A role for zinc in regulating hypoxia-induced contractile events
in pulmonary endothelium
Paula J. Bernal,*1Eileen M. Bauer,*2Rong Cao,2Salony Maniar,2Mackenzie Mosher,2Jun Chen,3
Qiming Jane Wang,3Joseph C. Glorioso,4Bruce R. Pitt,2Simon C. Watkins,1and Claudette M. St. Croix2
Departments of1Cell Biology and Physiology,2Environmental and Occupational Health,3Pharmacology and Chemical
Biology, and4Microbiology and Molecular Genetics, The University of Pittsburgh, Pittsburgh, Pennsylvania
Submitted 15 September 2010; accepted in final form 2 March 2011
Bernal PJ, Bauer EM, Cao R, Maniar S, Mosher M, Chen J,
zinc in regulatinghypoxia-inducedcontractileeventsinpulmonaryendothe-
lium. Am J Physiol Lung Cell Mol Physiol 300: L874–L886, 2011. First
published March 4, 2011; doi:10.1152/ajplung.00328.2010.—We previ-
ously reported that zinc thiolate signaling contributes to hypoxic
contraction of small, nonmuscularized arteries of the lung. The pres-
ent studies were designed to investigate mechanisms by which hy-
poxia-released zinc induces contraction in isolated pulmonary endo-
thelial cells and to delineate the signaling pathways involved in
zinc-mediated changes in the actin cytoskeleton. We used fluores-
cence-based imaging to show that hypoxia induced time-dependent
increases in actin stress fibers that were reversed by the zinc chelator,
N,N,N=,N=-tetrakis-(2-pyridylmethyl)-ethylenediamine (TPEN). We
further showed that hypoxia-induced phosphorylation of the contrac-
tile protein myosin light chain (MLC) and assembly of actin stress
fibers were each TPEN sensitive. Hypoxia and zinc-induced inhibition
of MLC phosphatase (MLCP) were independent of the regulatory
subunit (MYPT1) of MLCP, and therefore hypoxia-released zinc
likely inhibits MLCP at its catalytic (PP1) subunit. Inhibition of PKC
by Ro-31–8220 and a dominant-negative construct of PKC-ε attenuated
hypoxia-induced contraction of isolated pulmonary endothelial cells.
Furthermore, zinc-induced phosphorylation of MLC (secondary to inhi-
bition of MLCP) was PKC dependent, and hypoxia-released zinc pro-
moted the phosphorylation of the PKC substrate, CPI-17. Collectively,
these data suggest a link between hypoxia, elevations in labile zinc, and
activation of PKC, which in turn acts through CPI-17 to inhibit MLCP
activity and promote MLC phosphorylation, ultimately inducing stress
fiber formation and endothelial cell contraction.
pulmonary circulation; endothelial cells; actin stress fibers
OUR RECENT DATA SHOWED that acute hypoxia-induced increases
in nitric oxide (NO) biosynthesis result in increases in intra-
cellular free zinc that in turn contribute to vasoconstriction of
small, intra-acinar arteries of lung (8). Because this anatomic
site is composed primarily of endothelial cells with only
solitary or discontinuous smooth muscle-like cells (e.g., peri-
cytes) in their wall (13), we investigated the potential for
hypoxia-zinc-mediated contraction in pulmonary endothelium
and confirmed that isolated pulmonary (but not systemic)
endothelial cells constricted in hypoxia (8) and that these
contractile events were associated with hypoxia-induced in-
creases in labile zinc. The mechanism by which zinc can
induce vasoconstriction in the pulmonary vasculature is not
known. However, zinc-associated proteins account for roughly
10% of the human proteome (26), and many of these putative
targets for hypoxia-released zinc are involved in signaling
pathways regulating cellular contractility.
Endothelial cells contain all the molecular machinery required
to generate contractile force via the actomyosin motor. Contrac-
tion is initiated by phosphorylation of the 20-kDa regulatory
myosin light chain (MLC) at S19/T18 (17). The phosphorylation
of MLC is dependent on a balance between the activities of
calcium/calmodulin-dependent MLC kinase (MLCK) and MLC
phosphatase (MLCP). Remodeling of the endothelial actin cyto-
skeletal in response to either hypoxia or the procoagulant protein,
thrombin, involves MLC and actin-related proteins (16). One
potential connection between these signaling pathways and zinc is
the relationship between divalent metal ions and sulfhydryl resi-
dues in the activation of type 1 and type 2A serine/threonine
phosphoprotein phosphatases (PPases), with zinc having been
shown to be a potent inhibitor of ?-PPase (43). A further link is
provided by the indirect evidence supporting a role for zinc in
strengthening of focal adhesions (3) although the mechanism(s)
underlying this phenomenon are not clear. Likely participants in
and PKC isoforms, the latter of which are known to be tightly
regulated by zinc (27) and participate in hypoxic pulmonary
vasoconstriction (HPV) (24) (i.e., PKC-ε).
