Tumor Necrosis Factor/Sphingosine-1-Phosphate Signaling Augments Resistance Artery Myogenic Tone in Diabetes

Diabetes is a global health crisis: it currently afflicts 382 million people and is expected to become the seventh leading cause of death by 2030 (1,2). Much of the disease’s morbidity/mortality can be attributed to microvascular complications arising from functional and structural changes in resistance arteries (3–5). Indeed, the World Health Organization uses early microvascular manifestations (e.g., retinopathies) to determine diagnostic/therapeutic thresholds and to monitor therapeutic success (6). Controlling blood glucose is the primary clinical intervention for diabetes; however, glycemic control only manages the risk of complications: it does not restore normal metabolic activity. Consequently, patients with good glycemic control simply slow the progression of microvascular complications, rather than prevent them (7). Understanding the underlying molecular mechanisms that alter microvascular function in diabetes, therefore, is crucial to developing new strategies to prevent debilitating vascular complications.

By altering microvascular resistance artery function, diabetes complications impair blood flow control and organ perfusion (8). Resistance arteries are the key determinants of tissue perfusion: their behavior is primarily governed by the myogenic response, an intrinsic and dynamically regulated mechanism that permits highly localized control of blood flow. We have identified a signaling mechanism that augments resistance artery myogenic responses in several etiologically distinct diseases (9–12): it relies on vascular smooth muscle cell tumor necrosis factor (TNF) signaling and ultimately enhances sphingosine-1-phosphate (S1P)-mediated vasoconstriction. As a key molecular entry point into this signaling pathway, the increase in TNF activity triggered by diabetes (13) is positioned to augment resistance artery tone. Successfully demonstrating that this pathway operates in diabetes would provide new opportunities to interrupt the molecular events that alter resistance artery function, tissue perfusion, and disease progression. Mitigating microvascular dysfunction in patients with diabetes would undoubtedly improve clinical outcomes.

We titrated a standard high-fat diet (HFD)/streptozotocin (STZ) mouse model to induce a mild form of type 2 diabetes, characterized by insulin resistance and a relatively small increase in blood glucose. We hypothesize that this complex metabolic state induces TNF-dependent S1P signaling and consequently a reversible augmentation of myogenic vasoconstriction.

The use of human subjects in this study conforms to the principles outlined in the Declaration of Helsinki and was approved by the Research Ethics Board of St. Michael’s Hospital (approval no. 11-198). Study subjects who planned to have nonemergency (elective) coronary artery bypass graft surgery were recruited from the cardiac surgery clinic at St. Michael’s Hospital.

With informed patient consent, surgeons directly provided a small piece of thoracic wall skeletal muscle (3–4 cm³) to research staff inside the operating room. The specimen was immersed in room temperature MOPS-buffered saline, placed on ice, and transported to the laboratory; once the specimen cooled, it was washed with ice-cold MOPS buffer and placed in a Petri dish. Resistance arteries (100–200 μm in diameter) were carefully dissected from the surrounding tissue, with care taken to minimize vessel tension during the isolation process (14).

This investigation adheres to the Guide for the Care and Use of Laboratory Animals published by the National Institutes of Health (publication no. 85-23, revised 1996). All animal care and experimental protocols were approved by the Institutional Animal Care and Use Committees at the University of Toronto and the University Health Network, Toronto, and were conducted in accordance with Canadian animal protection laws.

All mouse strains used in this investigation are either commercially available or have previously been described. These strains include wild type C57BL/6N mice (Charles River Laboratories, Montreal, Quebec, Canada), germ-line TNF knockout mice (TNF^−/−) (Taconic Biosciences, Hudson, NY), germ line sphingosine kinase 1 (Sphk1) knockout mice (Sphk1^−/−) (a gift from Dr. Richard L. Proia, National Institutes of Health, Bethesda, MD) (15) and inducible, smooth muscle cell–targeted TNF knockout mice (12). All mice were housed under a standard 14 h:10 h light:dark cycle and had access to food and water ad libitum.

