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A novel swine model of subarachnoid hemorrhage-induced cerebral vasospasm
Correspondence Address: Source of Support: None, Conflict of Interest: None DOI: 10.4103/neuroindia.NI_357_16
Objective: One of the most serious complications following subarachnoid hemorrhage (SAH) is delayed cerebral ischemia (DCI) secondary to symptomatic vasospasm. An animal model mimicking post SAH vasospasm is essential for enabling the translation of newer technologies from the conceptual phase to animal studies, and eventually to clinical trials. Various animal models of DCI following SAH have been reported, with canine models being the most common. Due to the similarity of the swine cardiovascular system and its dimensions to the human's system, the main objective of this study was to establish a consistent and quantitatively representative model of SAH-induced vasospasm in swine. Keywords: Delayed cerebral vasospasm, subarachnoid hemorrhage, swine model, volume and timing of hemorrhage
Aneurysmal subarachnoid hemorrhage (aSAH) is the result of a ruptured intracranial aneurysm. The most devastating complication of aSAH is symptomatic vasospasm, which causes delayed cerebral ischemia (DCI) and may result in severe morbidity and mortality.[1] As effective treatment for cerebral vasospasm is limited,[2] there is a need for extensive research to develop reliable experimental models, and thereby effective treatment modalities. Animal models provide the basis for clarifying and studying the complex pathogenesis of vasospasm and DCI. In particular, it provides a tool for screening potential medical therapies. SAH can be simulated by endovascular puncture of a cerebral artery, subarachnoid clot placement (usually used in primate models), or injection of blood or other vasospastic substances into the subarachnoid space. These techniques have been validated in mouse, rat, rabbit, cat, dog, pig, goat, and primates.[3],[4],[5] Models of higher species such as canines and primates are preferred due to their resemblance to human models. The canine model described by Varsos et al.,[4],[6] has gained the widest acceptance in vasospasm research. This model involves double injection of blood into the basal cistern. However, preclinical canine and primate models are rarely used at present as they are expensive and infrequently approved for research.[7] Swine anatomy is similar to that of humans, specifically with respect to its cardiovascular system. In 1984, Takemae et al.,[8] first reported a swine experimental model of vasospasm. Since then, several studies have used swine as a model for SAH [Table 1].[9],[10],[11],[12],[13],[14],[15] However, reports to date have focused on a specific methodological issue, rather than comparing various parameters using the same experimental setup. The goal of the current study was to establish a reproducible, reliable, and quantitatively representative model of SAH-induced vasospasm in domestic swine. We examined, in a consistent cohort of pigs, the relationship between varying volumes and timing of hemorrhage, subsequent angiographic cerebral vasospasm, and clinical outcome. As the swine model is widely used for translational cardiovascular research, we believe this work constitutes an important benchmark for the development of pharmacological and medical devices and interventions.
Animals and anesthesia The study protocol was approved by the Sheba Medical Center Institutional Ethics Committee and was completed in accordance with the Guide for the Care and Use of Laboratory Animals.[16] The study was performed at the Neufeld Cardiac Research Institute, Sheba Medical Center (Tel Hashomer, Israel) and the Lahav C.R.O research unit (Kibbutz Lahav, Israel). Twelve 3.5-month-old domestic female pigs (Sus scrofa domestica), weighing 57 ± 3 kg were studied. The animals were divided into two groups. Group 1 (n = 5; animals 1–5) [Table 2] were injected with various arterial blood volume combinations. Group 2 (n = 7; animals 6–12) [Table 2] were used to implement the optimal parameters for SAH induction determined in Group 1.
