About this Journal Submit a Manuscript Table of Contents
Journal of Biomedicine and Biotechnology
Volume 2012 (2012), Article ID 721657, 13 pages
http://dx.doi.org/10.1155/2012/721657
Review Article

The Plasminogen Activation System and the Regulation of Catecholaminergic Function

1Department of Medicine, University of California San Diego, La Jolla, CA 92093, USA
2Veterans Administration San Diego Healthcare System, San Diego, CA 92161, USA

Received 20 March 2012; Accepted 17 May 2012

Academic Editor: David M. Waisman

Copyright © 2012 Hongdong Bai et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract

The local environment of neurosecretory cells contains the major components of the plasminogen activation system, including the plasminogen activators, tissue plasminogen activator (t-PA) and urokinase-type plasminogen activator (u-PA), as well as binding sites for t-PA, the receptor for u-PA (uPAR), and also the plasminogen activator inhibitor, PAI-1. Furthermore, these cells express specific binding sites for plasminogen, which is available in the circulation and in interstitial fluid. Colocalization of plasminogen and its activators on cell surfaces provides a mechanism for promoting local plasminogen activation. Plasmin is retained on the cell surface where it is protected from its inhibitor, 𝛼 2 -antiplasmin. In neurosecretory cells, localized plasmin activity provides a mechanism for extracellular processing of secreted hormones. Neurotransmitter release from catecholaminergic cells is negatively regulated by cleavage products formed by plasmin-mediated proteolysis. Recently, we have identified a major plasminogen receptor, Plg- R K T . We have found that Plg- R K T is highly expressed in chromaffin cells of the adrenal medulla as well as in other catecholaminergic cells and tissues. Plg- R K T -dependent plasminogen activation plays a key role in regulating catecholaminergic neurosecretory cell function.

1. Introduction

Chromaffin cells of the adrenal medulla and other neurosecretory cells contain specific binding sites for plasminogen [1, 2], which is available at high concentration in the circulation and in interstitial fluid. Furthermore, these cells secrete major components of the plasminogen activation system, including the plasminogen activators tissue plasminogen activator (t-PA) [37] and urokinase-type plasminogen activator (u-PA) [4, 8], and also the plasminogen activator inhibitors, PAI-1 [9] and neuroserpin [10]. Binding sites for t-PA [1, 8] and the receptor for u-PA (uPAR) are also expressed in these cells [8, 1113]. Thus, the major components of the plasminogen activation system are present in the local neurosecretory cell environment.

Colocalization of plasminogen and plasminogen activators on cell surfaces results in promotion of local plasminogen activation [14]. In addition, cell-associated plasmin is protected from its major inhibitor, 𝛼 2 -antiplasmin [15, 16]. In neurosecretory cells, localized plasmin activity provides a mechanism for extracellular processing of secreted hormones [1, 2]. For example, neurotransmitter release from catecholaminergic cells is negatively regulated by cleavage products formed by plasmin-mediated proteolysis of the major secretory vesicle core protein, chromogranin A (CgA) [1, 17]. Recently, we have identified a major plasminogen receptor, Plg-RKT [18]. In this paper we summarize the interplay between components of the local neurosecretory cell plasminogen activation system, the functional consequences of the interaction of plasmin with catecholaminergic cells, and our recent studies demonstrating the expression and function of Plg-RKT in catecholaminergic cells.

2. Components of the Local Chromaffin Cell Plasminogen Activation System: Plasminogen Activators, Plasminogen Activator Inhibitors, and Binding Sites for Plasminogen Activators

2.1. t-PA Expression and Its Targeting to the Regulated Secretory Pathway

The expression of t-PA, its subcellular localization, and its release in response to secretagogues have been investigated in several chromaffin cell sources. t-PA synthesis and expression were demonstrated in rat PC12 cells [47], bovine adrenal chromaffin cells [7], human pheochromocytoma (a catecholamine producing tumor of the adrenal gland) [7], and adrenal medulla [3]. A variety of methods including Northern blotting, Western blotting, metabolic labeling and immunoprecipitation with specific anti-t-PA antibody, immunoassays, and enzymatic activity assays have been used to demonstrate the presence of authentic t-PA in these cells [7].

Within neuroendocrine cells, secretory proteins are sorted into one of two pathways, either the regulated secretory pathway or the constitutive secretory pathway [19, 20]. Proteins that enter the constitutive pathway are not stored but are transported directly to the cell surface and secreted in the absence of an extracellular signal. In contrast, proteins that enter the regulated pathway are concentrated and stored in vesicles and subsequently released upon stimulation with a secretagogue or other specific extracellular stimuli. Catecholamine storage vesicles within the chromaffin cell are prototypic examples of regulated secretory vesicles [2023].

