Copyright ©The Histochemical Society, Inc.

Despite Transcriptional and Functional Coordination, Cyclooxygenase-2 and Microsomal Prostaglandin E Synthase-1 Largely Reside in Distinct Lipid Microdomains in WISH Epithelial Cells

William E. Ackerman, IV, John M. Robinson and Douglas A. Kniss

Department of Obstetrics and Gynecology (Laboratory of Perinatal Research and Division of Maternal-Fetal Medicine) (WEA,DAK), Center for Biomedical Engineering (DAK), and Department of Physiology and Cell Biology (JMR), The Ohio State University, Columbus, Ohio

Correspondence to: Douglas A. Kniss, Laboratory of Perinatal Research, Department of Obstetrics and Gynecology, The Ohio State University, 5th Floor Means Hall, 1654 Upham Drive, Columbus, OH 43210. E-mail: kniss.1{at}osu.edu


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Cytokine-induced prostaglandin (PG)E2 synthesis requires increased expression of cyclooxygenase-2 (COX-2) in human WISH epithelial cells. Recently, an inducible downstream PGE synthase (microsomal PGE synthase-1, mPGES-1) has been implicated in this inflammatory pathway. We evaluated cooperation between COX-2 and mPGES-1 as a potential mechanism for induced PGE2 production in WISH cells. Cytokine stimulation led to increased expression of both enzymes. Selective pharmacological inhibition of these enzymes demonstrated that induced PGE2 release occurred through a dominant COX-2/mPGES-1 pathway. Unexpectedly, immunofluorescent microscopy revealed that the expression of these enzymes was not tightly coordinated among cells after cytokine challenge. Within cells expressing high levels of both mPGES-1 and COX-2, immunolabeling of high-resolution semithin cryosections revealed that COX-2 and mPGES-1 were largely segregated to distinct regions within continuous intracellular membranes. Using biochemical means, it was further revealed that the majority of mPGES-1 resided within detergent-insoluble membrane fractions, whereas COX-2 was found only in detergent-soluble fractions. We conclude that although mPGES-1 and COX-2 show transcriptional and functional coordination in cytokine-induced PGE2 synthesis, complementary morphological and biochemical data suggest that a majority of intracellular mPGES-1 and COX-2 are segregated to discrete lipid microdomains in WISH epithelial cells. (J Histochem Cytochem 53:1391–1401, 2005)

Key Words: inflammation • cytokines • prostaglandin E2 • cyclooxygenase-2 • microsomal prostaglandin E • synthase-1 • lipid microdomains • epithelial cells


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AMONG THE FIRST OF THE EICOSANOIDS to be characterized, prostaglandin (PG)E2 is a ubiquitous lipid mediator regulating homeostatic and inflammatory functions in several major human systems (including reproductive, gastrointestinal, renal, neuroendocrine, and immune) (Serhan and Levy 2003Go). The biosynthesis of PGE2 involves the functional coordination of three major enzymatic reactions involving phospholipase A2, cyclooxygenase (COX), and PGE synthase (PGES). In this pathway, arachidonic acid liberated from membrane phospholipids by isoforms of phospholipase A2 is converted to intermediates (PGG2 and PGH2) by COX-1 or COX-2, which are subsequently isomerized by PGES isoforms to PGE2.

COX catalyzes the committing and rate-limiting step in PGE2 synthesis. COX-1 is constitutively expressed and contributes to immediate, low-amplitude PG release (Smith et al. 2000Go). The inducible COX-2 isoform is essential for high amplitude, delayed PG production, as is often seen under inflammatory conditions (Smith et al. 2000Go). Several distinct gene products bearing PGES activity have been cloned and characterized. As with COX-2, the expression of microsomal PGES-1 (mPGES-1, a member of the membrane associated proteins involved in eicosanoid and glutathione metabolism superfamily) is induced in response to cytokines and other inflammatory stimuli (Jakobsson et al. 1999Go; Forsberg et al. 2000Go; Thoren and Jakobsson 2000Go; Stichtenoth et al. 2001Go). In contrast, cytosolic PGES (cPGES) and microsomal PGES-2 (mPGES-2) show only limited inducibility in response to inflammatory mediators (Stichtenoth et al. 2001Go; Claveau et al. 2003Go; Puxeddu et al. 2003Go; Giannico et al. 2005Go). It has further been demonstrated that these PGES isoforms contribute unequally to the generation of PGE2 under inflammatory conditions. Cotransfection studies suggest that mPGES-1 is functionally coupled with COX-2 during delayed PG release (Murakami et al. 2000Go), whereas cPGES is preferentially association with COX-1 (Tanioka et al. 2000Go; Han et al. 2002Go). Microsomal PGES-2 appears to be COX nonselective, and current evidence suggests that the contribution of mPGES-2 to high-amplitude PGE2 release through COX-2 is limited (Watanabe et al. 1997Go; Murakami et al. 2003Go).

