RU2820288C2 - Use of cftr modulators for treating cerebrovascular conditions - Google Patents

Abstract

FIELD: medicine; pharmaceutics.

SUBSTANCE: invention relates to the field of medicine and pharmaceutics and the use of the CFTR (cystic fibrosis transmembrane conductivity regulator) corrector for the prevention and/or treatment of decreased or disturbed cerebral perfusion, where the CFTR corrector is a cyclopropane carboxamide derivative selected from the group consisting of lumacaftor, tezacaftor, C18 and their pharmaceutically acceptable salts and solvates.

EFFECT: use of the invention enables to effectively prevent and/or treat decreased or disturbed cerebral perfusion.

3 cl, 19 dwg, 1 ex

Description

The present invention relates to the use of CFTR (cystic fibrosis transmembrane conductance regulator) modulators for the prevention and/or treatment of cerebrovascular conditions and corresponding methods of treatment.

Among the older population, the number of people living with heart disease is growing rapidly. The main reasons are higher incidence rates among older people and advances in medicine that have significantly improved survival rates. Over the past decade, it has become clear that increasing life expectancy is associated with significant cognitive decline (1) and is fueling an emerging “silent epidemic” of cognitive impairment (2). The impact will be significant: beyond the direct medical needs, cognitive impairment seriously impacts patients' ability to cope with their underlying illness (eg, decreased adherence to treatment) and has significant social consequences for families, friends and society at large. Thus, the true burden of this emerging epidemic cannot be quantified by simple economic indicators. What is certain is that our medical and mental health networks are not structurally or financially equipped to cope with the coming crisis.

Cognitive decline in patients with heart failure (HF) is commonly attributed to “vascular cognitive impairment,” a term that describes multiple forms of cognitive impairment (eg, memory loss, loss of executive function, confusion) that occur due to vascular impairment. As an umbrella term, the etiology of vascular cognitive impairment is diverse and multifactorial; clinical diagnosis is complex and imprecise; and there are several distinct and complex subtypes of injury (1). Most cardiovascular and cerebrovascular pathologies (including hypertension, ischemic and hemorrhagic stroke, heart disease, and diabetes) are considered major and significant causes of cognitive decline due to vascular impairment (1). Although vascular cognitive impairment encompasses many complex random mechanisms, a unifying aspect is insufficient cerebral perfusion: in this regard, recent work by the present inventors has focused on improving cerebral perfusion in experimental models of HF and subarachnoid hemorrhage (SAH) (3–6).

The problem underlying the present invention is to provide a new regimen for improving cerebrovascular conditions.

A solution to the above technical problem is provided by the embodiments of the present invention disclosed in the claims, the present description and the accompanying drawings.

We have discovered key molecular mechanisms that reduce cerebral perfusion in HF and SAH (4-6). It is noteworthy that both pathologies alter the cerebrovascular autoregulation system and, consequently, cerebral blood flow (CBF) through the same fundamental mechanism. In particular, HF and SAH induce sustained expression of tumor necrosis factor (TNF) in the cerebral artery vessel wall: TNF, localized in smooth muscle cells, acts through an autocrine/paracrine mechanism to stimulate the production of sphingosine-1-phosphate (S1P), which then causes vasoconstriction via the S1P ₂ receptor subtype (3, 4, 6). We took advantage of this mechanistic insight and successfully validated two treatments: blocking TNF (etanercept) or antagonizing S1P ₂ receptor signaling in both models eliminated HF and SAH-mediated increases in cerebrovascular vasoconstriction (i.e., measured as increased myogenic tone) ( 3, 4, 6), normalizes brain perfusion (3, 4, 6) and, therefore, reduces neuronal damage (5, 6). However, the functionality of these two options may be limited: etanercept carries significant risks associated with immune suppression (7) and may compromise blood pressure control due to effects on skeletal muscle resistance arteries (8); while antagonism of S1P ₂ receptors will undoubtedly cause unpredictable secondary effects, since these receptors mediate important and diverse functions in several organs (9).

In the search for alternative therapeutic targets, the cystic fibrosis transmembrane conductance regulator (CFTR) has emerged as an interesting candidate. Previous work by the present inventors has demonstrated that CFTR protein expression in cerebral arteries is suppressed in HF and SAH: since CFTR is an important transporter involved in the degradation of S1P smooth muscle cells (4), suppression of CFTR should increase the bioavailability of S1P and its pro-constrictive actions ( 3, 4). Indeed, vascular TNF expression, CFTR expression, and myogenic reactivity are closely correlated (3, 4, 6), fueling the mechanistic proposal of the present invention that the TNF-dependent increase in cerebral artery myogenic tone is mediated by reduced CFTR protein expression (4). In this context, CFTR becomes an attractive and logical therapeutic target for correcting myogenic reactivity in the brain microcirculation, since CFTR therapeutics are known not to impair immune function and US Food and Drug Administration (FDA) approved therapeutics are available. which increase CFTR expression/activity (10).

The present invention relates to the use of CFTR modulator compounds, i.e. According to the present invention, one or more CFTR modulators can be used to treat and/or prevent a cerebrovascular condition.

According to the present invention, a “cerebrovascular condition” is a condition that impairs, impairs, or otherwise affects the cerebrovascular system of an individual, preferably a human patient. In particular, in cerebrovascular conditions, cerebrovascular perfusion is reduced or impaired. The term "cerebrovascular perfusion" is also known as and is synonymous with, respectively, "cerebral blood flow", "cerebral perfusion" and "cerebral vascular perfusion".

According to the present invention, the CFTR modulator exerts its effect on CFTR proteins present or expressed, respectively, in cerebral vascular smooth muscle cells. In this case, the CFTR modulator normalizes cerebrovascular Erk1/2 phosphorylation and sphingosine-1-phosphate signaling and/or cerebrovascular myogenic reactivity, and/or cerebrovascular blood flow, and/or neuronal apoptosis and cerebrovascular damage, and/or corrects behavioral and learning disorders in patients with cerebrovascular conditions, preferably reduced or impaired, respectively, cerebrovascular perfusion.

Typically, a patient having a cerebrovascular condition or disease state, respectively, has a primary disease condition, typically an underlying disease, which in turn causes the cerebrovascular condition, or at least a disease condition that is associated with the cerebrovascular condition. According to the present invention, preferred cerebrovascular conditions are treated or prevented, respectively, using said one or more CFTR modulators, where the condition(s) are caused by heart disease, heart failure (HF), subarachnoid hemorrhage (SAH), sudden sensorineural hearing loss (SSHL) ), vascular dementia, hypertension, ischemic stroke, hemorrhagic stroke, heart disease, diabetes and Alzheimer's disease.

A "CFTR modulator" of the present invention is a compound that modulates the function of the CFTR protein so that it can perform its primary function, namely, create a channel for chloride (a salt component) to pass through the cell surface.

Several useful CFTR modulators in the context of the present invention are known in the art and preferably include CFTR amplifiers, CFTR correctors and CFTR amplifiers. CFTR enhancers increase the activity of CFTR channels. CFTR correctors increase cell surface expression of CFTR. CFTR amplification agents increase the amount of CFTR expressed by increasing the amount of CFTR mRNA. More preferred CFTR modulators of the present invention are drugs that have a proteostatic effect on CFTR, in particular CFTR correctors and CFTR amplifiers, especially preferably CFTR correctors. CFTR correctors are also highly preferred for use in the present invention because they exhibit very good activity even at low CFTR content.

Particularly suitable CFTR corrector compounds for use in the present invention are cyclopropanecarboxamide derivatives such as those disclosed in WO-A-2005/075435, WO-A-2007/021982, WO-A-2008/127399, WO-A-2009/108657 and WO-A-2009/123896. Other preferred CFTR corrector compounds for use in the present invention are aminoheterocyclyl derivatives, such as pyrimidine derivatives, preferably the compounds disclosed in WO-A-2010/068863, WO-A-2010/151747 and WO-A-2011/008931 derivatives aminothiazole, preferably the compounds disclosed in WO-A-2006/101740, and quinoline/quinazoline derivatives, preferably the compounds disclosed in WO-A-2012/166654, coumarin derivatives, preferably the compounds disclosed in WO-A-2014/152213, and trimethylangelicin derivatives, preferably the compounds disclosed in WO-A-2012/171954. Additional suitable CFTR corrector compounds that can be used in the present invention are disclosed in WO-A-2014/081821, WO-A-2014/081820 and WO-A-2010/066912. Additional CFTR corrector compounds and compositions thereof for use in the present invention include transglutaminase 2 (TG2) inhibitors and casein kinase 2 (CK2) inhibitors. A preferred TG2 inhibitor for use in the present invention is cysteamine. A preferred CK2 inhibitor for use in the present invention is epigallocatechin gallate (EGCG). According to a preferred embodiment of the present invention, a TG2 inhibitor, preferably cysteamine, and a CK2 inhibitor, preferably EGCG, are used in combination. Such a combination can be provided by simultaneous administration, preferably in a single composition, but it is also contemplated to simultaneously administer two inhibitors separately, namely a TG2 inhibitor and a CK2 inhibitor, each inhibitor alone or as a component of a pharmaceutical composition. In an alternative embodiment, the TG2 inhibitor and the CK2 inhibitor can also be administered sequentially (either alone or in a suitable composition). A combination of a TG2 inhibitor and a CK2 inhibitor, preferably cysteamine and EGCG, is also disclosed in US-A-2013/0310329.

Specific examples of CFTR modulators for use in the present invention include, but are not limited to, C18 (VRT-534), lumacaftor (VX-809), tezacaftor (VX-661), 4,4',6-trimethylangelicin, VRT-768, VRT-422, VRT-325, CFpot-532, Copo-22, 002_NB_28 (DBM228), DBM_003_8Cl (DBM308), as well as any combination of such compounds. The most preferred CFTR modulators of the present invention are C18 and lumacaftor. It should be understood that the present invention also relates to the use of pharmaceutically acceptable salts, solvates, esters, salts of such esters, as well as any other adduct or derivative which, when administered to a patient in need of treatment, can provide directly or indirectly a CFTR modulator for use in the present invention, either a metabolite thereof or a residue thereof.

The present invention also relates to CFTR modulator(s), as described above, for use in the treatment and/or prevention of the conditions described above. The present invention also relates to the use of CFTR modulator(s), as described above, for the preparation of a medicament for the treatment and/or prevention of the conditions described above.

Furthermore, the present invention relates to a method of preventing and/or treating the conditions described above by administering at least one (i.e., one or more) CFTR modulator to a patient in need of such treatment, preferably a human patient, more preferably a patient suffering from a cerebrovascular condition associated with or caused, respectively, by one or more conditions selected from heart disease, heart failure (HF), subarachnoid hemorrhage (SAH), sudden sensorineural hearing loss (SSHL), vascular dementia, hypertension, ischemic stroke, hemorrhagic stroke , heart disease, diabetes and Alzheimer's disease.

According to the present invention, one or more CFTR modulator(s) can be used in its/their free form. In other embodiments, the CFTR modulator(s) (i.e., at least one CFTR modulator) used in the present invention are present in a pharmaceutical composition containing said at least one CFTR modulator, typically in combination with at least one one pharmaceutically acceptable excipient, diluent, carrier and/or excipient.

The effective amount of CFTR modulator required for use in the method and applications of the present invention, i.e. the specific effective dose level for any particular patient or organism will depend on many factors, including the disorder being treated and the severity of the disorder; the activity of the particular compound used; the specific composition used; the age, body weight, general health, gender and diet of the patient; timing of administration, route of administration, and rate of elimination of the particular compound used; duration of treatment; drugs used in combination or concurrently with the particular compound used, and similar factors well known in the medical field.

Preferred doses in the context of the present invention, particularly with respect to lumacaftor and C18, are from about 0.1 to about 10 mg per kg of body weight (hereinafter referred to as “mg/kg”), preferably from about 1 to about 8 mg/kg , more preferably from about 3 to about 5 mg/kg. The dose may be administered one or more times, for example two or three times a day. Preferably, doses are administered once or twice daily.

The route of administration of the CFTR modulator is not particularly important, and the route chosen depends on the particular CFTR modulator compound or compounds employed and the subject being treated. Preferably, the CFTR modulator(s) are administered systemically, for example by oral or intravenous administration, particularly preferably by oral administration. Topical administration may also be considered, with a particularly preferred local route of administration being injection into the cerebrospinal fluid.

As used herein, the term “patient” means an animal, preferably a mammal, and most preferably a human.