We examined the potential targeting of t-PA to the regulated secretory pathway by evaluating the subcellular localization of chromaffin cell t-PA in functional secretagogue release studies and using subcellular fractionation methods. In functional secretagogue-release studies, in which PC12 cells and bovine adrenal chromaffin cells were stimulated with a panel of secretagogues including nicotine (acting through nicotinic cholinergic receptors), KCl (a membrane depolarizing agent), and BaCl2 (a calcium agonist), each of which causes exocytotic release of catecholamine storage vesicle content, significant increases in t-PA secretion occurred (Figure 1). Moreover, t-PA release occurred in parallel with release of catecholamines, consistent with release from the same subcellular pool, the catecholamine storage vesicle. The release of t-PA from PC12 cells also occurs in response to membrane depolarization with tetraethylammonium chloride and depends on the influx of calcium ion [6]. This rapid release is also consistent with release of presynthesized t-PA from a storage pool because no changes in gene expression or protein synthesis are required [6].

fig1
Figure 1: Corelease of t-PA with norepinephrine from catecholaminergic cells. PC12 cells (a) or bovine adrenal chromaffin cells (b) were labeled with [3H]-norepinephrine and incubated at 37°C for 30 min in either the absence or presence of 60 μM nicotine, 55 mM KCl, or 2 mM BaCl2. The release buffer was aspirated, cells were harvested and lysed, and the release of t-PA antigen (open bars) and [3H]-norepinephrine (filled bars) was determined. [7]. ©The American Society for Biochemistry and Molecular Biology.

Subcellular fractionation studies using sucrose density gradients also demonstrated trafficking of t-PA to catecholamine storage vesicles [7]. Furthermore, catecholamine storage vesicle lysates isolated from human pheochromocytoma tumors were enriched 30-fold in t-PA antigen, compared with tumor homogenate [7]. The enrichment in t-PA antigen paralleled the enrichment in catecholamines, consistent with colocalization in the same subcellular fraction.

The localization of t-PA in secretory vesicles of PC12 cells has provided a key tool to study the mechanisms and kinetics of exocytosis. GFP-tagged t-PA has been used as a marker to understand axonal transport in nerve-growth-factor-(NGF-)treated PC12 cells. (When exposed to NGF, PC12 cells differentiate into cells that morphologically, biochemically, and electrophysiologically closely resemble sympathetic neurons [26].) Scalettar’s group demonstrated that GFP-t-PA was targeted for regulated secretion from growth cones of NGF-differentiated PC12 cells and released in response to the calcium ionophore A23187 or the cholinergic agonist, carbachol [27], and used GFP-t-PA to demonstrate that secretory granules are mobile in growth cones of these cells [28]. In other studies with undifferentiated PC12 cells, fluorophore-tagged t-PA has been used as a marker to demonstrate: that most granules in PC12 cells reseal after exocytosis, resulting in the differential release of cargo [29], that synaptotagmin VII modulates kinetics of dense-core vesicle exocytosis in PC12 cells [30], that actin rearrangement [31] and myosin II [32] influence the time course of secretory granule release, and that newly synthesized dense-core vesicle cargoes are released preferentially compared to aged vesicle cargo [33].

Thus, in response to specific secretagogue stimulation, chromaffin cells release t-PA into the extracellular space. In addition, t-PA can be rapidly released into the circulation in response to stress [3437]. Studies employing adrenergic stimulation and sympathectomy have demonstrated that sympathoadrenal and sympathoneural tissues may represent substantial sources contributing to changes in plasma t-PA concentrations [38, 39].

2.2. PAI-1 Is also Targeted to Catecholamine Storage Vesicles

As a potential mechanism for regulating t-PA activity, inhibitors of t-PA are present in catecholaminergic cells. We have recently demonstrated that plasminogen activator inhibitor-1 (PAI-1) is present in PC12 cells and bovine adrenal medullary chromaffin cells [9]. Secretagogue stimulation led to co-release of PAI-1 with catecholamines, consistent with storage in the same subcellular vesicle. Furthermore, immunoelectron microscopy and sucrose gradient fractionation studies demonstrated localization of PAI-1 in catecholamine storage vesicles [9]. In addition, parallel increases in plasma PAI-1 and catecholamines were observed in response to acute sympathoadrenal activation by restraint stress in mice in vivo [9]. Thus, the overall effect on plasminogen activation and fibrinolysis (both systemically and locally) from catecholamine storage vesicles will depend on a variety of factors that affect local t-PA/inhibitor balance, including the relative rates of synthesis of PAI-1 and t-PA, relative rates of trafficking to the vesicles, and the potential formation of t-PA/inhibitor complexes both within the vesicle and on release. Differential rates of exocytotic release of t-PA compared to PAI-1 could potentially take place as t-PA has been demonstrated to be released more slowly than other granule components [31, 32, 40]. In addition to PAI-1, the t-PA inhibitor, neuroserpin, is present in dense-core secretory vesicles within PC12 cells [10] and a targeting sequence for regulated secretion of neuroserpin has been identified [41].

