From the Department of Microbiology and Molecular Genetics, University of California, Irvine, California 92717-4025
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ABSTRACT |
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This study examined the signal transduction pathways involved in thrombin-induced neuroprotection and compares these results with those of a similar study of thrombin-induced neuronal death. In thrombin-induced protection of astrocytes from hypoglycemia, pretreatment of astrocytes with tyrosine or serine/threonine kinase inhibitors, cytochalasin D, or exoenzyme C3, a potent inhibitor of the small GTPase RhoA, attenuated thrombin-induced protection. These same inhibitors were previously shown to block thrombin-induced cell death, implying a similarity in the cell death and cell-protective pathways. Biochemical assays determined that thrombin increased available RhoA activity, although more slowly and to a lesser extent than occurs in thrombin-induced cell death. A clear difference in these pathways was revealed when a time course study of thrombin-induced cell death indicated that unlike thrombin-induced protection, cells must be exposed to thrombin for >16 h to irreversibly enter the cell death pathway. Addition of lower doses of thrombin every 24 h also induced cell death. These studies indicate that exposure of cells to micromolar concentrations of thrombin alone does not induce cell death, but the continued exposure to thrombin is required. Thus the cell death and protective pathways may share initial signaling proteins, but differences in the amplitude as well as the duration of the signal may result in different final pathways.
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INTRODUCTION |
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Thrombin is a multifunctional serine protease immediately produced
at sites of vascular injury. Although well known for its role in
hemostasis and thrombosis, thrombin also induces a wide range of
cellular inflammatory and proliferative responses associated with both
normal and disease processes (1, 2). Most recently, thrombin was shown
to induce two responses that indicate an important role for thrombin in
central nervous system disease and injury. Moderate concentrations of
thrombin protected hippocampal neurons and astrocytes from a variety of
cellular insults, including hypoglycemia, growth supplement
deprivation, oxidative stress, and -amyloid toxicity (3, 4), while
high concentrations of thrombin induced apoptosis in these same cells
(3, 5, 6). Both responses were shown to be due to activation of the
PAR-1 receptor (thrombin receptor) (3, 6), but many questions remain
concerning the mechanisms underlying these responses. How does thrombin
stimulate cells to survive toxic insults? What are the pathways
underlying this response, and as these pathways appear to be similar to
pathways recently reported to be involved in the apoptosis response,
how do cells distinguish between these signals?
The central nervous system is exposed to thrombin upon breakdown of the blood-brain barrier. This occurs in acute trauma such as head injury or stroke and may also occur in chronic neurodegenerative diseases such as Alzheimer's disease and cerebral-vascular dementia (7-10). Studies on the effects of thrombin on neuronal cells in culture have revealed a variety of responses to thrombin. Thrombin induces process retraction in neuroblastoma cells (11, 12), and human fetal neurons (13) and reverses stellate, process bearing astrocytes to a more flattened morphology (14, 15). Thrombin is mitogenic for astrocytes (14, 16, 17) and induces the synthesis and secretion of both nerve growth factor (18) and endothelin-1 (19). In one study, thrombin was directly infused into the rat caudate nucleus. This resulted in increased reactive gliosis, infiltration of inflammatory cells, proliferation of mesenchymal cells, and induction of angiogenesis, effects that resemble inflammation, reactive gliosis, and scar formation that occurs following injury to the central nervous system (20). These studies and the reports of thrombin-induced protection and apoptosis indicate the possible impact of thrombin exposure to central nervous system cells. Apoptosis is a common feature of both acute central nervous system injury and chronic neurodegenerative disorders such as Alzheimer's disease (21-23). It is of general interest to understand any mechanism that allows neuronal cells to survive toxic insults; thus a more clear understanding of how thrombin induces the protective and apoptotic responses is warranted.
Cellular responses induced by thrombin are due, in most cases, to activation of the PAR-1 receptor (24, 25). The PAR-1 receptor is a proteolytically activated, seven-transmembrane-spanning receptor that has been linked to a variety of cellular pathways, including hydrolysis of phosphoinositides, calcium mobilization, and activation of heterotrimeric G-proteins, tyrosine kinases, and monomeric G-proteins (for review, see Ref. 26). The PAR-1 receptor mRNA is also widely expressed throughout the central nervous system (27-29). In this study we questioned how thrombin induces both apoptosis and protection in astrocytes. One possibility is that different signal transduction pathways are involved in each response. Another possibility is that the pathways are similar, but that the different concentrations of thrombin required to induce each response result in a different level of stimulation of this pathway, so that the correct signal is ultimately generated. A third possibility is that the different culture conditions of the cells may also account for the different responses. Astrocytes experiencing hypoglycemia or another stress may express a different background of downstream effector proteins so that the connection to the protective pathway is made, rather the apoptosis pathway. We employed pharmacological and biochemical methods to help determine second messenger pathways involved in thrombin-induced protection of astrocytes from hypoglycemia and contrasted these results with a similar study of thrombin-induced apoptosis.
