Hypoxia Divergently Regulates Production of Reactive Oxygen Species in Human Pulmonary and Coronary...

1

Hypoxia Divergently Regulates Production of Reactive Oxygen Species in Human

Pulmonary and Coronary Artery Smooth Muscle Cells

Winnie Wu, Oleksandr Platoshyn, Amy L. Firth, and Jason X.-J. Yuan†

Division of Pulmonary and Critical Care Medicine, Department of Medicine, School of

Medicine, University of California, San Diego, La Jolla, CA 92093-0725

Running title: Hypoxia and ROS Production in PASMC and CASMC

†Address correspondence to: Jason X.-J. Yuan, M.D., Ph.D. Division of Pulmonary and Critical Care Medicine Department of Medicine, MTF-252 University of California, San Diego 9200 Gilman Drive, MC 0725 La Jolla, CA 92093-0725 Tel: (858) 822-6534 Fax: (858) 822-6531 E-mail: [email protected]

of 32Articles in PresS. Am J Physiol Lung Cell Mol Physiol (August 10, 2007). doi:10.1152/ajplung.00203.2007

Copyright © 2007 by the American Physiological Society.

2

ABSTRACT

Acute hypoxia causes pulmonary vasoconstriction and coronary vasodilation. The

divergent effects of hypoxia on pulmonary and coronary vascular smooth muscle cells

suggest that the mechanisms involved in oxygen sensing and downstream effectors are

different in these two types of cells. Since production of reactive oxygen species (ROS) is

regulated by oxygen tension, ROS have been hypothesized to be a signaling mechanism

in hypoxia-induced pulmonary vasoconstriction and vascular remodeling. Furthermore,

an increased ROS production is also implicated in arteriosclerosis. In this study, we

determined and compared the effects of hypoxia on ROS levels in human pulmonary

arterial smooth muscle cells (PASMC) and coronary arterial smooth muscle cells

(CASMC). Our results indicated that acute exposure to hypoxia (PO2=25-30 mmHg for

5-10 min) significantly and rapidly decreased ROS levels in both PASMC and CASMC.

However, chronic exposure to hypoxia (PO2=30 mmHg for 48 hrs) markedly increased

ROS levels in PASMC, but decreased ROS production in CASMC. Furthermore, chronic

treatment with endothelin-1, a potent vasoconstrictor and mitogen, caused a significant

increase in ROS production in both PASMC and CASMC. The inhibitory effect of acute

hypoxia on ROS production in PASMC was also accelerated in cells chronically treated

with endothelin-1. While the decreased ROS in PASMC and CASMC after acute

exposure to hypoxia may reflect the lower level of oxygen substrate available for ROS

production, the increased ROS production in PASMC during chronic hypoxia may reflect

a pathophysiological response unique to the pulmonary vasculature that contributes to the

development of pulmonary vascular remodeling in patients with hypoxia-associated

pulmonary hypertension.

of 32

3

Key words: ROS; membrane potential; vascular smooth muscle cells.

of 32

4

INTRODUCTION

Intracellular production of reactive oxygen species (ROS), such as superoxide

(O·−) and hydrogen superoxide (HO−) formed from sequential transfer of electrons from

molecular oxygen, in vascular smooth muscle cells plays an important role in the

physiological regulation of vascular tone and vascular remodeling (20, 22). These

reactive intermediates act as progenitors for other forms of ROS such as hydrogen

peroxide (H2O2), peroxynitrite (ONOO–) and the hydroxyl radical. There are currently

debates as to the nature of the dysregulation of ROS production that may contribute to

pathophysiological conditions, such as pulmonary hypertension and atherosclerosis, and

the disturbance of normal vascular function. Unsurprisingly, the mechanisms by which

hypoxia causes pulmonary vessels to constrict and coronary vessels to relax are therefore

an active area of research.

It is now known that hypoxic pulmonary vasoconstriction (HPV) is an intrinsic

property of the pulmonary vasculature and specifically resides in pulmonary arterial

smooth muscle cells (PASMC) (31, 34). It has been reported that hypoxia inhibits

voltage-gated K+ (Kv) currents in PASMC (39, 58) and the hypoxia-induced closure of

Kv channels appears to result from a change in cytoplasmic ROS or redox status in

PASMC (4). Channels comprising the Kv1.5 subunit are implicated in such mechanisms

(6, 26, 38). Despite extensive research into the mechanisms of HPV, there is currently no

single mechanism that can entirely explain the phenomena (41, 53). How O2 levels are

detected in PASMC and the subsequent signaling pathways contributing to pulmonary

arterial constriction remain debated. Since their production is dependent on cellular

oxygen concentration and redox status, changes in ROS have been hypothesized to be an

of 32

5

important signaling mechanism in HPV. Potentially, changes in cellular ROS levels can

sequentially modulate ion channel function and lead to pulmonary vasoconstriction.

It has been indicated that nicotinamide adenine dinucleotide phosphate-oxidase

(NADPH oxidase) is an important source of increased O2·− in response to hypoxia (32);

however, an increase in mitochondrial derived ROS is also widely supported (50). This

opposes the original hypotheses that favors a decrease in ROS predominantly due to a

reduction of O2·− production resultant of the decreased O2 supply to the mitochondria (5,

33) and furthermore, other groups still dispute that ROS changes at all in response to

hypoxia (11). NADPH oxidases have materialized as the prominent source of ROS in the

vasculature and cardiovascular system (2, 9, 16). Vascular smooth muscle cells from

resistant arteries are known to contain NOX2, a non-phagocytic NADPH oxidase protein

that consists of a membrane-bound cytochrome b558 including the catalytic gp91phox and

the p22phox subunits, with cytosolic components including p47phox, p67phox, and the small

Rho guanosine triphosphatase (GTPase) Rac (25). The specific sub-cellular localization

of NADPH oxidase may be an important determinant of the fate of NADPH-derived ROS

production and ROS-dependent signaling pathways. Indeed, recently NADPH has been

identified to be bound to the β-subunit of Kv channels (18, 19, 36) the oxidation of

NADPH to NADP+ confers a structural change to the channel complex conferring to

acceleration of the channel inactivation kinetics. Such data consolidates a link between

ROS and K+ channels and thus the excitability of cells.