PGE2 is generated at sites of inflammation in substantial amounts. Because newly synthesized PGs are released soon after they are produced, cells must be functionally adapted to generate substantial amounts of PGE2 in a relatively short time. We have previously demonstrated that PGE2 production in response to pro-inflammatory cytokines (e.g., interleukin 1ß [IL-1ß] and tumor necrosis factor-{alpha}) requires increased COX-2 expression in WISH epithelial cells (Albert et al. 1994Go; Perkins and Kniss 1997bGo). We hypothesized that cooperation between COX-2 and mPGES-1 might serve as an inducible pathway to facilitate this inflammatory cascade. In the present study, we used WISH cells to investigate the induced expression and subcellular localization of mPGES-1 in relation to COX-2. Furthermore, results from recent studies have indicated that a fraction of induced COX-2 may be localized to caveolar structures and may interact with caveolin-1 (Cav-1) (Liou et al. 2000Go,2001Go). As it has been speculated that PGE synthase isoforms could also be localized to these specific areas (Liou et al. 2001Go), we tested whether mPGES-1 and COX-2 were coordinated in Cav-1–containing lipid rafts.


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Cell Culture
Human WISH epithelial cells were obtained from the American Type Culture Collection (CCL-25) and maintained in Ham's F-12/DMEM (Invitrogen; Carlsbad, CA) supplemented with 2 mM L-glutamine, 1 mM sodium pyruvate, and 10% newborn calf serum. Cells were grown at 37C in an atmosphere of 95% air/5% CO2 and used for experiments between the 3rd and 25th passages.

Northern Blot Analysis
Total RNA was extracted using TRIZOL (Invitrogen) and prepared for Northern blotting as previously described (Ackerman et al. 2004Go). Bound transcripts were identified using digoxigenin-labeled cDNA probes (Roche Diagnostics; Indianapolis, IN). A 1.8-kb cDNA fragment encoding human COX-2 (a kind gift from Dr. Timothy Hla, University of Connecticut, Farmington, CT), a 0.4-kb cDNA fragment corresponding to the coding region of human mPGES-1 (Cayman Chemical; Ann Arbor, MI), and a 0.6-kb cDNA fragment encoding human glyceraldehyde-3-phosphate dehydrogenase were used for detection. Chemiluminescent signals were detected using the VersaDoc Imaging System and analyzed using Quantity One software (Bio-Rad Laboratories; Hercules, CA).

Immunoblot Analysis
Cellular proteins were extracted in PBS containing 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, 1 mM PMSF, and 10 µg/ml each of leupeptin, aprotinin, and antipain. Proteins (30 µg/lane) were resolved by SDS-PAGE and transferred to nitrocellulose. Nonspecific binding was blocked by incubation in Tris-buffered saline (pH 8.0) containing 0.1% Tween-20 (Sigma; St Louis, MO) and 5% nonfat dry milk. Membranes were then probed with polyclonal antibodies directed against mPGES-1 (160140; Cayman Chemical) or COX-2 (sc-1745; Santa Cruz Biotechnology, Santa Cruz, CA). After washing, the membranes were exposed to horseradish peroxidase-conjugated secondary antibodies and immune complexes were revealed by enhanced chemiluminescence. As a control for loading precision, membranes were reprobed with a monoclonal antibody directed against glyceraldehyde-3-phosphate dehydrogenase (MAB374; Chemicon International, Temecula, CA). Immunoreactive proteins were visualized using the VersaDoc Imaging System and analyzed using Quantity One software (Bio-Rad).

Immunofluorescence
For whole-mount specimens, WISH cells were grown in monolayer culture on flame-sterilized glass cover slips. After stimulation with 10 ng/ml of recombinant human IL-1ß (R & D Systems; Minneapolis, MN) for 6 hr, cells were fixed for 1 hr in 4% paraformaldehyde PBS and made permeable by the addition of 0.2% Triton X-100 (Sigma) in PBS for 15 min at room temperature. Alternatively, cells were incubated in the presence of 0.05% saponin (Sigma) as a means for permeabilization (Goldenthal et al. 1985Go). Cells were blocked in 1% nonfat dry milk/5% normal goat serum in PBS before the addition of antibodies. For colocalization experiments, rabbit polyclonal antibodies directed against mPGES-1 (160140; Cayman Chemical) were applied simultaneously with mouse monoclonal COX-2 antibodies (clone 33; BD Biosciences, San Jose, CA). To control for specificity, antibodies were preincubated with a 10-fold excess of either blocking peptide (for mPGES-1, Cayman Chemical) or recombinant human COX-2 (Oxford Biomedical Research; Oxford, MI) before application. Additional negative controls were conducted in which primary antibodies were omitted. After stringent washing in PBS, the cover slips were exposed to Alexa Fluor–conjugated secondary antibodies (Molecular Probes; Eugene, OR) and nuclei were counterstained with 5 µg/ml 4',6-diamidino-2-phenylindole (DAPI; Sigma). Specimens were then mounted on glass slides using the ProLong Antifade Kit (Molecular Probes) and visualized using standard epifluorescence (Nikon Instruments; Melville, NY) or confocal microscopy (Zeiss 510 META; Carl Zeiss Inc., Thornwood, NY). Images from whole-mount specimens were captured with MetaVue version 5.0 image analysis software (Universal Imaging Corp.; Downingtown, PA). Maximum fluorescence intensity measurements for structures labeled with anti-COX-2 and anti–mPGES-1 in each cell were obtained using the line scan function drawn through the long axis of the cell. Background fluorescence adjacent to each positive structure was subtracted from the peak fluorescence to obtain a fluorescence intensity measurement for each fluorochrome in each cell.

