Departments of 1 Cellular and Molecular Medicine and Kidney Research Centre and of 2 Biochemistry, Microbiology and Immunology, University of Ottawa, Ottawa, Ontario, Canada K1H 8M5
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ABSTRACT |
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Our present study has investigated the effect of cyclooxygenase-2 (COX-2) inhibition on prostaglandin E2 (PGE2) receptor expression in M-1 cortical collecting duct cells and measured their response to PGE2. Using a semiquantitative titration analysis method, we show that following the addition of the COX-2-specific inhibitor NS-398, E-prostanoid receptor subtype (EP3 and EP4) mRNA expression was found to increase threefold each vs. the vehicle-treated control. We also observed that EP1 but not EP2 is expressed in M-1 cells and EP2 levels are not induced by NS-398. To determine the status of the PGE2 response on exposure to NS-398, we measured cAMP levels in cells after stimulation with varying concentrations of PGE2, then pretreated the cells with 10 µM NS-398 before PGE2 exposure and found a significant rise in the stimulatory effect of PGE2 on cAMP production. Finally, Western blot analysis of the levels of the EP4 receptor protein in control vs. NS-398-treated cells revealed an induction in protein levels in these cells, correlating with the induction in EP4 mRNA. We conclude that NS-398 upregulates the expression of EP3 and EP4 mRNA in M-1 cells. Also, EP4 protein levels are increased, resulting in an increased stimulation of cAMP production by PGE2.
adenosine 3',5'-cyclic monophosphate measurements; cyclooxygenase-2; E-prostanoid receptor subtypes; semiquantitative reverse transcriptase-polymerase chain reaction; Western blotting; cortical collecting duct; prostaglandin E2
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INTRODUCTION |
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PROSTAGLANDIN E2 (pge2) is by far the most prevalent product of the arachidonic acid cascade in the kidney. It is highly produced in the glomerular regions and in the distal nephron, particularly in the collecting duct (5). PGE2 is clearly an important regulator of salt and water transport along the nephron (6). Recent advances in the study of prostaglandin physiology have identified at least four subtypes of G-protein-coupled cell surface receptors that bind PGE2 (8, 23), the E-prostanoid receptor subtypes (EP1, 2, 3, 4). For example, the EP1 receptor couples to Gq and activates protein kinase C. It is highly expressed in the medullary region of the kidney and is thought to be involved in the inhibition of sodium reabsorption by PGE2 in the collecting duct (11, 13). The EP2 receptor couples to Gs and activates adenylate cyclase; to date it remains unclear whether this receptor is expressed in renal tubules and what role it plays in renal physiology (6). The EP3 receptor couples to Gi and inhibits adenylate cyclase. By this mechanism, PGE2 inhibits arginine vasopressin-dependent water transport in the collecting duct (14). And, finally, the EP4 receptor couples to Gs and stimulates adenylate cyclase, resulting in increased water reabsorption in the collecting duct due to elevations in cellular cAMP levels (6). However, the signaling linked to PGE2 is not limited to these four subtypes of EP receptors. Alternatively spliced forms of the EP3 receptor exist (15) and, to various extents PGE2, are capable of activating other prostanoid receptors as well, i.e., the TP, IP, DP, and FP receptors (8, 23, 24), maybe even eliciting its effects through other indirect means within cells, adding to the complexity of its actions. In addition, nuclear signaling linked to PGE2 pathways is presently being explored, because nuclear EP1,3,4 receptors have been identified (3, 4).
PGE2 is produced via the cyclooxygenase (COX) pathway, catalyzed by the enzyme prostaglandin endoperoxide H synthase (also known as cyclooxygenase). There are two known isoforms for this enzyme, COX-1 and -2. Whereas the first is constitutively expressed in most regions of the kidney, the latter is an inducible form, upregulated for instance during an inflammatory response (17). However, studies in the M-1 cell line have shown that the intercalated cells of the collecting duct constitutively express both COX isoforms and that COX-2 contributes to the majority of the PGE2 produced in these cells (10). A widespread use of COX inhibitors, commercially known as nonsteroidal anti-inflammatory drugs (NSAIDs), has lead to extensive research in this field. In addition to their beneficial effects in many conditions, this inhibition of COX enzyme activity through NSAID abuse is associated with various adverse effects, including not only renal toxicity linked to a decrease in prostaglandin production but also abnormalities in water and sodium metabolism, resulting in edema and hypertension (7). Moreover, studies in COX-2 null mice revealed a major deficiency in postnatal renal development (20). This clearly demonstrates the significance of PGE2 in many biological processes. In addition, the sale of aspirin in itself is a billion-dollar industry worldwide (9). This recognition has led to the development of isoform-specific inhibitors, which now facilitate the study of the cyclooxygenase pathway and its usefulness within the cell, illustrated by alterations in PGE2 actions.
