (2 mm) over the guidewire into the right artery and secure the position with blood aspiration.
14. Use the Seldinger technique to position the central venous line into the right femoral vein. Aspirate all ports and flush them with saline solution.
15. 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).
16. 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.
17. 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.
18. Switch all infusions for maintaining anesthesia from the peripheral vein to the central venous line.
19. Take baseline values (hemodynamics, spirometry, NIRS (see section 4) and PiCCO (see section 3) after 15 min of recovery.
20. Initiate hemorrhagic shock (see section 5).

3. PiCCO Measurement

NOTE: For the PiCCO equipment, see the Table of Materials.

1. 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.
2. 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.
3. Connect the venous measuring unit of the PiCCO 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 Mayer and Suttner²¹.
4. Turn on the PiCCO system and confirm that a new patient is measured.
5. Enter the animal's size and weight and switch the category to adults.
6. Enter the protocol name and ID and enter Exit.
7. 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.
8. Enter the central venous pressure.
9. Open the three-way stopcock to the atmosphere, click on Zero for system calibration, and click on Exit.
10. Calibrate the continuous cardiac output measurement as described next and click on TD (Thermodilution). Prepare physiological saline solution with a temperature of 4 °C in a 10 mL syringe and click on Start.
11. 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.
12. Repeat this procedure until three measurements are completed.
13. Let the system calculate the mean of all parameters and click on Exit.
14. After complete calibration, immediately start the measurement. To monitor shock induction, focus on the PiCCO-derived parameter cardiac index.

4. Cerebral Regional Oxygenation Saturation

NOTE: For the equipment to monitor cerebral regional oxygenation, see the Table of Materials.

1. 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.
2. Connect the preamplifier to the monitor and connect the sensor cable connectors color-coded to the preamplifier.
3. 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.
4. Switch on the monitor, click on New Patient, enter the study name, and click on Done.
5. 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.
6. 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.

5. Hemorrhagic Shock Induction

1. 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.
2. 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·kg^-1.
3. Aspirate 50 mL of blood into the syringe and switch the three-way stopcock. Push the blood into the empty bottle.
4. Note the removed blood volume.
5. 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).
6. 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.

6. End of the Experiment and Euthanasia

1. Inject 0.5 mg of fentanyl into the central venous line and wait for 5 min.
2. Inject 200 mg of propofol into the central venous line and euthanize the animal with 40 mmol potassium chloride.

Results

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...

Discussion

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...

Materials

Name	Company	Catalog Number	Comments
3-way-stopcock blue	Becton Dickinson Infusion Therapy AB Helsingborg, Sweden	394602	Drug administration
3-way-stopcock red	Becton Dickinson Infusion Therapy AB Helsingborg, Sweden	394605	Drug administration/Shock induction
Atracurium	Hikma Pharma GmbH , Martinsried	AM03AC04*	Anesthesia
Canula 20 G	Becton Dickinson S.A. Carretera Mequinenza Fraga, Spain	301300	Vascular access
Datex Ohmeda S5	GE Healthcare Finland Oy, Helsinki, Finland	-	Hemodynamic monitor
Desinfection	Schülke & Mayr GmbH, Germany	104802	Desinfection
Heidelberger Verlängerung 75CM	Fresenius Kabi Deutschland GmbH	2873112	Drug administration/Shock induction
INVOS 5100C Cerebral	Medtronic PLC, USA	-	Monitore for cerebral regional oxygenation
INVOS Cerebral/Somatic Oximetry Adult Sensors	Medtronic PLC, USA	20884521211152	Monitoring of the cerebral regional oxygenation
Endotracheal tube	Teleflex Medical Sdn. Bhd, Malaysia	112482	Intubation
Endotracheal tube introducer	Wirutec GmbH, Sulzbach, Germany	5033062	Intubation
Engström Carestation	GE Heathcare, Madison USA	-	Ventilator
Fentanyl	Janssen-Cilag GmbH, Neuss	AA0014*	Anesthesia
Gloves	Paul Hartmann, Heidenheim, Germany	9422131	Self-protection
Incetomat-line 150 cm	Fresenius, Kabi GmbH, Bad Homburg, Germany	9004112	Drug administration
Ketamine	Hameln Pharmaceuticals GmbH, Zofingen, Schweiz	AN01AX03*	Sedation
Laryngoscope	Teleflex Medical Sdn. Bhd, Malaysia	671067-000020	Intubation
Logical pressure monitoring system	Smith- Medical GmbH,  Minneapolis, USA	MX9606	Hemodynamic monitor
Logicath 7 Fr 3-lumen 30cm	Smith- Medical GmbH,  Minneapolis, USA	MXA233x30x70-E	Vascular access/Drug administration
Masimo Radical 7	Masimo Corporation, Irvine, USA	-	Hemodynamic monitor
Mask for ventilating dogs	Henry Schein, Melville, USA	730-246	Ventilation
Original Perfusor syringe 50ml Luer Lock	B.Braun Melsungen AG, Melsungen, Germany	8728810F	Drug administration
PICCO Thermodilution. F5/20CM EW	MAQUET Cardiovascular GmbH, Rastatt, Germany	PV2015L20-A	Hemodynamic monitor
Percutaneous sheath introducer set 8,5 und 9 Fr, 10 cm with integral haemostasis valve/sideport	Arrow international inc., Reading, USA	AK-07903	Vascular access/Shock induction
Perfusor FM Braun	B.Braun Melsungen AG, Melsungen, Germany	8713820	Drug administration
Potassium chloride	Fresenius, Kabi GmbH, Bad Homburg, Germany	6178549	Euthanasia
Propofol 2%	Fresenius, Kabi GmbH, Bad Homburg, Germany	AN01AX10*	Anesthesia
Pulse Contour Cardiac Output (PiCCO2)	Pulsion Medical Systems, Feldkirchen, Germany	-	Hemodynamic monitor
Sonosite Micromaxx Ultrasoundsystem	Fujifilm, Sonosite Bothell, Bothell, USA	-	Vascular access
Stainless Macintosh Size 4	Teleflex Medical Sdn. Bhd, Perak,  Malaysia	670000	Intubation
Sterofundin	B.Braun Melsungen AG, Melsungen, Germany	AB05BB01*	balanced electrolyte infusion
Stresnil 40mg/ml	Lilly Germany GmbH, Wiesbaden, Germany	QN05AD90	Sedation
Syringe 10 mL	Becton Dickinson S.A. Carretera Mequinenza Fraga, Spain	309110	Drug administration
Syringe 2 mL	Becton Dickinson S.A. Carretera Mequinenza Fraga, Spain	300928	Drug administration
Syringe 20 mL	Becton Dickinson S.A. Carretera Mequinenza Fraga, Spain	300296	Drug administration
Syringe 5 mL	Becton Dickinson S.A. Carretera Mequinenza Fraga, Spain	309050	Drug administration
venous catheter 22G	B.Braun Melsungen AG, Melsungen, Germany	4269110S-01	Vascular access
*ATC:  Anatomical Therapeutic Chemical / Defined Daily Dose Classification