Characterization and Functional Activity of Thrombin Receptors in the Human Lens

Abstract

purpose. To investigate the expression of thrombin receptors in the human lens, the activation of downstream signaling pathways, and the ability of thrombin to regulate lens cell growth.

methods. Thrombin receptor function in the human lens was determined first by measuring changes in intracellular calcium in response to thrombin and protease-activated receptor–activating peptides (PAR-APs). In the human capsular bag model, cell growth was assessed by phase microscope inspection of the cell coverage of the posterior capsular surface. In the human lens cell line FHL124, it was assessed by [³H]thymidine incorporation. Changes in p42/p44 ERK phosphorylation (p-ERK) and protein kinase B (PKB/Akt) phosphorylation (p-Akt) were monitored by Western blot. Reverse transcription–polymerase chain reaction (RT-PCR) applied to isolated lens epithelia and ex vivo capsular bag preparations as well as FHL124 cells determined expression of mRNA for the PARs.

results. Brief exposures to thrombin (10 nM) and PAR1-AP (10 μM) induced an increase in cytosolic calcium in both anterior and equatorial lens cells, but activating peptides for PAR2, -3, and -4 failed to produce responses. Repeated exposure to thrombin produced a significant increase in cell coverage in the capsular bag model and increased [³H]thymidine incorporation into FHL124 cells. In the latter, exposure to thrombin (10 nM) and PAR1-AP (10 μM) induced biphasic increases in the phosphorylation of p42/p44 (p-ERK), with peak responses at 20 minutes and 12 hours. Thrombin also produced a 20-fold increase in p-Akt at 12 hours compared with the control, whereas PAR1-AP (10 μM) induced a much smaller response. PAR1-AP did not induce a significant increase in [³H]thymidine incorporation and PAR2-AP, PAR3-AP, and PAR4-AP failed to reproduce any of the thrombin-stimulated effects. mPAR1 and -3 were expressed in native lens cells, and this expression was conserved in ex vivo capsular bag preparations as well as in FHL124 cells.

conclusions. This study identifies thrombin receptors coupled to calcium, ERK, and Akt signaling that modulate growth in native lens tissue and cultured cells, and it appears that the PAR1 subtype is mainly responsible. PAR3 mRNA was also detected, but the receptor itself, if present, was not coupled to the above signaling elements.

The serine protease thrombin is an essential element in the blood coagulation cascade. ¹ However, it is now emerging that thrombin has additional functions in inflammation, wound healing and tissue remodeling, growth factor activation, embryogenesis, and both normal and aberrant growth control. ^{2 3 4 5 6} The most probable store of thrombin is the blood in the precursor form prothrombin, and introduction of the blood contents to a site of injury greatly influences the wound healing response. ^{7 8} Although prothrombin activation is best understood in terms of blood coagulation, it appears that the aqueous humor has coagulation properties. It has been shown that human aqueous applied to standard ear lobe punctures shortens the bleeding time. ⁹ This property may reside in a novel membrane-associated, prothrombin-activating protease that has been shown to be present in cells that are separated from direct contact with blood. ¹⁰  

Thrombin exerts its effect through a family of G-protein-coupled receptors, known as the protease-activated receptors (PARs), which use a novel mechanism of activation. Thrombin binds to the extracellular N terminus domain of the receptor, cleaving the peptide bond between the residues arginine-41 and serine-42. This proteolytic event reveals a new N terminus with a sequence, unique for each receptor subtype, that acts as a tethered ligand. The newly exposed sequence then binds to the body of the receptor, leading to irreversible activation. Currently, there are four known PARs (PAR1–4). PAR1, -3, and -4 are activated by thrombin, whereas PAR2 is activated by trypsin and mast cell tryptase. ¹¹ The PARs are known to couple to several downstream signaling pathways that include inhibition of adenylyl cyclase ^{12 13} and activation of the mitogen-activated protein kinase (MAPK) ⁴ and PI3K/Akt ¹⁴ pathways. However, in most cases, the preferred coupling mechanism appears to be via phospholipase-Cβ (PLCβ), generating the second-messenger inositol trisphosphate (InsP₃). This in turn activates InsP₃ receptors and releases calcium (Ca²⁺) from endoplasmic reticulum (ER) stores into the cytosol. 