2.3. t-PA Binding Sites on Catecholaminergic Cells

An additional key mechanism by which local t-PA function is regulated is by the presence of binding sites for t-PA on catecholaminergic cells, as initially demonstrated by Pittman and colleagues [8]. We found that the interaction of t-PA with PC12 cells was saturable and of high capacity [1]. Furthermore, the lysine analog, ε-aminocaproic acid (EACA), also inhibited the interaction [1]. Thus, the recognition specificity of t-PA for chromaffin cells is likely to be dependent on receptors that express C-terminal lysines. Thus, t-PA, released in response to secretagogue stimulation, can bind to its receptors for further amplification of plasminogen activator activity in the local environment of the chromaffin cell.

2.4. uPA and uPAR Expression in Chromaffin Cells

The synthesis and secretion of uPA by chromaffin cells has not been addressed extensively in the literature. uPA secretion by PC12 cells has been detected but accounts for only 5% of the plasminogen activator activity in the long-term conditioned media of these cells; the remaining 95% of plasminogen activator activity is contributed by t-PA [4]. In studies of sympathetic neurons, uPA appears to be released as soon as it is synthesized [8]. Thus, the constitutive release of u-PA may be the predominant source of plasminogen activator activity under basal conditions (on unstimulated chromaffin cells) while the action of t-PA, released in response to secretagogue stimulation, may predominate following its exocytotic release and subsequent binding to the chromaffin cell surface.

The urokinase receptor (uPAR) is also present on PC12 cells. We characterized the binding of single chain u-PA (scu-PA) to untreated PC12 cells. Human scu-PA bound specifically and saturably to PC12 cells (Figure 2(a)) with a Kd of 4 . 3 ± 0 . 5 4  nM and 1 . 1 ± 0 . 2 6 × 1 0 4  sites/cell [11]. A representative binding isotherm and Scatchard analysis are shown in Figures 2(a) and 2(b). Thus, the species specificity that was initially observed (human u-PA does not interact with murine cells and murine u-PA does not interact with human uPAR [42, 43]) does not apply to all interspecies interactions. (Human scu-PA also bound specifically to hamster CHO cells, but minimally to bovine aortic endothelial cells [11, 44]). The affinity of scu-PA for the PC12 cell surface that we determined is consistent with the presence of a functional uPAR on the surface of undifferentiated PC12 cells. In support of this, uPAR mRNA was prominently expressed in PC12 cells as assessed by Northern blotting (Figure 2(c)). Furthermore, we cloned rat uPAR from PC12 cells [11] (Figure 2(d)), and the sequence of the clone we obtained was identical to the sequence published for rat osteoclast uPAR [45]. In addition, using a specific uPAR ELISA, we found substantial expression of uPAR in human pheochromocytoma tissue samples [ 4 8 ± 8 . 2  pg/mg protein ( 𝑛 = 6 )] [11].

fig2
Figure 2: Expression of uPAR by PC12 cells. (a) Binding isotherm of 125I-scu-PA to PC12 cells. Cells in suspension were incubated with 125I-scu-PA, and bound ligand was separated from free by centrifugation over sucrose. Nonspecific binding was determined in the presence of unlabeled scu-PA. (b) Scatchard analysis of the binding isotherm in Panel A. (c) Northern blot analysis showing expression of uPAR in PC12 cells. Poly(A)-enriched RNA was isolated from PC12 cells and Northern blotted with a 458 bp SMaI + BamHI (nuc. 41–493) human uPAR cDNA probe. Northern blotting of U937 cell poly(A) RNA is shown for comparison. (d) cDNA sequence of PC12 uPAR. The rat uPAR sequence was obtained by screening a PC12 cDNA library using a 609 bp EcoRI + NcoI (nuc. 110–718) human uPAR cDNA probe labeled with [α-32P] dCTP.

With NGF-treated PC12 cells, binding sites for u-PA that did not require an active site in u-PA for the interaction were first detected immunochemically by Pittman and colleagues on the bottom surface of the cells [8]. Subsequently, Herschman and colleagues identified uPAR as a neurotrophin responsive gene (preferentially induced by NGF versus EGF) in PC12 cells [12, 46] and demonstrated that NGF-induced uPAR expression is required for NGF-driven PC12 cell differentiation [12, 46, 47]. Induction of uPAR requires ERK/JNK signaling [48], binding of specific Fos and Jun family members to a specific AP-1 site [49, 50], and is regulated by SH2B1β [51].

Thus, taken together, these studies suggest that multiple plasminogen activator receptors can be present simultaneously on the neurosecretory cell surface.