The at least one CFTR modulator, preferably present in the pharmaceutical composition as described above, for use in accordance with the present invention can be administered using any amount and any route of administration effective for treating the cerebrovascular condition. In accordance with the present invention, the term "treatment" or "treatment" is to be understood to mean that the severity of the condition is at least not progressive compared to the condition for which no treatment was given, preferably the severity of the condition is not progressing, more preferably the severity of the condition is decreasing, even more preferably, the condition is substantially reduced, and especially preferably, the condition is substantially cured. Preferably, the severity of the condition in accordance with the present invention is reduced by at least 30%, more preferably by at least 50%, even more preferably by at least 70%, especially preferably by at least 90% until the condition is completely cured, which represents is the most preferred outcome of treatment in accordance with the present invention.

With respect to the cerebrovascular condition that is the target of the application and the method of the present invention, the degree of reduction in severity or the degree of cure, respectively, the cerebrovascular condition is usually assessed by neurological evaluation of the subject receiving treatment, in accordance with medical and diagnostic methods known to one skilled in the art. field of technology, preferably before treatment in accordance with the present invention, preferably at normal intervals after initiation of treatment in accordance with the present invention. Such assessments preferably include measuring cerebral perfusion using MRI (magnetic resonance imaging), for example, as described in the example below (optionally tailored to the patient being examined, eg when the patient is a human).

The present invention also relates to the use of CFTR modulators, preferably as disclosed herein, for the prevention and/or treatment of pulmonary disorders, preferably pulmonary hypertension, more preferably secondary pulmonary disorders, especially secondary pulmonary hypertension associated with or caused, respectively, by certain primary disorders such as atherosclerosis, cardiovascular diseases (cardiovascular diseases such as coronary artery disease (CHD) (angina and myocardial infarction (commonly known as heart attack)), stroke, heart failure, hypertensive heart disease, rheumatic heart disease, cardiomyopathy, cardiac arrhythmias, congenital heart disease, valvular heart disease, carditis, aortic aneurysms, peripheral arterial disease, thromboembolic disease and venous thrombosis. The present invention also provides a method for preventing and/or treating pulmonary disorders, preferably pulmonary hypertension, more preferably secondary pulmonary disorders such as secondary pulmonary hypertension associated or caused, respectively, by certain primary disorders such as atherosclerosis, cardiovascular diseases (cardiovascular diseases such as coronary artery disease (CHD) (angina and heart attack) myocardium (commonly known as a heart attack)), stroke, heart failure, hypertensive heart disease, rheumatic heart disease, cardiomyopathy, cardiac arrhythmias, congenital heart disease, valvular heart disease, carditis, aortic aneurysms, peripheral arterial disease, thromboembolic disease and venous thrombosis, comprising the step of administering an effective amount of at least one CFTR modulator, preferably CFTR modulators as disclosed herein, to a patient in need thereof.

DESCRIPTION OF GRAPHIC MATERIALS

Figure 1. Cerebral blood flow is reduced in ^CFTRΔF508 mice.

(A) Myogenic vasoconstriction is greater in posterior cerebral arteries (PCAs) obtained from cystic fibrosis transmembrane conductance regulator (CFTR) mutant ΔF508 mice compared with control wild-type (WT) littermates. (B) Posterior cerebral arteries obtained from ^CFTRΔF508 mice show an upward shift in their phenylephrine dose response. (C) However, once responses to phenylephrine are normalized to baseline tone ( _active tone - _resting tone, where _active tone represents tone at a given phenylephrine concentration and _resting tone represents tone immediately before stimulation), the dose-response relationship of phenylephrine in WT and CFTR ^ΔF508 are almost identical (two-way ANOVA P=NS). Average maximum vessel diameters at 45 mm Hg. Art. (diameter _max ) are: CFTR ^ΔF508 : 186 ± 2 µm, n = 6 from 4 mice and WT: 169 ± 8 µm, n = 5 from 3 mice; (Mann-Whitney test P=NS for diameter _max ). (D) Magnetic resonance imaging was used to measure cerebral blood flow in predefined cortical forebrain regions. Representative forebrain perfusion maps of WT and ^CFTRΔF508 mice are shown. Cerebral blood flow is significantly lower in ^CFTRΔF508 mice (n = 10) compared to wild-type littermates (n = 11); however, neither (E) cardiac volumetric flow rate (CO) (n = 5 for both groups) nor (F) total peripheral vascular resistance (TRP) (n = 5 for both groups) differed between the two genotypes. (G) In WT mice, CFTR mRNA expression is significantly higher in the posterior cerebral arteries (n = 5) compared to the skeletal muscle arteries of the cremaster (the muscle that supports the testis) (Cre; n = 6). (H) Myogenic tone of the cremaster skeletal muscle resistance artery is not altered by CFTR inhibition in vitro (100 nmol/L CFTR _(inhibitor) -172 for 30 minutes). (I) Deletion of the CFTR gene (CFTR KO), however, causes a mild but significant attenuation of myogenic tone. Average maximum vessel diameters at 60 mm Hg. Art. (diameter _max ) are: WT (panel G): 72 ± 3 µm, n = 5 from 4 mice; CFTR KO (panel H): 88 ± 4 μm, n = 6 from 4 mice; and wild-type littermates (panel H): 78 ± 4 μm, n = 5 from 2 mice. All data are presented as mean ± standard error of the mean. * denotes P < 0.05 for unpaired comparisons using the Mann-Whitney test. Abbreviations: CFTR, cystic fibrosis transmembrane conductance regulator; Cre - cremaster; KO - knockout; PCA - posterior cerebral artery; WT - wild type.

Figure 2. C18 increases wild-type CFTR protein expression and function through a proteostatic mechanism.

(A) Cerebral arteries obtained from naïve mice treated with C18 (3 mg/kg per day for 2 days; n = 5) have higher expression of cystic fibrosis transmembrane conductance regulator (CFTR) protein compared with arteries obtained from from control carriers (n = 5). (B) C18 does not affect cerebral artery CFTR mRNA expression (control n = 6, C18 n = 5). (C) C18 (6 μmol/L; 24 h) increases CFTR protein expression in baby hamster kidney fibroblast cells stably expressing human CFTR (n = 12 for both groups); (D) C18 does not affect CFTR mRNA expression in this system (n = 6 for both groups). (E) Tumor necrosis factor (TNF; 10 ng/ml for 48 hours) suppresses CFTR protein expression in primary vascular smooth muscle cells of the mesenteric artery (control: n = 8; TNF: n = 8); Co-incubation of TNF with C18 (6 μmol/L; 24 h) after 24 h of TNF incubation (i.e., 48 h TNF + 24 h C18) completely restored CFTR protein expression (n = 7). Restoration of CFTR protein expression in vascular smooth muscle cells correlates with normalization of attenuated (F)FITC-labeled sphingosine-1-phosphate uptake (measured as an increase in fluorescence intensity at 525 nm using a standard fluorescence-activated cell sorting assay; control: n = 7; TNF: n = 10; TNF + C18: n = 6) and (G) forskolin-stimulated iodide efflux (control: n = 7; TNF: n = 6; TNF + C18: n = 6). All data are presented as mean ± standard error of the mean. In panels A to D, * denotes P < 0.05 for unpaired comparisons using the Mann-Whitney test; in panels E to G, * denotes P < 0.05 for unpaired comparisons with controls using the Kruskal-Wallis test and Dunnett's post hoc test. Abbreviations: Cont - control; CFTR - cystic fibrosis transmembrane conductance regulator; TNF - tumor necrosis factor.

Figure 3. C18 restores cerebral perfusion in heart failure.

(A) Cerebral arteries obtained from mice with heart failure (HF; 6 weeks after ligation of the left anterior descending coronary artery) have reduced expression of cystic fibrosis transmembrane conductance regulator (CFTR) protein (n = 5) compared with arteries obtained from sham-operated control animals (n = 6). In vivo treatment with C18 (3 mg/kg daily for 2 days) reversed this reduction in CFTR protein expression in the cerebral artery (n = 8). (B) In vivo C18 treatment reduces myogenic tone in posterior cerebral arteries isolated from HF mice, an effect (C) not observed in posterior cerebral arteries isolated from CFTR knockout (KO) mice. Average maximum vessel diameters at 45 mm Hg. (diameter _max ) are: sham operated = 146 ± 9 µm, n = 5 from 3 mice; CH = 149 ± 8 µm, n = 6 from 4 mice; CH + C18 = 155 ± 8 µm, n = 8 from 6 mice (Kruskal-Wallis test P=NS for diameter _max ); and CFTR KO = 142 ± 5 μm, n = 6 from 3 mice; CFTR KO + C18 = 145 ± 5 µm, n = 6 from 3 mice (Mann-Whitney test P=NS for diameter _max ). Compared with sham-operated controls, mice with HF have (D) reduced cardiac volume velocity (sham-operated: n = 6; HF: n = 8; HF + C18: n = 8), (E) increased total peripheral vascular resistance ( n = 6 for all groups) and (F) reduction in mean arterial pressure (MAP) (n = 6 for all groups). (G) Shown are representative magnetic resonance imaging perfusion maps that were used to determine cerebral blood flow in the forebrain cortex. HF provokes a decrease in cerebral perfusion; C18 treatment significantly improved brain perfusion in mice with HF (sham-operated: n = 6; HF: n = 8; HF + C18: n = 8). All data are presented as mean ± standard error of the mean. For panels A and G, * denotes P < 0.05 for unpaired comparisons with sham-operated animals using the Kruskal-Wallis test and Dunnett's post hoc test; for panel B, * denotes P < 0.05 for unpaired comparisons with sham-operated animals using two-way ANOVA and post hoc analysis with Tukey's test; in panel C, groups were statistically compared using two-way analysis of variance (P=NS); and in panels E to G, * denotes P < 0.05 for unpaired comparisons with HF using the Kruskal-Wallis test and Dunnett's post hoc test. All samples in the typical Western blot image displayed in panel A are from the same film, but the CH+C18 sample was not located side by side on that film as shown in this figure. Abbreviations: Cont - control; CFTR - cystic fibrosis transmembrane conductance regulator; diameter _max is the maximum diameter of the vessel; HF - heart failure; KO - knockout.

Figure 4. C18 does not cause cerebral edema.

Shown are typical quantitative T ₂ maps that estimate brain water content as T ₂ relaxation time on magnetic resonance images. A total of nine T ₂ maps were assessed for each mouse, covering the anterior, middle and posterior brain regions. Representative images show sections of the anterior (left), middle (center), and posterior (right) brain regions of sham-operated mice (top), mice with heart failure (HF; center), and mice with HF treated with C18 (3 mg/kg per day for 2 days) (below). Neither CH nor C18 treatment in CH mice caused changes in _T2 relaxation times in any of the regions of interest in any of the slices assessed. Thus, the data were combined to obtain the mean _T2 relaxation time for the cortical and subcortical regions and presented graphically (sham: n = 7; CH: n = 7; CH + C18: n = 6; Mann-Whitney test P=NS between groups in the cortical and subcortical regions, respectively). All data are presented as mean ± standard error of the mean. Groups were statistically compared using the Kruskal-Wallis test (P = NS). Abbreviations: HF - heart failure.

Figure 5. C18 restores cerebral perfusion in subarachnoid hemorrhage.

(A) Cerebral arteries obtained from mice with subarachnoid hemorrhage (SAH; 2 days after SAH induction) have reduced expression of cystic fibrosis transmembrane conductance regulator (CFTR) protein (n = 6), compared with arteries obtained from sham-operated controls animals (n = 6). Treatment with C18 in vivo (3 mg/kg per day for 2 days) reversed this decrease in arterial CFTR protein expression (n = 6). (B) In vivo C18 treatment reduces myogenic tone in olfactory arteries obtained from SAH mice, an effect (C) not observed in olfactory arteries obtained from CFTR knockout (KO) mice. Average maximum vessel diameters at 45 mm Hg. (diameter _max ) are: sham operated = 113 ± 3 µm, n = 5 from 3 mice; SAC = 109 ± 6 μm, n = 6 from 6 mice; SAC + C18 = 104 ± 12 µm, n = 5 from 4 mice (Kruskal-Wallis test P=NS for diameter _max ); and CFTR WT = 98 ± 6 μm, n = 8 from 4 mice; CFTR KO = 110 ± 8 μm, n = 5 from 4 mice; CFTR KO + C18 = 96 ± 6 µm, n = 6 from 3 mice (Kruskal-Wallis test P=NS for diameter _max ). (D) Representative magnetic resonance imaging perfusion maps used to determine cerebral blood flow in the forebrain cortex are shown. SAH stimulates decreased cerebral perfusion; C18 treatment significantly improved brain perfusion in SAH mice (sham-operated: n = 10; SAH: n = 5; SAH + C18: n = 9). All data are presented as mean ± standard error of the mean. In panels A and D, * denotes P < 0.05 for unpaired comparisons with SAH using the Kruskal-Wallis test and Dunnett's post hoc test; in panel B, * denotes P < 0.05 for unpaired comparisons with SAH using two-way ANOVA and post hoc ANOVA with Tukey's test; in panel C, groups were statistically compared using two-way analysis of variance (P = NS). Abbreviations: CFTR, cystic fibrosis transmembrane conductance regulator; diameter _max - maximum diameter of the vessel; HF - heart failure; KO - knockout; SAH - subarachnoid hemorrhage.

