TIMAP is a positive regulator of pulmonary endothelial barrier function

Abstract

TGF-β-inhibited membrane-associated protein, TIMAP, is expressed at high levels in endothelial cells (EC). It is regarded as a member of the MYPT (myosin phosphatase target subunit) family of protein phosphatase 1 (PP1) regulatory subunits; however, its function in EC is not clear. In our pull-down experiments, recombinant TIMAP binds preferentially the β-isoform of the catalytic subunit of PP1 (PP1cβ) from pulmonary artery EC. As PP1cβ, but not PP1cα, binds with MYPT1 into functional complex, these results suggest that TIMAP is a novel regulatory subunit of myosin phosphatase in EC. TIMAP depletion by small interfering RNA (siRNA) technique attenuates increases in transendothelial electrical resistance induced by EC barrier-protective agents (sphingosine-1-phosphate, ATP) and enhances the effect of barrier-compromising agents (thrombin, nocodazole) demonstrating a barrier-protective role of TIMAP in EC. Immunofluorescent staining revealed colocalization of TIMAP with membrane/cytoskeletal protein, moesin. Moreover, TIMAP coimmunoprecipitates with moesin suggesting the involvement of TIMAP/moesin interaction in TIMAP-mediated EC barrier enhancement. Activation of cAMP/PKA cascade by forskolin, which has a barrier-protective effect against thrombin-induced EC permeability, attenuates thrombin-induced phosphorylation of moesin at the cell periphery of control siRNA-treated EC. On the contrary, in TIMAP-depleted EC, forskolin failed to affect the level of moesin phosphorylation at the cell edges. These results suggest the involvement of TIMAP in PKA-mediated moesin dephosphorylation and the importance of this dephosphorylation in TIMAP-mediated EC barrier protection.

protein phosphorylation and dephosphorylation are known to be the key signaling events affecting the status of vascular endothelial barrier (11). Cytoskeletal and intercellular junctional proteins are regulated via reversible phosphorylation of serine (Ser), threonine (Thr), or tyrosine (Tyr) side chains. Based on many recent data, it is apparent that several types of protein phosphatases are intimately involved in the regulation of endothelial barrier function (10, 17, 27–29). However, their regulation is not yet completely understood.

Protein phosphatase 1 (PP1) is a multimeric phosphoserine/phosphothreonine-specific phosphatase. One of the four different isoforms, α, β, γ₁, or γ₂, of the catalytic subunit (PP1c) binds to one (or two) protein from a pool of regulatory subunits (R). The holoenzyme forms possess diverse cellular functions. A common structural element of R proteins is a short, conserved PP1c binding motif, (R/K)VXF (3, 9, 10). Different R subunits may direct PP1 holoenzymes to distinct subcellular locations and increase or suppress the activity toward specific substrates (3, 9). Myosin light chain phosphatase (or myosin phosphatase, MP), for example, is composed of PP1cβ and two regulatory subunits, namely, a larger targeting/regulatory subunit (myosin phosphatase target subunit, MYPT) and a small regulatory subunit (M20) (2, 10, 14). The activity of MP holoenzyme is increased toward phosphorylated myosin compared with the activity of the PP1c monomer (15).

It was recently shown that MP function is not limited to myosin dephosphorylation. The MP regulatory subunit MYPT1 can directly bind to F-actin binding proteins including ERM proteins (ezrin-radixin-moesin family). These proteins could be phosphorylated by either protein kinase Cθ or Rho kinase (12, 20); phosphorylation renders unfolded ERM protein, enabling its interaction with actin and membrane proteins (20, 21). ERM dephosphorylation by MP seems to affect ERM conformation and cytoskeletal/membrane binding capacities (12, 20). These data indicate that MP not only dephosphorylates myosin, but it is also involved in the regulation of F-actin cytoskeleton.

Recently, other proteins of the MYPT family, namely MYPT3, TIMAP (TGF-β-inhibited membrane-associated protein), and myosin binding subunit 85 (MBS85), were identified and characterized from different sources (8, 25, 26). They share some structural features with MYPT1, e.g., all of these proteins contain the PP1c binding motif followed by ankyrin repeats. On the other hand, MYPT3, TIMAP, and MBS85 have their own special features as well. For example, both TIMAP and MYPT3 have COOH-terminal prenylation motif suggesting possible membrane association. The high level of homology with MYPT1 implies that TIMAP, MYPT3, and MBS85 may be regulatory subunits of PP1; however, their physiological significance is not known.

