A Mouse Model of Spinal Cord Hypoperfusion with Immediate Paralysis Caused by Endovascular Repair of Thoracic Aortic Aneurysm

Abstract

Background:

A clinically relevant mouse model of thoracic endovascular aortic repair-induced ischemic spinal cord injury has been lacking since the procedure was first employed in 1991. We hypothesized that ligation of mouse intercostal arteries would simulate thoracic endovascular aortic repair-induced ischemic spinal cord injury and behavioral deficit. We aimed to create a mouse model of thoracic endovascular aortic repair-induced spinal cord hypoperfusion by ligating five pairs of mouse intercostal vessels.

Methods:

Mice were divided into sham (n=53) and ligation (n=60) groups. We double ligated three pairs and single ligated two pairs of thoracic intercostal arteries in adult C57BL/6 mice. A laser doppler probe was used in vivo on the spinal cords and intercostal arteries to document the extent of arterial ligation and spinal cord hypoperfusion. The Basso Mouse Scale for Locomotion, histological studies, and electron microscopy demonstrated post-ligation locomotive and histopathological changes.

Results.

Ligation induced a significant and instantaneous drop in blood flow in the intercostal arteries (% change) (mean=−63.81, 95%CI: −72.28 to −55.34) and the thoracic spinal cord (% change) (mean=−68.55, 95%CI: −80.23 to −56.87).

Paralysis onset was immediate and of varying degree, with behavioral deficit stratified into three groups: 9.4% exhibited severe paralysis, 37.5% moderate paralysis, and 53.1% mild paralysis at day 1 (n=32; p<0.001). Mild and moderate paralysis was transient, gradually improving over time. Severe paralysis showed no improvement and exhibited a higher mortality rate (83%; n=15/18) compared to moderately (33%; n=6/18) and mildly (24%; n=6/25) paralyzed mice (p<0.001). The overall ligation group survival rate (84%; n=46/55) was significantly lower than the sham group (100%; n=48/48) with p=0.003.

Conclusion:

Our mouse model generates reproducible spinal cord hypoperfusion and accompanying histopathological ischemic spinal cord damage. The resulting anatomical changes and variable behavioral deficits mimic the variability in radiological and clinical findings in human patients.

Introduction:

Left untreated, thoracic aortic aneurysm disease (incidence, 0.0053%; prevalence, 0.16%) can be lethal.¹ However, both repair methods—traditional open repair and thoracic endovascular aortic repair—cause the severe complication of ischemic spinal cord injury. Although the thoracic stent endograft deployed during thoracic endovascular aortic repair, a less-invasive alternative for abdominal aortic aneurysm since 1991² and thoracic aortic aneurysm since 1994,³ gained Food and Drug Administration approval in 2005,⁴ a clinically relevant mouse model for mechanistic study of thoracic endovascular aortic repair-induced ischemic spinal cord injury has been lacking for over 30 years.

Moulakakis et al. estimated the open repair-induced ischemic spinal cord injury complication rate as 8.26% (95%CI: 6.95 to 9.67) with paraparesis and paraplegia rates of 3.61% (95%CI: 2.25 to 5.25) and 5% (95%CI: 4.36 to 5.68), respectively.⁵ Scali et al. estimated the thoracic endovascular aortic repair-induced ischemic spinal cord injury incidence rate as 3.7% (n=422/11473; transient, 1.6% [n=179]; permanent, 2.1% [n=243]).⁶ Thoracic endovascular aortic repair-induced Ischemic spinal cord injury cases will continue to rise with the increased use of this preferred less-invasive procedure.

Existing open repair models provide pathophysiological evidence of post-ischemia reperfusion spinal cord injury.^7–11 However, high cost, difficulty of replicating large animal models (e.g., pigs, dogs), high morbidity and mortality rates, and the inability to genetically manipulate the animal¹² limit their utility. Lang-Lazdunski et al. attempted to resolve these limitations with the first open repair-induced ischemic spinal cord injury mouse model.¹³ However, the challenging surgical approach, high mortality, and limited survivability (< one week) precluded universal utilization. Awad et al. resolved these drawbacks with a simplified, reproducible, clinically relevant open repair mouse model enabling consistent use.¹²

Awad et al. later deployed stents into three canine aortas to examine the histopathological differences between open repair- and thoracic endovascular aortic repair-induced ischemic spinal cord injury and proved that the ischemic spinal cord injury mechanism differs between repair methods—reinforcing the need for a thoracic endovascular aortic repair unique mouse model.¹⁴ However, until now, simulation of thoracic endovascular aortic repair-induced ischemic spinal cord injury was confined to rabbits,¹⁵ dogs,¹⁶ pigs,¹⁷ and sheep¹⁸ with the same utility issues found in large-animal open repair models, making them ill-suited for the elucidation of the neuropathological mechanisms of thoracic endovascular aortic repair-induced post-ischemia hypoperfusion.

We aimed to create a simple, reproducible, clinically relevant mouse model of ischemic spinal cord hypoperfusion to facilitate mechanistic studies. We hypothesized that double-ligating three pairs and single-ligating two pairs of C57BL/6 adult mouse intercostal arteries would simulate thoracic endovascular aortic repair-induced ischemic spinal cord hypoperfusion with paralysis severity variability and mortality rates comparable with human patients. This configuration resulted from a pilot study of varying ligation strategies.

Radiological, behavioral, and histopathological studies demonstrate that our model induces spinal cord hypoperfusion and causes spinal cord histopathological ischemic damage resulting in variable behavioral deficit mimicking human patient findings. Our model paves the way for mechanistic study of thoracic endovascular aortic repair-induced spinal cord hypoperfusion and in vivo development of neuroprotective drugs and therapeutics.

Materials and Methods:

Animals

Adult C57BL/6 mice (age range between 16–20 weeks for both males and females) were purchased from Jackson Laboratories (Bar Harbor, ME). The mice were randomly assigned to the sham and ligation groups in each of the pilot and final configurations: pilot using 3 double ligation (sham, n=5; ligation, n=5); pilot using 5 double ligation (sham, n=17; ligation, n=22); pilot using 6 double ligation (sham, n=2; ligation, n=2); final configuration using 3 double ligation with 2 single ligation (sham, n=48; ligation, n=55). In the final configuration, one sham mouse was excluded from the study due to hind limb injury unrelated to the surgery. Supplemental Digital Content 1 illustrates the experimental design animal tree with total number of animals and the detailed number of animals in each group. Neither body weight nor distribution of each sex differed significantly across the groups. All male mice (n=58/58) were > 23 g; 59% (n=31/53) of the female mice were ≤ 23 g and 42% (n=22/53) were > 23 g. Mice were randomly chosen for either the sham or ligation surgery using the procedure described infra in Random Assignment and Blind Testing.