Supplementary Fig. 1 displays the scheme used to prepare the diabetic mouse model. Mice (8 weeks of age) were randomized into three groups and fed either normal chow (NC) (10% calories from fat) or an HFD (45% calories from fat, category no. TD.06415; Teklad Custom Research Diets, Harlan Laboratories, Madison, WI). After 4 weeks, mice were fasted for 5 h. After fasting, one group of HFD mice was injected with a single dose of STZ (HFD/STZ) (100 mg/kg i.p. injection in 0.1 mmol/L sodium citrate, pH 5.5); the other two groups (HFD and NC) received a sodium citrate vehicle injection. Tail vein blood samples at 72 h post–STZ injection displayed elevated blood glucose, consistent with β-cell injury. Mice were maintained on their respective diets for 12 additional weeks, at which point resistance arteries were isolated for assessment. Mesenteric (207 ± 3 μm in diameter, n = 178), olfactory cerebral (113 ± 2 μm, n = 140), cremaster skeletal muscle (79 ± 2 μm, n = 15), and posterior cerebral arteries (PCAs) (147 ± 3 μm, n = 28) were microsurgically isolated as previously described (10–12,16).

Resistance arteries were carefully cannulated onto micropipettes, pressurized, and warmed to 37°C in MOPS-buffered saline as previously described (10–12,16). Mouse olfactory and PCAs were pressurized to 45 mmHg; mouse mesenteric and human skeletal muscle resistance arteries were pressurized to 60 mmHg; and all treatments were diluted in MOPS-buffered saline.

Myogenic tone was calculated as the percent steady-state constriction after stepwise (20 mmHg increments) increases in transmural pressure in relation to the maximal diameter: tone (% of dia_max) = ([dia_max − dia_active]/dia_max) × 100, where dia_active is the vessel diameter in MOPS buffer containing Ca²⁺ and dia_max is the diameter in Ca²⁺-free MOPS buffer—both at the specified transmural pressure. Vasomotor responses (e.g., to phenylephrine [PE]) used the same calculation; only, in this case, dia_active represents the vessel diameter at steady state after application of the given agent and dia_max represents the maximal diameter (measured under Ca²⁺-free conditions); both of these measures were obtained at the transmural pressure set at the beginning of the experiment (i.e., 45 mmHg for olfactory and PCAs and 60 mmHg for mesenteric, cremaster, and human arteries).

Random fed blood glucose levels were measured with a glucometer (Bayer Healthcare, Mississauga, Ontario, Canada), using blood drawn from the tail vein. Glycated hemoglobin (HbA_1c) was measured using an A1C Now⁺ kit (Bayer Healthcare). Plasma insulin was measured by ELISA (Crystal Chem, Downers Grove, IL). For oral glucose tolerance tests, mice were fasted for 16 h and administered 1.5 mg/g body wt glucose by oral gavage.

A standard technique (10,17) was used to invasively measure blood/intraventricular pressures (Millar SPR-671 microtip mouse pressure catheter; Inter V Medical, Inc., Montreal, Quebec, Canada). High-field preclinical MRI was used for noninvasive measurement of cerebral perfusion using an established arterial spin-labeling approach as previously described (11,18). Specific methodological details are provided in Supplementary Data.

A high-sensitivity ELISA (Quantikine HS; R&D Systems, Minneapolis, MN) was used to measure TNF in patient plasma samples. Standard procedures were used for resistance artery mRNA isolation, conversion to cDNA, and subsequent quantitative RT-PCR. Specific methodological details have previously been described (14) and are provided in Supplementary Data.

All data are expressed as means ± SEM, where n is the number of independent experiments. For comparison of multiple independent groups, a nonparametric one-way ANOVA with exact P value computation was performed (Kruskal-Wallis), followed by a Dunn multiple comparison correction. For the assessment of myogenic responses, dose-response relationships, and passive diameters, data were analyzed using a two-way ANOVA. Differences were considered significant at error probabilities of P < 0.05.