TIMAP is a 64-kDa protein expressed at high levels in endothelial cells (EC). As TIMAP mRNA synthesis is strongly downregulated by TGF-β1 (8), it is likely to assume that TIMAP may be an important component of endothelial response to TGF-β1, including apoptosis, capillary morphogenesis, and barrier dysfunction. It is highly homologous to MYPT3 (∼45% amino acid homology) and shares its structural features, i.e., PP1c binding motif, ankyrin repeats, prenylation motif, and possible nuclear localization signals (8). Yeast and bacterial two-hybrid screening revealed several potential protein partners for TIMAP (1, 16). For instance, TIMAP interacts with the 37/67-kDa laminin receptor (LAMR1). It was suggested that TIMAP targets PP1c to LAMR1, and LAMR1 is a TIMAP-dependent PP1c substrate (16). Although protein-protein interaction between TIMAP and PP1c was shown by immunoprecipitation, its role in regulating PP1c activity is not clarified yet. In the present work, we present evidence for specific interaction between TIMAP and PP1cβ. Furthermore, we show that TIMAP has a barrier-protective role in human pulmonary artery endothelial cells (HPAEC), and we propose that ERM proteins are among its targets.

MATERIALS AND METHODS

Reagents.

Thrombin from human plasma and sphingosine-1-phosphate (S1P) were purchased from Sigma (St. Louis, MO). Forskolin was from Calbiochem (La Jolla, CA). Antibodies were purchased from various companies, as follows: custom-made rabbit polyclonal anti-TIMAP anti-peptide (NGDIRETRTDQENK) antibody was produced and purified by Zymed Laboratories (San Francisco, CA), rabbit polyclonal anti-PP1cβ was purchased from Upstate (Lake Placid, NY), rabbit polyclonal anti-PP1cα and anti-phosphoERM were from Cell Signaling Technology (Beverly, MA), mouse monoclonal anti-PP1c was from R&D Systems (Minneapolis, MN), goat polyclonal phospho-moesin was from Santa Cruz Biotechnology (Santa Cruz, CA), and mouse monoclonal anti-moesin antibody was from BD Biosciences Pharmingen (San Jose, CA). Alexa 488-, Alexa 594-conjugated secondary antibodies, Texas Red-phalloidin, and ProLong Gold Antifade medium were purchased from Molecular Probes (Eugene, OR). Bacterial expression vector pGEX-4T-3 (4.9 kb) was from GE Healthcare (Piscataway, NJ). Protease Inhibitor Cocktail Set III was purchased from EMD Biosciences (San Diego, CA). All other chemicals were from Sigma.

Cell line, cell culture, and cell treatment.

HPAEC (Cambrex Bio Science, Walkersville, MD) were cultured in Endothelial Cell Basal Medium-2 (EBM-2, Cambrex) supplemented with 10% FBS and EGM-2 SingleQuots (Cambrex). Cells were maintained at 37°C in a humidified atmosphere of 5% CO₂ and 95% air and were utilized at passages 6–10 (28).

Immunofluorescence.

HPAEC were plated onto glass coverslips and grown to confluence. The cells were washed once with 1× PBS (137 mM NaCl, 2.7 mM KCl, 4.3 mM Na₂HPO₄, 1.47 mM KH₂PO₄, pH 7.4) and fixed in 3.7% paraformaldehyde in 1× PBS for 10 min at room temperature (28). Between each step, the cells were rinsed three times with 1× PBS. The cells were permeabilized with 0.25% Triton X-100 in TBST (Tris buffered saline with Tween; 25 mM Tris·HCl, pH 7.5, 150 mM NaCl, 0.1% Tween 20) at room temperature for 30 min, blocked with 2% BSA in TBST (for 30 min at room temperature), and incubated with primary, then with secondary, antibodies diluted in blocking solution for 1 h at room temperature. Cover slips were rinsed and mounted in ProLong Gold Antifade and observed with a ×60 objective on a Nikon Eclipse TE300 microscope. Nonspecific binding of the secondary antibodies was checked in control experiments (data not shown). Images were processed using PhotoShop Imaging software.

Cloning and isolation of GST-TIMAP fusion protein.

Wild-type and truncated [Δ1-71 amino acid] mutant of TIMAP coding sequences were produced and subcloned into pGEX-4T-3 bacterial expression vector using standard molecular biological techniques. Namely, TIMAP coding sequences were amplified from HPAEC as described before (1). Forward primers 5′-TGGGATCCATGGCCAGTCACGTGG-3′ and 5′-TGGGATCCCTGCTGGAGGCCTCG-3′ for wild-type and truncated mutant of TIMAP, respectively; and reverse primer 5′-CGCTCGAGTCCTAGGAGATACGGCAAC-3′ for both forms were utilized. The primers were synthesized at Integrated DNA Technologies (Coralville, IA) and contained appropriate restriction sites for subcloning. The DNA sequences of the constructs were confirmed by sequencing (Sequencing Facility, Cancer Research Center, University of Chicago, Chicago, IL).