Notably, endothelin-1 (ET-1), which is shown to be upregulated during hypoxia

(29), has been demonstrated to mediate pulmonary vasoconstriction via NADPH oxidase-

derived superoxide and is an important mediator contributing to the pathogenesis of

of 32

6

chronic hypoxia-induced pulmonary hypertension (30, 51). ET-1 is additionally

implicated in the elevation of ROS in cardiac myocytes (57) via ETA receptor-mediated

regulation of NADPH oxidase activity and in NADPH oxidase dependent smooth muscle

cell proliferation (51). Another established activator of NADPH oxidases is angiotensin II

for which its incongruous effects are attributed to the excessive production of ROS (15,

40).

The objective of this study was therefore to investigate if there is a putative role

for hypoxia-induced elevation of ROS in the mechanisms of HPV by: a) determining if

and how ROS levels change during hypoxia in PASMC and b) comparing how hypoxia

affects ROS levels in pulmonary versus coronary vasculature, which vasoconstricts and

vasodilates in response to hypoxia, respectively.

of 32

7

MATERIALS AND METHODS

Cell preparation and culture. Cell lines of normal human PASMC and CASMC were

purchased from Clonetics (now Lonza Walkersville, Inc., MD). Cells were cultured in

75-cm2 flasks with smooth muscle growth medium (SmGM, Cambrex) supplemented

with 10% fetal bovine serum (FBS), 0.5 ng/ml human epidermal growth factor, 2 ng/ml

human fibroblast growth factor, 5 µg/ml insulin and 50 µg/ml gentamycin sulfate. Cells

were maintained in an incubator under a humidified atmosphere of 5% CO2-95% air at

37°C and used between passages 4 and 8. The medium was changed after 24 hrs and

every 48 hr thereafter. Cells were sub-cultured onto 25-mm2 cover slips using trypsin-

EDTA buffer (Cambrex) at 60% confluence and allowed to adhere for a minimum of 12

hrs. Cells were then fed with fresh media and grown for 24 hrs to allow recovery from

trypsinization before commencing experiments.

ROS detection. Intracellular ROS was detected via the dye dihydroethidium (DHE;

Molecular Probes). Unreacted DHE permeates live cells to accumulate as blue

fluorescence in the cytoplasm. DHE can be oxidized by superoxide to a specific

fluorescent product (ethidium/oxyethidium) which intercalates with DNA, accumulating

as red fluorescence in the nucleus. Oxyethidium has an additional oxygen atom in its

molecular structure compared to ethidium (59). Cells grown on cover slips were mounted

on a Nikon Eclipse TE-200 fluorescence microscope and superfused with normoxic

physiological salt solution (PSS) during the initial 30 sec of imaging. The PSS includes

(in mM): 141 NaCl, 4.7 KCl, 1.8 CaCl2, 1.2 MgCl2, 10 HEPES, and 10 glucose (pH 7.4).

Images were captured every 2 sec to minimize photobleaching on a Nikon Eclipse TE-

of 32

8

200 fluorescence microscope linked to a Photometrix digital camera and the average

fluorescence intensities for individual cells were quantified using NIH software ImageJ.

The fluorescence intensity of cells loaded with DHE is expressed as arbitrary units (a.u.)

and changes in the fluorescence are expressed as changes in arbitrary unit.

Hypoxic exposure. Acute hypoxia was achieved as described previously (58). Briefly,

cells were incubated with 80 µM DHE in the medium at 37°C for 15 min prior to hypoxic

treatment. Cells were perfused with normoxic PSS during the initial 30 sec of imaging.

Hypoxic conditions were established by bubbling the superfusion solution with N2

(100%) for 15 min to achieve oxygen tension (PO2) ranging from 25-30 mmHg at 24°C.

PO2 was quantified using an oxygen electrode (Microelectrodes, Londonderry, NH)

positioned in the cell chamber on the microscope stage to continuously monitor the PO2

as previously shown (58). For the chronic hypoxic experiments, cells cultured in serum-

free smooth muscle basal medium (SmBM, Cambrex) were incubated in a chamber for

48 hrs at 37°C in normoxia (PO2=140 mmHg) or hypoxia (PO2=30 mmHg). 80 µM DHE

was added to the medium and also incubated for 15 min prior to imaging. In the positive

control experiments, PASMC and CASMC were treated with 200 nM angiotensin II for 8

hrs to induce superoxide production (15). 100 U/ml polyethylene glycol-superoxide

dismutase (SOD) was used to catalyze the conversion of superoxide to H2O2, verifying

the fluorescence signal as superoxide (13). In experiments using 1 µM ET-1, cells were

incubated for 48 hrs.

of 32

9

Statistical analysis. The combined data are expressed as mean±SE. Statistical analysis

was performed using the unpaired Student's t test. Statistical differences were deemed to

be significant at P<0.05.

of 32

10

RESULTS

To validate the protocol for the detection of intracellular changes in the oxidative state in

PASMC using DHE, positive controls were established. Over a period of 15 min,

oxidation of DHE was observed by the change in the predominantly cytosolic blue DHE

fluorescence to red ethidium/oxyethidium fluorescence concentrated in the nucleus

representative of the basal ROS concentration in the cell (Fig. 1A). The dye was therefore

left to equilibrate for 15 min preceding the commencement of any experiments. Changes

in fluorescence intensity in a single PASMC after treatment were used in this study to

demonstrate an increase or a decrease of ROS production in response to different stimuli.

Using this method, we found that extracellular application of angiotensin II (A-II)

significantly increased ROS production in CASMC, while SOD partially inhibited the A-

II-mediated increase in ROS level (Fig. 1B).

