1 Departments of Physiology, Pediatrics, and Obstetrics & Gynecology and 2 Department of Anatomy and Neurobiology, University of Tennessee Health Science Center, Memphis, Tennessee 38163; and 3 Institute of Cytology, Russian Academy of Sciences, St. Petersburg, Russia 194064
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ABSTRACT |
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We investigated the relationships among
expression, activity, and spatial organization of cyclooxygenase (COX-1
and COX-2) in endothelial cells from porcine and human cerebral
microvessels and from human umbilical vein. In quiescent cells, COX-1
was detected in the perinuclear zone and the cytoplasm, while COX-2 was
mainly a nuclear resident possibly connected with the nuclear matrix. COX-2 immunogold labeling was situated in the nuclear envelope, at the
nuclear pores, and in connection with the perichromatin regions of the
nucleus, considered to be the sites of active transcription. In human
endothelial cells transcriptionally activated by interleukin (IL)-1,
the nucleus remained a major COX-2 localization site during the first
12 h of stimulation, when COX-2 expression was maximally induced.
The continuous rise in prostanoid synthesis at 17-23 h of
stimulation was associated with COX-2 relocation from the nucleus to
the nuclear envelope and the cytoplasm. IL-1
did not affect COX-1
expression, activity, and localization. COX-2 nuclear localization
sites and trafficking between the nucleus and the cytoplasm in
endothelial cells may indicate a novel function of COX-2 in regulating
gene expression.
prostanoids; endothelial cell; interleukin-1; nucleus
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INTRODUCTION |
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CYCLOOXYGENASE (COX-1 and COX-2), by catalyzing the first committed step in prostanoid biosynthesis, plays an important role in endothelial functioning, including vascular reactivity and angiogenesis (12, 13, 25, 38). COX-1 is a housekeeping gene product, while COX-2 was originally described as an inducible isoform, a product of primary response/immediate-early genes. However, COX-2 also is expressed under resting conditions in brain (4), cerebral microvessels (3, 28, 31), and cultured endothelial cells from cerebral microvessels (28). Vascular endothelium from the brain cortex is a physiologically significant model in which to study COX functioning, because endothelium-derived prostaglandins regulate cerebral blood flow in newborn piglets and human neonates (17, 18).
Despite similarity in structure and catalytic properties, COX-1 and
COX-2 use different pools of arachidonic acid (AA) (32) and have distinct functions in cell proliferation and differentiation (8, 12, 13, 25, 38). Distinct spatial organization of the
COX isoforms could provide a plausible explanation for these
observations. COX-1 and COX-2 molecules consist of several major
domains, including an NH2-terminal epidermal growth
factor-like domain, a monotopic membrane-binding motif, and the
COOH-terminal catalytic domain (16, 39). The catalytic
center is highly conserved (87%), while the amino acid sequences of
the membrane binding domains and the amino- and carboxy-terminal ends
of COX-1 and COX-2 molecules are largely different (16, 25,
39). Although the differences in the amino acid sequences
suggest that COX-1 and COX-2 might segregate into unique cellular
compartments (25), experimental data on the intracellular
localization of COX-1 and COX-2 remain controversial. Initial studies
indicated that in murine 3T3 cells, both COX-1 (in starved cells) and
COX-2 (in serum-stimulated cells) have identical localization sites: the nuclear envelope and the endoplasmic reticulum (33).
Further reports indicated that COX-2 is preferentially associated with the nuclear envelope (21). However, a later immunoelectron
microscopy study conducted by the same group of investigators in
serum-stimulated murine 3T3 cells and human monocytes and in
interleukin (IL)-1-stimulated human umbilical vein endothelial cells
(HUVEC) concluded that COX-2 is evenly distributed between the nuclear
envelope and the endoplasmic reticulum (40).
COX-2 may also have intranuclear localization sites. COX-1 and COX-2
were present on both the inner and outer nuclear membranes in
serum-stimulated human monocytes and NIH/3T3 cells (40). In a human colon cancer cell line, COX-2 appearance in the nucleus was
observed upon stimulation with transforming growth factor- (5). The possibility of nuclear localization of COX-2 is
especially intriguing because COX-2 has been implicated in the
pathogenesis of proliferative diseases and cancer by cell-specific
prostanoid-dependent and prostanoid-independent actions (4, 13,
44). In endothelial cells from cerebral microvessels of newborn
pigs in primary culture, COX-2 is localized mainly intranuclearly
(28). Under stimulated conditions associated with COX-2
induction in porcine cerebral microvascular endothelial cells, we
observed strong COX-2 immunofluorescence signal in the nucleus that was
followed by COX-2 accumulation in the nuclear envelope and the
cytoplasm (28). These findings may indicate a possible
physiological role of COX-2 in the nuclear functions in endothelial cells.
