Expression of TNF-α, a pleiotropic cytokine, is elevated during stroke and cerebral ischemia. TNF-α regulates arterial diameter, although mechanisms mediating this effect are unclear. In the present study, we tested the hypothesis that TNF-α regulates the diameter of resistance-sized (∼150-μm diameter) cerebral arteries by modulating local and global intracellular Ca2+ signals in smooth muscle cells. Laser-scanning confocal imaging revealed that TNF-α increased Ca2+ spark and Ca2+ wave frequency but reduced global intracellular Ca2+ concentration ([Ca2+]i) in smooth muscle cells of intact arteries. TNF-α elevated reactive oxygen species (ROS) in smooth muscle cells of intact arteries, and this increase was prevented by apocynin or diphenyleneiodonium (DPI), both of which are NAD(P)H oxidase blockers, but was unaffected by inhibitors of other ROS-generating enzymes. In voltage-clamped (−40 mV) cells, TNF-α increased the frequency and amplitude of Ca2+ spark-induced, large-conductance, Ca2+-activated K+ (KCa) channel transients ∼1.7- and ∼1.4-fold, respectively. TNF-α-induced transient KCa current activation was reversed by apocynin or by Mn(III)tetrakis(1-methyl-4-pyridyl)porphyrin (MnTMPyP), a membrane-permeant antioxidant, and was prevented by intracellular dialysis of catalase. TNF-α induced reversible and similar amplitude dilations in either endothelium-intact or endothelium-denuded pressurized (60 mmHg) cerebral arteries. MnTMPyP, thapsigargin, a sarcoplasmic reticulum Ca2+-ATPase blocker that inhibits Ca2+ sparks, and iberiotoxin, a KCa channel blocker, reduced TNF-α-induced vasodilations to between 15 and 33% of control. In summary, our data indicate that TNF-α activates NAD(P)H oxidase, resulting in an increase in intracellular H2O2 that stimulates Ca2+ sparks and transient KCa currents, leading to a reduction in global [Ca2+]i, and vasodilation.
- cerebrovascular circulation
- ryanodine-sensitive Ca2+ release channel
- Ca2+-activated K+ channel
- reactive oxygen species
stroke and transient brain ischemia are vasculature-borne brain pathologies accompanied by inflammatory changes in blood vessels and parenchymal tissues (15, 46). TNF-α, a pleiotropic cytokine, is a proinflammatory agent generated by glial cells, neurons, macrophages, and vascular endothelium during brain injury. Clinical data demonstrate a correlation between TNF-α levels and the severity of ischemic brain damage, but controversy exists regarding the functional role of TNF-α, with both beneficial and detrimental roles having been proposed (3, 8, 9, 15, 36, 46). Chronic TNF-α elevation is associated with risk factors for other vascular pathologies, including coronary artery disease and atherosclerosis (26). During stroke, TNF-α may cause neuronal damage directly and by accelerating ischemia-induced local inflammatory responses. In contrast, studies also indicate that TNF-α may be cytoprotective (8, 15, 36, 46).
TNF-α acts in a paracrine manner by binding to two membrane receptors, p55 and p75. Besides its actions on neurons, astrocytes, and microglia, TNF-α also regulates arterial contractility (15), an effect that may modulate injury by altering local blood supply. However, the regulation of arterial diameter and blood flow by TNF-α is complex. Intracranial injection of TNF-α in vivo constricted pial arterioles and reduced cerebral blood flow (29, 38, 44). TNF-α also induced procontractile effects in coronary arteries (23, 51). In contrast, TNF-α dilated cerebral, mesenteric, and cremaster muscle arterioles and relaxed endothelium-denuded aortas (4, 7, 16, 22, 32, 37). In bronchial arteries, TNF-α initially induced dilation, followed by constriction 2 h later (45). TNF-α-mediated vasoregulation can also occur through both endothelium-dependent and endothelium-independent mechanisms (4, 7, 16, 22, 23, 29, 32, 37, 38, 44, 45, 51). Collectively, data suggest that TNF-α can induce both vasoconstriction and vasodilation and that its effects may be time dependent. Regardless of these data, the mechanisms that mediate vasoregulation by TNF-α are poorly understood.
