Dynamics of nuclear localization sites for COX-2 in vascular endothelial cells

Helena Parfenova1, Vladimir N. Parfenov3, Boris V. Shlopov2, Vladimir Levine1, Sheryl Falkos1, Massroor Pourcyrous1, and Charles W. Leffler1

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


    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)-1beta , 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-1beta 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-1beta ; nucleus


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ABSTRACT
INTRODUCTION
METHODS
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DISCUSSION
REFERENCES

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)-1beta -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-alpha (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-1beta .


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INTRODUCTION
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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-1beta 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-1beta (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-1beta 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 beta -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 alpha -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 alpha -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-PGF1alpha and PGE2 from exogenous AA as described previously (28). Control or IL-1beta -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-PGF1alpha 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-1beta (see IL-1beta 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).

To investigate the association of the COX isoforms with the detergent-resistant cytoskeleton, we extracted endothelial cells by gentle agitation with PBS containing 1% Nonidet P-40 (NP-40), 0.5% sodium deoxycholate, and 0.1% SDS for 30 min on ice and rinsed them with PBS. Our data indicate that ~80% of total cell proteins were solubilized. The fraction of detergent-resistant endothelial cytoskeleton that remained attached to coverslips after the extraction was fixed and immunostained for the COX isoforms and F-actin as described above.

Slides were viewed with the use of an Image Deconvolution System consisting of a Nikon Diaphot microscope with fluorescein and rhodamine filters coupled to a Macintosh Quadra 950 computer system with a Power Mac processor 601. Five consequent images were taken from each slide. Images were deconvolved by using a Vaytech software for deconvolution and IPLab Spectrum software for image collection in conjunction with a cooled charge-coupled device camera (model 250 CH, Photometric). Digital processing of the images was done using Adobe Photoshop (Adobe Systems).

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-1beta was obtained from R&D Systems (Minneapolis, MN). All other reagents were from Sigma unless otherwise indicated.


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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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Fig. 1.   Cyclooxygenase (COX-1 and COX-2) detection by RT-PCR (A) and Western immunoblotting (B) in primary cultures of cerebral microvascular endothelial cells from newborn piglets (pCMVEC) or human brain (hCMVEC) and in human umbilical vein endothelial cells (HUVEC). Representative samples are shown.

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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Fig. 2.   COX-1 localization by indirect immunofluorescence in cerebral microvascular endothelial cells from newborn piglet brain (A) and human brain (B and D) and in HUVEC (C, E, and F). Quiescent cells were immunostained with anti-COX-1 antiserum raised against a human COX-1-specific peptide (1:50 dilution). As a control, preimmune rabbit serum (1:50 dilution) was used instead of primary antibodies (C). Bar, 20 µm.

We compared the COX-1 localization pattern with the distribution of cPLA2 in cerebral microvascular endothelial cells from newborn pigs. cPLA2 immunofluorescence was detected as a bright ring around the nucleus (nuclear envelope) and as a granular pattern in the cytoplasm (Fig. 3A). No cPLA2 immunofluorescence was observed in the nucleus or at the plasma membrane (Fig. 3A). Von Willebrand factor, an endothelium-specific antigen residing in the endoplasmic reticulum (36), showed exclusively cytoplasmic immunofluorescence, with many large granules symmetrically distributed around the nucleus in mono- and binuclear endothelial cells from cerebral microvessels (Fig. 3B).


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Fig. 3.   Cytosolic phospholipase A2 (cPLA2) and von Willebrand factor localization by indirect immunofluorescence in cerebral microvascular endothelial cells from newborn piglets. Quiescent cells were immunostained with monoclonal anti-cPLA2 (1:100 dilution; Santa Cruz) (A) or polyclonal anti-von Willebrand (1:100 dilution; Sigma) (B). Bar, 20 µm.

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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Fig. 4.   COX-2 localization by indirect immunofluorescence in cerebral microvascular endothelial cells from newborn piglet brain (A and F) and human brain (B and D) and in HUVEC (C and E). Quiescent cells were immunostained with anti-COX-2 antiserum raised against a human COX-2-specific peptide (1:100 dilution) (A, C, and E) or with anti-COX-2 (1:50 dilution; Cayman) (B and D). In the control (F), the COX-2 antiserum preadsorbed with the cognate peptide was used. Bar, 20 µm.

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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Fig. 5.   Effect of detergent extraction on cerebral microvascular endothelial cells from newborn piglets. Quiescent cells were untreated (A and C) or extracted with PBS containing 1% Nonidet P-40 (NP-40), 0.5% sodium deoxycholate, and 0.1% SDS for 30 min on ice (B and D). A and B: phase-contrast images. C and D: F-actin staining with Texas red-phalloidin (1:40 dilution). Bar, 20 µm.

In detergent-extracted endothelial cells, COX-2 immunofluorescence in the nucleus remained clearly seen, while cytoplasmic labeling was eliminated (Fig. 6D). Although COX-1 immunofluorescence was removed from the cytoplasm (Fig. 6B), COX-1 in the nuclear envelope area appeared to be partially resistant to the detergent treatment (Fig. 6B).


