The authors would like to thank Lena Brix, V.M.D, Institute of Animal Research, Hannover Medical School, as well as Mrs. Jutta Dammann, Facility of Research Animal Care, University Medical Center Hamburg-Eppendorf, Germany, for providing pre- and perioperative animal care and their technical assistance on animal handling. The authors would further like to thank Dr. Daniel Manzoni, Department of Vascular Surgery, H&#244;pital Kirchberg, Luxembourg, for his technical assistance.

The study was approved by the Governmental Commission on the Care and Use of Animals of the City of Hamburg (Reference-No. 60/17). The animals received care in compliance with the &#39;Guide for the Care and Use of Laboratory Animals&#39; (NIH publication No. 86-23, revised 2011) as well as FELASA recommendations and experiments were carried out according to the ARRIVE guidelines24,25. This study was an acute trial, and all animals were euthanized at the end of protocol.
NOTE: The study was performed in six three-month-old male and female pigs (German Landrace) weighing approximately 40 kg. Animals were brought to the animal care facilities at least 7 days prior to the experiments and were housed in accordance to animal welfare recommendations. Animals were provided food and water ad libitum, and their health status was regularly assessed by the responsible veterinarian. A fasting time of 12 h was maintained prior to the experiments. The entire experimental procedure and handling of the animals was supervised by the responsible veterinarian.
1. Anesthesia induction and maintenance of anesthesia
<ol>
	<li>For anesthesia induction and maintenance of anesthesia, premedicate the animals and deeply sedate them using an intramuscular injection followed by intravenous injections, if necessary, to perform endotracheal intubation. Thereafter, induce and maintain anesthesia by using a combination of a volatile anesthesia agent with a continuous opioid application complemented with an additional opioid bolus injection.</li>
	<li>Perform intramuscular injections of ketamine 20 mg&#183;kg-1, azaperone 4 mg&#183;kg-1, and midazolam 0.1 mg&#183;kg-1 for premedication and sedation.</li>
	<li>Place a venous catheter in an ear vein, secure proper fixation, and assess functionality by fast application of 10 mL of saline.</li>
	<li>Place the animal in a supine position on a warming blanket to prevent heat loss.</li>
	<li>Establish basic monitoring with electrocardiography (ECG) and pulse oximetry to monitor the cardio-pulmonary state of the animals, and connect it to the basic monitoring hardware.</li>
	<li>Administer 15 L&#183;min-1 of oxygen via a pig-shaped mask for preoxygenation.</li>
	<li>Inject intravenous boli of 0.1 mg&#183;kg-1 of 1% propofol, if necessary, and perform endotracheal intubation.</li>
	<li>Secure correct placement with end tidal capnography and auscultation, administer 0.1 mg&#8226;kg-1&#160;of pancuronium, and ensure proper fixation of the endotracheal tube.</li>
	<li>Establish volume-controlled ventilation using tidal volumes of 10 mL&#183;kg-1 bodyweight-1, a positive end-expiratory pressure of 10 cmH2O, and a fraction of inspired oxygen (FiO2)of 0.3 using the anesthesia machine. Adjust the ventilator frequency to maintain an end-expiratory carbon dioxide tension (etCO2) of 35-45 mmHg.</li>
	<li>Introduce a gastric tube, perform suction of gastric fluids, properly fix the tube, and connect it to a collection bag. Carefully close the animal&#39;s eyes to prevent dryness of eyes during anesthesia.</li>
	<li>Maintain anesthesia by continuous infusion of fentanyl (10 &#181;g&#183;kg-1&#183;h-1) and sevoflurane (3.0% expired concentration, delivered by the vapor). Ensure adequate level of anesthesia by careful observation of vital signs and ventilation parameters as well as by absence of any movements during the entire protocol, paying special attention to the phases of surgical stimulus. Give additional bolus doses of fentanyl (50 &#181;g) if there is any indication of pain or distress. 
	NOTE: Ensure the presence of researchers who are experienced in animal anesthesia during the entire procedure, and use supervision by an experienced veterinarian to secure proper anesthesia.</li>
	<li>Administer a baseline infusion rate of 10 mL&#183;kg-1&#183;h-1 balanced crystalloids to compensate for fluid losses during anesthesia, surgical preparation, and execution of the experimental protocol. Use a fluid warmer to prevent heat loss.</li>
	<li>Gently clean the pig&#39;s skin using soap water. Use a skin disinfection solution containing povidone-iodine to decrease skin contamination. Use sterile gloves for surgical preparations. Apply 300 mg of clindamycin as antimicrobial prophylaxis, and repeat the dosage after 6 h.</li>
</ol>
2. Probe placement
<ol>
	<li>Place the animal in the right lateral position, and flex the animal&#39;s back to widen the space between the vertebrae.</li>
	<li>Surgically expose the paravertebral area for the preparation of spinous processes and vertebral arches (Figure 1A).</li>
	<li>Place a vascular 14 G peripheral vein catheter paramedian into the spinal cord at the level of thoracic vertebra (Th) 13/14 or lumbar vertebra (L) 1/2 between two vertebral arches (Figure 1B).</li>
	<li>Remove the needle, insert the laser/Doppler needle probe over the vein catheter (Figure 1C), and test the signal quality by connection to the designated hard- and software. Ensure that there is a stable signal with moderate pulsatility.</li>
	<li>Carefully fix the probe with sutures (Figure 1D) and use padding to prevent dislocation or kinking of the probe.</li>
	<li>For percutaneous placement of cerebrospinal fluid drainage for measuring and controlling cerebrospinal pressure, identify the level of L 4/5 or L 5/6, puncture the skin and the subcutaneous space with the introducer needle, and remove the inlay needle.</li>
	<li>Place a saline-filled syringe on the needle, and carefully introduce the needle with constant pressure on the fluid-filled syringe.</li>