A major regulator of arterial diameter is the smooth muscle cell global intracellular Ca2+ concentration ([Ca2+]i). In smooth muscle cells, global [Ca2+]i is regulated by extracellular Ca2+ influx and intracellular Ca2+ release (19). An elevation in global [Ca2+]i induces constriction, whereas a decrease in global [Ca2+]i leads to vasodilation. Another Ca2+ signal, termed a Ca2+ spark, is a rapid, localized intracellular Ca2+ transient that occurs as a result of the activation of several ryanodine-sensitive Ca2+ release (RyR) channels on the sarcoplasmic reticulum (SR) (19, 33). In arterial smooth muscle cells, Ca2+ sparks occur close to the plasma membrane and activate several large-conductance Ca2+-activated K+ (KCa) channels, resulting in a transient KCa current. The resulting membrane hyperpolarization reduces voltage-dependent Ca2+ channel activity, leading to a decrease in Ca2+ entry, a reduction in global [Ca2+]i and vasodilation (19, 33). In contrast, Ca2+ spark inhibition leads to vasoconstriction due to depolarization-induced voltage-dependent Ca2+ channel activation (19). Alterations in Ca2+ sparks and effective coupling to KCa channels have been implicated in hypertension and diabetes (1, 6, 19, 31). Another intracellular Ca2+ signal, termed a Ca2+ wave, is a propagating [Ca2+]i elevation that occurs as a result of the activation of SR RyR channels and/or inositol 1,4,5-trisphosphate-gated Ca2+ channels. Although the physiological functions of Ca2+ waves have been less well described, Ca2+ waves, when stimulated, contribute Ca2+ for contraction (35).
The goal of the present study was to investigate whether TNF-α regulates cerebral artery diameter by modulating smooth muscle Ca2+ signaling and, if so, to determine the underlying mechanisms. Such information would improve the understanding of cytokine-mediated changes in arterial diameter during vascular pathologies, including stroke.
Tissue Preparation and Single-Cell Isolation
Sprague-Dawley rats (∼250 g) of either sex were euthanized by administering an intraperitoneal injection of a pentobarbital sodium overdose (150 mg/kg body wt) in accordance with the policies and procedures of the Animal Care and Use Committee at the University of Tennessee. The brains were removed and placed into ice-cold (4°C) oxygenated (95% O2-5% CO2) physiological salt solution (PSS) containing (in mM) 119 NaCl, 4.7 KCl, 24 NaHCO3, 1.2 KH2PO4, 1.6 CaCl2, 1.2 MgSO4, 0.023 EDTA, and 11 glucose. The posterior cerebral, middle cerebral, and cerebellar arteries (50–200 μm in diameter) were removed, cleaned of connective tissue, and maintained in ice-cold PSS. When appropriate, the endothelium was removed by introducing an air bubble into the arterial lumen for 2 min, followed by a 30-s wash with H2O. Individual smooth muscle cells were dissociated from cerebral arteries using enzymes and an isolation solution composed of (in mM) 55 NaCl, 80 sodium glutamate, 5.6 KCl, 2 MgCl2, 10 HEPES, and 10 glucose (pH 7.3 with NaOH) as previously described (18). Cells were stored on ice and used for experiments between 1 and 6 h after isolation.