The present study was designed to address three hypotheses. First,
COX-2 is expressed and has nuclear localization sites in unstimulated
endothelial cells of human origin similar to those from piglets.
Second, nuclear COX-2 localization sites are associated with the
specific nuclear domains that spatially organize nuclear functions.
Third, newly synthesized COX-2 protein induced in cytokine-stimulated endothelial cells can be transported to the nucleus. To address these
hypotheses, we have investigated the relationships among expression,
activity, and spatial organization of COX-1 and COX-2 in vascular
endothelial cells of piglet and human origin at quiescence and upon
activation with the proinflammatory cytokine IL-1.
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METHODS |
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Cell cultures. Endothelial cells from cerebral microvessels were isolated and cultured as previously described (28). Brain cortex was obtained from newborn pigs (1-5 days of age) or from a human newborn (gestation age 27 wk). Postmortem collection of human brain tissue for these specific studies was approved by the Institutional Review Board. The sample was collected by the pathologist at autopsy.
The brain cortex tissue was homogenized in M199 medium and filtered through 300- and 60-µm nylon mesh screens consecutively. Cerebral microvessels (60-300 µm) collected on the last screen were digested by collagenase-dispase (2 mg/ml for 2 h at 37°C). Dissociated endothelial cells were separated by centrifugation in Percoll density gradient and plated on Matrigel-coated cell culture plates or glass coverslips. Endothelial cells from newborn piglet cerebral microvessels were cultured (37°C, 5% CO2-95% air) in Dulbecco's modified Eagle's medium (DMEM) with 20% fetal bovine serum (FBS), 30 µg/ml endothelial cell growth supplement, 1 U/ml heparin, and antibiotic/antimycotic mixture for 5-6 days until confluence. Endothelial cells from human cerebral microvessels were cultured in the growth medium for human microvascular endothelial cells (BioWhittaker, San Diego, CA) containing 7% FBS. To isolate HUVEC, we injected the collagenase-dispase solution (2 mg/ml in M199) intraluminally into freshly dissected umbilical cords and incubated for 15 min at 37°C. Dissociated endothelial cells were flushed out with M199, collected by centrifugation, plated to Matrigel-coated cell culture plates or glass coverslips, and cultured in the growth medium for human endothelial cells (BioWhittaker) containing 2% FBS for 5-6 days. For some experiments, HUVEC (passage 1) purchased from VEC Technologies (Rensselaer, NY) were used. All experiments were performed on confluent quiescent cells in primary cultures (porcine brain, HUVEC) or in cells at one to three passages (human brain, HUVEC). To achieve quiescence, we exposed cells to a serum-depleted medium (0.1% FBS) for 15-20 h before the experiment.IL-1 treatment.
Endothelial cells were incubated with the serum-depleted starvation
medium (0.1% FBS-DMEM) for 15-20 h to achieve quiescence. After
this period, human IL-1
(50 ng/ml) was added to the starvation medium for an additional 0 (control), 2, 4, 6, 8, 12, 16, 20, and
23 h as indicated in RESULTS. On the basis of our
preliminary data, at this concentration IL-1
most effectively
increases prostanoid production without affecting endothelial cell
viability. Media were collected at the end of each incubation period,
precleared by centrifugation (2,000 g, 5 min), and stored at
20°C for prostanoid detection. Cells were used for RNA isolation,
Western blots, and COX activity detection as described.
RNA isolation, RT-PCR, and RNase protection assay.
All samples were initially normalized to the cell count. Total RNA was
isolated from cells by guanidinium thiocyanate phenol-chloroform in a
single-step extraction method using an RNA isolation kit (Stratagene,
La Jolla, CA). RNA concentration was determined by absorbance at 260 nm. The integrity of RNA was analyzed by agarose gel electrophoresis
and ethidium bromide staining. mRNA levels were assessed by reverse
transcriptase-polymerase chain reaction (RT-PCR). Total RNA was
converted to cDNA by incubation at 37°C for 1 h with 50 units of
reverse transcriptase and random primers (100 ng/ml) using a RT-PCR kit
(Stratagene). PCR primers for COX-1 and COX-2 were purchased from
Biomed (Oxford, MI), and primers for -actin were from Ambion
(Austin, TX). Samples were amplified for 40 cycles of denaturation at
94°C for 1 min, annealing at 53°C for 1 min, and extension at
72°C for 2 min in a PTC100 Programmable Thermal Controller (MJ
Research, Watertown, MA). Negative control without cDNA added was run
in parallel for all primer pairs in each experiment. Amplified cDNA was
analyzed by 1.2% agarose gel electrophoresis and visualized with SYBR
Green I nucleic acid gel stain (30-min incubation at 37°C) (Molecular
Probes, Eugene, OR). Gels were scanned and analyzed using a Storm Image
detection system (Molecular Dynamics, Sunnyvale, CA). RNase protection
assays were performed using the RiboQuant Multi-Probe RNase Protection Assay System and the custom-made template sets for human COX-1 and
COX-2 detection containing housekeeping genes L-32 and
glyceraldehyde-3-phosphate dehydrogenase (GAPDH; PharMingen, San Diego,
CA). Briefly, the DNA templates were collectively transcribed under the
direction of the DNA-dependent RNA T7 polymerase into an
-32P-labeled antisense RNA probe set. Labeled probes
(3.3 × 105 cpm) were hybridized for 16 h at
56°C to 15 µg of total RNA. To digest single-stranded RNA, we
treated RNA hybrids with ribonuclease A and purified by
phenol-chloroform extraction and ammonium acetate precipitation. The
protected
-32P-labeled RNA fragments were resolved on
5% polyacrylamide-urea sequencing gel. The undigested probes (1,000 cpm/lane) were used as the molecular weight markers. The gel was placed
on filter paper and dried under a vacuum. The protected RNA bands were
visualized and quantified by PhosphorImager (Molecular Dynamics) and
normalized to the L-32 and GAPDH levels.