Acute hypoxia significantly reduced superoxide production in human PASMC and

CASMC. Acute hypoxia, established by superfusing with N2-equilibrated PSS (PO2=30

mmHg), rapidly reduced ROS production in both PASMC (n=14) and CASMC (n=30)

(Fig. 2A & B). The decreased intensity over a 14-min period is expressed as raw whole-

cell fluorescence (left panels) and normalized to the basal fluorescence at time 0 min

(right panels) for PASMC (Fig. 2A) and CASMC (Fig. 2B) to facilitate comparison

between the cell types. The average kinetic profiles of the acute hypoxia-induced

decreases in fluorescence intensity were comparable in both PASMC and CASMC (Fig.

2C).

of 32

11

ET-1, an endothelium-derived contracting factor proposed to be an important

trigger in the development of HPV and hypoxia-associated pulmonary hypertension (30),

is also known to stimulate intracellular superoxide production (51). A significant increase

in PASMC ROS was stimulated by chronic treatment (48 hrs) with ET-1 (from 28.9±2.6

a.u., n=14 in control to 38.3±1.7 a.u., n=31 after chronic ET-1 treatment, P<0.01) (Fig.

3A & B). Subsequent exposure to acute hypoxia caused a significant (P<0.001) time-

dependent decrease in ROS levels in ET-1-treated PASMC (in which superoxide

concentration was higher in both the nuclei and cytosol) (Fig. 3A & C-F). This decrease

occurred at a significantly enhanced rate when compared to control (Fig. 3D). These

results consolidate that acute hypoxia reduces the level of ROS in PASMC, either under

basal or physiological conditions, or under pathophysiological conditions when

superoxide production is increased by ET-1. On the other hand, ET-1 had no effect upon

ROS production in CASMC with only a small insignificant decrease in whole cell

fluorescence observed (data not shown).

Chronic hypoxia significantly increases ROS levels in human PASMC, but decreases

ROS levels in human CASMC. Exposure of PASMC to chronic hypoxia (CH, 48 hrs),

however, significantly increased superoxide production (27.6±1.2 a.u., n=111, control;

47.1±3.0 a.u., n=49, CH; P<0.01) (Fig. 4A). The increase in ROS can be accredited to the

observed rightward shift to increased values in the average fluorescent intensity, as

indicated in the histograms featured in Figure 4B. In addition, 48-hr CH treatment of

PASMC also blocked the ET-1-mediated increase in ROS production that was observed

in the normoxic control (Fig. 4C). It is worth noting that the mean whole cell

of 32

12

fluorescence was significantly reduced after chronic ET-1 treatment in CH (32.1±1.7 a.u.,

n=110) when compared to the normoxic control (39.8±1.7 a.u., n=105, P<0.01) (Fig. 4C).

However, the addition of SOD in conjunction with ET-1, catalyzing the formation of

H2O2 from superoxide and which is known to prevent the formation of oxyethidium from

the interaction of DHE and superoxide (59), reduces whole cell fluorescence to similar

levels in both normoxic conditions (18.8±3.0 a.u., n=120) and after CH treatment

(21.3±1.0 a.u., n=100) (P<0.01 for both when compared to normoxic control).

On the contrary, incubation in CH decreased ROS production in CASMC from

39.1±2.9 a.u., n=57, in normoxia to 27.7±1.3 a.u., n=77, in CH; P<0.01) (Fig. 5A and B).

Moreover, ET-1 had no significant effect on superoxide production in either normoxic or

hypoxic condition in CASMC (Fig. 5C), dissimilar to PASMC where ET-1 significantly

increased superoxide (Fig. 4). Angiotensin II (A-II) is a known activator of superoxide

production in CASMC (15), incubation with A-II instead of ET-1 did cause a significant

increase in mean whole cell fluorescence (18.6±1.2 a.u. and 34.9±2.8 a.u., in the absence

and presence of A-II, respectively, P<0.001) and this increase included a substantial SOD

dependent component reflecting the catalysis of reactive intermediates such as superoxide

to more stable H2O2 (also see Fig. 1C) (59). Interestingly, SOD did significantly decrease

whole cell fluorescence in both normoxic and hypoxic conditions in comparison to the

control fluorescence (P<0.01), indicative of a decrease in superoxide from basal/control

levels.

of 32

13

DISCUSSION

A putative involvement of increased ROS in CH-induced pulmonary hypertension

is consolidated in the study where the data indicate that: a) chronic hypoxia induced

opposite effects on superoxide production in PASMC and CASMC; b) agonist ET-1 only

increased superoxide levels in PASMC, but not in CASMC. The differences in

superoxide production in response to hypoxia (and ET-1 treatment) between the

pulmonary and coronary circulation may have significant physiological implications in

terms of the effect of acute and chronic hypoxia on the contractility of different vascular

beds. Furthermore, the data also provide evidence for a differential regulation on

superoxide production during acute and chronic hypoxia in PASMC: acute hypoxia

decreases basal and agonist (ET-1)-induced superoxide production, whereas chronic

hypoxia increased superoxide production and additionally blocked ET-1-mediated

increases in superoxide.

Acute hypoxia reduces superoxide in both PASMC and CASMC. The data from this

study indicate that a decrease in superoxide production is involved in acute hypoxia-

induced pulmonary vasoconstriction. The change in fluorescence in our experiments can

be largely attributed to altered superoxide production converting DHE to oxyethidium,

though ethidium is also produced reflecting the more general cellular redox state (59).

The production of oxyethidium and subsequent change in fluorescence is known to be

inhibited by SOD and is not sensitive to changes in H2O2 concentration; therefore the

SOD-dependent component of DHE fluorescence can be attributed to a single electron

reduction of molecular oxygen to superoxide (13). In our experiments, the change in ROS

of 32

14

detected by DHE oxidation should therefore have a SOD sensitive component more

selective for superoxide and SOD insensitive component likely to be H2O2 dependent

(12).