Figure 6: Lumacaftor increases wild-type CFTR protein expression through a proteostatic mechanism and restores cerebral perfusion in subarachnoid hemorrhage.

(A) Cerebral arteries obtained from naïve mice treated with lumacaftor (Lum; 3 mg/kg daily for 2 days; n = 6) have higher expression of cystic fibrosis transmembrane conductance regulator (CFTR) protein compared with arteries obtained from control carriers (n = 7). (B) Lumacaftor did not affect CFTR mRNA expression in the cerebral artery (n = 5 for both groups). (C) lumacaftor (6 μmol/L; 24 h) increases CFTR protein expression in baby hamster kidney fibroblast cells stably expressing human CFTR (n = 13 for both groups); (D) Lumacaftor did not affect CFTR mRNA expression in this system (n = 6 for both groups). (E) Cerebral arteries obtained from mice with subarachnoid hemorrhage (SAH; 2 days after SAH induction) have reduced CFTR protein expression (n = 5) compared with arteries obtained from sham-operated controls (n = 6). In vivo treatment with lumacaftor (3 mg/kg daily for 2 days) reversed this reduction in CFTR protein expression in the cerebral artery (n = 6). (F) In vivo treatment with lumacaftor reduces myogenic tone in olfactory arteries isolated from mice with SAH. Average maximum vessel diameters at 45 mm Hg. Art. (diameter _max ) are: sham operated = 91 ± 3 µm, n = 7 from 4 mice, SAC = 86 ± 2 µm, n = 6 from 3 mice; NAC + Lum = 87 ± 4 µm, n = 7 of 4 mice (Kruskal-Wallis test P=NS for diameter _max ). (G) Representative magnetic resonance imaging perfusion maps used to determine cerebral blood flow are shown. SAH provokes a decrease in cerebral perfusion; treatment with lumacaftor significantly improves brain perfusion in mice with SAH (sham-operated: n = 6; SAH: n = 5; SAH + Lum: n = 6). All data are presented as mean ± standard error of the mean. In panels A to D, * denotes P < 0.05 for unpaired comparisons using the Mann-Whitney test; in panels E to G, * denotes P < 0.05 for unpaired comparisons with SAH using the Kruskal-Wallis test and Dunnett's post hoc test. In panel F* denotes P < 0.05 for unpaired comparisons with SAH using two-way ANOVA and post hoc analysis with Tukey's test. Abbreviations: CFTR, cystic fibrosis transmembrane conductance regulator; diameter _max - maximum diameter of the vessel; Lum - lumacaftor; SAH - subarachnoid hemorrhage.

Figure 7: Correction of CFTR reduces neuronal damage in subarachnoid hemorrhage.

Shown are representative images of cortical cells stained for cleaved caspase-3 expression (top) and fluorinephritis (bottom) for cohorts including treatment regimens of (A) C18 (3 mg/kg daily for 2 days) and (B) lumacaftor ( Lum; 3 mg/kg daily for 2 days). Subarachnoid hemorrhage (SAH; n = 13 mice) increases the number of (C) caspase-3 and (D) FJ-positive cells compared with sham-operated controls (n = 12); both C18 (n = 6) and lumacaftor (n = 5) abolished the enhancement of caspase-3 and FJ staining. (E) SAH mice (n = 11) have a lower modified Garcia score (maximum score = 18; blinded assessments made 2 days after SAH) compared with sham-operated mice (n = 11); treatment with C18 (n = 10) restored neurological scores to the level of sham-operated animals. All data are presented as mean ± standard error of the mean. * denotes P < 0.05 for unpaired comparisons with sham-operated animals using the Kruskal-Wallis test and Dunnett's post hoc test. In panels C and D, the + sign indicates P < 0.05 compared with untreated SAH using the Kruskal-Wallis test and Dunnett's post hoc test. Abbreviations: Lum - lumacaftor; SAH - subarachnoid hemorrhage.

Figure 8: Placement of region of interest on FAIR-EPI images and analyses.

Shown are representative anatomical T2-weighted maps, T1-weighted flow-dependent inversion-recovery (FAIR) maps, and cerebral blood flow (CBF) FAIR maps for mice undergoing a surgical procedure for heart failure (top row) or subarachnoid hemorrhage (SAH; bottom row). ). Regions of interest were manually drawn on FAIR images of forebrain slices acquired at an inversion time (TI) of 825 ms using MIPAV software. As shown, the regions of interest span approximately 1 mm ³ of cortical volume within one hemisphere. Regions of interest were then copied directly onto CB maps obtained from FAIR images at individual inversion time points. SAH mice and sham-operated controls showed varying degrees of echo-planar imaging (EPI) distortion (best visualized on the FAIR image); EPI distortion was minimal in mice with heart failure.

Abbreviations: CB - cerebral blood flow; EPI - echo planar imaging; FAIR - flow-dependent inversion-recovery sequence; SAH - subarachnoid hemorrhage.

Figure 9: Mean arterial pressure in CFTR knockout mice

Mean arterial pressure is lower in cystic fibrosis transmembrane conductance regulator knockout (CFTR KO; CFTRtm1Unc; n = 6) mice compared with wild-type (WT; n = 6) littermates. * denotes P < 0.05 for unpaired comparison using t test.

Abbreviations: CFTR, cystic fibrosis transmembrane conductance regulator; KO - knockout; WT - wild type

Figure 10: Responses to phenylephrine in mouse cremaster arteries

Phenylephrine stimulates dose-dependent vasoconstriction in cremaster skeletal muscle arteries obtained from wild-type and cystic fibrosis transmembrane conductance regulator knockout (CFTR KO; CFTRtm1Unc) mice. The dose response of phenylephrine is not altered by (A) inhibition of CFTR in vitro (100 nmol/L CFTR(inhibitor)-172 for 30 minutes) or (B) deletion of the CFTR gene. Average maximum vessel diameters at 60 mm Hg. Art. (diameter _max ) are: wild type (panel A): 72 ± 3 µm, n = 5 from 4 mice; CFTR KO (panel B): 88 ± 4 μm, n = 6 from 4 mice; and wild-type littermates (panel B): 79 ± 4 μm, n = 5 from 2 mice. The dose-response curves in panel A are compared using paired two-way ANOVA (P=NS); curves in panel B are compared using unpaired two-way ANOVA (P=NS). For panel B, diameter _max KO and wild type values do not differ (t-test P=NS).

Abbreviations: CFTR, cystic fibrosis transmembrane conductance regulator; diameter _max - maximum diameter of the vessel; KO - knockout; NS - not significant.

Figure 11: Responses to phenylephrine in mouse posterior cerebral arteries following C18 treatment in vivo

(A) Phenylephrine stimulates dose-dependent vasoconstriction in posterior cerebral arteries obtained from heart failure (HF) mice and heart failure mice treated with C18 in vivo (3 mg/kg i.p. daily for 2 days). Statistical analysis reveals a significant difference between the two curves (i.e., significantly higher arterial tone from HF mice compared to HF + C18 mice). However (B), when the data are normalized to baseline tone ( _active tone - _resting tone, where _active tone represents the tone at a given phenylephrine concentration and _resting tone represents the tone immediately before stimulation), dose-response relationships are not significantly different. Average maximum vessel diameters at 45 mm Hg. (diameter _max ) are: CH: 137 ± 7 µm, n = 5 from 3 mice and CH + C18: 140 ± 8 µm, n = 5 from 4 mice (t-test P=NS). In both panels, * denotes P < 0.05 using two-way ANOVA.

Abbreviations: diameter _max - maximum diameter of the vessel; HF - heart failure; NS - not significant.

Figure 12: In vivo treatment with C18 does not alter myogenic tone or responses to phenylephrine in posterior cerebral arteries obtained from sham-operated mice.

In vivo C18 treatment (3 mg/kg i.p. daily for 2 days) did not alter (A) myogenic tone or (B) phenylephrine response in posterior cerebral arteries obtained from sham-operated mice. Average maximum vessel diameters at 45 mm Hg. (diameter _max ) in panel A: sham-operated: 146 ± 9 μm, n = 5 from 3 mice and sham-operated + C18: 138 ± 9 μm, n = 7 from 4 mice (t-test P=NS); and in panel B: sham-operated: 148 ± 7 μm, n = 6 from 3 mice and sham-operated + C18: 130 ± 10 μm, n = 5 from 3 mice (t-test P=NS). The curves in both panels are compared using two-way analysis of variance (P=NS).

Abbreviations: diameter _max - maximum diameter of the vessel; NS - not significant.

Figure 13: Phenylephrine responses in posterior cerebral arteries from CFTR knockout mice treated with C18

Phenylephrine causes dose-dependent vasoconstriction in posterior cerebral arteries obtained from cystic fibrosis transmembrane conductance regulator knockout (CFTR KO; CFTRtm1Unc) mice. Treatment with C18 in vivo (3 mg/kg i.p. daily for 2 days) did not alter the dose-response relationship of phenylephrine. Average maximum vessel diameters at 45 mm Hg. (diameter _max ) are: CFTR KO: 142 ± 5 μm, n = 6 from 3 mice and CFTR KO + C18: 145 ± 5 μm, n = 6 from 3 mice (t-test P=NS). Curves are compared using two-way analysis of variance (P=NS).

Abbreviations: CFTR, cystic fibrosis transmembrane conductance regulator; diameter _max - maximum diameter of the vessel; KO - knockout; NS - not significant.

Figure 14: Sholl analysis of apical and basal dendrites.

(A) Shown are Sholl analysis histograms plotting the number of dendritic crossings (i.e., dendritic branching) as a function of dendrite length (i.e., distance from the neuron soma) for apical dendrites. There were no differences in branching morphology between groups of sham-operated mice (n = 13 neurons from N = 4 mice), heart failure (HF; n = 11; N = 4) and HF + C18 (3 mg/kg i.p. daily for 2 weeks ; n = 11; N = 4); however, apical dendritic length is shorter in CH mice compared to sham-operated control and CH+C18 groups. Accordingly, (B) the average apical dendrite length (i.e., maximum radius) is significantly reduced in CH mice compared with sham-operated mice; this effect is normalized by treatment with C18. Similarly, (C) Sholl analysis histograms for basal dendrites show no differences in branching morphology in the groups of sham-operated animals (n = 12; N = 4), CH (n = 13; N = 4), and CH + C18 (n = 11; N = 4); however, the length of basal dendrites is shorter in CH mice compared with sham-operated mice and the CH+C18 group. Accordingly, (D) the average basal dendrite length (i.e., maximum radius) is significantly reduced in CH mice compared with sham-operated mice, an effect that is normalized by C18 treatment. * denotes P < 0.05 for unpaired comparisons with sham-operated animals using one-way ANOVA and Dunnett's post hoc test.

Abbreviations: HF - heart failure.

Figure 15: Analysis of spine density of apical dendrites

(A) In sham-operated (n = 8 neurons from N = 4 mice), heart failure (HF; n = 6; N = 3), and HF+C18 mice (3 mg/kg i.p. daily for 2 weeks; n = 10; N = 4), apical dendritic spine density (i.e., a measure of synaptic density) is relatively constant along the length of the dendrite, yielding slopes that are not statistically different from zero. (B) Although trends are present, the average density of apical dendritic spines measured between 60 and 100 μm from the soma does not reach statistical significance. Groups were statistically compared using one-way analysis of variance (P=N.S.).

Abbreviations: HF - heart failure; NS - not significant.

Figure 16: Responses to phenylephrine in mouse olfactory cerebral arteries after C18 treatment in vivo.