Western blotting. Cells were scraped in cold PBS, collected by centrifugation (10 min at 2,000 g), and lysed in Laemmli buffer (0.125 mM Tris · HCl, pH 6.8, containing 10% glycerol, 2.5% SDS, 0.006% bromphenol blue, and 0.1 M dithiothreitol) for 10 min at 100°C. The amount of total protein in the samples was quantified by dot-blot staining with amidoblack as previously described (28). Proteins (30 µg protein/lane) were separated by 7.5% SDS-PAGE and transferred to nitrocellulose membranes. The membranes were blocked for 1 h with 5% bovine serum albumin (BSA) buffer solution containing 0.1% Tween 20. The membrane was probed with polyclonal antibody to COX-2 peptide of human origin (1:10,000 dilution; Cayman Systems, Stoneham, MA) or polyclonal antibody to COX-1 peptide (1:1,000 dilution; Oxford Biomed, Oxford, MI), followed by peroxidase-conjugated donkey anti-rabbit IgG (1:10,000 dilution; Jackson Immunoresearch, West Grove, PA) as described previously (28). To normalize the COX proteins to a major housekeeping gene product, we reprobed the membranes with monoclonal antibodies against a highly conserved region of actin (1:20,000 dilution; Roche Molecular Biochemicals, Indianapolis, IN), followed by peroxidase-conjugated donkey anti-mouse IgG (1:10,000 dilution; Jackson Immunoresearch). The immunoreactive bands were visualized with the Renaissance chemiluminescence kit (NEN, Boston, MA) and quantified by the digital densitometry using NIH Image 1.60.
COX activity.
COX activity in whole endothelial cells was detected as production of
6-keto-PGF1 and PGE2 from exogenous AA as
described previously (28). Control or IL-1
-treated
cells were incubated with 10 µM AA in Krebs buffer (in mM: 5.0 KCl,
0.6 MgSO4, 1.8 CaCl2, 120 NaCl, 6.0 glucose,
and 10.0 HEPES, pH 7.4) for 15 min at 37°C. To investigate relative
activities of the COX isoforms, we pretreated cells for 20 min with
10
5 M NS-398 (a COX-2-selective inhibitor) or
10
5 M indomethacin (a nonselective COX inhibitor)
immediately before the detection of the COX activity. After the
incubation with AA, the medium was aspirated for prostanoid detection,
and cells were collected for protein determination. The amount of
6-keto-PGF1
and PGE2 in the medium was
detected by radioimmunoassay (28). COX activity was
normalized to the amount of total cell protein quantified by the
micro-BCA (bicinchoninic acid) protein assay from Pierce (Rockford, IL).
Antibodies for COX-1 and COX-2 immunostaining. For COX-2 immunostaining, the COX-2-specific antisera were prepared in rabbits by immunization with ovalbumin-conjugated COX-2 peptide according to standard techniques (10). As a COX-2-specific peptide, a unique 11-amino acid peptide (amino acids 582-592) at the COOH-terminal region of human COX-2 protein (2) was selected (100% difference from COX-1 of human origin). Anti-COX-2 antisera did not cross-react with the COX-1-specific peptide (human) or COX-1 protein (ovine). For immunofluorescence staining, anti-COX-2 antisera was used at 1:100 dilution. COX-2 (human) polyclonal antiserum (1:50 dilution; Cayman) was also used for immunostaining. COX-1-specific antisera were prepared in rabbits by immunization with ovalbumin-conjugated COX-1 peptide (10). As a COX-1-specific peptide, a unique 18-amino acid peptide (amino acids 582-599) at the COOH-terminal region of human COX-1 protein (41) was selected (88% difference from human COX-2). For immunofluorescence staining, anti-COX-1 antisera was used at 1:50 dilution. Anti-COX-1 antiserum did not cross-react with the COX-2-specific peptide (human) or COX-2 proteins of either ovine and human origin. Both COX-1 and COX-2 antisera have higher affinity to native vs. denatured cognate peptides and proteins and were used in immunostaining protocols.