3. Characteristics of the Interaction of Plasminogen with Chromaffin Cells

As a mechanism for concentrating the activity of plasminogen in the plasminogen activator-rich local environment of the chromaffin cell, we found that plasminogen bound to PC12 and bovine adrenal chromaffin cells in a specific, saturable, and reversible manner [1, 2]. Plasminogen activation is markedly enhanced when plasminogen is bound to chromaffin cells. In the presence of PC12 cells, plasminogen activation by t-PA is markedly increased in a cell-dependent fashion, and at the saturating cell density, the enhancement caused by the presence of cells is 6.5-fold [24]. Primary bovine chromaffin cells also markedly stimulate t-PA-dependent plasminogen activation up to 10-fold in a cell concentration-dependent manner [24].

The interactions of plasminogen with substrates and regulatory molecules depend on the lysine binding sites within the disulfide-bonded kringle structures of plasminogen (reviewed in [52]). The interaction of plasminogen with PC12 and bovine adrenal chromaffin cells is blocked by the lysine analogues, EACA and 8-aminooctanoic acid [1, 2], suggesting that proteins with C-terminal lysines, exposed on the cell surface serve as plasminogen binding sites. To explore the relationship between plasminogen binding and stimulation of plasminogen activation, PC12 cells were treated with increasing concentrations of carboxypeptidase B (CpB) that has a preference for cleavage at the carboxyl side of basic amino acids. A dose-dependent decrease in plasminogen binding to PC12 cells was observed, reaching a plateau at 70% inhibition in the presence of CpB (Figure 3(a)) [24]. In addition, CpB treatment decreased the stimulating effect of the cells on plasminogen activation in a dose-dependent manner, approaching a plateau at 88% inhibition (Figure 3(b)) [24]. These results suggest that the chromaffin cell plasminogen receptors that are sensitive to CpB (i.e., proteins exposing C-terminal basic residues on the extracellular face of the cell membrane) are primarily responsible for the promotion of plasminogen activation by these cells.

fig3
Figure 3: Effect of treatment of PC12 cells with CpB on the interaction with plasminogen. PC12 cells were incubated with increasing concentrations of CpB and specific binding of 0.2 μM 125I-plasminogen (a) and plasminogen activation (b) were assessed as described in [24].

4. Role of the Local Chromaffin Cell Plasminogen Activation System in Prohormone Processing

Plasmin functions as a prohormone-processing protease within the neuroendocrine system [1, 17, 5357]. We used the prototypical prohormone chromogranin A (CgA) to study the role of the local chromaffin cell plasminogen activation system in prohormone processing. CgA is the major soluble protein in the core of catecholamine storage vesicles and is released with catecholamines in response to secretagogue stimulation of chromaffin cells of the adrenal medulla and from sympathetic neurons [58]. CgA is present ubiquitously in secretory vesicles of neuroendocrine cells [59] and serves as a precursor of several small, biologically active, secretion-inhibitory peptides that play a key autocrine regulatory role in neuroendocrine secretion from a variety of cell types [6067]. In the case of chromaffin cells and other catecholaminergic cells, CgA serves as a precursor from which peptides are released that modulate catecholamine secretion, contributing to an autocrine, homeostatic, negative-feedback mechanism for regulating catecholamine responses during stress [6567].

We found that plasmin processing of CgA produces regulatory secretion-inhibiting peptides. Plasmin specifically cleaved CgA and decreased the trichloroacetic acid (TCA) precipitability of 125I-CgA, indicating that fragments with Mr < 6,000 had been produced [1, 17]. Most importantly, the peptide fragments generated by plasmic cleavage of CgA inhibited secretagogue-stimulated catecholamine release from PC12 and primary bovine adrenal cells [1]. We identified the specific bioactive CgA peptide produced by plasmin proteolysis [using matrix-assisted laser desorption/ionization mass spectrometry (MALDI)] as a major peptide with a mass/charge ratio (m/z) of 1546 corresponding uniquely to hCgA-(360–373) [17]. In functional studies the hCgA-(360–373) peptide markedly inhibited nicotine-stimulated catecholamine release from pheochromocytoma cells [17].

Chromaffin cells had a marked effect on CgA cleavage by plasmin [1]. In the presence of PC12 cells, the TCA precipitability of 125I-CgA was markedly decreased, indicative of extensive CgA processing (Figure 4) [1]. EACA inhibited processing in the presence, but not in the absence of cells (Figure 4) [1]. Because EACA inhibits plasminogen binding to chromaffin cells, these data suggest a significant contribution of cellular plasminogen receptors to CgA processing.