(A) Phenylephrine stimulates dose-dependent vasoconstriction in olfactory cerebral arteries obtained from mice with subarachnoid hemorrhage (SAH) and SAH mice treated with C18 in vivo (3 mg/kg i.p. for 2 days). Despite the clear tendency to separate, the two curves are not statistically different. (B) When data are normalized to baseline tone ( _active tone- _resting tone, where _active tone represents tone at a given phenylephrine concentration and _resting tone represents tone immediately before stimulation), the separation of dose-response relationships is minimal. Average maximum vessel diameters at 45 mm Hg. (diameter _max ) are: SAC: 109 ± 6 μm, n = 6 from 6 mice and SAC + C18: 105 ± 12 μm, n = 5 from 4 mice (t-test P=NS). Curves are compared using two-way analysis of variance.

Abbreviations: diameter _max - maximum diameter of the vessel; NS - not significant; SAH - subarachnoid hemorrhage.

Figure 17: C18 does not affect myogenic tone or responses to phenylephrine in sham-operated mice.

In vivo C18 treatment (3 mg/kg i.p. daily for 2 days) did not alter (A) myogenic tone or (B) phenylephrine dose response in olfactory cerebral arteries obtained from sham-operated mice. Average maximum vessel diameters at 45 mm Hg. (diameter _max ) are: sham-operated: 113 ± 3 µm, n = 5 from 3 mice; and sham-operated + C18: 110 ± 8 μm, n = 6 from 4 mice (t-test P=NS). Curves are compared using two-way analysis of variance.

Abbreviations: diameter _max - maximum diameter of the vessel; NS - not significant.

Figure 18: Responses to phenylephrine in olfactory cerebral arteries obtained from CFTR knockout mice

Phenylephrine provokes dose-dependent vasoconstriction in olfactory cerebral arteries obtained from cystic fibrosis transmembrane conductance regulator knockout (CFTR KO; CFTRtm1Unc) mice, which is similar to that of wild-type (WT) littermates. Treatment with C18 in vivo (3 mg/kg i.p. daily for 2 days) did not alter the dose-response relationships of phenylephrine. Average maximum vessel diameters at 45 mm Hg. (diameter _max ) are: WT: 98 ± 6 µm, n = 8 from 4 mice; CFTR KO: 110 ± 8 μm, n = 5 from 4 mice and CFTR KO + C18: 96 ± 6 μm, n = 6 from 3 mice (one-way ANOVA P=NS). Curves are compared using two-way analysis of variance.

Abbreviations: CFTR, cystic fibrosis transmembrane conductance regulator; diameter _max - maximum diameter of the vessel; KO - knockout; NS - not significant; WT - wild type.

Figure 19. Responses to phenylephrine in mouse olfactory cerebral arteries following lumacaftor treatment in vivo.

Treatment with lumacaftor (Lum) in vivo (3 mg/kg intraperitoneally daily for 2 days) did not alter phenylephrine dose-response relationships in olfactory cerebral arteries obtained from mice with subarachnoid hemorrhage (SAH). Average maximum vessel diameters at 45 mm Hg. (diameter _max ) are: SAC: 86 ± 2 µm, n = 6 from 3 mice; and SAH + Lum: 87 ± 4 μm, n = 7 from 4 mice (t-test P=NS). Curves are compared using two-way analysis of variance.

Abbreviations: diameter _max - maximum diameter of the vessel; Lum - lumacaftor; NS - not significant; SAH - subarachnoid hemorrhage.

The present invention is further illustrated by the following non-limiting example.

EXAMPLE

Methods and reagents

Reagents

Human cystic fibrosis transmembrane conductance regulator (CFTR) antibody was used to detect CFTR in baby hamster kidney fibroblast cells ("antibody 596"), CFTR therapeutic corrector "C18"(CF-106951; see WO 2007/021982; 1-( benzo[d][1,3]dioxol-5-yl)-N-(5-((2-chlorophenyl)(3-hydroxypyrrolidin-1-yl)methyl)thiazol-2-yl)cyclopropanecarboxamide) was purchased under the programs to distribute reagents and antibodies from the Cystic Fibrosis Foundation (http://www.cff.org/research). Dr John Riordan (University of North Carolina - Chapel Hill) provided “antibody 596 CFTR”, and Dr Robert Bridges (Rosalind Franklin University of Medicine and Science, Chicago, USA) provided compound C18 . Antibody 596 CFTR targets amino acids 1204 to 1211 (i.e., located in nucleotide-binding domain 2) (1). Fluorescein-labeled S1P (FITC-S1P) was purchased from Echelon Biosciences (via Cedarlane Laboratories; Burlington, Canada); lumacaftor was purchased from Selleck Chemicals (Cedarlane Laboratories); recombinant tumor necrosis factor was purchased from Sigma-Aldrich Canada (Oakville, Canada; catalog number T6674); and protease inhibitor cocktail tablets (Complete®; used in lysis buffers for Western blots) were purchased from Roche (Mississauga, Canada). All other chemicals were purchased from Sigma-Aldrich. MOPS balanced salt solution contained (mmol/L): NaCl 145, KCl 4.7, CaCl ₂ 3.0, MgSO ₄ ⋅7H ₂ O 1.17, NaH ₂ PO ₄ ⋅2H ₂ O 1.2, pyruvate 2, 0, ethylenediaminetetraacetic acid (EDTA) 0.02, 3-morpholinopropanesulfonic acid (MOPS) 3.0, and glucose 5.0.

CFTR mutant and knockout mice

Male mice homozygous for the ΔF508 CFTR mutation (CFTRtm1EUR; referred to as “ΔF508” herein) ( 2 ), a deletion of the CFTR gene (CFTRtm1Unc; referred to as CFTR −/− ), and complemented wild-type littermate controls were obtained from a stabilized colony in Hospital for Sick Children, Toronto (all CFTR-/-, ^CFTRΔF508 and wild-type littermates are mixed strains). The present inventors have discovered that CFTR-/- mice are highly susceptible to death under anesthesia. Although this mortality was not a problem for cerebral artery isolation experiments (e.g., Figs. 3C and 5C), it significantly interfered with cerebral blood flow and systemic hemodynamic measurements. Unlike CFTR −/− mice, CFTR ^ΔF508 mutant mice were not as prone to death during anesthesia. Because the CFTR ^ΔF508 mutation significantly reduces delivery of CFTR to the plasma membrane ( 3 ), CFTR activity is very low in the CFTR ^ΔF508 mouse model ( 2 , 4 , 5 ). Thus, the CFTR ^ΔF508 model provided a suitable alternative to CFTR −/− mice for cerebral blood flow and systemic hemodynamic measurements. The present inventors have not investigated the underlying cause of death in CFTR-/-. Both CFTR−/− and ^CFTRΔF508 variants are well known to respond adversely to stress. For example, housing and movement conditions are two prominent stressors that can significantly increase mortality (6). Interestingly, ^CFTRΔF508 mice exhibit a less severe phenotype than CFTR−/− mice (5, 6): this is thought to be due to the small amount of residual CFTR activity that is present in ^CFTRΔF508 animals (2). In this context, we speculate that CFTR−/− mice may have been more sensitive to external environmental stressors in the experimental animal facility (e.g., housing, noise, and grooming), which ultimately resulted in increased mortality under anesthesia . As an experimental model, ^CFTRΔF508 mice have several abnormalities associated with loss of CFTR function. Intestinal complications are the most significant pathological effect of the CFTR mutation and are also a leading cause of postpartum mortality (6, 7). Common gastrointestinal problems include retention of thick mucus, impaired motility, and a strong tendency toward intestinal obstruction. CFTR ^ΔF508 mice are 40–50% smaller in weight/size compared to wild-type counterparts, although they develop well into adulthood ( 6 , 7 ). Several other differences from wild-type mice were observed; however, they appear to be relatively small (6, 7). For example, altered chloride currents and/or sodium transport in certain tissues (eg, kidney, gallbladder, nasal epithelium) have been noted, but the pathological significance of these differences is unclear since the tissues do not appear histologically defective (6, 7). It should be noted that CFTR ^ΔF508 mice (and CFTR mutant mice in general) have a rather mild cystic fibrosis phenotype compared to human cystic fibrosis. The lower respiratory tract of CFTR ^ΔF508 mice is essentially normal: there are no lower respiratory tract epithelial abnormalities, and pulmonary inflammation does not develop spontaneously without infection ( 6 , 7 ). The pancreas, gallbladder, liver, bile duct, male reproductive tract, lacrimal gland, and submandibular glands do not have obvious histopathology (5–7). The lack of a severe cystic fibrosis phenotype is explained in part by the expression of a CFTR-free calcium-activated chloride channel (CACC) in certain mouse tissues, which compensates for the loss of CFTR ( 7 , 8 ).

Global hemodynamic parameters

As described previously (9–11), echocardiographic measurements were obtained using a 30 MHz mechanical sector transducer (Vevo 770; Visual Sonics, Toronto, Canada) combined with mean arterial pressure (MAP) measurements (mouse microtip manometric catheter Millar SPR-671; Inter V Medical Inc., Montreal, Canada). Aortic velocity integral (VTI) was measured using pulsed wave Doppler just above the aortic root. Aortic cross-sectional area (CSA) was calculated from aortic root size (ARD): CSA = π(ARD/2)2. Stroke volume (SV), cardiac volume velocity (CO), and total peripheral vascular resistance (TPR) were calculated using the formula: SV = CSA × VTI; CO = SV × HR; TPR = MAP/CO.

Measuring brain perfusion using MRI

As previously described (10-12), the present inventors used magnetic resonance imaging (MRI) flow-dependent inversion recovery (FAIR) (13) to assess cerebral blood flow (CBF). During imaging, mice were immobilized with 1.8% isoflurane delivered via a nasal cone. To maintain body temperature, the MRI has built-in tubes that drain water from an external pump heater. Respiration was controlled using a pneumatic bag (SA Instruments, Stony Brook, USA); Isoflurane levels were adjusted to maintain breathing at approximately 50 breaths per minute.

FAIR images were acquired using a 7 Tesla micro-MRI system (BioSpec 70/30 USR, Bruker BioSpin, Ettlingen, Germany), including a B-GA12 gradient insert, a 72 mm ID linear cavity resonator for RF transmission, and a front-mounted head coil for receiving radio frequencies. FAIR highlights perfusion as an accelerated relaxation of the T1 signal after slice-selective versus non-selective inversion preparation according to the following equation: CC = l (1/T1, ss - 1/T1, ns) (ml/(100 g*min), where " ss" and "ns" denote slice-selective and non-selective measurements, and l is the blood-brain partition coefficient, defined as the ratio between the water concentration per gram of brain tissue and per ml of blood. This coefficient is approximately 90 ml/100 g y mice (14). The FAIR optimization used in the present invention was a single-pulse echo-planar imaging (EPI) method with adiabatic inversion preceding. Parameters included an echo time of 12.5 ms, a repetition time of 17 s, and 18 inversion times from. 25 to 6825 ms in 400 ms increments, 3 mm slice-selective inversion plate, 18 x 18 mm scan area with 72 x 72 matrix for 250 µm in-plane resolution, 1 mm slice thickness and 10 min 12 s acquisition time. Data acquisition was repeated in vertical sections of the forebrain, midbrain, and hindbrain, respectively, in the anterior, mixed, and posterior zones of the blood supply. FAIR images were assessed in words (MIPAV, NIH, Bethesda, MD; http://mipav.cit.nih.gov) subhemispheric regions of interest (ROIs) called “global” and local ROIs corresponding to the cortical and subcortical parenchyma in the anterior brain; cortical and paraventricular parenchyma in the middle sections; and cortical and paraventricular parenchyma of the midbrain in the hindbrain. Regions of interest are drawn directly onto T1-weighted signal images to allow manual motion correction within the scan. ROIs were recorded using parametric CBF maps to verify the absence of displacement from highly perfused vessels and meninges. T1 regressions and CC calculations were performed using Matlab (Mathworks, Natick, MA). All reported CBF values fall within the published range of CBF measurements in mice for isoflurane levels according to the present invention (15). EPI is particularly susceptible to magnetic susceptibility distortion in the phase encoding direction (16), which corresponds to the vertical direction in this FAIR protocol. As can be seen in FIG. 8, correction of magnetic field inhomogeneity does not sufficiently limit EPI distortion in a surgical model of subarachnoid hemorrhage (SAH) due to air-tissue boundaries immediately adjacent to the brain at postoperative imaging moments. In some cases, the distortion was so severe that the regions of interest had to be placed more laterally in the cortex (Fig. 8). However, since the distortion is the same in all T1-weighted images, it does not affect the T1 measurement provided that the distorted region does not overlap with another region. The relatively consistent ns-T1 values in each cohort (i.e., 7% standard deviation between cohorts, no treatment group differences within cohorts) provide supporting evidence that confounding bias had little impact on CBF measurements.