Although the main mechanism of action of NSAIDs is to inhibit COX
activity, which in turn leads to decreases in PGE2
production, there may also be another regulatory level in terms of the
cellular responsiveness to PGE2. This is conceivable if,
for instance, there is a resultant alteration in the expression of
PGE2 binding sites within the cells on exposure to NSAIDs.
This would imply, for example, that there is a compensatory
upregulation of EP receptors to counteract the downregulation in
prostaglandin production. Therefore, the main hypothesis of this work
is to determine whether the expression of EP receptor subtypes is
altered by NSAID exposure. The purpose of this study was to investigate
the effect of a COX-2-specific inhibitor, NS-398 (9), on
the expression of EP receptor subtypes in the M-1 cell line. The M-1
mouse cortical collecting duct (CCD) cell line is a heterogeneous
population of principal, -, and
-intercalated cells derived from
an SV-40 transgenic mouse. These cells have been shown to possess the
characteristics of the in vivo CCD, both in terms of morphology and
physiological properties (28). Expression of mRNA for the
specific EP receptors was examined in these cells by RT-PCR, and the
relative abundance in NS-398-treated cells was compared with control by
semiquantitative RT-PCR. Next, to examine the changes at the protein
level, Western blotting was performed. And finally, to assess the
change in cellular responsiveness to PGE2, cAMP levels
within the cells were measured.
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MATERIALS AND METHODS |
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M-1 cell culture. M-1 mouse CCD cells (CRL-2038, ATCC, Rockville, MD) were grown to confluence in 100-mm petri dishes at 37°C and 5% CO2. The culture media consisted of DMEM/F-12 media, pH 7.4, containing 5% fetal bovine serum and 1% penicillin-streptomycin-fungizone (GIBCO-BRL). The cells were grown at passages 19-41 for 2-3 days, and before the experiments they were serum deprived for 18 h. To assess the effect of the COX-2-specific inhibitor NS-398 (Cayman, Ann Arbor, MI) on the expression of EP receptors, 10 µM NS-398 was added to the respective cells.
RNA isolation and RT-PCR. M-1 cells were grown to confluence in 100-mm petri dishes and were scraped off and centrifuged at 1,100 rpm. The collected pellet was resuspended in 1 ml TRIzol reagent (GIBCO), and total RNA was isolated using the TRIzol method, as described by the manufacturer (GIBCO), and was DNase treated (Boehringer Mannheim) to eliminate genomic DNA. The RNA was reverse transcribed into cDNA using MuLV RT and random hexamers, provided in the Gene-AMP RNA PCR core kit (PerkinElmer). Samples were prepared in duplicate for each reaction, the duplicates serving as controls for the reverse transcription because the RT was omitted in these tubes. The upstream and downstream primers used for PCR amplification of each cDNA are EP1 receptor (336 bp; nucleotides 865-1201) EP1.1 5'-CGCAGGGTTCACGCACACGA-3' and EP1.2 5'-CACTGTG CCGGGAACTACGC-3'; EP2 receptor (401 bp; nucleotides 757-1158) EP2.1 5'-AGGACTTCGATGGCAGAGGAGAC-3' and EP2.2 5'-CAGCCCCTTACACTTCTCC AATG-3'; EP3 receptor (206 bp; nucleotides 86-291) EP3.1 5'-TGGCCTTTCCCAT CACCATGATGGTCA CTG-3' and EP3.2 5'-CCAGCGTCGCTGTGACAGGTACAC GAGG-3'; and EP4 receptor (423 bp; nucleotides 941-1364) EP4.1 5'-TTCCGCTCG TGGTGCGAGTTTC-3' and EP4.2 5'-GAG GTGGTGTCTGCT TGGGTCAG-3' (1). The DNA amplification was performed using a thermal cycler, the PerkinElmer Gene-AMP PCR System 2400. Depending on the primers used, the PCR reaction consisted of the following parameters: denaturing at 94°C for 240 s, followed by 35 cycles of denaturing at 94°C for 15 s; annealing at 63°C for 30 s; extension at 72°C for 30 s, then 72°C for 10 min; and, finally, cooldown to 4°C. The given amplification products were then separated by gel electrophoresis, on a 1.5-2% agarose gel, for size determination with standards, and visualized under ultraviolet light using ethidium bromide. Spleen total RNA served as positive control for the amplification of EP2. To verify the identity of EP1, a PstI (GIBCO) restriction digest was performed generating two products of 207 and 129 bp from the initial 336-bp EP1 product.
