Female mice display sex-specific differences in cerebrovascular function and subarachnoid haemorrhage-induced injury

Summary

Background

In male mice, a circadian rhythm in myogenic reactivity influences the extent of brain injury following subarachnoid haemorrhage (SAH). We hypothesized that female mice have a different cerebrovascular phenotype and consequently, a distinct SAH-induced injury phenotype.

Methods

SAH was modelled by pre-chiasmatic blood injection. Olfactory cerebral resistance arteries were functionally assessed by pressure myography; these functional assessments were related to brain histology and neurobehavioral assessments. Cystic fibrosis transmembrane conductance regulator (CFTR) expression was assessed by PCR and Western blot. We compared non-ovariectomized and ovariectomized mice.

Findings

Cerebrovascular myogenic reactivity is not rhythmic in females and no diurnal differences in SAH-induced injury are observed; ovariectomy does not unmask a rhythmic phenotype for any endpoint. CFTR expression is rhythmic, with similar expression levels compared to male mice. CFTR inhibition studies, however, indicate that CFTR activity is lower in female arteries. Pharmacologically increasing CFTR expression in vivo (3 mg/kg lumacaftor for 2 days) reduces myogenic tone at Zeitgeber time 11, but not Zeitgeber time 23. Myogenic tone is not markedly augmented following SAH in female mice and lumacaftor loses its ability to reduce myogenic tone; nevertheless, lumacaftor confers at least some injury benefit in females with SAH.

Interpretation

Female mice possess a distinct cerebrovascular phenotype compared to males, putatively due to functional differences in CFTR regulation. This sex difference eliminates the CFTR-dependent cerebrovascular effects of SAH and may alter the therapeutic efficacy of lumacaftor compared to males.

Funding

Brain Aneurysm Foundation, Heart and Stroke Foundation and Ted Rogers Centre for Heart Research.

Research in context.

Evidence before this study

• •Subarachnoid haemorrhage (SAH) is a devastating type of stroke with few treatment options.
• •In male mice, the cystic fibrosis transmembrane regulator (CFTR) is a potent regulator of cerebral artery vasoconstriction.
• •In male mice, a circadian rhythm in CFTR expression influences both cerebrovascular reactivity and the degree of brain injury that follows SAH.
• •In male mice, the CFTR therapeutic lumacaftor substantially improves brain injury following SAH.

Added value of this study

• •Female mice possess a distinctly different cerebrovascular phenotype compared to males.
• •Ovariectomy does not unmask a cerebrovascular rhythm, indicating that female sex hormones do not explain the phenotypic difference.
• •The CFTR expression profile in males and females is similar; CFTR's capacity to modulate cerebovascular reactivity, however, appears to differ between males and females.
• •The effect of SAH on cerebrovascular constriction and the effects of lumacaftor as an SAH intervention differ between males and females.

Implications of all the available evidence

• •Biological sex differences may impact the therapeutic value of CFTR correctors as an intervention for attenuating brain injury following SAH.

Introduction

Aneurysmal subarachnoid haemorrhage (SAH) is a devastating type of stroke with high mortality and morbidity.¹ As with other brain injuries,² the time of day that an SAH occurs significantly impacts the extent of injury that subsequently ensues.^3,4 Defining the diurnal/circadian mechanisms that drive these time-of-day differences is vital since, in addition to impacting injury severity and outcome, these mechanisms are also likely to influence the efficacy of therapeutic interventions.²

Our recent work in male mice demonstrates that: (i) cerebral resistance arteries possess a circadian rhythm in myogenic reactivity and (ii) the level of myogenic tone at the time of SAH strongly associates with the degree of SAH-induced injury.⁴ At the molecular level, circadian oscillations in cystic fibrosis transmembrane conductance regulator (CFTR) channel expression appear to drive the myogenic reactivity rhythm in male cerebral arteries and, by association, the level of SAH-induced injury.⁴ CFTR is a prominent modulator of cerebrovascular reactivity: in addition to its role as a chloride channel, its sphingolipid transporter function⁵ sequesters the myogenic signalling mediator sphingosine-1-phosphate away from its receptors, thereby preventing the activation of the pro-constrictive signalling pathways that augment myogenic vasoconstriction.6, 7, 8, 9

Given that our previous work focused solely on male mice,⁴ one key question to resolve is whether the male phenotype is generalizable to females. Remarkably, there are several reasons to propose that females will possess distinctly different vascular and SAH injury phenotypes compared to their male counterparts: (i) biological sex differences in cerebrovascular function, cerebral blood flow autoregulation and cerebrovascular pathologies are well-documented¹⁰; (ii) female sex hormones are known to modulate CFTR expression,^11,12 inflammatory processes¹⁰ and cellular stress responses¹⁰; and (iii) females appear to have a higher incidence of delayed cerebral ischemia at 3–5 days post-SAH.¹³ Circadian time is not frequently incorporated into experimental plans and thus, most studies addressing sex differences have been conducted during daylight hours. Unfortunately, this hampers the generalizability of the reported observations, as any identified difference (or lack thereof) could be sensitive to the circadian time of data collection.

This investigation examined whether biological sex differences exist in cerebral artery myogenic reactivity and how this influences injury following experimental SAH. To understand the role of sex hormones in females, we conducted assessments in both naïve and ovariectomized female mice. Our previous work establishes a strong association between cerebral resistance artery myogenic reactivity and SAH-induced injury⁴: thus, we hypothesized that this association persists in females. Indeed, our data show that this association is preserved in females, although naïve and ovariectomized female mice possess a distinctly different vascular phenotype compared to males.

Methods

Ethics

This investigation conforms to the National Research Council’s 2011 Guide for the Care and Use of Laboratory Animals (ISBN: 0-309-15400-6). All experimental procedures were approved by the Institutional Animal Care and Use Committee at the University of Toronto (Protocol ID# 20011424).

Mice

Wild-type male and female mice (C57BL/6N) were purchased from Charles River Laboratories (Montreal, Canada). Surgically ovariectomized female mice were professionally prepared by Charles River Laboratories. All mice were housed in a controlled climate (21 °C, 40–60% humidity) with a standard 12 h:12 h light–dark cycle, fed normal chow and had ad libitum access to water and food. Mice were used for experiments at 8–12 weeks of age and acclimatized to the light cycle for 2 weeks prior to experimentation. Upon reaching experimental endpoints, animals were deeply anesthetized with 5% isoflurane and euthanized by decapitation (tissue collection) or exsanguination/perfusion fixation (brain histology). To align with our previous work, the majority of data were collected at Zeitgeber time 11 (ZT11) and Zeitgeber time 23 (ZT23), since these time points define the peak and trough of the cerebrovascular myogenic rhythm in male mice.⁴

Induction of subarachnoid haemorrhage

We utilized a blood injection model of SAH, as previously described.⁴ Surgeries were completed within 30 min of ZT11, Zeitgeber time 19 (ZT19) or ZT23; surgeries at ZT19 and ZT23 were conducted under red light illumination to mitigate the impact of light on the entrained circadian rhythm. All mice received a single pre-operative dose of sustained release buprenorphine (1.0 mg/kg s.c.). Each mouse was anaesthetized (5% isoflurane induction, 1–2% maintenance) and fixed in a stereotactic 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 advanced to the prechiasmatic cistern and 80 μl of arterial blood from a syngeneic donor mouse, obtained by cardiac puncture, was injected into the intracranial space over 10 s. Following injection, the scalp incision was closed and the mice recovered in heated cages. Humane endpoints (dyspnea, vocalizing, dehydration, lethargy, weight loss >20%) were monitored twice daily; no animals required humane euthanasia. Experimental endpoints were collected at 48 h post-SAH induction. Mortality associated with the SAH procedure (5 out of 89 surgeries; ∼5%) occurred acutely following blood injection.

