We determined the effects of TNF-α on the expression of transient receptor potential channel (TRPC) homologues in human vascular endothelial cells and the consequences of TRPC expression on the endothelial permeability response. We observed that TNF-α exposure increased TRPC1 expression without significantly altering expression of other TRPC isoforms in human pulmonary artery endothelial cells (HPAEC). Because TRPC1 belongs to the store-operated cation channel family, we measured the Ca2+ store depletion-mediated Ca2+ influx in response to thrombin exposure. We observed that thrombin-induced Ca2+ influx in TNF-α-stimulated HPAEC was twofold greater than in control cells. To address the relationship between store-operated Ca2+ influx and TRPC1 expression, we overexpressed TRPC1 by three- to fourfold in the human dermal microvascular endothelial cell line (HMEC) using the TRPC1 cDNA. Thrombin-induced store Ca2+ depletion in these cells caused approximately twofold greater increase in Ca2+ influx than in control cells. Furthermore, the inositol 1,4,5-trisphosphate-sensitive store-operated cationic current was increased greater than twofold in TRPC1-transfected cells compared with control. To address the role of Ca2+ influx via TRPC1 in signaling endothelial permeability, we measured actin-stress fiber formation and transendothelial monolayer electrical resistance (TER) in the TRPC1 cDNA-transfected HMEC and TNF-α-challenged HPAEC. Both thrombin-induced actin-stress fiber formation and a decrease in TER were augmented in TRPC1-overexpressing HMEC compared with control cells. TNF-α-induced increased TRPC1 expression in HPAEC also resulted in marked endothelial barrier dysfunction in response to thrombin. These findings indicate the expression level of TRPC1 in endothelial cells is a critical determinant of Ca2+ influx and signaling of the increase in endothelial permeability.
- tumor necrosis factor-α
- store-operated calcium ion influx
- transient receptor potential channel 1
- endothelial barrier dysfunction
microvascular endothelial cells regulate the vessel wall permeability of solutes, liquid, and macromolecules and produce numerous autocrine and paracrine factors, such as NO, that modulate the contractility of the underlying vascular smooth muscle (4, 5, 21). Endothelial cells thus contribute to the maintenance of vascular tone and essential barrier property of the vessel wall (4, 5, 19, 21). Proinflammatory mediators such as thrombin and histamine and reactive oxygen species can increase vascular permeability by activating Ca2+-sensitive signaling pathways (35). In recent studies, we showed that Ca2+ entry through plasma membrane cation channels activated by Ca2+ store depletion is a critical determinant of increased endothelial permeability (30, 31, 33).
We have also shown that activation of endothelial cell surface protease-activated receptor-1 (PAR-1) by thrombin caused a rapid and transient increase in cytosolic Ca2+ concentration ([Ca2+]i) because of the release of stored Ca2+ and a subsequent Ca2+ entry triggered by store depletion (7, 20, 30). In endothelial cells, the plasma membrane cation channels known as store-operated cation channels (SOCs) mediate the entry of Ca2+ into the cell (2, 27, 32). Several studies have shown clearly influx of Ca2+ through SOCs in human vascular endothelial cells (1, 13, 25, 27, 33). Studies have also identified that the mammalian homologues of the transient receptor potential (TRP) gene family of channels function as SOCs (14, 18, 27, 29, 33, 42). These TRP genes encode a superfamily of proteins with six transmembrane helixes that are divided into the following four subfamilies: TRPC (canonical or classical), TRPV (vaniloid receptor-related), TRPM (melastatin-related), and TRPP (PKD-type) (1, 24). Members of the TRPC subfamily contain 700–1,000 amino acids, and seven isoforms (TRPC1–7) are expressed in mammalian cells. Mammalian TRPCs are grouped into the following four subfamilies. 1) One group consists of TRPC4 and TRPC5. Their activation is dependent on Ca2+ store depletion, and they have high Ca2+ selectivity, as assessed by their sensitivity to La3+ (13, 27). TRPC4 and TRPC5 are activated by G protein-coupled receptors and receptor tyrosine kinases coupled to phospholipase C. 2) TRPC1 is closely related to TRPC4 and TRPC5; although it forms SOCs, it is a less selective Ca2+ channel. 3) TRPC3, TRPC6, and TRPC7 form store-independent nonselective cation channels that may be activated by diacylglycerol (3, 24, 27); however, a store-dependent activation mechanism has been described for human TRPC3 (18). 4) TRPC2 function is unclear, and it may function as a pseudogene in humans (24).