Semiquantitative PCR analysis.
To measure the relative expression of EP3 and
EP4 mRNA in NS-398 vs. vehicle-treated M-1 cells, a
semiquantitative RT-PCR analysis was performed. For each experiment, a
standard curve was constructed, whereby 100, 200, 400, and 600 ng of
control total RNA were amplified by PCR. The signal density was then
measured with the Storm PhosphorImager system and was plotted as log
(signal density) against log [starting total RNA (ng)]. The
measurements with the PhosphorImager were performed by mixing 15 µl
of the amplified PCR products with Vistra green (Amersham) at a final dilution of 1:5,000, followed by a 15-min incubation at room
temperature. The samples were then electrophoresed on a 2% agarose
gel. Using linear regression, a straight line was obtained and the
slope and y-intercept were used to interpolate the relative
strength of the signals for NS-398-treated samples (starting RNA was
200 ng) compared with control. For amplification of the EP receptor fragment by PCR, the following parameters were employed: 94°C for
240 s, followed by 26 cycles of 95°C for 30 s and 72°C
for 30 s, and then 240 s at 72°C. To determine the adequate
number of cycles, log (signal density) was plotted against the cycle number, and the number was chosen within the linear range. The signal
was then normalized using the respective -actin signal, which was
amplified by 94°C for 240 s, followed by 27 cycles of 95°C for
15 s, 63°C for 30 s, and 72°C for 30 s, and then
240 s at 72°C. To minimize the variability between samples and
adequately normalize the quantity of starting RNA with
-actin, the
same RT sample was divided equally between EP and
-actin for
subsequent PCR amplification. For the EP2 receptor
transcript, the cultured M-1 cells were treated with 10 µM NS-398 for
3, 6, 12, and 24 h. Then PCR amplification was performed to
determine whether NS-398 could induce the expression of EP2
receptor mRNA in these cells, which clearly do not constitutively
express it.
cAMP radioassays. M-1 cells were grown to confluence in 24-well plates for 3 days and serum starved 24 h. The cells were then pretreated for 15 min in DMEM/F-12 containing 0.5 mM 3-isobutyl-1-methylxanthine and 10 µM indomethacin (Sigma). At timed intervals, the cells were then stimulated with 1 nM to 1 µM of arginine vasopressin (AVP) and PGE2. The samples were all prepared in duplicate. To stop the reaction, 300 µl of 10% trichloroacetic acid (TCA) were added to each well. After a 30-min incubation on ice, the samples in TCA were transferred to Eppendorf tubes and centrifuged for 10 min at 4,000 g. Next, 250 µl of each sample were transferred to glass test tubes, and four ether extractions of TCA were performed using 4 times the volume of H2O-saturated diethyl-ether per extraction. One molar Tris · HCl was used to bring the pH of the samples to 7-8. By using the cAMP radioassay kit (Intermedico), cAMP levels in each sample were then measured in 100 µl of sample according to the manufacturer's instructions. To assess whether treatment of the cells with NS-398 alters the cell response to PGE2, the cells were incubated for 12 h with 10 µM NS-398 and then assayed for cAMP levels on stimulation with PGE2, as described above.
