Regulation of the thrombin/protease-activated receptor 1 axis by chemokine (CXC motif) receptor 4

Abstract

The chemokine receptor CXCR4, a G protein–coupled receptor (GPCR) capable of heteromerizing with other GPCRs, is involved in many processes, including immune responses, hematopoiesis, and organogenesis. Evidence suggests that CXCR4 activation reduces thrombin/protease-activated receptor 1 (PAR1)-induced impairment of endothelial barrier function. However, the mechanisms underlying cross-talk between CXCR4 and PAR1 are not well-understood. Using intermolecular bioluminescence resonance energy transfer and proximity ligation assays, we found that CXCR4 heteromerizes with PAR1 in the HEK293T expression system and in human primary pulmonary endothelial cells (hPPECs). A peptide analog of transmembrane domain 2 (TM2) of CXCR4 interfered with PAR1:CXCR4 heteromerization. In HTLA cells, the presence of CXCR4 reduced the efficacy of thrombin to induce β-arrestin-2 recruitment to recombinant PAR1 and enhanced thrombin-induced Ca²⁺ mobilization. Whereas thrombin-induced extracellular signal-regulated protein kinase 1/2 (ERK1/2) phosphorylation occurred more transiently in the presence of CXCR4, peak ERK1/2 phosphorylation was increased when compared with HTLA cells expressing PAR1 alone. CXCR4-associated effects on thrombin-induced β-arrestin-2 recruitment to and signaling of PAR1 could be reversed by TM2. In hPPECs, TM2 inhibited thrombin-induced ERK1/2 phosphorylation and activation of Ras homolog gene family member A. CXCR4 siRNA knockdown inhibited thrombin-induced ERK1/2 phosphorylation. Whereas thrombin stimulation reduced surface expression of PAR1, CXCR4, and PAR1:CXCR4 heteromers, chemokine (CXC motif) ligand 12 stimulation reduced surface expression of CXCR4 and PAR1:CXCR4 heteromers, but not of PAR1. Finally, TM2 dose-dependently inhibited thrombin-induced impairment of hPPEC monolayer permeability. Our findings suggest that CXCR4:PAR1 heteromerization enhances thrombin-induced G protein signaling of PAR1 and PAR1-mediated endothelial barrier disruption.

Chemokine (CXC motif) receptor 4 (CXCR4) is essential during embryonic development and plays multifaceted roles in biology after birth (1–4). CXCR4 is also involved in numerous disease processes, and the CXCR4 antagonist AMD3100 has already reached the clinical arena as a drug approved by the United States Food and Drug Administration for mobilization of stem cells in cancer patients (5–7).

Previously, CXCR4 agonists have been reported to reduce inflammation-induced vascular hyperpermeability in animal models and to inhibit impairment of endothelial cell barrier function induced by the thrombin/protease-activated receptor 1 (PAR1) axis (8–18). The molecular mechanisms by which CXCR4 regulates the thrombin/PAR1 axis in endothelial cells, however, are not well-understood.

The majority of research on the roles of CXCR4 in health and disease has focused on CXCR4-mediated downstream signaling events to understand the mechanisms underlying its pleiotropic functions. Nevertheless, several lines of evidence suggest that the formation of heteromeric complexes with other receptors is another molecular mechanism through which CXCR4 regulates cell function. CXCR4 has been reported to form heteromeric complexes with multiple other G protein–coupled receptors (GPCRs), such as chemokine (C-C motif) receptor 2 (CCR2), CCR5, CXCR3, atypical chemokine receptor 3 (ACKR3), chemerin receptor 23, α_1A/B/D-adrenergic receptors (ARs), β₂-AR, δ-opioid receptor, vasopressin receptor 1A, cannabinoid receptor 2, or the virally encoded GPCR of Herpesvirus saimiri, leading to altered pharmacological properties of the interacting receptor partners (19–33). In the present study, we tested whether heteromerization of CXCR4 with PAR1 could be a molecular mechanism that contributes to the previously observed cross-talk between the receptors. Here we provide evidence suggesting formation of CXCR4:PAR1 heteromers in recombinant systems and in human primary pulmonary endothelial cells (hPPECs), through which CXCR4 regulates PAR1 signaling and function upon thrombin activation.

Results and discussion

PAR1 heteromerizes with CXCR4

We utilized intermolecular bioluminescence resonance energy transfer (BRET) titration assays to test whether CXCR4 heteromerizes with PAR1 (Fig. 1A). Consistent with nonspecific bystander BRET signals, BRET signals in cells transfected with CXCR4-RLuc and YFP were low and increased linearly with increasing energy acceptor/donor ratios. BRET signals in cells transfected with CXCR4-Rluc and PAR1-YFP showed hyperbolic progression with increasing energy acceptor/donor ratios (Fig. 1A). BRET was independent of the concentrations of BRET partners when tested at fixed acceptor/donor ratios (Fig. 1B), suggesting constitutive heteromerization between CXCR4 and PAR1 in a recombinant cell system (34).

Figure 1.