Primary endothelial cells in culture express TRPC1–TRPC6 to varying degrees (13, 14, 33). Mouse aortic and lung endothelial cells express TRPC1, TRPC3, TRPC4, and TRPC6, but TRPC4 expression is most pronounced in mouse endothelial cells (14, 33). Deletion of TRPC4 in mouse caused a marked impairment in store-operated Ca2+ current, and the Ca2+ store release activated Ca2+ influx in aortic and lung endothelial cells (14, 33). In TRPC4 knockout mice, ACh-induced endothelium-dependent smooth muscle relaxation (an NO-mediated effect) was also drastically reduced (14). In addition, we showed that the increased microvascular permeability induced by PAR-1 activation was impaired in TRPC4 knockout mice (33). This finding suggests that the TRPC4-mediated Ca2+ influx is an important determinant of the increase in microvascular permeability. Although TRPC4 can function as SOC in mouse endothelia, recent studies have shown that expression of TRPCs in human vascular endothelial cells differs from mice (15, 25, 28, 33). However, in general, the expression profile of TRPC isoforms and their contributions in human endothelial cell signaling are not well understood.
TNF-α released during bacterial infection, such as gram-negative septicemia, contributes to the mechanism of increased microvascular permeability associated with the septicemia (8–10, 23). TNF-α also promotes increased procoagulant activity by inducing the activation of the coagulation factor thrombin (8, 9, 10, 23, 26). Recently, we showed that short-term exposure of human umbilical vein endothelial cells (HUVEC) with TNF-α augmented the thrombin-induced Ca2+ influx by increasing Src tyrosine kinase activity (36). We also showed that TNF-α increased the expression of TRPC1 by de novo protein synthesis as a result of activating the nuclear factor-κB (NF-κB) signaling pathway in HUVEC (28). This resulted in increased TRPC1 expression that in turn was associated with augmented Ca2+ influx after intracellular Ca2+ store depletion (28). Because sepsis induces both TNF-α and thrombin production, in the present study, we further addressed the relationship between TNF-α-induced expression of TRPC isoforms and the endothelial response to thrombin. We observed using RT-PCR that, in human primary endothelial cells [HUVEC and human pulmonary artery endothelial cells (HPAEC)] and the human dermal microvessel endothelial cell line [human dermal microvascular endothelial cells (HMEC)], the expression profile showed the following pattern: TRPC1 and TRPC6 expression > TRPC3 and TRPC4 > TRPC2, TRPC5, and TRPC7. TNF-α exposure selectively increased TRPC1 expression without altering the expression of other TRPC isoforms. Furthermore, the expression of TRPC1 with either TNF-α treatment or TRPC1 cDNA transfection augmented the thrombin-induced Ca2+ influx and endothelial barrier dysfunction. These results point to the important role of level of TRPC1 expression as being a key determinant of the endothelial permeability response.
MATERIALS AND METHODS
Human α-thrombin was obtained from Enzyme Research Laboratories (South Bend, IN). Anti-TRPC1 polyclonal antibody was raised in rabbits using hTRPC1 COOH-terminal synthetic peptide (QLYDKGYTSKEQKDC; see Ref. 40). TRIzol reagent, Taq DNA polymerase, LipofectAmine, LipofectAmine Plus reagent, MCDB 131, Opti-MEM I, TOPO cloning kit, and restriction enzymes were from Invitrogen (Carlsbad, CA). FuGENE 6 transfection reagent and protease inhibitor cocktail were from Roche Molecular Biochemicals (Indianapolis, IN). Fura-2 AM, Alexa-488-labeled phalloidin, and Prolong Antifade mounting medium were from Molecular Probes (Eugene, OR). PCR primers were custom synthesized from IDT (Coralville, IA). FBS was from HyClone (Logan, UT). Endothelial growth medium-2 was obtained from Cambrex Bio Science (Walkersville, MD). Human recombinant TNF-α was obtained from Alexis Biochemicals (San Diego, CA) with biological activity of 100 U/ng protein. Anti-TNF-α monoclonal antibody (mAb) was obtained from R&D Systems (Minneapolis, MN).
Primary culture human vascular endothelial cell types (HUVEC and HPAEC) obtained from Cambrex Bio Science were grown in endothelial cell growth medium-2 supplemented with 10% FBS. Both HUVEC and HPAEC were grown to confluence on tissue culture dishes precoated with 0.1% gelatin and were used for experiments between three and six passages. HMEC were grown in endothelial basal medium MCDB131 supplemented with 10% FBS, 10 ng/ml epidermal growth factor, 2 mM l-glutamine, and 1 μg/ml hydrocortisone (6). COS-7 cells were grown in DMEM supplemented with 10% FBS.