All procedures used aseptic technique and all mice were housed in HEPA-filtered Bio-clean units. All procedures complied with the National Institute of Health Guide for the Care and Use of Laboratory Animals and were approved by The Ohio State University’s Animal Care and Use Committee (IACUC approval number 2007A0195).

Anesthesia preparation:

The core temperature for all mice was maintained between 33.0 – 33.5°C on a temperature-controlled surgical platform (World Precision Instruments, Sarasota, FL) (see Supplemental Digital Content 2, discussing the choice of core body temperature during anesthesia preparation). Mice were anesthetized using 4% isoflurane (100 mL/minute O₂) in 96% O₂ for induction, then maintained at 2% isoflurane (100 mL/minute O₂) in 98% O₂ throughout all surgical procedures. Two layers of disinfection, first with liquid surgical scrub and second with 70% ethanol were performed to clean the surgical site after shaving the hair. A lubricated mouse tracheal intubation cannula was inserted into the trachea (Hugo Sachs VK32) through the mouth after exposing the trachea with a vertical ventral midline incision in the neck extending slightly past the ear pinna. Then the larynx and trachea were exposed by retracting the overlaying fat, muscle, and submaxillary gland. A mouse ventilator (Hugo Sachs-Harvard Apparatus Minivent, Hollinston, MA) (tidal/stroke volume = 250 μL; rate = 230 ventilations/min) was used for ventilation. The throat incision was closed with surgical glue and a single suture, and the tracheal cannula was fixed in place to the lip of the mouse with a surgical clip. Then the mouse was injected subcutaneously at the site of the incision with bupivacaine (0.1 mg/mouse in 0.02 mL) and gentamycin (0.1 mg/mouse in 0.1 mL). Five to 10 minutes later, the chest wall was opened.¹²

Surgical procedure

Intubated mice were placed in a right lateral position and the left forelimb was extended beneath the mandible and secured to the surgical platform with adhesive tape to expose the lateral thoracic cage beneath the left scapula. A small transverse incision was made underneath the left shoulder, then a blunt dissection of the subcutaneous fat to expose the underlying rib cage was performed. Using scissors, skin was cut longitudinally (rostral to caudal) to the lower costal cartilage, then the fat overlying the rib cage was bluntly dissected and moved to either side exposing the entire left thoracic cage. The intercostal muscles between the 8th and 9th ribs were cut with scissors, exposing the lateral pleura. With the incision wide open, the lower lobe of the lung was pushed out of the field using retractors while the skin and fat on each side were held away from the incision site using lateral hooks. The subcutaneous fat was kept moist throughout the procedure using normal saline. Using forceps while viewing through a 10x power microscope lens, the fully exposed descending aorta was bluntly dissected in a rostral to caudal direction while carefully exposing the intercostal arteries (8th to 10th). A second incision was made in the muscle between the 6th and 7th rib and retractors were used to hold it wide open and to push the lungs away from the field. While viewing through a 16x power lens, the exposed descending aorta was bluntly dissected a second time in a rostral to caudal direction to increase the exposure and visibility of the intercostal arteries (6th through 10th).

Continuing with the 16x power lens, the intercostal arteries were sutured (ligated) bilaterally using 9–0 nonabsorbable nylon sutures (ETHILON) starting with the lower-most vessel (10th intercostal pair) and ending with the 6th intercostal pair; the 8th, 9th, and 10th artery pairs were double ligated, and the 6th and 7th pairs were single ligated. This configuration was chosen after conducting a pilot study where we first tried double ligating the 11th-6th (in the six double-ligation group), the 10th-6th (in the five double ligation group), and the 10th-8th (in the three double ligation group) (see Supplemental Digital Content 3, discussing the rationale for the use of double ligation). The rib cage and intercostal muscles were then sutured at the two incision sites using 6–0 polypropylene after absorbing any fluid or blood leak at the incision site. Finally, subcutaneous fat was put back after being moistened, and the skin was sutured using 5–0 polypropylene sutures. Figure 1a illustrates an intercostal ligation and ligature. Sham control mice underwent the exact same surgical procedure without ligation of the intercostal arteries.

Figure 1.

(A) Intercostal vessel ligation. Arrows indicate the ligature. (B) Laser doppler probe measuring blood flow directly from the exposed mouse spinal cord. Arrow indicates placement of the probe over the spinal cord.

Post-surgical care

All animals recovered spontaneously from anesthesia within 10–20 minutes on the surgical platform while their core temperatures were maintained. The mice were extubated and placed into new clean cages maintained on a warmer set at 34.5°C for two to five days. Bladder care by a gentle manual evacuation of the bladder was performed every 12 hours for the duration of the experiment to prevent urine retention and infection. Mice were given 0.5 mL 10% dextrose in water subcutaneously twice daily and were also given prophylactic antibiotics—gentamycin (0.1 mg/mouse in 0.1 mL) and Baytril (0.2 mg/mouse in 0.2 ml) twice daily subcutaneously through the first seven days post-surgery. Mice were kept on a regular diet supplemented by a Stat high caloric diet (PRN Pharmaceutical, Pensacola, FL) throughout the study. Animal welfare, post-operative pain control, and humane endpoints were ensured per the protocols established by the Animal Care and Use Committee at The Ohio State University (see Supplemental Digital Content 4, discussing post-surgery pain management and post-operative pain control agents, concentration, dose, and frequency).

Tissue Perfusion Measurements

Tissue perfusion data was obtained from five male mice using a Transonic Type N24 needle probe connected to a BLF22 laser doppler system (Transonic Systems Inc., Ithaca, NY). Twenty-four hours before measuring tissue perfusion, the mice underwent dorsal laminectomy of T8-T10, the skin was stapled and the mice left to recover after an injection of gentamycin (0.1 mg/mouse in 0.1 mL). The next day, the mice were anesthetized with 2% isoflurane in 100% oxygen, the spinal cord was exposed, and the laser doppler probe was stabilized over a marked location on the spinal cord using a micromanipulator (Fig. 1b). Spinal cord perfusion measurements were obtained before ligation. Mice were then placed in a left lateral position, the chest was opened, and the intercostal arteries were surgically ligated (as discussed supra in Surgical Procedure).

Intercostal artery blood flow measurements were taken before and after ligation by applying the laser doppler probe tip on the surface of the vessels. The mouse was then switched back to the prone position and the tip of the laser doppler probe was placed on the spinal cord at the same marked location to acquire post-ligation spinal cord tissue perfusion measurements. In accordance with the protocol approved by The Ohio State University Institutional Lab Animal Care and Use Committee, all mice were sacrificed at the end of the tissue perfusion measurement. Powerlab data acquisition system and LabChart software (ADInstruments Inc., Colorado Springs, USA) were used to record the output signals from the laser doppler system with output signals set to the tissue perfusion flow unit (mL/min/100 g tissue) and blood velocity unit (m/s/100 g tissue).