Pressure myography

Vascular reactivity was assessed by pressure myography, as previously described.⁴ Mouse olfactory cerebral arteries (a first branch of the anterior cerebral artery) were carefully dissected, cannulated onto micropipettes, stretched to their in vivo lengths, pressurized to 45 mmHg and warmed to 37 °C. The arteries were imaged with a CCD camera at 40× magnification during myography measurements, with luminal diameter measured using a Crescent Electronics (Windsor, Canada) video edge detector and logged using Photon Technology International FeliX32 analysis software (Horiba Canada Inc.; London, Canada).

All functional experiments were conducted in 3-morpholinopropanesulfonic acid (MOPS) buffered saline, with no perfusion ([mmol/L]: NaCl 147.0, KCl 4.7, CaCl₂ 1.5, MgSO₄ 1.2, NaH₂PO₄ 1.2, pyruvate 2.0, EDTA 0.02, MOPS 3.0 and glucose 5.0; all salts from BioShop Canada Inc., Burlington Canada). Vasomotor responses to 10 μmol/L phenylephrine (Millipore Sigma; Oakville, Canada) provided an assessment of vessel viability at the beginning of each experiment: arteries failing to show ≥30% constriction were considered damaged/compromised and excluded. A total of 26 out of 200 cerebral arteries were excluded for failing the viability test, which computes to a technical success rate of 87%.

Myogenic responses were elicited by stepwise 20 mmHg increases in transmural pressure from 20 mmHg to 80 mmHg. At each pressure step, vessel diameter (dia_active) was measured once a steady state was achieved. Following completion of all dia_active measurements, the MOPS buffer was replaced with a Ca²⁺-free version and maximal passive diameter (dia_max) was recorded at each pressure step. Myogenic tone was calculated as the percent constriction in relation to the maximal diameter at each respective transmural pressure: tone (% of dia_max) = [(dia_max-dia_active)/dia_max]×100, where dia_active is the vessel diameter in MOPS containing Ca²⁺ and dia_max is the diameter in Ca²⁺-free MOPS. Analyses of vasomotor responses to phenylephrine used the same calculation, only in this case, dia_active represents the vessel diameter at steady state following application of phenylephrine.

Histology

The histological procedures have been described previously.^4,14 Briefly, brains were fixed via aortic perfusion with 10% buffered formalin. Samples were then embedded in paraffin blocks. Standard procedures were utilized to prepare slides with 5 μm thick coronal slices starting at −2 mm from bregma, which includes the left and right temporal and parietal lobes.⁴ The slides were subsequently deparaffinized with xylene and rehydrated and graded levels of ethanol in distilled water.

For Fluoro-Jade staining, slides were serially incubated with 1% NaOH/80% ethanol (5 min), 70% ethanol (2 min), distilled water (2 min) and 0.06% potassium permanganate (10 min). After washing with deionized water, brain slices were stained with 0.0004% Fluoro-Jade C in 0.1% acetic acid (15 min). The samples were then washed with deionized water, dried and mounted.

For cleaved caspase-3/NeuN double-staining, slides were treated with citrate-based Antigen Unmasking Solution at 96 °C for 30 min (Vector Laboratories; Burlingame, USA; cat# H-3300-250). The slides were subsequently permeabilized with 0.3% Triton X-100 in PBS for 1 h at room temperature and then blocked with 10% goat serum in PBS containing 1% bovine serum albumin for 30 min at room temperature. Slides were incubated with the anti-active cleaved caspase-3 antibody overnight at 4 °C (1:200 dilution in 1% BSA in PBS; BD Biosciences; Mississauga, Canada; cat# 559565, RRID:AB_397274) and subsequently conjugated with Alexa Fluor 488-labelled goat anti-rabbit IgG for 1 h at room temperature (1:500 dilution in 1% BSA in PBS; ThermoFisher Scientific cat# A-11008, RRID:AB_143165). The slides were then incubated with mouse monoclonal anti-NeuN antibody for 1 h at room temperature (1:200 dilution in 1% BSA in PBS; Millipore cat# MAB377, RRID:AB_2298772) and subsequently conjugated with Alexa Fluor 568-labelled goat anti-mouse IgG for 1 h at room temperature (1:500 dilution in 1% BSA in PBS; ThermoFisher Scientific cat# A-11004, RRID:AB_2534072). Cell nuclei were then stained with DAPI (2 μg/ml in PBS) for 15 min. After washing, the specimens were mounted with CC Mount (Millipore Sigma; cat# C9368).

For microglia staining, slides were treated with citrate-based Antigen Unmasking Solution and subsequently blocked with serum-free protein block for 10 min at room temperature (Agilent Technologies Canada Inc.; Mississauga, Canada; cat #X090930-2). Slides were then co-incubated overnight (4 °C) with rabbit anti-ionized calcium binding adaptor molecule 1 (Iba1; 1:1000 dilution; FujiFilm Wako Chemicals USA Corp; Richmond, USA; cat# 019–19741; RRID:AB_839504) and rat anti-F4/80 (1:100 dilution; Abcam Inc.; Toronto, Canada; cat# ab6640; RRID:AB_1140040) in background-reducing antibody diluent (Agilent Technologies cat# S302283). The slides were subsequently conjugated with Alexa Fluor 555-labelled goat anti-rabbit IgG (1:200 dilution; ThermoFisher Scientific; cat# A32732; RRID:AB_2633281) and Alexa-Fluor 647-labelled goat anti-rat IgG (1:200 dilution; ThermoFisher Scientific; cat# A48265; RRID:AB_2895299) for 1 h at room temperature in background-reducing antibody diluent. Cell nuclei were then stained with DAPI (2 μg/ml in PBS) for 5 min. After washing, the specimens were mounted with Vectashield Vibrance anti-fade mounting medium (Vector Laboratories; cat# H-1700).

Digital imaging and analysis

Digital images of the caspase-3 and Fluoro-Jade staining were acquired at 20× magnification using a FV3000 laser confocal microscope under constant settings (Olympus Life Science, Richmond Hill, Canada). Image overlays were constructed with freely available ImageJ 1.44p software (National Institutes of Health, USA): positively-stained neuronal cells were manually counted in the cortical region of the full coronal brain slice.

Microglia number and morphologic activation were analysed in the cortical region using HALO software (version 3.6.4134.137) from Indica Labs (Albuquerque, USA; RRID:SCR_018350). Each slide was digitally imaged at 20× magnification with an Olympus VS-120 slide scanner and loaded into the HALO software: the HALO Object Colocalization FL v2.1.4 module quantified the number of Iba/F4/80 double-positive cells and the HALO Microglial Activation FL v1.0.6 module assessed morphologic activation, using previously established threshold settings for differentiating activated from non-activated microglia.¹⁴

Neurological function in SAH mice

Neurological function was assessed utilizing the modified Garcia score, as previously described.⁴ The neurological assessment consists of 6 domains: spontaneous activity, spontaneous movement of all 4 limbs, forepaw outstretching, climbing, body proprioception and response to vibrissae touch. The maximum score is 18, indicative of normal neurological function. Modified Garcia assessments at ZT23 were conducted under red light conditions.