All procedures were performed under general anesthesia, induced by intramuscular (IM) injection of ketamine (10 mg/kg), xylazine (2.0 mg/kg), and intravenous diazepam (0.1 mg/kg). An endotracheal tube was utilized for mechanical ventilation (2.5% isoflurane, oxygen and air mixture). Electrocardiogram (ECG), heart rate (HR), blood pressure (BP), O2 saturation, and body temperature were continuously monitored throughout the procedure. Post-procedure, the animals were treated with IM marbofloxacin 120 mg/day for 3 days and a single dose of IM dipirone 1000 mg. Animal #7 was treated with dexamethasone 40–100 mg/day IM for 3 days due to suspected cerebral edema. Angiography Baseline angiography was performed on day 0 (prior to SAH induction), and repeated at least once between day 7 and 14 following SAH induction [Table 2]. A 23-cm 4 Fr sheath (Brite Tip, Cordis, Ireland) was inserted into the femoral artery using the Seldinger technique, followed by a 100-cm 4 Fr diagnostic catheter (MP A1 tempo 4, Cordis, Ireland) that was advanced into the ascending pharyngeal artery. A single arterial phase 0° anteroposterior, 17° caudal baseline angiogram of the circle of Willis and basilar artery (BA) was done with maximal magnification (Philips Integris H5000F, Philips Medical Systems North America Co., Washington, USA). A total of 8 mL of diluted (1:3) contrast medium (Urografin, Scherring AG, Germany) was injected over 3 seconds. The 4 Fr catheter was used as a reference for calibrating radiographic magnification. Induction of subarachnoid hemorrhage Nonheparinized autologous blood was injected intrathecally into the subarachnoid space twice, i.e., at day 0 (following baseline angiography) and on day 2 using dissimilar injection volumes. The animal was placed in a recumbent “fetal” position with an approximate 30° head down tilt. The femoral sheath was left in place and an 18-gauge 3.5” spinal needle (KDL Ltd. Shanghai, China) was inserted intervertebrally between C2 and C3 to access the subarachnoid space. Cerebrospinal fluid (CSF) drainage allowed space for blood injection. Autologous, nonheparinized arterial blood drawn from the femoral artery was injected into the subarachnoid space at a rate of 10 ml/min. Group 1 was injected twice with equal blood volumes ranging from 6–18 ml, with a 2-day interval between the injections. Group 2 was injected with 12 ml on day 0 followed by 15 ml on day 2 [Table 2]. The needle was removed and the animal was moved to a prone position with an approximate 30° head down tilt for 30 minutes to ensure cephalic distribution of the blood in the subarachnoid space.[8] The animal was then moved to a supine position, the femoral sheath was removed with manual compression, and anesthesia was terminated. Arterial diameter measurement The diameters of the intracranial internal carotid artery (ICA), external carotid artery (ECA), and basilar artery (BA) were measured using an image analysis software (Philips Integris H5000F, Philips Medical Systems North America Co., Washington, USA) with a 4 Fr diagnostic catheter placed in the ascending pharyngeal artery as a reference. The mean diameter was obtained from five consecutive measurement points along the selected artery segment. Vasoconstriction in all arteries was measured as percentage change in diameter relative to baseline. We used a 5% decrease in diameter as the threshold for vasospasm. Student's t-test was used for statistical evaluation and P < 0.05 was considered to be statistically significant. Clinical evaluation Clinical behavior scores were recorded daily from day 0 to day 14 using a modified scoring table based on clinical outcome analysis used in previous studies.[17] Points were assigned based on scales for appetite, behavior, posture, walking, and eye movement. Appetite and behavior was scored as active and finished meal = 2 points; active only with stimulation, left meal unfinished = 1 point; lying down, scarcely ate = 0 points. Posture and walking was scored as standing and walking straight and stable = 2 points; seated position, unstable walking due to ataxia (side deviation) = 1 point; prone position, unable to walk, paresis = 0 points. Eye movement was scored as normal, parallel = 2 points; nystagmus = 1 point; unilateral paralysis = 0 points.
ICA vasoconstriction was recorded with the following injected volume combinations: 10 ml in each of the two injections, which yielded 34% and 12% arterial narrowing (left and right, respectively); 12 ml in each of the two injections, which yielded 14% arterial constriction on both sides; 12 ml followed by 15 ml injections, which yielded 24% and 18% arterial constriction (left and right, respectively). The maximal arterial constriction ratio of the left ICA and right ICA in Group 1 animals is illustrated in [Figure 1]a.