	<li>Once a loss of resistance is felt as evidence for epidural position, re-introduce the inlay needle, and introduce the needle 2-3 mm further to puncture the dura mater and remove the inlay needle.</li>
	<li>Verify intrathecal position by fast dripping of clear liquor. Introduce the drainage up to 20 cm depth, attach the Luer-lock adapter, and verify the position by careful aspiration of liquor.</li>
	<li>Carefully fix the drainage with sutures, and connect it to the cerebrospinal fluid drainage system.</li>
	<li>Expose the skull behind the left ear, and carefully perform a drill hole trepanation of the skin using a 6 mm drill attachment.</li>
	<li>Introduce a second laser doppler probe directly into the brain. Carefully fix the probe with sutures, and test the signal quality by connection to designated hard- and software. Again, make sure that there is a stable signal with moderate pulsatility.</li>
	<li>Disconnect all probes, carefully place the animal in a supine position, ensuring unaffected probe position. Ensure that at least 4-5 researchers perform this maneuver.</li>
	<li>Reconnect the probes, and re-check signal quality.</li>
	<li>Connect the output channels of the laser-Doppler hardware to the amplifier and synchronic acquisition hardware and software to additionally record laser/Doppler Flux simultaneously with macrohemodynamic signals.</li>
	<li>Calibrate Flux as per unit (PU) with 2-point calibration.
	<ol>
		<li>Press Enter to open the menu and select the analogue output setting.</li>
		<li>Use the displayed conversion factor (5.0 V = 1000 PU) to calibrate Flux with 2-point calibration for use with the synchronic acquisition software.</li>
		<li>Select Return to return to the previous menu, and select Measurement to continue with measurement.</li>
		<li>Open the synchronic acquisition software. Select zero all inputs from the Setup menu. Connect all inputs with the used devices and probes.</li>
		<li>Perform 2-point calibration for Flux by clicking on the dropdown menu of the Flux channel. Select 2-point calibration. Set units-conversion to on and select BPU as units. For point 1, set 0 V to 0 BPU. For point 2, set 5.0 V to 1000 BPU. Select set units for all and new data. Press OK to close the menu.</li>
	</ol>
	</li>
	<li>Start continuous cerebrospinal fluid drainage with a target pressure of 10 mmHg and drainage volume of 20 mL&#183;h-1.</li>
</ol>
3. Catheter placement
<ol>
	<li>Expose both femoral arteries.</li>
	<li>Ligate the distal part of the right femoral artery, temporarily occlude the proximal lumen of the artery using a vessel loop, perform a 2 mm cut of the vessel using a Potts&#39; scissor, and introduce the guide wire.</li>
	<li>Introduce the guide wire further, ensuring resistance-free insertion and avoiding any kinking of the wire; introduce the catheter over the wire.</li>
	<li>Fix the catheter with sutures.</li>
	<li>Ensure correct position by aspiration of arterial blood verified with blood gas analysis and arterial signal measurement after proper connection to the blood pressure and trans-cardiopulmonary monitoring hard- and software.</li>
	<li>Place a 5 mm flow-probe on the left femoral artery, and test the signal quality by connection to the flowmeter.</li>
	<li>Close both groins with sutures.</li>
	<li>Expose the right carotid artery as well as the right internal jugular vein for placement of 8 Fr. introducer sheaths.</li>
	<li>For catheter placement, proceed in the same manner as described in 3.2-3.4.</li>
	<li>Connect the side-lumen of the carotid artery introducer sheath to the basic pressure monitoring and pulmonary thermodilution hardware for arterial pressure measurement.</li>
	<li>Introduce a pressure-tip catheter into the ascending aorta, and verify the position by connection to the amplifier and synchronic acquisition hard- and software.</li>
	<li>Place a Swan-Ganz pulmonary artery catheter via the venous sheath in the pulmonary artery by inflating the balloon with air at 20 cm depth and gently inserting it until a wedge pressure is seen in the hemodynamic curve. Deflate the balloon and pull the catheter back 2 cm. Ensure satisfying signal quality of pulmonary artery pressure. Connect the thermistors to basic pressure monitoring and pulmonary thermodilution hardware.</li>
	<li>Use sonographic guidance for percutaneous placement of a 12 Fr. 5-Lumen central venous catheter for drug administration and central venous pressure measurement into the external right jugular vein. Use the 6 step-approach for sonographic placement38</li>
	<li>Connect the distal lumen of the catheter to the blood pressure and trans-cardiopulmonary monitoring hard- and software. Switch all drugs and infusions to the central venous catheter. Use different lumen for analgesics, fluids, and catecholamines, and spare the large lumen for administration of colloids during volume-loading steps.</li>
</ol>
4. Surgical preparation
<ol>
	<li>Perform a mini-laparotomy, mobilize the bladder, insert a foley catheter for urine drainage, inflate the balloon with saline, and fix the catheter with pouch sutures.</li>
	<li>Connect the catheter to a urine collection bag displaying the urine amount in mL.</li>
	<li>Increase the FiO2 to 1.0, and re-administer 0.1 mg&#183;kg-1 pancuronium intravenously.</li>
	<li>Perform a median sternotomy by using electrocautery for prepping down to the sternum. Gently dissect the sternum from the surrounding tissue. Perform retrosternal placement of a compress to prevent injuries.</li>
	<li>Stop ventilation and divide the bone with an oscillating saw. Continue ventilation and reduce FiO2 to 0.3. Use electrocautery to reduce bleeding, and seal the sternum with bone wax.</li>
	<li>Carefully mobilize the apex of the left lung, and divide the left lateral part of the diaphragm to facilitate surgical exposure.</li>
	<li>Expose the descending aorta proximal to the celiac trunk by gentle retraction of the left lung, ensuring undisturbed ventilation and avoiding trauma to the left lung (Figure 2A) and divide the surrounding tissue (Figure 2B). Administer 7 mL&#183;kg-1 hydroxyethyl starch colloid if hemodynamic stabilization is needed.</li>