Immunofluorescence.
For immunofluorescence studies, endothelial cells were grown on
Matrigel-covered glass coverslips. Confluent cells were starved overnight or treated with IL-1 (see IL-1
treatment). Cells rinsed with PBS were fixed with 3.7%
paraformaldehyde solution in PBS (pH 8.4; 15 min at room temperature)
and were permeabilized by 0.1% Triton X-100 solution in PBS (10 min at
room temperature). To block the nonspecific binding sites, we incubated
cells for 1 h at room temperature with PBS containing 5% BSA.
Cells were incubated with the COX isoform-specific primary polyclonal
antibody dissolved in PBS containing 0.5% BSA for 1 h at 37°C.
To visualize antigen-antibody complexes, we incubated cells with
FITC-conjugated anti-rabbit IgG (1:100 dilution; Vector Laboratories,
Burlingame, CA) for 1 h at 37°C (28). For negative
controls, cells were incubated with preimmune serum followed by
secondary antibody. As additional controls to determine the specificity
of the antigen detection, we used antisera preadsorbed with the
corresponding cognate peptide (1 mg/ml) before staining; neither of
these controls showed significant labeling. For F-actin detection,
coverslips with fixed/permeabilized endothelial cells were incubated
with Texas red-phalloidin (1:40 dilution; Molecular Probes) for 1 h at room temperature. To detect the endothelium-specific antigen, we
used polyclonal antibodies against von Willebrand factor (1:100 dilution; Sigma, St. Louis, MO). For cytosolic phospholipase
A2 (cPLA2) immunofluorescence, we used
monoclonal antibodies developed against the amino-terminal domain of
human cPLA2 (1:100 dilution; Santa Cruz Biotechnology,
Santa Cruz, CA), followed by FITC-conjugated anti-mouse IgG (1:100
dilution; Vector Laboratories). Coverslips were mounted on glass slides
using anti-fade mounting medium (Vector Laboratories).
Immunogold electron microscopy. Endothelial cells grown on Matrigel-covered cell culture plates were rinsed with PBS, gently scraped into PBS containing 4% formaldehyde and 0.1% glutaraldehyde, and precipitated for 10 min at 3,000 g. Precipitated cells were fixed in 4% formaldehyde and 0.1% glutaraldehyde in PBS for 1 h on ice. After dehydration, cells were embedded into L. R. White resin and sectioned. Immunogold labeling was performed as described previously (27). Silver-gray sections placed on nickel grids were blocked for 30 min at room temperature with 5% BSA in PBS containing 0.05% Tween 20 and 0.5 M NaCl (buffer A). The grids were incubated with primary antibody (polyclonal anti-COX-1 or anti-COX-2 as in Antibodies for COX-1 and COX-2 immunostaining) diluted 1:20 in buffer A containing 0.1% BSA for 60 min at 37°C, followed by incubation with 10-nm gold-conjugated anti-rabbit IgG (1:20 dilution; Electron Microscopy Sciences, Fort Washington, PA) for 60 min at 37°C. After each step, the grids were thoroughly washed in buffer A. Cells were postfixed in 2% glutaraldehyde in PBS for 20 min, briefly rinsed in H2O, and air dried. For contrasting, the grids were stained in 4% aqueous uranyl acetate and viewed in a Jeol 1200 electron microscope (Tokyo, Japan). Appropriate controls were maintained by omitting the primary antibody or by using preimmune serum; neither of these controls showed significant labeling.
Materials.
Cell culture reagents were purchased from Life Technologies
(Gaithersburg, MD), VEC Technologies, and Sigma. Percoll was from Amersham Pharmacia Biotech (Piscataway, NJ). Matrigel (growth factor
reduced) was from Becton Dickinson (Bedford, MA). Human recombinant
IL-1 was obtained from R&D Systems (Minneapolis, MN). All other
reagents were from Sigma unless otherwise indicated.
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RESULTS |
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COX-1 and COX-2 expression in quiescent endothelial cells.
COX-1 and COX-2 expression was detected by RT-PCR and Western
immunoblotting. Both COX-1 and COX-2 (mRNA and protein) were expressed
in quiescent endothelial cells from cerebral microvessels of porcine
and human origin and in primary HUVEC cultures (Fig. 1). COX-2 protein was also immunodetected
in quiescent endothelial cells using commercial antibodies from Cayman
(Fig. 1B). Although some variations in COX-1 and COX-2
expression were observed (Fig. 1B, representative samples of
cultured cells), all examined cells expressed both COX-1 and COX-2.