Cell culture

Procedures for isolating and culturing mesenteric artery smooth muscle cells were performed as described elsewhere (10). Briefly, mesenteric artery segments were isolated, cut into small pieces, and exposed to trypsin, collagenase, and elastase. The resulting cell suspension was washed several times with phosphate-buffered saline and placed in Dulbecco's modified Eagle's medium (DMEM) containing 10% fetal bovine serum and 1% penicillin and streptomycin. Cell cultures were maintained at 37°C and 5% CO ₂ and separated at a seeding density of 10 ⁶ cells. Baby hamster kidney fibroblast cells stably expressing wild-type human CFTR ( 10 , 17 ) were maintained in DMEM/F12 containing 5% fetal bovine serum and 250 μmol/L methotrexate (which activates the CFTR transgene promoter). Cells were maintained under standard cell culture conditions at 37°C and 5% CO ₂ .

FACS-based FITC-S1P absorbance measurement

As described previously ( 10 ), cell monolayers (treated or untreated) were incubated with 1 μmol/L S1P-FITC for 60 min; cells were then separated by trypsinization, washed twice with ice-cold phosphate-buffered saline, filtered through a 35-μm cell filter, and analyzed using Becton-Dickinson FACS Canto using FACS DIVA software version 6.1. Cell monolayers treated with unlabeled S1P served as background controls. During the analysis procedure, the average fluorescence intensity (arbitrary units) of each cell population, which is a measure of absorption, was determined.

Assessing iodide efflux

Iodide efflux measurements were performed as described previously (18). Briefly, monolayers of confluent cells were loaded with iodide by incubating them in HEPES-based loading buffer (136 mmol/L NaI, 3 mmol/L _KNO3 , 2 mmol/L Ca( _NO3 ) ₂ , 11 mmol/L glucose and 20 mmol/L l HEPES; pH 7.4) for 1 hour (under standard cell culture conditions at 37°C in 5% CO ₂ ). After loading with iodide, cells were washed 4 times with iodide-free efflux buffer (where NaNO ₃ was replaced by NaI). Iodide levels in the supernatant were quantified using an iodide-selective electrode (Lazar Research Laboratories; Los Angeles, USA) calibrated with NaI standards. Prestimulation iodide levels were determined, and cells were then stimulated with a mixture containing 10 μmol/L forskolin, 1 mmol/L isobutylmethylxanthine, and 100 μmol/L cpt-cAMP (in 1% v/v DMSO). The stimulating mixture strongly activates protein kinase A and therefore maximally phosphorylates/activates CFTR. Iodide levels in the supernatant were determined at seven consecutive 1-min intervals after stimulation. Consistent with previous results (18), outflow was evident within the first minute and was reliably maximal between 60 and 120 seconds. Thus, the maximum outflow rate was calculated using the measured outflow rate between 60 and 120 seconds after stimulation (18).

Western blotting

Western blotting for CFTR was performed as previously described ( 12 , 18 ). Cerebral artery lysates were prepared by grinding arterial samples in lysis buffer containing 50 mM Tris (pH 7.3), 150 mM NaCl, 2 mM EDTA, 0.1% Triton-X-100, 0.1% SDS, and protease inhibitors; the same lysis buffer was used to prepare cell culture lysates. After lysis, samples were centrifuged (10 minutes at 13500 g; 4°C) to remove insoluble material. Immediately before polyacrylamide electrophoresis, additional SDS (to a final concentration of 2%), glycerol (to a final concentration of 2%), β-mercaptoethanol (to a final concentration of 2%), and dithiothreitol (to a final concentration of 2 mM) were added. Proteins were separated electrophoretically on 7% acrylamide gels and transferred to polyvinylidene difluoride (PVDF) membranes. The membranes were blocked for 30 to 40 minutes in 5% skim milk (phosphate-buffered saline containing 1% Tween 20 (PSBT); 137 mM NaCl, 2.7 mM KCl, 10 mM Na ₂ HPO ₄ , 1.76 mM K ₂ HPO ₄ ; pH 7.4). Primary antibody treatments included: (i) rabbit polyclonal anti-CFTR antibody (1:1000 in 5% milk/PBST; Cell Signaling Technology from New England Biolabs Canada; Whitby, Canada; catalog no. 2269); lysates of cerebral arteries and vascular smooth muscle cells.

(ii) Mouse monoclonal antibodies to human CFTR (1:20,000 dilution in 5% milk/PBST; “antibody 596”); lysates of fibroblast cells from the kidneys of baby hamsters.

(iii) Mouse monoclonal antibody to α-tubulin (1:5000 in 5% milk/PBST; clone DM1A; Cell Signaling Technology from New England Biolabs Canada; catalog number 3873).

Primary antibodies were conjugated to either peroxidase-labeled secondary donkey anti-rabbit IgG (GE Healthcare Amersham (Piscataway, USA), catalog no. NA934) or secondary peroxidase-labeled goat anti-mouse IgG (GE Healthcare Amersham, catalog no. NA931). A standard chemiluminescence method (Westar ETA C; VPQ Scientific, Toronto, Canada) (ChemiDoc; Bio-Rad Laboratories; Mississauga, Canada) was used to expose X-ray film or collect digital images. Developed films were assessed by densitometry using Image J software (freely available from the NIH); digital images were assessed using Image Lab software (Bio-Rad).

RNA isolation and reverse transcription

Resistance artery RNA was isolated using Norgen Biotek (Thorold, Canada) "Total RNA Purification Micro" spin columns using proteinase K digestion and DNA removal methods as specified in the manufacturer's instructions. Eluted RNA was quantified using an Agilent Technologies RNA 6000 Pico kit and bioanalyzer; analysis confirmed the receipt of high-quality RNA from arterial tissue samples (RNA integrity [RIN]: brain = 8.4 ± 0.1, n = 5; cremaster = 8.2 ± 0.4, n = 6). RNA isolation from baby hamster kidney fibroblast cells was performed using the same method, minus the proteinase K digestion step. In all cases, cDNA was prepared from RNA using a Superscript III reverse transcription kit (Invitrogen Life Technologies; Burlington, Canada) according to the manufacturer's instructions. Residual RNA was removed by incubating the resulting cDNA with RNase H (0.125 U/μl; New England Biolabs Canada; Whitby, Canada).

Quantitative PCR

Quantitative PCR was performed using an Applied Biosystems ViiA™ 7 Real-Time PCR System and Power SYBR® Green PCR Master Mix (both from Invitrogen Life Technologies). Each primer set was carefully tested to ensure specificity and comparable performance. Target genes were assessed in triplicate using 1 ng of cDNA obtained by reverse transcription; Water was used as negative controls. PCR amplification consisted of 10 min of denaturation at 95°C, followed by 40 cycles of amplification (15 s at 95°C and 60 s at 60°C). After amplification, the amplicons were melted: the resulting dissociation curve confirmed the formation of a single product. Transcript expression levels in mouse tissues were calculated from DCt values relative to the standard housekeeping gene hydroxymethylbilane synthase (HMBS). To confirm the reliability of HMBS for normalization, transcript expression levels were also calculated from DCt values relative to the glucose-6-phosphate dehydrogenase (G6PD) gene, obtaining similar results. In experiments with baby hamster kidney fibroblast cells, primers specific for human CFTR and hamster GAPDH were used, the latter being a housekeeping gene suitable for normalization.

Fixation of the brain and preparation of sections for immunohistochemistry and fluorine nephritis staining

The development of delayed blood vessel constriction and brain damage occurs more rapidly in the experimental mouse model of SAH (2 to 5 days post-SAH) (12, 19-21) compared to what is observed clinically (4 to 12 days post-SAH). (22, 23). We chose the time point of 2 days after SAH induction to assess damage markers based on previous work demonstrating significant neuronal damage and behavioral deficits at this time point ( 12 ). 2 days after SAH induction, animals were anesthetized with isoflurane; their brains were perfused with phosphate-buffered saline (PBS) and then perfusion-fixed with paraformaldehyde in phosphate buffer (4%; pH 7.4), both solutions injected through the ascending aorta. The brain was immediately dissected and cut into coronal sections 1 mm posterior to the bregma. These brain sections were then fixed in 4% paraformaldehyde (pH 7.4) for 48 hours at 4°C and then incubated in 10% sucrose (3 hours) and then 30% sucrose overnight (at 4°C ) for cytoprotection. The brain sections were then placed in OCT reagent (Sakura Finetek USA; Torrance, USA) on isopentane and dry ice. Cryostat sections (5-μm-thick coronal sections) were collected on Tissue Path Superfrost Plus Gold slides (Fisher Scientific; Whitby, Canada) and stored at −80°C until use.

Fluorescent immunohistochemistry

All samples were pretreated for 10 minutes with proteinase (20 μg/ml; Promega; Madison, USA) and then blocked for 60 minutes in a standard manner with 10% normal donkey serum (Jackson ImmunoResearch Laboratories; West Grove, USA) in PBS or 10% goat serum. serum (Invitrogen Life Technologies; Burlington, Canada) in PBS containing 1% bovine serum albumin. Brain sections were incubated with monoclonal rabbit anti-human active cleaved caspase-3 antibody (clone C92-605; 1:1000 dilution; BD Biosciences Canada; Mississauga, Canada) and conjugated to goat anti-rabbit IgG antibody Alexa Fluor 488 (Invitrogen). Both types of antibodies were diluted in Can Get Signal® immune response enhancer solution and an incubation time of 1 hour at room temperature was used. Samples were washed and mounted using CC MountÔ.

Fluorine jade staining

Brain sections were sequentially incubated with 1% NaOH/80% ethanol (5 minutes), 70% ethanol (2 minutes), distilled water (2 minutes), and 0.06% potassium permanganate (10 minutes). After rinsing with deionized water, brain sections were stained with 0.0004% Fluoride-Jade B (Histo-Chem Inc., Jefferson, USA) in 0.1% acetic acid (15 min). The samples were then rinsed with deionized water, dried, and cleaned by immersion in xylene for 1 minute. Slides were mounted using DPX section mounting medium (Sigma).

Digital imaging and assessment of brain damage by cell counting

Digital immunofluorescence images were obtained using a Zeiss LSM 710 confocal laser scanning microscope coupled with Zeiss ZEN 2010 software (vascular imaging). Overlays were created using freely available ImageJ 1.44p software (National Institutes of Health, USA). Caspase-3 and FJ-positive cells were scored by two independent reviewers under blinded conditions as previously described ( 12 ). For each animal, counts were performed for a cortical region on a single coronal section of the brain 1 mm posterior to bregma (magnification 200×), which included the left and right temporal and parietal lobes. The number of positive cells counted by each examiner was averaged to obtain the average number of positive cells per animal, which was then used for statistical analysis.

Histological analysis of dendritic morphology:

Brains were perfused with heparinized saline via transcardial perfusion, isolated, and processed/stained using a commercial, commercially available Rapid GolgiStain kit (FD NeuroTechnologies Inc.; Columbia, USA). The kit uses a modified Golgi-Cox staining method originally described by Glaeser and van der Loos (24). Briefly, isolated brain tissue samples were immersed in the kit's solution containing mercuric chloride, potassium dichromate, and potassium chromate in the dark for 8 days; then incubated for 6 days with a tissue protection solution. Brain samples were then washed, mounted in 4% low-melting agarose, sectioned in the coronal plane (150 μm thick; Campden Instruments 7000smz-2 vibrating microtome), and mounted on gelatin-coated glass slides. The staining procedure was then completed using the development/staining reagents provided in the kit according to the manufacturer's instructions. Neurons and dendritic segments were imaged using NIS Elements AR stereology software on a Nikon Eclipse Ti2 microscope with motorized X, Y, and Z focus for high-resolution imaging (Nikon Instruments Europe; Amsterdam, the Netherlands). We analyzed both basal and apical dendritic networks of pyramidal cortical neurons in the frontal cortex using a quantitative approach originally described by Scholl ( 25 ). Image J software was used in combination with the semi-automated plugin Simple Neurite Tracer (both freely available from the NIH) to identify and digitally extract the dendritic networks of a single cortical neuron from a hyperprobe image at 200× magnification. The center of the soma was marked with an arrow; dendritic morphology was then characterized by Sholl analysis (see https://imagej.net/Sholl_Analysis, version 3.7.4) using 5-μm intervals up to a maximum radius of 300 μm. This analysis characterizes dendritic morphology in terms of dendritic intersections (i.e., branching) and dendrite length. For each treatment group, 2 to 4 neurons per mouse from 4 mice were analyzed in a blinded manner. To assess dendritic spine density (i.e., the number of small projections found on dendrites), 3rd order arbor dendritic segments were imaged at 100× magnification. Spine density was measured as the number of spines per segment for the following segments: 20 to 50 μm from the soma; from 30 to 60 µm from the soma; from 50 to 80 µm from the soma; and from 60 to 100 µm from the soma). This “sliding measurement” approach allows us to assess how spine density varies along the length of a dendritic arbor. For each treatment group, spine density was measured in 2 to 4 pyramidal cortical neurons (2 to 4 third-order branches per neuron) per mouse in 3 to 4 mice per group under blinded conditions.