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EXPERIMENTAL PROCEDURES |
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Materials--
Highly purified, high specific activity, human
-thrombin was purchased from Calbiochem and from Sigma. Because of
variation in specific activity between thrombin supplies, all thrombin
treatments were carried out using units/ml with 200 units/ml roughly
equivalent to 1 µM thrombin. Herbimycin A, cytochalasin
D, genistein, pertussis toxin, HA 1004, H-7, wortmannin, and exoenzyme
C3 were purchased from Calbiochem. Leupeptin, aprotinin,
phenylmethylsulfonyl fluoride, and soybean trypsin inhibitor were
purchased from Sigma. Monoclonal antibody to glial fibrillary acidic
protein was purchased from Boehringer Mannheim.
Isolation of Primary Cultures of Rat Astrocytes-- Primary cultures of type 1 astrocytes were prepared from the brains of 1- to 2-day-old rat pups (Sprague-Dawley) using a modification of previously described procedures (40). Briefly, the frontoparietal cortex from eight pups was isolated, stripped of meninges, and dissociated by a combination of trypsin digestion and mechanical trituration. A single cell suspension was prepared from this dissociated tissue by passage though Nitex nylon screens. The cells were plated into 75-cm2 plastic flasks and grown in Dulbecco's modified Eagle's medium containing 25 mM glucose, 7.5 mM sodium bicarbonate, 20 mM HEPES, 2 mM glutamine, 1 mM sodium pyruvate, 100 units/ml penicillin, 100 mg/ml streptomycin sulfate, and 10% fetal bovine serum. Once the cultures reached confluence (approximately 10-12 days), the flasks were shaken at 260 rpm for 24 h at 37 °C to remove nonadherent cells. The remaining type 1 astrocytes were trypsinized and reseeded into 75-cm2 flasks. Once these cells had reached confluence they were replated for experiments as detailed below. The purity of the cultures was confirmed by immunofluorescent staining with a monoclonal antibody to the type 1 astroglial specific marker glial fibrillary acidic protein; >95% of the cells were immunoreactive for this marker.
Experimental Treatment of Astrocytes-- Astrocytes were removed from flasks by trypsinization and plated at a density of 1 × 104 cells/cm2. After growth to 70-80% confluence (2-3 days after plating), the cells were used in experiments. In all experiments cells were rinsed three times with serum-free Dulbecco's modified Eagle's medium and then incubated in this medium for 16-18 h prior to experimental treatments. Cells were then differentiated for 24 h with dibutyryl cyclic AMP. Dibutyryl cyclic AMP was present in all media throughout the experiments. For experiments measuring the effect of pharmacological agents on thrombin-induced protection, astrocytes were serum-starved, differentiated with dibutyryl cyclic AMP (1.5 mM) for 24 h, and then pretreated for 3 h with each agent. For experiments involving pertussis toxin, cells were pretreated for 24 h. Cells were then washed three times and incubated in medium containing 0.5 mM glucose, and each agent in the presence or absence of 0.2 unit/ml (1 nM) thrombin. Cell viability was determined 72 h later. In all experiments, cell viability was calculated relative to its control wells, which were maintained in either low glucose medium alone or low glucose medium containing the pharmacological agent throughout the experiment. In experiments examining the time course of thrombin-induced cell death, cells were serum-starved overnight, washed, then incubated in serum-free medium plus bovine serum albumin (0.1%), in the presence or absence of 200 units/ml thrombin. After the indicated duration of exposure, both control and thrombin-treated wells were washed three times with serum-free medium, then incubated in this medium for the remainder of the experiment. In experiments examining whether lower doses of thrombin could induce cell death, cells were serum-starved overnight, washed, then incubated in serum-free medium plus bovine serum albumin (0.1%), in the presence or absence of the indicated concentrations of thrombin. Thrombin was then added directly to this medium 24 h from the start of the experiment. Cell viability was measured 72 h following initial exposure to thrombin. In all experiments, cell viability was determined by assaying the medium from each well for lactate dehydrogenase (LDH)1 activity using a diagnostic kit according to the manufacturer's instructions (Sigma). Released LDH is a stable enzymatic marker that correlates linearly with cell death. To determine total LDH activity, cells from control cultures were lysed in 0.5% Triton X-100, centrifuged at 16,000 × g for 1 min, and the supernatants were assayed for LDH activity. This cell-associated LDH activity was then added to the LDH activity released from control cultures, and the total activity was considered to represent 100% cell death. The amount of LDH present in the medium was then calculated as a percentage of the total, which determines the percent cell death in that sample. For clarity, results were then presented as the percent cell viability. In some experiments, SYTO 11 (Molecular Probes) was added to the control and thrombin-treated cultures to monitor viability microscopically. All results are expressed as the mean ± S.E. of triplicate samples. Data were statistically examined by one-way analysis of variance followed by pairwise comparisons using the Fisher procedure. All studies were repeated in at least three independent experiments.