Escherichia coli BL-21 (DE3; Bioline, Randolph, MA) transformed with pGEX-4T-3 containing glutathione S-transferase (GST) alone or wild-type or truncated mutant cDNA of TIMAP fused with NH₂-terminal GST were induced with 0.1 mM isopropyl β-d-thiogalactoside and grown at room temperature with shaking for 2 h. Cells were harvested by centrifugation, and the fusion proteins were isolated from the sonicated lysates by affinity chromatography on glutathione Sepharose 4B (GE Healthcare, Piscataway, NJ) according to the manufacturer's protocol. Eluted GST fusion proteins were tested by SDS-PAGE and confirmed by Western blotting (data not shown).

In vitro GST pull-down assay.

Confluent HPAEC monolayers grown in 60-mm culture flasks were washed twice with 1× ice-cold PBS, scraped, and lysed in 600 μl of 50 mM Tris·HCl, pH 7.4, buffer containing 0.1% 2-mercaptoethanol, and protease inhibitors. The lysates (corresponding to ∼1 mg of total protein) were incubated with 1 μmol each of GST/GST-TIMAP fusion proteins coupled to glutathione Sepharose beads (corresponding to 1 Microspin column, GE Healthcare, ∼0.6 ml of glutathione Sepharose slurry) for 1 h at 4°C. Afterward, the beads were washed three times with 1× PBS, resuspended in 150 μl of 5× SDS sample buffer, boiled for 5 min, and cleared by centrifugation, and 20 μl each of the supernatants were analyzed by Western blotting.

Depletion of TIMAP in HPAEC.

HPAEC were treated with four pooled SMARTselection-designed TIMAP-specific siRNA duplexes (SMARTpool reagent) obtained from Dharmacon Research (Lafayette, CO) to decrease the amount of endogenous TIMAP. Both TIMAP-specific SMARTpool reagent and nonspecific, nontargeting siRNA duplex #1 were ordered from Dharmacon Research in ready-to-use, desalted, and 2′-deprotected form. HPAEC were transfected at 60–70% confluence with 50 nM final concentration of siRNA using DharmaFECTI transfection reagent (Dharmacon) and used for further experiments at 48-h posttransfection as described before (4).

Measurement of transendothelial electrical resistance.

Transendothelial electrical resistance (TER) was measured dynamically across confluent monolayer using an electrical cell-substrate impedance sensing system (ECIS; Applied Biophysics, Troy, NY) as described previously (23). Decreases in monolayer resistance to electrical current flow, which correlated with paracellular gap formation, were measured according to the method described by Giaever and Keese (13). HPAEC grown to 60% confluency on golden electrodes in ECIS chambers were transfected with non-silencing or TIMAP siRNA (final concentration 50 nM) using Dharmafect 1 transfection reagent (Dharmacon). An equal amount of Dharmafect was used to study the effect of transfection reagent in sham transfection experiments. Forty-eight hours after transfection, the media was changed to serum-free, and basal TER was measured. The results are presented as means + SE of three independent transfection experiments, where four chambers were analyzed for each treatment.

Immunoprecipitation.

HPAEC monolayers in 100-mm tissue culture dishes were rinsed three times with 1× PBS and then lysed with 600 μl of immunoprecipitation (IP) buffer (20 mM Tris·HCl, pH 7.4, 150 mM NaCl, 2 mM EDTA, 2 mM sodium vanadate, 1% Nonidet P-40) containing protease inhibitors. The cells were scraped, and the lysate was centrifuged with 10,000 g for 15 min at 4°C. To avoid nonspecific binding, the supernatants were precleared with 60 μl of protein G Sepharose (GE Healthcare) at 4°C for 3 h with gentle rotation. Protein G Sepharose was removed by centrifugation at 4°C for 3 min, and the supernatant was incubated with the appropriate volume of antibody at 4°C for 1 h and then with 60 μl of fresh protein G Sepharose at 4°C overnight with gentle rotation. The resin was washed three times with 300 μl of IP buffer and then resuspended in 160 μl of 1× SDS sample buffer, boiled, and microcentrifuged for 5 min. The supernatant was further analyzed by Western blotting.

RESULTS

TIMAP specifically binds β-isoform of PP1c.

The presence of PP1c binding motif in the sequence of TIMAP and its further structural similarities to MYPT led us to suggest that TIMAP is able to interact with PP1c. To study the specificity of such interaction, we created GST-tagged bacterial expression constructs of TIMAP, originally cloned from HPAEC (1). As negative control, we used TIMAP truncated mutant missing a region that contains the PP1c binding motif and a part of the first ankyrin repeat. HPAEC cell lysates were incubated with the immobilized wild-type and truncated mutant recombinants of TIMAP. Analysis of the bound proteins revealed preferential binding of the β-isoform of PP1c to wild-type TIMAP compared with the weak binding of the α-isoform (Fig. 1). As we expected, the truncated form of TIMAP was not able to render significant binding of PP1c. Indeed, bands, corresponding to GST-bound and mutant TIMAP-bound PP1c, are approximately of the same intensity, and we regard them as bands representing nonspecific binding (Fig. 1).