PAR1 heteromerizes with CXCR4. A, HEK293T cells were transfected with a fixed amount of CXCR4-RLuc and increasing amounts of PAR1-EYFP (gray squares) or EYFP (open circles). 48 h after transfection, EYFP fluorescence and luminescence were read as described under “Experimental procedures.” Net BRET (528 nm/460 nm) was plotted against EYFP fluorescence/luminescence (EYFP/Lum). The graph is representative of three independent experiments. B, HEK293T cells were transfected with increasing amounts of both CXCR4-RLuc and PAR1-EYFP at a fixed ratio (1:10) in quadruplicate. Raw BRET (528 nm/460 nm) was plotted against total DNA amounts transfected. n = 3. C, hPPECs were transfected with PAR1 siRNA (red line) or nontargeting siRNA (blue line) and stained with anti-PAR1/Alexa 488–conjugated anti-rabbit. Black line, hPPECs labeled with Alexa 488–conjugated anti-rabbit (ctrl.). The graph is representative of n = 3. D, hPPECs were incubated with nontargeting siRNA, PAR1 siRNA, or PAR3 siRNA. PLA was performed for the detection of PAR1. Left, representative PLA images for the detection of PAR1. Images show merged PLA/4′,6-diamidino-2-phenylindole dihydrochloride signals. Scale bar, 25 µm. Right, quantification of PLA signals. n = 3 with n = 10 images/condition and experiment. *, p < 0.05 versus cells transfected with nontargeting and PAR3 siRNA. E, representative images for the detection of PAR1, CXCR4, and PAR1:CXCR4 heteromers by PLA in hPPECs. Images show merged PLA/4′,6-diamidino-2-phenylindole dihydrochloride signals. Ctrl, hPPECs were incubated with anti-PAR1 alone, and PLA was performed for the detection of PAR1:CXCR4 heteromers. Scale bar, 25 µm. Images are representative of n = 5 experiments. Error bars, S.E.

To assess whether endogenously expressed CXCR4 and PAR1 heteromerize in hPPECs, we employed proximity ligation assays (PLAs) to visualize individual receptors and proximity between CXCR4 and PAR1. To confirm selectivity of anti-PAR1 for its GPCR target (35), we first analyzed staining of hPPECs with anti-PAR1 by flow cytometry and PLA. We observed >50% reduction of signals in hPPECs after incubation with PAR1 siRNA, as compared with cells incubated with nontargeting or PAR3 siRNA (Fig. 1, C and D), suggesting sufficient selectivity of the antibody. We have validated anti-CXCR4 that we employed in PLA previously (27, 36). As shown in Fig. 1E, we detected positive PLA signals corresponding to PAR1 and CXCR4 individually and PLA signals suggesting localization of both receptors within a distance that is likely to permit direct interactions. In combination with the observed BRET signals for interactions between PAR1 and CXCR4, these data suggest that both receptors heteromerize with each other in hPPECs.

A peptide analog of transmembrane domain 2 of CXCR4 interferes with CXCR4:PAR1 heteromerization

We have shown previously that a peptide analog of transmembrane domain 2 (TM2) of CXCR4 disrupts CXCR4:α₁-adrenoceptor heteromers in an HEK293T expression system and in human vascular smooth muscle cells (27, 28, 31). Thus, we tested whether the TM2 peptide would also interfere with CXCR4:PAR1 heteromerization. As shown in Fig. 2 (A and B), incubation of hPPECs with the TM2 peptide did not significantly affect PLA signals corresponding to CXCR4 and PAR1 alone but reduced PLA signals corresponding to CXCR4:PAR1 heteromers by 57 ± 8%, as compared with cells incubated with vehicle. Incubation of cells with a control TM peptide did not significantly affect PLA signals corresponding to individual receptors or CXCR4:PAR1 heteromers in hPPECs. To confirm that the TM2 peptide interferes with CXCR4:PAR1 heteromerization, we transfected HEK293T cells, which endogenously express PAR1 (37, 38), with HA-CXCR4 and performed PLA with anti-PAR1 and anti-HA. As shown in Fig. 2 (C and D), the TM2 peptide did not affect PLA signals for individual receptors but reduced PLA signals corresponding to HA-CXCR4:PAR1 heteromers by 45 ± 4%. Because disruption of TM domains of GPCRs with TM-derived peptides can affect receptor heteromerization through interference with the correct assembly of the target membrane protein (29–31, 39, 40), our findings suggest similarity of the CXCR4 interactions site for PAR1 and α_1A/B-adrenoceptors, which appear to be distinct from the interaction sites of CXCR4 for ACKR3 (28).

Figure 2.

A peptide analog of TM2 of CXCR4 interferes with PAR1:CXCR4 heteromerization. A, hPPECs were incubated with a 50 μm concentration of the TM2 or control TM peptides at 37 °C for 30 min, followed by PLA for the detection of CXCR4, PAR1, and PAR1:CXCR4 heteromers. Images show merged PLA/4′,6-diamidino-2-phenylindole dihydrochloride signals. Scale bar, 25 µm. Images are representative of three independent experiments. B, quantification of PLA signals from three independent experiments, as in A. *, p < 0.05 versus vehicle and TM control. C, HEK293T cells were transfected with HA-CXCR4. 24 h after transfection, cells were incubated with a 50 μm concentration of the TM2 peptide or vehicle at 37 °C for 30 min, followed by PLA for the detection of HA-CXCR4, PAR1, and PAR1:HA-CXCR4 heteromers. Images show merged PLA/4′,6-diamidino-2-phenylindole dihydrochloride signals. Scale bar, 25 µm. Images are representative of three independent experiments. D, quantification of PLA signals from three independent experiments, as in C. *, p < 0.05 versus vehicle. Error bars, S.E.