The present studies were designed to investigate the mech-
anisms by which hypoxia-released zinc induces contraction in
pulmonary endothelium and to delineate the signaling path-
ways involved in zinc-mediated changes in the actin cytoskel-
eton. We hypothesized that hypoxia-induced alterations in zinc
homeostasis promote endothelial cell contraction via changes
in the phosphorylation status of MLC and increased formation/
stabilization of actin stress fibers.
MATERIALS AND METHODS
Chemicals and materials. All reagents were purchased from Sigma-
Aldrich (www.sigmaaldrich.com, St. Louis, MO) unless otherwise
noted. Details of the dominant-negative PKC-ε herpes simplex virus
were provided elsewhere (34). The enhanced green fluorescent protein
(EGFP) actin construct was a gift from Michael Davidson at The
Florida State University.
Cell culture. Cells were grown at 37°C with 5% CO2. Details
regarding the culture of sheep pulmonary artery endothelial cells
(SPAEC) are described elsewhere (36). Rat pulmonary microvascular
endothelial cells (RPMVEC) were purchased from VEC Technologies
(VEC Technologies, Rensselaer, NY) and grown in complete MCDB-
131 media (Lonza, Allendale, NJ). Rat pulmonary artery endothelial cells
(RPAEC) were grown in DMEM (Fisher Scientific, Waltham, MA) with
10% fetal bovine serum, 2 mM glutamine, and 100 U/ml penicillin/
* P. Bernal and E. Bauer contributed equally to this work.
Address for reprint requests and other correspondence: C. M. St. Croix,
Dept. of Environmental and Occupational Health, Univ. of Pittsburgh Graduate
School of Public Health, BRIDG, 100 Technology Drive, Pittsburgh, PA
15219 (e-mail: firstname.lastname@example.org).
Am J Physiol Lung Cell Mol Physiol 300: L874–L886, 2011.
First published March 4, 2011; doi:10.1152/ajplung.00328.2010.
1040-0605/11 Copyright © 2011 the American Physiological Society http://www.ajplung.orgL874
Immunofluorescence. Cells were seeded on glass coverslips coated
with laminin (11.34 ?g/ml, Fisher Scientific). Hypoxic exposures
were performed inside a hypoxic chamber (Coy Laboratory Products,
Grass Lake, MI). Following treatment, cells were fixed, and perme-
abilized in 2% paraformaldehyde with 0.1% Triton X-100. Alexa
Fluor488 Phalloidin was used for F-actin staining of stress fibers.
Sequential XYZ-sections (1024X1024 pixels, at Nyquist axial fre-
quency) were obtained using an Olympus Fluoview 1000 confocal
microscope (Bethlehem, PA) equipped with a ?60 oil-immersion
optic (NA, 1.43). Three-dimensional images were reconstructed using
Metamorph (Molecular Devices, Downingtown, PA), and quantifica-
tion of actin stress filaments was determined by volume rendering in
Imaris (Bitplane, Saint Paul, MN).
Live cell imaging. Cells were seeded on 35-mm laminin-coated
glass-bottom dishes (MatTek, Ashland, MA) and imaged in a closed,
thermocontrolled (37°C) stage-top incubator (Tokai-Hit, Tokyo, Ja-
pan). Images were obtained using a Nikon TE2000E (Melville, NY)
microscope equipped with a ?40 oil-immersion objective (Nikon,
CFI PlanFluor, NA 1.3) and Q-Imaging Retiga EXI camera (Burnaby,
BC, Canada). MetaMetamorph (Molecular Devices) was used to
collect and analyze data and to drive the microscope. Hypoxic
conditions were obtained by bubbling the media with hypoxic gas
(90% N2-5% CO2-1.5% O2), which reduced oxygen tension to 15 ?
2 mmHg, as measured using a Clarke electrode.
