Hif1α-dependent mitochondrial acute O2 sensing and signaling to myocyte Ca2+ channels mediate arterial hypoxic vasodilation

Moreno-Domínguez, Alejandro; Colinas, Olalla; Arias-Mayenco, Ignacio; Cabeza, José M.; López-Ogayar, Juan L.; Chandel, Navdeep S.; Weissmann, Norbert; Sommer, Natascha; Pascual, Alberto; López-Barneo, José

doi:10.1038/s41467-024-51023-3

Hif1α-dependent mitochondrial acute O₂ sensing and signaling to myocyte Ca²⁺ channels mediate arterial hypoxic vasodilation

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Published: 05 August 2024

Volume 15, article number 6649, (2024)
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Hif1α-dependent mitochondrial acute O₂ sensing and signaling to myocyte Ca²⁺ channels mediate arterial hypoxic vasodilation

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Abstract

Vasodilation in response to low oxygen (O₂) tension (hypoxic vasodilation) is an essential homeostatic response of systemic arteries that facilitates O₂ supply to tissues according to demand. However, how blood vessels react to O₂ deficiency is not well understood. A common belief is that arterial myocytes are O₂-sensitive. Supporting this concept, it has been shown that the activity of myocyte L-type Ca²⁺channels, the main ion channels responsible for vascular contractility, is reversibly inhibited by hypoxia, although the underlying molecular mechanisms have remained elusive. Here, we show that genetic or pharmacological disruption of mitochondrial electron transport selectively abolishes O₂ modulation of Ca²⁺ channels and hypoxic vasodilation. Mitochondria function as O₂ sensors and effectors that signal myocyte Ca²⁺ channels due to constitutive Hif1α-mediated expression of specific electron transport subunit isoforms. These findings reveal the acute O₂-sensing mechanisms of vascular cells and may guide new developments in vascular pharmacology.

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Introduction

It has been known for over a century that oxygen (O₂) is a major regulator of blood flow^1,2,3,4,5,6; (see^7,8 for reviews and references). O₂ is a potent vasoconstrictor^6,9,10,11,12 and in most systemic arterial territories, particularly in skeletal muscle, coronary, and cerebral vessels, a decrease in O₂ tension (hypoxia) leads to rapid vasodilation^{8,13,14,15,16,17}. In this manner, O₂ supply to tissues (e.g., the working heart) is swiftly adapted to demand. This essential vasomotor response also contributes to neurovascular coupling, a dilation of small blood vessels whereby O₂ availability is increased in the most active brain regions^18,19,20. Details of the cellular mechanisms governing hypoxic vasodilation (HVD) remain controversial and the underlying molecular processes are largely unknown^7,8,21.

The vascular endothelium and neighboring tissues can release relaxing factors that act on smooth muscle^{8,22,23,24,25,26,27,28,29}. In addition, hemoglobin desaturation and release of vasodilators from red blood cells has been proposed to induce increases in blood flow in hypoxic capillaries^30,31. However, numerous studies have suggested that myocytes are a primary site for vascular acute O₂ sensing^8,21,32. HVD has been recorded in arteries cleaned of surrounding tissue, denuded of endothelium, and bathed in cell-free external solutions equilibrated with low O₂ tension (PO₂)^33,34,35. Moreover, hypoxic relaxation is observed in arteries depolarized and precontracted with high extracellular K⁺^14,35,36,37, indicating that PO₂ directly modulates transmembrane Ca²⁺ influx in vascular smooth muscle (VSM)^36,38. In support of this concept, hypoxia, within physiological values, has been shown to elicit fast, and highly reversible inhibition of L-type voltage-gated Ca²⁺channels (the main ion channel type responsible for vascular contractility) in myocytes from several systemic rodent, porcine, and human arterial models^{9,35,37,39,40,41}. However, the molecular mechanism underpinning Ca²⁺ channel regulation by PO₂ has been elusive⁴². Here, we report that a constitutively expressed hypoxia inducible factor 1α (Hif1α)-dependent mitochondrial acute O₂ sensing and signaling pathway, in systemic arterial myocytes, mediates the hypoxia-induced inhibition of Ca²⁺ channel activity and HVD.

Results

Disruption of mitochondrial complex I in arterial myocytes abolishes acute O₂ modulation of Ca²⁺ channel activity and hypoxic vasodilation

Inhibition of mitochondrial complex (MC) I activity in carotid body chemoreceptor glomus cells, the prototypical acute O₂ sensors in mammals, suppresses responsiveness of these cells to hypoxia^43,44,45. Hence, we investigated whether a similar phenomenon occurred in arterial myocytes. Disruption of MCI was achieved by targeting the gene encoding Ndufs2, a core subunit essential for ubiquinone (CoQ) binding and NADH dehydrogenase activity⁴⁶. Cells lacking Ndufs2 show almost complete abolition of MCI function^44,47. In the present study, mice (of both sexes) carrying Ndufs2 floxed and minus alleles⁴⁵ were bred with mice carrying the tamoxifen (TMX)-activated Cre recombinase gene driven by the smooth muscle-specific SM22 promoter^48,49. These adult mice, with conditional deficiency of MCI in VSM cells (MCI-SM mice), were studied in comparison with wild type (WT) mice. Two-month-old mice received a TMX diet for one month and were left to recover for at least one additional month on a normal diet (Fig. 1a). TMX-treated MCI-SM mice ( ≥ 4-months old) showed a strong and selective decrease in Ndufs2 mRNA obtained from the femoral artery and aorta (Fig. 1b, c). Ablation of Ndufs2 in VSM cells resulted in an ~80% decrease in MCI activity in the aorta (Fig. 1d). Except for an increase in spleen size, MCI-SM mice did not show any alteration in survival, body weight or anatomical features (Supplementary Figs. 1a–c). They had a normal hematocrit and arterial histology (Supplementary Figs. 1d–g), with normal heart rate and hypoxic ventilatory responses to hypoxia, a homeostatic reflex largely dependent on O₂-sensing by carotid body glomus cells^44,50, and hypercapnia (Supplementary Figs. 1h–j), However, in comparison with WT mice, MCI-SM mice showed a marked hyperlactatemia (Fig. 1e), probably as a consequence of the up-regulation of glycolysis in MCI-deficient myocytes⁴⁵. They also showed a significant decrease in systolic and diastolic blood pressures (Fig. 1f), a phenomenon that could be due to the decreased vascular sensitivity to O₂ tension and the blunted vasoconstrictor effect of O₂ on aorta and peripheral arteries (see below). These changes indicated that even under basal physiological conditions the loss of MCI activity in VSM cells alters fundamental homeostatic mechanisms.