Cell Fractionation and [32P]ADP Ribosylation-- Cell fractionation and in vitro ribosylation were performed as described previously (41). Briefly, cells were grown in 10-cm culture dishes for 3-4 days, serum-starved, and differentiated as described above, then exposed to hypoglycemic conditions in the presence or absence of the indicated concentrations of thrombin for 0-48 h. Cells were washed twice with phosphate-buffered saline, scraped into ice-cold 20 mM Tris/HCl (pH 8.0) in the presence of a protease inhibitor mixture (0.4 mM phenylmethylsulfonyl fluoride, 20 µM leupeptin, 0.05 unit/ml aprotinin, 20 µg/ml soybean trypsin inhibitor), and homogenized using a tight pestle Dounce homogenizer (30 strokes). Lysates were centrifuged at 500 × g for 10 min at 4 °C. The supernatants were centrifuged at 25,000 × g for 30 min to separate the cytosolic from the crude membrane fraction. Protein concentration was determined using the Bio-Rad Bradford protein assay. Ribosylation assays were carried out using 30 µg of protein from each fraction that was heat-inactivated at 65 °C for 5 min and then taken up in reaction buffer (90 mM Tris/HCl (pH 8.0), 2.6 mM MgCl2, 1 mM EDTA, 10 mM thymidine, 10 mM dithiothreitol, 1 mM ATP, 100 µM GTP, 10 µCi/ml [32P]NAD). Reaction was started by addition of exoenzyme C3 (5 µg/ml) and allowed to proceed at 37 °C for 1 h. Proteins were acid-precipitated (10% trichloroacetic acid, w/v), centrifuged (15,000 × g for 15 min), and washed with ether. Samples were then subject to SDS-polyacrylamide electrophoresis (12.5%) and ribosylated proteins visualized by autoradiography.
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RESULTS |
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Pharmacological Characterization of the Signal Transduction Cascade Underlying Thrombin-induced Neuroprotection: Involvement of Tyrosine Kinases, Serine/Threonine Kinases, and the Actin Cytoskeleton-- PAR-1 receptor activation can result in induction of a variety of different second messengers, including hydrolysis of phosphoinositides, calcium mobilization, and activation of heterotrimeric G-proteins, tyrosine kinases, and monomeric G-proteins (26). To identify second messenger pathways involved in thrombin-induced neuroprotection, we tested several pharmacological agents for their ability to block thrombin-induced protection of astrocytes from hypoglycemia. Astrocytes were serum-starved, differentiated with dibutyryl cyclic AMP (1.5 mM) for 24 h, and then pretreated for 3 h with each agent, except pertussis toxin for which the cells were pretreated for 24 h. Cells were washed three times and incubated in medium containing 0.5 mM glucose, dibutyryl cyclic AMP, and each agent in the presence or absence of 0.2 unit/ml (1 nM) thrombin. Cell viability was measured using the LDH assay 72 h following exposure to hypoglycemic conditions. Consistent with the previous study (3), untreated hypoglycemic cultures showed 31% viability, whereas hypoglycemic cultures treated with thrombin were protected and showed greater than 75% viability (Fig. 1A). Two broad spectrum serine/threonine kinase inhibitors, HA 1004 and H7, were used to determine the involvement of serine/threonine kinases. H7, at 10 and 100 µM, blocked thrombin-induced protection. HA 1004, at concentrations up to 100 µM, failed to inhibit thrombin-induced protection. The primary difference in the in vitro inhibition profile of these two inhibitors is that H7 is a more potent inhibitor of protein kinase C than is HA 1004, suggesting a possible role for protein kinase C in thrombin-induced protection from hypoglycemia. The tyrosine kinase inhibitor, herbimycin A, also blocked thrombin-induced protection with both thrombin-treated and -untreated cultures retaining only 38 and 47% viability, respectively. Another broad spectrum tyrosine kinase inhibitor, genistein, did not block thrombin-induced protection; however, genistein treatment alone attenuated hypoglycemic death, making the effect on thrombin signaling difficult to determine (Fig. 1A).