The presence of CXCR4 reduces thrombin-induced β-arrestin-2 recruitment to PAR1

Because GPCR heteromerization may lead to signaling complexes with pharmacological properties distinct from the receptor protomers, we tested whether heteromerization of PAR1 with CXCR4 would affect thrombin-induced β-arrestin-2 recruitment to PAR1 utilizing the PRESTO-Tango (parallel receptorome expression and screening via transcriptional output, with transcriptional activation following arrestin-2 translocation) system (41). In this assay, HTLA cells stably expressing a tetracyclin transactivator (tTA)-dependent luciferase reporter and a β-arrestin-2–tobacco etch virus protease (TEV) fusion gene, are transfected with DNA encoding a GPCR that contains the sequences for a TEV cleavage site followed by tTA at the 3′-end (GPCR-Tango). β-Arrestin-2 recruitment upon receptor activation leads to cleavage of the TEV site and the release of tTA, which results in the transcription of luciferase, thus permitting luminescence measurements upon the addition of luciferase substrate. We transfected HTLA cells with PAR1-Tango plus pcDNA3 or CXCR4 and confirmed similar expression of PAR1-Tango and expression of CXCR4 by flow cytometry (Fig. 3A). Whereas the presence of CXCR4 did not affect the potency of thrombin to induce β-arrestin-2 recruitment to PAR1 (EC₅₀: PAR1-Tango/pcDNA3: 153 ± 78 nm; PAR1-Tango/CXCR4: 77 ± 81 nm, p > 0.05), the efficacy of thrombin to induce β-arrestin-2 recruitment was significantly reduced by the presence of CXCR4 (Fig. 3B).

Figure 3.

CXCR4 reduces β-arrestin-2 recruitment to PAR1 and enhances thrombin-induced Ca²⁺ fluxes. A, HTLA cells were transfected with FLAG-PAR1-Tango plus pcDNA3 (blue line) or HA-CXCR4 (red line), and flow cytometry was performed for the detection of FLAG-PAR1 with anti-FLAG (top) and for the detection of HA-CXCR4 with anti-HA (bottom). Gray area, unstained cells. B, PAR1 PRESTO-Tango β-arrestin recruitment assay upon stimulation with thrombin in cells transfected as in A. RLU, relative luminescence units. n = 5. *, p < 0.05 versus cells transfected with PAR1-Tango/pcDNA3. C, PAR1 PRESTO-Tango β-arrestin recruitment assay upon stimulation with thrombin after preincubation with a 20 μm concentration of the TM2 peptide (gray symbols) or the control TM peptide (open symbols) at 37 °C for 30 min. Cells were transfected as in A and B. n = 3. RLU%, relative luminescence units in percentage of cells transfected with PAR1-Tango/pcDNA3, pretreated with the control TM peptide and stimulated with 1 μm thrombin (=100%). *, p < 0.05 versus cells transfected with PAR1-Tango/pcDNA3 pretreated with the control TM peptide. D, HTLA cells transfected with FLAG-PAR1-Tango were incubated with vehicle (open circles, ctrl.) or a 50 μm concentration of the TM2 peptide (gray squares) at 37 °C for 30 min, and calcium fluxes upon thrombin stimulation (arrows) were measured. RFU, relative fluorescence units, expressed as percentage of baseline (=100%). n = 3. E, HTLA cells were transfected with FLAG-PAR1-Tango plus pcDNA3 (open bars) or FLAG-PAR1-Tango plus CXCR4 (gray bars), as in A. Cells were pretreated with a 20 μm concentration of the TM2 peptide or the control TM peptide at 37 °C for 30 min as indicated (+/−), and calcium fluxes upon thrombin stimulation (arrows) were measured as in D. Ca²⁺(dF/F0), maximal RFU minus baseline RFU (=dF) divided by baseline RFU (F0). n = 4. *, p < 0.05 versus cells transfected with FLAG-PAR1-Tango plus pcDNA3 pretreated with the control TM peptide. Error bars, S.E.

Next, we tested whether the effect of CXCR4 on thrombin-induced β-arrestin-2 recruitment to PAR1 can be reversed by interference with CXCR4:PAR1 heteromerization with the TM2 peptide. The TM2 peptide is known to function as a biased CXCR4 antagonist, which inhibits G protein signaling of CXCR4 but permits β-arrestin-2 recruitment to the receptor (40, 42, 43). After pretreatment of cells with the control peptide, the efficacy of thrombin to induce β-arrestin-2 recruitment to PAR1 was significantly reduced in cells co-expressing PAR1-Tango and CXCR4, as compared with cells transfected with PAR1-Tango alone (Fig. 3C). The presence of CXCR4 did not affect the potency of thrombin to induce β-arrestin-2 recruitment to PAR1 in cells pretreated with the control peptide (EC₅₀: PAR1-Tango/pcDNA3: 52 ± 8 nm; PAR1-Tango/CXCR4: 81 ± 14 nm, p > 0.05). These findings reproduce our observations in cells not exposed to the control peptide (Fig. 3B). Whereas pretreatment of cells with the TM2 peptide did not affect thrombin-induced β-arrestin-2 recruitment to PAR1 in cells expressing PAR1-Tango alone, it restored the efficacy of thrombin to induce β-arrestin-2 recruitment to PAR1 in cells co-expressing PAR1-Tango and CXCR4. These findings suggest that CXCR4 within CXCR4:PAR1 heteromers reduces β-arrestin-2 recruitment to PAR1 and that the TM2 peptide abolishes this effect via interference with CXCR4:PAR1 heteromerization.