Currently, human studies have not directly assessed the impact of diabetes on myogenic responses in resistance artery beds critical to organ perfusion or blood pressure control. We found that pressure-stimulated myogenic vasoconstriction (i.e., myogenic tone) is augmented in skeletal muscle resistance arteries isolated from cardiac surgery patients with type 2 diabetes relative to arteries isolated from cardiac surgery patients without diabetes (Fig. 1); PE responses and passive diameters were not different (Fig. 1). TNF mRNA expression was higher in skeletal muscle resistance arteries isolated from patients with diabetes; circulating plasma TNF levels, however, did not differ between the two groups (Supplementary Fig. 2). Profiles for the two patient cohorts are displayed in Supplementary Table 1; the mean HbA_1c level in the cohort of patients with diabetes was 7.2 ± 0.3% (range 5.1–9.7) (n = 15), with the normal range clinically defined as 4.6–6%.

We established a type 2 diabetes mouse model for mechanistic study. The combination of HFD/STZ reliably increases blood glucose and HbA_1c levels (Fig. 2); notably, blood glucose increases within a relatively narrow range (total range 10–15 mmol/L, n = 100 [∼1.5-fold to 2.0-fold increase]). Both HFD and HFD/STZ mice displayed insulin resistance, based on insulin levels and oral glucose tolerance tests (Fig. 2). Body weights are provided in Supplementary Table 2.

We strategically examined myogenic tone in mesenteric, olfactory cerebral, PCAs, and cremaster skeletal muscle resistance arteries, as we have previously characterized signaling mechanisms that pathologically augment myogenic tone in these arteries (10–12,16). HFD/STZ augments mesenteric and olfactory cerebral artery myogenic tone without altering PE responsiveness or passive diameter; myogenic tone in PCAs and cremaster skeletal muscle resistance arteries is not affected (Fig. 3). More than 4 weeks is required to augment mesenteric and olfactory artery myogenic tone (Supplementary Fig. 3); this progressive nature indicates that the augmented myogenic tone is not caused by STZ exerting direct toxic effects on resistance arteries under these conditions. HFD alone has no effect on myogenic tone, PE responses, or passive diameter (Fig. 3); even after 10 months on an HFD, mesenteric arteries display myogenic and PE responses similar to those of NC controls (Supplementary Fig. 4). STZ treatment in NC mice increases blood glucose but does not increase mesenteric or olfactory artery myogenic tone (Supplementary Fig. 5). Acetylcholine (ACh) and sodium nitroprusside (SNP) responses are preserved in HFD/STZ mesenteric and olfactory arteries, thereby excluding endothelial dysfunction and/or impaired vasodilator responsiveness as the cause of increased myogenic responsiveness (Fig. 4). Mean arterial pressure, cardiac function, and cerebral perfusion measures are similar in all groups (Supplementary Table 2 and Supplementary Fig. 6).

As in resistance arteries isolated from patients with diabetes, TNF mRNA is elevated in cerebral and mesenteric arteries isolated from HFD/STZ mice (Supplementary Fig. 7). We attempted to pharmacologically neutralize the resultant TNF protein in vitro (1 mg/mL etanercept [ETN] for 30 min): surprisingly, ETN abolishes, rather than normalizes, mesenteric artery myogenic tone in both NC and HFD/STZ mice without impacting PE responses (Supplementary Fig. 8).

Germ line TNF gene deletion does not abolish myogenic tone, but it prevents the HFD/STZ-stimulated myogenic tone augmentation in both mesenteric and olfactory arteries (Fig. 5); PE responses are unchanged (Fig. 5), as is the HFD/STZ-stimulated blood glucose elevation (Supplementary Table 3). Interestingly, HFD/STZ treatment in TNF^−/− mice increases passive diameter (Fig. 5A and D), an effect not observed in wild-type HFD/STZ mice (Fig. 3).

