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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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,&#160;</td>
			<td rowspan="2">Transonic Systems Inc., Ithaca, NY, USA</td>
			<td rowspan="2">MA16PAU</td>
			<td rowspan="2">Flow-Probe Aorta</td>
		</tr>
		<tr>
			<td>16 mm liners</td>
		</tr>
		<tr>
			<td>DIVA Sevoflurane Vapor</td>
			<td>Dr&#228;ger Medical, L&#252;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&#228;ger Medical, L&#252;beck, Germany</td>
			<td></td>
			<td>Basic Monitoring Hardware</td>
		</tr>
		<tr>
			<td>Infinity Hemo</td>
			<td>Dr&#228;ger Medical, L&#252;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&#246;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&#246;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&#246;teborg, Sweden</td>
			<td></td>
			<td>Thermistor-tipped Arterial Line&#160;</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 rowspan="2">SPSS statistical software package&#160;</td>
			<td rowspan="2">IBM SPSS Statistics Inc., Armonk, New York, USA</td>
			<td rowspan="2">v. 27</td>
			<td rowspan="2">Statistical Software</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>Twinwarm Warming System</td>
			<td>Moeck &#38; Moeck GmbH, Hamburg, Germany</td>
			<td>12TW921DE</td>
			<td>Warming System</td>
		</tr>
		<tr>
			<td>Universal II Warming Blanket</td>
			<td>Moeck &#38; 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&#228;ger Medical, L&#252;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. 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.

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

Acute and Chronic Tactile Sensory Testing after Spinal Cord Injury in Rats

Spinal cord injury (SCI) impairs sensory systems causing allodynia1-8. To identify cellular and molecular causes of allodynia, sensitive and valid sensory testing in rat SCI models is needed. However, until recently, no single testing approach had been validated for SCI so that standardized methods have not been implemented across labs. Additionally, available testing methods could not be implemented acutely or when severe motor impairments existed, preventing studies of the development of SCI-induced allodynia3. Here we present two validated sensory testing methods using von Frey Hair (VFH) monofilaments which quantify changes in tactile sensory thresholds after SCI4-5. One test is the well-established Up-Down test which demonstrates high sensitivity and specificity across different SCI severities when tested chronically5. The other test is a newly-developed dorsal VFH test that can be applied acutely after SCI when allodynia develops, prior to motor recovery4-5. Each VFH monofilament applies a calibrated force when touched to the skin of the hind paw until it bends. In the up-down method, alternating VFHs of higher or lower forces are used on the plantar L5 dermatome to delineate flexor withdrawal thresholds. Successively higher forces are applied until withdrawal occurs then lower force VFHs are used until withdrawal ceases. The tactile threshold reflects the force required to elicit withdrawal in 50% of the stimuli. For the new test, each VFH is applied to the dorsal L5 dermatome of the paw while the rat is supported by the examiner. The VFH stimulation occurs in ascending order of force until at least 2 of 3 applications at a given force produces paw withdrawal. Tactile sensory threshold is the lowest force to elicit withdrawal 66% of the time. Acclimation, testing and scoring procedures are described. Aberrant trials that require a retest and typical trials are defined. Animal use was approved by Ohio State University Animal Care and Use Committee.

We describe two tactile sensory testing methods for acute or chronic periods of spinal cord injury in rats. These validated procedures can detect the development and maintenance of allodynia-like sensations.

Spinal cord injury (SCI) impairs sensory systems causing allodynia1-8. To identify cellular and molecular causes of allodynia, sensitive and valid sensory ...

Real-time Digital Imaging of Leukocyte-endothelial Interaction in Ischemia-reperfusion Injury (IRI) of the Rat Cremaster Muscle

Ischemia-reperfusion injury (IRI) has been implicated in a large array of pathological conditions such as cerebral stroke, myocardial infarction, intestinal ischemia as well as following transplant and cardiovascular surgery.1 Reperfusion of previously ischemic tissue, while essential for the prevention of irreversible tissue injury, elicits excessive inflammation of the affected tissue. Adjacent to the production of reactive oxygen species, activation of the complement system and increased microvascular permeability, the activation of leukocytes is one of the principle actors in the pathological cascade of inflammatory tissue damage during reperfusion.2, 3 Leukocyte activation is a multistep process consisting of rolling, firm adhesion and transmigration and is mediated by a complex interaction between adhesion molecules in response to chemoattractants such as complement factors, chemokines, or platelet-activating factor.4
While leukocyte rolling in postcapillary venules is predominantly mediated by the interaction of selectins5 with their counter ligands, firm adhesion of leukocytes to the endothelium is selectin-controlled via binding to intercellular adhesion molecules (ICAM) and vascular cellular adhesion molecules (VCAM).6, 7 
Gold standard for the in vivo observation of leukocyte-endothelial interaction is the technique of intravital microscopy, first described in 1968.8
Though various models of IRI (ischemia-reperfusion injury) have been described for various organs, 9-12 only few are suitable for direct visualization of leukocyte recruitment in the microvascular bed on a high level of image quality.8 
We here promote the digital intravital epifluorescence microscopy of the postcapillary venule in the cremasteric microcirculation of the rat 13 as a convenient method to qualitatively and quantitatively analyze leukocyte recruitment for IRI-research in striated muscle tissue and provide a detailed manual for accomplishing the technique. We further illustrate common pitfalls and provide useful tips which should enable the reader to truly appreciate, and safely perform the method.
In a step by step protocol we depict how to get started with respiration controlled anesthesia under sufficient monitoring to keep the animal firmly anesthetized for longer periods of time. We then describe the cremasteric preparation as a thin flat sheet for outstanding optical resolution and provide a protocol for leukocyte imaging in IRI that has been well established in our laboratories.