Total RNA from HPAEC, HUVEC, and HMEC was isolated using TRIzol reagent (28). RT was performed using oligo(dT) primers and superscript RT (Invitrogen) following the manufacturer's instructions. Human TRPCs and GAPDH were amplified using the following primer sets: TRPC1 (forward, 5′-GATTTTGGAAAATTTCTTGGGATGT-3′ and reverse, 5′-TTTGTCTTCATGATTTGCTATCA-3′); TRPC2 (forward, 5′-CATCATCATGGTCATTGTGCTGC-3′ and reverse, 5′- GGTCTTGGTCAGCTCTGTGAGTC-3′), TRPC3 (forward, 5′-GACATATTCAAGTTCATGGTCCTC-3′ and reverse, 5′-ACATCACTGTCATCCTCAATTTC-3′); TRPC4 (forward, 5′-GCTTTGTTCGTGCAAATTTCC-3′ and reverse, 5′-CTGCAAATATCTCTGGGAAGA-3′); TRPC5 (forward, 5′-CAGCATTGCGTTCTGTGAAAC-3′ and reverse, 5′-CAGAGCTATCGATGAGCCTAAC-3′), TRPC6 (forward, 5′-GACATCTTCAAGTTCATGGTCATA-3′ and reverse, 5′-ATCAGCGTCATCCTCAATTTC-3′), TRPC7 (forward, 5′-CAGAAGATCGAGGACATCAGC-3′ and reverse, 3′-GTGCCGGGCATTCACGTGGTA-3′) and GAPDH (forward, 5′-TATCGTGGAAGGACTCATGACC-3′ and reverse, 5′-TACATGGCAACTGTG AGGGG-3′). RT product (2 μl) was amplified in a 50-μl volume containing 100 pmol primers and 2.5 units Taq DNA polymerase. Reaction conditions were as follows: 95°C for 30 s, 55°C for 30 s, 72°C for 1 min for 35 cycles, and then 72°C for 7 min. PCR products were resolved using 1.2% agarose gel and identified by ethidium bromide staining.
Relative quantitative RT-PCR was carried out to quantify the expression of TRPC1 in control and in TRPC1 cDNA-transfected HMEC. Measurement of GAPDH expression level was used to determine the exponential phase of amplification (39). For each group, a PCR master mix was prepared with normal PCR reaction components and a corresponding primer set. After 15 cycles, samples were collected out of the thermocycler. The next samples were collected with two-cycle intervals. Nine samples from cycles 15–31 were collected for GAPDH, and 15 samples were collected between 15 and 43 cycles to determine TRPC1 transcript expression. DNA from each PCR reaction was isolated using the QIAquick PCR purification kit (Qiagen, Valencia, CA) and quantified using an ultraviolet spectrophotometer (12).
TRPC1 cDNA transfection.
Human TRPC1 (hTRPC1) expression constructs were prepared as described by us (28) and transfected in HMEC using LipofectAmine Plus transfection reagent (6). We determined transfection efficiency by using green fluorescent protein expression plasmids. We observed that ∼50% of HMEC were fluorescent 48 h after transfection (6). COS-7 cells were transfected with TRPC1 cDNA using FuGENE 6 transfection reagent. At 48 h after tranfection, cells were used for experiments.
Immunoblotting experiments were performed to detect both endogenous and overexpressed TRPC1 protein in HMEC. At 72 h after transfection with hTRPC1 cDNA expression construct, cells were lysed with lysis buffer (50 mM Tris·HCl, pH 7.5, 150 mM NaCl, 1 mM EGTA, 1% Triton X-100, 0.25% sodium deoxycholate, 0.1% SDS, and protease inhibitor cocktail), and the cell lysates were used for immunoblotting. Proteins were separated on SDS-PAGE and transferred to polyvinylidene difluoride membrane strips. Nonspecific binding was blocked with 5% nonfat dry milk in TBST (10 mM Tris·HCl, pH 7.5, 0.15 M NaCl, and 0.05% Tween 20) for 1 h at 22°C and then incubated with anti-TRPC1 antibody in TBST containing 5% nonfat dry milk overnight at 4°C. After being washed with TBST, the strips were incubated with horseradish peroxidase-conjugated goat anti-rabbit secondary antibody for 1 h. The strips were washed, and the protein bands were detected by the enhanced chemiluminescence method (Amersham Biosciences).
Cytoplasmic Ca2+ measurement.