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COX-1 localization in quiescent endothelial cells by indirect
immunofluorescence.
COX-1 has two major sites of localization: the perinuclear zone
(including the nuclear envelope) and the cytoplasm (Fig.
2, A-F). COX-1 labeling
in the perinuclear zone had distinct asymmetrical distribution (Fig. 2,
A and F), especially evident in binuclear cells
(Fig. 2B). COX-1 immunofluorescence was also detectable in
the cytoplasm (Fig. 2, A, B, and D).
Although cytoplasmic COX-1 staining mainly had a granular character
(Fig. 2, B and D), a distinctive fibrillar
pattern was often observed, with fibrils spreading from the nuclear
envelope toward the cell periphery (Fig. 2, A and
E). We also observed a weak COX-1 immunofluorescence within
the nucleus (Fig. 2, A and E). COX-1 had an
identical pattern of intracellular localization in all three
endothelial cell types used in our study (in endothelial cells from pig
and human fetal brain and in HUVEC) (Fig. 2, A-F).
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COX-2 localization in quiescent endothelial cells by indirect
immunofluorescence.
COX-2 is localized mainly within the cell nucleus (Fig.
4, A-E) but not in the
nucleoli (Fig. 4, C and E). In some cells, a more
intense granular staining at the nuclear periphery was observed (Fig.
4E). Intranuclear localization of COX-2 was revealed by
using COX-2 antiserum developed in our laboratory (Fig. 4, A, C, and E) or commercial antibodies
from Cayman (Fig. 4, B and D). COX-2
immunostaining in the cytoplasm was also found, with the COX-2 granules
around the nucleus and at the cell periphery (Fig. 4, A and
B). Intranuclear localization of COX-2 was detected in
quiescent endothelial cells from piglet and human cerebral microvessels
and in HUVEC, both in primary cultures and at one to three passages.
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COX-1 and COX-2 immunofluorescence in cerebral microvascular
endothelial cells extracted with detergents.
We have previously demonstrated that a variety of endothelial proteins,
including cPLA2 and mitogen-activated protein kinases (extracellular signal-regulated kinases 1/2), are tightly
associated with the endothelial cytoskeleton (29). To
investigate whether COX-1 and COX-2 are detergent soluble, we extracted
cerebral microvascular endothelial cells with PBS containing 1% NP-40,
0.5% sodium deoxycholate, and 0.1% SDS for 30 min on ice. Although
~80% of total cell proteins were solubilized under these conditions,
the nuclei appeared largely intact under the microscope (Fig.
5, A and B). To
visualize the detergent-resistant scaffolding, we used Texas
red-conjugated phalloidin (Fig. 5, C and D). In
control quiescent cells, numerous F-actin stress fibers were observed
in the cytoplasm and along the cell periphery; multiple focal adhesions
were clearly seen (Fig. 5C). In cells extracted with
detergents, F-actin remained clearly seen as a granular and fibrillar
material around the nucleus but not at the cell-cell contact areas
(Fig. 5C).
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Nuclear localization sites for COX-1 and COX-2 in quiescent
endothelial cells.
We used immunogold electron microscopy to get a deeper insight into the
nuclear localization sites for COX-1 and COX-2. Our immunogold data
confirmed that in endothelial cells from the newborn pig cerebral
microvessels, COX-1 and COX-2 are nuclear residents (Figs.
7 and 8). Several COX localization sites
were revealed in the nucleus. Anti-COX-2 immunogold labeling was
present at the nuclear envelope, at the nuclear membranes, and in the
proximity of nuclear pores (Fig. 8,
asterisks), indicating possible transport places. Similarly, we
observed anti-COX-1 labeling at the nuclear envelope (Fig. 7,
arrowheads) and in the proximity of nuclear pores (Fig. 7, asterisks).
COX-1 and COX-2 labeling was also observed within the nucleus. As for
COX-1, only a few immunogold granules were observed within the cell
nucleus; the granules have a dispersed distribution, without any
connection with known intranuclear entities (Fig. 7). In contrast,
COX-2 labeling in the nucleus revealed a unique localization pattern,
with the immunogold granules observed mainly in the perichromatin zone
in connection with the border of condensed chromatin, both on the
periphery of the nucleus and around the nucleoli (Fig. 8, arrowheads).
It must be emphasized that practically all intranuclear anti-COX-2
immunogold labeling is associated with the clumps of opaquely condensed
chromatin. No COX-2 was observed in such intranuclear entities as
clusters of interchromatin granules and coiled bodies. Very few COX-2
immunogold particles were observed within the nucleoli in the areas
other than the border of condensed chromatin surrounding the nucleoli (Fig. 8).
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Time course of IL-1 effects on COX expression and activity.