	<li>Place an overhold around the descending aorta to ensure proper exposure (Figure 2C).</li>
	<li>Attach a flow probe around the descending thoracic aorta (Figure 2D). Ensure proper signal quality by connection to the flow module and synchronic acquisition hard- and software. Use contact gel to improve signal quality if needed.</li>
	<li>Attach a vessel loop around the descending aorta, distal to the flow probe to mark the area of aortic cross clamping.</li>
</ol>
5. Assessment and data acquisition
<ol>
	<li>Zero all catheters and level catheters using fluid-filled lines placed at the right atrial level.</li>
	<li>Place needle ECG electrodes and connect them to the synchronic acquisition hard- and software.</li>
	<li>Assessment of trans-cardiopulmonary thermodilution as well as aortic flow and pressure measurements have been previously described 34.</li>
	<li>For cardiac output measurement using pulmonary artery thermodilution, perform 3 injections with 10 mL of cold saline, and note the mean value displayed by basic monitoring hardware.</li>
	<li>Start the laser-Doppler software by simply pressing Start, and set a mark for each measurement step by carefully labeling the steps as M0 to M5.</li>
</ol>
6. Experimental protocol
<ol>
	<li>Perform baseline measurements (M0).</li>
	<li>Perform hemodynamic optimization using volume-loading steps of 7 mL&#183;kg-1 hydroxyethyl starch colloid. Perform each volume-loading step over 5 min using pressurized infusions. After completion of each volume-loading step, allow 5 min for equilibration. Commence volume loading until the increase in cardiac output is &#60;15%.</li>
	<li>Repeat measurements (M1) after completion of hemodynamic optimization.</li>
	<li>Induce ischemia/reperfusion for a total of 48 min of supra-celiac aortic cross-clamping by placing an aortic clamp at the marked area.</li>
	<li>Apply aortic clamping in ascending order of 1-, 2-, 5-, 10-, and 30-min intervals to improve the survival of the animals during the study protocol.</li>
	<li>Continue aortic cross-clamping after each interval after a maximum of 5 min or after normalization of femoral artery flow.</li>
	<li>Perform manual inflow occlusion of the inferior vena cava to prevent blood pressure increases of &#62; 100 mmHg mean arterial pressure.</li>
	<li>Administer bolus injections of norepinephrine or epinephrine during the clamping phase, if needed, to prevent decreases in mean arterial pressure below 40 mmHg.</li>
	<li>Repeat measurements at the end of the 30-min clamping interval prior to reperfusion (M2).</li>
	<li>Gradually open the clamp to ensure hemodynamic stability. Close the clamp if blood pressure drops too quickly and allow stabilization.</li>
	<li>Administer 7 mL&#183;kg-1 of hydroxyethyl starch colloids as well as additional bolus injections of 10-20 &#181;g of norepinephrine and/or epinephrine for stabilization. Administer 2 mL kg-1 of 8.4% sodium bicarbonate if the pH drops below 7.1. Ensure proper adjustment of the respiratory rate to ensure normocapnia.</li>
	<li>Repeat measurements 1 h after reperfusion (M3).</li>
	<li>Repeat hemodynamic optimization as described under 6.2, and repeat measurements (M4).</li>
	<li>Perform final measurements 4.5 h after the induction of ischemia/reperfusion (M5).</li>
</ol>
7. Euthanasia
<ol>
	<li>Administer 40 mmol of potassium chloride intravenously for euthanasia to induce ventricular fibrillation and asystole.</li>
	<li>Terminate ventilation and remove all catheters.</li>
</ol>
8. Organ harvesting
<ol>
	<li>Place the animal in a prone position, and remove the needle probes as well as the drainage.</li>
	<li>Expose the spine by skin incision and removal of muscle tissue using a scalpel and forceps.</li>
	<li>Use an oscillating saw to divide the vertebral arch paramedian on both sides, and remove the dorsal part of the vertebral bone by carefully moving the spinous process sideways to loosen the remaining connections.</li>
	<li>Use forceps to carefully lift the spinal cord from the caudal to cranial ends, and use a scalpel to cut the spinal nerves to remove the spinal cord.</li>
	<li>Store the spinal cord in 4% formalin until further utilization for histopathological evaluation or microsphere quantification.</li>
</ol>
9.&#160;Statistical analysis
<ol>
	<li>Use statistical software.</li>
	<li>Ensure normal distribution by inspection of histograms and log-transform variables if necessary.</li>
	<li>Subject the dependent variables-spinal cord Flux, cardiac output, heart rate, stroke volume, systolic arterial pressure, mean arterial pressure, diastolic arterial pressure, central venous pressure, systemic vascular resistance - as well as upper and lower spinal cord microperfusion as assessed with fluorescent microspheres if desired - to general linear mixed model analyses, using the routine GENLINMIXED for continuous data with an identity link function.</li>
	<li>Use baseline adjustments.</li>
	<li>Specify models with fixed effects for variable baseline and measurement point. Consider measurement point as repeated measures within animals.</li>
	<li>Report p-values of fixed effects for measurement point for each parameter.</li>
	<li>For spinal cord fluorescent microsphere analysis, use region (lower spinal cord, upper spinal cord) in addition as fixed effect and interaction between region and measurement point to evaluate interactions between regions and measurement point, and report p-values of fixed effects for interaction as well.</li>
	<li>Compute baseline adjusted marginal means with 95% confidence interval (CI) for all dependent variables at measurement points M1-M5, followed by pairwise comparisons via least significant difference tests.</li>
	<li>Express variables as mean (95% CI). Express animal weight as mean &#177; standard deviation.</li>
	<li>Present unadjusted p-values.</li>
</ol>