In the second set of experiments, to verify the ability of PGE2 to inhibit the AVP-stimulated cAMP, the cells were pretreated for 15 min with the above-mentioned concentrations of agonist and then treated with 0.1 µM AVP in the presence of PGE2 for 7 min. Similarly, to verify the effect of treatment of M-1 cells with NS-398 on this inhibitory response, the cells were incubated for 12 h with 10 µM of NS-398.Western blotting.
M-1 cells were grown to confluence in 100-mm petri dishes, after which
10 µM NS-398 was added for either 1 or 12 h, except for
controls, to which serum-free DMEM/F-12 was added. The M-1 protein
samples were prepared by lysing cells in 100 mM Tris · HCl (pH
7.4), containing 1 mM EDTA and 1 mM EGTA, followed by sonication for
5 s using an Ultrasonics cell disrupter. The cell lysates were
then centrifuged at 10,000 g for 10 min, and the supernatant
was removed. The Bio-Rad reagent was added to analyze the
protein content using the Bradford method (Bio-Rad, Mississauga, ON). After the protein concentrations were determined, 25 or
50 µg of protein were combined with the Laemmli buffer and denatured by boiling for 3 min followed by a 2-min incubation on ice. The samples
were then loaded onto a polyacrylamide gel (4% stacking and 10%
resolving layers) and resolved by SDS-PAGE using a Mini-PROTEAN II
apparatus (Bio-Rad). The protein was then transferred to a Hybond
enhanced chemiluminescence (ECL) nitrocellulose membrane (Amersham)
using the Mini-Trans Blot system (Bio-Rad). To minimize background
signals before incubation with the primary antibody, the membranes were
incubated overnight at 4°C in a solution of 10% fat-free dried milk
(Carnation) in Tris-buffered saline containing 0.1% Tween 20 (TBS-T).
The membranes were then incubated for 90 min at room temperature with
the primary antibody, a human polyclonal -EP4 IgG
(Cayman), and diluted 1:5,000 in 10% milk in TBS-T. To test the
specificity of the EP4 antibody, it was preincubated for
24 h at 4°C with its respective immunizing peptide. Before incubation with the secondary antibody, 3- to 20-min washes in TBS-T
were performed. Next, the membranes were incubated with horseradish
peroxidase (HRP)-conjugated goat anti-rabbit IgG (Promega, Madison, WI)
and diluted at 1:2,000 for 60 min at room temperature. After 60 min the
membranes were washed for (3 × 20 min), ECL was used to visualize
the results as directed by the manufacturer (Amersham). To normalize
for protein loading, the membranes were stripped and incubated with the
-mouse
-actin monoclonal antibody (Sigma) at a dilution of
1:5,000 for 30 min at room temperature. After being washed as above,
the HRP-conjugated
-mouse IgG secondary antibody was incubated with
the membranes for 30 min at room temperature and diluted at 1:2,000.
Results were visualized as described above. To assess protein loading,
densitometry was carried out using the Kodak Digital Science ID image
analysis software, and EP signal density was compared with the
corresponding
-actin density.
Statistics.
For semiquantitative RT-PCR, data were presented as means ± SE.
To analyze the statistical significance of the data, the Prism Graphpad
2.01 software was used and consisted of repeated measures of ANOVA,
followed by the Bonferroni posttest. Statistical significance was
determined at P < 0.05 with n 3. For cAMP and Western blot analysis, experiments were performed using
duplicate samples and repeated three to six times each. The SigmaPlot
software for windows version 4.01 (1986-1997) was used to analyze
data. Results are expressed as means ± SE. A Student's
t-test and a one-way ANOVA were performed to assess the
statistical significance between data points, followed by Tukey's test
for comparison of values, P < 0.05.
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RESULTS |
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Detection of EP receptor mRNA by RT-PCR.
Because PGE2 is a major renal product of the arachidonic
acid cascade clearly involved in modulating the transport of sodium and
water in the collecting duct (5, 6), this study analyzed the expression of EP receptors in the M-1 mouse CCD cell line. As shown
in Figs. 1, A and
B, 2A, and
3A, with the exception of EP2, mRNA for all three other EP receptors,
EP1, 3, 4 are detectable in M-1
cells. The 206- and 423-bp bands, representing the EP3 and
EP4 receptors, respectively, are visible in Figs.