Exposure to thrombin has been shown to initiate proliferation in several cell types, for example in rat astrocytes ⁴ and canine tracheal smooth muscle cells. ¹⁵ Furthermore, it has been shown in a mouse model that thrombin and thrombin receptor agonists can accelerate wound healing by promoting fibroblast and epithelial cell proliferation. ⁸ Other studies have started to identify specific PAR receptors involved in growth responses. A functional PAR1 exists in colonic cells and, when activated by thrombin, enhances proliferation and motility. ⁵ PAR4 has been reported to contribute to microglial proliferation, and inhibition of PAR4 could be a therapeutic target to prevent neuronal trauma. ¹⁶ Recently, Lang et al. ¹⁷ have shown that exposure of human corneal epithelial cells to thrombin induces secretion of the proinflammatory cytokines interleukin (IL)-6, IL-8, and TNFα through activation of PAR1 and -2 receptors. In the porcine lens, thrombin has been found to inhibit sodium-potassium (Na⁺,K⁺) transport ¹⁸ and, in an earlier study, thrombin was found to stimulate cell division in the cultured rabbit lens. ¹⁹ Also, in a human lens cell line, thrombin appears to initiate actin reorganization and increase sites of focal adhesion. ²⁰ These studies ^{18 19 20} confirm that lens cells are responsive to thrombin, but do not identify either the PAR subtypes involved or the signaling pathways that mediate the responses. 

Levels of blood proteins in the aqueous humor of the eye are normally low, and passage of proteins across the blood–aqueous barrier are likely to be minimal. In a diseased state, however, aqueous humor protein levels can be elevated due to disruption or weakening of the blood–aqueous barrier. ²¹ Furthermore, the presence of thrombin has been identified in samples of subretinal fluid removed from patients with perforated retinal detachment. ²² During cataract surgery, the blood–aqueous barrier can be breached, and this can introduce blood proteins, including (pro)thrombin, into the ocular environment. Thrombin entry into the interior of the eye via the iris is believed to play a critical role in lens regeneration. ²³  

The current work was undertaken to investigate the expression of thrombin receptors in human lens cells, the activation of downstream signaling pathways, and the ability of thrombin to regulate lens cell proliferation. Functional thrombin receptors were identified in the intact human lens, and activation of these receptors resulted in an increase in intracellular calcium ([Ca²⁺]_i) and an increase in cell growth across the posterior capsule. An upregulation of phosphorylated ERK and Akt proteins and an increase in lens cell proliferation were also observed in FHL124 cells. Furthermore, the data provide evidence that mRNA for PAR1 and -3 was expressed in native lens cells, and this expression was conserved in ex vivo capsular bag preparations, as well as in the human lens cell line. 

Methods

Thrombin, PAR1 (TFLLRN) and -2 (SLIGRL) activating peptides were obtained from Tocris Cookson, Ltd. (Avonmouth, UK). PAR3 (TFRGAP) and -4 (AYPGKF) activating peptides were obtained from CN Biosciences, Ltd. (Nottingham, UK). Unless otherwise indicated, all other chemicals were supplied by Sigma-Aldrich, Ltd. (Poole, UK). 

After removal of corneoscleral discs for transplant purposes, human donor eyes were obtained from the East Anglian or Bristol Eye Banks. In this study donors were aged between 18 and 86 years. The use of human tissue was in accordance with the provisions of the Declaration of Helsinki. 

Native Human Lens Preparations

For whole-lens studies, the lens was removed from the globe by dissecting it from the surrounding tissue and was placed anterior surface down on a plastic Ca²⁺-imaging chamber (described by Collison and Duncan ²⁴ ). It was then perifused at 35°C, the optimum temperature for the lens, ^{25 26} with artificial aqueous humor (AAH) of the following composition (in mM): KCl, 5; NaHCO₃, 5; glucose, 5; HEPES, 20; NaCl, 130; MgCl₂, 0.5; and CaCl₂, 1. The pH was adjusted to 7.25 by the addition of 4 M NaOH. In some cases, isolated anterior epithelial preparations were generated. The lens capsule, with its adherent epithelium was dissected from the fiber mass as described by Collison et al. ²⁷ and pinned to the base of the Ca²⁺-imaging chamber. 