As described above, the C18 CFTR corrector and anti-human CFTR antibody (designated “596”) were purchased through the Cystic Fibrosis Foundation reagent and antibody distribution programs. All other reagents used are commercially available as described above.

Animals: The Institutional Animal Care and Use Committees at the University of Toronto and the University Health Network (UHN) approved all animal care and experimental protocols. Commercially available male wild-type mice (2 to 3 months old; C57BL/6N) were purchased from Charles River Laboratories (Montreal, Canada). Male mice homozygous for the ΔF508 CFTR mutation (CFTR ^tm1EUR ; referred to as “ΔF508” herein) ( 11 ), a deletion of the CFTR gene (CFTR ^tm1Unc ; designated CFTR ^−/− ), and complemented wild-type littermate controls were obtained from a stabilized colony in Hospital for Sick Children, Toronto (all CFTR ^-/- , CFTR ^ΔF508 and littermates are mixed strains). All mice were housed on a standard 14-h:10-h light-dark cycle, fed regular chow, and provided with ad libitum access to water.

Myocardial infarction: Heart failure was induced by surgical ligation of the left anterior descending coronary artery (3). Briefly, mice were anesthetized with isoflurane, intubated with a 20-gauge angiocatheter, and ventilated with room air. Under sterile conditions, the chest and pericardium were opened, and the left anterior descending coronary artery was permanently ligated with 7-0 silk suture (Deknatel; Fall River, USA). In the control group, the chest and pericardium were opened in sham-operated animals, but the left anterior descending coronary artery was not ligated. After the procedure, the chest was closed and the mice were extubated while breathing spontaneously. Posterior cerebral arteries (PCAs) were isolated 4 to 6 weeks after infarction.

Induction of subarachnoid hemorrhage: A well-studied model of experimental SAH was used (6). Briefly, each mouse was anesthetized (isoflurane) and head fixed in a stereotaxic frame; A 7 mm incision was made along the midline of the anterior scalp and a 0.9 mm hole was drilled into the skull 4.5 mm anterior to the bregma. A spinal needle was inserted into the chiasmatic cistern: 80 μl of arterial blood was injected into the intracranial space over 10 seconds. The administered blood was obtained from an individual wild-type donor mouse (via cardiac puncture) immediately before injection and did not contain anticoagulants. After injection, the scalp incision was sutured. Buprenorphine (0.05 mg/kg; volume 0.5 to 1.0 mL) was administered twice daily (immediately after the SAH surgical procedure). Sham-operated animals underwent an identical surgical procedure using sterile saline instead of blood. Olfactory cerebral arteries were isolated 2 days after SAH induction.

Isolation and functional assessment of resistance arteries: The mouse olfactory arteries (first branch of the anterior cerebral artery) and PCA were carefully dissected, cannulated into micropipettes, stretched to their in vivo length, and pressurized to 45 mmHg as described previously (3, 6) ); The resistance arteries of mouse skeletal muscles were separated from the cremaster muscle, cannulated, and pressure was applied to 60 mmHg. All functional experiments were performed in 3-morpholinopropanesulfonic acid (MOPS) saline at 37°C without perfusion. Vasomotor responses to phenylephrine (5 μmol/L for posterior cerebral arteries, 10 μmol/L for cremaster skeletal muscle arteries) provided an assessment of vascular viability at the beginning and end of each experiment. Arteries that did not show a constriction response to phenylephrine greater than or equal to 25% were excluded.

Myogenic reactions were induced by a stepwise increase in transmural pressure by 20 mm Hg. from 20 mm Hg up to 80 mm Hg (olfactory arteries) or 100 mm Hg. (posterior cerebral arteries and resistive arteries of the skeletal muscles of the cremaster). At each pressure stage, the diameter of the vessel ( _active diameter) was measured after reaching a steady state (5 min). Vessels requiring treatment (eg, CFTR( _inhibitor )-172) were incubated with the reagent in MOPS for 30 minutes, and then myogenic responses in the presence of this reagent were assessed. After all _active diameter measurements were completed, the MOPS buffer was replaced with a Ca ²⁺ -free version, and the maximum passive diameter (diameter _max ) was recorded at each pressure step.

Myogenic tone was calculated as the percentage of constriction relative to the maximum diameter at each corresponding transmural pressure: tone (%diameter _max ) = [(diameter _max - diameter _active )/diameter _max ] × 100, where diameter _active is the diameter of the vessel in MOPS, containing Ca ²⁺ , and diameter _max is the diameter in MOPS without Ca ²⁺ . When analyzing vasomotor responses to phenylephrine, the same equation was used, but in this case, _active diameter represents the diameter of the vessel at steady state after administration of the agent, and _max diameter represents the maximum diameter measured in the absence of Ca ²⁺ .

Magnetic resonance imaging-based cerebral blood flow measurements: The non-invasive magnetic resonance imaging (MRI) method (FAIR method) was used to assess cerebral perfusion as previously described (6). Briefly, the FAIR method highlights perfusion as an accelerated relaxation of the T1 signal. MRI signals (Bruker Corporation Biospec 70/30 USR; Ettlingen, Germany) were obtained from vertical sections of the forebrain, midbrain, and hindbrain, which correspond to the anterior, mixed, and posterior circulation. FAIR images were assessed for designated regions of interest (location of regions shown in Fig. 8) using standardized image processing algorithms and techniques (MIPAV; National Institutes of Health, Bethesda, USA; http://mipav.cit.nih.gov/).

Magnetic resonance imaging-based edema measurement: Edema was assessed using quantitative _T2 mapping (12) using a 7 Tesla micro-MRI system (Bruker Corporation Biospec 70/30 USR; Ettlingen, Germany). T ₂ mapping acquisition produced quantitative T ₂ maps in 9 contiguous 1-mm-thick two-dimensional axial slices covering the volume from the forebrain to the hindbrain. T ₂ maps were generated from T ₂ -weighted images with echo times ranging from 12 to 384 ms using built-in Bruker software through linear regression of log-transformed signal magnitudes and echo times for each voxel. _T2 values were extracted using MIPAV software using manually drawn regions of interest placed in the forebrain, midbrain, and hindbrain _T2 maps.

Assessment of neurological function: Neurological function was assessed using the modified Garcia score as previously described (6). The neurological assessment consists of 6 domains: spontaneous activity, spontaneous movement of all 4 limbs, forelimb extension, climbing ability, body proprioception and response to whisker touch. Two observers performed a blinded neurological examination 2 days after SAH. The maximum score is 18, indicating normal neurological function.

Statistics: All data are presented as means ± standard error of the mean, where n represents the number of independent measurements (i.e., ratings of specimens, vessels, or test subjects). For statistical comparisons, the nonparametric Mann-Whitney test was used, with an exact p-value calculated to compare two independent groups. To compare several independent groups, nonparametric one-way analysis of variance (Kruskal-Wallis test) was used, followed by Dunnett's post hoc test. To evaluate myogenic responses and dose-response relationships, data were analyzed using two-way ANOVA followed by post hoc analysis with Tukey's test. Differences were considered significant at P < 0.05. All data presented in Fig. 6, were collected under blind conditions; all other data presented herein did not require blinding and were not collected under blinded conditions.

The following references are provided in the Methods and Reagents section above:

1. Cui L, Aleksandrov L, Chang XB, et al. Domain interdependence in the biosynthetic assembly of CFTR. J Mol Biol 2007; 365:981-94.

2. van Doorninck JH, French PJ, Verbeek E, et al. A mouse model for the cystic fibrosis delta F508 mutation. EMBO J 1995; 14:4403-11.

3. Van Goor F, Straley KS, Cao D, et al. Rescue of DeltaF508-CFTR trafficking and gating in human cystic fibrosis airway primary cultures by small molecules. Am J Physiol Lung Cell Mol Physiol 2006; 290:L1117-30.

4. French PJ, van Doorninck JH, Peters RH, et al. A delta F508 mutation in mouse cystic fibrosis transmembrane conductance regulator results in a temperature-sensitive processing defect in vivo. J Clin Invest 1996; 98:1304-12.

5. Zeiher BG, Eichwald E, Zabner J, et al. A mouse model for the delta F508 allele of cystic fibrosis. J Clin Invest 1995; 96:2051-64.

6. Wilke M, Buijs-Offerman RM, Aarbiou J, et al. Mouse models of cystic fibrosis: phenotypic analysis and research applications. J Cyst Fibros 2011; 10 Suppl 2:S152-71.

7. Lavelle GM, White MM, Browne N, McElvaney NG, Reeves EP. Animal Models of Cystic Fibrosis Pathology: Phenotypic Parallels and Divergences. Biomed Res Int 2016; 2016:5258727.

8. Nilius B, Droogmans G. Amazing chloride channels: an overview. Acta Physiol Scand 2003; 177:119-47.

9. Hoefer J, Azam MA, Kroetsch JT, et al. Sphingosine-1-phosphate-dependent activation of p38 MAPK maintains elevated peripheral resistance in heart failure through increased myogenic vasoconstriction. Circ Res 2010; 107:923-33.

10. Meissner A, Yang J, Kroetsch JT, et al. Tumor Necrosis Factor-α-Mediated Downregulation of the Cystic Fibrosis Transmembrane Conductance Regulator Drives Pathological Sphingosine-1-hosphate Signaling in a Mouse Model of Heart Failure. Circulation 2012; 125:2739 50.

11. Yang J, Hossein Noyan-Ashraf M, Meissner A, et al. Proximal Cerebral Arteries Develop Myogenic Responsiveness in Heart Failure via Tumor Necrosis Factor-α-Dependent Activation of Sphingosine-1-Phosphate Signaling. Circulation 2012; 126:196-206.

12. Yagi K, Lidington D, Wan H, et al. Therapeutically Targeting Tumor Necrosis Factor-α/Sphingosine-1-Phosphate Signaling Corrects Myogenic Reactivity in Subarachnoid Hemorrhage. Stroke 2015; 46:2260-70.

13. Kim S.G. Quantification of relative cerebral blood flow change by flow-sensitive alternating inversion recovery (FAIR) technique: application to functional mapping. Magn Reson Med 1995; 34:293-301.

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15. Foley LM, Hitchens TK, Kochanek PM, Melick JA, Jackson EK, Ho C. Murine orthostatic response during prolonged vertical studies: effect on cerebral blood flow measured by arterial spinlabeled MRI. Magn Reson Med 2005; 54:798-806.

16. Foltz WD, Porter DA, Simeonov A, et al. Readout-segmented echo-planar diffusion-weighted imaging improves geometric performance for image-guided radiation therapy of pelvic tumors. Radiother Oncol 2015; 117:525-31.

17. Sharma M, Benharouga M, Hu W, Lukacs GL. Conformational and temperature-sensitive stability defects of the delta F508 cystic fibrosis transmembrane conductance regulator in post-endoplasmic reticulum compartments. J Biol Chem 2001; 276:8942-50.

18. Malik FA, Meissner A, Semenkov I, et al. Sphingosine-1-Phosphate Is a Novel Regulator of Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) Activity. PLoS One 2015; 10:e0130313.

19. Lin CL, Calisaneller T, Ukita N, Dumont AS, Kassell NF, Lee KS. A murine model of subarachnoid hemorrhage-induced cerebral vasospasm. J Neurosci Methods 2003; 123:89-97.

20. Parra A, McGirt MJ, Sheng H, Laskowitz DT, Pearlstein RD, Warner DS. Mouse model of subarachnoid hemorrhage associated cerebral vasospasm: methodological analysis. Neurol Res 2002; 24:510-6.

21. Aum DJ, Vellimana AK, Singh I, et al. A novel fluorescent imaging technique for assessment of cerebral vasospasm after experimental subarachnoid hemorrhage. Sci Rep 2017; 7:9126.

22. Brilstra EH, Rinkel GJ, Algra A, van Gijn J. Rebleeding, secondary ischemia, and timing of operation in patients with subarachnoid hemorrhage. Neurology 2000; 55:1656-60.