The presence of CXCR4 enhances thrombin-induced G protein signaling of PAR1

To test whether heteromerization of PAR1 with CXCR4 also modulates thrombin-induced G protein signaling of PAR1, we utilized intracellular Ca²⁺ fluxes and extracellular signal–regulated kinase 1/2 (ERK1/2) phosphorylation (Thr-202/Tyr-204) as readouts. We first confirmed that the TM2 peptide does not interfere with intracellular Ca²⁺ mobilization upon thrombin stimulation in HTLA cells expressing PAR1-Tango (Fig. 3D). We then compared maximal thrombin-induced Ca²⁺ fluxes in HTLA cells expressing PAR1-Tango with or without CXCR4 after pretreatment with the TM2 or the control peptide (Fig. 3E). Ca²⁺ fluxes in HTLA cells expressing PAR1-Tango alone were indistinguishable in cells pretreated with the TM2 or control peptide. In combination with our observation that the TM2 peptide does not affect Ca²⁺ fluxes in HTLA cells expressing PAR1-Tango, when compared with vehicle-treated cells (Fig. 3D), these data indicate that both peptides do not interfere directly with PAR1 signaling.

The presence of CXCR4 significantly increased maximal Ca²⁺ responses in cells pretreated with the control peptide, but not in cells pretreated with the TM2 peptide (Fig. 3E). Similarly, thrombin-induced ERK1/2 phosphorylation in HTLA cells expressing PAR1-Tango alone was indistinguishable between cells pretreated with the TM2 or the control peptide (Fig. 4, A and B). The presence of CXCR4, however, increased peak ERK1/2 phosphorylation in cells pretreated with the control peptide, but not in cells pretreated with the TM2 peptide (Fig. 4, C and D). These observations imply that CXCR4 within CXCR4:PAR1 heteromers enhances G protein signaling of thrombin-activated PAR1, which can be prevented by disrupting PAR1:CXCR4 heteromers with the TM2 peptide. Whereas G protein–mediated ERK1/2 phosphorylation has been shown to occur rapidly and transiently, β-arrestin–mediated ERK1/2 phosphorylation occurs delayed and is sustained over longer time periods (44, 45). As shown in Fig. 4D, we observed that thrombin-induced ERK1/2 phosphorylation in cells co-expressing CXCR4 and PAR1 was significantly reduced at later time points after pretreatment with the TM control peptide, as compared with cells pretreated with the TM2 peptide. In combination with the reduced efficacy of thrombin to recruit β-arrestin-2 to PAR1 in the presence of CXCR4, our observations point toward reduced thrombin-induced β-arrestin-2–mediated signaling from the CXCR4:PAR1 heteromer.

Figure 4.

CXCR4 modulates thrombin-induced ERK1/2 phosphorylation. A and C, HTLA cells were transfected with FLAG-PAR1-Tango plus pcDNA3 (A) or with FLAG-PAR1-Tango plus CXCR4 (C) as in Fig. 3. 48 h after transfection, cells were serum-starved for 6 h and then pretreated with a 20 μm concentration of the TM2 peptide or the control TM peptide (TM ctrl.) at 37 °C for 30 min, followed by stimulation with 33 nm thrombin for 0–30 min. Cells were lysed, and lysates were used for the detection of phospho-ERK1/2 (top) and total ERK1/2 (bottom) by immunoblotting. Images are representative of five independent experiments. The migration position of molecular weight standards is indicated. B and D, densitometric quantification of the band densities as in A and C. The phospho-ERK1/2/total ERK1/2 ratio is expressed as a percentage of unstimulated cells (=100%, ctrl.). Open circles, cells pretreated with the control TM peptide. Gray squares, cells pretreated with the TM2 peptide. n = 5. *, p < 0.05 versus cells pretreated with the control TM peptide. Error bars, S.E.

Interference with CXCR4:PAR1 heteromerization inhibits thrombin-induced signaling in human primary pulmonary endothelial cells

To assess whether the TM2 peptide modulates thrombin-induced signaling in hPPECs, we measured ERK1/2 phosphorylation by Western blotting and Ras homolog gene family member A (RhoA) activation utilizing Rho-GTP pulldown assays (46, 47). Fig. 5A shows representative images from Western blotting experiments for the detection of pERK1/2 and total ERK in cell lysates from hPPECs, and Fig. 5B shows the densitometric quantification of ERK1/2 phosphorylation from five independent experiments. Treatment of hPPECs with the TM2 peptide and the control peptide did not affect ERK1/2 phosphorylation. Whereas thrombin stimulation increased ERK1/2 phosphorylation in cells pretreated with vehicle and the TM control peptide, thrombin-induced ERK1/2 phosphorylation was inhibited in cells pretreated with the TM2 peptide. Because these data suggest that disruption of endogenously expressed CXCR4:PAR1 heteromers inhibits thrombin-induced ERK1/2 phosphorylation, we tested whether depletion of CXCR4 from the cell surface of hPPECs by RNAi would also affect thrombin-induced signaling. We utilized flow cytometry to confirm knockdown of CXCR4 and to document unchanged expression of PAR1 in cells incubated with CXCR4 siRNA, when compared with cells incubated with nontargeting siRNA (Fig. 5C). Fig. 5D shows a representative image from Western blotting experiments for the detection of pERK1/2 and total ERK in cell lysates from hPPECs after incubation with NT or CXCR4 siRNA, and Fig. 5E shows the densitometric quantification of ERK1/2 phosphorylation from three independent experiments. Consistent with our observations on the effects of the TM2 peptide, depletion of CXCR4 from the cell surface of hPPECs significantly reduced thrombin-induced ERK1/2 phosphorylation.

Figure 5.