Constantin J. C. Trepte has received an honorary award for lectures by Maquet. All other authors declare no conflicts of interest.This study was supported by the European Society of Anaesthesiology Young Investigator Start-Up Grant 2018.

SCI induced by spinal cord ischemia is a major complication of aortic repair with tremendous impact on patient outcome1,2,3,4,10,11,12. Microcirculation-targeted therapies to prevent and treat SCI are most promising. The protocol provides a reproducible method for real-time spinal cord micro...

Spinal cord injury induced by ischemia/reperfusion (SCI) is one of the most devastating complications of aortic repair associated with reduced outcome1,2,3,4. Current prevention and treatment options for SCI include the optimization of macrohemodynamic parameters as well as the normalization of cerebrospinal fluid pressure (CSP) to improve spinal cord perfusion pressure2,5,6,7,8,9. Despite the implementation of these maneuvers, incidence of SCI still ranges between 2% and 31% depending on the complexity of aortic repair10,11,12.
Recently, microcirculation has gained increased attention13,14. Microcirculation is the area of cellular oxygen uptake and metabolic exchange and therefore, plays a critical role in organ function and cellular integrity13. Impaired microcirculatory blood flow is a major determinant of tissue ischemia associated with increased mortality15,16,17,18,19. Impairment of spinal cord microcirculation is associated with reduced neurological function and outcome20,21,22,23. Therefore, optimization of microperfusion for the treatment of SCI is a most promising approach. Persistence of microcirculatory disturbances, despite macrocirculatory optimization, has been described26,27,28,29. This loss of hemodynamic coherence occurs frequently in various conditions including ischemia/reperfusion, emphasizing the need for direct microcirculatory evaluation and microcirculation-targeted therapies26,27,30.
So far, only few studies have used laser-Doppler probes for real-time assessment of spinal cord microcirculatory behavior20,31. Existing studies have often used microsphere injection techniques, which are limited by intermittent use and post-mortem analysis32,33. The number of different measurements using microsphere injection technique is limited by the availability of microspheres with different wavelengths. Moreover, in contrast to Laser-Doppler techniques, real-time assessment of microperfusion is not possible, as post-mortem tissue processing and analysis is needed for this method. Here, we present an experimental protocol for the real-time assessment of spinal cord microcirculation in a porcine large animal model of ischemia/reperfusion.
This study was part of a large animal project combining a randomized study comparing the influence of crystalloids vs. colloids on microcirculation in ischemia/reperfusion as well as an explorative randomized study on the effects of fluids vs. vasopressors on spinal cord microperfusion. Flow probe 2-point calibration as well as pressure-tip catheter calibration has been previously described34. In addition to the reported protocol, fluorescent microspheres were used for the measurement of spinal cord microperfusion, as previously described, using 12 samples of spinal cord tissue for each animal, with samples 1-6 representing the upper spinal cord and 7-12 representing the lower spinal cord35,36. Microsphere injection was performed for each measurement step after the completion of Laser-Doppler recordings and macrohemodynamic evaluation. Histopathological evaluation was performed using the Kleinman-Score as previously described37.