Several factors were addressed before use of the laser doppler tissue perfusion probe. The probe was immobilized during the entire measurement to ensure proper measurement of reflected light. This was ensured by surgical implantation of the probe via micromanipulator. A mark for doppler measurement location ensured consistent pre- and post-ligation probe placement. Care was taken to avoid arterial occlusion by the probe and duplicate readings were taken to ensure consistent measurement upon placement. Additionally, the same ambient room lighting was maintained during the experiment to prevent confounding the results.

Behavioral assessment

Bilateral hindlimb function was blindly assessed with the Basso Mouse Scale for Locomotion¹⁹ at six hours, days one, two, three, five, eight and two weeks after injury (see infra Random Assignment and Blind Testing for a discussion of the blind procedure used for the behavioral assessment).The Basso Mouse Scale, a 10-point scale (0–9) with operational definitions of key locomotor features, quantifies the rate and extent of functional impairments in mice with neurovascular injury¹⁹ (see Supplemental Digital Content 5, explaining the Basso Mouse Scale assessment method).

Tissue harvesting

Mice were anesthetized with a mixture of 120–150 mg/kg ketamine (which is 1.5 times the regular dose of ketamine for sedation) and 5–10 mg/kg of xylazine (which is the regular dose for sedation), and then transcardially perfused with 25 ml 0.1 M phosphate-buffered saline, followed by 50 ml of 10% formalin (see infra Random Selection and Blind Testing for a discussion of the randomization procedure used to select mice for tissue harvesting). Spinal cords were removed and postfixed in 10% formalin for five days.

Histopathology

Spinal cords from an equal number of male and female mice subjected to sham surgery or surgical ligation were collected at 2 days (sham, n=6; ligated, n=6) and 8 days (sham, n=6; ligated, n=6). The Comparative Pathology and Digital Imaging Shared Resource (CPDISR) of The Ohio State University Comprehensive Cancer Center performed all pathology procedures. An experienced prosection technician trimmed spinal cords in the area of experimental interest (lower thoracic and upper lumbar segments where the hypoperfusion was expected) to identify regions of ischemia. Additional spinal cord tissue immediately proximal to the areas of expected ischemia was also trimmed and processed. Spinal cords were further post-fixed for 48 hours in 10% formalin. Tissues were routinely processed for histopathology on a Leica Peloris 3 Tissue Processor (Leica Biosystems, Buffalo Grove, IL) and embedded in paraffin. The spinal cords were sectioned at an approximate thickness of 4–5 micrometers to produce multiple cross sections per region of interest, and batch stained with hematoxylin and eosin (H&E) on a Leica ST5020 autostainer (Leica Biosystems, Buffalo Grove, IL) using a routine and quality-controlled protocol.

Immunohistochemistry to detect the presence of astrocytes positive for Glial Fibrillary Acidic Protein (GFAP, Dako Product #Z0334, Agilent, Santa Clara, CA; 1:5000 = 0.58 μg/mL), microglia positive for Ionized Calcium Binding Adaptor Molecule 1 (Iba1, Novus Biologicals Product # NB100–1028, Centennial, CO; 1:1000 = 0.5 μg/mL), oligodendrocytes positive for oligodendrocyte transcription factor (Olig2, Abcam Product #109186, Boston, MA; 1:400 = 0.325 μg/mL), and apoptotic cells and bodies positive for cleaved caspase 3 (Cell Signaling Technologies Product #9661, Danvers, MA; 1:180 = 0.26 μg/mL) was performed using a Lab Vision 360 Automatic Immunostainer (Thermo Scientific, Kalamazoo, MI) with optimized and quality controlled protocols specific to each primary antibody. All immunohistochemistry procedures follow optimized, validated protocols for paraffin embedded tissues with confirmed quality control for antibody specificity, including positive, negative, no primary, and isotype controls for all antibodies.

All slides were evaluated by an American College of Veterinary Pathologists board-certified comparative pathologist (Dr. Corps) using a Nikon Eclipse Ci-L Upright Microscope (Nikon Instruments, Inc., Melville, NY). Representative photomicrographs were taken using an 18 megapixel Olympus SC180 microscope-mounted digital camera and cellSens imaging software (Olympus Life Science, Center Valley, PA).

Electron microscopy

Fresh spinal cord samples (n=3) were fixed for a minimum of two hours at room temperature using 2.5% glutaraldehyde in 0.1M phosphate buffer, pH 7.4 in a volume of fixative that was greater than ten times the total volume of the tissue sample being fixed. Samples were stained using a LYNX tissue processor (Electron Microscopy Sciences) according to the following protocol: rinsed three times for five minutes each in 0.1M phosphate buffer, stained for one hour in 1% osmium tetroxide in 0.1M phosphate buffer, rinsed three times in distilled water, stained in aqueous 1% uranyl acetate for one hour, rinsed three times in distilled water, then dehydrated in an ethanol series (50%, 70%, 80%, 95%, 95%, 100%, 100%, 100%) where each step from 50% to 95% was 10 minutes long and each 100% step was 15 minutes.²⁰ Samples were then changed to acetone for ten minutes and the resin infiltration series began according to the following schedule: 1:1 acetone:resin mix without accelerant for one hour, 1:2 acetone:resin mix without accelerant for 1 hour, two changes of 100% resin mix with accelerant over two to six hours. The resin mix used was mixed according to the manufacturer’s recommendations using the Eponate 12 kit (Ted Pella, Inc.).

Samples were transferred into silicone molds containing freshly made resin mix and cured for 24 to 48 hours at 60˚C. Samples were then sectioned using an LC7 ultramicrotome (Leica) at 90 nm thickness and post-stained using aqueous 1% uranyl acetate for three minutes, rinsed in water, and stained with Reynold’s lead citrate²¹ for two minutes. Samples were then rinsed in water a second time and imaged using an FEI Spirit G2 Biotwin transmission electron microscope (FEI).

Blood pressure and blood gas measurement

To study changes in the blood pressure and blood gases during surgery, 15 adult C57BL/6 mice were randomly assigned to three groups (baseline group, n=5; ligation group, n=5; and sham group, n=5), with three males and two females in each group. The baseline group underwent intubation and ventilation using the same anesthesia procedure and the same ventilator settings as the ligation and sham groups but did not receive any surgical intervention.

Intraoperative blood pressure measurement using a noninvasive tail cuff method in mice.