CXCR4 regulates thrombin-induced signaling in hPPECs. A, hPPECs were grown to 90% confluence and incubated with 20 μm TM2 or control TM peptides at 37 °C for 30 min, followed by treatment with 33 nm thrombin for 5 min. Cells were lysed, and lysates were used for the detection of phospho-ERK1/2 (top) and total ERK1/2 (bottom) by immunoblotting. Images are representative of five independent experiments. The migration position of molecular weight standards is indicated. B, densitometric quantification of the band densities as in A. The phospho-ERK1/2/total ERK1/2 ratio is expressed as a percentage of unstimulated cells (=100%, ctrl.). n = 5. *, p < 0.05 as indicated. C, CXCR4 gene silencing by RNAi. hPPECs were incubated with nontargeting or CXCR4 siRNA. Surface expression of CXCR4 (left) and PAR1 (right) was measured by flow cytometry. Gray, unstained cells. Blue line, cells incubated with NT-siRNA. Red, cells incubated with CXCR4 siRNA. D, hPPECs after incubation with NT-siRNA or CXCR4 siRNA, as in C, were treated with vehicle (−) or 33 nm thrombin (+) at 37 °C for 5 min. Detection of phospho-ERK1/2 (top) and total ERK1/2 (bottom) was performed by immunoblotting as in A. The image is representative of three independent experiments. The migration position of molecular weight standards is indicated. E, densitometric quantification of the band densities as in D. The phospho-ERK1/2/total ERK1/2 ratio is expressed as a percentage of unstimulated cells (=100%, ctrl.). n = 3. *, p < 0.05 for cells incubated with NT-siRNA versus CXCR4 siRNA. F, RhoA-GTP pulldown assays. hPPECs were incubated with a 50 μm concentration of the TM2 peptide or 50 nm CXCL12 at 37 °C for 30 min and then treated with 33 nm thrombin for 5 min. Cells were lysed, and lysates were used for RhoA pulldown assays. Top, immunoblot analysis (WB) for the detection of RhoA in cell lysates. Bottom, immunoblot analysis for the detection of RhoA after RhoA-GTP pulldown. Images are representative of three independent experiments. The migration position of molecular weight standards is indicated. G, densitometric quantification of the band densities as in F. The RhoA-GTP/total RhoA ratio is expressed as a percentage of unstimulated cells (=100%, ctrl.). n = 3. *, p < 0.05 versus control. Error bars, S.E.

Fig. 5F shows representative images from Western blots after RhoA-GTP pulldown from hPPEC lysates, and Fig. 5G shows the densitometric quantification of the relative amounts of GTP-bound RhoA from three independent experiments. Pretreatment of cells with the TM2 peptide or with chemokine (CXC motif) ligand 12 (CXCL12) for 30 min did not affect the RhoA-GTP/total RhoA ratios in cell lysates. The latter is consistent with the rapid and transient nature of CXCR4-mediated RhoA activation (48). Whereas the RhoA-GTP/total RhoA ratio was increased in hPPECs 5 min after thrombin stimulation, this effect was inhibited in cells pretreated with the TM2 peptide or CXCL12. Collectively, our observations in hPPECs suggest that interference with PAR1:CXCR4 heteromerization inhibits thrombin-induced G protein–meditated signaling of PAR1, which is consistent with our findings in expression systems.

CXCL12 has previously been reported to enhance normal endothelial cell barrier function through CXCR4-mediated activation of the phosphoinositide 3-kinase/Ras-related C3 botulinum toxin substrate 1 (Rac1) pathway (9). The signaling events through which CXCL12 antagonizes thrombin-induced impairment of endothelial cell barrier function, however, are unclear. Because RhoA activation is a signaling event critical to PAR1-mediated endothelial barrier disruption upon thrombin stimulation (49–51), the inhibitory effect of CXCL12 on thrombin-induced RhoA activation that we observed implies that inhibition of thrombin-induced RhoA activation contributes to protective effects of various natural and synthetic CXCR4 agonists that have been observed previously (8, 9, 12). Inhibition of ERK1/2 phosphorylation and of RhoA activation has recently been described for the agonist-bound heterodimer between CXCR4 and cannabinoid receptor 2 (26, 48). Therefore, it appears possible that the CXCR4:PAR1 heterodimer exhibits a similar pharmacological behavior.

CXCL12 stimulation reduces expression of CXCR4 and CXCR4:PAR1 heteromers but does not affect PAR1 expression

Whereas CXCR4 is known to internalize upon CXCL12 binding in a β-arrestin–mediated mechanism, we provided evidence that the ACKR3:AVPR1A and CXCR4:α_1B-AR heteromers internalize upon binding to only one of the agonists (30, 31, 52, 53). Thus, as an alternative explanation for the effects of CXCL12 on thrombin-induced RhoA activation and endothelial cell barrier function, depletion of the CXCR4:PAR1 heteromer from the cell surface upon binding of CXCL12 to CXCR4 could contribute to these effects. To test this possibility, we pretreated hPPECs with the TM2 or the TM control peptide, followed by stimulation with thrombin or CXCL12. Quantification of CXCR4 and PAR1 expression by flow cytometry (Fig. 6, A–C) and by PLA (Fig. 7, A–D) demonstrated consistently that agonist stimulation of hPPECs reduced expression of the corresponding receptors. This effect was not affected by the presence of the TM peptides. Accordingly, quantification of PLA signals showed reduction of CXCR4:PAR1 heteromer levels after stimulation with both agonists (Fig. 7E). Whereas CXCL12 stimulation did not affect PAR1 expression (Figs. 6C and 7D), thrombin stimulation reduced expression of CXCR4 in cells pretreated with the TM control peptide, but not in cells pretreated with the TM2 peptide (Figs. 6B and 7C). These data suggest that thrombin activation of PAR1 within CXCR4:PAR1 heteromers results in co-internalization of both receptors, whereas CXCL12 binding to CXCR4 within CXCR4:PAR1 heteromers selectively depletes CXCR4 from the cell surface. Because PAR1 is known to internalize via a dynamin- and clathrin-dependent pathway that is independent of β-arrestins (51, 54), differences between the mechanisms that regulate CXCR4 and PAR1 internalization likely account for such asymmetrical agonist-induced effects on the CXCR4:PAR1 heteromer. It is of note that asymmetrical agonist- and antagonist-induced effects have previously been described for other GPCR heteromers (30, 31, 55, 56). Our observation that CXCL12-induced CXCR4 internalization reduces expression of CXCR4:PAR1 heteromers without affecting PAR1 expression suggests that CXCL12 stimulation increases the proportion of PAR1 that is not in contact with CXCR4, which will enhance β-arrestin-2 recruitment to PAR1 and inhibit G protein-mediated signaling of PAR1 upon thrombin stimulation.