Vasoconstriction progressed linearly for most volume combinations. The baseline angiographic (day 0) ICA diameter in all animals was 1.3–1.4 mm. Seven days following the first injection, all volume combinations tested showed a reduction in vessel diameter with the exception of the 12 ml double injection, which yielded an increase in vessel diameter on day 10. All other volume combinations (two 10 ml or the 12 ml followed by 15 ml) demonstrated a continuous decrease in vessel diameter until day 14. As the 12 and 15 ml volume combination showed the maximum linear decrease in vessel diameter, we contemplated it as the optimal volume combination. Thus, this injected volume combination was used in Group 2. Utilizing this injection volume, the maximal vasospasm was recorded on day 12 ± 2. Although the reduction in vessel diameter did not reach a plateau at day 14, we did not evaluate it beyond 14 days to maintain relevance to the time course in humans where vasospasm is mostly resolved during this period. The progression of vasoconstriction along time course of the study is illustrated in [Figure 1]b. The optimal volume and time parameters described above (12 ml injection on day 0 followed by 15 ml injection 2 days later) were implemented in Group 2. Vasospasm was assessed on day 12 ± 2. [Figure 2]a shows the ICA vessel diameters of the individual animal and the mean constriction [Figure 2]b. Most animals demonstrated a decrease in absolute vessel diameter bilaterally. The magnitude of constriction was progressive, showing 16% constriction on day 7 and 22% on day 12 ± 2 in the left ICA, and 6% and 16%, respectively, in the right ICA.
Evaluating the changes in vessel diameter of the anterior cerebral artery (ACA) revealed a significant decrease on day 12 ± 2, compared to day 0 (0.57 mm vs. 0.87 mm; P < 0.05, left ACA; and, 0.57 vs. 0.83, P < 0.05, right ACA; [Figure 3]a and representative angiography image [Figure 4]). The magnitude of mean vasoconstriction was 34% and 27% on day 12 ± 2 (left and right ACA, respectively; [Figure 3]b. The basilar artery demonstrated variable results and did not exhibit a significant reduction in diameter probably due to its initial small diameter (average of 0.54 mm). This data suggest that vasoconstriction, as induced by the testing parameters chosen for Group 2, is not restricted to a definite vessel or side, but rather the spasmogenic effect is widespread throughout the circle of Willis.
Behavioral monitoring following experimental SAH was used to assess clinical outcome [Table 3]. There was no direct correlation between the volume of blood injected and the neurological impact, as demonstrated in animals #1–5. When performed under optimal conditions (animals 5–12; 12 ml injection followed by 15 ml injection), there was no neurological effect in 50% of the animals. Of the remaining 4 animals, two (#5 and 6) demonstrated normal appetite and eye movement, unstable posture, and left side deviation of the head while walking from day 3 after the first injection. One animal (#7) demonstrated a decreased appetite until day 5, which resulted in 13% weight loss, and the animal was unable to stand or walk due to hind limb paresis on both the sides from day 3. The animal was treated with steroids on day 3 and exhibited slight improvement on day 8. The last animal (#11) had a permanent decreased appetite from day 4 and unstable walking due to forelimb weakness beginning from day 8. No correlation was found between the severity of the neurological outcome and the magnitude of the angiographic vasospasm.
Subarachnoid hemorrhage-related vasospasm induction in animal models Cerebral vasospasm is a major determinant of the prognosis of SAH.[15] An efficient and reliable animal model of SAH-induced vasospasm is highly valuable as a platform for understanding the underlying mechanisms of pathophysiology and for developing pharmacological agents and medical interventional therapies. Consistent and feasible models were published and reviewed in the dog, mouse, rabbit, rat, and primate,[3],[4],[18] but not in the cat, swine, or goat. Several studies have been performed using swine as an animal model for SAH.[8],[9],[10],[11],[12],[13],[14],[15] Nevertheless, comparative studies examining parameters of injected blood volume, timing of angiographic imaging, laterality of the effect, and clinical comparators have not been published and require further exploration.[3] Only three of the above mentioned 8 swine studies used angiographic measurements of the induced vasoconstriction in the intracranial arteries [Table 1].[10],[11],[15] These studies used different (1) injection profiles (single or double); (2) injected blood volumes; and, (3) timing of vasospasm assessment. Thus, no uniform protocol was adopted in the abovementioned studies.[3] Optimization of the experimental technique is likely to increase the clinical relevance of the vasospasm model for future experimental studies. We used a double intrathecal injection of nonheparinized autologous arterial blood into the subarachnoid space to induce SAH. As the technique used in our study was uniformly controlled (i.e., the volume of hemorrhage was the same as the injected dose, thereby eliminating uncontrolled hemorrhage secondary to arterial puncture), it has the potential to produce consistent results. Repeated intrathecal injections of blood in a double-hemorrhage canine model have been shown to produce prolonged and more intense vasospasm.