Western blots

CFTR Western blots were conducted as previously described.⁴ Briefly, cerebral arteries mechanically ground 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. Lysate proteins were electrophoretically separated using 7% acrylamide gels and transferred onto polyvinylidene difluoride membranes. Membranes were blocked for 45 min in 5% non-fat skim milk in phosphate-buffered saline containing 1% Tween 20 (PBST) and subsequently incubated overnight at 4 °C with primary antibody. The 100–280 kDa weight range of the blot was incubated with rabbit polyclonal anti-CFTR primary antibody (1:1000 in 5% milk/PBST; Alomone Labs cat# ACL-006, RRID:AB_2039804), while the remaining range (35–100 kDa) was incubated with mouse monoclonal anti-α-tubulin primary antibody (1:5000 in 5% milk/PBST; Cell Signalling Technology cat# 3873, RRID:AB_1904178). Following primary antibody incubation, the blots were washed with PBST and then conjugated with peroxidase-labelled secondary antibody (1:2000 in 5% milk/PBST) for 2 h at room temperature: the CFTR blot received peroxidase-labelled goat anti-rabbit IgG antibody (Cell Signalling Technology Cat# 7074, RRID:AB_2099233) and the tubulin blot received peroxidase-labelled horse anti-mouse IgG antibody (Cell Signalling Technology Cat# 7076, RRID:AB_330924). After washing, the blots were treated with chemiluminescence reagent. Digital images were collected with a Bio-Rad Laboratories (Mississauga, Canada) ChemiDoc system and analysed with Bio-Rad Laboratories Image Lab software. Uncropped blots for all Western blot experiments are presented in Supplemental Fig. S1.

Quantitative PCR

We have previously described the procedures for preparing cerebral artery cDNA and performing quantitative PCR.⁴ Briefly, Cerebral artery RNA was isolated with Norgen Biotek (Thorold, Canada) “Total RNA Purification Micro” spin columns (cat #35300), using the proteinase K digestion and DNA removal procedures, as directed by the manufacturer’s instructions. The RNA was converted to cDNA using a “Superscript III” reverse transcription kit (ThermoFisher Scientific; Mississauga, Canada; cat# 18080044), according to the manufacturer’s directions. Residual RNA was removed by incubating the resulting cDNA with RNAse H (0.125 U/μl).

Quantitative PCR was performed using a Bio-Rad Laboratories CFX384 Real Time PCR Detection System. Each PCR reaction contained Power SYBR® Green PCR master mix (ThermoFisher Scientific) and rigorously validated primer sets (400 nmol/L in each reaction). Gene targets and negative controls (water) were assessed in triplicate. The PCR amplification consisted of 10 min denaturation at 95 °C, followed by 40 cycles of amplification (15 s at 95 °C + 60 s at 60 °C). Following amplification, the amplicons were melted: the resulting dissociation curve confirmed the production of single products. Transcript expression levels in mouse tissues were calculated from the ΔCt values relative to the standard housekeeping gene glucose-6-phosphate dehydrogenase (G6PD). To confirm that G6PD was reliable for normalization, transcript expression levels were also calculated from the ΔCt values relative to hydroxymethylbilane synthase (HMBS) and glyceraldehyde 3-phosphate dehydrogenase (GAPDH). The PCR primers are described in Supplemental Table S1.

Pharmacological interventions

Lumacaftor (AdooQ Bioscience cat # A10986 via Cedarlane Laboratories; Burlington, Canada) was prepared as a 10x stock in DMSO and stored as frozen aliquots. These stocks were diluted in corn oil (Sigma–Aldrich) to working concentrations and delivered in vivo by intraperitoneal injection as a 50 μl bolus; vehicle controls received 10% DMSO in corn oil. CFTR_(inh)-172 (Sigma–Aldrich cat #C2292) and JTE013 (Cayman Chemical Co. cat # 10009458 via Cedarlane Laboratories) were prepared in DMSO, stored as frozen aliquots and diluted 1000-fold for in vitro application.

Antibody characterization

All antibodies used in the present study were purchased from commercial sources and have been previously validated for specificity in western blotting or immunohistochemistry applications by the manufacturer. We have not undertaken specific antibody validation experiments in the present study.

Statistics

This study conforms to the ARRIVE 2.0 guidelines.¹⁵ The data supporting the findings of this study are available from the corresponding author upon reasonable request. This study utilized 241 experimental mice and 77 blood donor mice over a 2-year period. A complete cohort breakdown and experimental timeline appears in Supplemental Tables S2 and S3.

Upon arrival to the animal facility, mice were arbitrarily distributed into cages and acclimatized without a randomization methodology. For interventions, acclimatized animals were randomly assigned to blocks with defined sizes (http://www.randomizer.org). We used previously published data to estimate the experimental group sizes necessary to provide an 80% power level for the detection of the anticipated differences between groups with a two-tailed alpha level of 0.05. For our myography experiments, we presumed that the previously described ZT11/ZT23 differential in males (μ1 = 34.2, σ1 = 5.0, μ2 = 25.4, σ2 = 2.6, n1/n2 = 1)⁴ would adequately estimate group sizes for all myography assessments/interventions (calculated group size n = 5). Our other calculations included: (i) ZT11/ZT23 differences in histological positive cell counts (μ1 = 80, σ1 = 45, μ2 = 162, σ2 = 62, n1/n2 = 1; calculated group size n = 8), modified Garcia scores (x∼1 = 16.0, σ1.0 = 1, x∼2 = 14.3, σ2 = 1.3, n1/n2 = 0.86; calculated group size n = 8) and CFTR protein expression (μ1 = 1.0, σ1 = 0.2, μ2 = 0.6, σ2 = 0.2, n1/n2 = 1.1; calculated group size n = 6)⁴; and (ii) the effect of CFTR therapeutics on histological positive cell counts (μ1 = 166, σ1 = 52, μ2 = 67, σ2 = 47, n1/n2 = 1.2; calculated group size n = 6) and modified Garcia scores (x∼1 = 15.0, σ1.0 = 1.4, x∼2 = 17.0, σ2 = 1.5, n1/n2 = 0.85; calculated group size n = 10).⁶

Histological assessments were conducted under blinded conditions (a non-assessor concealed the identity of the stained slides and revealed them post-assessment). All other endpoints were collected without blinding, as the assessment ZT times could not be concealed. There were no exclusions for primary endpoints; exclusions related to technical failure (non-viable vessels) were conducted as stated above in the pressure myography subsection.

Data were analysed with Graphpad Prism 9 software (Graphpad Software; San Diego, USA). Modified Garcia scores (ordinal data) are presented as box and whisker plots displaying median, upper and lower interquartile range and upper and lower extremes; all other data are presented as means ± standard deviation. In both cases, n represents the number of independent measures. Prior to conducting statistical comparisons, we assessed data normality (Shapiro–Wilk test and QQ-plots) and variance homogeneity (F test or Brown–Forsythe test). Normal data with equal variances were analysed with a two-tailed independent t-test or one-way ANOVA; normal data with unequal variances were analysed with a two-tailed Welch’s t-test or a Welch’s ANOVA with a Dunnet T3 post-hoc test; non-normal data were analysed with a two-tailed Mann–Whitney U Test or a Kruskal–Wallis Test with Dunn’s post-hoc test. Differences were considered significant at p < 0.05. Data statistics, including statistical comparison outcomes and mean difference with 95% confidence intervals are presented in Supplemental Tables S4 and S5.