**Fig. 1: Acute hypoxic inhibition of Ca²⁺ channel currents is abolished in mitochondrial complex I-deficient femoral arterial myocytes.**

Next, we studied acutely dispersed femoral myocytes, as a representative example of conduit systemic myocytes^39,51, from MCI-SM mice which, despite having a selective absence of Ndufs2 immunoreactivity, appeared healthy and with the characteristic elongated shape of WT VSM cells (Fig. 1g). However, they showed an increase in mitochondrial number, similar to what we have previously seen in MCI-deficient carotid body glomus cells⁴⁴, and aberrant ultrastructural changes that resemble those reported in skeletal muscle with mitochondrial myopathies⁵² (Supplementary Figs. 2a–i). Patch clamped MCI-deficient myocytes exhibited normal passive electrical parameters and density of voltage-gated Ca²⁺ and K⁺ channels (Fig. 1h; and Supplementary Figs. 3a–e), as well as responses to vasoactive agents (see below), indicating that as it occurs with O₂-sensitive carotid body glomus cells⁴⁴ arterial myocytes can survive mitochondrial dysfunction possibly due to compensatory up-regulation of glycolysis⁴⁵ (see Fig. 1e). WT mouse femoral myocytes were exposed to hypoxia by switching from PO₂ of 150 mmHg to ~15 mmHg in the solution bathing the cells³⁹, covering in this manner the full range of O₂ tensions existing in most tissues during normoxia and physiological hypoxia⁵³. As observed previously in several types of systemic myocytes^39,51, this treatment produced a fast, progressive, and reversible decrease in the amplitude of whole-cell L-type Ca²⁺ channel currents without altering their activation or deactivation kinetics (Fig. 1i–k and Supplementary Figs. 4a–d). However, in contrast to their normal histological appearance and basic electrophysiology, VSM cells from MCI-SM mice exhibited a practically abolished modulation of Ca²⁺ channel activity by PO₂ (Fig. 1l, m). In these patch clamp experiments cells were dialyzed with intracellular MgATP to prevent activation of ATP-dependent K⁺ channels, despite the participation of these channels in VSM responses to physiological hypoxia is questioned^{21,32,35,41,54,55}. Voltage-dependent K⁺ currents were unaffected by hypoxia (Supplementary Figs. 4e–g), suggesting that in femoral vascular myocytes voltage-gated K⁺ channels lack sites of interaction with the mitochondrial signals.

Femoral myocytes did not show any evidence of voltage-gated Na⁺ currents, despite we used a high-Na⁺ external solution in our patch clamp experiments. In addition, the size and time course of inward currents were unaffected by complete replacement of extracellular Na⁺ with the impermeant cation N-methyl-D-glucamine (Supplementary Figs. 5a, c). Modulation of the Ca²⁺ currents (recorded without Na⁺ in the internal solution) by hypoxia was also unaltered in the absence of extracellular Na⁺ (Supplementary Figs. 5b, c), thus making it unlikely any relevant role of Na⁺ control of mitochondrial signaling⁵⁶ in the response to hypoxia of myocyte Ca²⁺ channels.

In agreement with the electrophysiological data, a strong inhibition of transmembrane Ca²⁺ influx triggered by either methyl 2,5-dimethyl-4-[2-(phenylmethyl)benzoyl]-1H-pyrrole-3-carboxylate (FPL), an L-type Ca²⁺ channel activator^57,58, or depolarization with high extracellular K⁺^15,36 was produced by hypoxia in intact Fura 2-loaded femoral myocytes. These effects of hypoxia were eliminated in MCI-SM myocytes (Fig. 2a–c and Supplementary Figs. 6a–d). Rotenone, a highly membrane-permeant MCI blocker, mimicked hypoxia and reversibly inhibited FPL-induced transmembrane Ca²⁺ influx, an effect that was markedly attenuated in femoral myocytes from MCI-SM mice (Fig. 2d, e). Preincubation of WT myocytes with rotenone not only produced a dose-dependent inhibition of the amplitude of FPL-induced Ca²⁺ signals but it occluded any further effect of hypoxia (Supplementary Figs. 7a–d). Caffeine-evoked Ca²⁺ release from the sarcoplasmic reticulum⁵⁹ was unaltered in MCI-deficient myocytes (Supplementary Figs. 8a, b) and unaffected by rotenone (Supplementary Figs. 8c, d). Similar to hypoxia, blockers of the MCI (rotenone or piericidin A), MCIII (antimycin A) and MCIV (cyanide) decreased the amplitude of the Ca²⁺ currents and prevented any further effect of hypoxia (Fig. 2f left, g; and Supplementary Fig. 9a–f). Inhibition of the current with rotenone also occluded any further effect of cyanide (Fig. 2f, right, g).

**Fig. 2: Acute hypoxic inhibition of transmembrane Ca²⁺ influx is lost in mice with mitochondrial complex I-deficient arterial myocytes.**

As responsiveness to hypoxia may change in different arterial segments and territories^8,17, we also studied cerebral arterial VSM cells, as a representative example of resistance myocytes, which have a smaller size (Fig. 3a) and capacitance (Supplementary Figs. 3a and 10a) than femoral myocytes but similar O₂-sensitive Ca²⁺ currents (Fig. 3b, c). Passive electrical parameters and Ca²⁺ current amplitudes of cerebral myocytes were unaffected by MCI deficiency (Supplementary Fig. 10a); however, the modulation of Ca²⁺ channel activity by hypoxia was abolished (Fig. 3b, c; Supplementary Fig. 10b, c). We also tested that modulation of FPL- or high extracellular K⁺-induced transmembrane Ca²⁺ influx by hypoxia was abolished in MCI-deficient myocytes from middle or posterior cerebral arteries (example of resistance VSM cells) (Fig. 3d–f) and from aorta (example of myocytes from conduit and more elastic artery) (Supplementary Figs. 11a–c). Consistent with the findings in single femoral and cerebral myocytes, endothelium-denuded femoral arteries resected from MCI-SM mice showed a normal passive diameter and contractile responses to FPL, high K⁺ and caffeine, although they had strongly inhibited hypoxia-induced vasodilation (Fig. 3g–j).

**Fig. 3: Mitochondrial complex I disruption abolishes hypoxic inhibition of transmembrane Ca²⁺ influx in cerebral myocytes and arterial hypoxic vasodilation.**

Taken together, these data indicate that mice can survive MCI disruption in VSM, although they exhibit hyperlactatemia, decreased systemic blood pressure, and strong inhibition of HVD. Remarkably, MCI-deficient arterial myocytes have a normal morphology, electrical parameters, voltage-gated ionic currents, and response to caffeine but lack the hypoxic inhibition of transmembrane Ca²⁺ influx. These functional alterations appear to be due to the loss of L-type Ca²⁺ channel modulation by O₂, a physiological process whose underlying mechanism has remained unknown and is now shown to be strictly dependent on MCI function.

Interaction of O₂-dependent mitochondrial signals with Ca²⁺ channels

The production of intracellular reactive oxygen species (ROS) of mitochondrial origin has been reported in several VSM cell types exposed to hypoxia^60,61,62. On the other hand, in O₂ sensing carotid body glomus cells, hypoxia induces rapid and reversible increases in NADH and H₂O₂, which are abolished by MCI-deficiency⁴⁴. It has recently been proposed that acute O₂ sensing in glomus cells relies on a slowdown of the mitochondrial electron transport chain (ETC) in hypoxia and a subsequent backlog of electrons with the accumulation of NADH and ROS (H₂O₂) in the mitochondrial intermembrane space (IMS) (Fig. 4a). After equilibration with the cytosol, these mitochondrial signals, which are also produced by the rotenone-induced blockade of MCI, modulate membrane ion channels to induce glomus cell depolarization and transmitter release^44,47. To test whether similar mitochondrial-to-membrane signaling mechanisms operate in VSM cells, we studied the effects of the intracellular application of NADH and H₂O₂, as well as the reducing agent N-acetylcysteine (NAC), on Ca²⁺ currents in dialyzed patch-clamped WT myocytes (Fig. 4b). We also assessed the effect of pharmacological modulation of the intracellular NADH/NAD⁺ ratio by extracellular application of highly membrane permeable α-hydroxybutyrate or α-ketobutyrate (αHB/αKB)^47,63 (Fig. 4b). Both internal NADH and H₂O₂ at concentrations near estimated physiological levels produced a clear dose-dependent inhibition of Ca²⁺ currents, with the effects of the two agents found to be additive (Fig. 4c, d; and Supplementary Fig. 12a, b). Similar to hypoxia (see Figs. 1i–l), NADH and H₂O₂ consistently decreased Ca²⁺ current amplitude without affecting their activation or deactivation kinetics (Fig. 4e, f). Modulation of the Ca²⁺ current by hypoxia was blunted or even abolished in cells already loaded with relatively low doses of NADH plus H₂O₂ (Fig. 4c, g).