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The Available RhoA Activity Is Increased by Thrombin during Thrombin Protection from Hypoglycemia-- The small GTP-binding protein RhoA has previously been linked to thrombin-induced cellular responses, including neurite retraction in NEI-115 neuroblastoma cells (41) and, most recently, thrombin-induced apoptosis in astrocytes and hippocampal neurons (6). RhoA has been shown to regulate the actin cytoskeleton and is thought to contribute to mitogen-associated protein kinase cascades (47-49). To investigate the possibility of RhoA involvement in thrombin-induced protection, we assayed for the level of available RhoA, RhoA that is not inactivated by the cellular ADP-ribosyltransferase enzyme, under hypoglycemic conditions in the presence or absence of thrombin. Cells were cultured and differentiated as described, washed twice, and incubated in medium containing 0.5 mM glucose and 0 or 20 units/ml (10 nM) thrombin. At 3, 12, and 24 h post-treatment, cells were lysed and membrane and cytosolic fractions were prepared. These fractions were assayed for protein concentration and 30 µg of each sample were tested for the level of available, nonribosylated RhoA by in vitro ribosylation assay (41). Proteins were ADP-ribosylated by exoenzyme C3 and [32P]NAD, precipitated, and electrophoresed though 12.5% SDS-polyacrylamide gels. Gels were dried and ribosylated proteins visualized by autoradiography. Thrombin increased available RhoA activity in the membrane fraction, but only after at least a 3-h incubation with thrombin (Fig. 2A). This thrombin-induced increase in available RhoA continued and increased over the time course of the experiment (Fig. 2, B and C). As a control for endogenous ribosylation activity, 30 µg of a thrombin-treated sample was assayed without the addition of exoenzyme C3; no RhoA is labeled (lane 1, all panels). This result indicates that although nanomolar concentrations of thrombin do not immediately increase available RhoA, prolonged exposure to thrombin, even under hypoglycemic conditions, increases available RhoA activity. This raises the possibility that RhoA is involved in transducing thrombin protective signals.
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Exoenzyme C3, a Selective and Potent Inhibitor of RhoA, Blocks Thrombin-mediated Protection from Hypoglycemia-- Since prolonged treatment with thrombin showed a small but consistent increase in available RhoA activity, we questioned whether the activity of RhoA was necessary for thrombin-mediated protection from hypoglycemia. Cells were cultured as described and pretreated with exoenzyme C3 (25 µg/ml) for 3 h. Cells were then incubated in medium containing 0.5 mM glucose and exoenzyme C3 in the presence or absence of 0.2 unit/ml (1 nM) thrombin. Cell viability was measured using the LDH assay 72 h following exposure to hypoglycemic conditions. Pretreatment with exoenzyme C3 completely blocked thrombin-induced protection (Fig. 3). Exoenzyme C3 did not affect cell viability as compared with control untreated hypoglycemic cultures, indicating it was not increasing the toxicity of the hypoglycemic treatment. Thus, RhoA activity appears to be necessary for thrombin-induced protection in astrocytes.