Circadian rhythms were identified using JTK_CYCLE (version 3.1) with R software (version 3.4.0), a non-parametric test designed to reliably identify rhythmicity (a significant rhythm required p < 0.05).¹⁶ The JTK_CYCLE algorithm combines the Jonckheere-Terpstra (JT) non-parametric test for detecting monotonic orderings across ordered independent groups and Kendall’s tau, a measure of rank correlation between two measures.¹⁶ JTK_CYCLE applies the JTK algorithm to alternative hypothesized group orderings, effectively fitting multiple cosine curves to the data in order to determine the optimal combination of period and phase that minimizes the exact p-value of Kendall’s tau correlation.¹⁶ The resulting minimal p-value is Bonferroni-adjusted for multiple testing. All JTK_CYCLE analysis results are found in Supplemental Table S6.

Role of funders

The study sponsors played no role in the study design; the collection, analysis and interpretation of the data; manuscript preparation; or the decision to submit for publication.

Results

Female mice do not display circadian variations in myogenic tone and SAH-induced injury

In contrast to our previous observations from male cerebral arteries,⁴ the JTK_CYCLE analysis software did not identify a 24-h rhythm in female cerebral artery myogenic vasoconstriction (Fig. 1a; Supplemental Table S6). Strikingly, when the female tracing is qualitatively compared to our previously published male data,⁴ the female profile has the appearance of a “dampened rhythm”, with the female tone level aligning with the trough level of tone observed in male mice (i.e., the male ZT11 level; Supplemental Fig. S2). Phenylephrine-stimulated vasoconstriction is not identified as rhythmic in female cerebral arteries, which matches our previous observations in male arteries (Fig. 1b; Supplemental Table S6).⁴ Consistent with the lack (or very low amplitude) of rhythmicity, a comparison of female cerebral arteries isolated at ZT11 and ZT23 shows that Zeitgeber time has a very small effect of on myogenic tone and agonist-stimulated vasoconstriction (Fig. 1c; Supplemental Table S4); in contrast, a ZT11/ZT23 myogenic tone comparison for male cerebral arteries assessed in parallel (Fig. 1d) shows data compatible with a much larger effect size (Supplemental Table S4).

Fig. 1.

Female mice do not display circadian variations in myogenic tone.Panels a and b display wild-type female olfactory cerebral artery (a) myogenic tone and (b) phenylephrine-stimulated vasoconstriction plotted over Zeitgeber time (n = 5 arteries from 3 mice at Zeitgeber times 3, 11, 15 and 23, n = 6 arteries from 3 mice at Zeitgeber time 7 and n = 6 arteries from 4 mice at Zeitgeber time 19). Data are double-plotted for visualization purposes; white shading indicates “lights on” and dark shading indicates “lights off”. Neither myogenic tone nor phenylephrine-stimulated vasoconstriction display a statistically significant circadian rhythm according to JTK_CYCLE analysis. (c) Female olfactory cerebral artery myogenic tone and phenylephrine-stimulated vasoconstriction measured in arteries isolated at Zeitgeber time 11 (ZT11; n = 5 arteries from 3 mice) and Zeitgeber time 23 (ZT23; n = 5 arteries from 3 mice). (d) Male olfactory cerebral artery myogenic tone measured in arteries isolated at ZT11 (n = 8 arteries from 4 mice) and ZT23 (n = 6 arteries from 4 mice). All data are means ± standard deviation. In panels a and b, the Bonferroni-adjusted JTK_CYCLE p value for the given transmural pressure or PE concentration is shown below the respective graph. Panel c used an independent t-test (40–80 mmHg and 100 nmol/L), Welch’s corrected independent t-test (10 μmol/L) or a Mann–Whitney U test (20 mmHg, 1–10 nmol/L and 1 μmol/L) for statistical comparisons. Panel d used an independent t-test (40–80 mmHg) or a Mann–Whitney U test (20 mmHg) for statistical comparisons. Comparison test p-values are shown above the error bars.

We confirmed that the core molecular clock rhythm in female cerebral arteries is comparable to that of males: the mRNA expression patterns for Bmal1 (positive arm) and Per2 (negative arm) in female cerebral arteries are circadian (Supplemental Fig. S3) and have comparable calculated amplitudes/acrophases as those previously observed in male cerebral arteries (Supplemental Table S6).⁴ Thus, the apparent lack myogenic rhythmicity in female mice is not attributable to differences in core molecular clock timing.

Fig. 2a shows representative images of activated and non-activated microglia cells, with quantifications of cell counts and activation levels; sub-analyses of average nuclear area, average process length, and average process area are provided in Supplemental Fig. S4. In contrast to our previous work in male mice,⁴ the data in Fig. 2a and Supplemental Fig. S4 are not compatible with the hypothesis that there are meaningful differences in microglia number and/or their activation state when SAH is induced at ZT23 versus ZT11 (Supplemental Table S4). Representative images of cortical cells stained with/for Fluoro-Jade and activated caspase-3 are shown in Fig. 2b, with quantifications in Fig. 2c and d (2-channel images of the activated caspase-3 staining are provided in Supplemental Fig. S5). Since the female myogenic tone tracing reasonably aligns with the low ZT11 level of tone observed in male arteries (Supplemental Fig. S2), we expected that the SAH-induced injury in females would align with the ZT11 injury level previously described in male mice.⁴ Indeed, the caspase-3 positive staining and neurofunctional scores observed in females largely match the previously described ZT11 injury levels in male mice (Supplemental Fig. S2). However, the data do not support the hypothesis that SAH-induced injury in female mice is more severe when SAH is induced at ZT23 versus ZT11 (Supplemental Table S4), as previously reported in males.⁴ Modified Garcia scores are slightly higher (indicating better neurofunctional outcome) when SAH is induced at ZT23 versus ZT11 (Fig. 2e).

Fig. 2.

Female mice do not display circadian variations SAH-induced injury. Subarachnoid haemorrhage (SAH) was induced in wild-type female mice at Zeitgeber Time 11 (ZT11) or Zeitgeber Time 23 (ZT23); behavioural assessments and tissue collection were conducted 48 h afterwards. (a) Representative images of microglia double-stained with ionized calcium binding adaptor molecule 1 (Iba) and F4/80 are shown on the left. The top panel shows a highly fenestrated cell, while the bottom panel shows a rounded morphology (bar = 20 μm, pink = Iba, green = F4/80, blue = DAPI). To the right of the images are quantifications of double-positive cell counts and percent activated cells (based on morphology) when SAH is induced at ZT11 (n = 8 brain slices from 8 mice) or ZT23 (n = 8 brain slices from 8 mice). (b) Representative images of cortical cells stained for/with Fluoro-Jade (left) and activated caspase-3 (right); arrows point to positively stained neuronal cells (red = NeuN, green = activated caspase-3, cyan = DAPI). For all images, bar = 60 μm. Positive neuronal cell counts for (c) Fluoro-Jade staining and (d) activated caspase-3 when SAH is induced at ZT11 (n = 8 brain slices from 8 mice) or ZT23 (n = 8 brain slices from 8 mice). (e) Modified Garcia scores when SAH is induced at ZT11 (n = 8 mice) or ZT23 (n = 8 mice). Data in Panel e (modified Garcia scores) are ordinal data and are presented as box and whisker plots displaying median, upper and lower interquartile range and upper and lower extremes; all other data are presented as means ± standard deviation. Statistical comparisons were completed with an independent t-test in panels a and d; a Welch’s-corrected independent t-test in panel c; and a Mann–Whitney U test in panel e. Comparison test p-values are shown above the error bars.

Collectively, the data presented in Fig. 1, Fig. 2 and Supplemental Fig. S2 strongly reinforce the association that we previously described between myogenic tone at the time of SAH and the subsequent SAH-induced injury level.⁴ Of note, we did not compare Fluoro-Jade staining across different experiments: we have found that the method is highly-sensitive to experimental variations and consequently, are not confident that separate data sets are directly comparable.