**Fig. 4: Effect of changes in intracellular levels of NADH and/or H₂O₂ on Ca²⁺ currents in arterial myocytes.**

Dialysis of myocytes with NAC (0.5 mM) did not appreciably change the Ca²⁺ current amplitude (Fig. 5a, b), but did strongly inhibit the hypoxic modulation of these channels (Fig. 5a, c). Moreover, the application of cell-permeant αHB, which increases intracellular NADH levels, decreased, as seen with hypoxia, the amplitude of the Ca²⁺ current (Fig. 5d, e). In contrast, αKB, which reduces intracellular NADH^47,63, had little effect by itself but decreased the hypoxic modulation of the current (Fig. 5d, f). These data indicate that the increase in intracellular NADH and H₂O₂ levels produces a robust inhibition of myocyte Ca²⁺ currents similar to that elicited by hypoxia or rotenone, and that lowering PO₂ does not have any further effect on the current once NADH and H₂O₂ are interacting with the Ca²⁺ channels. Loading the cells with NAC can prevent the action of hypoxia-generated H₂O₂, while the increase in NADH alone (application of extracellular αHB) partially mimics hypoxia. These findings strongly suggest that the fast, reversible, and MCI-dependent hypoxic modulation of myocyte L-type Ca²⁺ channel activity is mediated by mitochondrial NADH and H₂O₂.

**Fig. 5: Effect of NADH and redox reagents on Ca²⁺ currents in arterial myocytes.**

Transgenic expression of yeast NADH dehydrogenase in MCI-deficient arterial myocytes restores O₂ modulation of Ca²⁺ channels

Given the essential role played by MCI in acute hypoxia signaling in arterial myocytes, we tested whether transgenic expression of Ndi1, a single-molecule yeast NADH dehydrogenase, can rescue Ca²⁺ channel sensitivity to hypoxia in MCI-deficient VSM cells. Ndi1 is attached to the inner mitochondrial membrane and transfers electrons from NADH to CoQ, but unlike MCI this activity is not coupled to a proton pump⁴⁵ (Fig. 6a). In this way, Ndi1 expression allowed us to assess if the recovery of NADH dehydrogenase activity by itself, without the electrochemical gradient generated by MCI, is sufficient to induce functional recovery in MCI-null myocytes. To this end, we generated two new conditional adult mouse models (named WT/NDI1-SM and MCI/NDI1-SM) in which TMX treatment induced Ndi1 expression in WT and MCI-SM mice, respectively (Fig. 6b). We showed immunocytochemical evidence of Ndi1 expression in VSM cells (Fig. 6c) and the appropriate selective down-/up-regulation of Ndufs2 and Ndi1 mRNAs, respectively, in arteries of the various mouse strains (Fig. 6d, e). Ndi1 expression did not induce any major alteration in survival, body weight or organ size in WT mice; however, the presence of Ndi1 rescued practically all the systemic structural and functional alterations observed in MCI-SM mice (Supplementary Figs. 13a–d). Notably, increased lactatemia and low systolic and diastolic blood pressures, which were characteristic features of MCI-SM mice (see Fig. 1e, f), showed full recovery in MCI/NDI1-SM mice (Fig. 6f, g).

**Fig. 6: Transgenic expression of yeast Ndi1 rescues O₂ modulation of Ca²⁺ channel activity in MCI-deficient arterial myocytes.**

While Ndi1 expression in WT myocytes did not produce any change in Ca²⁺ current amplitude or modulation by O₂ (Supplementary Fig. 13e), Ndi1 did induce an almost complete recovery of the inhibition of Ca²⁺ current by hypoxia in MCI/NDI1-SM myocytes (Fig. 6h, i, and Supplementary Fig. 13f), which was abolished in MCI-null VSM cells (see Figs. 1i, m). In agreement with these data, inhibition of FPL-induced transmembrane Ca²⁺ influx by hypoxia, which was blunted by Ndufs2 deficiency (see Fig. 2a–c), was also rescued in Ndi1-expressing myocytes (Fig. 7a, left, b, c). In contrast, the responsiveness of FPL-induced transmembrane Ca²⁺ influx to rotenone (see Fig. 2d, e) was not recovered in MCI/NDI1-SM myocytes, where it is well known that Ndi1 is insensitive to this drug⁴⁵ (Fig. 7a right, d).

**Fig. 7: Transgenic expression of yeast Ndi1 rescues O₂ modulation of cytosolic Ca²⁺ and NADH signals in MCI-deficient arterial myocytes.**

To further demonstrate the recovery of hypoxia signaling in Ndi1 expressing MCI-deficient myocytes, we recorded reversible changes in NADH autofluorescence induced by hypoxia in single dispersed myocytes using two photon microfluorimetry. As shown previously in O₂-sensitive glomus cells⁴⁷, hypoxia induced rapid and reversible increases in NADH levels in WT myocytes. This signal was practically abolished in cells from MCI-SM mice and was fully recovered in MCI/NDI1-SM myocytes (Fig. 7e, f). In all these genetically different cell types, a clear decrease in NADH levels was observed after addition of αKB (1 mM) (Fig. 7e), which permeates to the cell interior and consumes NADH during its conversion to αHb (see Fig. 5d). These data further support the view that the essential role played by MCI during hypoxia is to generate the signaling molecules (NADH and H₂O₂) that regulate Ca²⁺ channels. In this regard, Ndufs2 deficiency can be fully rescued by the transgenic expression of yeast NADH dehydrogenase.

Acute O₂ sensitivity of arterial myocytes relies on the constitutive activity of Hif1α and expression of atypical cytochrome c oxidase subunits

The hypoxia inducible transcription factors Hif1α and Hif2α are key mediators of cellular transcriptional responses to hypoxia⁶⁴. We have shown that Hif2α, which is expressed at constitutively high levels in the carotid body^65,66,67, is required for induction of a transcriptional program that determines the expression of atypical MCIV subunit isoforms conferring acute responsiveness of chemoreceptor glomus cells to physiological hypoxia^68,69,70. Given that Hif2α is expressed in systemic arteries, although at lower levels than in the lung (Supplementary Fig. 14a), we investigated whether conditional deletion of the Hif2α gene (Epas1) also alters myocyte acute O₂ sensing. To this end, we analyzed the EPAS1-ESR mouse model with a TMX-induced loss of Epas1 alleles in adulthood⁶⁸ (Supplementary Fig. 14b). We found a marked selective decrease of Epas1 mRNA in the aortas of EPAS1-ESR mice, however no changes in mRNA levels of relevant MCIV subunit isoforms were observed (Supplementary Fig. 14c). Modulation of Ca²⁺ channel activity (Supplementary Figs. 15a–d) and FPL-mediated Ca²⁺ influx (Supplementary Figs. 15e–g) during hypoxia were also unaltered in Hif2α-deficient myocytes. These findings indicated that Hif2α is not responsible for the acute O₂ sensitivity of systemic VSM cells. They also led us to hypothesize that the expression of MCIV subunit isoforms presumably involved in acute O₂ sensing could be under the transcriptional control of Hif1α, which is expressed in arterial smooth muscle at higher levels than those seen for Hif2α (Supplementary Fig. 16a).