[6],[19] Once exposed to the first bolus of injected blood, cerebral vessels become hypersensitive to the vasoactive stimulus in blood, which is likely due to the changes in the neurovascular plexus that chronically alters the normal reactivity of the cerebral arteries during SAH.[19] Hence, we hypothesized that the second injection was warranted in our model (two-hit hypothesis). Furthermore, larger volume of blood in the second injection is likely to generate a stronger vasospastic response. We modified the protocol of Xu et al.,[15] who used a double injection (day 1 and 3) of 6 ml (average of 0.22 ml/kg) autologous arterial blood into the cisterna magna. They obtained 33–37% vasoconstriction in the right ICA and BA 10 days after the first injection, with no neurological deficit. We defined the relationship between various injection volumes (0.1–0.3 ml/kg) and times of measurement (7–14 days after injection) affecting the severity of angiographic cerebral vasospasm on both the left and right ICA and ACA, with a rigorous clinical outcome follow-up in a swine model of SAH-induced vasospasm. We found that intrathecal injection of 12 ml followed by 15 ml of nonheparinized autologous arterial blood (2 days apart) into the subarachnoid space was the most effective and consistent hemorrhage volume for inducing bilateral mild (10–30%) to moderate (31–50%) vasospasm 12 ± 2 days after the first injection, with no associated neurological changes. Blood volume Experimental and clinical data suggested that the volume and/or the duration of exposure of blood to cerebral arteries are important determinants of vasospasm.[12],[19] Imaging studies based on computed tomographic (CT) scans in humans have shown that a larger volume of hemorrhage is a major factor in chronic vasospasm.[20] Although our angiographic data demonstrate a direct correlation between the blood volume and the severity of vasospasm of the right ICA, the left ICA did not demonstrate such a correlation. Even the highest blood volume (double dose of 18 ml each) did not induce left ICA constriction [Figure 1]a. This side variability may be due to the inability to control the spread of blood injected between the two hemispheres, and thereby lateralization of the resultant hematoma on one side. Based on the results, the blood volumes chosen for the time course analysis were the double injection of 10 ml, double injections of 12 ml, and an injection of 12 ml followed by 15 ml. Among these tested volumes, the injection combination of 12 ml followed by 15 ml of blood demonstrated a linear decrease in the right and left ICA diameter on both sides [Figure 1]b. Thus, this was considered the optimum volume combination for this model in our study. Timing of angiographic measurements Applying the above-mentioned parameters to the second group of animals (n = 7), followed by angiographic measurements of the ICA and ACA, verified that the time course of vasospasm in this swine model is consistent with the expected clinical course of angiographic vasospasm.[21],[22] Specifically, progressive vasospasm was observed between the first and second week following SAH, with the maximal vasospasm occurring at day 12 ± 2, similar to a previously reported cerebral vasospasm swine model.[15] Our model did not study the resolution of the spasm, which normally occurs within 2–3 weeks of hemorrhage. Degree of vasospasm The vasoconstriction produced in various reported SAH models was not as significant as that seen in clinically symptomatic patients.[3],[19],[23] The degree of angiographically-measured vasospasm reported in previous swine studies was 19% constriction in the basal intrathecal cerebral arteries on day 7,[11] 25% constriction in the ICA,[9] and moderate vasospasm (over 30% decrease) in the right ICA and BA on day 10 post-SAH.[15] Our vasospasm swine model yielded mild-to-moderate vasoconstriction (16–34% reduction) in the right and left ICA and ACA on day 12 ± 2 after the first injection,[9],[11],[15] which represents a comparable but slightly higher degree of vasoconstriction. The vasonstriction yielded in our swine model was limited due to the limited range of the injected blood volumes in relation to the brain and CSF volume. As we aimed to minimize other pathologies, such as increased intracranial pressure, the blood injection was not performed against pressure and the amount of blood injected was limited to the CSF volume drained spontaneously from the subarachnoid space. Another reason for the limited vasoconstriction demonstrated in our model was the absence of arterial puncture in this model with lack of endothelial cell damage, thus avoiding the release of potent vasoconstrictor, peptide endothelin-1 from endothelial cells.[24] The insignificant vasoconstriction of the right ICA (P > 0.05) may be because of the laterality of the intrathecal blood injection. In all animals, vasospasm was induced while the animal was lying on its left side. The flow of blood to the left side may explain the greater extent of angiographic vasospasm of the left ICA. In a canine model of SAH-induced vasospasm, higher reductions in vessel diameter (up to 71%) were angiographically measured in the BA.