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Time Course Study of Thrombin-induced Cell Death in Astrocytes Reveals That Prolonged Exposure to Thrombin Is Necessary to Signal Apoptosis-- The results of these studies on the signal transduction cascade underlying thrombin-induced protection when compared with the previous study of thrombin-induced apoptosis (6) indicate a similar pathway is stimulated in both cases. How does thrombin cause these very different effects in these cells? One possibility is that although the signal pathway is similar, perhaps identical in the beginning, the different concentrations of thrombin required for each response result in a different level of stimulation of this pathway so that a different signal is ultimately generated. Another possibility is that duration of signal plays a part, that the cells must be continuously exposed to elevated levels of thrombin in order to induce cell death. To test the latter hypothesis, astrocytes were exposed to 200 units/ml thrombin for 8, 12, 16, 20, or 72 h, after which cells were washed three times, and serum-free medium was added back to the culture well. Control cells were maintained in serum-free medium and washed with media exactly as thrombin-treated cells. Cell viability was then measured 72 h from the addition of thrombin. Astrocytes exposed to thrombin for up to 16 h showed no loss in cell viability; however, exposure of cells to thrombin for 20 h or longer resulted in nearly as much cell death as a full 72-h thrombin exposure (Fig. 4A). This demonstrated that duration of exposure to thrombin contributes to the cellular response to thrombin. It is also noteworthy that the control culture not treated with thrombin had a consistent decrease in cell viability (approximately 30%). This is most likely due to the removal of the cells from the incubator every 4 h of the experiment, because control cells not removed from the incubator showed no significant loss in cell viability over a similar 72 h time course. Importantly, thrombin treatment up to the 16-h time point actually protected the cells from the deleterious stresses of this experiment, showing that not only were the high doses of thrombin not toxic, but actually protected the astrocytes (Fig. 4A). These results, in combination with the pharmacological studies, suggest the possibility that thrombin activates a common pathway when inducing either protection or apoptosis and that in the case of apoptosis, the persistence of the signal contributes to the altered response of the cell. In the initial report of thrombin-induced neuroprotection, a time course study determined that no pretreatment of astrocytes with thrombin was necessary to protect these cells from hydrogen peroxide toxicity (3). Thus the protective response appears to be a much more rapid response than thrombin-induced cell death.
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DISCUSSION |
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The recent report that thrombin can protect neuronal cells from toxic insults raises the tantalizing possibility of understanding the mechanism behind this protection. This in turn may provide insight into general mechanisms and signals that can be used to increase neuronal survival following central nervous system injury or decrease losses in chronic neurodegenerative disorders such as Alzheimer's disease. In this study, we presented findings indicating the involvement of tyrosine and serine/threonine kinases and the small GTP-binding protein RhoA in the signal transduction pathways of thrombin-induced protection of astrocytes. We also examined the time course of thrombin-induced apoptosis and re-evaluated the thrombin concentrations needed to induce cell death. These results, taken into context with what is known about PAR-1 receptor activation and other thrombin-induced neuronal responses, help to explain how thrombin induces both apoptosis and protection in the same cell type and shed new light on the possible result of thrombin exposure to the central nervous system.
The results of our investigation into the signal transduction pathways underlying thrombin-induced protection indicate a strong similarity to pathways implicated in thrombin-induced cell death. As was true for the cell death pathway, tyrosine and serine/threonine kinases and an intact actin cytoskeleton appear to be necessary for thrombin-induced protection of astrocytes from hypoglycemia. The serine/threonine kinase H7 completely blocked thrombin-induced protection, while HA-1004, another broad spectrum serine/threonine kinase inhibitor, had no effect. Although both agents have been shown to inhibit many of the same kinases, H7 is a more potent inhibitor of protein kinase C, at least in vitro, thus indicating a possible role for protein kinase C in thrombin-induced protection. The tyrosine kinase inhibitor herbimycin A also significantly blocked thrombin-induced protection, and these data support the many previous reports that PAR-1 receptor activation involves tyrosine kinase activity (30-33). PAR-1 receptor activation has also been linked to phospholipase D, phospholipase A2, and phosphoinositol 3-kinase activity, but the inhibitor wortmannin failed to block the protection, making the involvement of these kinases unlikely. This inhibition profile for thrombin-induced protection is strikingly similar to the profile reported for thrombin-induced apoptosis (6).