While the debate continues over decreased (52) or a paradoxical increase in ROS

(24, 48) during hypoxia, it is possible that ROS are not involved in acute HPV. In our

experiments, acute hypoxia decreased superoxide in both PASMC and CASMC whilst it

is known to induce pulmonary vasoconstriction but coronary vasodilatation. This notion

is strengthened by, as it is previously reported, that increased ROS takes place over

several minutes whilst the onset of HPV occurs within 7 seconds (27). Although more

evidence is needed, it is possible that decreased superoxide triggers a mechanism causing

PASMC contraction (e.g., via inhibition of Kv channels and/or activation of transient

receptor potential cation channels), but causes CASMC relaxation [e.g., via increased

cGMP and nitric oxide (NO) (42), Ca2+ desensitization (49), and/or a lowering of

cytosolic NADPH levels (21)].

Previous work in our laboratory, supported by others, has demonstrated a

decrease in Kv1.5 currents in response to acute hypoxia specific to the pulmonary

circulation or PASMC; no response was observed in the mesenteric arterial SMCs (26,

37, 58), which is also difficult to attribute to the similar decreased superoxide production

in PASMC and CASMC, observed in this paper. However, more recently, a Kv channel β

subunit that resembles a functional NADPH-dependent enzyme to catalyze redox

reactions has been identified (8, 36, 55). It is possible that the presence of this β subunit

in PASMC associated with the “hypoxic-sensitive” Kv channel α subunits may account

for the selectivity of the effect of hypoxia in pulmonary but not systemic (or coronary)

of 32

15

arteries. On the other hand, it is also possible, and perhaps more likely, that the acute

hypoxic changes in superoxide are not directly linked to the contractility of the

pulmonary vessels. Acute hypoxia is also known to increase cytosolic free Ca2+

concentration ([Ca2+]cyt) in PASMC (but not in aortic SMC) via capacitative Ca2+ entry

(CCE) through transient receptor potential (TRP) cation channels, a mechanism that may

or may not be dependent on cellular redox state (35, 45). The decrease in superoxide in

response to hypoxia most likely reflects the lower level of oxygen available as a substrate

for oxygen radical production and leading to a reduced cellular redox state. This is

consistent with the expected chemical effects of lower oxygen tension on ROS levels.

Clarification of these mechanisms specific to HPV requires further experimentation.

Chronic hypoxia divergently affects ROS production in PASMC and CASMC. The

increased superoxide in PASMC during chronic hypoxia may, however, reflect a

pathophysiological response unique to pulmonary circulation that contributes to sustained

pulmonary vasoconstriction and excessive pulmonary vascular remodeling in patients

with hypoxia-associated pulmonary hypertension as such increased levels were not

observed in CASMC (Figs. 4 & 5).

There is a large body of evidence supporting an increase in PASMC ROS during

chronic hypoxia (32, 50). In support of our data, a FRET-based intracellular sensor has

recently shown a hypoxia-induced increase in ROS (23). It is proposed that this increase

in ROS is unlikely to act directly on Kv channels, but may influence their activity or

expression indirectly by regulating ET-1 signaling (47). Endothelin-1, which is shown to

be upregulated during hypoxia (29), has been demonstrated to mediate pulmonary

of 32

16

vasoconstriction via NADPH oxidase-derived superoxide and is importantly proposed to

contribute to HPV and the pathogenesis CH-induced pulmonary hypertension (30, 51).

ET-1 also stimulates PASMC proliferation, an effect likely to be caused by its

augmentation of ROS production (51), which may contribute to the vascular remodeling

observed in pulmonary hypertension. Furthermore, in ovine fetal PAMSC (17), inhibition

of NADPH oxidase induces apoptosis and prevents ET-1-mediated SMC proliferation. In

our experiments, ET-1 did increase superoxide concentration, mimicking the effects

observed with chronic hypoxia in PASMC. The opposite effects was observed in

CASMC, which cannot be attributed to damage to cells or intracellular mechanisms as A-

II was still able to increase ROS in these cells (Fig. 5D). Therefore, increased ET-1 by

hypoxia has very different effects on ROS in coronary and pulmonary vascular smooth

muscle cells and this, in part, may account for the opposite responses (coronary

vasodilation vs. pulmonary vasoconstriction) to CH. Moreover, differences between

systemic and pulmonary artery ROS/superoxide production in response to sustained

hypoxia may be due to differential regulation of NADPH oxidases (or there may be

differences in SOD activity). In pulmonary artery, both NOX1 and NOX4 are present

(10) and superoxide production by these NOX isoforms is dependent upon direct

interaction with p22phox (3). p22phox-based NADH/NADPH oxidase is also implicated in

the pathogenesis of atherosclerotic coronary artery disease (7, 56). However, it is possible

that similar interaction is differentially regulated by hypoxia in the coronary and

pulmonary circulation (or CASMC and PASMC). Both coronary and pulmonary arteries

have been shown to express NOX isoforms at similar levels, however increased

superoxide production may be due to higher basal cytosolic NADPH in pulmonary

of 32

17

arteries (21) or the subcellular localization of NOX isoforms may be crucial to the

activation of downstream signaling in response to increased ROS. Such pathways remain

to be fully defined.

Other possibilities exist for the divergent effects of hypoxia on ROS and vascular

function. For example, TRP expression and function are also known to be upregulated in

PASMC during chronic hypoxia. Effects may, in part, be attributed to the increased

superoxide production that we and others have observed. Firstly, ROS are known to

activate adenylate cycalse that can increase cADPR, activate ryanodine receptors and

deplete intracellular Ca2+ stores (e.g., sarcoplasmic/endoplasmic reticulum); sustained

store depletion then activate store-operated TRP channels (notably TRPC 1, 4 and 6) and

cause increase in [Ca2+]cyt (44). Furthermore, increased TRP channel expression induced

by hypoxia can be regulated by hypoxia-inducible factor -1 (HIF-1) an hypoxia-induced

transcription factor which is known to be upregulated by increased superoxide (1, 46) and

may, therefore, have profound effects on hypoxia-associated pulmonary hypertension. In

addition, chronic hypoxia-mediated pulmonary vascular remodeling may at least partly

result from (or relate to) the observed increase in production of superoxide; this may

subsequently activate the MAPK pathway and cause cell proliferation, as reported in

many cell types such as; human, rat and mouse pulmonary, but not systemic, fibroblasts