Figure 6.

Effects of agonist stimulation on CXCR4 and PAR1 expression in hPPECs. A, hPPECs were pretreated with 20 μm TM2 or control TM (TM ctrl.) peptides at 37 °C for 30 min, followed by stimulation with vehicle (red lines), 50 nm CXCL12 (green line), or 33 nm thrombin (blue line) at 37 °C for 30 min. Surface expression of CXCR4 (left) and PAR1 (right) was measured by flow cytometry. Gray, unstained cells. B, quantification of CXCR4 expression, as in A. CXCR4 expression is expressed as a percentage of hPPECs pretreated with the control TM peptide and stimulated with vehicle (=100%, ctrl.). n = 3. *, p < 0.05 versus control. C, quantification of PAR1 expression, as in A. PAR1 expression is expressed as a percentage of hPPECs pretreated with the control TM peptide and stimulated with vehicle (=100%, ctrl.). n = 3. *, p < 0.05 versus control. Error bars, S.E.

Figure 7.

Effects of agonist stimulation on CXCR4, PAR1 and CXCR4:PAR1 heteromer expression in hPPECs. A and B, hPPECs were pretreated with a 50 μm concentration of the control TM (TM ctrl., A) or TM2 (B) peptides at 37 °C for 30 min, washed with PBS stimulated with vehicle, 50 nm CXCL12, or 33 nm thrombin for 30 min at 37 °C. Cells were then used for PLA for the detection of CXCR4, PAR1, and PAR1:CXCR4 heteromers. Images show merged PLA/4′,6-diamidino-2-phenylindole dihydrochloride signals. Scale bar, 25 µm. Images are representative of three independent experiments. C–E, quantification of PLA signals (CXCR4 (C), PAR1 (D), and CXCR4:PAR1 heteromers (E)) from three independent experiments, as in A and B. *, p < 0.05 versus cells pretreated with the control TM peptide and stimulated with vehicle. #, p < 0.05 versus cells pretreated with the TM2 peptide and stimulated with vehicle. Error bars, S.E.

Interference with CXCR4:PAR1 heteromerization inhibits thrombin-induced disruption of endothelial barrier function

Because the TM2 peptide inhibited thrombin-induced ERK1/2 phosphorylation and RhoA activation in hPPECs, we tested whether these effects on intracellular signaling events correspond to functionally relevant effects on endothelial cell barrier function. Whereas the TM2 peptide (50 µm) did not affect hPPEC monolayer permeability for FITC-dextran, thrombin-induced hPPEC monolayer hyperpermeability could be inhibited with the TM2 peptide in a dose-dependent manner (Fig. 8A). The control TM peptide did not affect thrombin-induced hyperpermeability (Fig. 8B).

Figure 8.

A peptide analog of TM2 of CXCR4 inhibits thrombin-induced impairment of hPPEC barrier function. hPPECs were grown to a confluent monolayer on collagen-coated permeable membranes. Cells were pretreated with TM2 (A) or control peptides (B) for 10 min and then exposed to thrombin (33 nm), followed by the addition of FITC-labeled dextran. Endothelial permeability was assessed by monitoring the amount of FITC-labeled dextran that permeated through the cell monolayer by measuring fluorescence in the solution underneath the membrane insert at different time points. RFU, relative fluorescence units. n = 3 in quadruplicate. *, p < 0.05 versus thrombin alone. #, p < 0.05 versus cells pretreated with 50 μm TM2 peptide followed by thrombin exposure. Error bars, S.E.

The apparent paradox that the CXCR4 agonist CXCL12 as well as the TM2 peptide, which inhibits G protein–mediated signaling and function of CXCR4, inhibit thrombin-induced G protein–mediated signaling of PAR1 and endothelial barrier function impairment can be explained by the similarity of their effects on CXCR4:PAR1 heteromerization. Both molecules reduce expression of CXCR4:PAR1 heteromers on the cell surface without affecting PAR1 expression levels.

Whereas thrombin-induced PAR1 activation is known to impair endothelial barrier function predominantly via G protein–mediated signaling, such as ERK1/2 and RhoA activation, occupancy of endothelial protein C receptor by activated protein C has been shown to bias PAR1 signaling upon activated protein C and thrombin activation toward β-arrestin-2–mediated cytoprotective signaling (50, 51, 57). Our findings suggest that CXCR4 within CXCR4:PAR1 heteromers enhances thrombin-induced G protein–mediated signaling of PAR1 and facilitates impairment of endothelial barrier function, which can be prevented by interference with CXCR4:PAR1 heteromerization.