CODA High Throughput Noninvasive Blood Pressure system (Product # CODA-HT8, Kent Scientific Corporation) was used with the following settings: maximum occlusion pressure of 250 mmHg; deflation time at 20s; minimum volume at 15 μL; display style “One Channel per Graph;” acclimation cycles at 5; number of sets at 1; time between sets at 30s; cycles/set at 20; time between cycles at 5s.

CODA Dual Channel Blood Pressure System (Kent Scientific Corporation) was used with the following setting: maximum occlusion pressure: 250 mmHg, deflation time at 20s. Animals were fixed on the heating surgery stage set at 33.5°C. Five to 15 acclimation cycles were performed prior to start of surgery until reaching tail volume ≥ 15 μl. During the procedure, the room temperature was stable (20–23°C) and the experiment took place well away from air conditioning vents or fans. Eight intraoperative blood pressure readings for each mouse were taken: two readings at the beginning of the experiment (after intubation and ventilation in the baseline group; after opening the chest in the ligation and sham groups); four readings mid-experiment (one hour after intubation and ventilation in the baseline group; during ligation of the intercostal vessels in the ligation group; and 30 minutes after opening the chest in the sham group); and two readings at the end of the experiment (90 minutes after intubation and ventilation in the baseline group; after complete ligation of the intercostal vessels in the ligation group; and one hour after opening the chest in the sham group). An example intraoperative blood pressure reading from this experiment is shown in Supplemental Digital Content 6.

Intraoperative blood gas analysis.

At the end of the experiment and after recording the last two blood pressure readings, a cardiac puncture was performed by advancing a 25G needle into the apex of the heart for blood sampling. 0.2 ml of blood was collected and used to measure pH, the partial pressure of oxygen (PaO₂) and partial pressure of carbon dioxide (PaCO₂), bicarbonate concentration (HCO₃⁻), hematocrit (Hct) and hemoglobin concentration (Hb) using an Abbott i-Stat hematology blood gas analyzer.

Random Selection and Blind Testing

All mice were subjected to preoperative behavioral assessment using the Basso Mouse Scale five days after arrival to The Ohio State University from Jackson laboratory. Any mouse with pre-operative abnormal behavior was not included in the study. Then mice were randomly given numbers, and all mice were subjected to the same preoperative care.

Each surgery day, four mice were randomly selected to undergo surgery as follows: one cage for females and one cage of males were taken to the surgery station. Then one mouse (of either sex) was randomly selected and then underwent surgery (ligation or sham), then another mouse of the opposite sex was subjected to the same surgery. This pattern was then repeated for two additional mice to receive the other type of surgery, such that four mice were operated upon on the same day under the same conditions to have two ligated mice (one male and one female) and two sham mice (one male and one female).

The behavioral assessments were performed blindly. Prior to the performance of each behavioral assessment, all living mice to be tested that date (ligated and sham) at all timepoints (e.g., one day, three days, one week, etc.) were randomly assigned alphabetical letters (A, B, C, etc.). The behavioral assessments were performed, and Basso Mouse Scale scores assigned with the observers blind and unaware of the status of each mouse. Once all assessments were complete at the end of the testing session, the blinding was resolved by matching each mouse’s number to the randomized letter and the corresponding Basso Mouse Scale score.

A random selection process was followed when sacrificing the mice and for tissue harvesting and processing. Histopathology, electron microscopy, and statistical analysis were each performed by different team members that were blindly given the samples labeled as “group A” and “group B,” and were therefore unaware of which group was the sham or ligation.

Statistical analysis

For surgical results, categorical variables are summarized as frequencies (percentage) and compared between groups using chi-squared tests or Fisher’s exact tests where relevant. Continuous variables are summarized as means (standard deviation) and compared between groups using Student’s independent t-tests. Mouse blood flow and velocity changes were compared between time points using paired t-tests. Overall survival was compared between study groups using a Kaplan-Meier survival plot and Log rank test. Hypothesis testing was 2-sided and p-values < 0.05 were considered statistically significant. SAS version 9.4 (SAS Institute, Cary, NC) was used to conduct the statistical analysis.

For the Basso Mouse Scale data, the statistics could not be run in a single repeated measure, as animals were deducted at planned endpoints as the study progressed. Using an established analysis strategy for multiple endpoints,²² we applied a repeated measures ANOVA for the intervals of 6h, 1d and 2d and a second ANOVA for 3d, 5d and 8d. Tukey’s Post Hoc was used to compare the sham vs. ligand at each timepoint. The figures discussed infra in the Results display all animals tested at each timepoint. The use of parametric statistics for the Basso Mouse Scale and similar locomotor rating scales is supported by Abelson and Tukey²³ and confirmed to be statistically appropriate for spinal cord injury.²⁴

For laser doppler tissue perfusion, all statistics were calculated by paired t-test. All data were calculated using GraphPad Prism 8.4.3. One mouse and all associated data were excluded (Mouse 3) from the study due to abnormal velocity detected with doppler ultrasound prior to ligation resulting in a positive outlier using the Grubbs Test.

Results:

Survival probability post-surgery

In the pilot study, there was a significant difference in paralysis outcomes and mortality among the three ligation strategies at day 1. (Table 1). Increasing the number of arteries ligated and/or the number of ligations on each artery increased the severity of paralysis—but at a cost of survivability. (Table 1). Double ligation of three pairs of intercostal arteries produced only mild paralysis (Basso Mouse Scale ≥ 7.5) at 24 hours with 0% mortality (n=0/5; p<0.001). Double-ligation of six pairs of intercostal arteries caused 100% severe paralysis (Basso Mouse Scale ≤ 4) at 24 hours with 100% mortality (n=2/2; p<0.001). Double-ligation of five pairs of intercostal arteries produced 59% (n=13/22) of mice with severe paralysis at 24 hours, but the mortality rate was still very high at 72% (n=16/22; p<0.001).

Table 1. Comparison of severity of paralysis at day 1 based on Basso Mouse Scale score between ligation groups.

Severe paralysis (Basso Mouse Scale ≤ 4), moderate severity paralysis (Basso Mouse Scale >4 & < 7.5), and mild severity paralysis (Basso Mouse Scale ≥ 7.5) at 24 hours. Also shown are the mortality rates based on severity of paralysis for all ligated mice and comparison of mortality by ligation group.