Fig. 1.

Specific interaction between TGF-β-inhibited membrane-associated protein (TIMAP) and PP1cβ. Bacterially expressed glutathione S-transferase (GST), GST-tagged wild-type TIMAP (wtTIMAP), and GST-tagged truncated mutant of TIMAP (mTIMAP), each bound to glutathione-Sepharose, were incubated with human pulmonary artery endothelial cell (HPAEC) lysates as described in materials and methods. After extensive washes to remove unbound material, protein complexes were eluted and probed with PP1cβ- (top) and PP1cα- (middle) specific rabbit polyclonal antibodies in Western blot experiments. HPAEC cell lysate was also loaded as positive control. The bottom panel shows Coomassie staining of GST, recombinant wtTIMAP, and mTIMAP eluted from glutathione-Sepharose in a separate experiment. Shown are representative data of at least 3 independent experiments.

Interaction of endogenous TIMAP with PP1c in HPAEC.

To assess probable colocalization of TIMAP with PP1c in HPAEC, we costained endothelial monolayers with anti-TIMAP and anti-PP1c antibodies. TIMAP appears to be present mainly at the cell membrane and in the nucleus (Fig. 2A), with some concomitant staining of the perinuclear region. On the other hand, PP1c shows more homogeneous distribution with some highlights of filamentous structure (Fig. 2C). In these experiments, we had to use monoclonal anti-PP1c antibody, which recognizes both α- and β-isoforms of the protein.

Fig. 2.

Localization of TIMAP and PP1c in HPAEC. Immunofluorescence staining of confluent HPAEC monolayers without (A, C, E) or with (B, D, F) thrombin treatment (20 nM, 30 min) was performed using anti-TIMAP (A, B; green) and anti-PP1c (C, D; red) primary antibodies. A and C, B and D are both parallel images of the same set of double-stained cells. E and F are merged images of A and C, and B and D, respectively. Arrows point to the same positions in the parallel images. No nonspecific binding of the secondary antibodies was detected in control experiments (not shown). Shown are representative data of at least 3 independent experiments.

Next, we assessed how the barrier-disruptive agent thrombin affects the intracellular distribution of TIMAP and PP1c. Thrombin treatment is known to induce reorganization of the cytoskeleton structure of EC and cell contraction. We detected certain changes in the distribution of PP1c after thrombin as the staining of cell boundaries became more prominent (Fig. 2D). To our surprise, the pattern and intensity of TIMAP staining did not change upon thrombin treatment (Fig. 2B). Merge of PP1c and TIMAP images emphasizes the overlap of staining in the nuclei of both treated and untreated cells (Fig. 2, E and F). However, at the edges of the cell, overlap became more obvious after thrombin treatment (Fig. 2, E and F).

To confirm the interaction of PP1c with TIMAP in HPAEC, we employed immunoprecipitation. Figure 3 shows that in the absence of treatment, these proteins coimmunoprecipitate with each other. Next, we tested if barrier-protective or barrier-disruptive agents change the amount of PP1c or TIMAP in coimmunoprecipitates. We have not been able to detect significant changes in either the amount of PP1c in TIMAP precipitates or in the amount of TIMAP in PP1c precipitates in the presence of the barrier-disruptive agents nocodazole and thrombin and the barrier-protective agents ATP and S1P.

Fig. 3.

Treatments enhancing or weakening barrier function do not influence TIMAP-PP1cβ interaction. TIMAP (A, C) and PP1cβ (B, D) were immunoprecipitated from lysates (precleared with protein G Sepharose) of untreated control, ATP- (10 μM, 30 min), nocodazole- (ND; 0.1 μM, 30 min), sphingosine-1-phosphate- (S1P; 1 μM, 3 h), or thrombin- (THR; 20 nM, 30 min) treated HPAEC as described in materials and mehtods. The immunoprecipitation (IP) complexes were probed for TIMAP (A, D) and PP1cβ (B, C). HPAEC cell lysate was also loaded as positive control of Western blot. Shown are representative data of at least 3 independent experiments. Bottom: represents a quantitative analysis of the Western blots performed by scanning densitometry of the membranes and expressed as average of relative density units (RDU) ± SE.

TIMAP regulates EC barrier function.

Our next question was whether TIMAP is involved in EC barrier regulation. Its specific binding to PP1cβ implies the possibility that it may regulate PP1c activity toward cytoskeleton/cytoskeleton-related proteins and control EC cytoskeleton structure. Using TIMAP-specific siRNA duplexes, we depleted TIMAP in HPAEC and measured TER of the monolayers. It was shown that changes in TER reflect changes in EC barrier integrity: increased or decreased resistance of a monolayer reflects increased barrier stability, or barrier compromise, respectively (23). Figure 4A shows that non-silencing RNA treatment did not have an effect on TIMAP protein level. Importantly, TIMAP siRNA significantly suppressed TIMAP protein content as evidenced by Western blot and immunofluorescence (Fig. 4, A and C). Measurement of basal resistance across HPAEC monolayer revealed that transfection reagent alone did not have a significant effect on TER value; however, both non-silencing and silencing RNAs slightly but noticeably reduced the basal resistance (Fig. 4B).