The capsular bag model has been described in detail elsewhere. ²⁸ Briefly, a sham cataract operation was performed, and the resultant capsular bag was dissected from the zonules and secured on a sterile 35-mm polymethylmethacrylate Petri dish. Entomology pins (D1; Watkins and Doncaster, Ltd., Kent, UK) were inserted through the edge of the capsule to retain its circular shape. Experiments were performed on matched pairs of capsular bags that were maintained in Eagle’s modified essential medium (EMEM) or EMEM supplemented with 10 nM thrombin. The media were replaced every 48 hours, and ongoing observations of cell coverage across the posterior capsule were performed by phase contrast microscopy (TE-200 Eclipse microscope fitted with a Coolpix 950 digital camera and MDC lens; Nikon Industries, Tokyo, Japan). Examples of cell coverage maps generated in this way are given in Liu et al. ²⁸ For ex vivo specimens, donor eyes contained capsular bags containing intraocular lenses (IOLs) that had been generated by cataract surgery. The capsular bag was dissected from the zonules and snap frozen before RNA extraction. All specimens showed signs of cell growth and of posterior capsule opacification (PCO). ²⁹  

Cell Line

FHL124 cells (see Wormstone et al. ³⁰ for details) were suspended in EMEM supplemented with 5% fetal calf serum (FCS) and gentamicin (50 μg/mL), and 10⁴ cells (in 400 μL EMEM) were seeded onto the center of round, glass coverslips (diameter 16 mm, No. 0; BDH, Poole, UK). The coverslips were placed in 35-mm Petri-dishes and incubated for 4 hours at 35°C in a 5% CO₂ atmosphere to allow the cells to adhere before adding 1 mL of the same medium. After the cells were cultured for 48 hours, the medium was exchanged for serum-free EMEM for a further 24 hours of culture at 35°C in 5% CO₂. The glass coverslip eventually formed the base of a modified Ca²⁺-imaging chamber. 

Measurement of [Ca²⁺]_i

The technique for measuring [Ca²⁺]_i has been described in detail. ²⁴ All cell and tissue preparations were loaded with 3 μM Fura-2 AM (the acetoxymethylester form) for 40 minutes at room temperature. The preparations were then washed in AAH for 20 minutes to allow complete de-esterification of the dye and to remove any extraneous dye before measuring [Ca²⁺]_i levels. An epifluorescence microscope (Eclipse TE-200; Nikon) fitted with a ×20 objective was used for real-time ratiometric imaging of [Ca²⁺]_i. Cultured cells were large enough to be imaged individually, but native cells were imaged as regions of interest containing approximately 15 to 20 cells. Lens preparations were bathed in warmed (35°C) AAH in which the various agonists were dissolved when required. Cells were alternately excited at 340- and 380-nm wavelengths, with the resultant fluorescent emissions collected at 510 nm. For technical reasons (see Ref. ²⁴ ) the Ca²⁺ data are presented in ratio, rather than calibrated, form. 

Reverse Transcription–Polymerase Chain Reaction

FHL124 cells were grown on 35-mm tissue culture dishes to 85% to 90% confluence in EMEM supplemented with 5% FCS and gentamicin (50 μg/mL) and then cultured in serum-free EMEM for 24 hours. Human lens epithelial cell RNA was collected (RNeasy Mini Kit; Qiagen Ltd., Crawley, UK). RNA (1 μg) was reverse transcribed in a 20-μL reaction mixture (Superscript II RT; Invitrogen Ltd., Paisley, UK). A 0.2-μL portion of the cDNA was amplified by PCR in a 100-μL reaction buffer in the following conditions: 0.2 μM each primer (Invitrogen Ltd., Paisley, UK), 0.2 mM deoxy-nucleoside trisphosphate mixture (Bioline Ltd., London, UK), 10 mM Tris-HCl, 1.5 mM MgCl₂, 50 mM KCl, and 2.5 U Taq DNA polymerase (Roche, Lewes, UK). The PCR was performed by using the following program with a thermal controller (MJ Research Inc., Reno, NV): initial denaturation 94°C for 4 minutes, denaturation at 94°C for 1 minute, annealing at 55°C for 1 minute, and extension at 72°C for 1 minute. Steps 2 through 4 were cycled 30 times for PAR1, -2, and -4; 33 times for PAR3; and 27 times for GAPDH, with a final extension at 72°C for 10 minutes. The oligonucleotide primer (5′–3′) sequences specific for PAR types 1 to 4 ³¹ and GAPDH ³⁰ are listed in Table 1 . PCR products, together with DNA base pair markers (Invitrogen-Life Technologies, Gaithersburg, MD), were run on a 0.8% agarose gel, and images were captured and analyzed (1D, ver. 3.5 software; Kodak Scientific Imaging Systems, Rochester, NY). 