Ovariectomy does not unmask a cerebrovascular rhythm in females

We hypothesized that ovariectomy would “unmask” a hidden cerebrovascular rhythm in female mice that drives higher levels of myogenic tone and SAH-induced injury at ZT23: data collected from ovariectomized female mice, however, do not support the hypothesis. Both myogenic reactivity (Fig. 3a) and phenylephrine-stimulated vasoconstriction (Supplemental Fig. S6) in cerebral arteries isolated from ovariectomized female mice remain non-rhythmic, according to JTK_CYCLE analysis (Supplemental Table S6). Intriguingly, the new myogenic reactivity profile displays a prominent “isolated peak” in myogenic tone at ZT19 (Fig. 3a). When qualitatively compared to the naïve female data, the myogenic tracing from ovariectomized females displays reasonable alignment with the naïve myogenic tone level, except for the aforementioned ZT19 peak (Supplemental Fig. S7). Vascular reactivity data from arteries isolated from ovariectomized female mice at ZT11 and ZT23 (Fig. 3b) indicate that Zeitgeber time has a very small effect on vascular reactivity; contrary to our hypothesis, the data are more compatible with ZT11 possessing higher tone compared ZT23 than vice versa (Supplemental Table S4). In terms of SAH-induced injury in ovariectomized female mice at 2 days post-SAH (representative images of cortical cells stained with/for Fluoro-Jade and activated caspase-3 are shown in Fig. 3c; 2-channel images of the activated caspase-3 staining are provided in Supplemental Fig. S5), cortical Fluoro-Jade (Fig. 3d) and activated caspase-3 (Fig. 3e) positive cell counts are more compatible with either a non-substantive effect or reduced injury when SAH is induced at ZT23 versus ZT11 than vice versa (Supplemental Table S5); likewise, modified Garcia score data (Fig. 3f) are more compatible with an improved outcome when SAH is induced at ZT23, rather than a detrimental outcome. Taken together, we conclude that ovarian sex hormones do not explain the differences between males and females with respect to circadian variations in cerebral artery myogenic tone and time-of-day differences in SAH-induced injury.

Fig. 3.

Ovariectomy does not unmask circadian variations in myogenic tone or SAH-induced injury. (a) Myogenic tone in cerebral arteries isolated from ovariectomized female mice (OVX) is plotted over Zeitgeber time (n = 5 arteries from 3 mice at Zeitgeber times 7 and 19, n = 6 arteries from 3 mice at Zeitgeber times 3 and 15, n = 7 arteries from 4 mice at Zeitgeber time 11 and n = 8 arteries from 5 mice at Zeitgeber time 23). Data are double-plotted for visualization purposes; white shading indicates “lights on” and dark shading indicates “lights off”. No statistically significant circadian rhythm is present according to JTK_CYCLE analysis. (b) Myogenic and phenylephrine-stimulated vasoconstriction in olfactory cerebral arteries isolated from OVX mice at Zeitgeber time 11 (ZT11; n = 7 arteries from 4 mice) and Zeitgeber time 23 (ZT23; n = 8 arteries from 5 mice). In Panels c-f, subarachnoid haemorrhage (SAH) was induced in OVX mice at ZT11 or ZT23; behavioural assessments and tissue collection were conducted 48 h afterwards. (c) Representative images show cortical cells stained for Fluoro-Jade (left) and activated caspase-3 (right); arrows point to positively stained neuronal cells (Bar = 60 μm; red = NeuN, green = activated caspase-3, cyan = DAPI). Positive neuronal cell counts for (d) Fluoro-Jade staining and (e) activated caspase-3 when SAH is induced at ZT11 (n = 10 brain slices from 10 mice) or ZT23 (n = 8 brain slices from 8 mice). (f) Modified Garcia scores when SAH is induced at ZT11 (n = 10 mice) or ZT23 (n = 8 mice). Data in Panel f (modified Garcia scores) are ordinal data and are presented as box and whisker plots displaying median, upper and lower interquartile range and upper and lower extremes; all other data are presented as means ± standard deviation. In panel a, the Bonferroni-adjusted JTK_CYCLE p value for the given transmural pressure is shown below the respective graph. Statistical comparisons in panel c used a Welch’s corrected independent t-test (20 mmHg) or an independent t-test (all other comparisons). Statistical comparisons in panel d used a One-Way ANOVA (data analysed in conjunction with data presented in Supplemental Fig. S8); panel e used a Kruskal–Wallis Test with Dunn’s post-test (data analysed in conjunction with data presented in Supplemental Fig. S8); and panel f used a Mann–Whitney U test. Comparison test p-values are shown above the error bars.

The “isolated peak” observed at ZT19 in ovariectomized female mice provides a unique opportunity to further examine the relationship between cerebrovascular tone and SAH-induced injury. Our vascular data predicts that, despite only 4 h of time separation, SAH-induced injury should be higher when the SAH ictus occurs at ZT19, relative to ZT23. As shown in Supplemental Fig. S8 and Table S5, data for activated caspase-3 positive cell counts and Fluoro-Jade positive cell counts are compatible with a medium-to-large effect of Zeitgeber time, with higher injury occurring at ZT19, as hypothesized. This Supplementary analysis strongly reinforces our previously documented association between vascular tone at the time of SAH and the degree of injury that subsequently ensues.⁴

Since the effect of Zeitgeber time on myogenic tone, activated caspase-3 staining and modified Garcia scores in both naïve/non-ovariectomized and ovariectomized mice appears to be small, we pooled the ZT11/ZT23 data collected in Fig. 1, Fig. 2, Fig. 3. This yielded additional statistical power to examine whether ovariectomy alters myogenic reactivity and/or SAH-induced injury. Given the vasodilative and neuroprotective properties widely associated with oestrogen,¹⁰ we hypothesized that ovariectomy would increase myogenic tone, increase positive cell counts and decrease modified Garcia scores. Surprisingly, ovariectomy had very small effects on myogenic tone (Fig. 4a) and activated caspase-3 positive cell counts (Fig. 4b); in contrast to our hypothesis, these data are more compatible with either a non-substantive effect or lower myogenic tone/positive cell counts in the ovariectomized group, than vice versa (Supplemental Table S4). However, consistent with our hypothesis, ovariectomized mice display a clear tendency for reduced modified Garcia scores (Fig. 4c and Supplemental Table S4).

Fig. 4.

Ovariectomy does not alter myogenic tone or SAH-induced injury. The myogenic tone, activated caspase-3 and modified Garcia score data presented in Fig. 1, Fig. 2, Fig. 3 were pooled (i.e., Zeitgeber time 11 and Zeitgeber time 23 data were combined) into non-ovariectomized (No OVX) and ovariectomized (OVX) groups. Shown are No OVX/OVX comparisons for (a) myogenic tone (No OVX n = 10 arteries from 6 mice; OVX n = 15 arteries from 9 mice), (b) caspase-3 positive cell counts at 48 h post-subarachnoid haemorrhage (No OVX n = 16 brain slices form 16 mice; OVX n = 18 brain slices from 18 mice) and (c) modified Garcia scores at 48 h post-subarachnoid haemorrhage (No OVX n = 16 mice; OVX n = 18 mice). Data in Panel c (modified Garcia scores) are ordinal data and are presented as box and whisker plots displaying median, upper and lower interquartile range and upper and lower extremes; all other data are presented as means ± standard deviation. Statistical comparisons in panel a used an independent t-test (40 and 80 mmHg) or Mann–Whitney U test (20 and 60 mmHg). A Mann–Whitney U test was used for statistical comparisons in panels b–c. Comparison test p-values are shown above the error bars.