To address this question, we generated adult HIF1α-SM mice with a conditional loss of Hif1α alleles in VSM cells (Fig. 8a) following the same breeding protocol and TMX treatment described for MCI-SM mice (see Fig. 1a). We confirmed a marked selective decrease in Hif1α mRNA levels in aortic tissue (with unchanged Epas1 mRNA levels) paralleled by a manifest mRNA down-regulation of relevant atypical MCIV subunit isoforms, such as Cox4i2 or Cox8b⁶⁸, as well as Pgk1, a well-known Hif1α-dependent gene encoding a glycolytic enzyme^71,72. Other mRNA species encoding MCIV subunits (Ndufa4l2, Ndufa4), enzymes (Pcx), or transcription factors suggested to regulate the expression of MCIV subunits (Rbpj, Chchd2, Cxxc5)⁷³ were either less affected or unaltered (Fig. 8b, c). Remarkably, HIF1α-SM mice did not show any major alteration in body weight or gross modification of organ size, hematocrit or heart rate (Supplementary Figs. 16b–e). However, similar to MCI-SM mice (see Fig. 1f), HIF1α-SM mice showed a clear decrease in systolic and diastolic blood pressures (Fig. 8d). Hif1α-deficient femoral artery myocytes appeared normal, with an unchanged density of L-type Ca²⁺ channels (Supplementary Figs. 16f, g). However, in these myocytes, modulation of Ca²⁺ channel activity by hypoxia, as evidenced both by direct measurements of whole-cell currents and by FPL-induced transmembrane Ca²⁺ influx in Fura-2-loaded isolated cells, were practically abolished (Fig. 8e–i, Supplementary Figs. 16f, g). In contrast, the rise in cytosolic [Ca²⁺] induced by either caffeine or phenylephrine (an α₁ receptor agonist)⁷⁴, was unaffected (Fig. 8j). These data indicated that O₂ modulation of Ca²⁺ channel activity in arterial myocytes depends on Hif1α, possibly due to the expression of Hif1α-dependent mitochondrial MCIV subunit isoforms.

**Fig. 8: Responsiveness to acute hypoxia is selectively abolished in *Hif1α*-deficient arterial myocytes.**

Among the MCIV subunits whose mRNA expression was modulated by Hif1α in arterial myocytes (see Fig. 8b, c), we focused on Cox4i2, a tissue-specific Cox4 isoform that is highly expressed in the lung⁷⁵ and the carotid body^66,67, and which is induced by hypoxia in several cell classes in a Hif1α-dependent manner⁷⁶. Expression of Cox4i2 instead of Cox4i, the most common subunit isoform, alters cytochrome c oxidase (CCO) activity^75,76,77 and is essential for ROS production and depolarization of cultured pulmonary VSM cells in response to hypoxia⁷⁸. Mice lacking Cox4i2 show abolition of the acute phase of hypoxic pulmonary vasoconstriction⁷⁸ and strong inhibition of the acute O₂-sensing properties of CB glomus cells⁶⁸. To analyze the participation of Cox4i2 in the acute O₂-modulation of Ca²⁺ channel activity in systemic arterial myocytes, we generated adult conditional Cox4i2-deficient mouse models using either TMX-activatable generalized Ubiquitin (UBC)-Cre or smooth muscle-specific SM-Cre (COX4I2-UBC and COX4I2-SM mice, respectively) (Fig. 9a). In the two mouse models, we observed a clear selective decrease in Cox4i2 mRNA in aortic myocytes (Fig. 9b, c). In addition, we showed a decrease in Cox4i2 immunostaining in Cox4i2-deficient femoral myocytes (Fig. 9d). COX4I2-SM mice showed normal body weight, organ size and hematocrit (Supplementary Figs. 17a–c). However, we observed a strong inhibition of the hypoxic modulation of Ca²⁺ currents measured in dispersed Cox4i2-deficient femoral VSM cells (Fig. 9e, f; Supplementary Figs. 17d, e). Moreover, endothelium-denuded femoral arteries removed from these mice had normal passive diameters and contractile responses to FPL and high K⁺, but were, nevertheless, practically insensitive to hypoxia (Fig. 9g–j). These observations suggest that the Hif1α-dependent expression of Cox4i2 in vascular myocytes plays a fundamental role in the acute modulation of Ca²⁺ channel activity by PO₂ and in HVD.

**Fig. 9: Acute O₂ modulation of Ca²⁺ channel activity and hypoxic vasodilation are lost in *Cox4i2*-deficient arterial myocytes.**

Discussion

Two main conclusions can be drawn from the studies presented in this paper. The first is that acute modulation of L-type Ca²⁺ channels by PO₂ in systemic arterial myocytes depends on the integrated functions of mitochondria working as O₂ sensors and effectors. The O₂ modulation of Ca²⁺ channels is abolished in MCI-deficient cells, but can be rescued by the transgenic expression of a yeast NADH dehydrogenase, and mimicked by the intracellular application of NADH and H₂O₂. Remarkably, modulation of Ca²⁺ channels by O₂ depends on dynamic changes in the mitochondrial ETC, and requires a permanent, constitutive activation of Hif1α, which in turn determines the expression of at least one atypical MCIV subunit isoform. It must be also stressed that our electrophysiological studies, done on VMS cells dialyzed with 4 mM MgATP, strongly suggest that changes in intracellular ATP levels^{21,32,35,41,54,55} are not involved in inhibition of Ca²⁺ channel activity and depolarization-induced Ca²⁺ influx during hypoxia. These findings support a mitochondrial-to-membrane signaling (MMS) model of acute O₂ sensing^44,68 in which the decrease in MCIV activity under hypoxia leads to a backlog of electrons along the ETC and an increase in the reduced ubiquinone (CoQH₂) pool. This results in a slowdown of MCI activity and the accumulation of NADH and H₂O₂ which interact with Ca²⁺ channels (Fig. 10). This model provides a mechanistic explanation for the rapid and reversible inhibition of Ca²⁺ currents by hypoxia, a phenomenon observed in myocytes from several mammalian systemic conduit and resistance arteries whose underlying mechanisms have remained unknown^{9,35,37,39,40,41}.