<table><tbody><tr><td>CardioMed Flowmeter</td><td>Medistim AS, Oslo, Norway</td><td>CM4000</td><td>Flowmeter for Flow-Probe Femoral Artery</td></tr><tr><td>CardioMed Flow-Probe, 5mm</td><td>Medistim AS, Oslo, Norway</td><td>PS100051</td><td>Flow-Probe Femoral Artery</td></tr><tr><td>COnfidence probe,&amp;nbsp;</td><td>Transonic Systems Inc., Ithaca, NY, USA</td><td>MA16PAU</td><td>Flow-Probe Aorta</td></tr><tr><td>16 mm liners</td><td></td><td></td><td></td></tr><tr><td>DIVA Sevoflurane Vapor</td><td>Dr&amp;auml;ger Medical, L&amp;uuml;beck, Germany</td><td></td><td>Vapor</td></tr><tr><td>Hotline Level 1 Fluid Warmer</td><td>Smiths Medical Germany GmbH, Grasbrunn, Germany</td><td>HL-90-DE-230</td><td>Fluid Warmer</td></tr><tr><td>Infinity Delta</td><td>Dr&amp;auml;ger Medical, L&amp;uuml;beck, Germany</td><td></td><td>Basic Monitoring Hardware</td></tr><tr><td>Infinity Hemo</td><td>Dr&amp;auml;ger Medical, L&amp;uuml;beck, Germany</td><td></td><td>Basic Pressure Monitoring and Pulmonary Thermodilution Hardware</td></tr><tr><td>LabChart Pro</td><td>ADInstruments Ltd., Oxford, UK</td><td>v8.1.16</td><td>Synchronic Laser-Doppler, Blood Pressure, ECG and Blood-Flow Aquisition Software</td></tr><tr><td>LiquoGuard 7</td><td>M&amp;ouml;ller Medical GmbH, Fulda, Germany</td><td></td><td>Cerebrospinal Fluid Drainage System</td></tr><tr><td>Millar Micro-Tip Pressure Catheter (5F, Single, Curved, 120cm, PU/WD)</td><td>ADInstruments Ltd., Oxford, UK</td><td>SPR-350</td><td>Pressure-Tip Catheter Aorta</td></tr><tr><td>moor VMS LDF</td><td>moor Instruments, Devon, UK</td><td></td><td>Designated Laser-Doppler Hardware</td></tr><tr><td>moor VMS Research Software</td><td>moor Instruments, Devon, UK</td><td></td><td>Designated Laser-Doppler Software</td></tr><tr><td>Perivascular Flow Module</td><td>Transonic Systems Inc., Ithaca, NY, USA</td><td>TS 420</td><td>Flow-Module for Flow-Probe Aorta</td></tr><tr><td>PiCCO 2, Science Version</td><td>Getinge AB, G&amp;ouml;teborg, Sweden</td><td>v. 6.0</td><td>Blood Pressure and Transcardiopulmonary Monitoring Hard- and Software</td></tr><tr><td>PiCCO 5 Fr. 20cm</td><td>Getinge AB, G&amp;ouml;teborg, Sweden</td><td></td><td>Thermistor-tipped Arterial Line&amp;nbsp;</td></tr><tr><td>PowerLab</td><td>ADInstruments Ltd., Oxford, UK</td><td>PL 3516</td><td>Synchronic Laser-Doppler, Blood Pressure, ECG and Blood-Flow Aquisition Hardware</td></tr><tr><td>QuadBridgeAmp</td><td>ADInstruments Ltd., Oxford, UK</td><td>FE 224</td><td>Four Channel Bridge Amplifier for Laser-Doppler and Invasive Blood Pressure Aquisition</td></tr><tr><td>Silverline</td><td>Spiegelberg, Hamburg, Germany</td><td>ELD33.010.02</td><td>Cerebrospinal Fluid Drainage</td></tr><tr><td>SPSS statistical software package&amp;nbsp;</td><td>IBM SPSS Statistics Inc., Armonk, New York, USA</td><td>v. 27</td><td>Statistical Software</td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>Twinwarm Warming System</td><td>Moeck &amp;amp; Moeck GmbH, Hamburg, Germany</td><td>12TW921DE</td><td>Warming System</td></tr><tr><td>Universal II Warming Blanket</td><td>Moeck &amp;amp; Moeck GmbH, Hamburg, Germany</td><td>906</td><td>Warming Blanket</td></tr><tr><td>VP 3 Probe, 8mm length (individually manufactured)</td><td>moor Instruments, Devon, UK</td><td></td><td>Laser-Doppler Probe</td></tr><tr><td>Zeus</td><td>Dr&amp;auml;ger Medical, L&amp;uuml;beck, Germany</td><td></td><td>Anesthesia Machine</td></tr></tbody></table>

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.