Total internal reflectance fluorescence imaging. Cells were imaged
on a Nikon TiE inverted microscope with a 1.49 NA oil-immersion
objective capable of both epifluorescence and total internal reflectance
fluorescence (TIRF) microscopy illumination through the objective.
EGFP was excited with a 488-nm Coherent Sapphire laser (Coherent,
Santa Clara, CA). Laser intensities were controlled using a Neos
AOTF mounted on a Prairie Technologies laser bench (Madison,
WI). To ensure correct image registration, a triple-pass filter cube
(488 nm, 561 nm, 638 nm) and matched emitter filters were used
(Chroma, Brattleboro, VT). Images were collected using a Photo-
metrics HQ2 Coolsnap camera (Photometrics, Tucson, AZ) at full
resolution. Data were collected and analyzed using NIS-Elements
Western blotting. Mouse monoclonal PKC-ε antibody (E-5) was
purchased from Santa Cruz Biotechnology (Santa Cruz, CA) and used
at a 1:500 dilution. MLC2, phospho-MLC (Thr18/Ser19), MYPT1,
and phospho-MYPT1 (Thr853) antibodies were purchased from Cell
Signaling Technology (Beverly, MA) and used at 1:1,000 dilutions.
CPI-17 and phospho-CPI-17 antibodies were purchased from Santa
Cruz Biotechnology and used at 1:100 dilutions. Total protein levels
were normalized to the level of ?-actin for each sample, and phos-
phorylated protein expressed relative to total protein levels.
Membrane fractionation. Cells were lysed (100 mM Tris·HCl, pH
7.4, 1%, vol/vol Nonidet P-40, 10 mM NaF, 1 mM vanadate, 10 ?g/ml
of aprotinin, 10 ?g/ml of leupeptin) and centrifuged for 10 min at 14,000
revolution/min. The supernatant was further centrifuged at 43,000 g for
30 min (21) to separate cytoplasmic and membranes fraction.
PKC-? immunoprecipitation and enzyme activity assay. Cells were
lysed in modified RIPA buffer (100 mM Tris·HCl, pH 7.4, 1%,
vol/vol, Nonidet-P40 10 mM NaF, 1 mM vanadate, 10 ?g/ml of
aprotinin, 10 ?g/ml of leupeptin). Insoluble material was removed by
centrifugation, and protein concentrations were determined using the
Bio-Rad DC protein assay (Bio-Rad, Hercules. CA). Equal amounts
of protein were precleared with protein A-Sepharose and incubated
with antibody for 2 h at 4°C. The immune complexes were isolated
with Protein A-Sepharose, washed, and eluted. Equal amounts of
immunocomplex were then subjected to PKC-ε kinase assay, as
described previously (6).
Statistical analysis. Data are presented as means ? SD. Compar-
isons between more than two groups were done using ANOVA
followed by Dunnett’s posttest. A value of P ? 0.05 was considered
Hypoxia induces zinc-dependent changes in the actin cyto-
skeleton of isolated pulmonary microvascular endothelial
cells. We previously reported that hypoxia induced increases in
labile zinc in small intra-acinar arteries of the isolated perfused
mouse lung (8). The observation that hypoxic vasoconstriction
was blunted in the lungs of mice in which the major zinc binding
protein (metallothionein; MT) was knocked out (MT?/?mice), or
in wild-type mice perfused with the zinc chelator, N,N,N=,N=-
tetrakis-(2-pyridylmethyl)-ethylenediamine (TPEN), led us to
hypothesize that observed increases in intracellular zinc con-
tribute to constriction in the pulmonary microvasculature. The
anatomic site in question was shown to be composed primarily
of endothelial cells (8), and these initial investigations confirmed
the potential for hypoxia-zinc-mediated contraction in isolated
primary cultures of pulmonary endothelium. In the present report,
in the actin cytoskeleton in isolated rat pulmonary microvascular
endothelial cells (RPMVEC). Hypoxic exposure increased the
abundance or total volume of actin per cell, as well as the
alignment of actin stress fibers (Fig. 1, mean data Fig. 2B)
compared with normoxia. Treatment with TPEN reduced the
effects of hypoxia on the actin cytoskeleton (Figs. 1 and 2B),
suggesting that hypoxia-induced changes in intracellular
zinc contribute to the formation or stabilization of actin
stress filaments in RPMVEC. Although early reports suggest
that zinc can alter skeletal muscle contractility (19), little is
known about the effects of zinc on the intracellular contrac-
tile apparatus in either muscle or nonmuscle cells. Our data
in fixed pulmonary endothelial cells showed that exogenous
zinc (in the presence of the zinc ionophore, pyrithione) also
increased the abundance and altered the distribution of actin
stress fibers (Fig. 2, A and B), consistent with a contractile
We next used TIRF microscopy of EGFP-tagged actin to
examine hypoxia- and zinc-induced contractile events in live
cultures of RPMVEC. TIRF relies on an evanescent wave
generated perpendicular to the optical axis when light reflects
off of a surface at an incident angle. The intensity of the
evanescent wave decays exponentially and effectively pene-
trates only 100–150 nm beyond the coverslip into the cell,
thereby confining the excitation of fluorophores to an ex-
tremely thin axial slice (20). As a result the signal-to-noise
ratio in TIRF is extremely high. Although the approach will
only image events at the basal surface of the cell, one can be
certain that emitted signal is only derived from this region of
the cell, as opposed to other methods that are not specifically
constrained in the z-axis to a single plane (e.g., confocal).