721657.fig.004
Figure 4: Effect of cells on cleavage of 125I-CgA by plasmin. Either PC12 cells (filled bars) or buffer (open bars) was incubated with unlabeled plasminogen. Then, 125I-CgA and t-PA were added in either the presence or the absence of EACA. The reactions were stopped by addition of aprotinin and centrifuged. The supernatants were analyzed for % TCA precipitability. Results are as mean + SEM; 𝑛 = 6 for each experimental group. ** 𝑃 < 0 . 0 0 1 compared to each of the experimental groups. [1].

We examined the effect of local modulation of the plasminogen activation system on nicotine-mediated catecholamine secretion. Overexpression of t-PA in PC12 cells resulted in a marked ( 8 1 ± 2 %) inhibition of nicotine-stimulated catecholamine release compared with control cells [1]. In the presence of anticatalytic antiplasminogen mAb, catecholamine secretion was markedly increased, compared with isotype control [1]. Thus, the effect of the anticatalytic anti-plasminogen mAb was to restore nicotine-mediated secretion in these cells.

We present a working model for the function of the local chromaffin cell plasminogen activation system in the processing of CgA in Figure 5 [1]. Upon stimulation of the chromaffin cell by a secretagogue, CgA and catecholamines are coreleased by exocytosis. Plasminogen (from circulating sources) and t-PA (synthesized and secreted from the chromaffin cell) bind to the chromaffin cell surface, resulting in activation of plasminogen to plasmin. CgA is subsequently cleaved by plasmin to liberate a peptide fragment that provides a negative feedback loop to modulate subsequent catecholamine release.

721657.fig.005
Figure 5: Working model for a local (autocrine/paracrine) chromaffin cell plasminogen/plasmin system and its interactions with CgA. Upon stimulation of the chromaffin cell by a secretagogue, CgA and catecholamines (cats) are coreleased by exocytosis. Plasminogen (from circulating sources) and its activator t-PA (synthesized and secreted from the chromaffin cell) bind to the chromaffin cell surface, with resultant activation of plasminogen to plasmin. CgA subsequently is cleaved by plasmin to liberate peptide fragments, which provides a negative feedback loop to modulate subsequent catecholamine release. [1].

5. Expression and Subcellular Localization of Plg-R 𝐊 𝐓 in Chromaffin Cells

In earlier studies of chromaffin cell plasminogen receptors, we identified catecholaminergic plasminogen receptors required for enhancing plasminogen activation using targeted specific proteolysis with CpB and a proteomics approach using two-dimensional gel electrophoresis, radioligand blotting, and tandem mass spectrometry. Two major plasminogen-binding proteins that exposed C-terminal lysines on the cell surface contained amino acid sequences corresponding to β/γ actin [24]. An antiactin monoclonal antibody inhibited cell-dependent plasminogen activation and also enhanced nicotine-dependent catecholamine release [24], suggesting that cell-surface-expressed forms of actin (with a processed C-terminus to generate a C-terminal lysine) bind plasminogen, thereby promoting plasminogen activation and increased prohormone processing, leading to inhibition of neurotransmitter release. Although cell surface actin accounted for a substantial fraction of plasminogen binding and activation, a critical role for other cell surface plasminogen-binding proteins with C-terminal lysines was also suggested on these cells. Notably, in plasminogen ligand blotting of 2D gels of PC12 membrane fractions, we also detected a major unknown CpB-sensitive protein migrating with an Mrapp of 17,200 [24].

Recently, we isolated a structurally unique plasminogen receptor from monocyte progenitor cells, the novel protein, Plg-RKT [18]. The Plg-RKT protein is composed of 147 amino acids and has a molecular mass of 17,261 Da. Plg-RKT is synthesized with and exposes a C-terminal lysine on the cell surface, in an orientation to bind plasminogen and to promote plasminogen activation (Figure 6) [18]. Therefore, we investigated expression of Plg-RKT in human and murine adrenal tissues. Prominent staining with anti-Plg-RKT mAb was observed in adrenal medullary chromaffin cells in human (Figure 7(a)) and murine (Figure 7(c)) adrenal tissue [25]. Plg-RKT was also prominently expressed in PC12 cells, bovine adrenal chromaffin cells, and human pheochromocytoma and also in murine hippocampus, representing a nonadrenal source of catecholaminergic cells [25].

721657.fig.006
Figure 6: Structural model of Plg-RKT. Green indicates amino acids within the predicted primary transmembrane helix. Orange indicates amino acids within the predicted secondary transmembrane helix. Red indicates basic amino acids. [18].
fig7
Figure 7: Expression of Plg-RKT in adrenal medulla. Human (a, b) and mouse (c, d) adrenal medulla tissue sections were stained with anti-Plg-RKT mAb, in the absence (a, c) or the presence (b, d) of the decapeptide used for immunization, CEQSKFFSDK (corresponding to the nine C-terminal amino acids of rat Plg-RKT with an amino terminal cysteine added for coupling). Positive tissues are indicated by brown staining with DAB. Scale bar = 100 μm. [25]. ©The American Society for Biochemistry and Molecular Biology.