Western Blot Detection of Phosphorylated ERK and Akt Proteins

Cells were grown on 35-mm tissue culture dishes to 85% to 90% confluence in EMEM supplemented with 5% FCS and gentamicin (50 μg/mL), incubated in serum-free EMEM for 24 hours, and treated with EMEM supplemented with thrombin (10 nM), activating peptides for PAR types 1 to 4 (10 μM) or serum-free EMEM alone for 20 minutes. Cells were then lysed on ice in buffer (50 mM HEPES [pH 7.5], 150 mM NaCl, 1% Triton X-100, 1 mM EDTA, 10% glycerol, 10 mM sodium pyrophosphate, 2 mM sodium orthovanadate, 10 mM sodium fluoride, 1 mM phenylmethylsulfonyl fluoride and 10 μg/mL aprotinin). ³² Lysates were precleared by centrifuging at 13,000 rpm at 4°C for 10 minutes and the protein content assayed by the bicinchoninic acid (BCA) assay (Perbio Science Ltd, Chester, UK) so that equal amounts of protein per sample were loaded into 10% SDS-PAGE gels for electrophoresis and transferred onto nitrocellulose membrane with a transfer cell (Trans-Blot Semidry Transfer Cell; Bio-Rad, Hercules, CA). Proteins were detected with a blot analysis system (ECL; Amersham Pharmacia Biotech, Little Chalfont, UK) with total and phosphorylated MAPK (ERK 1/2) antibodies (Upstate Biotechnology, La Jolla, CA). The corresponding antibodies for Akt were obtained from Cell Signaling Technology (Beverly, MA). 

Growth Assays for FHL124 Cells

The Cell division rate was monitored by [³H]thymidine incorporation. A 1-mL portion of a 1 × 10⁴-cell/mL suspension in 5% FCS-EMEM was added to each well of a 24-well tissue culture plate (BD Labware, Bedford, MA) and cultured for 24 hours. The medium was then removed and replaced with serum-free EMEM and cultured for a further 48 hours. After replenishing this medium, the cells were placed in experimental conditions for a further 24 hours. During the final 4 hours of the culture period the cells were exposed to final concentrations of 1 μCi/mL [³H]thymidine (Amersham International, Amersham, UK), with 1 μM cold thymidine. Each well was then washed twice with 1 mL EMEM to remove residual radioactive [³H]thymidine. One milliliter of 5% trichloroacetic acid (TCA) was added to each well. After 30 minutes at room temperature, the TCA was removed and 1 mL 250 mM sodium hydroxide (NaOH) was added to each well and left overnight at 4°C, before 0.5 mL of this NaOH was sampled. Scintillation fluid (10 mL; HiSafe Supermix; PerkinElmer, Wellesley, MA) was added to each sample and appropriate controls. Measurements were obtained using a scintillation counter (EG&G Wallac, Cambridge, UK). 

Data Analysis

Unless otherwise specified, in all cases n = 4; data are expressed as the mean ± SEM. Statistical analyses of the results were evaluated by ANOVA with the Dunnett test, with the exception of determination of capsular bag cell coverage, for which the t-test was used. P < 0.05 was considered to be statistically significant. 

