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

Summary

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.

Abstract

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.

Introduction

Spinal cord injury induced by ischemia/reperfusion (SCI) is one of the most devastating complications of aortic repair associated with reduced outcome^1,2,3,4. 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 pressure^2,5,6,7,8,9. Despite the implementation of these maneuvers, incidence of SCI still ranges between 2% and 31% depending on the complexity of aortic repair^10,11,12.

Recently, microcirculation has gained increased attention^13,14. 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 mortality^{15,16,17,18,19}. Impairment of spinal cord microcirculation is associated with reduced neurological function and outcome^20,21,22,23. Therefore, optimization of microperfusion for the treatment of SCI is a most promising approach. Persistence of microcirculatory disturbances, despite macrocirculatory optimization, has been described^26,27,28,29. This loss of hemodynamic coherence occurs frequently in various conditions including ischemia/reperfusion, emphasizing the need for direct microcirculatory evaluation and microcirculation-targeted therapies^26,27,30.

So far, only few studies have used laser-Doppler probes for real-time assessment of spinal cord microcirculatory behavior^20,31. Existing studies have often used microsphere injection techniques, which are limited by intermittent use and post-mortem analysis^32,33. 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 cord^35,36. 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.

Protocol

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 'Guide for the Care and Use of Laboratory Animals' (NIH publication No. 86-23, revised 2011) as well as FELASA recommendations and experiments were carried out according to the ARRIVE guidelines^24,25. 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

1. 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.
2. Perform intramuscular injections of ketamine 20 mg·kg^-1, azaperone 4 mg·kg^-1, and midazolam 0.1 mg·kg^-1 for premedication and sedation.
3. Place a venous catheter in an ear vein, secure proper fixation, and assess functionality by fast application of 10 mL of saline.
4. Place the animal in a supine position on a warming blanket to prevent heat loss.
5. 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.
6. Administer 15 L·min^-1 of oxygen via a pig-shaped mask for preoxygenation.
7. Inject intravenous boli of 0.1 mg·kg^-1 of 1% propofol, if necessary, and perform endotracheal intubation.
8. Secure correct placement with end tidal capnography and auscultation, administer 0.1 mg•kg^-1 of pancuronium, and ensure proper fixation of the endotracheal tube.
9. Establish volume-controlled ventilation using tidal volumes of 10 mL·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.
10. Introduce a gastric tube, perform suction of gastric fluids, properly fix the tube, and connect it to a collection bag. Carefully close the animal's eyes to prevent dryness of eyes during anesthesia.
11. Maintain anesthesia by continuous infusion of fentanyl (10 µg·kg^-1·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 µg) 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.
12. Administer a baseline infusion rate of 10 mL·kg^-1·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.
13. Gently clean the pig'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.

2. Probe placement

1. Place the animal in the right lateral position, and flex the animal's back to widen the space between the vertebrae.
2. Surgically expose the paravertebral area for the preparation of spinous processes and vertebral arches (Figure 1A).
3. 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).
4. 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.
5. Carefully fix the probe with sutures (Figure 1D) and use padding to prevent dislocation or kinking of the probe.
6. 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.
7. Place a saline-filled syringe on the needle, and carefully introduce the needle with constant pressure on the fluid-filled syringe.
8. 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.
9. 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.
10. Carefully fix the drainage with sutures, and connect it to the cerebrospinal fluid drainage system.
11. Expose the skull behind the left ear, and carefully perform a drill hole trepanation of the skin using a 6 mm drill attachment.
12. 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.
13. 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.
14. Reconnect the probes, and re-check signal quality.
15. 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.
16. Calibrate Flux as per unit (PU) with 2-point calibration.
    1. Press Enter to open the menu and select the analogue output setting.
    2. Use the displayed conversion factor (5.0 V = 1000 PU) to calibrate Flux with 2-point calibration for use with the synchronic acquisition software.
    3. Select Return to return to the previous menu, and select Measurement to continue with measurement.
    4. Open the synchronic acquisition software. Select zero all inputs from the Setup menu. Connect all inputs with the used devices and probes.
    5. 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.
17. Start continuous cerebrospinal fluid drainage with a target pressure of 10 mmHg and drainage volume of 20 mL·h^-1.