To assess the subcellular localization of Plg-RKT in catecholaminergic cells and to determine whether the Plg-RKT protein contains a dominant plasma membrane trafficking signal, we transfected PC12 cells with pAcGFP-Plg-RKT (an expression vector in which the Plg-RKT cDNA was inserted in-frame for expression of a GFP-Plg-RKT fusion protein with Plg-RKT at the C-terminus). In confocal microscopy of nonpermeabilized fixed cells, GFP-Plg-RKT showed membrane localization that was highly colocalized with wheat germ agglutinin (WGA) (a well-established cell surface marker) (Figure 8) [25]. In Triton X-114 phase separation experiments [42, 68] Plg-RKT was detected in the detergent phase, but was not detected in the aqueous phase (Figure 9(a)) [25], consistent with Plg-RKT behaving as an integral membrane protein in these cells. Furthermore, anti-Plg-RKT antibody immunoprecipitated both Plg-RKT and uPAR from membrane fractions of PC12 cells (Figure 9(b)) [25], providing further demonstration of the cell membrane localization of Plg-RKT. The physical association of these receptors suggests a key mechanism for promoting plasminogen activation via colocalization of uPA activity (bound to uPAR) with the substrate, plasminogen (bound to Plg-RKT). Furthermore, t-PA also binds to the C-terminus of Plg-RKT [18]. Thus, binding of t-PA and plasminogen to adjacent Plg-RKT molecules may also serve as a means to promote plasminogen activation on the neurosecretory cell surface.

fig8
Figure 8: Plg-RKT re-directs GFP to the cell membrane. PC12 cells were transiently transfected with pAcGFP-Plg-RKT (in which the Plg-RKT cDNA was inserted in frame for expression of a GFP-Plg-RKT fusion protein, with Plg-RKT at the C-terminus) or with control pAcGFP vector. (a) Cell lysates were Western blotted with polyclonal anti-Plg-RKT IgG (lanes 1 and 2) or anti-GFP IgG (lanes 3 and 4). (b) The transfected cells were stained with a combination of DAPI and WGA-rhodamine. The first and third panels of images represent maximum projections of a series of optical slices through the cells. The GFP-Plg-RKT signal was localized primarily to the plasma membrane. The second and fourth panels of images represent sagittal (apical-basal) slices through the same cells along the white dotted line indicated. Here the peripheral plasma membrane localization of GFP-Plg-RKT is also evidentthroughout the vertical stacks of images that were acquired. Scale bar in bottom right corner of image represents 10 microns. [25]. ©The American Society for Biochemistry and Molecular Biology.
fig9
Figure 9: Plg-RKT is an integral plasma membrane cell surface protein. (a) Phase partitioning of Plg-RKT. An immunoreactive band corresponding to the Mrapp of Plg-RKT was detected in the detergent phase, but not in the aqueous phase. (b) Coimmunoprecipitation of Plg-RKT with uPAR. (c) FACS analysis of Plg-RKT expression on intact PC12 cells. Viable cells were gated from nonviable cells and histogram plots of viable cells are shown. Dotted tracings: anti-Plg-RKT mAb IgG. Black tracings: isotype control IgG. Dashed tracings: autofluorescence. [25]. ©The American Society for Biochemistry and Molecular Biology.

In additional studies to address the cell surface orientation of the C-terminus of Plg-RKT, fluorescence activated cell surface (FACS) analysis with anti-Plg-RKT mAb (raised against a synthetic peptide corresponding to the C-terminal peptide of Plg-RKT) demonstrated prominent and specific binding of the mAb to the PC12 cell surface, indicating both cell membrane localization and exposure of the C-terminus of Plg-RKT on the cell surface (Figure 9(c)).

The above studies do not exclude the possibility that other plasminogen receptors with C-terminal lysines (including α-enolase [69], S100A10 [70, 71], Histone H2B [72], and TIP49a [73]) may participate in plasminogen binding on these cells.

6. Regulation of Catecholaminergic Neurosecretory Cell Function by Plg-R 𝐊 𝐓

Based on our studies reviewed above (Section 3), we investigated whether Plg-RKT played a role in promoting plasminogen activation. We stably overexpressed Plg-RKT in PC12 cells, which resulted in prominent expression of Plg-RKT on the cell surface that was markedly greater than the expression of endogenous Plg-RKT (Figure 10) [25]. When cells were stably transfected with Plg-RKT, plasminogen activation was markedly enhanced compared to cells transfected with empty vector (Figure 10(c)) [25]. These results are consistent with a major role for Plg-RKT in cell surface-dependent stimulation of plasminogen activation.