Using TIRF, we observed that hypoxia caused time-dependent
and reversible (Fig. 3) increases in the abundance and align-
ment of basal stress fibers (Figs. 3 and 4, Supplemental movies
S1 and S2; supplemental material for this article is available
online at the American Journal of Physiology Lung Cellular
and Molecular Physiology website). Consistent with the data in
fixed cells (Fig. 1), the addition of the zinc chelator, TPEN,
during hypoxia, resulted in the rapid disassembly of actin stress
filaments (Fig. 4, Supplemental movie S2). We previously
showed that isolated RPMVEC that were embedded in a
flexible collagen matrix actively contracted in response to
hypoxic stimuli (8). The resultant thickness of this collagen gel
HYPOXIA, ZINC, ENDOTHELIUM, AND CELL CONTRACTION
AJP-Lung Cell Mol Physiol • VOL 300 • JUNE 2011 • www.ajplung.org
contractile regulatory protein, MLC, was PKC dependent (Fig.
5A). We further showed that hypoxia-released zinc promoted
the phosphorylation of the PKC substrate, CPI-17 (Fig. 5C).
PKC acts through CPI-17 to stabilize the phosphorylation of
MLC by inactivating MLC phosphatase, ultimately inducing
stress fiber formation and cell contraction (Fig. 10). Such
signaling has been shown to regulate the reorganization of
cytoskeletal proteins and affect changes in barrier function in
pulmonary endothelium (21). Although there is compelling
evidence supporting a critical role for zinc in maintaining
epithelial barrier function in the context of inflammatory stress
(9) and it is not unreasonable to propose that zinc may also play
a role in regulating endothelial barrier function, further studies
are required to determine the downstream effects of the hyp-
oxia/zinc pathway on endothelial permeability in the lung.
Investigating a role for individual PKC isoforms in the
described zinc signaling pathways regulating endothelial con-
tractility is complicated by the lack of specificity of the
pharmacological inhibitors and activators of the enzyme. Ro-
31–8220 has an IC50of 0.024 ?M for PKC-ε (42); however, at
high concentrations it can also inhibit PKC-?, PKC-?II, and
PKC-?. The difficulties in using such inhibitors are further
demonstrated by observations that Ro-31–8220 can activate
c-Jun expression and inhibit MAPK phosphatase-1 expression
(1, 7). Although we cannot eliminate the possibility that mul-
tiple isoforms of PKC play a role in modulating the response to
hypoxia observed in pulmonary endothelium, the observation
that the dominant-negative approach to specifically inhibit
PKC-ε also significantly attenuated the hypoxia-induced con-
traction of isolated cells confirmed the importance of the
PKC-ε isoform in mediating pulmonary endothelial cell con-
traction. Similarly, although zinc-induced changes in MLC
phosphorylation were abrogated by pharmacological inhibition
of Rho kinase, it has been reported that Y-27632 can exert
nonspecific inhibitory effects on PKC-mediated vascular re-
sponses at concentrations in excess of 10 ?M (10). Therefore,
future studies incorporating specific knockdown of the genes in
question are required to delineate the precise pathways regu-
lating these events.