[Ca2+]i in single endothelial cells was measured by fura-2 fluorescence imaging (37). Cells grown on 25-mm-diameter glass coverslips were washed two times in Hanks' balanced salt solution (HBSS) and loaded with 3 μM fura 2-AM for 20 min at 37°C. Cells were then washed two times in HBSS and imaged by using an Attofluor RatioVision digital fluorescence microscopy system (Atto Instruments, Rockville, MD) equipped with a Zeiss Axiovert S100 inverted microscope and F-Fluar ×40, 1.3 numeric aperture oil immersion objective. Regions of interest in individual cells were marked and excited at 334 and 380 nm with emission at 520 nm at 5-s intervals. The 334- to-380-nm fluorescence ratio has been used to represent changes in [Ca]i.
Patch-clamp recording of SOC current.
Whole cell patch-clamp was performed on single endothelial cells adhering to glass coverlips (14). Patch electrodes were made from 1.5-mm (OD) borosilicate glass tubing. The resistance of the pipettes was ∼5 MΩ when filled with internal solution (composition given below). Membrane currents were measured with a patch-clamp amplifier (Axopatch 200B; Axon Instruments, Foster City, CA) under control of pClamp 8.1 software. The currents were filtered at a corner frequency of 2 kHz with a low-pass Bessel filter and sampled at 10-ms intervals. Capacitive transients were subtracted from current records. Coverslips with cells attached were perfused at a rate of 2 ml/min; solution changes were complete within 10 s. The standard extracellular solution contained (in mM) 135 sodium glutamate, 1 MgCl2, 1 CaCl2, 10 glucose, and 10 HEPES buffer, pH 7.4. Internal solution contained (in mM) 135 N-methyl-d-glucamine, 10 CsCl, 1 BAPTA, 1 MgCl2, 1 ATP, and 10 HEPES, pH 7.2 (CsOH). All experiments were performed at room temperature.
Distribution of actin stress fibers.
Endothelial cells were grown to confluence on gelatin-coated glass cover slips. Cells were incubated with 1% FBS-supplemented growth medium for 2 h before exposure to thrombin (50 nM) for different time intervals. Cells were washed and fixed for 15 min with 2% paraformaldehyde in HBSS containing 10 mM HEPES buffer (pH 7.4) at 22°C and then permeabilized with 0.1% Triton X-100 for 30 min at 22°C. Cells were washed and stained with Alexa-568-labeled phalloidin for 30 min at 22°C. Cells were then washed three times with HBSS, mounted with Prolong Antifade mounting medium, and viewed through the ×60 objective of a Zeiss LSM 510 confocal microscope.
Transendothelial electrical resistance.
The real-time change in endothelial monolayer resistance was measured to assess endothelial barrier function, as described by us (34). In brief, HMEC or HPAEC were grown to confluence on a small gold electrode (4.9 × 10−4 cm2). The small electrode and the larger counter electrode were connected to a phase-sensitive lock-in amplifier. An approximate constant current of 1 μA was supplied by a 1-V, 4,000-Hz alternating current signal connected serially to a 1-MΩ resistor between the small electrode and the larger counter electrode. The voltage between the small electrode and large electrode was monitored by a lock-in amplifier, stored, and processed by a personal computer. The same computer controlled the output of the amplifier and switched the measurement to different electrodes in the course of an experiment. Before the experiment, the confluent endothelial monolayer was kept in 1% FBS-containing medium for 2 h, and then thrombin-induced change in resistance of the endothelial monolayer was measured. The data are presented in resistance normalized to its value at time 0, as described (7, 34). In some experiments, endothelial cells grown on the electrodes were used for transfection with TRPC1 cDNA.
Statistical comparisons were made using two-tailed Student's t-test. Experimental values were reported as means ± SE. Differences in mean values were considered significant at P < 0.05.
Effects of TNF-α on Expression of TRPC Homologs in Human Endothelial Cells
We have shown in recent studies that TRPC1 expression was increased in HUVEC by TNF-α exposure (28); however, it is not clear whether TNF-α also increases the expression of other TRPC isoforms in these endothelial cells. In this study, we determined expression of different TRPC isoforms in HUVEC, HPAEC, and HMEC (see Methods for details). We observed that TRPC1 and TRPC6 transcripts were expressed at higher levels in all three human endothelial cell types (Fig. 1A). The presence of TRPC3, TRPC4, and TRPC7 was detectable in these endothelial cells; however, TRPC2 and TRPC5 expression was not measurable (Fig. 1A).