Results

Figure 1Ashows that thrombin (10 nM) induced a significant increase in cytosolic Ca²⁺ in anterior lens epithelial cells of the intact lens. The response was clearly biphasic, with a characteristic initial increase in [Ca²⁺]_i, followed by a smaller second phase. A second, similar response to 10 nM thrombin was obtainable, but only after the lens had been perifused for at least 45 minutes in control medium alone. Identical responses to thrombin were also observed in the isolated lens epithelium (data not shown). Similarly, in the equatorial region of the intact lens, the Ca²⁺ response was biphasic but with a more prolonged second phase (Fig. 1B)and again approximately 45 minutes had to elapse before a second response of similar magnitude was obtained. It is interesting that thrombin produced large Ca²⁺ responses in both anterior and equatorial cells, because acetylcholine (ACh), for example, produces a response only in anterior epithelial cells. ²⁴ It appears that thrombin signals through the ER store, as the specific ER Ca²⁺-ATPase inhibitor thapsigargin totally abolished the response to thrombin (Fig. 1C , Tg). Application of 10 μM lanthanum chloride ³³ (LaCl₃), a Ca²⁺-channel blocker, returned [Ca²⁺]_i to basal levels, indicating that the sustained thapsigargin response was due to Ca²⁺ influx through store-operated channels and was not simply a toxic effect. To identify further whether the second phase produced by thrombin was the result of Ca²⁺ influx, we applied thrombin (10 nM) to isolated lens epithelia in the presence of Ca²⁺-free medium (Fig. 1D) . Thrombin produced a transient increase in [Ca²⁺]_i, which rapidly returned to unstimulated levels, and there was no evidence of a second phase in the response (cf. Figs. 1A 1B ). Return to control medium containing 1 mM CaCl₂ produced a significant increase in internal Ca²⁺, which returned to basal levels when Ca²⁺-free medium was reapplied (Fig. 1D) . In control medium and in the absence of thrombin, there was no large change in internal Ca²⁺, either on switching from control to Ca²⁺-free medium (Fig. 1D)or on readmission of Ca²⁺ to the medium (our unpublished data, 2003). 

To determine which PAR receptor subtype(s) were responsible for initiating the Ca²⁺-signaling events, the lens was exposed to specific activating peptides (PAR-APs) for each receptor subtype. Activating peptides for PAR2, -3, and -4 failed to induce any change in Ca²⁺ levels either in the anterior or equatorial regions of the lens, whereas PAR1-AP induced a Ca²⁺ response in both regions (Fig. 2) . Note that the PAR1 responses in both regions are larger and have a more rapid time course than the corresponding thrombin-induced responses. FHL124 cells loaded with Fura2 responded to thrombin and PAR1, but not PAR2, -3, and -4, in a manner similar to that described for native cells (our unpublished data, 2003). 

Native lens epithelia possessed mRNA for PAR1 and -3 as assessed by bands at the appropriate molecular size (Fig. 3A) . However, mRNA for PAR2 and -4 was not detected. This expression profile was conserved in capsular bags removed from donor eyes that had undergone cataract surgery (Fig. 3B)and the human lens cell line FHL124 (Fig. 3C) . 

Cell coverage on the central posterior capsule was observed in both EMEM controls and in capsular bags cultured in medium supplemented with thrombin (10 nM). In both cases the cells continued to grow across the previously cell-free posterior capsule in a progressive manner, but the rate of coverage in thrombin-treated preparations was significantly faster than the rate in the control serum-free EMEM (Fig. 4) . 

Native donor eye tissue is in limited supply, and therefore FHL124 cells were used for more detailed studies. The presence of mRNA for PAR1 and -3 indicated that the FHL124 cell line is a good model for such studies. Addition of thrombin (10 nM) and PAR1-AP (10 μM) induced an increase in phosphorylation of p42/p44 ERK (p-ERK) proteins in FHL124 cells. Activating peptides for PAR2, -3, and -4 failed to stimulate p-ERK (Figs. 5A 5B) . Significantly, both thrombin and PAR1-AP induced multiphasic responses in p-ERK. p-ERK levels began to increase significantly within 5 minutes, reached a plateau at 10 to 30 minutes, then returned to near baseline levels by 60 minutes (Figs. 5B 5C) . When the exposure time was increased, both thrombin and PAR1-AP induced further significant increases in p-ERK after 8 hours (Fig. 5C) . 

Because both agonists ultimately induced significant increases in p-ERK, we expected both to stimulate growth of FHL124 cells in a similar manner. However, when [³H]thymidine incorporation into DNA was assayed (Fig. 6) , incorporation was significantly increased only in cells cultured in EMEM supplemented with thrombin (10 nM). Supplementing the medium with PAR1-AP failed to induce a statistically significant increase in incorporation under these conditions. Furthermore, cells cultured in the presence of activating peptides for PAR2, -3, and -4 also showed no significant increase over the control (James C, Duncan G, unpublished data, 2004). 