	6 double ligation (n=2)	5 double ligation (n=22)	3 double & 2 single ligation (n=32)	3 double ligation (n=5)	p-value	Mortality by Severity	p-value
Basso Mouse Scale ≤ 4 at day 1 (Severe paralysis)	2 (100%)	13 (59%)	3 (9%)	0 (0.0%)	<0.001	15/18 (83%)	<0.001
Basso Mouse Scale > 4 & <7.5 at day 1 (Moderate paralysis)	0 (0.0%)	6 (27%)	12 (38%)	0 (0.0%)		6/18 (33%)
Basso Mouse Scale ≥ 7.5 at day 1 (mild paralysis)	0 (0.0%)	3 (14%)	17 (53%)	5 (100%)		6/25 (24%)
Mortality by Ligation	2(100%)	16 (72%)	9 (28%)	0 (0%)	<0.001

Based on these results, our final model used the configuration of double-ligation of three pairs of intercostals (8th-10th) and single ligation of two pairs of intercostals (6th and 7th) which resulted in three stratified groups of behavioral deficits, 9% (n=3/32) severe paralysis, 38% (n=12/32) moderate paralysis, and 53% (n=17/32) mild paralysis and an overall mortality rate of 28% (n=9/32) at day 1 (p<0.001). (Table 1).

Overall, considering both the pilot and final configurations, we found that severely paralyzed mice (Basso Mouse Scale ≤ 4) had the highest mortality rate (83%; n=15/18) compared to moderately paralyzed mice (4 < Basso Mouse Scale < 7.5) (33%; n=6/18) and mildly paralyzed mice (Basso Mouse Scale ≥ 7.5) (24%; n=6/25) with p<0.001. (Table 1).

In the final model configuration, we also found a significantly lower overall survival rate for the ligation group (84%; n=46/55) compared to the sham group (100%; n=48/48) with p=0.003. No significant difference in survival probability between the ligation and sham groups occurred at 6-hours and 12-hours post-operation but was apparent at post-operative days 2, 8, and 14 (see Supplemental Digital Content 7, table showing survival probability by study group and time). Figure 2 illustrates the overall survival comparison between ligation and sham groups over a two-week period. Mice that died in the first three days post-surgery expired due to respiratory difficulties. Mice that died after three days post-surgery expired due to excessive weight loss or infection.

Figure 2.

Overall survival comparison between the ligation group (blue; n=55) and the sham group (red; n=48) over a two-week period using a Kaplan-Meier survival plot and Log rank test (Log-Rank test p= 0.004).

Tissue Perfusion Measurement

Intercostal artery and spinal cord blood flow (ml/min/100g tissue) and red blood cell velocity (m/s/100g tissue) was measured in male C57BL/6 mice by laser doppler tissue perfusion (n=4). Baseline intercostal arteries blood flow (ml/min/100g tissue) (mean=67.66, 95%CI: 59.32 to 76, p=<0.001) and velocity (m/s/100g tissue) (mean=4.57, 95%CI: 2.52 to 6.62, p=0.030) were higher than the thoracic spinal cord blood flow (ml/min/100g tissue) (mean=31.83, 95%CI: 27.13 to 36.53, p=<0.001) and velocity (m/s/100g tissue) (mean=1.30, 95%CI: 1.19 to 1.41, p=<0.001). After intercostal arteries ligation, a significant and instantaneous drop in blood flow (% change) (mean=−63.81, 95%CI: −72.28 to −55.34) and velocity (% change) (mean=−78.14, 95%CI: −91.15 to −65.13) occurred in the arteries (Fig 3A-D). The intercostal artery occlusion also caused significant instantaneous drop in blood flow in the thoracic spinal cord (% change) (mean=−68.55, 95%CI: −80.23 to −56.87) and an instantaneous reduction in blood flow velocity (m/s/100g tissue) in the thoracic spinal cord (mean=−62.43, 95%CI: −78.2 to −46.66) (Fig 3E-H). (See Supplemental Digital Content 8, table showing mean blood flow and velocity in the intercostal arteries and spinal cord pre- and post-ligation and the percent change in the intercostal arteries arteries and spinal cord blood flow pre- and post-ligation).

Figure 3.

Localized blood hemodynamics measured in n=4 mice by Laser Doppler tissue perfusion pre- and post-arterial ligation for intercostal artery and spinal cord blood flow (A, E) with calculated % change in flow from pre- to post-ligation (B, F); intercostal artery and spinal cord blood velocity (C, G) with calculated % change in flow from pre- to post-ligation (D, H). Data presented as mean ± S.D. and analyzed by paired T-test. *p<0.05; **p<0.001

Ligation of five pairs of intercostal arteries induced spinal cord hypoperfusion resulting from a variable but significant drop in spinal cord blood flow. Figure 3 and Supplemental Digital Content 8 show the mean blood flow and velocity in the intercostal arteries and spinal cord pre- and post-ligation and the percent change in the intercostal and spinal cord blood flow and velocity pre-and post-ligation (n=4). Figure 4A shows an example of the change in the intercostal artery blood flow and velocity pre- and post-ligation in one mouse. Figures 4B and 4C show examples of the change in the spinal cord blood flow and velocity pre- and post-ligation in two other mice. For example, one mouse experienced an 83.6% decrease in spinal cord blood flow post-ligation largely related to low spinal cord baseline blood flow levels (Fig. 4B, mouse 4). In contrast, another mouse had comparatively higher spinal cord baseline blood flow and experienced only a 55.1% decrease in spinal cord blood flow post-ligation (Fig. 4C, mouse 2).

Figure 4. LabChart data from Laser Doppler Analysis of Intercostal Artery and Spinal Cord blood flow.

(A) Mouse 5 intercostal artery change in blood flow (red) and velocity (green) pre- and post-intercostal ligation. (B) Mouse 4 spinal cord flow (red) and velocity (green) pre- and post-intercostal ligation. Notice the marked drop in spinal cord blood flow. (C) Mouse 2 spinal cord change in blood flow (red) and velocity (green) pre- and post-intercostal ligation. The drop in spinal cord blood flow is noticeably less than that in Mouse 4.

Histopathology

Spinal cords from both male and female mice undergoing ligation were clearly distinguished from those undergoing sham surgical procedure without ligation. Within 2-day and 8-day ligation groups there was variability in the severity and distribution of lesions, and changes in the white matter were less pronounced than those in the grey matter unless necrosis was present. (Fig. 5).

Figure 5. Ligation induces changes in the grey and white matter of the spinal cord at 2 and 8 days post-surgery.