Laser-Scanning Confocal Ca2+ Imaging
Artery segments (1–2 mm in length) were placed into HEPES-buffered PSS composed of (in mM) 134 NaCl, 6 KCl, 2 CaCl2, 1 MgCl2, 10 HEPES, and 10 glucose (pH 7.4, NaOH) containing fluo-4 AM (10 μM) and Pluronic F-127 (0.05%) for 60 min at 22°C. To allow for indicator deesterification, the arteries were subsequently placed into HEPES-buffered PSS for 30 min at 22°C. Smooth muscle cells within the arterial wall were scanned using an OZ laser-scanning confocal microscope (Noran Instruments, Middleton, WI) and a ×60 magnification water-immersion lens objective (numerical aperture 1.2) attached to a TE300 microscope (Nikon). Cells were illuminated using a krypton-argon laser at 488 nm, and emitted light >500 nm was collected. Planar images (56.3 × 52.8 μm) were recorded every 16.7 ms (i.e., 60 images s−1). To compare laser-scanning confocal Ca2+ imaging data with electrophysiological recordings obtained in this study, Ca2+ sparks in smooth muscle cells of arterial segments were measured in an extracellular solution containing 30 mM K+, which depolarizes smooth muscle cells to approximately −40 mV. This membrane potential is similar to that of cerebral arteries pressurized to 60 mmHg (see Ref. 20 for similar procedure). The 30 mM K+ bath solution contained (in mM) 110 NaCl, 30 KCl, 10 HEPES, 2 CaCl2, 1 MgCl2, and 10 glucose (pH 7.4 maintained using NaOH). At least two different representative areas of each artery were scanned for at least 10 s for each condition. The same area of each artery was scanned only once to avoid any laser-induced changes in Ca2+ signaling, and the effects of drugs were measured in paired experiments. Ca2+ sparks were detected in smooth muscle cells using custom analysis software written in IDL 5.2 (Research Systems, Boulder, CO), kindly provided by Drs. M. T. Nelson and A. D. Bonev (University of Vermont, Burlington, VT). Automated and manual detection of Ca2+ sparks were performed by dividing an area of 1.54 μm (7 pixels) × 1.54 μm (7 pixels) (i.e., 2.37 μm2) in each image (F) by a baseline level (F0) that was determined by averaging 10 images without Ca2+ spark activity. The entire area of each image was analyzed to detect Ca2+ sparks. A Ca2+ spark was defined as a localized increase in F/F0 >1.2 (10). Arterial Ca2+ spark frequency (measured in Hz) was calculated by averaging values from at least two different areas. Ca2+ waves were defined as an elevation in F/F0 >1.2 that propagated for at least 20 μm. Global Ca2+ fluorescence was calculated from the same images used for Ca2+ spark analysis and was the mean pixel value of 100 different images acquired during a 10-s period. To calculate whether TNF-α altered global [Ca2+]i, global Ca2+ fluorescence in TNF-α was divided by the corresponding control value.
K+ currents were measured using either the conventional whole cell or perforated configuration of the patch-clamp technique with an Axopatch 200B amplifier (Axon Instruments, Union City, CA). Bath solution was 6 mM K+ HEPES-buffered PSS (composition described above). For perforated patch-clamp experiments, the pipette solution contained (in mM) 110 potassium aspartate, 30 KCl, 10 NaCl, 1 MgCl2, 10 HEPES, and 0.05 EGTA (pH 7.2 maintained using KOH). For conventional whole cell experiments, the pipette solution contained (in mM) 140 KCl, 1.9 MgCl2, 0.037 CaCl2, 0.1 EGTA, 10 HEPES, and 2 Na2ATP (pH 7.2 maintained using KOH); the calculated free Ca2+ and free Mg2+ concentrations of this solution were 100 nM and 1 mM, respectively (WEBMAXC; Stanford University, Palo Alto, CA). In some experiments, 200 U/ml catalase was included in the whole cell pipette solution. When appropriate, catalase was heat inactivated by incubation at 92°C for 25 min. TNF-α was applied to the bath solution 10 min after formation of the whole cell configuration to ensure exchange of the pipette solution with the cytosol. All experiments were performed using a holding potential of −40 mV. Membrane currents were recorded using a sample rate of 2.5 kHz and filtered at 1 kHz. Transient KCa current analysis was performed offline. A transient KCa current was defined as the simultaneous opening of three or more KCa channels.
Reactive Oxygen Species Measurements
Reactive oxygen species (ROS) were measured in isolated intact arterial segments using 2′,7′-dichlorodihydrofluorescein diacetate (H2DCFDA). Isolated endothelium-denuded cerebral artery segments (1–2 mm) were incubated for 45 min at room temperature in 10 μM H2DCFDA. H2DCFDA is hydrolyzed by intracellular esterases to the nonfluorescent derivative dichlorodihydrofluorescein (H2DCF). ROS oxidize H2DCF to fluorescent dichlorofluorescein (DCF). DCF fluorescence was excited with 488 nm of light, and emitted light >525 nm was collected. Planar images were acquired using a Zeiss LSM5 laser-scanning confocal microscope. Images were recorded 10 min after TNF-α or H2O2 application. Time controls were also performed in which agents were not applied. Because DCF fluorescence increases spontaneously upon exposure to excitation light (12), arterial regions were scanned only once. Images were background corrected. ImageJ software (National Institutes of Health, Bethesda, MD) was used for analysis.