Because all three human endothelial cell types showed a similar TRPC expression profile, we next determined the effects of TNF-α on the expression of TRPC isoforms in HPAEC. In TNF-α-stimulated HPAEC, only TRPC1 (an SOC) transcript expression increased (an increase of ∼4-fold compared with control; Fig. 1B), whereas TRPC6 expression was unaltered (Fig. 1B). Exposure of TNF-α to HPAEC in the presence of neutralizing anti-TNF-α mAb prevented the TNF-α-induced TRPC1 expression (Fig. 1B). We did not observe any change in their expression profile of TRPC2, TRPC3, TRPC4, TRPC5, and TRPC7 isoforms in response to TNF-α compared with control (data not shown). We also determined TRPC1 protein expression in HPAEC after TNF-α challenge by immunoblot. We observed that TRPC1 protein expression was increased more than twofold at 18 h after TNF-α challenge (data not shown), which is similar to what we observed in HUVEC (28).
Because our results indicated that the TRPC1 isoform was selectively increased in TNF-α-stimulated HPAEC, we addressed the possible role of TRPC1 in the mechanism of thrombin-induced increase in [Ca2+]i. In the presence of extracellular Ca2+ (1.26 mM), thrombin produced an increase in [Ca2+]i followed by a decline to baseline within 4 min after thrombin stimulation (Fig. 1C). In TNF-α-stimulated cells, however, the thrombin-induced increase in [Ca2+]i was greatly prolonged to >8 min (Fig. 1C) and returned to baseline only at 15 min after thrombin stimulation. We also compared the thrombin-induced Ca2+ influx in control and TNF-α-stimulated cells. In this experiment, cells were first stimulated with thrombin in the absence of extracellular Ca2+ to deplete endoplasmic reticulum (ER) stored Ca2+, and then Ca2+ was reapplied to assess the Ca2+ influx. In both control and TNF-α-treated cells, the thrombin-induced increase in the initial peak was similar; however, Ca2+ addition to the medium produced a twofold greater increase in Ca2+ influx in TNF-α-treated cells compared with control cells (Fig. 1D). To address the specificity of Ca2+ influx, we determined the effects of La3+ on thrombin-induced Ca2+ influx in both the control and TNF-α-treated cells. La3+ addition did not influence the initial peak Ca2+ response, but Ca2+ entry resulting from thrombin was blocked in both the control and TNF-α-treated cells (Fig. 1E). We also determined the TNF-α effect in the presence of anti-TNF-α mAb and observed that the anti-TNF-α mAb prevented the TNF-α effect (Fig. 1C).
TRPC1 cDNA Expression Increases Store-operated Ca2+ Influx in Endothelial Cells
Because TRPC1 was the predominant SOC isoform expressed in human endothelial cells and its level was selectively increased in response to TNF-α stimulation, to study the relationship between TRPC1 activation and endothelial permeability, we ectopically expressed TRPC1 in the HMEC line. To determine TRPC1 transcript copy number by relative quantitative RT-PCR, equal amounts of total RNA from the vector (pCR3.1) and TRPC1 cDNA-transfected HMEC were reverse transcribed, products were PCR amplified at different cycles, and the total amount of amplified DNA was plotted against the number of cycles to evaluate the shift in the amplification curve (Fig. 2, A and B). Relative expression levels of TRPC1 and GAPDH are shown in Fig. 2, A and B. The first detectable band of GAPDH was visible at cycle 15 for both vector (control) and TRPC1-transfected cells (Fig. 2, A and B), remained identical, and reached a plateau after 31 cycles. In the case of TRPC1-transfected endothelial cells, the amplified DNA was detectable at cycle 27, whereas in control cells TRPC1 was detectable at cycle 29 (Fig. 2, A and B). The amplification curve showed that GAPDH level was unchanged, whereas a two-cycle shift occurred in the case of TRPC1-transfected HMEC, indicating at least a fourfold increase in TRPC1 mRNA (Fig. 2B). We also determined TRPC1 protein expression in TRPC1 cDNA-transfected and control cells. At 72 h after transfection, cell lysates were immununoblotted with anti-TRPC1 antibody to assess TRPC1 protein expression. TRPC1 protein expression was increased approximately threefold (Fig. 2C) in TRPC1 cDNA-transfected HMEC compared with control (i.e., vector alone transfected).