Confluent human endothelial cells were exposed to human recombinant
IL-1
(50 ng/ml) for 2-20 h (Fig.
9). IL-1
selectively and transiently
stimulated COX-2 expression in HUVEC (Fig. 9) and in human cerebral
microvascular endothelial cells (data not shown). An 80- to 100-fold
induction in COX-2 mRNA observed in 1-4 h was followed by a slow
decline in the message level despite the continuous presence of IL-1
(Fig. 9). In contrast, only moderate (1.5- to 2-fold) elevation in
COX-1 mRNA was observed in IL-1
-treated cells (Fig. 9). Western
immunoblotting data demonstrated the linear increase in the amount of
COX-2 protein during 4-12 h of the stimulation; maximal (40- to
50-fold) induction at 12-16 h was followed by a gradual decline in
the protein level (Fig. 9). Time-dependent increases in prostanoid
accumulation in the media were detected during a 4- to 20-h period of
the IL-1
stimulation (Fig. 9). NS-398, a COX-2-selective inhibitor,
reduced IL-1
-stimulated COX activity by 90% (Fig.
10), thus indicating a major
contribution of COX-2 to induced prostanoid synthesis. Surprisingly,
the continuous rise in prostanoid production was extended far beyond
the plateau of the COX-2 protein accumulation (12-16 h) and was
observed even during its decline (16-20 h) (Fig. 9).
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Time-dependent effects of IL-1 on COX-2 and COX-1 localization.
To investigate whether the induced COX-2 expression is associated with
changes in spatial architecture of the COX, we investigated time-dependent effects of IL-1
(50 ng/ml) on COX-2
immunofluorescence in human cerebral microvascular endothelial cells.
In control untreated cells, COX-2 immunofluorescence was observed
mainly in the nucleus (Fig.
11A), consistent with our
previous observations. In the cells treated with IL-1
for 5 h,
a strong nuclear COX-2 immunofluorescence signal was supplemented with
the granular labeling diffusely distributed throughout the cytoplasm
(Fig. 11B). In 9-12 h of the treatment, multinuclear
endothelial cells were formed; at this stage, COX-2 immunofluorescence
was also localized mainly in the nucleus, although it was supplemented
by intense cytoplasmic labeling (Fig. 11, C and
D). Striking changes in COX-2 localization were observed in
endothelial cells treated with IL-1
for 17-23 h: COX-2
immunofluorescence was detectable almost exclusively in the nuclear
envelope and in the cytoplasm; only weak labeling still remained in the
nucleus (Fig. 11, E and F). COX-2 accumulation in
the nuclear envelope and in the cytoplasm was coincident with the
continuous rise in prostanoid synthesis, although COX-2 expression had
already reached its maximum. As for COX-1 localization, no changes in
COX-1 immunofluorescence were revealed in microvascular endothelial
cells stimulated by IL-1
for 10-14 h (Fig.
12, B and C). As
in control cells (Fig. 12A), COX-1 immunofluorescence
remained highly visible in the nuclear envelope and in the perinuclear area of the cytoplasm; a fibrillar pattern of COX-1 immunofluorescence was often present in the cytoplasm (Fig. 12, B and
C).
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DISCUSSION |
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Although COX-2 was discovered as an immediate-early gene inducible by growth factors and tumor promoters (3, 12, 13, 19, 25, 38), only recent experimental evidence has indicated that COX-2 is involved in the regulation of nuclear functions. COX-2 overexpression has been linked to the cell cycle progression, proliferation, colorectal cancer, and apoptosis (3, 13, 37, 44). Unknown mechanisms that mediate nuclear functions of COX are cell specific and, most importantly, are not necessarily mediated by prostanoids (3, 13, 22, 44). Increased proliferation of the colon cancer cells was sensitive to the COX inhibitors and coincident with the enhanced COX-2 expression (37). In human endothelial cells stimulated by inflammatory cytokines, induction of COX-2 and prostanoid synthesis was accompanied by inhibition of cell growth (19). Transient overexpression of COX-2 in a variety of cell types, including human and bovine endothelial cells, resulted in cell cycle arrest in S phase, changes in the nuclear architecture, and cell death; these effects were not reversed by the COX inhibitors (44). COX-1 overexpression caused transformation of immortalized HUVEC, leading to the formation of indomethacin-resistant tumors in vivo (22). Spatial organization of COX isoforms suggests its possible relation to the nuclear functions (5, 21, 22, 28, 33, 40).
The present study investigated expression and spatial architecture of the COX isoforms in endothelial cells from three different sources: cerebral microvessels of both human and porcine origin and human umbilical vein. These cell types were selected for several reasons. First, endothelium-derived prostanoids are physiological regulators of the blood flow to the brain in newborn pigs and human neonates (17, 18). Second, COX-1 and COX-2 are expressed in unstimulated cerebral microvascular endothelial cells from newborn piglets (28, 31), while no data in human neonates had been reported previously. Third, HUVEC express low levels of COX-2 under resting conditions (9). Using RT-PCR and Western immunoblotting, we have demonstrated here that COX-1 and COX-2 are expressed in quiescent endothelial cells from cerebral microvessels of perinatal porcine and human origins and from human umbilical vein.