Real-time Digital Imaging of Leukocyte-endothelial Interaction in Ischemia-reperfusion Injury IRI of the Rat Cremaster Muscle

Digital intravital epifluorescence microscopy of postcapillary venules in the cremasteric microcirculation is a convenient method to gain insights into leukocyte-endothelial interaction in vivo in ischemia-reperfusion injury (IRI) of striated muscle tissue. We here provide a detailed protocol to safely perform the technique and discuss its applications and limitations.

Ischemia-reperfusion injury (IRI) has been implicated in a large array of pathological conditions such as cerebral stroke, myocardial infarction, ...

A Contusion Model of Severe Spinal Cord Injury in Rats

The translational potential of novel treatments should be investigated in severe spinal cord injury (SCI) contusion models. A detailed methodology is described to obtain a consistent model of severe SCI. Use of a stereotactic frame and computer controlled impactor allows for creation of reproducible injury. Hypothermia and urinary tract infection pose significant challenges in the post-operative period. Careful monitoring of animals with daily weight recording and bladder expression allows for early detection of post-operative complications. The functional results of this contusion model are equivalent to transection models. The contusion model can be utilized to evaluate the efficacy of both neuroprotective and neuroregenerative approaches.

A contusion model of severe spinal cord injury is described. Detailed pre-operative, operative and post-operative steps are described to obtain a consistent model.

The translational potential of novel treatments should be investigated in severe spinal cord injury (SCI) contusion models. A detailed methodology is ...

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 ...

Controlled Cortical Impact Model for Traumatic Brain Injury

Every year over a million Americans suffer a traumatic brain injury (TBI). Combined with the incidence of TBIs worldwide, the physical, emotional, social, and economical effects are staggering. Therefore, further research into the effects of TBI and effective treatments is necessary. The controlled cortical impact (CCI) model induces traumatic brain injuries ranging from mild to severe. This method uses a rigid impactor to deliver mechanical energy to an intact dura exposed following a craniectomy. Impact is made under precise parameters at a set velocity to achieve a pre-determined deformation depth. Although other TBI models, such as weight drop and fluid percussion, exist, CCI is more accurate, easier to control, and most importantly, produces traumatic brain injuries similar to those seen in humans. However, no TBI model is currently able to reproduce pathological changes identical to those seen in human patients. The CCI model allows investigation into the short-term and long-term effects of TBI, such as neuronal death, memory deficits, and cerebral edema, as well as potential therapeutic treatments for TBI.

Traumatic brain injuries (TBIs) remain a serious health problem. Using the controlled cortical impact surgery model, research on the effects of TBI and possible treatment methods may be performed.

Every year over a million Americans suffer a traumatic brain injury (TBI).  Combined with the incidence of TBIs worldwide, the physical, emotional, ...

A Radio-telemetric System to Monitor Cardiovascular Function in Rats with Spinal Cord Transection and Embryonic Neural Stem Cell Grafts

High thoracic or cervical spinal cord injury (SCI) can lead to cardiovascular dysfunction. To monitor cardiovascular parameters, we implanted a catheter connected to a radio transmitter into the femoral artery of rats that underwent a T4 spinal cord transection with or without grafting of embryonic brainstem-derived neural stem cells expressing green fluorescent protein. Compared to other methods such as cannula insertion or tail-cuff, telemetry is advantageous to continuously monitor blood pressure and heart rate in freely moving animals. It is also capable of long term multiple data acquisitions. In spinal cord injured rats, basal cardiovascular data under unrestrained condition and autonomic dysreflexia in response to colorectal distension were successfully recorded. In addition, cardiovascular parameters before and after SCI can be compared in the same rat if a transmitter is implanted before a spinal cord transection. One limitation of the described telemetry procedure is that implantation in the femoral artery may influence the blood supply to the ipsilateral hindlimb.

We present a protocol for using a radio-telemetric system to record cardiovascular parameters in T4 spinal cord transected rats eight weeks after embryonic brainstem neural stem cell grafting into the lesion site. Telemetry is an advanced technique to accurately evaluate cardiovascular function in conscious freely moving spinal cord injured rats.

High thoracic or cervical spinal cord injury (SCI) can lead to cardiovascular dysfunction. To monitor cardiovascular parameters, we implanted a catheter ...

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 ...