3. Catheter placement

1. Expose both femoral arteries.
2. 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' scissor, and introduce the guide wire.
3. Introduce the guide wire further, ensuring resistance-free insertion and avoiding any kinking of the wire; introduce the catheter over the wire.
4. Fix the catheter with sutures.
5. 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.
6. Place a 5 mm flow-probe on the left femoral artery, and test the signal quality by connection to the flowmeter.
7. Close both groins with sutures.
8. Expose the right carotid artery as well as the right internal jugular vein for placement of 8 Fr. introducer sheaths.
9. For catheter placement, proceed in the same manner as described in 3.2-3.4.
10. Connect the side-lumen of the carotid artery introducer sheath to the basic pressure monitoring and pulmonary thermodilution hardware for arterial pressure measurement.
11. Introduce a pressure-tip catheter into the ascending aorta, and verify the position by connection to the amplifier and synchronic acquisition hard- and software.
12. 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.
13. 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
14. 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.

4. Surgical preparation

1. 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.
2. Connect the catheter to a urine collection bag displaying the urine amount in mL.
3. Increase the FiO₂ to 1.0, and re-administer 0.1 mg·kg^-1 pancuronium intravenously.
4. 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.
5. 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.
6. Carefully mobilize the apex of the left lung, and divide the left lateral part of the diaphragm to facilitate surgical exposure.
7. 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·kg^-1 hydroxyethyl starch colloid if hemodynamic stabilization is needed.
8. Place an overhold around the descending aorta to ensure proper exposure (Figure 2C).
9. 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.
10. Attach a vessel loop around the descending aorta, distal to the flow probe to mark the area of aortic cross clamping.

5. Assessment and data acquisition

1. Zero all catheters and level catheters using fluid-filled lines placed at the right atrial level.
2. Place needle ECG electrodes and connect them to the synchronic acquisition hard- and software.
3. Assessment of trans-cardiopulmonary thermodilution as well as aortic flow and pressure measurements have been previously described ³⁴.
4. 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.
5. 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.

6. Experimental protocol

1. Perform baseline measurements (M0).
2. Perform hemodynamic optimization using volume-loading steps of 7 mL·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 <15%.
3. Repeat measurements (M1) after completion of hemodynamic optimization.
4. Induce ischemia/reperfusion for a total of 48 min of supra-celiac aortic cross-clamping by placing an aortic clamp at the marked area.
5. 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.
6. Continue aortic cross-clamping after each interval after a maximum of 5 min or after normalization of femoral artery flow.
7. Perform manual inflow occlusion of the inferior vena cava to prevent blood pressure increases of > 100 mmHg mean arterial pressure.
8. Administer bolus injections of norepinephrine or epinephrine during the clamping phase, if needed, to prevent decreases in mean arterial pressure below 40 mmHg.
9. Repeat measurements at the end of the 30-min clamping interval prior to reperfusion (M2).
10. Gradually open the clamp to ensure hemodynamic stability. Close the clamp if blood pressure drops too quickly and allow stabilization.
11. Administer 7 mL·kg^-1 of hydroxyethyl starch colloids as well as additional bolus injections of 10-20 µg 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.
12. Repeat measurements 1 h after reperfusion (M3).
13. Repeat hemodynamic optimization as described under 6.2, and repeat measurements (M4).
14. Perform final measurements 4.5 h after the induction of ischemia/reperfusion (M5).

7. Euthanasia

1. Administer 40 mmol of potassium chloride intravenously for euthanasia to induce ventricular fibrillation and asystole.
2. Terminate ventilation and remove all catheters.

8. Organ harvesting

1. Place the animal in a prone position, and remove the needle probes as well as the drainage.
2. Expose the spine by skin incision and removal of muscle tissue using a scalpel and forceps.
3. 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.
4. 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.
5. Store the spinal cord in 4% formalin until further utilization for histopathological evaluation or microsphere quantification.

9. Statistical analysis

1. Use statistical software.
2. Ensure normal distribution by inspection of histograms and log-transform variables if necessary.
3. 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.
4. Use baseline adjustments.
5. Specify models with fixed effects for variable baseline and measurement point. Consider measurement point as repeated measures within animals.
6. Report p-values of fixed effects for measurement point for each parameter.
7. 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.
8. 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.
9. Express variables as mean (95% CI). Express animal weight as mean ± standard deviation.
10. Present unadjusted p-values.

Results

All six animals survived until the completion of the protocol. Animal weight was 48.2 ± 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...

Discussion

SCI induced by spinal cord ischemia is a major complication of aortic repair with tremendous impact on patient outcome^{1,2,3,4,10,11,12}. Microcirculation-targeted therapies to prevent and treat SCI are most promising. The protocol provides a reproducible method for real-time spinal cord micro...

Materials

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