Mild histologic lesions are evident in the spinal cord at 2 days post-ligation (E-H) compared to sham surgery (A-D). E. Hypocellularity in the dorsal horns (circle) is accompanied by multifocal swollen axon cylinders (spheroids, filled arrows) in adjacent white matter spinal tracts and hypercellularity in adjacent ventral grey matter. F. There are increased Iba-1+ microglia including large activated microglia near the margins of hypocellular and hypercellular grey matter. G. Numerous large GFAP+ astrocytes are present in the dorsal horn grey matter around the central canal. H. There is a slight increase in Olig2+ oligodendrocytes in the grey matter and ventral white matter. More severe lesions affecting most of the grey matter are present at 8 days post-ligation (M-P) compared to cords subjected to sham surgery (I-L). M. Grey matter in ligated mice is hypercellular with decreased neurons, increased glia and multifocal proliferative small caliber blood vessels (Q, open arrows). Surrounding white matter tracts contain numerous swollen axon cylinders (spheroids, arrows). N. Grey matter contains numerous Iba-1+ microglia that frequently exhibit large, round, activated morphology. O. Activated, GFAP+ astrocytes in the grey matter and surrounding white matter (arrows) with the exception of areas containing proliferative blood vessels, where there are few GFAP+ astrocytes (R). P. Increased Olig2+ oligodendrocytes in the spinal grey matter of ligated mice. Scale bars in A-P = 100 μm, taken at 100x total magnification. Insets in E-H and M-P taken at a total magnification of 200x with scale bars = 50 μm. Q and R taken at a total magnification of 400x with scale bars = 20 μm.

Grey Matter Lesions

Prominent hypercellularity was present in both dorsal and ventral horn grey matter in ligated mice, particularly at 8 days post-ligation. Cells contributing to hypercellularity included large numbers of Iba-1+ microglia (Fig. 5N), GFAP+ astrocytes (Fig. 5O) and fewer Olig2+ oligodendrocytes (Fig. 5P). Microglia frequently had small, dense, round to oval, peripheralized nuclei with large, round to oval cytoplasmic surface area. Microglia in sham animals exhibited the expected morphology of thin arborizations and a small cell body. Astrocytes in and around affected grey matter had large, open oval nuclei with increased cytoplasm and plump, shorter cytoplasmic processes rather than the small soma and small, central round nuclei observed in spinal cords from mice subjected to sham procedure. Frequently, increased numbers of microglia and astrocytes exhibiting these respective morphological changes were present in viable tissue along the margins of foci of cell death and tissue loss. These changes were more prominent in spinal cords examined at 8 days post-ligation compared to 2 days post-ligation (Fig. 5E-H). Interestingly, in 3 spinal cords collected 2 days following ligation, the dorsal horns were markedly hypocellular with almost no neurons, microglia, astrocytes or oligodendrocytes compared to sham spinal cords at the same time point (Fig. 5E-H). Unique in the ligated spinal cords examined at 8 days post-procedure were multifocal clusters of numerous small, proliferative capillaries with variable luminal diameter. These capillaries were frequently found in the dorsal horn in areas of marked hypercellularity and glial neuroinflammation (Fig. 5Q and R).

White Matter Lesions

Lesions in the white matter were highly variable in spinal cords examined at 2 days or 8 days post-ligation but were less pronounced than lesions in the grey matter. These changes included swollen axon cylinders (spheroids) in dilated myelin sheaths (Fig. 5E and M) and occasional small foci of neuroinflammation (including increased microglia and astrocytes, Fig. 5F and N, G and O) adjacent to blood vessels or foci of cell death. Spheroids were observed in all white matter tracts but often present in only 1–3 funiculi in an individual spinal cord. Variability in the size and severity of axon swelling was also common, with spheroids of varying size present in individual spinal cords at both time points.

Evaluation of cleaved caspase 3 immunohistochemistry at 2 and 8-days timepoints revealed rare, individualized cells with positive punctate cytoplasmic or defined nuclear staining. Positively labeled cells were rare, with a single positive cell present per spinal cord section (data not shown).

Electron Microscopy

In each sample, including the control, some fixation and expansion artifacts were present, therefore we did not include those criteria in our assessment of ischemia-related damage.²⁵ Rather, we relied on the relative severity of extracellular space and the presence of broken cell membranes relative to the control sample as our criteria for damage assessment. Because standard fixation induces cellular swelling²⁶ and extracellular space here was not preserved, we expect to see very little extracellular space. After intercostal ligation, extracellular space was substantially increased relative to the control 24-hour post-ischemia (Fig. 6a, b). In contrast, by 48 hours, less extracellular space was present (Fig. 6c), but more cellular damage, i.e., ruptured membranes, was observed.

Figure 6. Transmission electron microscopy of sections from ischemia-induced mouse spinal cord segments showing the degrees of damage after ischemia.

Ruptured cell membranes and an increase in extracellular space (arrowheads) are visible (b) 24 hours after ischemia, and (c) at 48 hours after ischemia, less extracellular space is visible, but more cellular damage, i.e., ruptured membranes, are visible,and (a) no ischemic damage is present in the control. Scale bar: 1um.

Behavioral assessment

In all ligated mice, hind limb motor deficit of a variable degree was observed immediately after recovery from anesthesia. The 6-hour time point showed the lowest Basso Mouse Scale score. Over the two-week period, there was gradual increase of locomotor function of the mice that began 24 hours post-ligation and continuously improved from 3d to 14d (Fig. 7). The ligation group had significantly lower Basso Mouse Scale scores relative to sham through 8 days (p < 0.01; Fig. 7A).

Figure 7. Basso Mouse Scale Locomotor Scores show improvement through time and wide variability.

(A) Basso Mouse Scale locomotor scores (average + standard deviation (SD)) for the sham and ligation groups averaged across all animals (sham, n=48; ligation, n=55) tested at each timepoint. The ligation group is significantly lower through 8d (*; p<.01 using separate repeated measures ANOVA for 6h-2d and 3d-8d with Tukey’s Post Hoc). (B) Wide variability in behavioral performance is displayed in a box plot, showing that mice scores are not tightly bunched at the mean. Black circles are individual values. Box edges show 25th and 75th percentile range, and whiskers show 10th and 90th percentile range. Points beyond that are shown in triangles and are designated as outliers. Dashed line is mean, and solid line is the median.

After ligation, a large range of locomotor deficits occurred at each time point. The greatest variability occurred early after ligation such that at 6 hours Basso Mouse Scale scores ranged from 0 (no hind limb movement) to 8 (nearly normal locomotion) (Fig. 7B). The large Basso Mouse Scale score drop in our ligated mice at 6 hours is non-attributable to anesthesia effects because sham mice returned to normal or near-normal behavior at this timepoint (Fig. 7A).

By 14 days, the variability had narrowed substantially. Variability occurred both as underperforming and outperforming the group mean. A single mouse scored Basso Mouse Scale 0 at the initial test and showed no improvement over time, maintaining full hind limb paralysis until sacrificed at the 8-day time point. The relationship between severity of deficits and mortality was examined for high severity with Basso Mouse Scale scores ≤4, moderate severity with Basso Mouse Scale scores >4 and <7.5, and mild severity with Basso Mouse Scale scores ≥ 7.5. The Fisher’s exact test showed that Basso Mouse Scale scores were significantly different across the four configurations (p<0.001). (Table 1). For example, 100% of the 6 double ligation group (n=2/2), 59% of the 5 double ligation group (n=13/22), 9% of the 3 double and 2 single ligation group (n=3/32), and 0% of the 3 double ligation group (n=0/5) had Basso Mouse Scale scores ≤ 4 at day 1 with p<0.001.