In tandem experiments, semithin cryosections were prepared for immunocytochemistry as previously described (Takizawa et al. 2003Go), with modifications. Briefly, confluent monolayer cultures grown in 175 cm2 flasks and stimulated for 6 hr with IL-1ß were fixed for 1 hr with 4% paraformaldehyde in 0.1 M sodium cacodylate (pH 7.4)/5% sucrose. Specimens were washed, collected by scraping, centrifuged briefly to pellet, and embedded in 10% gelatin made with 0.1 M sodium cacodylate (pH 7.4)/5% sucrose. Within the solidified gelatin, cell pellets were cut into small pieces, infiltrated with 2.3 M sucrose in 0.1 M sodium cacodylate (pH 7.4), and mounted on specimen pins. Specimens were then stored in liquid N2 until the time of sectioning. Semithin sections (~400 nm) were cut using a cryoultramicrotome, collected on droplets of 2 M sucrose containing 0.75% gelatin, and transferred to glass cover slips coated with 2% of 3-aminopropyltriethoxysilane (Sigma). Specimens were then exposed to anti–COX-2 and anti–mPGES-1 antibodies as before.

Prostaglandin E2 Enzyme Immunoassay
PGE2 content of culture media was quantified using a commercially available enzyme immunoassay (EIA) kit, according to the instructions of the manufacturer (Cayman Chemical). All treatments were conducted in the presence of 5 µM exogenous arachidonic acid (Cayman Chemical). The intra- and interassay coefficients of variation were <10%. Cells were lysed using 1 N NaOH and analyzed for protein content. PGE2 content was then normalized to total protein for each sample.

Preparation of Detergent-resistant Membrane Fractions
Detergent-resistant membrane (DRM) fractions were prepared by flotation on a sucrose step gradient as described previously (Liou et al. 2001Go), with modifications. After treatment with IL-1ß (10 ng/ml) or vehicle for 6 hr, confluent WISH cells were washed in ice-cold PBS and collected by scraping and centrifugation at 400 x g for 5 min at 4C. Cell pellets were lysed in ice-cold 50 mM Tris-HCl (pH 7.4), 150 mM NaCl, 5 mM EDTA buffer (TNE) containing 1% Triton X-100, 1 mM PMSF, and 10 µg/ml each of leupeptin, aprotinin, and antipain. Homogenization was achieved using 10 strokes of a loose-fitting Dounce homogenizer. The 0.6 ml homogenate was adjusted to 40% sucrose with the addition of 1.4 ml of 56% sucrose prepared in TNE buffer. This was placed in the bottom of an ultracentrifuge tube and overlaid with equal volumes of TNE buffer containing 30% and 5% sucrose, respectively. After centrifugation at 250,000 x g for 20 hr at 4C, nine 600-µl fractions from top to bottom were collected. Equivalent volumes from these fractions were separated by SDS-PAGE, transferred to nitrocellulose, and analyzed for COX-2, mPGES-1, and Cav-1 content by immunoblotting. The chicken anti-peptide Cav-1 antibody was described previously (Lyden et al. 2002Go).

Statistical Analysis
EIA data were expressed as PGE2 produced (ng/mg of protein) and represent the means ± SEM for quadruplicate determinations, which were repeated twice. The data were assessed by one-way ANOVA followed by the Tukey-Kramer multiple comparisons post hoc test. Spearman rank correlation analysis was used to assess the relation between COX-2–associated fluorescence intensity and that of mPGES-1 in a representative population of cells. Simple linear regression was used to model the relationship between these two variables. A p value of less than 0.05 was considered significant.


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Cytokines Induce the Expression of mPGES-1 and COX-2
Challenge of WISH cells with 10 ng/ml of recombinant human IL-1ß elicited increased expression of both COX-2 and mPGES-1. In Northern blots, a persistent 50-fold increase in the expression of the major 4.5 kb COX-2 transcript was observed (Figure 1A), which was consistent with previous studies (Albert et al. 1994Go). Concomitantly, a 2-fold increase in mPGES-1 mRNA expression was noted (Figure 1A). Levels of mPGES-1 mRNA rose above baseline by 1 hr poststimulation and peaked between 4 and 8 hr, declining thereafter. Although COX-2 mRNA was practically undetectable in unstimulated cells, the mPGES-1 message was readily detectable at 0 hr, indicating a limited amount of constitutive expression. In the presence of an inhibitor of RNA synthesis (10 µg/ml actinomycin D, a dose chosen based on preliminary experiments to give complete inhibition of RNA synthesis), COX-2 expression in response to two cytokines (IL-1ß or tumor necrosis factor-{alpha}) was significantly but incompletely attenuated (Figure 1B). In contrast, actinomycin D reduced cytokine-elicited mPGES-1 expression to levels observed in nonstimulated cells (Figure 1B, right panel, Lanes 4–6). These results suggest a limited role for transcript stabilization in COX-2, but not mPGES-1, mRNA accumulation.