Fig. 4.

Effect of TIMAP depletion on agonist-induced endothelial cell transendothelial resistance (TER) change. A: the results of TIMAP expression detected by Western blot. HPAEC were grown on a 6-well plate and transfected with small interfering RNA (siRNA) as described in materials and methods. Lysates of nontransfected (1), non-silencing RNA (2), or TIMAP-specific siRNA (3) transfected cells were analyzed by Western blot using TIMAP antibody and actin antibody as loading control. The amount of TIMAP signal was expressed as ratio of TIMAP:β-actin staining. The error bars correspond to standard errors from 4 independent transfections. B: the effect of sham transfection (2), non-silencing siRNA (3), and TIMAP siRNA (4) on the basal transendothelial resistance compared with untreated cells (1). Each experiment was performed 3 times; the error bars correspond to standard errors. No statistically significant differences were found between control siRNA-treated and TIMAP siRNA-treated cells. C: immunofluorescence staining (left) and a representative immunoblot (right) of control (nontransfected), non-silencing RNA-treated, and TIMAP-depleted HPAEC monolayers and cell lysates, respectively, using anti-TIMAP primary antibody. Western blots were also treated with anti-β-actin antibody as loading control. D: control (nontransfected), non-silencing RNA-treated (nsRNA) or TIMAP-depleted (TIMAP siRNA) HPAEC were treated either with vehicle (−), 1 μM S1P, 10 μM ATP, 0.1 μM nocodazole (ND), or 20 nM thrombin (Thr) (+). TER was monitored for 3 h. Initial resistance values varied between 800 and 1,200 Ω. Relative resistance detected at maximal TER increase or decline is shown in the presence of the effectors. Data are expressed as the average of n (n = 3 or 4) experiments ± SE. Statistically significant differences were determined utilizing an unpaired Student's t-test with significance defined as P < 0.02.

We next proceeded to study the effect of TIMAP depletion on HPAEC barrier properties in the presence of barrier disruptors and barrier enhancers. The effects of both S1P and ATP, two barrier-protective agents (18, 22), were attenuated, whereas the effects of thrombin and nocodazole, both evoking barrier dysfunction (11), were enhanced by TIMAP depletion (Fig. 4D). These results clearly indicate that TIMAP is involved in the regulation of EC barrier function as a barrier-protective protein.

Immunostaining with TIMAP-specific antibody showed that S1P treatment significantly enhances TIMAP localization within the area of cell-cell contacts (Fig. 5, A and B). We did not observe significant differences in the distribution of TIMAP when the cells were treated with thrombin (Fig. 5, E and F), ATP, or nocodazole (data not shown).

Fig. 5.

Localization of moesin, phospho-moesin, and TIMAP in HPAEC. Immunofluorescence staining of confluent HPAEC monolayers without (A, C) or with (B, D) S1P treatment (1 μM, 3 h); without (E, G, I, K, M) or with (F, H, J, L, N) thrombin treatment (20 nM, 30 min) was performed using anti-TIMAP (A, B: green; E, F: red), anti-moesin (C, D: red; G, H: green), and anti-phospho-moesin (K, L) primary antibodies. Actin microfilaments were stained with Texas Red-phalloidin (M, N). Arrows point to the straightened TIMAP staining after S1P treatment (B), increased membrane localization of moesin (H, J), and phospho-moesin (L) after thrombin treatment. A and C, B and D, E and G, F and H, K and M, L and N are parallel images of the same set of double-stained cells. I and J are merged images of E and G, and F and H, respectively. Shown are representative data of at least 3 independent experiments.

Moesin is a possible target for TIMAP.