The small GTP-binding protein RhoA also appears to be involved in both thrombin-induced protection and apoptosis. Astrocytes treated with 200 units/ml (1 µM) thrombin, concentrations that are known to induce apoptosis in these cells, show a rapid increase in available RhoA activity within minutes of thrombin treatment (6). Under the conditions of thrombin-induced protection in which cells are exposed to hypoglycemic conditions and 2 units/ml (10 nM) thrombin, a small but consistent increase in the available RhoA activity was measurable after 3 h of thrombin treatment (Fig. 2). This demonstrates a clear difference between the protective and apoptotic pathways, with the apoptotic pathway involving a much more rapid and robust change in available RhoA than the protective pathway. Because the difference in available RhoA levels in the protective response is small and requires >1 h to measure, it could be inferred that it is due to an effect of the hypoglycemic treatment slowing protein synthesis in the unprotected cells and not due to a thrombin-stimulated increase in available RhoA in the protected astrocytes. Although we have no direct evidence to contradict this, use of the RhoA inhibitor, exoenzyme C3, indicates otherwise. Pretreatment of astrocytes with exoenzyme C3 did not increase the cell death brought on by hypoglycemia, indicating it was not generally toxic, and completely blocked the protective effect of added thrombin, implying RhoA activity is required for protection. Because RhoA is thought to regulate the actin cytoskeleton, we also questioned whether treatment with exoenzyme C3 blocked thrombin responses in cells simply by disrupting the actin cytoskeleton such that any receptor-mediated signal would be blocked. Using biotin-conjugated phaloidin toxin, we stained the actin microfilaments of astrocytes prepared as in previous protection experiments. Astrocytes were treated with dibutyryl cyclic AMP alone or followed with a 3-h exoenzyme C3 or cytochalasin D treatment. Our studies indicated that although C3 treatment caused a relaxation of actin fibers compared with dibutyryl cAMP treatment alone, it did not cause a mass disappearance of actin filaments and, unlike cytochalasin D treatment, did not cause a clustering of actin monomers and loss of actin microfilament assembly.2 We concluded that the increase in available RhoA and the inhibition of protection by C3 treatment were due to involvement of RhoA in a specific PAR-1 receptor-mediated pathway and not due to a nonspecific breakdown of the actin cytoskeleton.
Although these results indicate that the same or similar second messenger pathways and proteins are involved in both thrombin-induced protection and apoptosis, there are differences in the culture conditions and thrombin concentration the cells are exposed to that may account for the different cellular responses. The difference in culture conditions, such as stressing the cells with hypoglycemic medium, may cause a shift in the expression of available downstream effector proteins, so that the background of effector proteins is different in a healthy unstressed cell from that of a stressed cell. PAR-1 receptor activation may induce a similar initial signal transduction cascade in each condition, but this cascade may interact with one set of effector proteins in a healthy cell and an entirely different set of effector proteins in a stressed cell. Our results thus far cannot rule out such a possibility.
It is also possible that the same cascade is stimulated in each response, but that the different concentrations of thrombin result in different levels of activation of this pathway, thus generating a different response in the cell. Thrombin, being an enzyme, does not activate its receptor in the way a classical ligand does (24, 35). Thrombin activates PAR-1 by cleaving the amino terminus; this generates a novel terminus that acts as a tethered ligand, binding and activating the cell. Therefore, unlike a classical ligand-receptor interaction, cells cannot use graded receptor occupancy to achieve graded and concentration-dependent responses to thrombin. Coughlin and associates addressed this issue in the following manner. Using expression of epitope-tagged PAR-1 receptors in Rat 1 cells, Ishii et al. (35) were able to measure PAR-1 receptor cleavage and correlate it to second messenger production by measuring phosphoinositide hydrolysis. They were able to demonstrate that for every PAR-1 receptor activated, a quantum of second messenger is produced, which in turn is cleared within a given time period. The authors concluded that cells give graded responses to thrombin by measuring the balance between rate of receptor activation and clearance of second messengers produced (35).
The thrombin-induced protective and cell death responses are highly concentration-dependent. Apoptosis requires at least 100 units/ml (500 nM) thrombin when added as a single addition, whereas the protective response is induced over a concentration range of 0.0002 to 20 units/ml (1 pM to 100 nM). The difference in concentration needed to induce each response may be due to a need to activate the PAR-1 receptor at different rates to achieve each signal. Studies using the PAR-1-activating peptide, SFLLRN, support this. Thrombin-induced protection was induced by 100 nM to 10 µM SFLLRN (3), whereas only very high concentrations of SFLLRN (15 mM) were able to induce the apoptosis pathway (6). The results of the pharmacological studies indicate that many of the same type of second messengers are involved in both pathways, suggesting it is the same pathway, but these studies cannot tell us the degree to which any actual second messenger is stimulated during each response. The biochemical assay measuring changes in available RhoA activity indicates that RhoA is involved in both responses, with a small increase seen in the protective response (Fig. 2), but a larger increase in the cell death response (6). This result suggests that the pathway is activated to a greater degree in the cell death response. It is possible that the same pathway is induced in both responses, but that the cell death response may be a result of overactivation of PAR-1 and overactivation of this pathway, driving the cells into apoptosis.