Cerebral arterial segments were incubated for 20 min in HEPES-buffered PSS without (time-matched control) or with TNF-α. Arteries were then embedded in OTC and snap-frozen in liquid N2. Transverse sections 30 μm thick were cut, mounted onto glass slides, and incubated with 10 μM dihydroethidium (DHE) in a 100% N2 atmosphere in the dark for 30 min at 37°C. Arterial sections were excited with 488 nm light, and emitted light >590 nm was collected using an OZ laser-scanning confocal microscope. Custom-written software was used to measure DHE fluorescence intensity. Fluorescence intensity in TNF-α was divided by that in time-matched controls to determine the effects of TNF-α on DHE intensity. Tissue processing did not elevate ROS, because the DHE fluorescence intensity of arteries preincubated with Mn(III)tetrakis(1-methyl-4-pyridyl)porphyrin (MnTMPyP) was 1.04 ± 0.06-fold of those that were left untreated (n = 5; P > 0.05).
Pressurized Artery Diameter Measurements
Cerebral arterial segments (100–200 μm diameter) were placed in a temperature-controlled perfusion chamber (Living Systems Instrumentation, Burlington, VT) and cannulated with glass pipettes at each end. The chamber was perfused continuously with PSS equilibrated with 5% CO2, 21% O2, and 74% N2, pH 7.4, and maintained at 37°C. Arteries were observed using a charge-coupled device camera attached to an inverted microscope (TS100F; Nikon). Arterial diameter was monitored using edge detection software (IonWizard; IonOptix, Milton, MA). Arterial intraluminal pressure was elevated to 60 mmHg, which resulted in the development of myogenic tone. After diameter stabilization, tested agents were applied using chamber perfusion. Arterial diameter was digitized at 1 Hz using the edge detection function of IonWizard software. Endothelial denudation was confirmed by the absence of a dilatory response to carbachol (10−5 M), an endothelium-dependent cerebral vasodilator. At the end of each experiment, passive arterial diameter was determined using a Ca2+-free bath solution containing 1 mM EGTA.
Data are expressed as means ± SE. Student's t-test was used to compare paired and unpaired data from two populations, and ANOVA and the Student-Newman-Keuls test were used for multiple-group comparisons. Evaluation of whether distributions were Gaussian using the method of Kolmogorov and Smirnov. P < 0.05 was considered significant.
Unless stated otherwise, all chemicals used in this study were obtained from Sigma (St. Louis, MO) or Calbiochem (San Diego, CA). Papain was purchased from Worthington Biochemical (Lakewood, NJ), fluo-4 AM and H2DCFDA were obtained from Molecular Probes (Eugene, OR), and rat recombinant TNF-α was purchased from BD Pharmingen (San Diego, CA).
TNF-α Activates Ca2+ Sparks and Ca2+ Waves and Reduces Global [Ca2+]i in Smooth Muscle Cells of Intact Cerebral Arteries
The regulation of local and global Ca2+ signals in smooth muscle cells by TNF-α was measured in cerebral artery segments. TNF-α (50 ng/ml) increased mean Ca2+ spark frequency from 1.53 ± 0.16 Hz to 2.6 ± 0.51 Hz, or ∼1.7-fold (Fig. 1, A and B). TNF-α did not change Ca2+ spark amplitude (control F/F0, 1.36 ± 0.01, n = 184 sparks, and TNF-α F/F0, 1.34 ± 0.01, n = 329 sparks; P > 0.05). In the same cells, TNF-α also elevated mean Ca2+ wave frequency from 0.32 ± 0.02 Hz to 0.48 ± 0.03 Hz, or ∼1.5-fold (Fig. 1C). In contrast, TNF-α decreased mean global [Ca2+]i to 0.85 ± 0.02 of control (n = 6 arteries, P < 0.05). These experiments suggest TNF-α elevates Ca2+ spark and wave frequency and reduces global [Ca2+]i in cerebral arterial smooth muscle cells.