The apparent presence of TIMAP at the cell membrane and its structural features suggest possible interaction of TIMAP with membrane and membrane-associated proteins. It was shown earlier that one of the ERM proteins, moesin, associates with MYPT1 in MDCK (Madin-Darby canine kidney) cells (12). ERM proteins mediate binding of actin filaments to membrane proteins. Their conformation and binding ability may change upon phosphorylation of a threonine side chain close to their COOH terminus (21). Since MDCK cells were shown lacking endogenous TIMAP (16), we speculated that in endothelial cells expressing TIMAP at high levels, ERM proteins can interact with TIMAP along with/instead of MYPT1. Coimmunoprecipitation and pull-down experiments supported our hypothesis (Fig. 6). We observed moesin in both TIMAP and PP1c immunoprecipitates, suggesting that TIMAP may target PP1c toward moesin to allow its dephosphorylation. Immunofluorescent staining of HPAEC with moesin-specific antibody untreated showed homogeneous distribution of moesin (Fig. 5, C and G). S1P did not change the distribution pattern of moesin (Fig. 5D). However, thrombin treatment markedly enhanced moesin staining along the cell membrane (Fig. 5H). Although we did not observe a significant change in the intensity of TIMAP staining at the membrane (Fig. 5, E and F), the merged images (Fig. 5, I and J) obtained in the presence of thrombin suggest possible interaction between moesin and TIMAP in the area along the cell membrane. We have also assessed the distribution of the phosphorylated form of moesin in HPAEC. Immunostaining revealed the significant level of phospho-moesin in thrombin-treated cells, as opposed to the low level in quiescent cells. Phospho-moesin pattern did not differ from the pattern of moesin staining (Fig. 5, G and H, and K–N).

Fig. 6.

Protein-protein interactions between TIMAP, moesin, and PP1cβ. A and B: moesin, PP1c, or TIMAP (A) and PP1c or moesin (B) were immunoprecipitated from HPAEC lysates as described in materials and methods, and the IP complexes were probed for moesin (A and B) or PP1cβ (B). HPAEC cell lysate was also loaded as positive control of Western blot. C: bacterially expressed GST-tagged wild-type TIMAP (TIMAP) and GST, each bound to glutathione-Sepharose, were incubated with HPAEC lysates without or with thrombin treatment (20 nM, 30 min) as described in materials and methods. After extensive washes to remove unbound material, protein complexes were eluted and probed with moesin, phospho-ERM, or PP1cβ-specific rabbit polyclonal antibodies in Western blot experiments. HPAEC cell lysate was also loaded as positive control. Shown are representative data of at least 3 independent experiments.

It was shown in a recent work that MYPT3, the closest homologous protein of TIMAP from MYPT family, can be phosphorylated by PKA (30). Interestingly, the phosphorylation resulted in PP1c activation. The corresponding region around the phosphorylation sites in TIMAP is highly homologous.

To study the effect of PKA activation on the phosphorylation of TIMAP plausible substrate, ERM, and contractile state of EC, we next treated HPAEC with adenylate cyclase activator forskolin and assessed phospho-ERM (Fig. 7, A–D), actin (Fig. 7, E–H), TIMAP (data not shown), and moesin (data not shown) distribution by immunofluorescent staining. We have shown before that PKA activation by forskolin inhibited nocodazole-induced stress-fiber formation and barrier compromise (5). Here we show that forskolin suppressed thrombin-induced stress fiber formation (Fig. 7, F and H). More importantly, forskolin completely diminished the phospho-ERM staining along the cell edges in both thrombin-treated and untreated cells (Fig. 7, C and D). Of note, forskolin pretreatment did not change TIMAP or moesin distribution (data not shown).

Fig. 7.

Forskolin pretreatment rescues ERM phosphorylation in HPAEC upon thrombin challenge. HPAEC confluent monolayers were double-stained with anti-phospho-ERM (A–D) primary antibody and Texas Red-phalloidin (E–H) without treatment (A, E) or following various treatments as follows: B, F: 20 nM, 30 min thrombin; C, G: 50 μM, 10 min forskolin; D, H: 50 μM, 10 min forskolin followed by 20 nM, 30 min thrombin. Arrows point to the increased level of phospho-ERM at the cell boundaries after thrombin treatment (B) and to the cell edges after forskolin and thrombin challenge (D). Shown are representative data of at least 3 independent experiments.

We next studied how TIMAP depletion will affect pERM distribution and contractile apparatus assembly in thrombin-treated HPAEC. We failed to observe the effect of TIMAP siRNA on pERM and actin distribution in the presence or absence of thrombin (Fig. 8, A–D and I–L) by immunofluorescence. However, forskolin could only prevent thrombin-dependent ERM phosphorylation in the cells treated with non-silencing RNA (Fig. 8O). In contrast, TIMAP siRNA-treated cells demonstrated a high level of thrombin-dependent ERM phosphorylation at the cell boundaries in the presence of forskolin (Fig. 8H). Western blot experiments showed a less prominent effect of forskolin (Fig. 9A) on ERM phosphorylation level. One plausible explanation could be that Western blot detects total ERM phosphorylation level, whereas the immunofluorescence approach lets us monitor the phosphorylation of membrane-associated fraction. Due to the failure to detect a significant effect of forskolin on thrombin-induced ERM phosphorylation by Western blot, we were unable to dissect the effect of TIMAP depletion on this process. We next attempted to assess the effect of TIMAP on forskolin-induced barrier enhancement and forskolin-attenuated, thrombin-induced barrier disruption. Although we failed to detect the effect of TIMAP depletion on forskolin-induced changes in TER, our data of the barrier properties indicate that TIMAP depletion leads to the exacerbation of thrombin effect in the presence or absence of forskolin (Fig. 9B).