Studies conducted in other cell types suggest that there would be little difference in the rate of receptor activation at the concentrations of thrombin used for the protective or the cell death response. Both concentrations are fairly high and probably result in very rapid receptor cleavage, thus our conclusion that the higher concentration needed to induce cell death is at least partially due to a difference in rate of receptor cleavage may not be true. We suggest, however, that it may be erroneous to assume that concentrations of thrombin that resulted in a particular rate of receptor cleavage in one cell type is going to be the same in another cell type. Astrocytes secrete a variety of potent thrombin inhibitors, most notably Protease Nexin-1, and also produce a variety of cell surface and extracellular matrix glycoproteins to which thrombin may bind and be sequestered from interacting with the thrombin receptor. Thus drawing conclusions as to how much thrombin truly cleaved how many PAR-1 receptors and at what rate is impossible to tell without directly measuring the rate of receptor cleavage. Unfortunately, the antibodies for this experiment are currently unavailable.
The results of our study of the duration of exposure to high concentrations of thrombin necessary to induce apoptosis and the ability of lower concentrations of thrombin to induce apoptosis, if added repeatedly, indicate that not only does the apoptotic pathway require a threshold level of thrombin, which translates into a rate of receptor activation and second messenger production, but that this rate of receptor activation must continue for a given period of time before the apoptotic pathway is irrevocably activated. This constitutes a requirement for duration of receptor activation to get a particular cellular response as well as a required thrombin concentration. This may also explain why such high concentrations of thrombin or SFLLRN are required to induce cell death in culture. Both the enzyme and peptide are added as a single bolus in these experiments and are subsequently degraded over the time course of the experiment. A high initial concentration may be necessary to obtain duration of signal.
Several studies report that the PAR-1 receptor undergoes homologous desensitization and internalization after activation (36-39). Replacement of receptors occurs within hours in some cells, but no trafficking of the PAR-1 receptor or desensitization studies have been performed in astrocytes as yet. Astrocytes treated to such high concentrations of thrombin as 200 units/ml or even 40 units/ml probably undergo desensitization and internalization. The experiments illustrating that only a continued presence of thrombin will induce cell death in astrocytes suggests that these cells recover sensitivity to thrombin, most likely though replacement of PAR-1 receptors.
Absolute concentrations of thrombin needed to induce apoptosis, or indeed any cellular response to thrombin, cannot be determined with cell culture assays. Cell culture studies present a variety of artifacts and uncontrollable variables in the study of thrombin-induced cellular responses. Thrombin added to culture medium is subject to degradation and inactivation, uptake by the cells, inhibition by cellular thrombin inhibitors over time, and binding to extracellular matrix proteins, sulfated proteoglycans on the cell surface, as well as binding to and activating the PAR-1 receptor. However, some physiological relevance can be derived from these experiments. These results indicate that relatively high concentrations of thrombin are needed to induce apoptosis, but what may be more important is the continued presence of thrombin that leads to apoptosis, not just a high concentration. It would appear from the results of our time course study that high concentrations of thrombin can stimulate the protective pathway in astrocytes, as long as the thrombin activity is greatly decreased or removed before the critical duration of signal is reached. This suggests that in cases of acute trauma or stroke, there may be a window of time during which central nervous system cells are exposed to a high local concentration of thrombin, but are not yet signaled to undergo apoptosis, that may lead to cell protection. In cases of chronic neurodegenerative disorders such as Alzheimer's disease, compromised blood-brain barrier function may allow focal regions of the brain to be exposed to thrombin, perhaps over periods of years, thus increasing the likelihood of thrombin-induced apoptosis contributing to such disorders.
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FOOTNOTES |
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* This work was supported in part by National Institutes of Health Grant AG00538.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Supported by National Institutes of Health Training Grant
AG00096.
§ To whom correspondence should be addressed. Tel.: 714-824-5267; Fax: 714-824-8598; E-mail: ddcunnin{at}uci.edu.
1 The abbreviation used is: LDH, low density lipoprotein.
2 F. M. Donovan and D. D. Cunningham, unpublished observation.
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REFERENCES |
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