TNF-α Activates NAD(P)H Oxidase Leading to a ROS Elevation in Smooth Muscle Cells of Intact Cerebral Arteries
TNF-α activates ROS generation in smooth muscle cells (13). Because RyR channels are redox sensitive (42), we tested the hypothesis that TNF-α regulates local Ca2+ signaling by elevating intracellular ROS. Several enzymes generate ROS including NAD(P)H oxidase, xanthine oxidase, cytochrome P450, and mitochondrial electron transport chain complexes (13, 28, 41, 49). The DCF and DHE fluorescence methods were used to determine the pathways through which TNF-α elevates ROS in the smooth muscle cells of small cerebral arteries. TNF-α increased DCF fluorescence intensity 2.7 ± 0.32-fold and DHE intensity 1.51 ± 0.03-fold in the smooth muscle cells of intact arteries (Fig. 2, A–D). During the same period, H2O2 elevated DCF fluorescence 4.7 ± 0.68-fold, whereas DCF fluorescence did not change in controls (absence of applied agents) (Fig. 2B). In the presence of apocynin or diphenyleneiodonium (DPI), which are NAD(P)H oxidase blockers, TNF-α did not elevate DCF fluorescence (Fig. 2B). MnTMPyP, a membrane-permeant O2−· and catalase mimetic, also prevented the TNF-α-induced DCF fluorescence elevation (Fig. 2B). Apocynin (1.07 ± 0.23-fold vs. control; n = 5), DPI (0.85 ± 0.1-fold vs. control; n = 4), and MnTMPyP (1.14 ± 0.17-fold vs. control; n = 4) did not change ROS when applied alone (P > 0.05 for each). TNF-α-induced DCF fluorescence elevations were not reduced by oxypurinol, a xanthine oxidase inhibitor; 17-octadecanoic acid, a cytochrome P450 blocker; or lonidamine, a mitochondrial permeability transition pore (PTP) opener (Fig. 2B). Oxypurinol (0.98 ± 0.13-fold vs. control; n = 4), 17-octadecanoic acid (1.09 ± 0.12-fold vs. control; n = 5), and lonidamine (0.86 ± 0.13-fold vs. control; n = 4) did not change DCF fluorescence when applied alone (P > 0.05 for each). These data indicate that TNF-α activates NAD(P)H oxidase, leading to ROS elevation in cerebral arterial smooth muscle cells.
TNF-α Activates Transient KCa Currents in Isolated Arterial Smooth Muscle Cells
In arterial smooth muscle cells, a Ca2+ spark activates several KCa channels, resulting in a transient KCa current that induces membrane hyperpolarization (19). To investigate the effects of TNF-α-induced changes in Ca2+ signaling, transient KCa current regulation by TNF-α was measured in isolated, voltage-clamped (−40 mV) arterial smooth muscle cells. When using the perforated patch-clamp configuration, TNF-α evoked a gradual increase in transient KCa current frequency and amplitude that reached a steady-state elevation of ∼1.7- and ∼1.4-fold, respectively, in ∼10 min (Fig. 3, A, C, and D). Similar observations were made when using the conventional whole cell configuration (Fig. 3, B–D). Apocynin did not alter transient KCa current frequency (0.89 ± 0.12-fold vs. control; n = 6) or amplitude (1.0 ± 0.06-fold vs. control; n = 6) when applied alone (P > 0.05 for each) but reversed the TNF-α-induced increase in transient KCa current frequency and amplitude (Fig. 3, B–D). These data suggest that TNF-α stimulates transient KCa currents by activating NAD(P)H oxidase.
TNF-α-induced intracellular signaling may involve mitochondrial PTP activation and ceramide generation (14, 22, 28, 30, 39, 41, 52). However, ceramide did not alter transient KCa currents, indicating that it is unlikely to mediate TNF-α-induced activation of these events (Fig. 3, C and D). Lonidamine decreased mean Ca2+ spark frequency from 1.45 ± 0.16 Hz to 0.73 ± 0.18 Hz, or ∼0.5-fold compared with control (n = 4; P < 0.05) and inhibited transient KCa currents (10). Similarly, atractyloside, another mitochondrial PTP opener, decreased transient KCa current frequency and amplitude (Fig. 3, C and D). These data demonstrate that PTP openers inhibit transient KCa currents, indicating that such an effect cannot mediate TNF-α-induced transient KCa current activation.