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Figure 1

Cytokine stimulation increases cyclooxygenase-2 (COX-2) and microsomal prostaglandin E synthase-1 (mPGES-1) expression. (A) Northern blot analysis of COX-2 and mPGES-1 mRNA in WISH cells treated with IL-1ß (with glyceraldehyde-3-phosphate dehydrogenase as a loading control). (B) Effect of actinomycin D (ActD) on cytokine-induced COX-2 and mPGES-1 mRNA expression at 1 and 4 hr after stimulation, respectively. Images of 28S and 18S rRNA (rRNA) bands demonstrate equivalent loading. C = control (vehicle only); I = IL-1ß treatment; T = tumor necrosis factor-{alpha} treatment. (C) Immunoblot analysis of COX-2 and mPGES-1 in WISH cells after stimulation IL-1ß (with glyceraldehyde-3-phosphate dehydrogenase as a control). These blots are representative of at least three separate experiments.

 
In Western blots, low levels of COX-2 protein were evident in unstimulated cells, and expression was increased 10-fold in response to IL-1ß addition (Figure 1C). Peak COX-2 immunoreactivity was attained 8 hr posttreatment and persisted through 24 hr. IL-1ß–elicited increases in mPGES-1 immunoreactivity paralleled that of COX-2, with maximal levels of both enzymes occurring between 8 and 16 hr (Figure 1C). Relative to COX-2, the accumulation of mPGES-1 protein was modest; at its peak, immunoreactivity increased an average of 2-fold over basal levels of expression. A more intense response for COX-2 induction might be anticipated, at least in part, from the phenomenon of suicide inactivation. During catalysis, reaction intermediates cause COX enzymatic activity to fall to zero within minutes (Smith et al. 2000Go). Such inactivation is not known to occur during mPGES-1–mediated isomerization of PGG2/H2 to PGE2. Based on these data, we conclude that cytokine stimulation results in a time-dependent accumulation of both COX-2 and mPGES-1 in WISH cells.

Inhibition of mPGES-1 Attenuates Cytokine-induced PGE2 Production
To investigate the contribution of COX-2 and mPGES-1 to cytokine-induced PGE2 production, cells were stimulated with IL-1ß (10 ng/ml) in the presence or absence of indomethacin (a nonselective COX inhibitor; Cayman Chemical), NS-398 (a COX-2 selective inhibitor; Cayman Chemical), or MK-886 (an inhibitor of mPGES-1 that also inhibits other membrane associated proteins involved in eicosanoid and glutathione metabolism proteins such as leukotriene C4 synthase [LTC4S] and 5-lipoxygenase activating protein [FLAP]; BIOMOL Research Laboratories, Plymouth Meeting, PA) (Mancini et al. 2001Go). Challenge with IL-1ß caused a 7-fold increase in PGE2 production relative to control cells (Figure 2, p<0.0001). Consistent with our previous work (Perkins and Kniss 1997aGo), synthesis of PGE2 was completely attenuated when stimulated with IL-1ß in the presence of 1 µM of indomethacin or 1 µM of NS-398 (both p<0.0001). MK-886 attenuated IL-1ß–induced PGE2 synthesis in a dose-dependent manner. At doses that inhibit mPGES-1 activity in vitro by ~70% (6.25 µM) or ~85% (12.5 µM) (Mancini et al. 2001Go), MK-886 reduced PGE2 production by 40% (p<0.01) and 49% (p<0.001), respectively. At higher doses (25–50 µM), which completely inhibit mPGES-1 in vitro (Mancini et al. 2001Go; Kamei et al. 2003Go), IL-1ß-induced PGE2 production was reduced to that observed in control cells (both p<0.0001). Treatment with vehicle alone (ethanol) did not affect IL-1ß–induced PGE2 production (not shown). No morphological evidence of cellular toxicity was evident at the doses of MK-886 tested. These results suggest that, despite the constitutive presence of cPGES and mPGES-2 in these cells (Ackerman and Kniss, unpublished data), cytokine-induced PGE2 production proceeds predominantly, although not exclusively, through a COX-2/mPGES-1 pathway.



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Figure 2

Cytokine-induced prostaglandin E2 (PGE2) release occurs through a dominant cyclooxygenase-2/microsomal PGE synthase-1 metabolic pathway. PGE2 production was measured by EIA in WISH cells treated for 6 hr in the absence or presence of IL-1ß with or without indomethacin (Indo), NS-398, or MK-886 (mean ± SEM for quadruplicate determinations, which were repeated twice). Statistical comparisons were performed with one-way ANOVA followed by the Tukey-Kramer post hoc test. *p<0.0001 versus control group; **p<0.0001 versus IL-1ß treatment alone; {dagger}p<0.001 versus IL-1ß treatment alone; #p<0.01 versus IL-1ß treatment alone.

 
Induced COX-2 Is Not Tightly Coordinated with mPGES-1 Expression among Individual Cells
Immunocytochemical analysis of whole-mount specimens revealed that the intercellular expression of mPGES-1 and COX-2 in response to IL-1ß was heterogenous (Figure 3A). In merged color images, cells in which COX-2 immunoreactivity predominated appeared green (arrows in Figure 3A), whereas those showing a preponderance of mPGES-1 appeared red (arrowheads in Figure 3A). To better describe this observation, fluorescence intensity analysis was performed in a representative population of cells; 628 cells were measured from 10 randomly selected fields at 40x magnification. When plotted on a histogram, the fluorescence intensity associated with mPGES-1 followed a symmetrical pattern of distribution (Figure 3B). The mean of the mPGES-1 histogram was used to divide the cells into two groups: those showing low mPGES-1 expression (51%) and those in which mPGES-1 was high (49%). Fluorescence intensity associated with COX-2 was skewed with a tail extending to the right (Figure 3C), indicating that individual cells showing very high COX-2 fluorescence intensity were among a minority of those sampled. Based on the anticipated 10-fold increase in immunoreactive COX-2 expression after IL-1ß treatment (from analysis of immunoblotting results, relative to control cells), 35% of the cells were classified as showing high COX-2 expression.