(54); alveolar epithelial cells (43) and cancerous cells (14). Additionally, p38 MAP

kinase plays an important role in mediating hypoxia-induced contraction of the rat PA

(28).

of 32

18

Concluding remarks. Amidst an active debate on the “ups and downs” of ROS during

hypoxia, the data presented in this paper provide a possible explanation for the regulation

and involvement of ROS in hypoxia. During acute hypoxic exposure, ROS, particularly

superoxide, decreases in both coronary and pulmonary vascular smooth muscle cells and

is thus unlikely to highly influential over the vessel contractility in both circulations that

are known to dilate and constrict to hypoxia, respectively. However, superoxide

regulation during more sustained, chronic periods of hypoxia is differentially regulated;

increasing in PASMC and decreasing in CASMC. The increased ROS production in

PASMC during chronic hypoxia may indeed reflect a pathophysiological response unique

to pulmonary circulation that contributes to hypoxia-induced pulmonary vasoconstriction

and vascular remodeling observed in patients with chronic hypoxia-mediated pulmonary

hypertension.

of 32

19

Acknowledgment

We thank Ann Nicholson, M.S. for her technical assistance. This work was supported in

part by grants from the National Heart, Lung, and Blood Institute of the National

Institutes of Health (HL054043, HL064945, and HL066012).

of 32

20

References

1. Aaronson PI. TRPC channel upregulation in chronically hypoxic pulmonary arteries: the HIF-1 bandwagon gathers steam. Circ Res 98: 1465-1467, 2006.

2. Al-Mehdi A, Zhao G, Dodia C, Tozawa K, Costa K, Muzykantov V, Ross C, Blecha F, Dinauer MC, and Fisher AB. Endothelial NADPH oxidase as the source of oxidants in lungs exposed to ischemia or high K+. Circ Res 83: 730-737, 1998.

3. Ambasta RK, Kumar P, Griendling KK, Schmidt HHHW, Busse R, and Brandes RP. Direct interaction of the novel Nox proteins with p22phox is required for the formation of a functionally active NADPH oxidase. J Biol Chem 279: 45935-45941, 2004.

4. Archer SL, Huang J, Henry T, Peterson D, and Weir EK. A redox-based O2sensor in rat pulmonary vasculature. Circ Res 73: 1100-1112, 1993.

5. Archer SL, Reeve HL, Michelakis E, Puttagunta L, Waite R, Nelson DP, Dinauer MC, and Weir EK. O2 sensing is preserved in mice lacking the gp91 phox subunit of NADPH oxidase. Proc Natl Acad Sci USA 96: 7944-7949, 1999.

6. Archer SL, Souil E, Dinh-Xuan AT, Schremmer B, Mercier JC, El Yaagoubi A, Nguyen-Huu L, Reeve HL, and Hampl V. Molecular identification of the role of voltage-gated K+ channels, Kv1.5 and Kv1.2, in hypoxic pulmonary vasoconstriction and control of resting membrane potential in rat pulmonary artery myocytes. J Clin Invest 101: 2319-2330, 1998.

7. Azumi H, Inoue N, Ohashi Y, Terashima M, Mori T, Fujita H, Awano K, Kobayashi K, Maeda K, Hata K, Shinke T, Kobayashi S, Hirata K-i, Kawashima S, Itabe H, Hayashi Y, Imajoh-Ohmi S, Itoh H, and Yokoyama M. Superoxide generation in directional coronary atherectomy specimens of patients with angina pectoris: Important role of NAD(P)H oxidase. Arterioscler Thromb Vasc Biol 22: 1838-1844, 2002.

8. Bahring R, Milligan CJ, Vardanyan V, Engeland B, Young BA, Dannenberg J, Waldschutz R, Edwards JP, Wray D, and Pongs O. Coupling of voltage-dependent potassium channel inactivation and oxidoreductase active site of Kvβsubunits. J Biol Chem 276: 22923-22929, 2001.

9. Cai H and Harrison DG. Endothelial dysfunction in cardiovascular diseases: the role of oxidant stress. Circ Res 87: 840-844, 2000.

10. Clempus RE, Sorescu D, Dikalova AE, Pounkova L, Jo P, Sorescu GP, Lassegue B, and Griendling KK. Nox4 is required for maintenance of the differentiated vascular smooth muscle cell phenotype. Arterioscler Thromb Vasc Biol 27: 42-48, 2007.

11. Evans AM, Mustard KJW, Wyatt CN, Peers C, Dipp M, Kumar P, Kinnear NP, and Hardie DG. Does AMP-activated protein kinase couple inhibition of mitochondrial oxidative phosphorylation by hypoxia to calcium signaling in O2-sensing cells? J Biol Chem 280: 41504-41511, 2005.

12. Fernandes DC, Wosniak J, Jr., Pescatore LA, Bertoline MA, Liberman M, Laurindo FRM, and Santos CXC. Analysis of DHE-derived oxidation products by HPLC in the assessment of superoxide production and NADPH oxidase activity in vascular systems. Am J Physiol Cell Physiol 292: C413-422, 2007.

of 32

21

13. Fink B, Laude K, McCann L, Doughan A, Harrison DG, and Dikalov S. Detection of intracellular superoxide formation in endothelial cells and intact tissues using dihydroethidium and an HPLC-based assay. Am J Physiol Cell Physiol 287: C895-902, 2004.

14. Giles GI. The redox regulation of thiol dependent signaling pathways in cancer. Curr Pharm Des 12: 4427-4443.

15. Griendling KK, Minieri CA, Ollerenshaw JD, and Alexander RW. Angiotensin II stimulates NADH and NADPH oxidase activity in cultured vascular smooth muscle cells. Circ Res 74: 1141-1148, 1994.

16. Griendling KK and Ushio-Fukai M. NADH/NADPH oxidase and vascular function. Trends Cardiovasc Med 7: 301-307, 1997.

17. Grobe AC, Wells SM, Benavidez E, Oishi P, Azakie A, Fineman JR, and Black SM. Increased oxidative stress in lambs with increased pulmonary blood flow and pulmonary hypertension: role of NADPH oxidase and endothelial NO synthase. Am J Physiol Lung Cell Mol Physiol 290: L1069-L1077, 2006.