Using a tissue-specific TNF gene deletion model (12), we next determined whether vascular smooth muscle cells are the relevant TNF source, as our mechanistic model predicts (11). Chronically removing smooth muscle cell TNF (i.e., treating SMMHC-Cre^ERT2/TNF^flox/flox mice with tamoxifen 17 weeks prior to functional assessment [denoted henceforth as TNF^smKO]) exerts an unexpected effect on myogenic responsiveness in mesenteric arteries. Specifically, we observed a bimodal distribution, with a subset of arteries displaying minimal myogenic tone. We exclude compromised viability as the cause, since contractile responses to PE were virtually identical to those in vessels with robust myogenic responses (i.e., PE maximal constriction and half-maximal effective concentration values were similar in arteries with and without myogenic responses [data not shown]). Since arteries isolated from Cre-expressing, tamoxifen-treated wild-type mice (HFD/STZ-treated SMMHC-Cre^ERT2/TNF^wt/wt mice [denoted henceforth as TNF^smWT]) all had normal myogenic responses (Fig. 6A–C), we attribute this loss in myogenic responsiveness to the long-term removal of smooth muscle cell TNF. When we compared the relevant NC TNF^smKO and HFD/STZ TNF^smKO treatment groups, similar proportions of arteries displayed the loss of myogenic tone phenotype (NC, 4 of 8; HFD/STZ, 5 of 7). Importantly, the removal of smooth muscle TNF clearly prevents the augmentation of myogenic tone: the NC TNF^smKO and HFD/STZ TNF^smKO groups did not differ in terms of myogenic tone, PE responses, or passive diameter (Fig. 6).

In contrast to mesenteric arteries, targeted smooth muscle cell TNF gene deletion did not cause a loss of myogenic responsiveness in olfactory arteries (Fig. 6). As predicted by our model, smooth muscle cell TNF gene deletion prevents the HFD/STZ-stimulated augmentation of myogenic tone in olfactory arteries; PE responses and passive diameter were similar to those in NC controls (Fig. 6).

We have previously shown that TNF augments myogenic tone by activating the S1P-generating enzyme Sphk1 and consequently enhancing proconstrictive S1P signaling (9–12). Accordingly, germ line Sphk1 gene deletion prevents the HFD/STZ-stimulated myogenic tone augmentation in both mesenteric and olfactory arteries (Fig. 7), consistent with this proposed mechanistic link. HFD/STZ does not affect PE responses or passive diameters in arteries isolated from Sphk1^−/− mice (Fig. 7).

STZ injection in HFD mice reliably increases blood glucose in all gene knockout models used in this investigation; mouse characteristics for each genotype are presented in Supplementary Table 3. Myogenic tone, PE responses, and passive diameters in mesenteric and olfactory arteries isolated from TNF^−/− and Sphk1^−/− mice on HFD alone are not different relative to NC controls (data not shown).

Our gene deletion data identify both TNF and S1P signaling as key elements in the pathological augmentation of resistance artery myogenic tone. We therefore validated the therapeutic viability of targeting either signaling pathway, using medications that are either clinically available (ETN, an anti-TNF medication) or in clinical trials (S1P receptor antagonists) (19).

Consistent with observations in TNF^−/− and Sphk1^−/− mice, neither anti-TNF therapy (ETN: 1 injection of 1 mg/kg s.c. twice per week 14 days prior to sacrifice [10,11]) nor S1P₂ receptor antagonism (JTE013: 3 mg/kg i.p. daily for 14 days prior to sacrifice) affects the HFD/STZ-induced blood glucose elevation (Supplementary Table 3). However, both interventions successfully reverse the STZ-augmented tone in HFD mice, normalizing it in both mesenteric and olfactory arteries to the level of ETN- or JTE013-treated NC controls (Fig. 8); neither ETN nor JTE013 altered PE responses or passive diameters (Fig. 8).

The current study demonstrates that a mild form of type 2 diabetes (i.e., an HFD/STZ mouse model incorporating insulin resistance and reduced insulin production) progressively augments resistance artery myogenic responsiveness, albeit heterogeneously. Microvascular smooth muscle cell dysfunction plays the pivotal role in this vasculopathy: our investigation characterizes the signaling processes triggered and identifies new therapeutic targets that may assist in managing diabetes-associated microvascular complications.