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Fig. 6.   Effect of detergent extraction on COX-1 and COX-2 immunofluorescence in cerebral microvascular endothelial cells from newborn piglets. Quiescent cells were untreated (A and C) or extracted with PBS containing 1% NP-40, 0.5% sodium deoxycholate, and 0.1% SDS for 30 min on ice (B and D). Cells were immunostained with anti-COX-1 antiserum raised against a specific peptide at the carboxy-terminal region of human COX-1 (1:50 dilution) (A and B) or with anti-COX-2 antiserum raised against a human COX-2-specific peptide (1:100 dilution) (C and D). Bar, 20 µm.

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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Fig. 7.   Anti-COX-1 immunogold labeling in quiescent cerebral microvascular endothelial cells from newborn piglets. Shown is a part of the endothelial cell with the nucleus (N) and the cytoplasm (C) after immunogold labeling with anti-COX-1 antiserum raised against a human COX-1-specific peptide (1:20 dilution). Note the presence of anti-COX-1 labeling in the nuclear envelope (arrowheads) at both the outer and inner nuclear membranes. Also note the aggregation of label in zone of nuclear pores (asterisks). Original magnification, ×60,000.



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Fig. 8.   Anti-COX-2 immunogold staining in quiescent cerebral microvascular endothelial cells from newborn piglets. Note the presence of anti-COX-2 labeling in the nuclear envelope (NE) at both the outer and inner nuclear membranes. Also note the aggregation of label in the zone of nuclear pores (asterisks). Note the localization of the label in the nucleus (N) on the boundary of the condensed chromatin (CH) at the peripheral part of the nucleus (left) and around the nucleoli (Nu) (arrowheads). No labeling is observed in the Nu. For immunogold labeling, anti-COX-2 antiserum raised against a human COX-2-specific peptide was used at a 1:20 dilution. Original magnification, ×50,000.

Time course of IL-1beta effects on COX expression and activity. Confluent human endothelial cells were exposed to human recombinant IL-1beta (50 ng/ml) for 2-20 h (Fig. 9). IL-1beta 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-1beta (Fig. 9). In contrast, only moderate (1.5- to 2-fold) elevation in COX-1 mRNA was observed in IL-1beta -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-1beta stimulation (Fig. 9). NS-398, a COX-2-selective inhibitor, reduced IL-1beta -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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Fig. 9.   Effect of interleukin (IL)-1beta on COX expression and prostanoid production in HUVEC. Cells (passage 3) were incubated with starvation medium (0.1% FBS-DMEM) for 15 h to achieve quiescence. After this period, human IL-1beta (50 ng/ml) was added to the starvation medium for the time intervals indicated. COX-1 and COX-2 mRNA were detected by RNase protection assay (15 µg RNA/lane) (A), quantitated by digital densitometry, and normalized to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (C, triangle ). COX-2 protein was detected by Western blot (30 µg protein/lane) (B), quantitated by digital densitometry, and normalized to actin (C, ). Prostanoid (6-keto-PGF1alpha ) accumulation in the medium was detected by radioimmunoassay (C, open circle ) (28).



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Fig. 10.   Effect of NS-398 on COX activity in HUVEC. Cells (passage 1) were incubated with starvation medium (0.1% FBS-DMEM) for 15 h to achieve quiescence. Human IL-1beta (50 ng/ml) was added to the starvation medium for an additional 0 (control) and 20 h. COX activity was detected as prostanoid production from 10 µM arachidonic acid (28). To determine the COX-2 contribution to the total enzyme activity, we pretreated cells for 20 min with NS-398 (10-5 M) or indomethacin (10-5 M). Concentrations of 6-keto-PGF1alpha (A) and PGE2 (B) were determined by radioimmunoassay (28).

Time-dependent effects of IL-1beta 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-1beta (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-1beta 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-1beta 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-1beta 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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Fig. 11.   Time-dependent effects of IL-1beta on COX-2 localization in human cerebral microvascular endothelial cells. Cells (passage 2) were incubated with starvation medium (0.1% FBS-DMEM) for 15 h to achieve quiescence. Human IL-1beta (50 ng/ml) was added to the starvation medium for an additional 0 (A, control), 5 (B), 9 (C), 12 (D), 17 (E), and 23 h (F). Control and treated cells were immunostained with polyclonal anti-COX-2 (1:50 dilution; Cayman). Note the presence of bright nuclear staining in control (A) and in cells treated with IL-1beta for 5-12 h (B-D). After longer durations of the treatment, immunofluorescence accumulated mainly in the nuclear envelope and in the cytoplasm (E and F). Bar, 20 µm.



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Fig. 12.   COX-1 localization in IL-1beta -treated human cerebral microvascular endothelial cells. Cells (passage 2) were incubated with starvation medium (0.1% FBS-DMEM) for 15 h to achieve quiescence. After this period, human IL-1beta (50 ng/ml) was added to the starvation medium for an additional 0 (A, control), 10 (B), and 14 h (C). Control and treated cells were immunostained with anti-COX-1 antiserum raised against a human COX-1-specific peptide (1:50 dilution).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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-1beta are accompanied by COX-2 trafficking between the nucleus and the cytoplasm in human endothelial cells. IL-1beta rapidly and transiently upregulated COX-2 mRNA, indicating transcriptional activation of endothelial cells. At the early stages of the IL-1beta 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-1beta 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-1beta 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-1beta 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-1beta 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-1beta , 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-1beta 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-1beta ; 5) decrease in transcriptional activity at later stages of the IL-1beta -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-1beta -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.


    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.


    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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