We detected COX-1 localization by using antisera developed against a specific peptide at the carboxy terminus of human COX-1. An identical pattern of immunostaining was observed in porcine and human endothelial cells. The most intense COX-1 immunofluorescence was in the perinuclear zone of the cytoplasm, including the nuclear envelope. COX-1 was also observed in the endoplasmic reticulum area marked by von Willebrand factor (36). The nuclear envelope and endoplasmic reticulum have been reported as COX-1 localization sites in other cell types, including 3T3 fibroblasts (33-35, 40), monocytes (40), megakaryoblasts (45), HUVEC (40, 45), and astrocytes (43). In addition, we have described two novel COX-1 sites in endothelial cells: 1) cytoplasmic fibrils and 2) nuclear sites. A fibrillar pattern of COX-1 immunofluorescence was often observed, with fibrils expanding from the nuclear envelope toward the cell periphery. This finding may indicate COX-1 association with the cytoskeletal filaments. As revealed by immunogold electron microscopy, the nucleus has multiple COX-1 localization sites that include not only the nuclear envelope but also nuclear pores and the nucleoplasm. COX-1 immunogold particles were diffusely distributed in the nucleoplasm with no connection to known nuclear domains.
We detected COX-2 localization by using antisera developed against a unique peptide at the carboxy terminus of human COX-2. In quiescent endothelial cells from newborn pigs, COX-2 is localized mainly within the nucleus, with much less COX-2 immunofluorescence in the cytoplasm. This confirms our previous findings in porcine endothelial cells using commercial COX-2 antiserum (28). We now report that the nucleus is also a major COX-2 localization site in quiescent endothelial cells from perinatal human cerebral microvessels and in HUVEC in primary culture and at one to three passages. Immunogold electron microscopy revealed multiple nuclear COX-2 sites. COX-2 immunogold labeling was accumulated in the nuclear envelope at both inner and outer nuclear membranes and in proximity with the nuclear pores. Within the nucleus, a very distinct pattern of COX-2 distribution was observed. Intranuclear COX-2 was found in the perichromatin zone in connection with the border of condensed chromatin, both on the periphery of the nucleus and around the nucleoli. Perichromatin zones containing perichromatin fibrils on the periphery of condensed chromatin are structural domains of the nucleoplasm that represent the sites of active transcription and RNA processing (6, 20). The nucleolus, considered the rRNA synthesis/processing nuclear compartment (6), did not show COX-2 immunolabeling. COX-2 immunogold granules were not observed in other nuclear domains such as clusters of interchromatin granules and coiled bodies that function as the storage sites for components of the transcription/splicing machinery (7). The specific pattern of intranuclear distribution indirectly indicates that COX-2 could bind to the sites of RNA synthesis and processing in the perichromatin zones.
COX-2 appears to be associated with the nuclear matrix. This is indicated by our observations that nuclear COX-2 labeling is resistant to ionic detergents. The nuclear matrix is a detergent-resistant nonchromatin scaffolding of the ribonucleoprotein network that is connected to the inner nuclear lamina (24). The nuclear matrix spatially organizes chromatin and has been implicated in essential nuclear activities, such as transcription, replication, and regulation of gene expression (24, 30). Nuclear matrix proteins that include various transcription factors are tissue and cell specific (23, 30, 42). Hormone receptors and components of signal transduction mechanisms, such as protein kinase C, are also found in association with the nuclear matrix (23, 42). Our finding that COX-2 is associated with the nuclear matrix, especially when considered in a context of COX-2 localization in the perichromatin zone, may suggest that COX could participate in regulation of gene expression. In contrast, intranuclear COX-1 was removed by detergent treatment, whereas COX-1 in the nuclear envelope was detergent resistant. Because the nuclear lamina is a part of the nuclear matrix (23, 24, 30, 42), it appears that COX-1 also could be a nuclear matrix component.
We investigated whether the transcriptional activation of endothelial
cells is associated with intracellular redistribution of COX-2.