CFTR expression is oscillatory in female mice, but is only a modest regulator of myogenic tone

Our previous report asserted that rhythmic CFTR expression underlies the circadian rhythm in myogenic reactivity observed in male mice.⁴ We therefore assessed whether CFTR oscillates in female cerebral arteries and subsequently, whether CFTR plays a significant role as a regulator of myogenic tone. Consistent with our previous observations in male mice⁴: (i) the JTK_CYCLE analysis algorithm found that CFTR mRNA expression in female cerebral arteries is rhythmic, with an acrophase at Zeitgeber 10 (Fig. 5a and Supplemental Table S6); and (ii) CFTR is more highly expressed at ZT11 relative to ZT23 at the protein level (Fig. 5b; Supplemental Table S4). However, as shown in Fig. 5c, specific CFTR channel inhibition (100 nmol/L CFTR_(inh)-172) at ZT11, when CFTR expression is high, has only a very small effect on myogenic tone (Supplemental Table S4). Not surprisingly, CFTR channel inhibition in female cerebral arteries isolated at ZT21, when CFTR expression is low, also has a very small effect on myogenic tone (Supplemental Fig. S9 and Table S4). These data support the conclusion that CFTR is not a significant regulator of female cerebral artery tone. However, the functional differences between male and female cerebral arteries⁴ are unlikely to be attributable to lower CFTR protein expression levels in female arteries, as CFTR expression data from male and female cerebral arteries at ZT11 (male = 1.00 ± 0.47 arbitrary units, n = 6 samples from 6 mice; female = 1.22 ± 0.26 arbitrary units, n = 6 samples from 6 mice; p = 0.336 by independent t-test) indicate that female arteries are more likely to have comparable or higher CFTR expression levels as male arteries, than they are to have lower CFTR expression levels (for all CFTR expression statistics, see Supplemental Table S4).

Fig. 5.

CFTR expression and function in female cerebral arteries. (a) CFTR mRNA expression in female cerebral arteries is plotted over Zeitgeber time (n = 3 samples from 3 mice at each Zeitgeber time point). Data are double-plotted for visualization purposes; white shading indicates “lights on” and dark shading indicates “lights off”. CFTR mRNA expression exhibits a statistically significant circadian rhythm, according to JTK_CYCLE analysis. (b) A comparison of CFTR protein expression in cerebral arteries isolated from female mice at Zeitgeber Time 11 (ZT11; n = 9 samples from 9 mice) versus Zeitgeber Time 23 (ZT23; n = 9 samples from 9 mice). (c) The effect of CFTR channel inhibition (100 nmol/L CFTR (inh)-172 for 30 min in vitro) on myogenic tone in female cerebral arteries isolated at ZT11 (n = 6 arteries from 3 mice; paired comparisons). (d) Myogenic tone and phenylephrine-stimulated vasoconstriction in female cerebral arteries isolated at ZT11 from mice treated with lumacaftor (3 mg/kg/day; n = 5 arteries from 3 mice) or vehicle (n = 6 arteries from 4 mice) for 2 days. (e) Myogenic tone in female cerebral arteries isolated at ZT23 from mice treated with 3 mg/kg/day lumacaftor (n = 5 arteries from 3 mice), 30 mg/kg/day lumacaftor (n = 7 arteries from 4 mice) or vehicle (n = 10 arteries from 5 mice) for 2 days. All data are means ± standard deviation. In panel a, the Bonferroni-adjusted JTK_CYCLE p value is shown below the graph. Statistical comparisons in panel b used an independent t-test; panel c used paired t-tests; and panel d used either an independent t-test (20 mmHg and 1–10 nmol/L) or a Mann–Whitney U test (40–80 mmHg and 100 nmol/L-10 μmol/L). Statistical comparisons in panel e used either a Kruskal–Wallis test (20 mmHg) or a One-Way ANOVA (40–80 mmHg). Comparison test p-values are shown above the error bars.

Given the modest regulatory role that CFTR plays in female cerebral arteries, we questioned whether in vivo lumacaftor treatment, which stabilizes and thereby increases CFTR protein levels at the plasma membrane,^4,17 modulates cerebrovascular reactivity. Lumacaftor treatment in naïve female mice (3 mg/kg/day for 2 days) profoundly reduces myogenic tone in cerebral arteries isolated at ZT11 (Fig. 5d; Supplemental Table S4); this was associated with reduced PE-stimulated vasoconstriction at the 1 μmol/L concentration (i.e., within the dynamic range of responsiveness), but otherwise, lumacaftor had a relatively small effect on all other PE concentrations tested (Fig. 5d; Supplemental Table S4). Accordingly, there is a clear rightward shift in the log EC₅₀ value (mean difference of 0.7 log units ± 0.4 log units error margin; Supplemental Table S4). Lumacaftor treatment in female mice does not meaningfully reduce myogenic tone at ZT23, even when the lumacaftor dose is increased to 30 mg/kg/day (Fig. 5e; Supplemental Table S4). This result contrasts our previous observations in male mice, where lumacaftor treatment (3 mg/kg/day) substantially attenuated myogenic tone at ZT23.⁴ Thus, as a positive control, we reproduced the previously observed lumacaftor treatment effect at ZT23 in male mice (Supplemental Fig. S10 and Table S4).

In male cerebral arteries, CFTR exerts its modulatory effect on myogenic vasoconstriction by sequestering S1P from its pro-constrictive sphingosine-1-phosphate subtype 2 receptor.^7,9 Using 1 μmol/L JTE013 (30 min incubation in vitro), we confirmed that female cerebral arteries are sensitive to sphingosine-1-phosphate receptor 2 antagonism (Supplemental Fig. S11 and Table S4), as previously demonstrated for male cerebral arteries.^8,9

The SAH phenotype differs in females compared to males

Wild-type female mice underwent the SAH or sham surgical procedure in the light phase between ZT7 and ZT11; functional assessments and histological specimen collection were conducted 48 h post-surgery. In parallel, we assessed myogenic reactivity in a small cohort of male sham and SAH mice. In stark contrast to male mice, which reproduced the large myogenic tone augmentation following SAH that we have previously documented (Supplemental Fig. S12 and Table S4),^4,6,9 the effect of SAH on cerebrovascular myogenic reactivity in female mice is rather small (Fig. 6a; Supplemental Table S5). Further, female SAH mice become “resistant” to lumacaftor treatment (3 mg/kg/day) and no longer display the large reduction in myogenic tone that is observed in naïve females at ZT11 (compare Figs. 6a to 5d; Supplemental Table S5).

Fig. 6.

Effect of Lumacaftor on cerebrovascular myogenic tone and brain injury following subarachnoid haemorrhage in female mice. Wild-type female mice underwent the subarachnoid haemorrhage (SAH) or sham surgical procedure in the light phase (between Zeitgeber Time 7 and Zeitgeber Time 11); behavioural assessments and tissue collection were conducted 48 h afterwards. (a) Myogenic tone in female cerebral arteries isolated from sham-operated mice (n = 5 arteries from 3 mice), SAH mice with vehicle treatment (n = 6 arteries from 3 mice) and SAH mice with lumacaftor treatment (Lum; 3 mg/kg i.p. daily for 2 days; n = 5 arteries from 3 mice). (b) Shown are representative images of cortical cells stained for Fluoro-Jade (top) and activated caspase-3 (bottom); arrows point to positively stained neuronal cells (bar = 60 μm; red = NeuN, green = activated caspase-3, cyan = DAPI). (c) Positive neuronal cell counts for Fluoro-Jade staining and activated caspase-3 in sham (n = 8 brain slices from 8 mice), SAH (n = 10 brain slices from 10 mice) and lumacaftor-treated SAH mice (n = 10 brain slices from 10 mice). (d) Modified Garcia scores for sham (n = 8), SAH (n = 10) and lumacaftor-treated SAH mice (n = 10). Data in Panel d (modified Garcia scores) are ordinal data and are presented as box and whisker plots displaying median, upper and lower interquartile range and upper and lower extremes; all other data are presented as means ± standard deviation. Statistical comparisons in panel a used a One-Way ANOVA; panel c used a Kruskal–Wallis test with Dunn’s post-test (Fluoro-Jade) or a Welch’s corrected ANOVA with Dunnet T3 post-test (Caspase-3); and panel d used a Kruskal–Wallis test with Dunn’s post-test. Comparison test p-values are shown above the error bars.