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Figure 3

Heterogeneous intercellular expression of cyclooxygenase-2 (COX-2) and microsomal prostaglandin E synthase-1 (mPGES-1) after cytokine challenge. WISH cells treated with IL-1ß for 6 hr were immunolabeled by simultaneous addition of antibodies directed against COX-2 (green) and mPGES-1 (red). (A) A representative photomicrograph showing apparent heterogeneity in COX-2 and mPGES-1 expression (bars = 20 µm). Arrows indicate cells showing predominant COX-2 immunoreactivity, whereas arrowheads denote cells in which mPGES-1 staining is superior. This image is representative of the results of at least four separate experiments. (B–D) Maximum fluorescence intensity measurements for structures labeled with anti–COX-2 and anti–mPGES-1 in each cell were assessed. Histograms of fluorescence intensity associated with mPGES-1 (B) and COX-2 (C) in this population of cells are shown. (D) Scattergram showing the relationship between COX-2 and mPGES-1 fluorescence intensity for each cell. The horizontal line indicates the demarcation between low and high COX-2–associated fluorescence intensity, whereas the vertical line denotes the demarcation between low and high mPGES-1–associated fluorescence intensity.

 
The overall intercellular expression of induced COX-2 and mPGES-1 showed a very modest but statistically significant positive correlation (Spearman r = 0.347; 95% confidence interval: 0.274–0.416; p<0.0001). Linear regression analysis was performed; however, the quality of fit to a linear model of correlation was poor (r2 = 0.12). Despite the statistical correlation, it was evident in a large proportion of cells that mPGES-1 and COX-2 were not coordinately induced. Based on the selected cutoff values between low and high fluorescence intensity (shown in Figure 3D as vertical and horizontal lines for mPGES-1 and COX-2, respectively), only 23% of the cells showed high expression of mPGES-1 and COX-2 concomitantly. Of the remainder, 39% of the cells showed low fluorescence intensity for both enzymes, 26% of the cells showed a pattern of high mPGES-1/low COX-2 expression, and 12% of the cells showed a pattern of low mPGES-1/high COX-2 expression.

Spatial Segregation of Induced COX-2 and mPGES-1 Within Cells
Having established the intercellular expression pattern for COX-2 and mPGES-1, we next evaluated the degree of intracellular colocalization of these proteins. In whole-mount specimens permeabilized with Triton X-100 and visualized by standard epifluorescence microscopy, examination of cells showing coordinate increases in both COX-2 and mPGES-1 immunoreactivity revealed apparent intracellular colocalization, most prominent in the perinuclear area (arrows in Figure 4C). Equivalent results were obtained when viewed by confocal microscopy (Figure 4D). Given that Triton X-100 might cause artifactual disruption of intracellular architecture, particularly when examining membrane-associated antigens (Goldenthal et al. 1985Go), we sought to confirm our initial observations in the presence of a second permeabilization agent (saponin). However, in the presence of saponin, the degree of apparent intracellular colocalization of COX-2 and mPGES-1 was less pronounced compared with specimens permeabilized with Triton X-100 when assessed either by standard epifluorescence (Figure 4G) or confocal microscopy (Figure 4H). Notably, in the presence of saponin, the pattern of mPGES-1–like immunoreactivity was altered. Specifically, perinuclear mPGES-1 immunostaining was reduced compared with that seen after Triton X-100 treatment, whereas punctate cytoplasmic staining was increased (Figure 4F).



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Figure 4

Intracellular localization of cytokine-induced cyclooxygenase-2 (COX-2) with microsomal prostaglandin E synthase-1 (mPGES-1). WISH cells were stimulated with IL-1ß 6 hr before fixation. Whole-mount specimens were fixed and permeabilized either with 0.2% Triton X-100 (A–D) or 0.05% saponin (E–H) before addition of anti–COX-2 (A,E) and anti–mPGES-1 (B,F) antibodies. (I–L) Intracellular localization of COX-2 (I) and mPGES-1 (J) in semi-thin cryosections. Photomicrographs were taken using the oil immersion objective of a standard epifluorescence microscope except (D) and (H), which were imaged by confocal microscopy (bars = 20 µm). Image overlay (C,D,G,H,K,L) demonstrates the degree of apparent colocalization in these images. Arrows in (C) and (D) denote a high degree of perinuclear COX-2/mPGES-1 colocalization, which was prominent in the whole-mount specimens permeabilized with Triton X-100. (L) Detail of WISH cell demonstrating mutually exclusive staining for COX-2 (green) and mPGES-1 (red) in the perinuclear area (bar = 5 µm). (M–Q) Controls for antibody specificity as visualized by standard epifluorescence microscopy (bars = 20 µm) in which staining for COX-2 (green) or mPGES-1 (red) is superimposed with that of DAPI (blue). (M,N) Typical staining patterns for COX-2 antibody alone (M) or when preabsorbed with excess recombinant protein (N). (O,P) Typical staining patterns for mPGES-1 antibody alone (O) or when preabsorbed with excess blocking peptide (P). (Q) Control in which primary antibodies were omitted.