The possible roles of β-arrestins in this process, however, are currently difficult to assess because β-arrestin-1 and -2 fulfill distinct roles in regulating PAR1 signaling, and our observations are limited to β-arrestin-2. Unlike β-arrestin-1, β-arrestin-2 has been reported not to play a significant role in PAR1 uncoupling from G protein signaling (54, 58). In contrast, cytoprotective β-arrestin–mediated signaling of PAR1 has been attributed to β-arrestin-2 (50, 51, 57). Our observations on the effects of CXCR4 on thrombin-induced β-arrestin-2 recruitment to PAR1 and on the temporal pattern of thrombin-induced ERK1/2 phosphorylation are consistent with reduced β-arrestin-2–mediated signaling from the CXCR4:PAR1 heteromer.

Thus, CXCR4 within CXCR4:PAR1 heteromers may bias PAR1 signaling by enhancing G protein and reducing β-arrestin-2 signaling. Because thrombin, however, is known to recruit β-arrestin-1 and -2 to PAR1, it appears possible that the efficacy of thrombin to recruit β-arrestin-1 to PAR1 within CXCR4:PAR1 heteromers is also reduced, which would imply reduced PAR1 uncoupling of G protein signaling as a mechanism by which CXCR4 enhances G protein signaling. Detailed studies on the functional roles of β-arrestins in thrombin-induced signaling from the CXCR4:PAR1 heteromer will be required to answer these questions in the future.

Conclusively, the findings of the present study identify PAR1 as another GPCR heteromerization partner of CXCR4 and provide evidence for pharmacologically relevant functions of the CXCR4:PAR1 heteromer. Our findings suggest a molecular mechanism that likely contributes to the previously described protective effects of CXCR4 agonists on thrombin-induced endothelial barrier function impairment in vitro and on inflammation-induced vascular hyperpermeability in preclinical disease models. Our observations further support the concept that the development of drugs targeting GPCR heteromers could provide new therapeutic opportunities in the future.

Experimental procedures

Proteins, peptides, and reagents

CXCL12 was purchased from Protein Foundry (Milwaukee, WI, USA) and human α-thrombin from Enzyme Research Laboratories (South Bend, IN, USA). The peptide analog of TM2 of CXCR4 was as described (28, 31, 42, 43, 59). A peptide analog of TM4 of α_1A-AR (IVNLAVADLLLTSTVLPFSAIFEVDDD) was used as a control peptide. Solid-phase synthesis on a 433A Applied Biosystems Peptide Synthesizer using Fmoc (N-(9-fluorenyl)methoxycarbonyl) amino acid derivatives was used for the production of both peptides. After cleavage with 87.5% (v/v) TFA containing 5% (v/v) water, 5% (v/v) thioanisol, and 2.5% (v/v) triisopropylsilane, the peptides were purified by reverse-phase HPLC using an Atlantis C3 column (Agilent Technologies). The peptide structure and purity were confirmed by ion-spray MS combined with HPLC. Accell PAR1, PAR3, CXCR4, and nontargeting siRNA were purchased from GE Dharmacon.

Plasmid construction

pIRES-cMyc-hCXCR4var2-Rluc (CXCR4-RLuc) was kindly provided by Dr. Michel Bouvier. The FLAG-tagged Tango plasmid (F2R-TANGO or PAR1-TANGO, accession no. 66276) was from Addgene, deposited by the laboratory of Dr. Bryan Roth (41). HA-tagged CXCR4 was as described (29, 31). To construct the PAR1-EYFP fusion protein, cDNA encoding EYFP was amplified from pEYFP (Clontech) and inserted at the restriction sites AgeI and XbaI of PAR1-TANGO and fused to the C terminus of PAR1. All plasmids were verified by DNA sequencing.

Cells and cell lines

hPPEC (ATCC (Manassas, VA), PCS-100-022) were cultured in vascular cell basal medium (ATCC, PCS-100-030) with endothelial cell growth kit-VEGF (ATCC, PCS-100-041). HEK293T cells (ATCC, CRL-11268) were cultured in DMEM. The HTLA cell line, a HEK293 cell line stably expressing a tTA-dependent luciferase reporter and a β-arrestin2-TEV fusion gene, was generously provided by the laboratory of Dr. Bryan Roth (41) and maintained in high-glucose Dulbecco's modified Eagle's medium supplemented with 10% fetal bovine serum, 100 units/ml penicillin, 100 μg/ml streptomycin, 100 µg/ml hygromycin B, and 2 µg/ml puromycin. All cells were cultured in a humidified environment at 37 °C, 5% CO₂.

Gene silencing by RNAi

hPPECs were transfected with nontargeting siRNA (siCTL), PAR1 siRNA, PAR3 siRNA or CXCR4 siRNA at a final concentration of 1 μm using Accell Delivery Media (GE Dharmacon) as previously described (27, 29, 30).

In vitro endothelial cell permeability assays

Permeability assays with hPPECs were performed as described previously (12). In brief, 96-well collagen-coated permeability assay plates were prehydrated for 15 min, and 5 × 10⁵ cells were seeded on each well and grown to a confluent monolayer for 48 h. FITC-dextran (20 μg/ml) was then added on top of the monolayer, and the amount of FITC-dextran that permeated through the monolayer was quantified by measuring fluorescence in a Synergy 2 multimode microplate reader (BioTek, Winooski, VT, USA) at various time points over a 255-min time period.