**Fig. 10: Schematic representation of the mitochondrial-to-membrane signaling model of acute hypoxic vasodilation in systemic arteries.**

The mitochondrial-based O₂-sensing system in arterial myocytes resembles in many aspects the MMS pathway supporting the acute O₂-dependent control of membrane excitability in carotid body glomus cells that is responsible for the ventilatory response to hypoxia^45,50,68. Although studied in less detail, similar mechanisms appear to mediate acute O₂ sensing and responsiveness to hypoxia in other peripheral chemoreceptors, such as cells in the embryonic/neonatal adrenal medulla and the abdominal paraganglia^44,79. Moreover, the mitochondria production of H₂O₂ in myocytes of resistance arteries has been suggested to mediate acute pulmonary artery vasoconstriction^60,61,62. It seems, therefore, that the MMS model described here for HVD could be a general mechanism that operates in organs of the homeostatic acute O₂ sensing system⁸⁰. To function as O₂ sensors within physiological ranges of PO₂, mitochondria require the expression of atypical MCIV subunit isoforms (Cox4i2 and others) that may alter the catalytic activity (affinity for O₂ and/or accessibility of substrates to the active site) of CCO^68,77. Remarkably, whereas in carotid body glomus cells and catecholaminergic peripheral chemoreceptors the expression of Cox4i2 depends on the constitutive high expression of Hif2α,^68,79 in systemic arterial myocytes the same process relies on the constitutive activity of Hif1α, which further indicates that the two transcription factors are not redundant but that they have tissue-specific specializations^81,82.

Our results strongly suggest that NADH and H₂O₂ are the mitochondrial hypoxic signals that inhibit Ca²⁺ channel activity in arterial myocytes. This observation fits well with previous data indicating that recombinant as well as native L-type Ca²⁺ channels are modulated by H₂O₂^42,83,84,85. Nicotine dinucleotides have been shown to bind and modulate several subtypes of K⁺ channels; however, their effect on other ion channel types is less known⁸⁶. Although in our studies HVD seemed to depend mainly on the inhibition of Ca²⁺ channel activity, it is conceivable that modulation of other channel types by PO₂ contributes to vasodilation in specific arterial segments, given the functional heterogeneity existing in vascular territories^8,17. The activation of voltage-gated K⁺ channels has been suggested to participate in HVD⁸⁷, with a recent study in myocytes from small-diameter mouse mesenteric arteries showing that intracellular NADH stimulates K⁺ channel activity and enhances the sensitivity of the channel to H₂O₂⁸⁸. NADH and H₂O₂ are promiscuous molecules whose actions probably depend on the ion channel subunits expressed in the various VSM cells and the existence of O₂ sensing microdomains formed by mitochondria placed near clusters of specific ion channel classes (e.g., L-type Ca²⁺ channels) in the plasma membrane⁸⁹. In addition to NADH and H₂O₂, it has been suggested that oxidation of the constitutively produced redox reagent hydrogen sulfide (H₂S) by the enzyme sulfide-quinone oxidoreductase (SQR) in VSM cells contributes to the CoQH₂ pool. In this way, accumulation of CoQH₂ during hypoxia leads to an increase in H₂S⁹⁰. We did not consider H₂S to be a relevant hypoxic signal in our studies because rotenone and piericidin A, selective inhibitors of MCI, completely inhibited the effect of hypoxia on Ca²⁺ channels.

The second far-reaching conclusion from our work is that the modulation of Ca²⁺ channels by PO₂ in arterial myocytes is a process of functional relevance. Circulatory blood flow is regulated by a complex ensemble of signaling pathways that show redundancy and compensation^7,8,91. However, we found that both modulation of Ca²⁺ channel activity by PO₂ in single myocytes and HVD of whole arteries ex vivo are selectively abolished in adult Ndufs2-, Cox4i2-, or Hif1α-deficient mice, despite the vessels and myocytes having a normal structure, electrophysiology, and responses to other vasoactive agents (FPL, high K⁺, caffeine, or phenylephrine). Moreover, adult MCI-SM or HIF1α-SM mice, which did not show O₂ modulation of myocyte Ca²⁺ channels, presented with a decrease in basal systemic blood pressure. These alterations in the control of tissue blood perfusion at rest could have severe consequences for the maintenance of homeostasis under challenging conditions (e.g., physical exercise or sojourn at high altitude) or for long-term adaptive brain and cardiac functions. In addition to their pathophysiological significance, our findings provide new scenarios (NADH mimetics, mitochondrial metabolism, and HIF inhibitors among others) of potential interest in cardiovascular pharmacology.

Methods

Genetically modified mouse models

MCI-SM mice

Mice carrying Ndufs2 floxed allele⁴⁴ were bred with those carrying tamoxifen-induced CRE recombinase under the control of smooth muscle cell-specific SM22α promoter⁴⁸ to generate NDUFS2 adult conditional knockout mice (Ndufs2 flox/-;SM22-Cre, named as MCI-SM in the text and figures).

MCI/NDI1-SM and WT/NDI1-SM mice

MCI-SM mice were bred with mice carrying a lox-STOP-lox site preceding yeast Ndi1 sequence targeted to the Rosa26 locus⁴⁵. The resulting mice (MCI/NDI1-SM mice) expressed NDI1 in NDUFS2-deficient VSM cells. Mice carrying yeast NADH dehydrogenase (NDI1) and green fluorescent protein (GFP) sequences were also bred with those carrying tamoxifen-induced CRE recombinase under the control of SM22α promoter to express NDI1 in VSM cells (named as WT/NDI1-SM).

HIF1α-SM mice

Mice carrying Hif1α floxed allele (The Jackson laboratory, Stock No: 007561)⁹² were bred with those carrying tamoxifen-induced CRE recombinase under the control of smooth muscle cell-specific SM22α promoter to generate HIF1α adult conditional knockout mice (HIF1α flox/-;SM22-Cre, named as HIF1α-SM in the text and figures).

EPAS1-ESR mice

Mice carrying Epas1 floxed allele (The Jackson laboratory, Stock No: 008407)^68,82 were bred with those carrying tamoxifen-inducible Cre recombinase transgene⁹³ to generate HIF2α adult conditional knockout mice (Epas1 flox/-;ESR-Cre, named as EPAS1-ESR in the text and figures).

COX4I2-SM and COX4I2-UBC mice

Mice carrying Cox4i2 floxed allele^68,78 were bred with those carrying tamoxifen-induced CRE recombinase under the control of either smooth muscle cell-specific SM22α promoter or human ubiquitin C (UBC) promoter⁹⁴ to generate COX4I2 adult conditional knockout mice (Cox4i2 flox/-;SM22-Cre and Cox4i2 flox/-;UBC-Cre mice which are named, respectively, as COX4I2-SM and COX4I2-UBC in the text and figures).

Animal maintenance and treatments

Mice were housed at regulated temperature (22 ± 1 °C) in a 12 h light/dark cycle with ad libitum access to food and drink. Both male and female mice were used in the current study. Mice strains were in mixed genetic background (129 Sv:C57BL/6 or 129 Sv:C57BL/6/Balb/c). To induce Cre-mediated recombination, two-months old wild type and knockout mice were treated with tamoxifen-containing diet (TAM400/CreER, 400 mg tamoxifen citrate/kg, Envigo) for a month followed by 1 to 1.5 months of normal diet before experiments. For in vitro studies, animals were sacrificed with intraperitoneal administration of a lethal dose of sodium thiopental (120-150 mg/Kg) before tissue dissection. Dissected tissues were used for either functional, biochemical or histological analyzes. For RNA isolation and measurement of mitochondrial complex I activity, tissues were flash-frozen in liquid N₂ and stored at −80 °C until use. All procedures were approved by the Institutional Committee for animal care and use at the University of Seville (PN2019 07/04/2020/051). Handling of the animals was conducted in accordance with the European Community Council directive of 22 September 2010 (Directive 2010/63/EU) and the implementations of 5 June 2019 (Regulation 2019/2010) for the Care and Use of Laboratory Animals. Mice used in the in vivo measurements were subsequently sacrificed for the in vitro experiments in order to minimize the number of animals used in the study.