The myogenic response is a prominent determinant of microvascular tone and consequently a key regulator of tissue perfusion. Compromised myogenic reactivity in patients with diabetes, therefore, is increasingly recognized to underlie microvascular dysfunction and insufficient organ perfusion (20). Remarkably, direct assessments in human arteries (i.e., pressure myography in isolated arteries) are sparse (21). We demonstrate, for the first time, that myogenic tone is augmented in skeletal muscle resistance arteries isolated from patients with diabetes, and this correlates with increased vascular TNF expression. Based on our previous work (11,12), we cautiously speculate that the diabetes-associated increase in vascular TNF expression drives the enhanced myogenic responses in these patients. Of note, the microvascular phenotype materializes despite reasonable blood glucose control: our patient cohort with diabetes had a mean HbA_1c level of 7%. Although this level is elevated compared with the 4–6% range found in patients without diabetes, it reasonably meets current therapeutic targets (22) and would not likely prompt further intervention.

Myogenic tone is a computed functional parameter that correlates with the activity of pressure-sensitive vascular smooth muscle cell signaling. Intriguingly, we observe enhanced myogenic tone in human skeletal muscle resistance arteries without obvious differences in either active or passive diameters (Fig. 1). Given the inherent variability in patient characteristics, comorbidities, and medications, the clear separation of the myogenic tone curves points to a highly consistent pathologic effect in patients with diabetes: we therefore conclude that the myogenic tone augmentation is genuine, albeit derived from small differences in the active and passive diameters. The functional effect we observe in the present investigation may be relatively small (Fig. 1B) because our diabetes patient cohort maintained relatively good glucose control.

We transitioned to a mouse diabetes model, since patient sample access and the general inability to manipulate the human system (e.g., systemic treatments/gene deletions) hinder mechanistic study. Although rodent models are widely used to study diabetes, relatively few investigations have published myogenic tone data. This is remarkable, given the clinical significance of microvascular complications.

To our knowledge, we are the first to assess myogenic tone in a wild-type HFD/STZ mouse model, which incorporates both insulin resistance and reduced insulin production. The present investigation demonstrates that in isolation, neither insulin resistance (HFD) nor mild blood glucose elevation (STZ alone) alters myogenic tone. Several other investigations concur with respect to the effect of HFD (23,24); however, hyperglycemia in type 1 diabetes models, which do not incorporate obesity, is associated with perturbed myogenic responsiveness (25–27). These type 1 diabetes models, however, typically induce more severe hyperglycemia than in the current study (i.e., 2.0- to 4.0-fold above baseline), which is a possible explanation for the discrepancy. We therefore conclude that the complex metabolic state induced by obesity/HFD sensitizes resistance arteries to hyperglycemia (as induced by STZ in this study); from a clinical perspective, this implies that in type 2 diabetes, even a small elevation in blood glucose should be considered hazardous.

The effect of HFD/STZ on myogenic responsiveness is heterogeneous across distinct microvascular beds: while mesenteric and olfactory cerebral arteries display augmented myogenic tone, PCAs and skeletal muscle resistance arteries do not. This heterogeneity explains why global cerebral blood flow and systemic hemodynamic parameters do not change at this time point in our model: in the brain, the microvascular effects are localized and likely compensated by collateral blood supply; systemically, skeletal muscle resistance arteries, the prominent regulators of total peripheral resistance, have not been recruited into an augmented myogenic phenotype. In principle, all four artery types are capable of increasing myogenic tone in pathological settings (11,12,16): augmenting myogenic responses in PCAs and skeletal muscle resistance arteries, therefore, may simply require more time or a more robust trigger (e.g., a higher glucose elevation). Indeed, even the enhancement of mesenteric and olfactory artery myogenic tone is far from rapid (i.e., requires >4 weeks): it is tempting to speculate, therefore, that hypertension (28) and cerebral perfusion deficits (27) emerge at later time points, as vascular beds are progressively recruited into an augmented myogenic tone phenotype.