23. Macdonald RL. Delayed neurological deterioration after subarachnoid haemorrhage. Nat Rev Neurol 2014; 10:44-58.

24. Glaser EM, Van der Loos H. Analysis of thick brain sections by obverse-reverse computer microscopy: application of a new, high clarity Golgi-Nissl stain. J Neurosci Methods 1981; 4:117-25.

25. Sholl DA. Dendritic organization in the neurons of the visual and motor cortices of the cat. J Anat 1953; 87:387-406.

RESULTS

Insufficient CFTR function leads to decreased cerebral perfusion

Previous work has demonstrated that CFTR significantly regulates cerebral artery myogenic tone and has identified a strong correlation between cerebral artery CFTR protein expression and cerebral perfusion ( 4 , 6 ). Thus, the mechanistic concept suggests that correction of insufficient cerebrovascular CFTR activity (i.e., reduction of S1P transport due to decreased cell surface expression of CFTR) is a viable therapeutic strategy to normalize the impaired cerebral myogenic response and hence cerebral blood flow (4).

CFTR mutant mice were used to establish the causal relationship between cerebral artery CFTR activity and cerebral perfusion. Because CFTR ^−/− mice were extremely sensitive to experimental stressors, this did not allow for accurate and reproducible echocardiographic/CBF measurements. Therefore, we used the more stable CFTR ^ΔF508 mice, which have minimal cell surface CFTR expression and activity (11). This supported the inventors' strategic decision to use CFTR ^ΔF508 mice to correlate cerebral artery myogenic tone with hemodynamic parameters in a loss-of-function phenotype.

Vasomotor reactivity in the posterior cerebral arteries (PCAs) was assessed to directly compare the ^CFTRΔF508 phenotype with a previously published characterization of PCA responses from CFTR knockout mice ( 4 ). As expected, the PCA of ^CFTRΔF508 mice exhibited increased myogenic tone compared with wild-type littermates ( Fig. 8A ); the magnitude of the increase is qualitatively similar to that found for CFTR ^−/− mice ( 4 ). Responses to phenylephrine show an upward shift due to increased myogenic tone (Fig. 8B); however, _EC50 values did not differ (log _EC50 WT: -5.83 ± 0.21, n = 5 from 3 mice; log _EC50 CFTR ^ΔF508 : -6.00 ± 0.04, n = 6 from 4 mice; Mann-Whitney test P=NS) and the difference between the curves is eliminated by adjusting for differences in basal tone (ie, myogenic tone at 45 mmHg; Fig. 8C). Thus, the ΔF508 mutation increases myogenic sensitivity but does not affect overall contractility. CBF is significantly lower in ^CFTRΔF508 mice ( Fig. 8D ); however, neither CO (Fig. 8E) nor TPR (Fig. 8F) were mutated. Based on these observations, it is clear that the cerebral perfusion deficiency is of vascular origin.

The CFTR ^ΔF508 mutation does not alter TPR, indicating that not all vascular beds are subject to CFTR-dependent regulation. Since TPR is primarily generated and regulated by skeletal muscle resistance arteries (14), it was analyzed whether CFTR influences myogenic tone in mouse skeletal muscle resistance arteries. CFTR mRNA expression is approximately 10-fold lower in skeletal muscle resistance arteries of wild-type mice compared with cerebral arteries ( Fig. 1G ). Inhibition of CFTR activity in vitro (100 nmol/L CFTR(inhibitor)-172, 30 minutes) did not affect the myogenic tone of the wild-type cremaster artery (Fig. 1H), which contradicts recent observations that CFTR(inhibitor)-172 enhances myogenic tone. tone of the olfactory cerebral artery (6). In contrast to PMA ( 4 ), deletion of the CFTR gene did not increase myogenic tone in the skeletal muscle cremaster resistance arteries ( Fig. 1I ); in fact, myogenic tone is slightly weakened and corresponds to a lower mean arterial pressure (MAP) (Fig. 9). The dose response of phenylephrine was not altered by either inhibition of CFTR in vitro or deletion of the CFTR gene (Fig. 10).

In summary, the present experiments with ^CFTRΔF508 establish: (i) a causal relationship between CFTR expression, PCA myogenic reactivity, and CD; (ii) CFTR modulates myogenic tone in specific vascular beds; and (iii) CFTR does not modulate the tone of skeletal muscle resistance arteries, which is noteworthy because these arteries make a significant contribution to TPR. Because HF and SAH suppress CFTR expression in cerebral arteries concurrently with microvascular dysfunction (4, 6), these data provide a strong mechanistic basis for the use of CFTR-modulating drugs to specifically correct cerebrovascular dysfunction and CBF deficiency in this disease.

C18 increases CFTR activity after TNF-dependent downregulation

Current CFTR-modulating drugs identified through high-throughput drug screening are being used experimentally and clinically to mitigate the effects of cystic fibrosis. In accordance with the present invention, C18 was chosen for studies because it has previously been demonstrated that it directly interacts with and stabilizes wild-type CFTR, thereby increasing CFTR expression at the plasma membrane (15).

In accordance with the present invention, it was first confirmed that C18 is able to increase the expression of CFTR in both mice and humans. Cerebral arteries obtained from naïve mice treated with C18 (3 mg/kg daily for 2 days) exhibited higher levels of CFTR protein expression compared with vehicle-treated controls ( Fig. 2A ); C18 did not affect CFTR mRNA expression (Fig. 2B). To confirm the effect of C18 on human CFTR, heterologous expression of a baby hamster kidney fibroblast cell system stably expressing a human CFTR construct was used. Treatment with C18 (6 μmol/L; 24 h) more than doubled CFTR protein expression in baby hamster kidney fibroblast cells (Fig. 2C); C18 did not affect CFTR mRNA expression in this system (Fig. 2D). Taken together, the data are consistent with previous observations that C18 has a direct stabilizing effect on CFTR protein expression (15) and demonstrates its effectiveness against murine and human CFTR.

We then assessed whether C18 treatment increases CFTR content under cell culture conditions that sufficiently mimic the relevant pathophysiological mechanisms and cellular environment. As previously demonstrated (4), TNF (10 ng/ml) reduces CFTR protein expression in primary vascular smooth muscle cells and thus mimics the TNF-dependent down-regulation of CFTR observed in HF (4) and SAH (Figure 2E) (6). As expected, decreased CFTR expression correlated with attenuated FITC-S1P uptake (i.e., a CFTR-dependent process (4); Fig. 2F) and forskolin-stimulated iodide efflux (i.e., a classic measure of CFTR activity (13); Fig. 2G). Treatment with C18 (6 μmol/L; 24-h co-incubation with 10 ng/ml TNF followed by 24-h incubation with 10 ng/ml TNF) reversed the decrease in CFTR expression (Fig. 2E) and completely restored FITC-S1P uptake (Fig. . 2F) and iodide efflux (Fig. 2G). These results are entirely consistent with the suggestion made herein that, under conditions where CFTR is decreased, the proteostatic effect of C18 increases overall CFTR expression and restores its activity.

C18 Treatment Reverses Cerebral Blood Flow Deficiency in Heart Failure

As previously published by Meissner et al. (4), induction of HF stimulates a significant decrease in CFTR protein expression in the cerebral artery (Fig. 3A), which coincides with a marked increase in PCA myogenic tone (Fig. 3B). As predicted by the data in FIG. 2, in vivo treatment with C18 (3 mg/kg daily for 2 days) restored CFTR expression in cerebral arteries obtained from mice with HF (Fig. 3A) and simultaneously normalized PCA myogenic tone (Fig. 3B). Decreased myogenic tone is associated with an expected shift in response to phenylephrine due to decreased basal tone; however, EC ₅₀ values do not differ (log EC ₅₀ sham-operated: -6.30 ± 0.14, n = 6 from 4 mice; log EC ₅₀ CH: -6.25 ± 0.07, n = 5 from 3 mice; log EC ₅₀ CH + C18: -5.94 ± 0.21, n = 5 from 4 mice; Kruskal-Wallis test P=NS), and the curves overlap after correction for differences in basal tone (Fig. 11). C18 treatment did not affect PCA myogenic tone or responses to phenylephrine in sham-operated mice (Fig. 12). Using PMA obtained from CFTR knockout mice, we confirmed that C18 mediates its attenuating effect on myogenic tone through its action on CFTR. As expected, PMAs from CFTR knockout mice exhibited increased myogenic tone that was insensitive to C18 treatment in vivo ( Fig. 3C ); treatment with C18 did not affect the response to phenylephrine (Fig. 13).

At the systemic level, CH induction significantly reduced CO compared with sham-operated controls ( Fig. 3D ; measured 6 weeks post-infarction); TPR increases as a compensatory response (Fig. 3E) to prevent a significant decrease in SBP (Fig. 3F) (3, 4). Despite the relatively small reduction in SBP, CBF decreased markedly (Fig. 3G). In vivo treatment with C18 restores CBF in mice with HF (Fig. 3G), closely correlating with normalization of PCA myogenic tone (Fig. 3B). C18 does not improve the cardiovascular damage caused by the left anterior descending coronary artery ligation procedure: therefore, the improvement in CBF can be clearly attributed to a vascular mechanism. Moreover, since C18 does not affect TPR or SBP (Fig. 3), C18-mediated CBF restoration must represent a localized microvascular effect that is independent of changes in systemic hemodynamic parameters. This therapeutic profile is ideally consistent with the previously described use of etanercept (3).

In addition to maintaining constant perfusion when systemic pressure fluctuates, the myogenic response also protects the fragile capillary bed from damaging pressure (16) and maintains capillary hydrostatic pressure at levels that minimize edema formation (17). In this context, therapeutic reduction of cerebral artery myogenic tone, even from the elevated levels observed in HF, could potentially cause the counterproductive side effect of cerebral edema formation. Thus, noninvasive edema imaging was used to confirm that neither HF nor C18 treatment in the context of HF caused overt edema in any brain region ( Fig. 4 ).

C18 Treatment Reverses Cerebral Blood Flow Deficiency in SAH

Previous work demonstrates that SAH has a similar cerebrovascular phenotype to HF in that (i) CFTR protein expression is reduced in the cerebral artery; (ii) increased myogenic tone of the cerebral artery; and (iii) cerebral perfusion is reduced (6). As in HF (Fig. 3), in vivo treatment with C18 (3 mg/kg daily for 2 days) restored CFTR expression in the cerebral arteries of SAH mice (Fig. 5A) and simultaneously normalized myogenic tone of the olfactory cerebral artery (Fig. 5B ). The decrease in myogenic tone is associated with an expected baseline shift in the response to phenylephrine due to a decrease in basal tone; however, EC ₅₀ values did not differ (log EC ₅₀ sham: -5.52 ± 0.23, n = 5 from 3 mice; log EC ₅₀ SAH: -6.01 ± 0.16, n = 6 from 6 mice; log EC ₅₀ SAH + C18: -6.01 ± 0.31, n = 5 from 4 mice; Kruskal-Wallis test P=NS), and the curves overlap after correction for differences in basal tone (Fig. 14). Treatment with C18 did not affect myogenic tone of the olfactory cerebral arteries or responses to phenylephrine in sham-operated mice (Fig. 15). Using olfactory cerebral arteries obtained from CFTR knockout mice, we confirmed that C18 mediates its attenuating effect on myogenic tone by targeting CFTR: as expected, olfactory cerebral arteries from CFTR knockout mice exhibit increased myogenic tone, which not treatable C18 in vivo. (Fig. 5C); C18 treatment did not affect phenylephrine sensitivity in these arteries (Fig. 16). Importantly, in vivo treatment with C18 restored CBF ( Fig. 5D ), which again correlated with normalization of olfactory medullary artery myogenic tone ( Fig. 5B ).