To determine whether TNF-α-induced transient KCa current activation occurs as a result of a ROS elevation, we measured transient KCa current regulation by MnTMPyP as well as by catalase that converts H2O2 to H2O. MnTMPyP reversed TNF-α-induced elevations in transient KCa current frequency and amplitude (Fig. 4, A–C). In contrast, MnTMPyP did not change transient KCa current frequency (1.0 ± 0.1-fold vs. control; n = 5) or amplitude (1.0 ± 0.07-fold vs. control) when applied alone (P > 0.05 for each). Similarly, when the conventional whole cell configuration was used, inclusion of catalase in the pipette solution (to allow intracellular access) prevented TNF-α-induced transient KCa current activation (Fig. 4, B and C). In contrast, when the pipette solution was supplemented with heat-inactivated catalase, TNF-α increased transient KCa current frequency and amplitude ∼1.8- and ∼1.4-fold, respectively (Fig. 4, B and C). In summary, these data indicate that TNF-α activates Ca2+ sparks and transient KCa currents by inducing an NAD(P)H oxidase-mediated ROS elevation.
TNF-α Dilates Pressurized Cerebral Arteries by Elevating ROS, SR Ca2+-ATPase Release, and KCa Channel Activity
TNF-α has been demonstrated to induce both vasoconstriction and vasodilation by endothelium-dependent and endothelium-independent pathways (4, 7, 16, 22, 23, 29, 32, 37, 38, 44, 51). The data reported herein suggest that TNF-α activates a dilatory Ca2+ signaling pathway in the smooth muscle cells of small, resistance-sized cerebral arteries. Therefore, vasoregulation by TNF-α was investigated using resistance-sized, pressurized (60 mmHg) cerebral arteries. In endothelium-intact arteries, from a mean active diameter of 142 ± 3 μm (0.64 ± 0.01-fold vs. passive diameter), TNF-α induced a mean dilation of 22 ± 1 μm (Fig. 5, A–C). In endothelium-denuded arteries, TNF-α increased mean diameter by 18 ± 3 μm from 130 ± 9 μm (0.57 ± 0.02-fold vs. passive diameter) (Fig. 5C). TNF-α-induced dilations were not significantly different in endothelium-intact or endothelium-denuded arteries (P > 0.05). TNF-α-induced dilations were fully reversible (Fig. 5A), and consecutive TNF-α applications resulted in dilations of similar magnitude (first application, 19 ± 2 μm; second application, 21 ± 2 μm) (n = 2).
To investigate the importance of ROS generation in TNF-α-induced vasodilation, responses to TNF-α were measured in control and then in the presence of MnTMPyP (10 μM) in the same artery. MnTMPyP reduced mean TNF-α-induced vasodilation to 7 ± 2 μm or to 0.34 ± 0.03 of control dilations (Fig. 5C). The same protocol was performed with thapsigargin (100 nM), a SR Ca2+-ATPase blocker that inhibits Ca2+ sparks, or iberiotoxin (100 nM), a selective KCa channel blocker (19). Thapsigargin reduced mean TNF-α-induced vasodilation to 4 ± 2 μm, or to 0.15 ± 0.01 of those obtained in controls in the same arteries (Fig. 5C). Similarly, iberiotoxin reduced mean TNF-α-induced vasodilation to 6 ± 1 μm, or to 0.29 ± 0.05 of control (Fig. 5, B and C). These data indicate that TNF-α dilates cerebral arteries by generating ROS that stimulate Ca2+ sparks and KCa channels in smooth muscle cells.
In the present study, we investigated the mechanisms by which TNF-α, a pleiotropic cytokine, regulates the diameter of small cerebral arteries. Our data indicate that TNF-α activates smooth muscle cell NAD(P)H oxidase, leading to the generation of ROS that activate Ca2+ sparks and thus transient KCa currents. The ensuing decrease in global [Ca2+]i leads to vasodilation. These findings show for the first time that an inflammatory mediator that is central to a variety of vascular diseases induces vasodilation through Ca2+ spark activation.