 
Given these discrepancies, we next assessed the topological arrangement of mPGES-1 and COX-2 in semithin (~400 nm) cryosections. In comparison to the whole-mount preparations, this methodology is advantageous in that it obviates the need for detergent permeabilization, because antigens are readily available for immunodetection upon sectioning. Furthermore, because of the limited z-axis resolution of conventional mounting and microscopy, artifacts often occur because of out-of-focus fluorochromes; thus, the degree of colocalization observed tends toward overestimation, particularly in areas where staining is intense (Robinson et al. 2001Go). By physically limiting the z-axis of the specimens, the amount of "glare" from out-of-focus chromophores is reduced and two-dimensional resolution is increased (Robinson et al. 2001Go). Using this latter method, we again found mPGES-1 and COX-2 to be spatially segregated in a vast majority of cells (Figure 4K). Although the reticular pattern of immunostaining for both proteins was consistent with localization to intracellular membranes, many areas showed mutually exclusive staining for either antigen. This is illustrated in Figure 4L, in which perinuclear mPGES-1 staining was observed in areas in which COX-2 immunoreactivity was absent.

Segregation of mPGES-1 and COX-2 within Intracellular Membrane Lipid Microdomains
According to the contemporary view of membrane organization, it is widely recognized that certain proteins may reside in, or be excluded from, specialized membrane microdomains (such as lipid rafts) within the context of the cellular environment (Munro 2003Go; Cohen et al. 2004Go). In this model, cholesterol- or sphingolipid-enriched membrane microdomains may develop tightly packed, liquid-ordered phases that are distinct from the fluid-phase properties of the membrane in general (Munro 2003Go; Zurzolo et al. 2003Go). Experiments in our laboratory have indicated that WISH cells contain lipid microdomain-enriched caveolae-like structures, and express the lipid raft-associated caveolar protein, Cav-1 (Ackerman, Robinson, and Kniss, unpublished observations). Furthermore, recent studies have indicated that a proportion of catalytically active COX-2 may be localized to caveolae-like structures, and may interact with Cav-1 (Liou et al. 2000Go, 2001Go). Given the mutual exclusivity of mPGES-1 and COX-2 immunostaining within intracellular membranes, we hypothesized that segregation of COX-2, but not mPGES-1, to lipid microdomains might provide an explanation for these observations.

The relative enrichment in tightly packed cholesterol and sphingolipids imparts a resistance of lipid microdomains to extraction in cold Triton X-100 (Munro 2003Go; Cohen et al. 2004Go). As a consequence of their low buoyant densities in sucrose gradients, lipid microdomain-enriched DRM fractions can be separated biochemically from other cellular components. To assess localization of COX-2 and mPGES-1 to such microdomains, DRMs from IL-1ß–treated WISH cells were prepared as described previously (Liou et al. 2000Go, 2001Go). As in previous studies, immunoreactive Cav-1 was associated with detergent-insoluble fractions (Figure 5A, fractions 2–6). The majority (56%) of immunoreactive Cav-1 was located within fractions 2 and 3 (containing ~5% sucrose), whereas 43% was located within fractions 4–6 (containing ~30% sucrose) (Figures 5A and 5B). Unexpectedly, 96% of immunoreactive COX-2 was located within the detergent-soluble fractions 7–9 (containing ~40% sucrose) (Figures 5A and 5B). Equivalent results were obtained when COX-2 was induced using phorbol 12-myristate 13-acetate (data not shown), indicating that this observation was not restricted to the mode of induction.



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Figure 5

Segregation of mPGES-1 and COX-2 within lipid microdomains. (A) Partitioning of cyclooxygenase-2, microsomal prostaglandin E synthase-1, and caveolin-1 within DRM fractions prepared from WISH cells treated with IL-1ß for 6 hr. Fractions 1–6 represent detergent-resistant membranes, which are enriched in liquid-ordered domains, including caveolae. Fractions 7–9 contain detergent-soluble membrane and cytosolic fractions. These images are representative of the results of four independent experiments. (B) Graphical representation of immunoreactive intensity for data in (A), expressed as a percentage of total immunoreactivity for each protein as assessed by densitometry.

 
Intriguingly, we also found that 68% of immunoreactive mPGES-1 was located in DRMs, with 11% in fractions 1–3 and 57% in fractions 4–6 (Figures 5A and 5B). Only a minority (32%) of immunoreactive mPGES-1 was located within the detergent-soluble fractions (7–9 in Figures 5A and 5B). In the absence of cytokine stimulation, the partitioning of mPGES-1 between detergent-soluble and DRM fractions was equivalent to that seen after IL-1ß treatment (not shown). This suggests that cytokine stimulation alone is unlikely to be the basis for the presence of mPGES-1 in DRMs. Collectively, these data suggest that the majority of mPGES-1 may reside in lipid microdomains that exclude COX-2.