Fig. 8.

TIMAP depletion inhibits the rescuing effect of forskolin in thrombin-treated HPAEC. TIMAP-depleted (A–H) and control, non-silencing RNA-treated (I–P) HPAEC monolayers were double-stained with anti-phospho-ERM (B, D, F, H, I, K, M, O) primary antibody and Texas Red-phalloidin (A, C, E, G, J, L, N, P) without treatment (A, B, I, J) or following various treatments as follows: 20 nM, 30 min thrombin (C, D, K, L); 50 μM, 10 min forskolin (E, F, M, N); 50 μM, 10 min forskolin followed by 20 nM, 30 min thrombin (G, H, O, P). Arrows point to the increased level of phospho-ERM at the cell boundaries after thrombin challenge in TIMAP-depleted (D) and non-siRNA-treated (K) HPAEC and after forskolin and thrombin treatment of TIMAP-depleted cells (H) and to the cell edges of non-siRNA-treated cells after forskolin and thrombin challenge (O). Shown are representative data of at least 3 independent experiments.

Fig. 9.

The effect of forskolin and thrombin on ERM phosphorylation and the barrier properties of TIMAP-depleted and control HPAEC. A: HPAEC grown on 6-well plate and transfected with siRNA as described in materials and methods were pretreated with 50 μM forskolin for 10 min and then challenged with 20 nM thrombin for 30 min. Cells were washed and scraped into SDS buffer containing 20 mM NaF. Lysates were analyzed by Western blot using pERM antibody, TIMAP antibody to confirm the efficiency of TIMAP depletion, and GAPDH antibody as loading control (left). The amount of phospho signal was expressed as ratio of pERM:GAPDH staining (right). The results from each independent transfection experiment (n = 3) were normalized to the value of vehicle control sample and summarized as means + SE. B: after basal TER monitoring, the cells were pretreated with 50 μM forskolin or DMSO vehicle control. When TER increase reached plateau (after ∼30 min), cells were challenged with 100 nM thrombin. The results are presented as means + SE of maximal TER increase/decrease from 3 independent transfection experiments. *Significant difference (P < 0.05) between TIMAP siRNA-pretreated samples and corresponding samples pretreated with non-silencing RNA.

DISCUSSION

TIMAP is highly abundant in endothelial cells; still, its physiological function is not clear. Because of its structural relationship to the PP1 regulatory subunit MYPT1, TIMAP is regarded as a potential regulator of PP1 activity. Recent work shows that LAMR1 dephosphorylation by a calyculin A-sensitive protein phosphatase is TIMAP-dependent, and TIMAP was shown to coimmunoprecipitate with both LAMR1 and PP1. Therefore, it was suggested that TIMAP, as a PP1 targeting subunit, regulates LAMR1 phosphorylation (16). Furthermore, TIMAP-associated PP1c regulation of filopodia formation was shown in glomerular endothelial (GEN) cells (19).

Using different experimental tools, we provide further evidence of the physical interaction between TIMAP and PP1c. For the first time, we studied specificity of TIMAP toward PP1c isoforms and showed that TIMAP binds primarily the β-isoform compared with the α-isoform of PP1c (Fig. 1).

The NH₂-terminal nuclear localization consensus sequences and the COOH-terminal CAAX box (prenylation motif) are unique structural features shared by TIMAP and MYPT3 (8) but not by the other MYPT family members. They predict nuclear and plasma membrane localization, respectively. Accordingly, we detected endogenous TIMAP in both the nuclei and plasma membrane regions of HPAEC. In GEN and MDCK cells, recombinant TIMAP also seems to be present in the nuclei (8). In contrast, endogenous TIMAP is missing from the nuclei of GEN cells (16). This difference in TIMAP localization between HPAEC and GEN could reflect different functions performed by TIMAP in glomerular and arterial endothelial cells. Although nuclear targets of TIMAP have not been characterized yet, our data suggest that TIMAP may be involved in the regulation of some nuclear processes.

Unlike nuclear targets, at least one membrane-associated target of TIMAP was already characterized (16, 19). We studied membrane localization of TIMAP and showed an increased amount of membrane-associated TIMAP after S1P treatment. Since S1P is a known barrier protector (22), we hypothesized that TIMAP may have a positive role in endothelial barrier regulation. We checked our hypothesis by comparing TER in control and TIMAP-depleted cells. Although TIMAP depletion did not have a significant effect on basal TER (Fig. 4B), it markedly attenuated the effects of barrier-protective agents (S1P and ATP) (18, 22) (Fig. 4D). Moreover, the effects of thrombin and nocodazole, evoking barrier dysfunction (11), were significantly enhanced in TIMAP-depleting cells (Fig. 4D). These data unequivocally and for the first time demonstrate the barrier-protective role of TIMAP.