The specificity of the proposed zinc pathways in mediating
hypoxia-induced contraction in the pulmonary vasculature is
not known. We have previously shown that both hypoxia-
induced changes in labile zinc and hypoxia-induced endothelial
cell contraction (8) were unique to cells derived from lung in
that aortic endothelium did not exhibit these responses to low
PO2. However, we do not anticipate that the tissue specificity
of HPV is conferred by activation of PKC-ε in that PKC has
been shown to be an important signaling molecule mediating
contractile responses in both the systemic and pulmonary
circulation (32, 41). In our hands, the hypoxia-induced changes
in PKC-ε activity were both L-NAME and TPEN sensitive,
arguing a role for NO-induced changes in labile zinc in
mediating enzyme function. NO donors have also been shown
to induce activation, translocation, and nitration of PKC-ε in
cardiac myocytes (5). In fact, both endothelial NOS (eNOS)
and inducible NOS proteins are associated with PKC-ε in the
heart (30) although data obtained in PKC-ε?/?mice suggested
a role for PKC-ε in the regulation of eNOS expression during
chronic hypoxic exposure rather than the direct regulation of
enzymatic activity (24). Our data suggest a role for PKC in
mediating the downstream effects of hypoxia-induced changes
in NO synthesis and intracellular zinc homeostasis to produce
contraction of pulmonary endothelium. Although data obtained
using dominant-negative approaches, and isoform-specific an-
tibodies, directly implicated the involvement of PKC-ε, we
cannot eliminate the possibility that other pathways, or other
isoforms of PKC, also contribute to HPV and/or the generation
of contractile force in isolated pulmonary endothelial cells.
The authors acknowledge the technical expertise of Christina Goldbach and
This work was funded in part by NIH HL081421 (C. St Croix), NIH R37
HL65697 (B. Pitt), R01CA142580-01 and R01CA129127-01 (Q. Wang), and
1 U54 RR022241-01 (S. Watkins).
No conflicts of interest, financial or otherwise are declared by the authors.
1. Alessi DR, Caudwell FB, Andjelkovic M, Hemmings BA, Cohen P.
Molecular basis for the substrate specificity of protein kinase B; compar-
ison with MAPKAP kinase-1 and p70 S6 kinase. FEBS Lett 399: 333–338,
2. An SS, Pennella CM, Gonnabathula A, Chen J, Wang N, Gaestel M,
Hassoun PM, Fredberg JJ, Kayyali US. Hypoxia alters biophysical
properties of endothelial cells via p38 MAPK- and Rho kinase-dependent
pathways. Am J Physiol Cell Physiol 289: C521–C530, 2005.
3. Avnur Z, Geiger B. Substrate-attached membranes of cultured cells
isolation and characterization of ventral cell membranes and the associated
cytoskeleton. J Mol Biol 153: 361–379, 1981.
4. Bain J, McLauchlan H, Elliott M, Cohen P. The specificities of protein
kinase inhibitors: an update. Biochem J 371: 199–204, 2003.
5. Balafanova Z, Bolli R, Zhang J, Zheng Y, Pass JM, Bhatnagar A,
Tang XL, Wang O, Cardwell E, Ping P. Nitric oxide (NO) induces
nitration of protein kinase Cepsilon (PKCepsilon ), facilitating PKCepsi-
lon translocation via enhanced PKCepsilon -RACK2 interactions: a novel
mechanism of no-triggered activation of PKCepsilon. J Biol Chem 277:
6. Bandyopadhyay G, Standaert ML, Zhao L, Yu B, Avignon A, Gallo-
way L, Karnam P, Moscat J, Farese RV. Activation of protein kinase C
(alpha, beta, and zeta) by insulin in 3T3/L1 cells. Transfection studies
suggest a role for PKC-zeta in glucose transport. J Biol Chem 272:
7. Beltman J, McCormick F, Cook SJ. The selective protein kinase C
inhibitor, Ro-31–8220, inhibits mitogen-activated protein kinase phospha-
tase-1 (MKP-1) expression, induces c-Jun expression, and activates Jun
N-terminal kinase. J Biol Chem 271: 27018–27024, 1996.
8. Bernal PJ, Leelavanichkul K, Bauer E, Cao R, Wilson A, Wasserloos
KJ, Watkins SC, Pitt BR, and St Croix CM. Nitric-oxide-mediated zinc
release contributes to hypoxic regulation of pulmonary vascular tone. Circ
Res 102: 1575–1583, 2008.