Proinflammatory cytokines, such as IL-1, induce immediate-early response genes, including COX-2, in a variety of cell types (11, 19, 15). We found that transcriptional activation and COX-2 induction in response to IL-1 are accompanied by COX-2 trafficking between the nucleus and the cytoplasm in human endothelial cells. IL-1
rapidly and transiently upregulated COX-2 mRNA, indicating transcriptional activation of endothelial cells. At the early stages of
the IL-1
stimulation (5-9 h) marked by increased
transcriptional activity, the nucleus remained a major COX-2
localization site. By this time, COX-2 protein was induced 20- to
30-fold higher than the control level. Because the cytoplasm is a site
of a new protein synthesis, COX-2 accumulation in the nuclei is
indicative of active COX-2 nuclear import in transcriptionally
activated cells. In cells exposed to IL-1
for 12 h, COX-2
accumulation in the cytoplasm was clearly observed, although the
nucleus remained a major COX-2 localization site. At this time point,
COX-2 mRNA level started to decline, reflecting a decrease in
transcriptional activity, whereas COX-2 protein reached its maximum
(50-fold higher than control level), indicating continuing
translational activation. Decline in the message and protein levels at
later stages of the IL-1
stimulation (17-23 h) was coincident
with COX-2 accumulation at the nuclear envelope and in the cytoplasm;
practically no COX-2 immunofluorescence in the nucleus was observed at
that time.
COX-2 accumulation in the cytoplasm and nuclear envelope in 12-23
h of IL-1 stimulation was associated with a steep increase in
prostanoid synthesis, although COX-2 protein expression was then
decreasing. Induced COX activity was completely abolished by NS-398,
indicating COX-2 contribution. Because cPLA2 is also localized in the nuclear envelope and the cytoplasm, it is possible that the formation of spatially efficient complexes between the two key
enzymes is essential for maximal prostanoid synthesis. However, when AA
was used to bypass phospholipase, a major increase in COX activity was
also observed in human endothelial cells stimulated with IL-1
for
17-24 h. This may indicate COX-2 activation in the cytoplasm.
Indeed, in porcine brain endothelial cells, COX-2 activity can be
rapidly increased by tyrosine phosphorylation (26). In
human endothelial cells stimulated by IL-1
, no changes in COX-1
expression, activity, and distribution were observed. COX-1 remained a
perinuclear and a cytoplasmic resident; both granular and fibrillar
patterns of COX-1 immunofluorescence were clearly seen. Under no
conditions did we observe COX-1 accumulation within the nucleus.
COX-2 accumulation in the nucleus at early stages of IL-1
stimulation is indicative of the nuclear import of a newly synthesized COX-2 coincidentally with the transcriptional activation of endothelial cells. Nuclear depletion of COX-2 along with its redistribution to the
nuclear envelope and the cytoplasm at the later stages may indicate
inhibition of the nuclear import and/or activation of nuclear export of
COX-2. Transport through the nuclear pore complexes is a leading
mechanism of the nuclear protein import (1, 14).
Accumulation of the labeled proteins at the nuclear pores is a distinct
morphological marker of the nuclear transport. Our electron microscopy
data revealed COX-2 immunogold granule accumulation near the nuclear
pores. We also detected COX-1 immunogold granules in proximity to the
nuclear pores. Therefore, it appears that COX-1 can be also transported
into the nucleus. Because both COX-1 and COX-2 are integral membrane
proteins, translocation to the nucleus would require some drastic
modifications of the monotopic membrane-binding domains.
In conclusion, our findings in vascular endothelial cells of porcine
and human origin are as follows: 1) COX-1 and COX-2 have nuclear and cytoplasmic localization sites; 2) nuclear sites
for COX-1 and COX-2 include the nuclear envelope, nuclear pores, and the nucleoplasm; 3) in the nucleoplasm, COX-2 is localized
in the perichromatin zone, recognized as the site of active
transcription, whereas COX-1 has a diffuse distribution; 4)
accumulation of a newly synthesized COX-2 within the nucleus was
coincident with the transient transcriptional activation of endothelial
cells with IL-1; 5) decrease in transcriptional activity
at later stages of the IL-1
-stimulation was accompanied by COX-2
relocation to the cytoplasm/nuclear envelope and the enzyme activation;
and 6) COX-1 activity and localization were not altered in
IL-1
-stimulated endothelial cells. Together, these results suggest
that the potential role for COX-2 in the regulation of the nuclear
functions and transcriptional activation should be further investigated.
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ACKNOWLEDGEMENTS |
---|
We thank D. Harder, J. Narayanan, and M. Aebly from the Medical College of Wisconsin for help in designing COX-1 and COX-2 peptides, Alex Fedinec and Mildred Jackson for excellent technical support, Donna Davis and Katherine J. Troughton for help in immunogold electron microscopy, and D. Morse for the illustrations.
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FOOTNOTES |
---|
This research was supported by the National Institutes of Health, the National Heart, Lung, and Blood Institute, and the Southeast Affiliate of the American Heart Association.
Address for reprint requests and other correspondence: H. Parfenova, Dept. of Physiology, Univ. of Tennessee Health Science Center, 894 Union Ave., Memphis, TN 38163 (E-mail: hparf{at}physio1.utmem.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 16 November 2000; accepted in final form 16 February 2001.
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