All six animals survived until the completion of the protocol. Animal weight was 48.2 &#177; 2.9 kg; five animals were male, and one animal was female. Spinal cord needle probe insertion as well as spinal cord Flux measurement was feasible in all animals.
Examples of real-time spinal cord microcirculatory recordings in combination with cerebral microcirculatory and macrohemodynamic recordings during aortic cross-clamping for isc...

Watch this Scientific Journal Video about Real-Time Assessment of Spinal Cord Microperfusion in a Porcine Model of Ischemia/Reperfusion at JoVE.com

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

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 ...

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3.4K Views.  University Medical Center Hamburg-Eppendorf. De microcirculatie van het ruggenmerg speelt een cruciale rol bij dwarslaesie. De meeste methoden laten geen real-time beoordeling van de microcirculatie van het ruggenmerg toe, wat essentieel is voor de ontwikkeling van microcirculatiegerichte therapieën. Hier stellen we een protocol voor met behulp van Laser-Doppler-Flow Needle-sondes in een groot diermodel van ischemie / reperfusie.

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

Using Laser Doppler Imaging and Monitoring to Analyze Spinal Cord Microcirculation in Rat

Laser Doppler flowmetry (LDF) is a noninvasive method for blood flow (BF) measurement, which makes it preferable for measuring microcirculatory alterations of the spinal cord. In this article, our goal was to use both Laser Doppler imaging and monitoring to analyze the change of BF after spinal cord injury. Both the laser Doppler image scanner and the probe/monitor were being employed to obtain each readout. The data of LDPI provided a local distribution of BF, which gave an overview of perfusion around the injury site and made it accessible for comparative analysis of BF among different locations. By intensely measuring the probing area over a period of time, a combined probe was used to simultaneously measure the BF and oxygen saturation of the spinal cord, showing overall spinal cord perfusion and oxygen supply. LDF itself has a few limitations, such as relative flux, sensitivity to movement, and biological zero signal. However, the technology has been applied in clinical and experimental study due to its simple setup and rapid measurement of BF.

Here we present a combination of laser Doppler perfusion imaging (LDPI) and laser Doppler perfusion monitoring (LDPM) to measure spinal cord local blood flows and oxygen saturation (SO2), as well as a standardized procedure for introducing spinal cord trauma on rat.

Laser Doppler flowmetry (LDF) is a noninvasive method for blood flow (BF) measurement, which makes it preferable for measuring microcirculatory ...

Behavior

Reproducable Paraplegia by Thoracic Aortic Occlusion in a Murine Model of Spinal Cord Ischemia-reperfusion

Background 
Lower extremity paralysis continues to complicate aortic interventions. The lack of understanding of the underlying pathology has hindered advancements to decrease the occurrence this injury.&#160;The current model demonstrates reproducible lower&#160;extremity paralysis following thoracic aortic occlusion.
Methods 
Adult male C57BL6 mice were anesthetized with isoflurane. Through a cervicosternal incision the aorta was exposed. The descending thoracic aorta and left subclavian arteries were identified without entrance into pleural space. Skeletonization of these arteries was followed by immediate closure (Sham) or occlusion for 4 min (moderate ischemia) or 8 min (prolonged ischemia). The sternotomy and skin were closed and the mouse was transferred to warming bed for recovery.&#160; Following recovery, functional analysis was obtained at 12 hr intervals until 48 hr.
Results 
Mice that underwent sham surgery showed no observable hind&#160;limb deficit. Mice subjected to moderate ischemia for 4 min had minimal functional deficit at 12 hr followed by progression to complete paralysis at 48 hr. Mice subjected to prolonged ischemia had an immediate paralysis with no observable hind-limb movement at any point in the postoperative period. There was no observed intraoperative or post&#160;operative mortality.
Conclusion 
Reproducible lower extremity paralysis whether immediate or delayed can be achieved in a murine model. Additionally, by using a median sternotomy and careful dissection, high survival rates, and reproducibility can be achieved.

The lack of mechanistic understanding of spinal cord ischemia-reperfusion injury has hindered further adjuncts to prevent paraplegia following high risk aortic operations. Thus, the development of animal models is imperative. This manuscript demonstrates reproducible lower extremity paralysis following thoracic aortic occlusion in a murine model.

Background
Lower extremity paralysis continues to complicate aortic interventions. The lack of understanding of the underlying pathology has hindered ...