fig10
Figure 10: Overexpression of Plg-RKT increases cell surface plasminogen activation. PC12 cells stably overexpressing Plg-RKT (pCIneo-Plg-RKT) (a) or vector alone (pCIneo) (b) were analyzed by dual color FACS analysis. Viable cells were gated from nonviable cells and histogram plots of viable cells are shown. Dotted tracings: anti-Plg-RKT mAb IgG. Black tracings: isotype control IgG. Dashed tracings: autofluorescence. (c) PC12 cells stably overexpressing Plg-RKT (pCIneo-Plg-RKT) or vector alone (pCIneo) were incubated with plasminogen and t-PA was added and plasminogen activation was measured. Cell-mediated plasminogen activation was substantially greater in Plg-RKT overexpressing cells than in control cells ( 𝑃 < 0 . 0 0 1 at each cell concentration tested) ( 𝑛 = 3 ). [25]. ©The American Society for Biochemistry and Molecular Biology.

Based on our studies reviewed above (Section 4), we tested the effect of overexpression of Plg-RKT on secretagogue-stimulated catecholamine release. Norepinephrine release in response to nicotine was markedly suppressed in cells overexpressing Plg-RKT when compared with release from control cells (Figure 11) [25]. This result is consistent with processing of prohormones by plasmin to produce peptides that feed back to inhibit catecholamine release as outlined in our model in Figure 5.

721657.fig.0011
Figure 11: Effect of overexpression of Plg-RKT on catecholamine release. PC12 cells stably overexpressing either Plg-RKT (filled bars) or vector alone (open bars) were treated with 60 μM nicotine (Nicotine) or buffer (Basal) at 37°C for 15 minutes and catecholamine release was measured by liquid scintillation counting. Percent release was calculated as percentage of secretion [amount released/(amount released + amount in cell lysate)] and results expressed as net release (% secretagogue-stimulated release minus % basal release). Results are represented as mean ± SEM, 𝑛 = 9 for each experimental group. 𝑃 < 0 . 0 0 1 for the Plg-RKT transfectants stimulated with nicotine compared with corresponding values for the vector control cells. [25]. ©The American Society for Biochemistry and Molecular Biology.

7. Conclusions

The plasminogen activation system plays a major role in catecholaminergic cell function by processing secreted hormones that feed back to regulate the neurosecretory characteristics of these cells. In this paper, we summarize results suggesting that Plg-RKT is a crucial molecular focal point in the regulation of the cell-surface-dependent mechanism underlying the ability of catecholaminergic cells to promote local plasminogen activation. Expression of Plg-RKT and additional binding sites for plasminogen [1, 24] and t-PA [1, 8], along with trafficking of t-PA to catecholamine storage vesicles [7, 74], constitute a local catecholaminergic cell plasminogen activation system that regulates cell surface-dependent neuroendocrine prohormone processing (after secretagogue-stimulated storage vesicle exocytosis) that plays a key role in the regulation of neurotransmitter release. In addition, constitutive release of low levels of uPA [4] and subsequent binding to the neurosecretory cell uPAR [11] may also contribute to prohormone processing and to the regulation of differentiation of neuronal cells [12, 46, 47]. These processes are, in turn, locally regulated by the presence of PAI-1 [9] and neuroserpin [10], which are secreted in a regulated fashion, concomitantly with t-PA.

Components of the plasminogen activation system are expressed broadly in neuroendocrine sites, including the cerebral cortex [75], cerebellum [7577], hippocampus [75, 7781], sympathetic neurons [39, 82] as well as the adrenal medulla [7, 77]. Notably, the transcript for Plg-RKT is expressed in each of these tissues (http://www.ebi.ac.uk/gxa/). Future studies to develop and characterize Plg-RKT deficient mice and studies with mice deficient in other plasminogen receptors are warranted to address the role of Plg-RKT and other plasminogen receptors in key plasminogen- and t-PA-dependent neuronal/neuroendocrine plasminogen-dependent processes, including neurite outgrowth [5, 83, 84]; synaptic transmission, NMDA receptor-mediated signaling and excitotoxin-induced neuronal degeneration [85, 86]; long-term potentiation, learning, and memory [56, 78, 80, 8790]; cleavage and activation of other neuroendocrine substrates such as the neurotrophin proBDNF (brain-derived neurotrophic factor) [56], β-endorphin, and α-melanocyte-stimulating hormone [57]; and systemic metabolic and cardiovascular physiologic responses under the control of sympathoadrenal and sympathoneuronal activities [1, 17, 39, 55].

Acknowledgments

This paper was supported by National Institutes of Health Grant HL50398 to R. J. Parmer and Department of Veterans Affairs to R. J. Parmer. S. Nangia was supported by NIH Physician-Scientist Research Training Grant T32DK069263. The authors thank Ms. Linda Bonafede for paper preparation.