Plethysmography

Plethysmography used to monitor ventilation in conscious unrestricted rodents has been previously described^45,50. Mice were placed inside plethysmographic chambers (EMKA Technologies) and continuously exposed at a constant flow rate (1 L/min) with a gas mixture containing 21% O₂, 0.04% CO₂ and 78.96% N₂ (normoxia); 10% O₂, 0.04% CO₂ and 89.96% N₂ (hypoxia); or 21% O_2, 5% CO₂ and 74% N₂ (hypercapnia). Data acquisition was performed using the software Iox2 (EMKA Technologies). The hermetic chambers were provided with O₂ and CO₂ sensors to monitor the gas composition in parallel with changes in respiratory frequency recorded by a pressure sensor during the experiment. Each animal was exposed to two cycles of hypoxia (5 min) followed by a final exposure to hypercapnia (2 min).

Measurement of arterial pressure and heart rate

Systolic and diastolic pressures and heart rate were determined by using a non-invasive tail-cuff transmission photoplethysmography (BP-2000) Blood Pressure Analysis System for mice and rats, (Bioseb, in vivo research instruments, RRID:SCR_02 2985). Conscious mice were placed in a restrained cage with automatic temperature control that gently warms the animals. Before taking the real pressure values, mice were recorded for 3 previous days to allow them to get used to the system and to avoid pressure increases due to stress produced by the technique. Values recorded during the 4th and 5th days were used to average systolic and diastolic pressures and heart rate in each mouse.

Measurement of hematocrit and blood lactate

Lactate Plus Meter was used to determine lactate plasma concentration (Nova Biomedical). Control solutions with 1–1.6 mM and 4–5.4 mM lactate, provided by the supplier, were used to calibrate the device. For measurements, a small drop of blood obtained from the mouse tail after a lancet puncture, was placed on the specific strip for lactate quantification. Hematocrit was measured by collecting blood from an incision of the carotid artery using a hematocrit tube allowing blood to flow by capillarity action into tube. The ends of the tube were sealed with bone wax and the tubes were placed in a microhematocrit centrifuge and span for 5 minutes at 2.500 g. The hematocrit is expressed as the percentage of blood volume occupied by erythrocytes.

Measurement of contractility in arterial rings

Femoral arteries were isolated, cleaned of connective and endothelial tissue, cut in rings ( ~ 2 mm in diameter) and mounted on a small vessel myograph (Danish Myo Technology) connected to a digital recorder (AcqKnowledge 3.8.1, BIOPAC System) to measure isometric tension. Endothelium-denuded rings were placed in a chamber with Kreb’s solution and bubbled with 95 % O₂ and 5 % CO₂ at pH = 7.4. The “hypoxic” solution was bubbled with 5% CO₂, and 95% N₂. Before the experiments, the segments were subjected to an optimal tension test (90 % tension was equivalent to an intramural pressure of 100 mmHg) and stabilized for at least 1 h, when basal tone was determined in unstimulated arteries. The drugs were added directly to the chamber while vessel tension was monitored. Experiments were performed at ~30 °C.

Histology

Complete femoral arteries (with endothelial and connective tissue) were fixed with 4% paraformaldehyde. Arteries were then cleared and embedded in paraffin. Paraffin sections (6 μm) were dewaxed, rehydrated, and stained with hematoxilin–eosin following standard procedures. Measures were done with ImageJ software (version 1.51 S) and each value was obtained from triplicate determinations of femoral artery from at least four mice in each condition.

Mitochondrial complex I activity dipstick assay

MCI activity in aorta was determined using complex I enzyme activity dipstick assay kit (Abcam) following the manufacturer’s instruction with minor modifications. Aorta arteries pooled from 2 mice were homogenized in 100 µl extraction buffer followed by incubation on ice for 20 minutes. The homogenate was then centrifuged at 16000 g for 30 minutes and the supernatant was collected for the enzymatic assay. 8 µg of protein extract, which was determined using the BioRad protein assay (BioRad), were applied to measure the MCI activity. The signal on the dipstick was scanned with an image analyzer (ImageQuant LAS 4000mini, GE Healthcare) and the signal intensity was quantified using the ImageQuant TL software (GE Healthcare).

Real-time quantitative PCR

Total RNA was isolated from adult mice ( ~ 4 months old) 1–1.5 months after the completion of tamoxifen treatment. Total RNA was isolated from aorta and femoral arteries using RNeasy Micro kit (Qiagen). Due to the small tissue size, each femoral artery replicate was pooled from 2 mice to obtain enough amount of RNA. The RNA quality was determined using an Agilent 2100 Bioanalyzer (Agilent). cRNA was then amplified from VSM total RNA using GeneChip WT PLUS Reagent Kit (Affymetrix). Total RNA from brain and lungs were isolated using Trizol (Ambion, Life Technologies). 500 ng of total RNA (or amplified cRNA in the case of aorta and femoral arteries) was copied to cDNA using QuantiTect Reverse Transcription Kit (Qiagen) in a final volume of 20 μl. Real-time quantitative PCR reactions were performed in a 7500 Fast Real Time PCR System (Life Technologies). PCR reactions were performed in duplicates in a total volume of 20 μl containing 1–4 μl of cDNA solution and 1 μl of Taqman probe of the specific gene (ThermoFisher Scientific) (Supplementary Table 1. Oligonucleotides). Gapdh was also estimated in each sample to normalize the amount of total RNA (or cRNA) input in order to perform relative quantifications.

Preparation of acutely dispersed arterial myocytes

Vascular smooth muscle (VSM) cells from femoral, middle and posterior cerebral arteries, and aorta were obtained after carefully dissection and cleaning of connective and endothelial tissues. Afterwards, small segments of endothelium-free arteries artery were placed in smooth muscle dissociation solution (SMDS, Ca²⁺ free) containing 1 mg/ml papain (Worthington), 1 mg/ml BSA (Sigma) and 1 mg/ml dithiothreitol (DTT; Sigma) and incubated at 37 °C for 15 min (femoral arteries) or 12 min (cerebral arteries) in a shaking water bath. After two washings in SMDS (with 10 μM Ca²⁺) a second 15 min (femoral arteries) or 12 min (cerebral arteries) incubation was performed with SMDS (10 μM Ca²⁺) in a 0.7 mg/ml collagenase F, 0.3 mg/ml collagenase H (Sigma) and 1 mg/ml BSA solution. Single cells were obtained by gentle trituration with a wide-bore glass pipette, plated on glass cover slips treated with poly-L-lysine, stored at 4◦C, and used within the same day. Ionic composition of SMDS was (in mM): NaCl 145; KCl 4.2; KH₂PO₄ 0.6; MgCl₂ 1.2; Hepes 10; glucose 11, pH adjusted to 7.4 with NaOH.