9. Besecker B, Bao S, Bohacova B, Papp A, Sadee W, Knoell DL. The
human zinc transporter SLC39A8 (Zip8) is critical in zinc-mediated
cytoprotection in lung epithelia. Am J Physiol Lung Cell Mol Physiol 294:
10. Budzyn K, Paull M, Marley PD, Sobey CG. Segmental differences in
the roles of rho-kinase and protein kinase C in mediating vasoconstriction.
J Pharmacol Exp Ther 317: 791–796, 2006.
11. Chou SS, Clegg MS, Momma TY, Niles BJ, Duffy JY, Daston GP,
Keen CL. Alterations in protein kinase C activity and processing during
zinc-deficiency-induced cell death. Biochem J 383: 63–71, 2004.
12. Csermely P, Szamel M, Resch K, Somogyi J. Zinc can increase the
activity of protein kinase C and contributes to its binding to plasma
membranes in T lymphocytes. J Biol Chem 263: 6487–6490, 1988.
13. Donoghue L, Tyburski JG, Steffes CP, Wilson RF. Vascular endothelial
growth factor modulates contractile response in microvascular lung peri-
cytes. Am J Surg 191: 349–352, 2006.
HYPOXIA, ZINC, ENDOTHELIUM, AND CELL CONTRACTION
AJP-Lung Cell Mol Physiol • VOL 300 • JUNE 2011 • www.ajplung.org
14. Eto M, Ohmori T, Suzuki M, Furuya K, Morita F. A novel protein
phosphatase-1 inhibitory protein potentiated by protein kinase C. Isolation
from porcine aorta media and characterization. J Biochem (Tokyo) 118:
15. Fukata Y, Amano M, Kaibuchi K. Rho-Rho-kinase pathway in smooth
muscle contraction and cytoskeletal reorganization of non-muscle cells.
Trends Pharmacol Sci 22: 32–39, 2001.
16. Gavard J, Gutkind JS. Protein kinase C-related kinase and ROCK are
required for thrombin-induced endothelial cell permeability downstream
from Galpha12/13 and Galpha11/q. J Biol Chem 283: 29888–29896,
17. Goeckeler ZM, Wysolmerski RB. Myosin light chain kinase-regulated
endothelial cell contraction: the relationship between isometric tension,
actin polymerization, and myosin phosphorylation. J Cell Biol 130: 613–
18. Gopalakrishna R, Jaken S. Protein kinase C signaling and oxidative
stress. Free Radic Biol Med 28: 1349–1361, 2000.
19. Isaacson A, Sandow A. Effects of zinc on responses of skeletal muscle.
J Gen Physiol 46: 655–677, 1963.
20. Kane LP, Watkins SC. Dynamic regulation of Tec kinase localization in
membrane-proximal vesicles of a T cell clone revealed by total internal
reflection fluorescence and confocal microscopy. J Biol Chem 280:
21. Kolosova IA, Ma SF, Adyshev DM, Wang P, Ohba M, Natarajan V,
Garcia JG, Verin AD. Role of CPI-17 in the regulation of endothelial
cytoskeleton. Am J Physiol Lung Cell Mol Physiol 287: L970–L980, 2004.
22. Kumar S, Maxwell IZ, Heisterkamp A, Polte TR, Lele TP, Salanga M,
Mazur E, Ingber DE. Viscoelastic retraction of single living stress fibers
and its impact on cell shape, cytoskeletal organization, and extracellular
matrix mechanics. Biophys J 90: 3762–3773, 2006.
23. Leung T, Chen XQ, Manser E, Lim L. The p160 RhoA-binding kinase
ROK alpha is a member of a kinase family and is involved in the
reorganization of the cytoskeleton. Mol Cell Biol 16: 5313–5327, 1996.
24. Littler CM, Morris KG Jr, Fagan KA, McMurtry IF, Messing RO,
Dempsey EC. Protein kinase C-epsilon-null mice have decreased hypoxic
pulmonary vasoconstriction. Am J Physiol Heart Circ Physiol 284:
25. Maekawa M, Ishizaki T, Boku S, Watanabe N, Fujita A, Iwamatsu A,
Obinata T, Ohashi K, Mizuno K, Narumiya S. Signaling from Rho to
the actin cytoskeleton through protein kinases ROCK and LIM-kinase.
Science 285: 895–898, 1999.
26. Maret W, Li Y. Coordination dynamics of zinc in proteins. Chem Rev
109: 4682–4707, 2009.
27. Murakami K, Whiteley MK, Routtenberg A. Regulation of protein
kinase C activity by cooperative interaction of Zn2?and Ca2?. J Biol
Chem 262: 13902–13906, 1987.