18. Gulbis JM, Mann S, and MacKinnon R. Structure of a voltage-dependent K+

channel β subunit. Cell 97: 943-952, 1999. 19. Gulbis JM, Zhou M, Mann S, and MacKinnon R. Structure of the cytoplasmic

beta subunit-T1 assembly of voltage-dependent K+ channels. Science 289: 123-127, 2000.

20. Gupte SA, Rupawalla T, Phillibert D, Jr., and Wolin MS. NADPH and heme redox modulate pulmonary artery relaxation and guanylate cyclase activation by NO. Am J Physiol 277: L1124-1132, 1999.

21. Gupte SA and Wolin MS. Hypoxia promotes relaxation of bovine coronary arteries through lowering cytosolic NADPH. Am J Physiol Heart Circ Physiol 290: H2228-H2238, 2006.

22. Gutterman DD. Mitochondria and reactive oxygen species: an evolution in function. Circ Res 97: 302-304, 2005.

23. Guzy RD, Hoyos B, Robin E, Chen H, Liu L, Mansfield KD, Simon MC, Hammerling U, and Schumacker PT. Mitochondrial complex III is required for hypoxia-induced ROS production and cellular oxygen sensing. Cell Metab 1: 401-408, 2005.

24. Guzy RD and Schumacker PT. Oxygen sensing by mitochondria at complex III: the paradox of increased reactive oxygen species during hypoxia. Exp Physiol 91: 807-819, 2006.

25. Hordijk PL. Regulation of NADPH Oxidases: The Role of Rac Proteins. Circ Res 98: 453-462, 2006.

26. Hulme JT, Coppock EA, Felipe A, Martens JR, and Tamkun MM. Oxygen sensitivity of cloned voltage-gated K+ channels expressed in the pulmonary vasculature. Circ Res 85: 489-497, 1999.

27. Jensen KS, Micco AJ, Czartolomna J, Latham L, and Voelkel NF. Rapid onset of hypoxic vasoconstriction in isolated lungs. J Appl Physiol 72: 2018-2023, 1992.

28. Karamsetty MR, Klinger JR, and Hill NS. Evidence for the role of p38 MAP kinase in hypoxia-induced pulmonary vasoconstriction. Am J Physiol Lung Cell Mol Physiol 283: L859-866, 2002.

of 32

22

29. Li H, Chen SJ, Chen YF, Meng QC, Durand J, Oparil S, and Elton TS. Enhanced endothelin-1 and endothelin receptor gene expression in chronic hypoxia. J Appl Physiol 77: 1451-1459, 1994.

30. Liu JQ, Zelko IN, Erbynn EM, Sham JSK, and Folz RJ. Hypoxic pulmonary hypertension: role of superoxide and NADPH oxidase (gp91phox). Am J Physiol Lung Cell Mol Physiol 290: L2-L10, 2006.

31. Madden JA, Vadula KS, and Kurup VP. Effects of hypoxia and other vasoactive agents on pulmonary and cerebral artery smooth muscle cells. Am J Physiol 263: L384-L393, 1992.

32. Marshall C, Mamary AJ, Verhoeven AJ, and Marshall BE. Pulmonary artery NADPH-oxidase is activated in hypoxic pulmonary vasoconstriction. Am J Respir Cell Mol Biol 15: 633-644, 1996.

33. Mohazzab KM, Fayngersh RP, Kaminski PM, and Wolin MS. Potential role of NADH oxidoreductase-derived reactive O2 species in calf pulmonary arterial PO2-elicited responses. Am J Physiol 269: L637-644, 1995.

34. Murray TR, Chen L, Marshall BE, and Macarak EJ. Hypoxic contraction of cultured pulmonary vascular smooth muscle cells. Am J Respir Cell Mol Biol 3: 457-465, 1990.

35. Ng LC, Wilson SM, and Hume JR. Mobilization of sarcoplasmic reticulum stores by hypoxia leads to consequent activation of capacitative Ca2+ entry in isolated canine pulmonary arterial smooth muscle cells. J Physiol (Lond) 563: 409-419, 2005.

36. Peri R, Wible BA, and Brown AM. Mutations in the Kvbeta 2 Binding Site for NADPH and Their Effects on Kv1.4. J Biol Chem 276: 738-741, 2001.

37. Platoshyn O, Brevnova EE, Burg ED, Yu Y, Remillard CV, and Yuan JX-J. Acute hypoxia selectively inhibits KCNA5 channels in pulmonary artery smooth muscle cells. Am J Physiol Cell Physiol 290: C907-C916, 2006.

38. Platoshyn O, Yu Y, Golovina VA, McDaniel SS, Krick S, Li L, Wang JY, Rubin LJ, and Yuan JX-J. Chronic hypoxia decreases KV channel expression and function in pulmonary artery myocytes. Am J Physiol Lung Cell Mol Physiol 280: L801-L812, 2001.

39. Post JM, Hume JR, Archer SL, and Weir EK. Direct role for potassium channel inhibition in hypoxic pulmonary vasoconstriction. Am J Physiol 262: C882-C890, 1992.

40. Rajagopalan S, Kurz S, Münzel T, Tarpey M, Freeman BA, Griendling KK, and Harrison DG. Angiotensin II-mediated hypertension in the rat increases vascular superoxide production via membrane NADH/NADPH oxidase activation. Contribution to alterations of vasomotor tone. J Clin Invest 97: 1916-1923, 1996.

41. Robertson TP, Aaronson PI, and Ward JPT. Hypoxic vasoconstriction and intracellular Ca2+ in pulmonary arteries: evidence for PKC-independent Ca2+ sensitization. Am J Physiol 268: H301-H307, 1995.