Immunohistochemistry of isolated arterial myocytes

Freshly isolated VSM cells were plated onto glass cover slips and let to sit for 20 min at room temperature before the immunofluorescence assay. Cells were fixed with 4% paraformaldehyde (PF) in phosphate buffer, pH 7.5, for 15 min at 20 °C, washed in PBTx (PBS, 0.1% Triton X-100), and blocked with PBTx-10 mg/ml BSA-2% normal goat serum for 10 min, Primary antibodies were diluted in blocking solution and incubated with the cells for 60 min at room temperature: COX4I2 (1:100 dilution, 11463-1-1AP, Proteintech); NDUFS2 (1:200 dilution, ab192022, Abcam); α-SMA (1:100 dilution, A2547, Sigma); GFP (1:200 dilution, A11122, Invitrogen). After washes in PBTx, cells were incubated with secondary antibodies for 60 min. The fluorescently labeled secondary antibodies: Alexa 568 or Alexa 488 (1:1000 dilution, A11011, A11015, Invitrogen). Nuclei were labeled with 4’, 6’-diamidino-2-phenylindole (DAPI) staining. Immunofluorescent images were obtained using Leica Stellaris 8 confocal microscopy (Leica) and Leica Application Suite X software (Leica Microsystems).

Electron microscopy

For Transmission Electronic Microscope (TEM) analysis, femoral arteries were fixed for 2 h at 4 °C in 2.5% glutaraldehyde prepared in 0.1 M sodium cacodylated buffer, pH 7.4. After three washing steps (5 min each one), samples were post fixed for 1 hour at 4 °C in 1% osmium tetroxide in the same buffer. After three washing steps (20 min each one), the samples were immersed in 2% uranyl acetate, dehydrated through a gradient acetone series (50%, 70%, 90% and 100%), and embedded in Spurr resin (Tedpella). Blocs were obtained by polymerization at 70 °C for 8 h. Thin sections were cut with a diamond knife in an ultramicrotome (Leica UC7) and examined with a transmission electron microscope (Zeiss Libra 120) operating at 80 kV. The electron microscopy images were taken at different magnifications with an EMCCD camera (TRS 2k × 2k).

Microfluorimetric measurements

For cytosolic Ca²⁺ measurements, dispersed myocytes from femoral and cerebral arteries, and aorta were incubated in SMDS containing 4 μM Fura 2-AM (F-1221, Molecular Probes) for 30 minutes at room temperature and subsequently incubated for 15 min in SMDS to remove excess Fura2-AM. After loading the cells, the coverslip was transferred to the recording chamber and continuously perfused with external solution. The set up consists of an inverted microscope (Nikon eclipse Ti) equipped with a 40x/0.60 NA objective, a monochromator (Polychrome V, Till Photonics), and a CCD camera, controlled by Aquacosmos software (version 2.6, Hamamatsu Photonics). Alternating excitation wavelengths of 340 and 380 nm were used, and background fluorescence was subtracted before obtaining the F340/F380 ratio. A dichroic FF409-Di03 (Semrock) and a band pass filter FF01-510/84 (Semrock) were used. Cytosolic [Ca²⁺] signals were digitized at a sampling interval of 500 ms. Experiments were performed at ~35 °C.

NAD(P)H microfluorimetric measurements in dispersed myocytes from femoral artery were performed using two-photon laser scanning microscope (2PLSM) containing a multiphoton galvanometer scanning system (Scientifica Ltd) with an Olympus Å~60/1.0 NA water-dipping objective lens. A Chameleon Ultra II (680–1080 nm), 3.5 W Ti: sapphire laser system (Coherent laser group) provided the 2 P excitation source. Laser power attenuation was achieved using two Pockels cell electro-optic modulators (S-MP-4700 and S-MP-6000-INT/UK) in series controlled by Labview. Labview software was written by Scientifica (version 12.0). Non-descanned emission photons were detected with GaAsP photomultiplier tube (PMT), S-MDU-PMT-50, green, 490 nm to 560 nm. To measure changes in NADH autofluorescence, cells were excited with 720 nm light using a non-ratiometric protocol. Green channel (490–560 nm) fluorescent emission signals were detected using a non-descanned Scientifica S-MDU-PMT-50-50 select GaAsP PMT. Continuous acquisition mode (rate of 0.76 f.p.s., 146 n pixels) was used for the selected wavelength. Time series analysis was conducted offline using ImageJ (version 1.51 S). Multiple cytosolic ROIs and a background ROI were measured, the background was subtracted for each ROI at each time point. As recording control, 1 mM αKB was added to the external solution (showing a clear decrease in the basal NADH autofluorescence level) at the end of each experiment.

Patch clamp recordings in dispersed myocytes

Macroscopic ionic currents were recorded from dispersed mouse arterial myocytes using the whole cell configuration. Patch clamp pipettes (2–3 MΩ) were pulled from capillary glass tubes (Kimax, Kimble Products) with a horizontal pipette puller (Sutter instruments model P-1000) and fire-polished with a microforge MF-830 (Narishige). Voltage-clamp recordings were obtained with an EPC-7 amplifier (HEKA Electronik). The signal was filtered (3–10 kHz), subsequently digitized with an analog/digital converter (ITC-16 Instrutech Corporation) and finally sent to the computer. Data acquisition and storage was performed using the PULSE/PULSEFIT software (HEKA, Electronics) at a sampling interval of 20 μs. Experiments were performed at room temperature.

Recording solutions

For microflourimetric measurements, dispersed myocytes were transferred to a recording chamber and continuously superfused with a control solution containing (in mM): 125 NaCl, 4.5 KCl, 23 NaHCO₃, 1 MgCl₂, 2.5 CaCl₂, 5 glucose and 5 sucrose, at ~35 °C. In 70 mM K⁺ solution, NaCl was replaced equimolarly with KCl. The “normoxic” solution was bubbled with a gas mixture of 20% O₂, 5% CO₂, and 75% N₂ (O₂ tension ~145–150 mmHg). Hypoxia in the chamber was obtained by switching to a “hypoxic” solution bubbled with 5% CO₂, and 95% N₂ to reach an O₂ tension of ~15 mmHg in the chamber.

Macroscopic inward Ca²⁺ (Ba²⁺), and outward K⁺ currents were recorded in dialyzed arterial myocytes. Solutions used to record whole-cell Ca²⁺ (Ba²⁺) currents contained (in mM): external solution: 140 NaCl, 10 BaCl₂, 4.7 KCl, 10 HEPES, and 10 glucose, pH 7.4 and osmolality 300 mOsm/Kg; and internal solution: 130 CsCl, 10 EGTA, 10 HEPES, and 4 ATP-Mg, pH 7.2 and osmolality 285 mOsm/Kg. In Na⁺-free solutions extracellular Na⁺ was completely equimolarly replaced with impermeant cation N-methy-D-glucamine (NMDG). Solutions used to record whole-cell K⁺ currents contained (in mM): external solution: 140 NaCl, 2.5 KCl, 4 MgCl₂, 2.5 CaCl₂, 10 HEPES, and 10 glucose, pH 7.4; and internal solution: 80 K⁺ glutamate, 50 KCl, 1 MgCl₂, 10 HEPES, 4 MgATP, and 5 EGTA, pH 7.2. The external solutions were bubbled and equilibrated with air or mixtures of N₂ and air to achieve the desired O₂ tension.