28. Naumanen P, Lappalainen P, Hotulainen P. Mechanisms of actin stress
fibre assembly. J Microsc 231: 446–454, 2008.
29. Orton EC, Raffestin B, McMurtry IF. Protein kinase C influences rat
pulmonary vascular reactivity. Am Rev Respir Dis 141: 654–658, 1990.
30. Ping P, Takano H, Zhang J, Tang XL, Qiu Y, Li RC, Banerjee S,
Dawn B, Balafonova Z, Bolli R. Isoform-selective activation of protein
kinase C by nitric oxide in the heart of conscious rabbits: a signaling
mechanism for both nitric oxide-induced and ischemia-induced precondi-
tioning. Circ Res 84: 587–604, 1999.
31. Rainteau D, Wolf C, Lavialle F. Effects of calcium and calcium analogs
on calmodulin: a Fourier transform infrared and electron spin resonance
investigation. Biochim Biophys Acta 1011: 81–87, 1989.
32. Rasmussen H, Takuwa Y, Park S. Protein kinase C in the regulation of
smooth muscle contraction. FASEB J 1: 177–185, 1987.
33. Somlyo AP, Somlyo AV. Ca2?sensitivity of smooth muscle and non-
muscle myosin II: modulated by G proteins, kinases, and myosin phos-
phatase. Physiol Rev 83: 1325–1358, 2003.
34. Srinivasan R, Wolfe D, Goss J, Watkins S, de Groat WC, Sculpto-
reanu A, Glorioso JC. Protein kinase C epsilon contributes to basal and
sensitizing responses of TRPV1 to capsaicin in rat dorsal root ganglion
neurons. Eur J Neurosci 28: 1241–1254, 2008.
35. St Croix CM, Stitt MS, Leelavanichkul K, Wasserloos KJ, Pitt BR,
Watkins SC. Nitric oxide-induced modification of protein thiolate clusters
as determined by spectral fluorescence resonance energy transfer in live
endothelial cells. Free Radic Biol Med 37: 785–792, 2004.
36. St Croix CM, Wasserloos KJ, Dineley KE, Reynolds IJ, Levitan ES,
Pitt BR. Nitric oxide-induced changes in intracellular zinc homeostasis
are mediated by metallothionein/thionein. Am J Physiol Lung Cell Mol
Physiol 282: L185–L192, 2002.
37. Takayama M, Ebihara Y, Tani M. Differences in the expression of
protein kinase C isoforms and its translocation after stimulation with
phorbol ester between young-adult and middle-aged ventricular cardio-
myocytes isolated from Fischer 344 rats. Jpn Circ J 65: 1071–1076, 2001.
38. Tang ZL, Wasserloos K, St Croix CM, Pitt BR. Role of zinc in
pulmonary endothelial cell response to oxidative stress. Am J Physiol Lung
Cell Mol Physiol 281: L243–L249, 2001.
39. Velasco G, Armstrong C, Morrice N, Frame S, Cohen P. Phosphory-
lation of the regulatory subunit of smooth muscle protein phosphatase 1M
at Thr850 induces its dissociation from myosin. FEBS Lett 527: 101–104,
40. Ward JP, McMurtry IF. Mechanisms of hypoxic pulmonary vasocon-
striction and their roles in pulmonary hypertension: new findings for an old
problem. Curr Opin Pharmacol 9: 287–296, 2009.
41. Weissmann N, Voswinckel R, Hardebusch T, Rosseau S, Ghofrani
HA, Schermuly R, Seeger W, Grimminger F. Evidence for a role of
protein kinase C in hypoxic pulmonary vasoconstriction. Am J Physiol
Lung Cell Mol Physiol 276: L90–L95, 1999.
42. Wilkinson SE, Parker PJ, Nixon JS. Isoenzyme specificity of bisindolyl-
maleimides, selective inhibitors of protein kinase C. Biochem J 294:
43. Zhuo S, Dixon JE. Effects of sulfhydryl regents on the activity of lambda
Ser/Thr phosphoprotein phosphatase and inhibition of the enzyme by zinc
ion. Protein Eng 10: 1445–1452, 1997.
HYPOXIA, ZINC, ENDOTHELIUM, AND CELL CONTRACTION
AJP-Lung Cell Mol Physiol • VOL 300 • JUNE 2011 • www.ajplung.org