Contrast Enhanced Ultrasound Imaging for Assessment of Spinal Cord Blood Flow in Experimental Spinal Cord Injury

Reduced spinal cord blood flow (SCBF) (i.e., ischemia) plays a key role in traumatic spinal cord injury (SCI) pathophysiology and is accordingly an important target for neuroprotective therapies. Although several techniques have been described to assess SCBF, they all have significant limitations. To overcome the latter, we propose the use of real-time contrast enhanced ultrasound imaging (CEU). Here we describe the application of this technique in a rat contusion model of SCI. A jugular catheter is first implanted for the repeated injection of contrast agent, a sodium chloride solution of sulphur hexafluoride encapsulated microbubbles. The spine is then stabilized with a custom-made 3D-frame and the spinal cord dura mater is exposed by a laminectomy at ThIX-ThXII. The ultrasound probe is then positioned at the posterior aspect of the dura mater (coated with ultrasound gel). To assess baseline SCBF, a single intravenous injection (400 &#181;l) of contrast agent is applied to record its passage through the intact spinal cord microvasculature. A weight-drop device is subsequently used to generate a reproducible experimental contusion model of SCI. Contrast agent is re-injected 15 min following the injury to assess post-SCI SCBF changes. CEU allows for real time and in-vivo assessment of SCBF changes following SCI. In the uninjured animal, ultrasound imaging showed uneven blood flow along the intact spinal cord. Furthermore, 15 min post-SCI, there was critical ischemia at the level of the epicenter while SCBF remained preserved in the more remote intact areas. In the regions adjacent to the epicenter (both rostral and caudal), SCBF was significantly reduced. This corresponds to the previously described &#8220;ischemic penumbra zone&#8221;. This tool is of major interest for assessing the effects of therapies aimed at limiting ischemia and the resulting tissue necrosis subsequent to SCI.

Contrast Enhanced Ultrasound imaging is a reliable in-vivo tool for quantifying spinal cord blood flow in an experimental rat spinal cord injury model. This paper contains a comprehensive protocol for application of this technique in association with a contusion model of thoracic spinal cord injury.

Reduced spinal cord blood flow (SCBF) (i.e., ischemia) plays a key role in traumatic spinal cord injury (SCI) pathophysiology and is accordingly an ...

Improvement of a Closed Chest Porcine Myocardial Infarction Model by Standardization of Tissue and Blood Sampling Procedures

Myocardial ischemia reperfusion (I/R) injury contributes to almost half of the necrotic area after myocardial infarction. To date there is no approved drug to prevent or reduce myocardial I/R injury. The study and understanding of the pathophysiological mechanisms of myocardial I/R injury is essential to develop successful treatments. Large animal experiments are an important step in translational methods. The porcine model of acute myocardial infarction has been established and described by ourselves and others. We aimed to further improve the value of the model by focusing in detail on the sampling techniques for use in future experiments. Furthermore, we emphasize small but important steps that can affect the quality of the final results. To mimic the clinical situation of myocardial I/R injury, a percutaneous coronary intervention (PCI) catheter was inserted into the left anterior descending coronary artery (LAD) of an anesthetized pig. &#176;&#176;&#176;This model mimics acute myocardial infarction and PCI treatment in humans with the possibility of accurately determining the area at risk as well as the necrotic- and viable ischemic tissue. Here the model was used to investigate the effect of a bicyclic peptide inhibitor of FXIIa. The model can also be modified to allow longer reperfusion times to study later effects of myocardial infarction.

Here we demonstrate a protocol to standardize sampling procedures of an established porcine model of acute myocardial infarction in order to increase its translational value in the understanding of the pathophysiology of myocardial ischemia/reperfusion injury and to test novel drug candidates.

Myocardial ischemia reperfusion (I/R) injury contributes to almost half of the necrotic area after myocardial infarction. To date there is no approved ...

Intestinal Stem Cell Isolation and Culture in a Porcine Model of Segmental Small Intestinal Ischemia

Intestinal ischemia remains a major cause of morbidity and mortality in human and veterinary patients. Many disease processes result in intestinal ischemia, when the blood supply and therefore oxygen is decreased to the intestine. This leads to intestinal barrier loss and damage to the underlying tissue. Intestinal stem cells reside at the base of the crypts of Lieberk&#252;hn and are responsible for intestinal renewal during homeostasis and following injury. Ex vivo cell culture techniques have allowed for the successful study of epithelial stem cell interactions by establishing culture conditions that support the growth of three-dimensional epithelial organ-like systems (termed "enteroids" and "colonoids" from the small and large intestine, respectively). These enteroids are composed of crypt and villus-like domains and mature to contain all of the cell types found within the epithelium. Historically, murine models have been utilized to study intestinal injury. However, a porcine model offers several advantages including similarity of size as well as gastrointestinal anatomy and physiology to that of humans. By utilizing a porcine model, we establish a protocol in which segmental loops of intestinal ischemia can be created within a single animal, enabling the study of differing time points of ischemic injury and repair in vivo. Additionally, we describe a method to isolate and culture the intestinal stem cells from the ischemic loops of intestine, allowing for the continued study of epithelial repair, modulated by stem cells, ex vivo.