There is some evidence from other cell systems to indicate that both p-ERK and p-Akt stimulation are required for thrombin-stimulated proliferation. ^{34 35} The data in Figure 7show that thrombin induced a 20-fold increase in p-Akt relative to control at 12 hours, whereas the increase produced by exposure to PAR1-AP, though still significant, was much lower. 

Discussion

Thrombin receptors have been identified in a variety of cell and tissue types. However, the expression pattern in the lens is unknown. The present study was therefore undertaken and showed that message for PAR1 and -3, but not for PAR2 and -4, was expressed in isolated lens epithelia, capsular bags removed from donors who had undergone cataract surgery and the human lens cell line FHL124. The expression profiles in other cell types reveal that several receptor subtype combinations exist. PAR1 and -4 are expressed in human astrocytoma cells, ³⁶ for example, whereas PAR1 and -2 are found in stromal fibroblasts. ³⁷ An important finding in this study is that the native lens epithelium and the human lens cell line FHL124 exhibit the same receptor pattern, as some other G-protein receptors (e.g., muscarinic) have been found to change in culture from their native lens counterparts. ²⁷ Analysis of signaling pathways showed that thrombin actively stimulated Ca²⁺ release and phosphorylation of ERK and Akt. Furthermore, application of specific peptide agonists to the PARs demonstrated active signal transduction only with PAR1-AP. These data, therefore, suggest that PAR1 is the only thrombin receptor subtype coupled to calcium signaling in human lens cells. The expression of PAR3 is unusual without joint expression of PAR4 as, at present, the only known role of PAR3 is as a cofactor for PAR4 activation. ³⁸ The role for PAR3, however, remains as poorly defined in the lens as it is in other tissues. ³⁹ There is a possibility that PAR3 mRNA is not translated into protein or, alternatively, that it mediates its effects via other signaling mechanisms not investigated in the present study. In contrast, PAR1 is clearly present and is activated by thrombin. 

In the anterior region of the lens, thrombin induced an initial increase in [Ca²⁺]_i, followed by a short second phase, whereas in equatorial cells the initial increase was followed by a more prolonged second phase. These biphasic profiles are characteristic of Ca²⁺ signaling in several cell types. ⁴⁰ The prolonged raised level elicited from equatorial cells was similar to the response obtained from growth-factor tyrosine kinase receptor–mediated Ca²⁺ influx. ^{24 41} In contrast, PAR1-AP elicited a larger but more transient increase in Ca²⁺, and these kinetics are more characteristic of other G-protein-coupled receptor agonists (e.g., ACh and adenosine triphosphate [ATP]). The second phase of the thrombin response was identified as an influx from the extracellular medium, as it was abolished by removing external Ca²⁺. Covic et al. ⁴² proposed that in platelets, the majority of the overall Ca²⁺ response to thrombin was due to influx through plasma membrane Ca²⁺ channels, which may also be true for cells of the equatorial region. 

In the porcine lens, thrombin has been shown to stimulate the release of endothelin 1 (ET-1) ⁴³ which in turn inhibits Na⁺,K⁺,ATPase activity through the release of Ca²⁺ from internal stores. ⁴⁴ However, ET-1 (100 nM) and ET-2 (10 μM) were applied to the whole human lens, and no change in [Ca²⁺]_i was detected (our unpublished data, 2003). Differences in cell type, species, or thrombin receptor subtype expression may account for these conflicting results. 