2A and 3A. Whereas the expression of
EP2 mRNA was clearly not detected in these cells, it is
abundantly expressed in preparations of mouse spleen total RNA,
confirming the quality of the primers used and the efficacy of the
parameters employed: cycle number, annealing temperature, and amount of
starting RNA. The 401-bp EP2 product is indeed only
detected in spleen preparations and not in the M-1 cells (Fig.
1B). This is consistent with previous findings, indicating
that it is the EP4 and not the EP2 receptor
that couples to Gs in the collecting duct to increase cAMP
(6). Furthermore, it is believed that very little, if any,
EP2 receptors are present within the kidney
(6). In accordance, previous studies in our laboratory
have also shown that the EP2 receptor is not present in the
rat kidney (25). As for the EP1 receptor mRNA,
a restriction digest using the PstI endonuclease was
performed to verify the identity of the PCR product obtained. As
expected, the digest generated two bands: 207 and 129 bp from the
initial 336-bp product (Fig. 1A), confirming the presence of
EP1 receptors in the M-1 cell line. The absence of the
129-bp band in Fig. 1A is possibly due to the fact that it
is concealed by the bromophenol blue from the loading buffer, which
runs at this size. In addition, the protein kinase N (PKN) long
transcript was consistently amplified using these EP1
primers, giving a band of 776 bp, as shown in Fig. 1A. The
identity of this 776-bp fragment as to PKN was demonstrated by
digestion with PstI, yielding three bands: 388, 202, and 186 bp.
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Effect of NS-398 on the relative expression of EP3 and EP4 in M-1 cells compared with control. A possible mechanism by which any given cell can compensate for a decreased production of prostaglandins is to increase the sensitivity of the cell to the agonist, for instance, by increasing the number of binding sites on its surface. Therefore to examine whether the COX-2-specific inhibitor NS-398 could alter the expression of the EP3 and EP4 receptor mRNA, a semiquantitative RT-PCR approach was employed, whereby a titration analysis for both EP3 and EP4 was performed. First, PCR conditions were optimized to ensure that alterations in the quantity of starting RNA would be reflected in the amount of PCR product amplified. In fact, there is a linear increase in signal intensity corresponding to increases in starting RNA from 100 to 600 ng; this is demonstrated in Figs. 2A (lanes 1-4) and 3A (lanes 1-4), for EP3 and EP4, respectively. As shown in Fig. 2A (lanes 5-7), when 200 ng of starting RNA were used, there is a gradual increase in EP3 mRNA, dependent on increasing the exposure time to NS-398 from 1.5 to 6 h, with significant differences reaching a plateau at levels threefold higher than control after 4-h exposure. The same is seen for EP4 mRNA in Fig. 3A (lanes 5-7), with significant increases also reaching a plateau at levels threefold higher than control after 4-h exposure to NS-398. Next, changes in EP2 receptor mRNA were assessed, to ensure that although the M-1 cells did not constitutively express this receptor, it is not inducible under certain conditions. Therefore, as described above, we treated the cells with the COX-2-specific inhibitor NS-398 for varying periods of time, from 6-24 h. However, as shown in Fig. 1B, lanes 4-7, treatment of the M-1 cells with NS-398 did not result in an induction of the EP2 receptor mRNA. As for the EP1 receptor subtype (Fig. 1A), due to the constant amplification of the PKN product described above, it would have been difficult to interpret the results of band densities due to the fact that there is differential amplification of the 336- and 776-bp bands; and accounting for the differences in specificity of the primers and Taq DNA polymerase for each transcript further complicates the analysis. Furthermore, a literature search indicates that the whole EP1 gene is entirely overlapped by an exon of the long transcript of PKN (2), preventing the selection of primers that would only amplify the EP1 fragment.
NS-398 increases cAMP stimulation by PGE2.