We compared the localization of TIMAP and PP1c in cells treated with the barrier disruptor thrombin. Although we observed TIMAP as highly abundant at the plasma membrane region regardless of thrombin treatment, we detected a definite increase of PP1c plasma membrane localization only after thrombin (Fig. 2). This observation suggested that TIMAP-PP1c interaction at the plasma membrane site might be subject to the regulation by the physiological state of the cell. As TIMAP is likely to be a targeting subunit of PP1, the transient colocalization with PP1c at the plasma membrane raises the question, what protein(s) might be the probable substrate(s) of PP1 at the plasma membrane?

The ERM proteins ezrin, radixin, and moesin are highly homologous in their primary structures and functions (7, 20). Serving as cross-linkers between actin filaments and plasma membranes, they are thought to be engaged in cell adhesion, microvilli formation, cell motility, etc. (6). Their structure allows variable protein-protein interactions. The COOH-terminal domain may form an intramolecular binding to the NH₂-terminal FERM domain or may bind to F-actin depending on the phosphorylation state of a conserved threonine residue (Thr567 in ezrin, Thr564 in radixin, Thr558 in moesin). On the basis of in vitro studies, it was suggested that phosphorylation of the Thr residue suppresses the intramolecular binding (21). In fibroblasts, ERM proteins are phosphorylated and relocalized to apical membrane protrusions in a RhoA-dependent manner (24). In endothelial cells, we were also able to detect ERM/moesin phosphorylation and plasma membrane association after treatment with thrombin (Figs. 5 and 7), a known Rho activator. Colocalization of TIMAP and phospho-ERM at the plasma membrane implies possible interaction between these proteins; therefore, we hypothesized that TIMAP might serve as a subunit targeting PP1c toward ERM and providing ERM dephosphorylation.

Recent work (30) showed phosphorylation of MYPT3 by PKA. It was proposed that there is a preference of PKA for sites Ser340 and Ser353 with a higher preference for Ser353 in the following region of MYPT3: ³³⁷RRTSSAGSRGKVVR RVSL³⁵⁴ (30). The corresponding region in TIMAP is almost identical: ³³⁴RRTSSAGSRGKVVRRASL³⁵¹ (the phosphorylation sites are bold, underlined amino acids are different in the 2 sequences). We detected ∼0.6 mol/mol phosphate incorporation into bacterially expressed wild-type TIMAP after in vitro PKA phosphorylation assay (data not shown). To activate PKA in HPAEC, we increased the level of cAMP by forskolin treatment. Although Western blot failed to detect a significant effect of forskolin on basal or thrombin-induced ERM phosphorylation, immunofluorescence staining showed that the membrane fraction of ERM became completely dephosphorylated both in nontransfected and transfected cells after forskolin treatment. Importantly, we detected attenuation of thrombin-induced phosphorylation of ERM in the presence of forskolin. The attenuation was obvious only in the cells with unsuppressed TIMAP expression. These seemingly conflicting effects of forskolin on phospho-ERM level detected by immunofluorescence and Western blot may suggest that PKA-phosphorylated TIMAP either regulates the localization of phospho-ERM at the membrane or contributes to localized ERM phosphorylation. Our ECIS data show that TIMAP depletion exacerbated the effect of thrombin in the presence or absence of forskolin. The fact that we could not observe the effect of TIMAP depletion on forskolin-induced barrier enhancement or forskolin-induced attenuation of the thrombin effect could be explained by the numerous pathways by which cAMP/PKA provide barrier protection. Nonetheless, we propose that TIMAP function and association with substrates, including ERM, is regulated by PKA phosphorylation, and this phosphorylation may contribute to the regulation of barrier function. Recent work published during the preparation of this paper supported our suggestion. Li and coauthors (19) demonstrated that TIMAP is phosphorylated in a PKA-dependent manner in endothelial cells. The phosphorylation status of TIMAP regulates its association with PP1c and the level of the PP1c activity associated with TIMAP.

In summary, our results show for the first time that TIMAP is important for barrier regulation as it clearly possesses barrier-protective properties. This is likely achieved via the targeting of PP1cβ rather than PP1cα toward the cytoskeletal substrates that require dephosphorylation. Our results suggest that dephosphorylation of ERM proteins is regulated by TIMAP in HPAEC. This regulation is affected by the state of PKA activation, implying the intersection of PKA-TIMAP and thrombin (RhoA or PKC)-ERM regulatory axis. Further work will be needed to demonstrate in vivo phosphorylation of TIMAP and to elucidate the signaling pathway of TIMAP-mediated regulation of ERM phosphorylation level.

GRANTS

This work was supported by National Heart, Lung, and Blood Institute Grants HL-58064, HL-67307, HL-80675, and HL-083327 (to A. D. Verin) and from the Hungarian Scientific Research Fund T043133 (C. Csortos).