PLAs

PLAs were performed as described (12, 27, 29, 31). The following primary antibodies were utilized for the detection of individual receptors and receptor-receptor interactions: rabbit anti-PAR-1 (Abcam (Cambridge, UK), ab63445), goat-anti-CXCR4 (Abcam, ab1670), and mouse anti-HA (Thermo Fisher Scientific, 26183). All primary antibodies were used in a 1:750 (v/v) dilution. Comparisons and statistical analyses were performed only when PLA assays were performed on the same day in parallel experiments, and fluorescence microscopy was performed with identical settings.

PRESTO-Tango β-arrestin-2 recruitment assay

The PRESTO-Tango assay was performed as described previously (12, 29–31, 41, 42). HTLA cells (2.5 × 10⁵/well) were seeded in a 6-well plate and transfected with 750 ng of each plasmid (PAR1-Tango plus pcDNA or PAR1-Tango plus HA-CXCR4) using Lipofectamine 3000 (Thermo Fisher Scientific). The following day, transfected HTLA cells (75,000 cells/well) were plated onto poly-l-lysine–precoated 96-well microplates and grown for 24 h. After cells were treated with thrombin overnight, culture medium was removed and replaced with a 100-μl 1:5 mixture of Bright-Glo (Promega) and 1× Hanks' balanced salt solution, 20 mm HEPES solution. Plates were then incubated at room temperature for 20 min before measuring luminescence on a Biotek Synergy II plate reader. When HTLA cells were not transfected with a Tango plasmid, no change in luminescence was detectable upon agonist treatment.

BRET assays

BRET assays were performed as described (30, 59). In brief, HEK293T cells were seeded in 12-well plates and transfected with the plasmids indicated using the Lipofectamine 3000 transfection reagent (Thermo Fisher Scientific). For BRET titration assays, CXCR4-RLuc at a fixed amount of 50 ng was transfected alone or with increasing amounts of EYFP or PAR1-EYFP. For BRET assays at a constant energy donor/acceptor ratio, increasing amounts of both CXCR4-RLuc and PAR1-EYFP were co-transfected at a ratio of 1:10. In all assays, empty vector pcDNA3 was added to maintain the total DNA amount for each transfection constant. After an overnight incubation, cells were seeded in poly-l-lysine–coated 96-well white plates and incubated again overnight. Cells were then washed with PBS, and fluorescence was measured in a Biotek Synergy HT4 plate reader (excitation 485 nm, emission 528 nm). For BRET measurements, coelenterazine H was added at a final concentration of 5 μm. After a 10-min incubation at room temperature, luminescence was measured at 460 and 528 nm. The BRET signal was calculated as the ratio of the relative luminescence units (RLU) measured at 528 nm over RLU at 460 nm. The net BRET is calculated by subtracting the BRET signal detected when CXCR4-RLuc was transfected alone. For titration experiments, net BRET ratios are expressed as a function of fluorescence/total luminescence.

Immunoblotting

Immunoblotting with anti-phospho-ERK1/2 (Thr-202/Tyr-204), total ERK1/2, and anti-RhoA (all from Cell Signaling Technology) was performed as described (27, 60, 61). Densitometric quantifications of the band densities were performed with the Quantity One software (Bio-Rad).

RhoA-GTP pulldown assays

RhoA-GTP pulldown assays were performed using the Active Rho Detection Kit (Cell Signaling Technology) according to the manufacturer's instructions. In brief, cells were serum-starved for 5 h; exposed to vehicle, the TM2 peptide, or CXCL12 for 30 min; and then treated with thrombin for 5 min. Cells were washed with ice-cold PBS and lysed in 25 mm Tris, pH 7.5, 250 mm NaCl, 0.05% Triton X-100, 0.25% sodium deoxycholate, 0.05% SDS, and 5 mm MgCl₂, supplemented with protease inhibitors. The cell lysate was incubated at 4 °C for 1 h with GST-rhotekin RBD attached to GSH-agarose beads. The beads were then washed three times with wash buffer (50 mm Tris, pH 7.5, 75 mm NaCl₂, 0.5% Triton X-100, and 5 mm MgCl₂) and boiled in SDS-PAGE sample buffer. Activated RhoA was estimated by comparing pulldown RhoA-GTP eluted from the agarose beads versus total RhoA in the cell lysates by immunoblotting.

Flow cytometry

Flow cytometry after labeling cells with phycoerythrin-conjugated anti-FLAG (BioLegend (San Diego, CA), 637310), FITC-conjugated anti-HA (Sigma-Aldrich, H7411), or rabbit anti-PAR1 (Abcam, ab63445) and corresponding secondary Alexa 647– or Alexa 488–conjugated antibodies was used to quantify receptor expression levels in HTLA cells and in hPPECs, as described previously (30, 31). At least 10,000 cells/sample were recorded and analyzed with FlowJo software (FlowJo LLC, Ashland, OR).

Ca²⁺ assay

Intracellular calcium was measured using the FLIPR Calcium 6 assay kit (Molecular Devices), as described previously (27, 62, 63).

Data analyses

Data are presented as mean ± S.E. of n independent experiments performed on different days. Data were analyzed with unpaired Student's t test and one- or two-way analyses of variance with Bonferroni's multiple-comparison post hoc test, as appropriate. Dose–response curves were generated using nonlinear regression analyses. All analyses were performed with GraphPad Prism 8, version 8.4.0 (GraphPad Software, Inc., La Jolla, CA, USA). A two-tailed p < 0.05 was considered significant.

Data availability

All data are contained within the article.