Numerous studies have demonstrated that PGE2, acting mainly
through the EP4 receptor, stimulates adenylate cyclase via
coupling to Gs and thereby increases cellular cAMP levels
within the collecting duct (6, 8, 23-24). Because in
this study we demonstrated an induction in EP4 receptor
mRNA by NS-398, we set out to decipher whether this is correlated to an
increase in cellular signaling via EP4. Therefore, these
experiments measured the cellular levels of cAMP in response to
stimulation of cultured M-1 cells with increasing concentrations of
PGE2. As shown in Fig. 4, a
stimulatory response significantly different from control was obtained
on treatment with PGE2 concentrations above 100 nM. The
basal cAMP concentrations were 11 ± 3 × 103
pmol/µg. In the presence of PGE2, cAMP levels ranged
between 17 ± 5 × 10
3 pmol/µg at 1 nM and
197 ± 30 × 10
3 pmol/µg at 10 µM, reaching
significance at 100 nM with cAMP levels at 109 ± 32 × 10
3 pmol/µg. In the presence of NS-398, basal levels
did not differ from nontreated cells, 12 ± 2 × 10
3 pmol/µg. But the stimulatory response to
PGE2 was significantly increased, ranging from 202 ± 27 × 10
3 pmol/µg at 100 nM to 375 ± 34 × 10
3 pmol/µg at 10 µM, P < 0.05.
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Altered EP4 protein levels on stimulation with NS-398.
To determine whether the induction in EP4 mRNA levels on
treatment with NS-398 is correlated to an increase in protein levels, Western blot analysis using a human -EP4 receptor
antibody was performed. As shown in Fig.
6A, EP4 protein is
detectable in cultured M-1 cells. Whereas no significant alteration in
EP4 protein was detectable after 1 h exposure of cells
to the COX-2-specific inhibitor NS-398, there was up to a sixfold
increase (Fig. 6D) in response to 12 h of NS-398
treatment. To verify the specificity of the antibody, it was
preincubated with the immunizing peptide used to generate it. This
resulted in an inhibition of binding of the antibody to the membrane;
hence no signal is visible in Fig. 6B. To perform the
densitometric analysis of the results, the
-actin levels were
detected in Fig. 6C. Overall, this compensatory increase in
EP4 receptor expression in response to inhibition of COX-2; i.e., prostaglandin synthesis is somewhat predictable. Previous work in
our laboratory demonstrated that in response to inhibition of COX
activity, there is a cellular upregulation in COX levels (10). The fact that the number of cell surface binding
sites for PGE2 is increased in the M-1 cells provides a
further level of compensation by the cell, ultimately increasing the
sensitivity to PGE2.
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DISCUSSION |
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The kidney collecting duct is a predominant site of renal
PGE2 synthesis (5). Tubular PGE2
plays an important role in regulating ion concentrations in the urine.
PGE2 has been shown to inhibit NaCl transport in the
collecting duct, favoring salt elimination in the urine together with
water (6). The significance of PGE2 to the
maintenance of salt and water homeostasis is clearly demonstrated by
the undesirable renal effects such as sodium and potassium retention
(26) associated with the use of NSAIDs, which inhibit the
production of prostaglandins. However, whether selective inhibition of
COX-2 produces less kidney toxicity than nonselective NSAIDs is poorly
documented. However, a study using the COX-2 inhibitor Celecoxib
revealed that sodium and potassium retention resulted from COX-2
selectivity and therefore does not spare the kidney (26).
To better appreciate the protective vs. toxic effects of COX inhibitors
within the renal cell, one must examine alterations at different levels
of the prostaglandin pathway, looking not only at prostaglandin
production but also cellular responsiveness to prostaglandins. The
latter was the focus of this work, investigating the effects of the
specific COX-2 inhibitor NS-398 on the levels of EP receptor subtypes
and the signaling pathways coupled to these receptors. Although work in
this area is limited, there are two distinct studies that have examined
the specific effect of each isoform on the regulation of prostaglandin
receptors. The first demonstrated the effects of different inhibitors
on receptor densities in response to NS-398, the specific COX-2
inhibitor, and to valeryl salicylate, the COX-1 inhibitor, in cerebral
vasculature of newborn pigs (19). They showed an increase
in PGE2 and PGF2 receptors only in the
presence of the COX-2 inhibitor, because they report that COX-2 is the
predominant contributor of PG production in the newborn pig brain
(19). Another study looking at the effects of COX
inhibition on the expression of PG receptors in retinal vessels of
neonatal pigs revealed a similar alteration in EP1,
EP3, and FP receptors in response to both COX-1 and
-2-selective inhibition (12).