42. Sato A, Sakuma I, and Gutterman DD. Mechanism of dilation to reactive oxygen species in human coronary arterioles. Am J Physiol Heart Circ Physiol 285: H2345-2354, 2003.

of 32

23

43. Sigaud S, Evelson P, and Gonzalez-Flecha B. H2O2-induced proliferation of primary alveolar epithelial cells is mediated by MAP kinases. Antioxidants Redox Signal 7: 6-13, 2005.

44. Wang J, Shimoda LA, and Sylvester JT. Capacitative calcium entry and TRPC channel proteins are expressed in rat distal pulmonary arterial smooth muscle. Am J Physiol Lung Cell Mol Physiol 286: L848-L858, 2004.

45. Wang J, Shimoda LA, Weigand L, Wang W, Sun D, and Sylvester JT. Acute hypoxia increases intracellular [Ca2+] in pulmonary arterial smooth muscle by enhancing capacitative Ca2+ entry. Am J Physiol Lung Cell Mol Physiol 288: L1059-L1069, 2005.

46. Wang J, Weigand L, Lu W, Sylvester JT, Semenza GL, and Shimoda LA. Hypoxia inducible factor 1 mediates hypoxia-induced TRPC expression and elevated intracellular Ca2+ in pulmonary arterial smooth muscle cells. Circ Res 98: 1528-1537, 2006.

47. Wang X, Tong M, Chinta S, Raj JU, and Gao Y. Hypoxia-induced reactive oxygen species downregulate ETB receptor-mediated contraction of rat pulmonary arteries. Am J Physiol Lung Cell Mol Physiol 290: L570-L578, 2006.

48. Ward JP. Point: Hypoxic pulmonary vasoconstriction is mediated by increased production of reactive oxygen species. J Appl Physiol 101: 993-995, 2006.

49. Wardle RL, Gu M, Ishida Y, and Paul RJ. Ca2+-desensitizing hypoxic vasorelaxation: pivotal role for the myosin binding subunit of myosin phosphatase (MYPT1) in porcine coronary artery. J Physiol 572: 259-267, 2006.

50. Waypa GB, Chandel NS, and Schumacker PT. Model for hypoxic pulmonary vasoconstriction involving mitochondrial oxygen sensing. Circ Res 88: 1259-1266, 2001.

51. Wedgwood W, Dettman RW, and Black SM. ET-1 stimulates pulmonary arterial smooth muscle cell proliferation via induction of reactive oxygen species. Am J Physiol Lung Cell Mol Physiol 281: L1058-L1067, 2001.

52. Weir EK and Archer SL. Counterpoint: Hypoxic pulmonary vasoconstriction is not mediated by increased production of reactive oxygen species. J Appl Physiol 101: 995-998, 2006.

53. Weissmann N, Sommer N, Schermuly RT, Ghofrani HA, Seeger W, and Grimminger F. Oxygen sensors in hypoxic pulmonary vasoconstriction. Cardiovas Res 71: 620-629, 2006.

54. Welsh D, Mortimer H, Kirk A, and Peacock A. The role of p38 mitogen-activated protein kinase in hypoxia-induced vascular cell proliferation: An interspecies comparison. Chest 128: 573S-574, 2005.

55. Weng J, Cao Y, Moss N, and Zhou M. Modulation of voltage-dependent Shaker family potassium channels by an aldo-keto reductase. J Biol Chem 281: 15194-15200, 2006.

56. Wolin MS. How could a genetic variant of the p22phox component of NAD(P)H oxidases contribute to the progression of coronary atherosclerosis? Circ Res 86: 365-366, 2000.

57. Xu FP, Chen MS, Wang YZ, Yi Q, Lin SB, Chen AF, and Luo JD. Leptin induces hypertrophy via endothelin-1-reactive oxygen species pathway in cultured neonatal rat cardiomyocytes. Circulation 110: 1269-1275, 2004.

of 32

24

58. Yuan XJ, Goldman WF, Tod ML, Rubin LJ, and Blaustein MP. Hypoxia reduces potassium currents in cultured rat pulmonary but not mesenteric arterial myocytes. Am J Physiol 264: L116-L123, 1993.

59. Zhao H, Kalivendi S, Zhang H, Joseph J, Nithipatikom K, Vasquez-Vivar J, and Kalyanaraman B. Superoxide reacts with hydroethidine but forms a fluorescent product that is distinctly different from ethidium: potential implications in intracellular fluorescence detection of superoxide. Free Rad Biol Med 34: 1359-1368, 2003.

of 32

25

Figure Legends:

Fig. 1. Verification of using DHE to measure changes in intracellular ROS production or

ROS levels in PASMC and CASMC. A: Representative PASMC showing cytosolic blue

DHE fluorescence and, after 15 min, the DHE has been oxidized by ROS to be visible as

red fluorescence predominantly localized in the nucleus where the oxidized

ethidium/oxyethidium intercalated with DNA. The level of oxidized

ethidium/oxyethidium fluorescence after 15 min was considered to be basal ROS levels

and corresponds to time point 0 min in all experiments. Overlay of the left and right

panels showing co-localization of the respective fluorescence. B: Summarized data (mean

±SE) shown mean whole-cell fluorescence in control CASMC (open bar, n=28) and

CASMC treated with angiotensin-II (A-II, solid bar, n=27) and A-II plus SOD (gray bar,

n=21). *** P<0.001 vs. control bar.

Fig. 2. Acute hypoxia decreases ROS in both PASMC and CASMC. Averaged, time-

dependent decrease in whole cell fluorescence (left panel) and normalized to basal

fluorescence in normoxia for comparison (right panel) in PASMC (A, n=14) and CASMC

(B, n=30). C: Comparison of the normalized average kinetics of the acute hypoxia-

mediated decrease in ROS over 15 min in PASMC (red) and CASMC (blue).