We measured store Ca2+ depletion-mediated Ca2+ influx in control and TRPC1-overexpressing HMEC. In vector-transfected cells, thrombin caused a transient increase of [Ca2+]i followed by a gradual decline to the basal level within 3 min (Fig. 3A), whereas, in TRPC1 cDNA-transfected cells, thrombin produced an initial peak increase followed by a markedly slow decline to baseline (Fig. 3A). Next, we measured Ca2+ influx in response to thrombin-induced ER stored Ca2+ depletion in TRPC1-transfected and control cells. In this experiment, cells were first stimulated with thrombin in the absence of extracellular Ca2+ to deplete ER-stored Ca2+, and then Ca2+ was readded to assess the Ca2+ influx. Thrombin produced an initial peak increase in [Ca2+]i that rapidly returned to the baseline value in control and TRPC1 cDNA-transfected cells (Fig. 3B). The addition of Ca2+ (1.5 mM) to the extracellular medium after thrombin-induced Ca2+ store depletion caused Ca2+ entry in both control and TRPC1-transfected HMEC (Fig. 3B); however, in TRPC1-transfected HMEC, Ca2+ influx was approximately twofold greater than control, indicating that TRPC1 expression in HMEC is responsible for the increased Ca2+ influx. To address the specificity of Ca2+ influx, we measured the effect of La3+ on thrombin-induced Ca2+ influx in control and TRPC1-transfected cells. Pretreatment of cells with 100 μM LaCl3 prevented the thrombin-induced Ca2+ influx in both control and TRPC1-transfected cells (data not shown). Moreover, we transfeced TRPC1 antisense construct into HMEC and measured the thrombin-induced increase in [Ca2+]i. In TRPC1 antisense-transfected cells, Ca2+ influx was markedly reduced compared with control (Fig. 3B).
To address whether the increased TRPC1 expression has any influence on basal Ca2+ influx (i.e., constitutive channel activity), we measured Ca2+ influx in the absence of ER-stored Ca2+ depletion. In this experiment, fura 2-loaded cells were incubated in the absence of extracellular Ca2+, and then Ca2+ was added back to the extracellular medium to assess Ca2+ entry. In TRPC1-transfected cells, Ca2+ entry was markedly increased compared with vector alone-transfected cells (Fig. 3C). Moreover, LaCl3 prevented the basal Ca2+ influx in TRPC1-transfected cells (Fig. 3C), indicating that TRPC1 also regulates the constitutive Ca2+ influx pathway in endothelial cells.
To further address TRPC1 function, we transfected Myc-TRPC1 cDNA in COS-7 cells and measured the thapsigargin-induced increase in [Ca2+]i. In Myc-TRPC1-transfected cells, thapsigargin produced an approximately twofold increase in [Ca2+]i compared with vector alone-transfected cells (Fig. 4A). In this experiment, we also measured the thapsigargin-induced Ca2+ influx, as described above in Fig. 3B. We observed an approximately twofold increase in Ca2+ influx in Myc-TRPC1 cDNA-transfected COS-7 cells compared with controls (Fig. 4B).
Relationship Between TRPC1 Expression and Store-operated Current
Because TRPC1 expression augmented Ca2+ influx through SOCs, we studied the magnitude of inward current on TRPC1 activation by store Ca2+ depletion. We presumed that cell currents (I) generated by Ca2+ influx through SOCs are a function of the number (N) of open TRPC1 channels and the current through the single TRPC1 channel based on the equation: I = N × i × Popen, where i is amplitude of single channel current and Popen is steady-state open probability of the channel; thus, it is predicted that, when TRPC1 channel expression is increased, the whole cell current through the channels should rise (27). To test this idea, we dialyzed HMEC with 30 μM inositol 1,4,5-trisphosphate (IP3) through the patch pipette to open the channels. Within 2 min of patch rupture, a sustained inward current developed at −50 mV that reached a stationary level within ∼1 min. The net inward current in control HMEC stimulated by IP3 was 2.09 ± 0.18 pA/pF (n = 5; Fig. 5, A–C), whereas in TRPC1-transfected HMEC, the net inward current was 4.3 ± 0.21 pA/pF. La3+ (100 μM) almost completely blocked this current (Fig. 5B). These results demonstrate the direct relationship between TRPC1 protein expression and store-operated current in endothelial cells.
TNF-α-induced TRPC1 Expression Augments Thrombin-induced Endothelial Barrier Dysfunction
To address the relationship between increased Ca2+ influx via TRPC1 and endothelial permeability, we measured thrombin-induced actin-stress fiber formation and transendothelial monolayer resistance (TER) in control and TNF-α challenged HPAEC. In control cells, thrombin induced actin-stress fiber formation, and these cells returned to normal 60 min after thrombin treatment (Fig. 6A); however, in TNF-α-challenged cells, thrombin-induced stress fiber formation persisted for >90 min and returned to normal 2 h after thrombin stimulation (Fig. 6A). Next, we measured the change in TER in control and TNF-α-stimulated HPAEC. In controls, thrombin produced an ∼33% decrease in TER that returned to normal within 2 h after thrombin challenge (Fig. 6, B and C), whereas in TNF-α-stimulated cells an ∼55% decrease in TER was observed (Fig. 6, B and C), and the TER returned to normal 4 h after thrombin challenge (Fig. 6B).