Our group has reported that both COX-1 and COX-2 enzymes localized to the intercalated cells of the mouse CCD and not to the principal cells. Also, and perhaps more importantly, the COX-2 enzyme catalyzes the majority of PGE2 synthesis in this nephron segment (10). The actions of PGE2 are mediated by G-protein-linked receptors termed EP. Acting through at least three subtypes of the EP receptors, EP1, EP3, and EP4 (6, 30), PGE2 is involved in the fine-tuning of salt and water transport in the collecting duct. Functional studies revealing that EP receptors mediate these effects have suggested that these receptors are present on the principal cells of the collecting duct (6, 11). Taken together, these studies imply that a paracrine interaction between the intercalated and principal cells where COX-2 localized to the intercalated cells would produce PGE2, which would stimulate EP receptors on the principal cells to alter water and sodium reabsorption.
Having demonstrated that PGE2 synthesis in the collecting duct is COX-2 dependent, we initiated experiments to unravel the mechanism by which COX-2-selective inhibitors interfere with the collecting duct functions. Previously, we have shown that NS-398, a COX-2-selective inhibitor, upregulates the levels of COX-2 protein as efficiently as the nonselective NSAID indomethacin (10). Present results indicate that the levels of EP3 and EP4 receptor mRNA also increase on exposure to NS-398 and that EP4 protein is upregulated. This finding is consistent with the work of Hardy et al. (12), showing an upregulation in EP3 receptors. However, our study is the first demonstration to our knowledge of an alteration in EP4 levels. Therefore, this regulatory effect of COX inhibition does seem to be true for most if not all prostaglandin receptors (EP1,3,4 and FP) and is independent of the inhibitors used (NS-398, valeryl salicylate, indomethacin), whether they are nonselective or isoform specific (12, 19). Because PGE2 production is important for the collecting duct physiology, and the adverse renal effects associated with a lack of PGE2 production have been clearly demonstrated (7, 20), it is highly conceivable that the cells possess many compensatory mechanisms to maintain homeostatic function. One possible regulatory level is the EP receptors on the cell surface, thereby altering the cellular responsiveness to PGE2. It is noteworthy that the two EP receptors, which we found to increase on exposure to NS-398, are coupled to opposing signaling pathways in the collecting duct; EP3 inhibits cAMP formation whereas EP4 stimulates cAMP production. There is a possibility that selective EP3 spliced variants are upregulated by exposure to NS-398, and the primers used in our study to amplify the EP3 fragment are not selective. EP3 spliced variants have been shown to couple to different G proteins and as a consequence can activate different signaling pathways (23, 24). Therefore we cannot as yet exclude the possibility that the inhibitory pathway is also altered by COX-2 inhibition.
To verify whether the compensatory upregulation in EP4 receptor expression in response to COX-2 inhibition is relevant and corresponds to changes in prostaglandin physiology in the collecting duct, we studied the cAMP signaling pathway in response to PGE2 in the absence and presence of NS-398. In addition to finding that PGE2 stimulates cAMP levels in M-1 cells, we show that the extent of stimulation is significantly increased in cells pretreated with NS-398. Because the EP2 receptor subtype was not detected in M-1 cells, we conclude that due to the upregulation in EP4 receptors in M-1 cells, we observe an increase in PGE2 signaling via this receptor subtype when COX-2 is inhibited. Because both EP2 and EP4 receptor subtypes couple to Gs protein to stimulate adenylate cyclase, use of the PGE2 analog butaprost, a selective agonist for the EP2 and not the EP4 receptor subtype (16), would provide further verification that the cAMP effect is mediated by the EP4 receptor and not EP2; however, due to the absence of EP2 receptors, we did not feel it would be necessary to make this distinction.