Thrombin receptor activation has been implicated in growth promotion (for a review, see MacFarlane et al. ³ ), and has been largely associated with inflammation, ¹¹ the repair of injured tissues, ⁴⁵ and neovascularization. ⁴⁶ Growth results obtained during the present study both with capsular bag cultures and FHL124 cells are in agreement with previous studies that have reported the induction of mitosis and proliferation by thrombin. ^{4 15 19} Both thrombin and PAR1-AP stimulated p-ERK production in FHL124 cells, results similar to those obtained by Molloy et al., ⁴⁷ who found that in normally quiescent rat aortic smooth muscle cells, both thrombin and PAR1-AP induce pronounced increases in tyrosine phosphorylation of MAPK proteins. However, PAR1-AP did not appear to induce an increase in thymidine incorporation in lens cells, although an increase has been observed in rat smooth muscle cells. It is interesting that the kinetics of calcium increase induced by thrombin in human lens cells are different from those induced by PAR1-AP. The influx phase of the Ca²⁺ response is more pronounced in the presence of thrombin compared with PAR1, and this is especially significant in view of the fact that it has recently been shown that capacitative Ca²⁺ entry (CCE) mediates the mitogenic response to EGF. ⁴⁸ It is interesting that, in the present study, the major downstream signaling differences between PAR1-AP and thrombin do not lie at the level of p-ERK stimulation, but rather at the level of p-Akt induction, which occurs at a later stage. 

Akt signaling has recently been a focus of attention in lens research, largely in terms of stimulating cell survival rather than cell growth. ^{49 50} However, there is evidence from other cell systems that sustained Akt phosphorylation is required for G1 phase progression when cell division is activated by thrombin. ¹⁴ It appears in lens cells that although PAR1-AP did indeed stimulate p-ERK and p-Akt, the level attained in the latter was insufficient to drive detectable growth. There are other systems in which the effects produced on exposure to thrombin and PAR1-AP are not identical. For example, in human brain microvascular endothelial cells, whereas both stimulated calcium release, only thrombin decreased transendothelial resistance. ⁵¹ The cleaved peptide from PAR1 is a more potent stimulant of platelet-endothelial adhesion than is thrombin. ⁵² Hence, although the activating peptides are useful in elucidating which thrombin receptor subtypes are functionally active, there is now a considerable body of data (the present study included) to indicate that their application has only a limited use in unraveling the intricacies of the downstream signaling events. 

In summary, the current study shows that lens epithelial cells express mRNAs for PAR1 and -3, and these are conserved in FHL124 cells and in the capsular bag in vivo. Of the two receptors expressed, only PAR1 appears to be functionally active in terms of Ca²⁺ signaling and downstream phosphorylation events. These data raise the possibility that thrombin has a role in wound healing after cataract surgery, and disruption of this signaling system may provide a means of reducing PCO. 

 Supported by the Biotechnology and Biological Sciences Research Council (United Kingdom), Fight for Sight, and the Humane Research Trust.

 Submitted for publication May 11, 2004; revised July 29, November 8, and November 22, 2004; accepted November 23, 2004.

 Disclosure: C. James, None; D.J. Collison, None; G. Duncan, None

 The publication costs of this article were defrayed in part by page charge payment. This article must therefore be marked “advertisement” in accordance with 18 U.S.C. §1734 solely to indicate this fact.

 

Table 1.

 

Gene-Specific Primer Sequences Used in RT-PCR Amplification

Gene		Primer Sequence* (5′–3′)	PCR Product Size (bp)
PAR1	Sense	2422TGTGAACTGATCATGTTTATG2442	708
	Antisense	3129TTCGTAAGATAAGAGATATGT3109
PAR2	Sense	44GCAGGTGAGAGGCTGACTTT63	334
	Antisense	377CAGTCGGTTCCGTCTAACCGG356
PAR3	Sense	147GAAAGCCCTCATCTTTGCAG166	599
	Antisense	745AGGTGAAAGGATGGACGATG726
PAR4	Sense	1121GGCAACCTCTATGGTGCCTA1140	244
	Antisense	1364TTCGACCCAGTACAGCCTTC1345
GAPDH	Sense	ACCACAGTCCATGCCATCAC
	Antisense	TCCACCACCCTGTTGCTGTA

*  Numbers refer to base positions on the known human gene sequence. ³¹

Figure 1.