[19] The diameter of the canine BA is larger than the circle of Willis arteries, whereas the diameter of the swine BA is relatively smaller than other circle of Willis arteries. Furthermore, a BA of ~25-kg weighing dog is larger than the BA of 60-80-kg weighing swine. Hence, a reliable angiographic measurement of the swine BA diameter was not feasible in the current study. In addition, evaluation of angiographic vasospasm of the anterior cerebral arterial system is more relevant to clinical vasospasm in humans. Clinical outcome Although there is a correlation between the severity of large artery vasospasm and symptomatic ischemia in humans, there are patients with severe vasospasm who never become symptomatic, and others with modest vasospasm who not only develop symptoms but tend to develop infarction.[2] In the current model, the clinical grading system subdivided the outcome into three categories, as described in the methods. Although several previous studies evaluated the neurological deficits in a swine model of SAH-induced vasospasm,[14],[15] there is no published data describing in detail, the evolution of clinical signs in a swine vasospasm model. In the current study, there was no direct correlation between the volume of hemorrhage and the neurological outcome. Moreover, there was no correlation between the magnitude of the angiographic vasospasm and the severity of the neurological outcome, with 50% of the animals exhibiting no neurological deterioration. In the remaining animals, the clinical outcome was mild-to-moderate, with left side impairment in all the cases. In 3 out of these 4 animals, clinical manifestations appeared the day after the second injection, and only one animal (#11) exhibited clinical signs on day 5 after the second injection. The lack of procedure-related clinical events in half of the animals may be a result of: (1) the inability to replicate the precise distribution of the subarachnoid clot subsequent to intrathecal blood injection; or, (2) random differences in the pathophysiological response of individual animals. Normal eye movements and mandible posture in all the animals pointed to the absence of cranial nerve injury. As most of the clinical signs appeared within the first post procedural day, it was reasonable to assume that they were not vasospasm-related. If the cause was vasospasm, we would expect it to appear 4–6 days later.[14] In addition, the effective steroid treatment in animal #7 suggests a reduction in edema and associated intracranial pressure rather than a resolution of vasospasm. This edema may have been due to the injection, specifically the case of multiple manipulations of the spinal needle and injection of blood, which may have caused increased spinal canal pressure. One animal (#11) among the 8 animals tested, however, demonstrated clinical signs that may be attributed to vasospasm.
The use of rats and mice for studying vasospasm has become increasingly common over the past several years because of advancements in the imaging techniques, new approaches in vessel morphometry, and reduced costs related to handling of small animals. Feline, swine, and canine models of SAH and vasospasm are generally more expensive. However, in these species, cerebral vessels are larger, and are more easily subjected to angiography and angioplasty; and, the results of these experiments can be more readily extrapolated to human vasospasm.[4] Canine models of SAH are still considered superior for vasospasm studies as the murine models to study human vasospasm are disputed.[3],[5] However, a major disadvantage of using canines for cerebrovascular studies is the relatively small diameter of the circle of Willis arteries. In contrast, swine models can yield better correlation with human cerebral arteries. The limitations of the swine SAH model are the relatively thick scalp bones that obligate the administration of a higher volume of contrast material, that itself, even with a gradual and moderate injection pressure, has a dilatory effect on the cerebral arteries. The pulsatility of these arteries has a relatively major effect on the variability of diameter as measured on angiography. The current study defines a reproducible, streamlined protocol for producing consistent mild-to-moderate vasospasm over a clinically relevant time course using a swine model. This model may enable the development of new technologies for treating SAH-induced cerebral vasospasm. Specifically, implementation of standardized experimental techniques may improve the comparability between laboratories, facilitate the interpretation of results, and increase the relevance of future preclinical studies. Acknowledgment The authors are grateful to David Castel DVM for his collaboration at the animal lab of Neufeld Institute, Sheba Medical Center, Israel. We thank Dr. Iris Kulbatski and Mrs. Avital Tamari for their assistance with the submission. Financial support and sponsorship Nil. Conflicts of interest There are no conflicts of interest.
[Figure 1], [Figure 2], [Figure 3], [Figure 4]
[Table 1], [Table 2], [Table 3]
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