Current HbA_1c thresholds are based primarily on the risk for retinopathies, a microvascular complication that possesses a strong relationship with blood glucose control (6,29). Yet, the association between HbA_1c and other microvascular complications is much weaker: the currently accepted thresholds, therefore, rely on a single microvascular complication that covers only a fraction of the morbidity burden in type 2 diabetes (29). This likely explains why intensive glucose control does not alleviate all diabetes complications (7,30,31). Supporting this premise, the present investigation identifies two vascular beds that are highly sensitive to impaired glucose control. Although the observed functional changes do not overtly perturb systemic hemodynamics or cerebral perfusion, they should not be underestimated: microvascular complications in diabetes are often asymptomatic during their early stages; once symptoms develop, they are generally difficult to reverse or clinically manage (32–34). Consequently, silent or asymptomatic microcirculatory dysfunction is now viewed as a key intervention point that can potentially alter the patient’s clinical course.

Many studies conclude that endothelial dysfunction, a broad range of abnormalities that disrupts the delicate balance between dilatory (most notably nitric oxide [35]) and constrictive mediators (36), is the mechanistic basis of enhanced microvascular contractility in diabetes (37,38). Our data do not support an endothelial basis for our model’s phenotype: 1) endothelial-stimulated vasodilation is preserved, and 2) the PE dose-response relationship does not left shift, which would be expected if constitutive vasodilatory signals were attenuated. Importantly, targeted TNF gene deletion localizes the primary mechanism that augments myogenic tone in this model to vascular smooth muscle cells. Since severe hyperglycemia is known to damage/alter endothelial function (39,40), we speculate that the apparent lack of endothelial dysfunction observed in the present HFD/STZ model is due to the relatively small blood glucose elevation in relation to other diabetes models.

Mechanistically, the present investigation identifies TNF as a central pathological mediator that augments mesenteric and olfactory resistance artery myogenic tone in our mouse model of diabetes. This conclusion is based on the observations that 1) resistance artery TNF expression is elevated in HFD/STZ mice and 2) interfering with TNF activity (with an antagonist or genetic deletion) abolishes the augmented myogenic tone. Expanding on this fundamental mechanistic insight, targeted smooth muscle cell TNF gene deletion demonstrates that the relevant TNF source is localized within the vascular wall and acts by an autocrine/paracrine mechanism. Interestingly, we observe that ETN abolishes, rather than normalizes, the enhanced myogenic tone in vitro; yet, ETN normalizes myogenic tone when systemically administered. Pursuing the mechanistic basis for discrepancy is beyond the scope of the current study, although compromised vessel viability/general contractility can be excluded as the cause.

S1P is a key regulator of rodent resistance artery myogenic tone (11,41). We have previously demonstrated that TNF enhances S1P signaling in multiple microvascular beds and thus, augments myogenic responsiveness (9–12). Even in nonpathological settings, Sphk1 is a key source of S1P and a significant regulator of myogenic responsiveness (41): this prompted an expectation that Sphk1 gene deletion should abolish myogenic tone. Although myogenic responses are retained in arteries isolated from Sphk1^−/− mice (most likely through compensatory mechanisms involving Sphk2 [15]), the absence of augmented myogenic tone in HFD/STZ-treated Sphk1^−/− mice confirms the central role Sphk1 plays in the pathologic enhancement of myogenic responsiveness in diabetes.

Our gene deletion models identify two targets that can be therapeutically exploited: they predict that antagonizing either TNF or S1P in vivo will interrupt the pathological signaling chain and thereby correct the microvascular defect. Indeed, both strategies (TNF antagonism via ETN and S1P₂ receptor antagonism with JTE013) fully reverse the HFD/STZ-induced myogenic tone enhancement. This is the first evidence that systemically administered TNF and S1P signaling antagonists can correct microvascular dysfunction in the early stages of diabetes. Full preservation of PE responsiveness under these conditions demonstrates that the effects of diabetes are specific and restricted to the enhancement of myogenic tone: thus, capitalizing on therapeutic targets that correct the diabetes-associated microvascular dysfunction (i.e., interfering with TNF and/or S1P signaling) is unlikely to overtly compromise normal vascular function.