For wire myography all measurements were performed in a final volume of 5 mL of Krebs solution, whose composition was (in mM): 119 NaCl, 4.7 KCl, 1.17 KH₂HPO₄, 24 NaHCO₃, 2.5 CaCl₂, 1.17 MgSO₄, 5.5 glucose, constantly bubbled with 95% O₂ and 5% CO₂ at pH 7.4. In 70 mM K⁺ solution, NaCl was replaced equimolarly with KCl. The “hypoxic” solution was bubbled with 5% CO₂, and 95% N₂.

All reagents used were added to the external (e.g., FPL or caffeine) or internal (e.g., MgATP) solutions at the indicated concentrations.

Statistical analysis

Statistical analyzes were carried out using Prism Version 8.2.1 (279) for MacOS. Normality of the data sets was tested with the Shapiro-Wilk test, D’Agostino and Pearson test, or Kolmogorov-Smirnov test. In cases, a log transformation was performed to normalize the distribution prior to parametric analysis. Data with normal distribution were described as mean ± standard error of the mean (SEM) with the number (n) of experiments indicated. For graphical representation of the data with normal distribution we used bar diagrams, with indication of the mean ± SEM, and a scatter plot of the data points superimposed. For graphical representation of non-parametric data, we used box plots with indication of median, quartiles, and whiskers (highest and lowest values). Comparison between two groups was performed as follows: For parametric data we used paired or unpaired t tests. For non-parametric data we used Mann Whitney unpaired test. All tests, parametric and non-parametric, were two-tailed. Comparison of several groups was done as follows: For parametric data we used one-way ANOVA followed by either Tukey’s, Sidak’s or Dunnett’s multiple comparisons test. For non-parametric data, we used Kruskal-Wallis test followed by Dunn’s multiple comparisons test to compare every group with each other. Statistical tests used are indicated in the figure legends. A P < 0.05 was considered statistically significant with the values indicated in the figures.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

The data supporting the findings from this study are available within the manuscript and its supplementary information. Any remaining raw data will be available from the corresponding author upon reasonable request. Source data are provided with this paper.

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Acknowledgements

This research was supported by the Spanish Ministries of Science and Innovation and Health (Grants SAF2016-74990-R, JL-B. and PID2019-106410RB-I00, J.L.-B. funded by MCIN/AEI/10.13039/501100011033, and the European Research Council (ERC Advanced Grant PRJ201502629, J.L.-B.). We thank Dr. Juan Luis Ribas (CITIUS, University of Seville) for help with electron microscopy. We also thank the staff of IBiS and “Centro de Producción y Experimentación Animal Oscar Pintado” for technical assistance.

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Instituto de Biomedicina de Sevilla (IBiS), Hospital Universitario Virgen del Rocío/CSIC/Universidad de Sevilla, Seville, Spain
Alejandro Moreno-Domínguez, Olalla Colinas, Ignacio Arias-Mayenco, José M. Cabeza, Juan L. López-Ogayar, Alberto Pascual & José López-Barneo
Departamento de Fisiología Médica y Biofísica, Facultad de Medicina, Universidad de Sevilla, Seville, Spain
Alejandro Moreno-Domínguez, Olalla Colinas & José López-Barneo
Centro de Investigación Biomédica en Red sobre Enfermedades Neurodegenerativas (CIBERNED), Madrid, Spain
Alejandro Moreno-Domínguez, Olalla Colinas, Alberto Pascual & José López-Barneo
Division of Pulmonary and Critical Care Medicine, Department of Medicine, Northwestern University, Chicago, IL, USA
Navdeep S. Chandel
Excellence Cluster Cardiopulmonary System, University of Giessen and Marburg Lung Centre (UGMLC), German Centre for Lung Research (DZL), Justus-Liebig-University, Giessen, Germany
Norbert Weissmann & Natascha Sommer

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Alejandro Moreno-Domínguez
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Olalla Colinas
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Ignacio Arias-Mayenco
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José M. Cabeza
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Juan L. López-Ogayar
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Navdeep S. Chandel
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Norbert Weissmann
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Natascha Sommer
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Alberto Pascual
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José López-Barneo
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A.M.-D., O.C., I.A.-M., J.M.C. J.L.L.-O., A.P. and J.L.-B. performed the experiments and/or analyzed the data. N.C., N.W. and N.S. provided essential material for the experiments. All authors contributed to the design of the study and to the discussion of the data. A.M.-D. and J.L.-B generated the draft of the paper. J.L.-B. coordinated the project and the writing of the paper.

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Correspondence to José López-Barneo.

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Moreno-Domínguez, A., Colinas, O., Arias-Mayenco, I. et al. Hif1α-dependent mitochondrial acute O₂ sensing and signaling to myocyte Ca²⁺ channels mediate arterial hypoxic vasodilation. Nat Commun 15, 6649 (2024). https://doi.org/10.1038/s41467-024-51023-3

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Received: 15 January 2024
Accepted: 23 July 2024
Published: 05 August 2024
DOI: https://doi.org/10.1038/s41467-024-51023-3
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Hif1α-dependent mitochondrial acute O2 sensing and signaling to myocyte Ca2+ channels mediate arterial hypoxic vasodilation

Abstract

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Introduction

Results

Disruption of mitochondrial complex I in arterial myocytes abolishes acute O2 modulation of Ca2+ channel activity and hypoxic vasodilation

Interaction of O2-dependent mitochondrial signals with Ca2+ channels

Transgenic expression of yeast NADH dehydrogenase in MCI-deficient arterial myocytes restores O2 modulation of Ca2+ channels

Acute O2 sensitivity of arterial myocytes relies on the constitutive activity of Hif1α and expression of atypical cytochrome c oxidase subunits

Discussion

Methods

Genetically modified mouse models

MCI-SM mice

MCI/NDI1-SM and WT/NDI1-SM mice

HIF1α-SM mice

EPAS1-ESR mice

COX4I2-SM and COX4I2-UBC mice

Animal maintenance and treatments

Plethysmography

Measurement of arterial pressure and heart rate

Measurement of hematocrit and blood lactate

Measurement of contractility in arterial rings

Histology

Mitochondrial complex I activity dipstick assay

Real-time quantitative PCR

Preparation of acutely dispersed arterial myocytes

Immunohistochemistry of isolated arterial myocytes

Electron microscopy

Microfluorimetric measurements

Patch clamp recordings in dispersed myocytes

Recording solutions

Statistical analysis

Reporting summary

Data availability

References

Acknowledgements

Author information

Authors and Affiliations

Contributions

Corresponding author

Ethics declarations

Competing interests

Ethics approval

Peer review

Peer review information

Additional information

Supplementary information

Source data

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Hif1α-dependent mitochondrial acute O₂ sensing and signaling to myocyte Ca²⁺ channels mediate arterial hypoxic vasodilation

Disruption of mitochondrial complex I in arterial myocytes abolishes acute O₂ modulation of Ca²⁺ channel activity and hypoxic vasodilation

Interaction of O₂-dependent mitochondrial signals with Ca²⁺ channels

Transgenic expression of yeast NADH dehydrogenase in MCI-deficient arterial myocytes restores O₂ modulation of Ca²⁺ channels

Acute O₂ sensitivity of arterial myocytes relies on the constitutive activity of Hif1α and expression of atypical cytochrome c oxidase subunits