TNF-α is produced in the brain during ischemia and is implicated in the etiology of several cardiovascular diseases, including stroke, cardiac failure, coronary artery disease, and atherosclerosis (15, 46). TNF-α is reported to induce both vasoconstriction and vasodilation. Topical application of TNF-α dilated cerebral arterioles (7, 37), whereas intracranial injection of TNF-α induced cerebrovascular constriction and reduced blood flow (29, 38, 44). TNF-α also dilated mesenteric and cremaster muscle arterioles (4, 32, 45). Earlier studies suggested that TNF-α promotes vasodilation by elevating nitric oxide (NO) and prostaglandin production in the arterial wall and in perivascular tissues (4, 7, 37). In contrast, in pulmonary arterial rings, a hypoxia-induced increase in TNF-α mRNA was associated with contraction (43). TNF-α also reduced bradykinin- and ACh-induced vasodilation in rat mesenteric arteries (48) and bradykinin- and A23187-induced vasodilation in bovine coronary arteries (51), blunted ACh-induced vasodilation in cat carotid arteries (2), and evoked endothelium-dependent vasoconstriction in human and rat coronary arteries (17, 23). In humans, intrabrachial TNF-α injection impaired bradykinin- and ACh-induced vasodilation (11). Data also suggest that TNF-α inhibits vasodilation by triggering ROS that scavenge endothelial NO (51). It is unclear what underlies differential regulation of arterial contractility by TNF-α, but several possibilities exist. Vasoregulatory actions of TNF-α may be vascular bed specific. Different experimental protocols used may also explain the diversity of observations reported in various studies. In addition, the effects of TNF-α on arterial contractility may be time dependent. For example, in sheep bronchial arteries, TNF-α initially induced dilation, followed by constriction 2 h later (45). Data also suggest that differential effects on contractility may occur when TNF-α acts primarily on either endothelial or smooth muscle cells. Conceivably, TNF-α-induced vasoconstriction might occur primarily as a result of an endothelium-dependent mechanism, whereas vasodilation may be mediated by a direct effect on smooth muscle cells. Here, we have shown that in isolated, pressurized, myogenic rat cerebral arteries, acute TNF-α application (<30 min) induced vasodilation that was endothelium independent and was markedly attenuated by an antioxidant and inhibitors of SR Ca2+ release and KCa channels.
TNF-α activates several signaling pathways in vascular smooth muscle cells. TNF-α stimulated NAD(P)H oxidase, leading to ROS production in rat aortic smooth muscle cells (13). TNF-α relaxed endothelium-denuded, phenylephrine-constricted aortic rings via PLA2 activation, implicating ceramide as a mediator (22). In cardiac and endothelial cells, signaling mechanisms for TNF-α involve mitochondria (20, 23–25, 35, 36). TNF-α disrupted the mitochondrial electron transport chain, caused release of cytochrome c, induced mitochondrial ROS generation, and triggered apoptosis (14, 28, 30, 41, 52). Although the downstream intramitochondrial target for TNF-α is unclear, PTP involvement was proposed in endothelial cells (52). PTP is also a downstream target for TNF-α-induced signaling in other cell types, and PTP opening may be triggered by electron transport chain inhibition and mitochondrial depolarization (5, 10, 39). Ceramide had no effect on transient KCa currents, and PTP openers inhibited these events, consistent with the findings reported in a previous study of arterial smooth muscle cells (10). Therefore, TNF-α is unlikely to activate Ca2+ sparks and transient KCa currents by elevating ceramide or by inducing PTP opening. In some cell types, including neutrophils, preincubation with apocynin is required to block association of NAD(P)H oxidase subunits (40). Here, we have shown that in cerebral artery smooth muscle cells, apocynin reverses TNF-α-induced transient KCa current activation within ∼10 min. This effect may reflect rapid subunit turnover in the presence of TNF-α in these cells. Our present data indicate that TNF-α stimulates NAD(P)H oxidase, generating ROS that activate Ca2+ sparks and transient KCa currents in cerebral artery smooth muscle cells. These findings are consistent with those described in a recent study that reported mitochondria-derived ROS activate Ca2+ sparks and transient KCa currents in arterial smooth muscle cells (50). Thus, ROS derived from different sources induce vasodilation through a common pathway, namely, Ca2+ spark activation.