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 Literature Cited
 
It has been established that cytokine-mediated PGE2 production requires de novo COX-2 synthesis in human WISH epithelial cells (Perkins and Kniss 1997aGo). Our current data suggest that, as in other models, high-amplitude PGE2 output depends on cooperation between COX-2 and mPGES-1 in carrying out the sequential reactions of PGG2/H2 generation and isomerization to PGE2 (Murakami et al. 2000Go; Thoren et al. 2003Go; Trebino et al. 2003Go). As evidence of this, we found COX-2 and mPGES-1 to be coinduced after challenge with the cytokine IL-1ß. Additionally, we found that treatment with the membrane-associated proteins involved in eicosanoid and glutathione metabolism inhibitor MK-886 abrogated cytokine-elicited PGE2 release in a dose-dependent manner. Because MK-886 inhibits mPGES-1 (but not mPGES-2 or cPGES) activity in vitro, this further suggests that cytokine-elicited PGE2 production proceeds preferentially through mPGES-1. This is consistent with the results of recent gene targeting studies, through which a critical role for mPGES-1 in COX-2–mediated PGE2 production has been established (Uematsu et al. 2002Go; Boulet et al. 2004Go). In light of recent findings that the maximum reaction velocity, turnover rate, and catalytic efficiency of purified mPGES-1 are orders of magnitude higher than those of other terminal PG synthases (including cPGES and mPGES-2), it is quite likely that mPGES-1 is the dominant catalyst for delayed PGE2 isomerization in human cells (Thoren et al. 2003Go). We are extending our results using RNA interference technology to examine the absolute requirement of mPGES-1 for PGE2 biosynthesis in WISH cells.

A fascinating finding in our current work was the unexpected observation of cell-to-cell variability in induced expression of mPGES-1 and COX-2. Despite a modest positive correlation, coordinate increases in immunoreactive mPGES-1 and COX-2 occurred in only 23% of cells (Figures 3A and 3D). Such an observation cannot be appreciated using methods in which cell lysates are pooled (such as with Northern or Western blotting). Although common signaling cascades and transcription factors (such as nuclear factor-kappa B) contribute to upregulation of both enzymes (Catley et al. 2003Go), our data suggest that the overall regulatory control for each of these genes is distinct, which is consistent with a number of recent reports (reviewed in Murakami and Kudo 2004Go). Our data additionally suggest that individual cells may contribute unequally to overall cytokine-induced PGE2 release, such that the most substantial PG production may occur in a limited number of cells. That this may be relevant in vivo is supported by our recent report of heterogeneous expression of COX-2 protein in human amnion epithelial cells from placental membranes in the setting of term human labor (Dunn-Albanese et al. 2004Go).

The role of intracellular compartmentalization between COX isoforms and terminal PGE isomerases is not well-delineated, but has profound functional consequences. It has been suggested that the enzymes involved in the liberation and conversion of arachidonic acid to PGE2 (including COX-1, COX-2, phospholipase A2 isoforms, and certain terminal PG isomerases) might colocalize to the same subcellular organelles, which might facilitate coordination of their metabolic activities (Naraba et al. 1998Go; Murakami and Kudo 2004Go). Our data indicate that, in WISH epithelial cells, COX-2 and mPGES-1 reside primarily within intracellular membranes, most probably within the endoplasmic reticulum (ER) and along the contiguous outer nuclear envelope. This has been corroborated by the finding of immunoreactive mPGES-1 and COX-2 in microsomal, but not cytosolic, fractions prepared from these cells (data not shown), which is consistent with previous reports (Jakobsson et al. 1999Go; Murakami et al. 2000Go). However, our data also indicate that discrete lipid microdomains may physically separate the two enzymes. In whole-cell immunocytochemical preparations permeabilized with Triton X-100, we found cytokine-induced COX-2 and mPGES-1 to be particularly concentrated (and apparently colocalized) within the perinuclear area (Figures 4C and 4D). Using saponin as alternate permeabilization agent, however, we noted that the degree of overlap between immunofluorescently stained COX-2 and mPGES-1 was significantly reduced (Figures 4G and 4H). Similar reagent-specific inconsistencies in antigen distribution have been noted previously (Goldenthal et al. 1985Go), suggesting that standard permeabilization methods may be inadequate to characterize accurately the intracellular arrangement of some membrane-associated proteins. Using a higher resolution methodology employing semithin physical sections (which circumvents the need for postfixation detergent treatment), we discovered that COX-2/mPGES-1 were largely noncolocalized, even in the perinuclear area (Figures 4K and 4L). This indicates that the extent of overlap observed by conventional analysis might be overestimated, suggesting that a majority of COX-2 and mPGES-1 are not in immediate spatial proximity.

In further support of a topological separation between COX-2 and mPGES-1, we found that the majority (68%) of immunoreactive mPGES-1 partitioned with Cav-1–enriched DRMs in which COX-2 was excluded (Figures 5A and 5B). Consequently, most intracellular COX-2 and mPGES-1 appear to be biochemically segregated, even when residing in the same subcellular compartment. The finding of COX-2 (together with other ER-associated membrane proteins, not shown) in soluble fractions constitutes an important negative control in these experiments, because it excludes incomplete solubilization of bulk membranes as a trivial explanation for the finding of mPGES-1 in DRMs (Schuck et al. 2003Go). These results are in contrast to those described by Liou et al. (2001)Go, who found that in human foreskin fibroblasts, a significant proportion of induced COX-2 was present in DRM fractions concomitantly with Cav-1. This suggests that COX-2/Cav-1 interactions may be cell type–specific.