We also measured thrombin-induced actin-stress formation and the decrease in TER in TRPC1-transfected HMEC and control HMEC. In control cells, thrombin increased the actin-stress fiber formation, and actin filamentous changes returned to normal within 2 h (Fig. 6D), whereas, in TRPC1-transfected cells, thrombin-induced actin-stress fiber showed prolonged recovery compared with control cells (Fig. 6D). Moreover, thrombin produced an ∼30% maximum decrease in TER, and TER returned to normal within 1 h after thrombin challenge in control cells (Fig. 6, E and F). However, in TRPC1-transfected cells, thrombin produced an ∼55% maximum decrease in TER, and TER returned to normal 2 h after thrombin challenge (Fig. 6, E and F).
Recent studies have shown that the members of the canonical TRPC isoform family provide the structural basis of SOCs in mammalian cells (2, 14, 27, 29, 33). Studies have also demonstrated that the pattern of expression of these TRPC isoforms differs between mouse and human endothelial cells (13, 14, 27, 33). Mouse endothelial cells express TRPC4 as the predominant isoform (14, 33), such that TRPC4 deletion [TRPC4 knockout (TRPC4−/−)] resulted in impaired SOC-mediated Ca2+ influx in endothelial cells (33). In the present study, we determined the expression of TRPC isoforms in human endothelial cells. We observed that, among the seven TRPC isoforms, TRPC1 and TRPC6 were predominantly expressed in HUVEC, HPAEC, and HMEC. Interestingly, we observed that TNF-α stimulation only increased the expression of TRPC1 in these human vascular endothelial cells. The TNF-α-induced TRPC1 expression also resulted in augmentation of Ca2+ influx in response to store Ca2+ depletion. In addition, we observed that ectopic expression of TRPC1 in HMEC mimicked the TNF-α effect in amplifying the influx of Ca2+ after store Ca2+ depletion.
Acute lung injury, acute respiratory distress syndrome (ARDS), and bacterial infection-induced sepsis are clinical manifestations of lung vascular inflammation, respiratory dysfunction, polymorphonuclear neutrophil (PMN) sequestration in the lung tissue, and the accumulation of protein-rich edema fluid in the air spaces as a result of increased permeability of both the capillary and alveolar barriers (23, 41). The increased production of inflammatory cytokines, such as TNF-α and interleukin (IL)-1β, and chemokines, including IL-8 and macrophage inflammatory protein-2α from the activated macrophages and monocytes, contributes to the vascular inflammatory process (11, 41). These mediators promote PMN sequestration by increasing the expression of endothelial cell adhesion molecules via the activation of the key inflammatory transcription factor NF-κB in endothelial cells (11, 41). TNF-α generation in the vascular bed specifically decreases the natural anticoagulant pathways (9, 10, 23) by downregulating the expression of thrombomodulin and protein C receptor in endothelial cells, and other molecules of the coagulation system that are necessary for the conversion of inactive protein C to activated protein C (9, 10, 23). The decrease in natural anticoagulant components in the vascular bed increases thrombin generation, which can activate protease-activated receptors, such as PAR-1, on endothelial cell surface (7, 38). We have shown in a number of studies that PAR-1 activation in endothelial cells increases permeability both in vitro and ex vivo (7, 31, 33, 38). Furthermore, we showed that store-operated Ca2+ influx signaling is critical in the mechanism of increased endothelial permeability (33, 35). In addition, recently, we showed that the canonical cation channel TRPC1, which is an essential component of SOC in human endothelial cells, is upregulated in response to TNF-α (28). TNF-α-induced TRPC1 expression in endothelial cells occurred via the NF-κB signaling pathway (28). Therefore, it is possible that NF-κB activation-dependent TRPC1 expression in microvascular endothelial cells may contribute to the pulmonary edema associated with ARDS.