Treatment with lumacaftor reverses cerebral blood flow deficits in SAH

Data on C18 are complemented by reports of treatment with lumacaftor, a CFTR corrector, a clinically relevant analogue of C18 approved by the US Food and Drug Administration for the treatment of cystic fibrosis, in combination with Ivacaftor (i.e., Orkambi®), a CFTR enhancer. For the first time, lumacaftor was confirmed to be able to increase CFTR expression in both mouse and human: as observed for C18 (Figure 2), lumacaftor increased CFTR protein expression in the cerebral arteries of mice (mice administered 3 mg/kg daily for 2 days ) and baby hamster kidney fibroblast cells stably expressing human CFTR (Fig. 6). In both cases, CFTR mRNA expression was unchanged, suggesting a nontranscriptional mechanism (Fig. 6). Consistent with the present data on C18 in SAH (Fig. 5), in vivo treatment with lumacaftor (3 mg/kg per day for 2 days) restores CFTR expression in the cerebral arteries of mice with SAH (Fig. 6) and simultaneously normalizes myogenic tone olfactory cerebral artery (Fig. 6) and cerebral perfusion (Fig. 6). Lumacaftor does not affect phenylephrine sensitivity (Fig. 17), and therefore EC ₅₀ values do not differ (log EC ₅₀ sham: -5.79 ± 0.09, n = 7 from 4 mice; log EC ₅₀ SAH: - 5.93 ± 0.19, n = 6 from 3 mice; log EC ₅₀ SAH + Lum: -5.90 ± 0.18, n = 7 from 4 mice; Kruskal-Wallis test P=NS).

C18 and lumacaftor reduce neuronal damage in SAH

Neuronal damage caused by SAH can be easily characterized by standard histological techniques (fluorinephritis and activated caspase-3 staining) and simple neurological testing (modified Garcia score) (6): these techniques were used to determine whether restoration of normal myogenic response correlated and CD with improved neurological outcome. Indeed, both C18 and lumacaftor dramatically reduced neuronal damage in SAH, as assessed by activated caspase-3 and FJ staining ( Fig. 7 ); the reduction in neuronal damage in C18-treated mice correlated with improved neurological function ( Fig. 7E ). Specifically, SAH mice scored lower on the Modified Garcia Test of Neurological Function than sham-operated mice; however, SAH mice treated with C18 had neurological function scores comparable to sham-operated mice.

Discussion

Previous work has refined the original finding that CFTR inhibition increases microvascular myogenic tone into a new mechanistic concept (4, 6): at its core, CFTR critically regulates S1P degradation and thus largely regulates cerebrovascular tone in health and disease. diseases (4). The present invention applies this knowledge to the treatment of HF and SAH. The present invention provides a new class of therapeutic agents for the treatment of diseases of the cardiovascular system. The present invention provides in vivo verification that CFTR is a cerebrovascular regulator and confirms that it can be used to improve cerebral perfusion in both HF and SAH. As the first new microvascular-targeted drug to emerge in recent years, therapeutics that enhance CFTR function may represent an untapped resource for managing a wide range of pathologies that cause cerebrovascular dysfunction and cerebral perfusion deficits.

The proximal PCA was strategically selected for the HF studies presented herein, and the olfactory cerebral arteries were selected for the SAH studies according to the present example to directly compare CFTR-targeted treatments with previous studies (3, 4, 6). Although the PCA and olfactory arteries originate from different regions of the cerebral microcirculation (i.e., posterior and anterior, respectively) and exhibit differences in baseline myogenic tone curves, they nevertheless behave similarly in terms of pathological signaling mechanisms that increase myogenic sensitivity ( 3, 4, 6). The comparable success of CFTR-targeted therapies in the PCA and olfactory arteries suggests that CFTR regulates vascular reactivity broadly throughout the cerebral microcirculation.

Clearly, CFTR does not regulate vascular responsiveness in all vascular beds, as demonstrated by comparisons of (i) cerebral and skeletal muscle resistance arteries, (ii) TPR measurements in CFTR mutant and wild-type mice, and (iii) the lack of effect of C18 treatment on TPR. Differences in CFTR expression measured by qPCR explain why CFTR inhibition regulates myogenic tone in cerebral arteries but has no apparent role in skeletal muscle resistance arteries. Taken together, the present data demonstrate that CFTR therapeutics will have limited effects that are specific to discrete microvascular beds (eg, cerebral microcirculation). This property is a prerequisite for effective redistribution of blood flow and provides a significant therapeutic advantage over common vasodilators, which can dangerously lower blood pressure by indiscriminately dilating blood vessels.

Interestingly, while acute/chemical inhibition of CFTR had no effect on cremaster resistance arteries, germline CFTR gene deletion slightly reduced myogenic tone in this specific microvascular bed. Because global deletion of CFTR results in a complex cardiovascular phenotype (18), this small discrepancy is explained by an indirect and/or adaptive response. The present inventors have previously demonstrated the effectiveness of a CFTR inhibitor in both cellular (4) and isolated arterial (6) systems. Thus, it can be concluded that CFTR plays a minimal role in modulating the myogenic tone of the cremaster artery.

The myogenic response is the basis of autoregulation of CBF, constant coordination of vascular resistance with the prevailing transmural pressure. Thus, interventions aimed at increasing pathologically decreased CBF should aim to restore normal myogenic reactivity rather than indiscriminately suppress vasoconstriction. By specifically normalizing myogenic reactivity in HF and SAH, the CFTR-targeted treatments of the present invention are expected to preserve both CBF autoregulation and responsiveness to local vasoactive stimuli (eg, neurovascular coupling) in vivo.

The myogenic response not only determines brain perfusion through its autoregulatory function, but also maintains capillary pressure at levels that minimize damage and edema formation (16, 17). Consequently, decreased myogenic reactivity may cause edema to form, allowing additional pressure to enter the capillary beds. In this regard, since S1P significantly regulates blood-brain barrier permeability (19), CFTR-dependent changes in S1P signaling may cause edema due to changes in barrier function. Thus, it was extremely important to exclude edema as a serious negative effect of the treatment method of the present invention. Indeed, treatment with CFTR corrector (C18) does not induce edema in mice with heart failure. This confirms that: (i) hydrostatic pressure remains within acceptable limits (this would be expected since CBF and SBP measurements are comparable to sham-operated animals); and (ii) blood-brain barrier function is maintained during treatment.

To date, neither cerebrovascular expression of CFTR in human cerebral arteries nor cerebral perfusion has been determined in patients with cystic fibrosis. Clinically, cystic fibrosis is not associated with cognitive impairment or ischemic injury; however, this does not exclude a decrease in cerebral perfusion: as has been demonstrated previously (3, 20), the brain can tolerate a moderate decrease in CBF without crossing an obvious damage threshold. Thus, cognitive impairment would not necessarily be expected in young and relatively healthy patients with cystic fibrosis. Although there is no doubt that hypoperfusion significantly contributes to cognitive decline (1) and cerebral damage (21), other “hits to the system,” including aging and inflammatory responses caused by heart disease, are required to overcome the threshold for damage caused by clinically insensible decreases in CBF. vascular system or SAH.

Access to human cerebral artery samples is difficult: thus, we relied on transfer data involving systematic comparisons of CFTR expression in mesenteric and skeletal muscle arteries in humans and mice (22) to determine whether CFTR could be expressed in human cerebral arteries . The CFTR protein is present in human mesenteric arteries, and its functional inhibition results in the expected increase in myogenic reactivity (22): CFTR is therefore confirmed as a modulator of human resistance artery myogenic reactivity. Unlike human mesenteric arteries, human skeletal muscle resistance arteries do not contain detectable CFTR protein and are therefore insensitive to CFTR inhibition (22). Because the functional profile of mesenteric and skeletal muscle arteries overlaps for humans and mice (i.e., CFTR modulates mesenteric but not skeletal muscle resistive arteries; Fig. 1 (4, 22)), it can be concluded that the functional profile also overlaps for cerebral microcirculation . In the absence of direct assessments, this assumption is currently the most reasonable basis for predicting the effect of CFTR modulators on the human cerebrovascular system.

Corrector compounds are currently used to treat cystic fibrosis caused by the CFTR ΔF508 mutation: these compounds help in escorting misfolded mutant CFTR proteins to the plasma membrane. Neither the pathology of deficient CFTR trafficking nor supportive therapeutic interventions are applicable to HF and SAH, where transcription-based downregulation of CFTR expression occurs. Thus, it was critical to confirm that CFTR modulators, preferably CFTR correctors, particularly C18 and lumacaftor, increase wild-type CFTR in experimental conditions where CFTR protein expression is reduced. Indeed, the present data indicate that CFTR modulators, preferably CFTR correctors, particularly C18 and lumacaftor, also increase the amount of wild-type CFTR through a nontranscriptional mechanism. This proteostatic effect stabilizes cell surface-localized CFTR from internalization and subsequent degradation (15). Over time, stabilization of CFTR at the cell surface increases overall levels of CFTR expression as less CFTR at the cell surface is internalized and processed by degradation mechanisms. Therapeutic restoration of CFTR expression in the disease models used in the present invention coincides with attenuation of myogenic tone, normalization of CBF and SAH, and reduction of neuronal damage.

Because CFTR is a critical regulatory protein in several organs, most notably the lungs, the effects of CFTR therapeutics on avascular tissues must also be considered. Notably, HF suppresses CFTR expression in the epithelial cells of the terminal bronchioles of the lung (4), and thus the reduction of CFTR is not limited to the microcirculation in these conditions (i.e., HF causes a “cystic fibrosis phenotype” that potentially causes multiorgan failure). Therefore, CFTR therapeutics may have broad and significant benefits for patients with heart failure beyond improving cerebral perfusion. In this context, the fact that several serious comorbidities are common to both HF and cystic fibrosis (eg, secondary pulmonary hypertension (24)) gives rise to the provocative but still controversial claim that certain secondary pathologies in HF either are caused or aggravated by insufficient CFTR activity and are therefore corrected by CFTR therapeutics.

conclusions

The present invention provides the first evidence that CFTR regulates cerebrovascular reactivity and therefore cerebral perfusion: this positions CFTR as the “master switch” in the control of cerebral perfusion. Both HF and SAH chronically reduce cerebral perfusion by suppressing CFTR protein expression in the cerebral artery, thereby impairing autoregulatory control. The present invention discloses that clinically available CFTR therapeutics can restore cerebral artery CFTR expression, vascular reactivity, and cerebral perfusion. It is noteworthy that this therapeutic effect is localized to the cerebral microcirculation, since CFTR does modulate the responsiveness of peripheral arteries involved in blood pressure control. Thus, CFTR therapy is emerging as a valuable clinical tool for the treatment of cerebrovascular dysfunction, impaired cerebral perfusion, and neuronal damage. The present invention discloses the effectiveness of CFTR therapeutics for preventing and improving cerebral perfusion deficits, particularly those caused by HF and SAH.

The following references are provided throughout this document, with the exception of the Methods and Reagents section above, which has a separate list of references.

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3. Yang J, Hossein Noyan-Ashraf M, Meissner A, et al. Proximal Cerebral Arteries Develop Myogenic Responsiveness in Heart Failure via Tumor Necrosis Factor-α-Dependent Activation of Sphingosine-1-Phosphate Signaling. Circulation 2012;126:196-206.

4. Meissner A, Yang J, Kroetsch JT, et al. Tumor Necrosis Factor-α-Mediated Downregulation of the Cystic Fibrosis Transmembrane Conductance Regulator Drives Pathological Sphingosine-1-Phosphate Signaling in a Mouse Model of Heart Failure. Circulation 2012;125:2739-50.

5. Meissner A, Visanji NP, Momen MA, et al. Tumor Necrosis Factor-α Underlies Loss of Cortical Dendritic Spine Density in a Mouse Model of Congestive Heart Failure. J Am Heart Assoc 2015;4:e001920.

6. Yagi K, Lidington D, Wan H, et al. Therapeutically Targeting Tumor Necrosis Factor-α/Sphingosine-1-Phosphate Signaling Corrects Myogenic Reactivity in Subarachnoid Hemorrhage. Stroke 2015;46:2260-70.

7. Murdaca G, Spanò F, Contatore M, et al. Infection risk associated with anti-TNF-α agents: a review. Expert Opin Drug Saf 2015;14:571-82.

8. Kroetsch JT, Levy AS, Zhang H, et al. Constitutive smooth muscle tumor necrosis factor regulates microvascular myogenic responsiveness and systemic blood pressure. Nat Commun 2017;8:14805.

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Claims (3)

1. The use of a CFTR corrector (cystic fibrosis transmembrane conductance regulator) to prevent and/or treat a decrease or impairment of cerebral perfusion, where the CFTR corrector is a cyclopropane carboxamide derivative selected from the group consisting of lumacaftor, tezacaftor, C18 and their pharmaceutically acceptable salts and solvates. 2. Use of a CFTR corrector according to claim 1, wherein the CFTR corrector is lumacaftor. 3. The use of a CFTR corrector according to claim 1 or 2, where the decrease in cerebral perfusion is associated with a condition selected from the group consisting of heart disease, heart failure, subarachnoid hemorrhage, sudden sensorineural hearing loss, vascular dementia, arterial hypertension, ischemic stroke, hemorrhagic stroke, diabetes and Alzheimer's disease.