Given that mPGES-1 partitioned into DRM fractions, it is tempting to speculate that this enzyme might reside in lipid rafts within the cell. According to the lipid raft model, cholesterol- and sphingolipid-enriched membrane microdomains may develop tightly packed, liquid-ordered phases that are distinct from the fluid-phase properties of the membrane in general (Munro 2003Go; Zurzolo et al. 2003Go). Transmembrane proteins can reside in, or be excluded from, such lipid rafts depending on their physical characteristics (Munro 2003Go). A widely used method to delineate protein content of lipid rafts is to extract membranes in cold Triton X-100, in which proteins associated with DRM fractions are not solubilized (Munro 2003Go; Schuck et al. 2003Go). As a consequence of their low buoyant densities, DRM fractions can be physically separated detergent soluble fractions by isopycnic centrifugation and analyzed for protein content by immunoblotting.

It is difficult to reconcile that mPGES-1 appears to localize to internal membranes while residing in largely in lipid microdomains. As traditionally conceived, lipid rafts are commonly present in the plasma membrane, but occur at low levels within most internal membranes (Munro 2003Go). However, given that the ER is the site of sterol production, and that high levels of sterols favor the development of liquid-ordered domains, it is conceivable that liquid-ordered domains may exist within ER membranes. Indeed, detergent-resistant ER lipid microdomains have been described, potentially associated with lipid body biogenesis (Hayashi and Su 2003Go; Tauchi-Sato et al. 2002Go). Hayashi and Su (2003)Go noted that these ER-associated DRM microdomains exhibited lower buoyancy in Triton X-100 sucrose flotation compared with plasma membrane DRM fractions. Similar to their results, we observed that peak Cav-1 immunoreactivity (as a marker for plasma membrane rafts) occurred in a more buoyant fraction (fraction 3, Figures 5A and 5B) than that of mPGES-1 (fractions 4–6, Figures 5A and 5B), suggesting that mPGES-1 may partition into a similar ER-associated microdomains. We are attempting to define further these putative ER-associated lipid microdomains in terms of lipid composition, protein content, and sensitivity to cholesterol depletion, as well as through the use of more refined extraction methodologies. Although we are sensitive to the realization that there is controversy regarding extrapolating data derived from DRMs to preexisting structures within living cells (Munro 2003Go; Zurzolo et al. 2003Go), we believe our results are sufficiently compelling to warrant further investigation.

In light of our present findings, the mechanism for preferential coupling between COX-2 and mPGES-1 may be more dynamic and complex than would be anticipated based on a simple compartmentalization model. Our current fluorescence imaging and biochemical data suggest either that immediate proximity may not be required for COX-2/mPGES-1 functional coupling or that, if required, it may occur only transiently. The finding of coincident partitioning of a minority of mPGES-1 and essentially all of COX-2 into detergent-soluble membrane fractions suggests that transient mPGES-1/COX-2 interactions may occur within liquid-disordered internal membranes of a given cell. To date, however, no direct interactions between COX-2 and mPGES-1 have been documented. Additionally, we speculate that intracellular COX-2/mPGES-1 coupling may be but one of the mechanisms through which PGE2 biosynthesis occurs. Our current immunofluorescence data suggest that transcellular arachidonic acid metabolism might also occur in WISH, such that cells expressing high levels of COX-2 may donate metabolic intermediates (PGG2/H2) to neighboring cells enriched in mPGES-1. A precedent for such a scenario stems from observations that endothelial cell-derived PGH2 can be used as a substrate for thromboxane production by platelets (Karim et al. 1996Go; Camacho and Vila 2000Go) or for PGI2 production by lymphocytes (Merhi-Soussi et al. 2000Go).

In summary, our results suggest that synergistic activity between COX-2/mPGES-1 is the principal means through which cytokine-elicited PGE2 production proceeds within WISH epithelial cells. The mechanisms through which mPGES-1 and COX-2 may functionally couple remain incompletely defined. Our data suggest that the role of intracellular compartmentalization in COX-2/mPGES-1 interactions may be more complex than current models indicate. Future efforts will be directed at exploring these and other potential mechanisms for mPGES-1/COX-2 coupling.


    Acknowledgments
 
This work was supported by National Institutes of Health Grant RO1 HD-35881 (to DAK), National Institutes of Health postdoctoral training Grant F32 HD-42910 (to WEA), and The Ohio State University Perinatal Research and Development Fund.

We gratefully acknowledge Dr. Toshihiro Takizawa, whose expert guidance was indispensable for the preparation of the semi-thin physical sections. We are indebted to Kathleen Wolken of the Campus Microscopy and Imaging Facility for her excellent technical assistance. We thank Drs. Eric J. Smart and William V. Everson of the University of Kentucky for critical review of this manuscript. Portions of this work were presented at the 50th Annual Meeting of the Society for Gynecologic Investigation, March 26–30, 2003, Washington, DC, and the 10th Annual Meeting of the International Federation of Placenta Associations, September 25–29, 2004, Asilomar, CA.


    Footnotes
 
Received for publication April 6, 2005; accepted May 18, 2005


    Literature Cited
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