Fig. 3. Chronic ET-1 treatment increases basal ROS production in PASMC. A:

Representative image of a PASMC under basal condition, after chronic (48 hr) exposure

to 1µM ET-1 and after a subsequent acute hypoxic episode (depicted left to right). B: ET-

1 increases PASMC ROS production indicated by increased whole cell fluorescence (**

of 32

26

P<0.01 vs. control, n=111 and 105 in control and ET-1 treated, respectively). Time-

dependent kinetics of the decrease in whole cell fluorescence (C, n=14 in control and

n=31 ET-1 treated) and normalized fluorescence (D, n=14 in control and n=31 ET-1

treated) in response to acute (15 min) hypoxia in PASMC. The effect of acute hypoxia

after ET-1 treatment is summarized for whole cell fluorescence and normalized

fluorescence in E and F, respectively (n=31 in normoxia and n=31 in hypoxia).

Significant changes compared to control (basal) levels are indicated as follows: **

P<0.01 and *** P<0.001 vs. Normoxia bars.

Fig. 4. Chronic hypoxia increases ROS production in PASMC. A: Representative image

of PASMC on a cover slip in basal (control) (top left), after chronic hypoxia (top right),

in normoxia after chronic ET-1 treatment (bottom left) and in normoxia after ET-1 and

SOD treatment (bottom right). B: Histograms reflecting the spread of individual cell

fluorescence in normoxia (top, n=111) and after chronic hypoxia (bottom, n=49). C:

Comparison of the effect of both ET-1 and SOD on ROS production in normoxic (left,

n=111, 105 and 120 for control, ET-1 and ET-1 + SOD, respectively) and chronic

hypoxia (right, n=49, 110 and 100 for control, ET-1 and ET-1 + SOD, respectively)

conditions. The dashed line indicates the basal fluorescence at the start of the

experiments. Significant changes compared to control (basal) levels are indicated as

follows: ** P<0.01 and *** P<0.001 vs. normoxic control.

Fig. 5. Chronic hypoxia decreases ROS production in CASMC. A: Representative image

of CASMC on a cover slip in basal (control), after chronic hypoxia, in normoxia after

of 32

27

chronic ET-1 treatment and in normoxia after ET-1 and SOD treatment (depicted

clockwise from top left). B: Histograms showing the spread of individual cell

fluorescence in normoxia (top, n=57) and after CH (bottom, n=82). C: Comparison of the

effect of both ET-1 and SOD on ROS production in normoxic (left, n=57, 72 and 38 for

control, ET-1 and ET-1 + SOD, respectively) and chronic hypoxia (right, n=82, 77 and

64 for control, ET-1 and ET-1 + SOD, respectively) conditions. The dashed line indicates

the basal fluorescence at the start of the experiments. Significant changes compared to

control (basal) levels are indicated as follows: ** P<0.01 and *** P<0.001 vs. normoxic

control.

of 32

Mea

n W

hole

-cel

lFl

uore

scen

ce

0

10

20

30

40

Control A-II A-II+SOD

***

Time (min)0 3 6 9 12 15

Fluo

resc

ence

16

18

20

22

24

26

28

Overlay

Cytosolic dihydroethidium Oxidized to ethidium

Fig. 1

A

PASMC

C

B

CASMC

of 32

Time in Hypoxia (min)0 2 4 6 8 10 12 14

Raw

Who

le-c

ell

Fluo

resc

ence

20

25

30

35


Nor

mal

ized

Who

le-c

ell

Fluo

resc

ence

(F/F

0)

0.7

0.8

0.9

1.0

PASMC

CASMC


Nor

mal

ized

Fluo

resc

ence

(F/F

0)

0.7

0.8

0.9

1.0


Raw

Who

le-c

ell

Fluo

resc

ence

20

25

30

35


Nor

mal

ized

Fluo

resc

ence

(F/F

0)

0.7

0.8

0.9

1.0

Fig. 2

(PASMC)A

(CASMC)B

C

of 32

Time in Hypoxia(min)

0 3 6 9 12 15

Raw

Who

le-c

ell

Fluo

resc

ence

20

25

30

35

40

ControlPASMC

ET-1-treated PASMC

Time in Hypoxia(min)

0 3 6 9 12 15

Mea

n N

orm

aliz

edFl

uore

scen

ce (F

/F0)

0.6

0.7

0.8

0.9

1.0

Raw

Who

le-c

ell

Fluo

resc

ence

20

25

30

35

40

Cont ET-1

**

PASMC

ET-1-treated PASMC

ControlPASMC

Raw

Who

le-c

ell

Fluo

resc

ence

0

10

20

30

40

***M

ean

Nor

mal

ized

Fluo

resc

ence

(F/F

0)

0.0

0.5

1.0

***

Nor Hyp Nor HypET-1-treated

PASMCET-1-treated

PASMC

B C D

E F

Fig. 3

A

PASMC

Normoxia Control ET-1 (48 hrs) ET-1+Hypoxia

of 32

Mea

n W

hole

-cel

lFl

uore

scen

ce

0

10

20

30

40

50

Cont ET-1 ET-1+SOD

Normoxia

Cont ET-1 ET-1+SOD

Hypoxia (48 h)

**

**

**

**

Fig. 4

Cel

l Num

ber

0369

12151821

Fluorescence Intensity0 10 20 30 40 50 60 70 80 90

Cel

l Num

ber

01234567

Normoxia control

Hypoxia (48 hrs)

Normoxia Control Hypoxia (48 hrs)

PASMC

Normoxia+ET-1+SODNormoxia+ET-1

A

B

C

of 32

Mea

n W

hole

-cel

lFl

uore

scen

ce

0

10

20

30

40

50

****

**

Cont ET-1 ET-1+SOD

Normoxia

Cont ET-1 ET-1+SOD

Hypoxia (48 h)

Cel

l Num

ber

012345678

Fluorescence Intensity0 10 20 30 40 50 60 70 80 90

Cel

l Num

ber

0369

121518

Normoxia control

Hypoxia (48 hrs)

Fig. 5

Normoxia Control Hypoxia (48 hrs)

CASMC

Normoxia+ET-1+SODNormoxia+ET-1

A

B

C

of 32

Hypoxia Divergently Regulates Production of Reactive Oxygen Species in Human Pulmonary and Coronary...

Documents

Transcript of Hypoxia Divergently Regulates Production of Reactive Oxygen Species in Human Pulmonary and Coronary...