Blood pressure and blood gas measurement

In contrast to the blood pressure and blood gases results of Awad et al.’s aortic cross clamp mouse model,¹² there was no significant difference in the intraoperative blood pressure or blood gases among the baseline, ligation, and sham groups.

The parameter of significance to determine whether the ligation surgery has any effect on blood pressure is the mean arterial pressure (MAP). Between the three groups, there was no significant difference in the mean intraoperative MAP. The mean intraoperative MAP was 76.2 mmHg (95% CI: 72.8 to 79.5) for the baseline group (n=5), 78.2 mmHg (95% CI: 74.8 to 81.7) for the ligation group (n=5), and 79.6 mmHg (95% CI: 76.1 to 83.0) for the sham group (n=5). The baseline-ligation difference was −2.0 mmHg (−6.8, 2.8) non-significant with p=0.40. The baseline-sham difference was −3.4 mmHg (−8.1, 1.4) non-significant with p=0.17. The ligation-sham difference was −1.3 mmHg (−6.2, 3.5) non-significant with p=0.59.

No significant difference was observed among the three groups in the mean pH, mean partial pressure of carbon dioxide (PaCO₂), mean bicarbonate concentration HCO₃^-, mean partial pressure of oxygen (PaO₂) or the mean hemoglobin concentration (Hb) (see Supplemental Digital Content 9, table showing the intraoperative blood gas measurements from the baseline, ligation and sham groups.). The mean pH was 7.4 (95% CI: 7.4 to 7.4) for the baseline group (n=5), 7.4 (95% CI: 7.4 to 7.4) for ligation group (n=5) and 7.4 (95% CI: 7.4 to 7.5) for the sham group (n=5). The baseline-ligation difference was −0.0 (95% CI: −0.0 to 0.0) non-significant, with p=0.97. The baseline-sham difference was −0.0 (−0.1, 0.0) non-significant with p=0.16. The ligation-sham difference was −0.0 (95% CI: −0.1 to 0.0) non-significant, with p=0.17.

The mean partial pressure of carbon dioxide (PaCO₂) was 35.9 mmHg (95% CI: 34.0 to 37.8) for the baseline group (n=5), 35.2 mmHg (95% CI: 33.3 to 37.2) for the ligation group (n=5), and 33.8 mmHg (95% CI: 31.8 to 35.7) for the sham group (n=5). The baseline-ligation difference was 0.7 mmHg (95% CI: −2.1 to 3.4), non-significant with p=0.60. The baseline-sham difference was 2.1 mmHg (−0.6, 4.9), non-significant with p=0.12. The ligation-sham difference was 1.4 mmHg (95% CI: −1.3 to 4.2), non-significant with p=0.27.

The mean bicarbonate concentration (HCO₃^-) was 21.9 mmol/l (95% CI: 20.7, 23.1) for the baseline group (n=5), 21.5 mmol/l (95% CI: 20.3 to 22.7) for the ligation group (n=5), and 21.9 mmol/l (95% CI: 20.7 to 23.1) for the sham group (n=5). The baseline-ligation difference was 0.4 mmol/l (−1.3, 2.1), non-significant with p=0.62. The baseline-sham difference was −0.0 mmol/l (−1.7, 1.7), non-significant with p=1.00. The ligation-sham difference was −0.4 mmol/l (95% CI: −2.1 to 1.3), non-significant with p=0.62.

The mean partial pressure of oxygen (PaO₂) was 300.2 mmHg (95% CI: 260.3 to 340.1) for the baseline group (n=5), 315.0 mmHg (95% CI: 275 to 354.9) for the ligation group (n=5), and 326.8 mmHg (95% CI: 286.9 to 366.7) for the sham group (n=5). The baseline-ligation difference was −14.8 mmol/l (95% CI: −71.3 to 41.7), non-significant with p=0.60. The baseline-sham difference was −26.6 mmol/l (95% CI: −83.1 to 29.9), non-significant with p=0.33. The ligation-sham difference was −11.8 mmol/l (95% CI: −68.3 to 44.7), non-significant with p=0.66.

The mean hemoglobin concentration (Hb) was 13.8 g/dl (95% CI: 13.4 to 14.2) for the baseline group (n=5), 13.5 g/dl (95% CI: 13.1 to 14.0) for the ligation group (n=5), and 12.9 g/dl (95% CI: 12.5 to 13.3) for the sham group (n=5). The baseline-ligation difference was 0.3 g/dl (95% CI: −0.3 to 0.9), non-significant with p=0.35. The baseline-sham difference was 0.9 g/dl (95% CI: 0.3 to 1.5), non-significant with p=0.009. The ligation-sham difference was 0.6 g/dl (95% CI: −0.0 to 1.2), non-significant with p=0.05.

Discussion:

We’ve demonstrated that the number and degree of ligated intercostal arteries is directly proportional to the severity of paralysis and inversely proportional to survivability. Our final configuration—3 double and two single ligations—caused clear motor deficits below the ligation level without high mortality for ≥14 days. Survival rate ranged between 100% (6-hour and 12-hour groups) to 63% (2-week group).

Our model demonstrates four behavioral similarities with humans. First, immediate paralysis occurring in our model is like the immediate paralysis occurring in humans after thoracic endovascular aortic repair,²⁷ and contrasts with the delayed paralysis occurring in the open repair mouse model⁹ and humans.²⁷

Second, the wide behavioral deficit variability (severe to mild) occurring in response to the same ligation procedure replicates the clinical profile in patients post-thoracic endovascular aortic repair.⁶ (Fig. 7).

Third, gradual paralysis improvement occurred throughout the two-week follow-up period with sustained improvement starting at day 3 (Fig. 7A). Variable degree of deficit and gradual improvement mimic the recovery process in humans who develop thoracic endovascular aortic repair-induced paralysis where mild paralysis was transient and improved with time, while severe paralysis was permanent.⁶

Deficit variability showing improvement may be attributed to extensive collateral circulation and adaptation to blockage. Etz et al. proved that in humans, spinal cord perfusion pressure drops markedly then recovers gradually during the first several hours post-extensive segmental artery sacrifice.²⁸ In the same study, they showed that all but 1 patient, with remarkably low postoperative spinal cord perfusion pressure and experienced paraparesis, regained normal spinal cord function. Etz, et al. also proved the existence of an extensive collateral network around pig spinal cords.²⁹ Griepp et al. demonstrated that the collateral network around the pig spinal cord enlarges within 24 hours after extensive ligation of segmental arteries, that maximum collateral circulation expansion is achieved by the fifth day post-ligation and correlated these results to Etz et al.’s findings in humans.³⁰ All three studies demonstrate similarities between human and pig spinal cord collaterals and human and pig temporal response to segmental artery ligation or sacrifice. We found similar behavioral/locomotor findings as those in humans with gradual improvement of motor function over the first few days in mildly and moderately paralyzed mice, and permanent motor function loss in severely paralyzed mice.^6,28 No known study demonstrates the existence of collateral circulation around the mouse spinal cord. However, similar dysfunctional findings in our mice with human and pig studies lead us to conclude that anatomical similarities—i.e., a collateral network around the spinal cord that gradually improves after intercostal artery ligation—should also exist in mice.