 

Increase in [Ca²⁺]_i in human lens cells on exposure to thrombin. (A) Anterior epithelial cells in the intact human lens: A 2-minute exposure to 10 nM thrombin induced an increase in fluorescence ratio, indicating a transient increase in [Ca²⁺]_i. The larger initial increase was followed by a smaller second phase. Subsequent exposure to thrombin elicited a similar Ca²⁺ response only after 45 minutes of perifusion with bathing medium alone, as indicated by the space between responses. (B) Equatorial cells in the intact human lens: The responses and the recovery time between them were similar to responses in (A), except that the second phase was more pronounced. (C) Anterior epithelial cells in the intact lens: exposure to Ca²⁺-ATPase inhibitor thapsigargin (1 μM; Tg) produced a characteristic increase in internal Ca²⁺ and abolished any subsequent response to thrombin. LaCl₃ restored cytosolic Ca²⁺ to the resting level by abolishing the capacitative entry pathway. ³³ (D) Anterior epithelial cells in the isolated epithelium: Thrombin induced a transient increase in [Ca²⁺]_i when the lens epithelium was bathed in Ca²⁺-free medium. When Ca²⁺ was reintroduced in control medium, a pronounced influx was observed that was reduced when the medium was returned to Ca²⁺-free. Note that the data are presented in ratio form, as it was not possible to obtain fully calibrated values for the equatorial cells. ²⁴ For the anterior cell response illustrated in (A) a full calibration was performed, and the internal Ca²⁺ increased from a starting level of 100 nM to a maximum of 255 nM.

Figure 2.

 

Effect of PAR-APs on the calcium response of the intact human lens. (A) In anterior epithelial cells, PAR1-AP (10 μM) induced a repeatable, transient increase in Ca²⁺, whereas PAR2-, PAR3-, and PAR4-AP (10 μM) failed to evoke a response. (B) Similar responses were produced in cells in the equatorial region. Note that ACh and ATP were used to give standard G-protein responses in the anterior and equatorial cells, respectively. ²⁴

Figure 3.

 

RT-PCR data for PAR1 (708 bp), PAR2 (334 bp), PAR3 (599 bp), and PAR4 (244 bp). (A) Isolated lens epithelium; (B) ex vivo capsular bag; and (C) FHL124 cells. For PAR1 and -3, but not for PAR2 and -4, products of the appropriate size were detected in all samples. M, DNA marker; G, GAPDH.

Figure 4.

 

Effect of thrombin on cell coverage of the posterior capsule in cultured lens capsular bags. Repeated exposure to thrombin (10 nM in serum-free EMEM, replaced every 48 hours for 12 days) significantly increased cell coverage compared with that of capsular bags cultured in serum-free EMEM alone (control). Note that 100% represents total cell coverage. The data were derived from four match-paired capsular bags, and each point represents the mean ± SEM.

Figure 5.

 

Immunoblot analysis of p-ERK response to thrombin and PAR-APs in FHL124 cells. (A) Representative Western blots for p42/p44 total and phosphorylated ERK protein after exposure to thrombin (10 nM) and PAR1 to -4-AP (10 μM) for 20 minutes. (B) Time course of the early phase of upregulation of p-ERK proteins in response to thrombin and PAR1 to -4-AP. Only thrombin and PAR1-AP gave responses that were significantly different from the control (**P < 0.05; ANOVA with the Dunnett test; n = 4). (C) Late phase of p-ERK stimulation in response to thrombin and PAR1-AP exposure. The early phase of the response declined to a minimum within 2 hours and then increased again to give a more sustained stimulation. **Significant difference from the control, as in (B).

Figure 6.

 

Effect of thrombin and PAR1-AP on thymidine incorporation in FHL124 cells. Cells cultured for 24 hours in EMEM supplemented with thrombin (10 nM) had a significantly increased level of [³H]thymidine (33%, P < 0.05) compared with cells cultured in EMEM alone, whereas the apparent slight increase observed with PAR1-AP (10 μM) supplementation (<10%) was not statistically significant. Data were analyzed by ANOVA with the Dunnett test; n = 4.

Figure 7.

 

Immunoblot analysis of the time-course of the p-Akt response to thrombin (10 nM) and PAR1-AP (10 μM) in FHL124 cells. The very large increase induced by thrombin was apparent after 12 hours of culture and was present for at least another 12 hours. **Significant difference from the corresponding control level (P < 0.05; ANOVA with the Dunnett test, n = 4).

The authors thank John Reddan (Eye Research Institute, Oakland University, Rochester, MI) for providing the FHL124 cells, and I. Michael Wormstone, Lixin Wang, Julie Eldred, and Diane Alden for help and technical assistance.