TNF-α-induced ROS may activate Ca2+ sparks and transient KCa currents as a result of a direct or indirect effect on RyR and KCa channels. Cardiac and skeletal muscle RyR channels are redox sensitive and are activated by oxidizing agents, including H2O2 (42). ROS also activate a number of other signal transduction pathways that may regulate RyR channel activity (42, 49). Although O2−· is the initial product of NAD(P)H oxidase, O2−· has a short half-life and rapidly dismutates to H2O2, a relatively stable oxidant. O2−· is unlikely to mediate the effects of TNF-α on Ca2+ sparks, particularly because catalase blocked TNF-α-induced transient KCa current activation. Data suggest that H2O2 formed from NAD(P)H oxidase-generated O2−· activates RyR channels, leading to a Ca2+ spark frequency elevation. Indeed, exogenous H2O2 activates transient KCa currents in cerebral artery smooth muscle cells (10). Another potential ROS mediating Ca2+ spark activation in this study is HO·, a highly reactive species produced in a transition metal-catalyzed reaction (the Fenton reaction) between H2O2 and O2−·. In conventional whole cell experiments, we used a pipette solution absent in transition metals, and dialysis of the pipette solution within the cell interior should have washed out endogenous cytosolic transition metals. Thus HO· generation in these experiments should have been minimal, arguing against a significant role for HO· in TNF-α-induced Ca2+ spark and KCa channel activation.
ROS production or inhibition of ROS degradation leads to vasoconstriction, high blood pressure, and vascular hypertrophy (24). However, ROS also stimulate vasodilation. For example, flow-mediated vasodilation occurs because of the generation of mitochondria-derived H2O2 (27). Furthermore, topical application of ROS dilates cerebral arteries in vivo (47). Thus ROS induce both vasoconstriction and vasodilation. Conceivably, there might be a delicate balance between ROS concentrations that regulate physiological functions and higher ROS concentrations that lead to pathologies (24). The data reported herein indicate that TNF-α regulates Ca2+ sparks by generating ROS from a single source, NAD(P)H oxidase. During disease, vasoconstriction may result from ROS produced in higher concentrations, from additional enzymes, from exogenous sources, and for prolonged periods. For example, oxyhemoglobin, which may contribute to vasoconstriction after subarachnoid hemorrhage, inhibits Ca2+ sparks in cerebral artery smooth muscle cells, and this effect can be blocked by antioxidants (21). TNF-α receptors were recently proposed to confine NAD(P)H oxidase-mediated redox signaling spatially in human microvascular endothelial cells (25). TNF-α may also activate Ca2+ sparks by elevating ROS in the local vicinity of Ca2+ spark sites. Through restriction, the global effects of ROS could be lessened and dilatory signaling could be maintained. Because Ca2+ spark sites are close to sarcolemmal KCa channels, local NAD(P)H oxidase-derived ROS should also regulate KCa channel activity. TNF-α did not alter Ca2+ spark amplitude but increased transient KCa current amplitude. NAD(P)H oxidase inhibitors and antioxidants reduced the TNF-α-induced transient KCa current amplitude elevation, indicating that the effect was mediated by NAD(P)H oxidase-derived ROS. Thus TNF-α also enhances the effective coupling of Ca2+ sparks to KCa channels, presumably by elevating KCa channel Ca2+ sensitivity within the micromolar concentration range generated by these Ca2+ transients (34). The combined effects of elevated Ca2+ spark frequency and transient KCa current amplitude could significantly increase KCa channel activity, leading to vasodilation (19, 33).
TNF-α elevated Ca2+ wave frequency ∼1.5-fold but reduced global [Ca2+]i. In the absence of receptor agonists, Ca2+ wave frequency in arterial smooth muscle cells is low, and Ca2+ waves contribute little to the global [Ca2+]i (18). When stimulated to high frequencies, Ca2+ waves can contribute to the global [Ca2+]i and thus induce smooth muscle contraction (35). Although TNF-α stimulated Ca2+ waves, the frequency elevation was insufficient to contribute significantly to the global [Ca2+]i. In contrast, TNF-α-induced Ca2+ spark activation significantly elevated KCa channel activity. The resulting membrane hyperpolarization would reduce voltage-dependent Ca2+ channel activity, leading to the observed decrease in global [Ca2+]i (19). Therefore, the net effect of TNF-α-induced Ca2+ spark and Ca2+ wave activation is a reduction in the global [Ca2+]i, leading to vasodilation.
In summary, in this study, we have defined a novel vasodilatory signaling pathway for an inflammatory cytokine mediated by Ca2+ spark and KCa channel activation in smooth muscle cells. After ischemia and stroke, TNF-α-induced Ca2+ spark activation and cerebrovascular dilation would be important in regulating local blood supply in the affected brain region.
This project was supported by grants from the National Institutes of Health and the American Heart Association National Center (to J. H. Jaggar). S. Y. Cheranov is a recipient of a postdoctoral fellowship from the American Heart Association-Southeast Affiliate.
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