Influx of Ca2+ via some of the TRPC isoforms (TRPC1, TRPC4, and TRPC5) is achieved by IP3-induced depletion of Ca2+ stores or preventing the Ca2+ refilling process by inhibiting the Ca2+ pump (Ca2+-ATPase) using thapsigargin. Ca2+ influx by other TRPC isoforms (TRPC3, TRPC6) is mediated by phosphatidylinositol 4,5-bisphosphate (PIP2) hydrolysis, likely via the involvement of IP3, diacylglycerol, or PIP2 (14, 17, 22). TRPC homologues can coassemble as tetramer or multimer complexes to form functional SOCs (32). Association of TRPC1 with TRPC4, or TRPC5 or TRPC1 itself was shown by coexpression and fluorescence resonance energy transfer analysis (16); however, in endothelial cells, it is unclear whether TRPC1 alone coassembles or oligomerizes with other isoforms to form functional SOCs. We have previously shown that the proinflammatory cytokine TNF-α induces TRPC1 expression in HUVEC (28). Also, the increased TRPC1 expression after TNF-α challenge was associated with augmented Ca2+ influx in response to ER-stored Ca2+ depletion (28). In the present study, we analyzed the effects of TNF-α on the expression of multiple TRPC isoforms in HPAEC. We observed that only TRPC1 expression was increased in response to TNF-α challenge. Importantly, the thrombin-induced Ca2+ influx was approximately twofold greater in TNF-α-stimulated cells than in control cells, indicating that TRPC1 expression is chiefly responsible for the enhanced Ca2+ influx.
To further address the effects of TRPC1 expression in regulating endothelial function, we also ectopically expressed TRPC1. We observed that, in TRPC1 cDNA-transfected HMEC, TRPC1 mRNA and protein levels were increased approximately fourfold and approximately threefold, respectively. We noted that the increased TRPC1 expression produced a markedly greater Ca2+ influx in response to ER-stored Ca2+ depletion. Expression of TRPC1 in COS-7 cells also showed enhanced Ca2+ influx when ER Ca2+ stores were depleted using thapsigargin. We observed additionally that the basal Ca2+ influx (i.e., constitutive channel activity) was increased in TRPC1-transfected HMEC compared with control cells. To address the relationship between TRPC1 expression and endothelial function, we measured IP3-induced store-operated current in control and TRPC1 cDNA-expressing HMEC. These results showed that the IP3-induced store-operated current was increased proportionately to the increased TRPC1 expression. Thus these results point to the role of the level of TRPC1 expression as being important in activating SOCs and mediating Ca2+ influx in human vascular endothelial cells.
We have shown previously that the increase in intracellular Ca2+ plays an essential role in signaling the increased endothelial permeability (30, 31, 33, 38). Agonists such as thrombin mediate endothelial barrier dysfunction by activating store-operated Ca2+ influx (30, 33). We showed that Ca2+ influx stimulates downstream signaling events, such as activation of myosin light chain kinase, to induce cytoskeletal reorganization, which is a prerequisite for interendothelial gap formation and the subsequent increase in endothelial permeability (35). Because we observed that TRPC1 was predominantly expressed in human endothelial cells and TNF-α challenge only increased TRPC1 expression, we addressed the relationship between Ca2+ influx through TRPC1 and the increased endothelial permeability response. We exposed HPAEC to TNF-α for 18 h [the time required to induce TRPC1 expression (28)] and assessed cytoskeletal and endothelial permeability by measuring acting stress fiber formation and TER. In control cells, thrombin induced actin-stress fiber formation, and the cells returned to normal 60 min after thrombin; however, in TNF-α-challenged endothelial cells, the thrombin-induced stress fiber formation persisted for >90 min and returned to normal levels only 2 h after thrombin stimulation. Thrombin produced an ∼33% decrease in TER, and this returned to normal levels within 2 h after thrombin challenge in control cells, whereas, in TNF-α-stimulated cells, the thrombin-induced decrease in TER was augmented, and the value returned to normal 4 h after thrombin challenge. We also determined actin-stress fiber formation and the decrease in TER in TRPC1 cDNA-transfected HMEC. In TRPC1 cDNA-transfected HMEC, the thrombin-induced actin-stress fiber formation and decrease in TER persisted for a prolonged time compared with control HMEC. Taken together, these results demonstrate that the level of TRPC1 expression is a key determinant of the Ca2+ influx via SOCs and that it plays an important role in signaling the increased endothelial permeability response.
In summary, the present study demonstrates that TRPC1 expression is selectively increased in response to TNF-α challenge of human vascular endothelial cells. We showed that increased TRPC1 expression resulted in the augmentation of store-operated Ca2+ influx, indicating the important role for TRPC1 as being an essential component of SOC in human endothelial cells. We further demonstrated a direct relationship between the store-operated Ca2+ influx through TRPC1 and the increase in endothelial permeability. These studies raise the interesting possibility that septic patients developing “leaky” vessel syndromes, such as the ARDS, may have an inappropriately elevated level of TRPC1 expression that may be capable of enhancing the store-operated Ca2+ influx and thereby increasing endothelial permeability.
This work was supported by National Institutes of Health Grants GM-58531 and P01 HL-60678.
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