Induction and Assessment of Ischemia-reperfusion Injury in Langendorff-perfused Rat Hearts

The biochemical events surrounding ischemia reperfusion injury in the acute setting are of great importance to furthering novel treatment options for myocardial infarction and cardiac complications of thoracic surgery. The ability of certain drugs to precondition the myocardium against ischemia reperfusion injury has led to multiple clinical trials, with little success. The isolated heart model allows acute observation of the functional effects of ischemia reperfusion injury in real time, including the effects of various pharmacological interventions administered at any time-point before or within the ischemia-reperfusion injury window. Since brief periods of ischemia can precondition the heart against ischemic injury, in situ aortic cannulation is performed to allow for functional assessment of non-preconditioned myocardium. A saline filled balloon is placed into the left ventricle to allow for real-time measurement of pressure generation. Ischemic injury is simulated by the cessation of perfusion buffer flow, followed by reperfusion. The duration of both ischemia and reperfusion can be modulated to examine biochemical events at any given time-point. Although the Langendorff isolated heart model does not allow for the consideration of systemic events affecting ischemia and reperfusion, it is an excellent model for the examination of acute functional and biochemical events within the window of ischemia reperfusion injury as well as the effect of pharmacological intervention on cardiac pre- and postconditioning. The goal of this protocol is to demonstrate how to perform in situ aortic cannulation and heart excision followed by ischemia/reperfusion injury in the Langendorff model.

The isolated rat heart is an enduring model for ischemia reperfusion injury. Here, we describe the process of harvesting the beating heart from a rat via in situ aortic cannulation, Langendorff perfusion of the heart, simulated ischemia-reperfusion injury, and infarct staining to confirm the extent of ischemic insult.

The biochemical events surrounding ischemia reperfusion injury in the acute setting are of great importance to furthering novel treatment options for ...

A Neonatal Mouse Spinal Cord Compression Injury Model

Spinal cord injury (SCI) typically causes devastating neurological deficits, particularly through damage to fibers descending from the brain to the spinal cord. A major current area of research is focused on the mechanisms of adaptive plasticity that underlie spontaneous or induced functional recovery following SCI. Spontaneous functional recovery is reported to be greater early in life, raising interesting questions about how adaptive plasticity changes as the spinal cord develops. To facilitate investigation of this dynamic, we have developed a SCI model in the neonatal mouse. The model has relevance for pediatric SCI, which is too little studied. Because neural plasticity in the adult involves some of the same mechanisms as neural plasticity in early life1, this model may potentially have some relevance also for adult SCI. Here we describe the entire procedure for generating a reproducible spinal cord compression (SCC) injury in the neonatal mouse as early as postnatal (P) day 1. SCC is achieved by performing a laminectomy at a given spinal level (here described at thoracic levels 9-11) and then using a modified Yasargil aneurysm mini-clip to rapidly compress and decompress the spinal cord. As previously described, the injured neonatal mice can be tested for behavioral deficits or sacrificed for ex vivo physiological analysis of synaptic connectivity using electrophysiological and high-throughput optical recording techniques1. Earlier and ongoing studies using behavioral and physiological assessment have demonstrated a dramatic, acute impairment of hindlimb motility followed by a complete functional recovery within 2 weeks, and the first evidence of changes in functional circuitry at the level of identified descending synaptic connections1.

This article describes a method for generating a reproducible spinal cord compression injury (SCI) in the neonatal mouse. The model provides an advantageous platform for studying mechanisms of adaptive plasticity that underlie spontaneous functional recovery.

Spinal cord injury (SCI) typically causes devastating neurological deficits, particularly through damage to fibers descending from the brain to the spinal ...

Murine Model of Intestinal Ischemia-reperfusion Injury

Intestinal ischemia is a life-threatening condition associated with a broad range of clinical conditions including atherosclerosis, thrombosis, hypotension, necrotizing enterocolitis, bowel transplantation, trauma and chronic inflammation. Intestinal ischemia-reperfusion (IR) injury is a consequence of acute mesenteric ischemia, caused by inadequate blood flow through the mesenteric vessels, resulting in intestinal damage. Reperfusion following ischemia can further exacerbate damage of the intestine. The mechanisms of IR injury are complex and poorly understood. Therefore, experimental small animal models are critical for understanding the pathophysiology of IR injury and the development of novel therapies.
Here we describe a mouse model of acute intestinal IR injury that provides reproducible injury of the small intestine without mortality. This is achieved by inducing ischemia in the region of the distal ileum by temporally occluding the peripheral and terminal collateral branches of the superior mesenteric artery for 60 min using microvascular clips. Reperfusion for 1 hr, or 2 hr after injury results in reproducible injury of the intestine examined by histological analysis. Proper position of the microvascular clips is critical for the procedure. Therefore the video clip provides a detailed visual step-by-step description of this technique. This model of intestinal IR injury can be utilized to study the cellular and molecular mechanisms of injury and regeneration.

Here we describe the detailed procedure of intestinal ischemia-reperfusion in mice which results in reproducible injury without mortality to encourage the standardization of this technique across the field. This model of intestinal ischemia-reperfusion injury can be utilized to study the cellular and molecular mechanisms of injury and regeneration.