This protocol will enable readers to successfully establish a porcine model of segmental intestinal ischemia and subsequently isolate and culture intestinal stem cells for the study of epithelial repair following injury.

Intestinal ischemia remains a major cause of morbidity and mortality in human and veterinary patients. Many disease processes result in intestinal ...

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

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 ...

Post-Myocardial Infarction Heart Failure in Closed-chest Coronary Occlusion/Reperfusion Model in G&#246;ttingen Minipigs and Landrace Pigs

The development of heart failure is the most powerful predictor of long-term mortality in patients surviving acute myocardial infarction (MI). There is an unmet clinical need for prevention and therapy of post-myocardial infarction heart failure (post-MI HF). Clinically relevant pig models of post-MI HF are prerequisites for final proof-of-concept studies before entering into clinical trials in drug and medical device development.
Here we aimed to characterize a closed-chest porcine model of post-MI HF in adult G&#246;ttingen minipigs with long-term follow-up including serial cardiac magnetic resonance imaging (CMRI) and to compare it with the commonly used Landrace pig model.
MI was induced by intraluminal balloon occlusion of the left anterior descending coronary artery for 120 min in G&#246;ttingen minipigs and for 90 min in Landrace pigs, followed by reperfusion. CMRI was performed to assess cardiac morphology and function at baseline in both breeds and at 3 and 6 months in G&#246;ttingen minipigs and at 2 months in Landrace pigs, respectively.
Scar sizes were comparable in the two breeds, but MI resulted in a significant decrease of left ventricular ejection fraction (LVEF) only in G&#246;ttingen minipigs, while Landrace pigs did not show a reduction of LVEF. Right ventricular (RV) ejection fraction increased in both breeds despite the negligible RV scar sizes. In contrast to the significant increase of left ventricular end-diastolic (LVED) mass in Landrace pigs at 2 months, G&#246;ttingen minipigs showed a slight increase in LVED mass only at 6 months.
In summary, this is the first characterization of post-MI HF in G&#246;ttingen minipigs in comparison to Landrace pigs, showing that the G&#246;ttingen minipig model reflects post-MI HF parameters comparable to the human pathology. We conclude that the G&#246;ttingen minipig model is superior to the Landrace pig model to study the development of post-MI HF.

The overall goal of the current study is to present the techniques of induction of myocardial infarction (MI) and post-myocardial infarction heart failure (post-MI HF) in closed-chest, adult G&#246;ttingen minipigs and the characterization of post-MI HF model in G&#246;ttingen minipigs as compared to Landrace pigs.

The development of heart failure is the most powerful predictor of long-term mortality in patients surviving acute myocardial infarction (MI). There is an ...

Coronary Artery Assessment During Ex-Situ Heart Perfusion

Coronary Angiography During Ex-Situ Heart Perfusion in a Porcine Model

Heart transplantation is the gold standard treatment for advanced heart failure. The procurement of extended criteria donors (ECD) increases due to the current organ shortage. Coronary angiography is recommended in ECD at risk for coronary artery disease but is not systematically performed. These hearts are, therefore, either declined for transplant or procured without screening for coronary artery disease. Coronary angiography during normothermic ex-situ heart perfusion (NESP) could be an interesting approach to enhance the rate of ECD procurement and to reduce the risk of primary graft failure in the absence of coronary angiography in ECD. The present protocol aims to provide material details along with optimal imaging views for coronary angiography during NESP. Reproducible angiographic views were observed, including one dedicated to the right coronary artery, two for the left anterior descending artery, two for the circumflex artery, and a spider view. Continuous lactate extraction was observed in all procedures with a final median concentration of 1.10 mmol/L (0.61-1.75 mmol/L) two hours after coronary angiography, consistent with myocardial viability. The median contrast agent volume used for ex-situ imaging of the isolated perfused heart was 48 mL (38-108 mL). This protocol was reproducible for coronary artery imaging and did not impair myocardial viability during NESP.

Author Spotlight: Innovative Technique for Coronary Angiography in Marginal Donors

Screening for coronary artery disease could increase the procurement of donor hearts with extended criteria. A protocol for performing reproducible coronary angiography during ex-situ heart perfusion is described herein in a porcine model.

Heart transplantation is the gold standard treatment for advanced heart failure. The procurement of extended criteria donors (ECD) increases due to the ...

Laser Doppler Needle Probe Insertion: A Technique for Real-time Quantification of Blood Flow within the Spinal Cord in a Porcine Model

Source: Behem, C. R. et al., <a href="https://www.jove.com/v/62047">Real-Time Assessment of Spinal Cord Microperfusion in a Porcine Model of Ischemia/Reperfusion.</a> J. Vis. Exp. (2020).
This video demonstrates the placement of a laser Doppler needle probe in the spinal cord of a porcine model. This helps in the continuous quantification of blood supply to the spinal cord, an important assessment in spinal cord injury.

Source: Behem, C. R. et al., Real-Time Assessment of Spinal Cord Microperfusion in a Porcine Model of Ischemia/Reperfusion. J. Vis. Exp. (2020).
This ...

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

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