Although we could not measure the levels of the EP3 receptor protein in response to NS-398, we attempted to examine whether signaling pathways via this receptor subtype are altered, by looking at the inhibitory effect of PGE2 on AVP-stimulated cAMP production. Unfortunately, we were unable to inhibit the AVP-dependent stimulation of cAMP by pretreating the cells with PGE2. Furthermore, the inhibitory response could not be induced by the NS-398 treatment of cells, although an upregulation of the EP receptor mRNA was obtained. In fact, PGE2 had an additive effect on AVP-stimulated cAMP, and this additivity was not reversed by NS-398. This lack of inhibitory response in cultured M-1 cells is consistent with previous work by Sonnenberg et al. (27) in rabbit CCD cells, suggesting the possibility that the Gi-protein signaling pathway is aberrant in rabbit CCD cultured cells. Whether this defect is due to a lack of receptor coupling to Gi protein, to an inactivation of one of the subunits, or to an absence of Gi protein due to a protein downregulation in response to immortalization procedures or culture conditions is not clear for this system. As for the signaling pathways present in the cultured M-1 cells, it is quite clear from various studies that these remain intact. For instance, Stoos (29) showed that bradykinin and atrial natriuretic factor together inhibit AVP-cAMP, but each of these compounds alone did not have this effect. Therefore it would seem that the inhibitory pathway in M-1 cells is functional, although we could not detect any inhibition in this study mediated by EP3 coupling to Gi protein. This is not due to technical difficulties, because previous work in our laboratory has shown an inhibitory effect of various prostaglandin analogs, cicaprost, iloprost, and PGE2, on AVP-stimulated cAMP production in freshly isolated rat inner medullary collecting duct (IMCD) cells (25) but not in cultured rat IMCD cells.
With respect to the EP receptors in M-1 cells, the lack of antibodies and of specific EP agonists did not allow us to study the regulation of all EP subtypes by NS-398. Nonetheless, the highest levels of EP1 are found in the collecting duct, and this receptor does couple to Gq protein to increase calcium levels (6). For example, previous work in rat terminal IMCD cells showed that PGE2 caused a rapid increase in intracellular calcium concentration ([Ca2+]i) (21). As mentioned above, EP receptors are also expressed on the nuclear envelope and can participate in the regulation of gene transcription. Furthermore, endothelin is known to increase calcium in microperfused rat terminal IMCD via the ETB receptor (22) and has been shown to increase [Ca2+]i in M-1 cells (18). Therefore, calcium signaling mechanisms are intact in M-1 cells. Because we did not pursue the analysis of EP1 receptor expression, we cannot yet dismiss the possibility that PGE2, acting through the EP1 receptor, also mediates its effect on calcium metabolism in these cells, and to what extent this response is regulated by NSAIDs.
In summary, this study examined the PGE2 signaling pathways in M-1 cells and described the postulated regulatory effects of COX-2 inhibition by NS-398 on EP receptor subtypes and their second messenger responses. As previously described, PGE2 production in the intercalated cells is enhanced by NS-398 exposure due to an increase in COX-2 (10). PGE2 can then elicit various effects in principal cells in a paracrine mode of action. In terms of cell surface EP receptor expression, we show that COX-2 inhibition by NS-398 increases the levels of EP4 and also enhances the stimulation of cAMP production by PGE2. Whether the same can be said for cell surface EP1 and EP3 receptor expression, and the signaling pathways mediated by these receptors, remains to be determined. In addition, we show that exposure of M-1 cells to NS-398 increases the expression of both EP3 and EP4 receptor mRNA. Because recent studies indicate the presence of nuclear EP1,3,4 receptors (3, 4), it would be of interest in future work to study the regulatory effect of COX-2 inhibitors on nuclear PGE2 signaling mechanisms and the regulation of gene transcription by NSAIDs. Therefore, in conclusion, this study showed the expression of EP1,3,4 receptor subtypes in M-1 mouse CCD cells. Specific inhibition of COX-2 by NS-398 revealed a threefold upregulation in both EP3 and EP4 mRNA. This was correlated with a sixfold increase in EP4 protein and an increase in PGE2-stimulated cAMP production.
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ACKNOWLEDGEMENTS |
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This research was supported by grants from the Kidney Foundation of Canada (to R. L. Hébert and O. Laneuville) and by the Medical Research Council of Canada (MT-14103; to R. L. Hébert).
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FOOTNOTES |
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Address for reprint requests and other correspondence: R. L. Hébert, Dept. of Cellular and Molecular Medicine, Faculty of Medicine, Univ. of Ottawa, 451 Smyth Rd., Rm. 1337, Ottawa, ON, Canada K1H 8M5 (E-mail: rlhebert{at}uottawa.ca).
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 5 December 2000; accepted in final form 27 February 2001.
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