Our previous work demonstrated that lumacaftor has a large beneficial effect on SAH-induced injury in male mice.⁶ However, based on our observation that lumacaftor has only a small effect on myogenic tone in female mice following SAH, we hypothesized that lumacaftor would confer minimal benefit in females. The data presented in Fig. 6b–d, Supplemental Fig. S5 and Table S5 do not support this hypothesis: indeed, the data are compatible with lumacaftor possessing at least some beneficial effect in female mice. Specifically, data for fluoro-jade positive staining (Fig. 6c) support a small-to-medium effect, although “no benefit” also falls well within the 95% confidence interval (−88 mean difference ± 189 error margin); data for caspase-3 positive staining (Fig. 6c) are consistent with a much larger effect (−229 mean difference ± 193 error margin) that qualitatively, compares well with our previous effect size in male mice⁶; finally, modified Garcia scores (Fig. 6d) are consistent with a small improvement in neurobehavioural outcomes.

Discussion

Our previous work demonstrated that (i) myogenic reactivity in olfactory cerebral arteries isolated from male mice oscillates with a circadian rhythm and (ii) male mice display a diurnal difference in SAH-induced injury, with the extent of injury associating with the level myogenic tone at the time of SAH.⁴ The present study demonstrates that female mice possess a distinctly different phenotype: there is no statistical rhythm in myogenic reactivity, nor is there a diurnal difference in SAH-induced injury.

Circadian rhythms are present in virtually all living organisms: their entrainment to the solar day/night cycle allows organisms to anticipate recurring environmental patterns and to adapt their physiology and behaviours accordingly.^18,19 Time of day is a crucial biological variable that, unfortunately, is rarely incorporated into experimental designs, despite a clear influence on both behaviour and experimentally-induced pathologies.^4,18,19 Within the circadian research field, female subjects remain underrepresented, despite clear evidence of physiological sex differences in circadian timing and animal behaviour.¹⁹ Incorporating these variables into scientific research is essential for generalizability, translation and ensuring safety, efficacy and benefit across the entire population. To emphasize the danger of ignoring female sex in drug development, a United States government audit of prescription drugs withdrawn from the market between 1997 and 2001 found that 8 out of 10 medications disproportionately affected women: 4 posed a greater health risk to women compared to men (terfenadine, mibefradil, astemizole and cisapride), while another 4 had higher use in women (fenfluramine, dexfenfluramine, troglitazone and alosetron).^20,21 Likewise, the Hygia Chronotherapy Trial has shown that timing matters, with the key finding that the “traditional” recommendation of taking anti-hypertension medications in the morning is actually less efficacious and carries higher risk than taking the medications in the evening.²²

In this context, the critical observation of the present study is that the cerebrovascular myogenic reactivity rhythm previously described in males⁴ is dampened to the extent that it is functionally absent in females. To the best of our knowledge, this is the first time such a profound difference between the sexes has been demonstrated at the circadian level. We initially postulated that CFTR expression differences may be explanatory, since our previous work indicated that a circadian rhythm in CFTR expression underpinned the myogenic rhythm observed in male mice.⁴ However, contrary to our expectations, we did not observe differences in molecular clock timing, the circadian CFTR mRNA expression pattern (both timing and magnitude) or CFTR protein expression.⁴ CFTR is clearly capable of modulating female artery myogenic tone: lumacaftor treatment, which increases cerebral artery CFTR expression,⁶ amplifies the non-significant “fingerprint of a rhythm” observed in Fig. 1a and imparts a phenotype comparable to males (i.e., reduced myogenic tone at ZT11 in relation to ZT23).⁴ We conclude, therefore, that there must be a sex difference in cerebrovascular CFTR channel regulation that requires females to have higher CFTR expression levels than males for a comparable modulatory influence.

There is precedence for sex differences in CFTR regulation: in a model of polycystic kidney disease, primary renal epithelial cells from male mice displayed CFTR-dependent currents, while those from females did not, despite similar CFTR expression levels.²³ This sex disparity was attributed to higher intracellular calcium levels in males relative to females.²³ Several other significant regulators of CFTR activity,²⁴ including Protein Kinase A (PKA),²⁵ Protein Kinase C (PKC)²⁶ and adenosine monophosphate-activated protein kinase (AMPK),²⁷ also show higher activity levels in males, compared to females. While differential CFTR activity/regulation may explain why CFTR lacks a modulatory influence in females, this mechanistic model creates an apparent paradox: reduced CFTR activity in female mice should be expected to yield the higher ZT23 level of myogenic tone observed in males, not the lower ZT11 level shown in Supplemental Fig. S2.⁴ This paradox can be reconciled by the fact that the same entities that regulate CFTR activity (e.g., calcium, PKC, PKA, etc.) are also important regulators of myogenic signalling and thus, we speculate that the reduced CFTR function in females is offset by dampened myogenic signalling.

Female sex hormones have well-known acute effects on vascular contractility²⁸ and thus, we were surprised to observe that ovariectomy has no effect on myogenic reactivity. Similarly, oestrogens are widely considered to be neuroprotective,¹⁰ and it was therefore surprising that ovariectomy did not enhance SAH-induced activated caspase-3 staining and elicited only a tendency for worse modified Garcia scores following SAH. Although classically considered an ovarian sex hormone, oestrogen is synthesized by astrocytes and neurons, providing a brain-specific source of oestrogen that serves neuroprotective and anti-inflammatory roles.²⁹ While ovariectomy undoubtedly reduces circulating oestrogen levels, it is not clear how ovariectomy affects brain oestrogen levels in our SAH model. Thus, the results stemming from our ovariectomy model must be viewed cautiously.

Consistent with the observations from our ovariectomized mouse model, several studies in eumenorrheic human female subjects also indicate that ovarian sex hormones minimally influence autoregulatory function.30, 31, 32 Within the field, however, studies evaluating the influence of sex hormones on vascular function have generated heterogeneous and frequently contradictory results^33,34: these discrepancies arise from many factors, including human subject characteristics (e.g., eumenorrheic, menopausal, use of contraception, etc.) and the interventional challenges utilized to perturb the system (e.g., hypercapnia, postural changes). Additionally, the in vitro system utilized in the present study to assess vascular reactivity washes away diffusible factors and therefore, likely eliminates any reversible effect during functional assessment. Therefore, caution must be applied when comparing our myogenic data to clinical measurements in vivo.