As a reported cause of insulin resistance (42) and diabetes complications (43), several small clinical trials have targeted TNF in diabetes (44–47). The vast majority of these investigations primarily focused on blood glucose control and insulin sensitivity as clinical end points (44–46); a few of these trials indirectly assessed vasodilator responses via forearm perfusion (46,47). Although anti-TNF therapy successfully lowers inflammatory markers (e.g., C-reactive protein), it largely fails to improve any other defined clinical end point. In the context of this study, we also observe that anti-TNF therapy fails to improve glucose control (Supplementary Table 1). The microvascular end points central to the present investigation, however, have not been systematically assessed in clinical trials: our data establish microvascular reactivity as an early disease marker and encourage its clinical examination.

Our study encourages future investigation on several issues. First, the TNF/S1P mechanism identified in mouse mesenteric/olfactory arteries must be translated to the human setting. The fact that the diabetes-induced myogenic tone augmentation associates with increased TNF expression in human skeletal muscle resistance arteries is encouraging; however, a causal relationship between TNF, S1P signaling, and myogenic responsiveness remains to be confirmed. Second, the current study has not determined the molecular basis for how diabetes increases microvascular TNF expression and how TNF enhances S1P signaling: further examination is warranted. Finally, although our data do not support a direct endothelial contribution to the myogenic tone enhancement (e.g., impaired vasodilator release), further assessments are required to conclusively exclude other forms of endothelial dysfunction that could indirectly contribute.

In conclusion, patients with diabetes have augmented skeletal muscle resistance artery myogenic tone, despite clinically acceptable glucose control. Our bed-to-bench approach endeavored to replicate this defect in a mouse model of diabetes. In our mouse model, both TNF signaling and S1P signaling augment myogenic tone; these pathways can be therapeutically targeted to specifically correct this defect. Our investigation adds diabetes to a diverse group of microvascular diseases influenced by pathological TNF/S1P signaling, which include heart failure, hearing loss, and stroke (9–12). These reactivity changes set the stage for more permanent effects (e.g., structural remodeling) and therefore potentially represent an untapped therapeutic target to prevent the cardiovascular complications associated with diabetes.

Acknowledgments. The authors thank Amy Cao (University of Toronto) for technical assistance with mouse husbandry, M. Hossein Noyan-Ashraf (Toronto General Research Institute) for technical assistance with the 10-month diet mice, and Eric Aki Shikatani (Toronto General Hospital Research Institute) for assistance with the analysis of the invasive hemodynamic data. The authors acknowledge the Spatio-Temporal Targeting and Amplification of Radiation Response (STTARR) program and its affiliated funding agencies.

Funding. M.S. received stipend support from a Canadian Institutes of Health Research (CIHR) doctoral research award (GSD-121812), the Banting and Best Diabetes Centre, the Natural Sciences and Engineering Research Council of Canada Collaborative Research and Training Experience Program (NSERC CREATE) in Microfluidic Applications and Training in Cardiovascular Health (MATCH), the Heart and Stroke/Richard Lewar Centre of Excellence for Cardiovascular Research, and an Ontario Graduate Scholarship. S.K.H. received stipend support from a CIHR doctoral research award in hypertension (DHY-121263), the Peterborough K.M. Hunter Foundation, and NSERC CREATE MATCH. The authors thank the institutions involved for the following funding: the Russian Science Foundation (14-50-00060 to S.A.N.), the CIHR (FRN-119345 to S.-S.B.), the Canada Foundation for Innovation and Ontario Research Fund (RI-11810 to S.-S.B.), the Heart and Stroke Foundation of Ontario (HSFO) (G13-0002813 to S.-S.B.), HSFO New (NIA-6581 to S.-S.B.), and Career (CIA-7432 to S.-S.B.) Investigator Awards and start-up funding from the University of Toronto (to S.-S.B.).

Duality of Interest. No potential conflicts of interest relevant to this article were reported.

Author Contributions. M.S. and S.-S.B. conceived of and designed the experiments. M.S., S.K.H., D.D.D., W.D.F., A.M., J.T.K., and D.L. performed the experiments. M.S., S.K.H., D.D.D., W.D.F., A.M., J.T.K., D.L., and S.-S.B. analyzed data. M.S., D.L., and S.-S.B. wrote the manuscript. W.D.F., S.A.N., S.O., M.H., and S.-S.B. contributed reagents/materials/analysis tools. S.-S.B. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.