A critical foundation of our model included restricted spinal cord blood flow confirmation. We extended laser doppler measurement of spinal cord blood flow from rats to mice³¹ to collect in vivo blood flow measurements from the exposed spinal cord and intercostal arteries. The percent change variability in spinal cord blood flow among our ligated mice is explained by the existence of collaterals around the spinal cord and the degree of patency of these collaterals that varies among the ligated mice.^29,30 Further, the degree of the collateral’s patency and the pre-ligation baseline spinal cord blood flow variability explain the variability in the post-ligation percent decrease in spinal cord blood flow, which explains spinal cord tissue damage variability and the resultant behavioral paresis among the ligated mice. We therefore postulate that baseline spinal cord blood flow evidences the extent of collateral network development around the spinal cord—where a relatively low baseline blood flow demonstrates a less developed network, and a comparatively higher baseline blood flow demonstrates a more extensively developed network.

Thus, mice with more developed and patent collaterals will be associated with a lesser drop in blood flow, mild behavioral deficit, and progressive recovery over time, whereas mice with less developed and patent collaterals will be associated with a marked drop in blood flow and severe behavioral deficit. Therefore, the pre-thoracic endovascular aortic repair degree of patency and development of collaterals around the spinal cord may predict post-procedure paralysis severity and permanence, which may prove to be important clinical screening characteristics.

Fourth, severely paralyzed mice had the highest mortality rate (83%; n=15/18) compared to moderately paralyzed (33%; n=6/18) and mildly paralyzed mice (24%; n=6/25) (p<0.001). This relationship aligns with 2020 national incidence and mortality data reported for thoracic endovascular aortic repair.⁶ Patients who developed reversible spinal cord injury symptoms had significantly worse 1-year survival rates than those without spinal cord injury (transient spinal cord injury, 80% [95% CI: 73% to 87%]; no spinal cord injury, 87% [95% CI: 86% to 88%]; log-rank p=0.1) but significantly better survival than those with permanent spinal cord injury symptoms (permanent spinal cord injury, 54% [95% CI: 47% to 61%]; transient spinal cord injury, 80% [95% CI: 73% to 87%]; p<.0001).⁶

Histopathological spinal cord changes in ligated mice also reflect ischemic change, severity variability, and precise localization of lesions. Apparent activation of astrocytes and microglia occurred post-ligation based on classic morphological changes in other sterile injury models.^32–34 A more modest increase in Olig2+ oligodendrocytes was present in spinal cords examined 8 days post-ligation compared to sham control spinal cords, but no appreciable change in Olig2+ cells was present in spinal cords examined 2 days post-ligation (Fig. 5). Electron microscopic images differentiated abnormal cellular processes—broken cellular membranes and diffuse tissue damage after ligation. This supports our conclusion that the ligated animal’s behavioral deficit is due to spinal cord ischemic damage and is not a mere side effect of the surgical approach.

Despite lesion variability, a pattern of changes suggesting a temporal continuum emerged. In the grey matter of spinal cords at 2 days post-ligation, there is a paucity of cells, particularly in the dorsal horn but also extending multifocally into the ventral horns. These areas are surrounded by activated microglia and astrocytes and are multifocally accompanied by white matter lesions including histological evidence of axonal injury. In spinal cords 8 days post-ligation, there is marked gliosis diffusely in the grey matter and in multifocal perivascular regions in the white matter composed predominantly of reactive microglia and astrocytes. This is frequently accompanied by proliferation of small capillaries (Fig 5). A single spinal cord was characterized by loss of architecture and paucity of cells with replacement by cellular debris in approximately 60% of the examined section. This spinal cord corresponded to severe paralysis and Basso Mouse Scale score of zero at 8 days post-procedure. Of interest is the lack of cleaved caspase 3 immunolabeling. It is possible that apoptosis is a component of the model but is missed via immunohistochemistry at the examined time points or based on the immunohistochemistry findings at 2 days, apoptosis may not represent the predominant mechanism of cell death in this ischemia model.

These spatially variable lesions are consistent with MRI findings in humans with thoracic endovascular aortic repair-induced spinal cord injury that showed variability in location of the lesion in the spinal cord across the anterior gray matter and anterior portion of posterior white column, central cord lesion, and posterior gray matter, and posterior white column.³⁵ This difference in the location and extent of infarction area can be explained by collateral conception.^29,30 The blood flow decrease in the anterior spinal artery caused by ligation will be partially compensated by flow from the collaterals in and around the spinal cord.^29,30 The area of the critical drop in tissue perfusion within the spinal cord varies among ligated mice due to the variability in degree of collateral development and patency. Collateral conception together with ischemia-induced inflammation can explain the proliferation of small capillaries at the infarct area margins.

Electron microscopy further demonstrates ischemic spinal cord tissue damage. Because hypoperfusion-induced tissue damage results in tissue that might contain abnormal (swollen or shrunken processes) or a diffuse type of damage not limited to cracking or breaks in the sample, we differentiated this type of damage from the mechanically based cracking damage (Fig. 6). Abnormal cellular processes, broken cellular membranes, and diffuse tissue damage were more prevalent in the ligated samples. These findings further support our conclusion that the ligated animal’s behavioral deficit is caused by spinal cord ischemic damage and is unrelated to the surgical approach.

Our study has three limitations. Our initial focus was development of a ligation strategy to achieve the desired paralysis phenotype with an acceptable mortality rate, precluding a pre-experiment sample size calculation (see Supplemental Digital Content 10). Histopathological results showed posterior spinal cord lesions, suggesting that sensory tests may have revealed additional mice demonstrating deficit; multi-modal sensory testing is recommended for future studies. Finally, no autopsies were performed on expired mice to establish actual cause of death.

We presented a mouse model with confirmed spinal cord hypoperfusion and spinal cord lesion histological topography caused by ligation of the intercostal arteries, which causes immediate paralysis that mimics in both severity and reversibility human paralysis following thoracic endovascular aortic repair. This model has good translational potential to support development of neuroprotective drugs and therapeutics for ischemic spinal cord injury following thoracic endovascular aortic repair.