Clinical studies have consistently shown that cerebral blood flow is 10–15% higher in females compared to males.35, 36, 37 Several mechanistic explanations have been proposed, including sex differences in brain metabolism, haematocrit/blood viscosity and hormones. However, these explanations are somewhat tenuous, as perfusion differences exist in the absence of oxygen consumption differences³⁷ and they are also present in prepubescent males/females, who have very low sex hormone levels and minimal differences in haematocrit.³⁶ Our data provide a more fundamental explanation for the observed perfusion differences: during the active/waking phase, females possess lower cerebrovascular myogenic tone compared to males. In perfect alignment with this explanation, two separate clinical studies concluded that human female subjects indeed possess lower cerebrovascular resistance than males, while maintaining comparable autoregulatory function.^31,36

Myogenic reactivity is the basis of cerebral blood flow autoregulation,⁷ a regulatory mechanism that maintains constant brain perfusion over a relatively wide range of cerebral perfusion pressures. This mechanism originates within the cerebral microcirculation, where resistance arteries actively match their level of constriction and hence, vascular resistance, to the prevalent perfusion pressure.³⁸ Cerebral autoregulation is clearly impaired in patients with SAH39, 40, 41, 42, 43 and it is a strong independent predictor of both DCI and negative outcome.41, 42, 43 Our present and previous work⁴ establishes a strong association between the level of myogenic tone at the time of SAH ictus and the degree of cerebral injury that subsequently ensues: this relationship predicts the effects of circadian time, smooth muscle cell-specific molecular clock disruption, intervention with CFTR-modifying therapeutics and biological sex differences.⁴ There is no doubt that perfusion levels within the acute phase of SAH play a significant role in the brain injury process^44,45: our myogenic tone measurements, therefore, presumably predict the perfusion level at the time of SAH and more importantly, the extent of ischemia/hypoperfusion within the early stages following SAH.

Only a handful of experimental studies have assessed the effect of sex on SAH-induced injury.^13,46, 47, 48, 49 A commonality across these previous studies is the use of vascular perforation/aneurysm rupture models that, compared to the blood injection model used in the present study, possess variable bleed severity, extended periods of elevated intracranial pressure and higher mortality. Additionally, these previous studies utilized vastly different time points for injury assessment (minutes to 21 days post-SAH) and none statistically compared male and female injury levels at 48 h post-SAH. Given these confounds, it is not practical to directly compare our study results to this previous body of work. However, the primary goal or our investigation was not to address sex differences in SAH-induced injury per se, but rather to identify potential sex differences in the cerebrovascular phenotype and assess its association to SAH-induced injury. In this regard, our vascular data predicted that female mice would display less brain injury than males when the SAH ictus occurs during the waking/active phase of the circadian cycle: a comparison of our current and previously published histological/neurofunctional data substantiate this prediction.⁴

From a translational perspective, this study identifies a critical sex difference in lumacaftor’s therapeutic efficacy in our mouse model of SAH. Our previous data shows that in male mice, lumacaftor normalizes vascular reactivity, improves cerebral perfusion and potently reduces neurological injury observed at 2 days post-SAH induction.⁶ These benefits are ostensibly derived from lumacaftor’s ability to prevent CFTR expression reductions during the early phase of the SAH pathology.⁶ The present study shows that CFTR is not an appreciable modulator of vascular tone in females, despite similar expression levels as males. Assuming that CFTR expression is down-regulated in female cerebral arteries following SAH, as is the case in male arteries,^6,9 lumacaftor would be hard pressed to elicit a vascular effect: it would not only have to counteract the pathological down-regulation, but also raise CFTR expression above normal levels. Indeed, lumacaftor lacks a vascular effect in females with SAH, suggesting that it would not improve cerebral perfusion in this setting. We did observe an obvious tendency for improved activated caspase-3 in lumacaftor-treated female SAH mice, which could indicate that reduced CFTR expression in non-vascular tissues also contributes to the SAH injury process. Neuronal cells would be a prime candidate, as we recently demonstrated that CFTR is neuroprotective in vitro.⁵⁰ In this scenario, males would benefit from the “dual actions” of improved perfusion and direct neuroprotection, while females would only benefit from the latter.

A broader translational implication of the present study is the danger of ignoring circadian time and biological sex in preclinical animal research studies. It is easy for researchers to overlook the fact that research animals and patients are very different biochemical systems at day and night; likewise, female biological sex is frequently neglected, often because the results from males are assumed to apply to females.⁵¹ Yet these factors can significantly influence the severity and outcome of not only SAH, but also other types of haemorrhagic stroke, including intracerebral haemorrhage.^52,53 If sex and circadian time indeed segregate patients into distinct subpopulations, we are unlikely to find a “one size fits all” therapeutic intervention. It is therefore crucial for pre-clinical animal studies to comprehensively assess these variables, in order to more effectively develop and guide clinical interventions.

There are several study limitations that must be acknowledged. First, differences in cerebral perfusion presumably underlie the observed association between myogenic reactivity and SAH-induced injury; however, we have not measured cerebral perfusion in the present study or our previous circadian study in males.⁴ Unfortunately, this imparts a level of uncertainty in the relationship, as myogenic reactivity is not the sole determinant of cerebral perfusion. Second, we have not characterized circulating or brain female sex hormone levels in our ovariectomized model. Given the unique phenotype that resulted (i.e., the ZT19 “myogenic peak”), we can presume that the ovariectomy procedure was successful; the mechanism of the ZT19 peak, however, is unknown and as discussed previously, the conclusions from our ovariectomized model must be tempered. Third, caution must be applied to the use of activated caspase-3 as an apoptosis marker, as activated caspase-3 has non-apoptotic functions that may be activated in brain injury models. As an example, positive activated/cleaved caspase-3 staining was found to be primarily non-apoptotic (i.e., did not overlap with TUNEL staining) in a permanent middle cerebral artery occlusion model.⁵⁴ Future studies will need to confirm the injury implications of activated caspase-3 staining in males and females with additional apoptosis markers. Finally, the relatively small group sample sizes in the present study imparts a risk of type II error and thus, our conclusions that the cerebrovascular and SAH-induced injury phenotypes differ between the sexes warrant a larger confirmative study, with contemporaneous endpoint comparisons in both males and females.

In summary, this investigation shows that female mice possess a distinctly different cerebrovascular phenotype than their male counterparts. Mechanistically, CFTR activity appears to be lower in females and consequently, CFTR fails to appreciably modulate cerebrovascular tone at any expression level within its circadian expression cycle. Female mice lack the diurnal difference in SAH-induced injury previously observed in males, fairing equal or better depending on SAH ictus. Notably, ovariectomy does not unmask a rhythmic phenotype. The lack of CFTR involvement in female vascular tone regulation strongly impacts the therapeutic value of CFTR correctors as an intervention for attenuating brain injury following SAH. Thus, translational studies clinically assessing CFTR therapeutics in cerebrovascular pathologies must be prepared to identify and quantify potential sex differences in efficacy.

Contributors

Conceptualization: DL and SSB; Investigation: DDD, HW and DL; Formal Analysis: DDD, HW and DL; Verified the Underlying Data: DDD, HW and DL; Data Curation: DDD, HW, DL and SSB; Writing - Original Draft: DL; Writing - Review and Editing: DDD, HW, DL and SSB; Project Administration: DL; Supervision: DL and SSB; Resources: SSB; Funding Acquisition: SSB. DDD and HW made an equal first-author contribution; DL and SSB made an equal senior author contribution. All authors have read and agreed to the published version of the manuscript.

Data sharing statement

The data supporting the findings of this study are available from the corresponding author upon reasonable request.

Declaration of interests

DL is a consultant for Qanatpharma AG and Aphaia Pharma AG. SSB is an executive board member of Qanatpharma and Aphaia Pharma. Neither Qanatpharma nor Aphaia Pharma had any financial or intellectual involvement in this article.