References

  1. R. J. Parmer, M. Mahata, Y. Gong et al., “Processing of chromogranin A by plasmin provides a novel mechanism for regulating catecholamine secretion,” Journal of Clinical Investigation, vol. 106, no. 7, pp. 907–915, 2000. View at Scopus
  2. L. A. Miles, S. B. Hawley, and R. J. Parmer, “Chromaffin cell plasminogen receptors,” Annals of the New York Academy of Sciences, vol. 971, pp. 454–459, 2002. View at Scopus
  3. P. Kristensen, D. M. Hougaard, L. S. Nielsen, and K. Danø, “Tissue-type plasminogen activator in rat adrenal medulla,” Histochemistry, vol. 85, no. 5, pp. 431–436, 1986. View at Publisher · View at Google Scholar · View at Scopus
  4. P. Leprince, B. Rogister, P. Delree, J. M. Rigo, B. Andre, and G. Moonen, “Modulation of proteolytic activity during neuritogenesis in the PC12 nerve cell: differential control of plasminogen activator and plasminogen activator inhibitor activities by nerve growth factor and dibutyryl-cyclic AMP,” Journal of Neurochemistry, vol. 57, no. 2, pp. 665–674, 1991. View at Scopus
  5. R. N. Pittman and A. J. DiBenedetto, “PC12 cells overexpressing tissue plasminogen activator regenerate neurites to a greater extent and migrate faster than control cells in complex extracellular matrix,” Journal of Neurochemistry, vol. 64, no. 2, pp. 566–575, 1995. View at Scopus
  6. A. Gualandris, T. E. Jones, S. Strickland, and S. E. Tsirka, “Membrane depolarization induces calcium-dependent secretion of tissue plasminogen activator,” Journal of Neuroscience, vol. 16, no. 7, pp. 2220–2225, 1996. View at Scopus
  7. R. J. Parmer, M. Mahata, S. Mahata, M. T. Sebald, D. T. O'Connor, and L. A. Miles, “Tissue plasminogen activator (t-PA) is targeted to the regulated secretory pathway: catecholamine storage vesicles as a reservoir for the rapid release of t-PA,” The Journal of Biological Chemistry, vol. 272, no. 3, pp. 1976–1982, 1997. View at Publisher · View at Google Scholar · View at Scopus
  8. R. N. Pittman, J. K. Ivins, and H. M. Buettner, “Neuronal plasminogen acivators: cell surface binding sites and involvement in neurite outgrowth,” Journal of Neuroscience, vol. 9, no. 12, pp. 4269–4286, 1989. View at Scopus
  9. Q. Jiang, N. A. Gingles, M. A. Olivier, L. A. Miles, and R. J. Parmer, “The anti-fibrinolytic SERPIN, plasminogen activator inhibitor 1 (PAI-1), is targeted to and released from catecholamine storage vesicles,” Blood, vol. 117, no. 26, pp. 7155–7163, 2011. View at Publisher · View at Google Scholar · View at Scopus
  10. P. K. Parmar, L. C. Coates, J. F. Pearson, R. M. Hill, and N. P. Birch, “Neuroserpin regulates neurite outgrowth in nerve growth factor-treated PC12 cells,” Journal of Neurochemistry, vol. 82, no. 6, pp. 1406–1415, 2002. View at Publisher · View at Google Scholar · View at Scopus
  11. B. J. Fowler, N. Mackman, R. J. Parmer, and L. A. Miles, “Molecular basis for the species specificity of single chain urokinase (Scu-PA) binding to the urokinase receptor (u-PAR),” Circulation, vol. 94, pp. I–512, 1996. View at Publisher · View at Google Scholar
  12. H. R. Herschman, G. D. Ferguson, J. D. Feldman, R. Farias-Eisner, and L. Vician, “Searching for depolarization-induced genes that modulate synaptic plasticity and neurotrophin-induced genes that mediate neuronal differentiation,” Neurochemical Research, vol. 25, no. 5, pp. 591–602, 2000. View at Publisher · View at Google Scholar · View at Scopus
  13. A. H. Chou, S. Zheng, T. Itsukaichi, and B. D. Howard, “Wnt-1 inhibits nerve growth factor-induced differentiation of PC12 cells by preventing the induction of some but not all late-response genes,” Molecular Brain Research, vol. 77, no. 2, pp. 232–245, 2000. View at Publisher · View at Google Scholar · View at Scopus
  14. L. A. Miles, S. B. Hawley, N. Baik, N. M. Andronicos, F. J. Castellino, and R. J. Parmer, “Plasminogen receptors: the sine qua non of cell surface plasminogen activation,” Frontiers in Bioscience, vol. 10, no. 2, pp. 1754–1762, 2005. View at Scopus
  15. E. F. Plow, D. E. Freaney, J. Plescia, and L. A. Miles, “The plasminogen system and cell surfaces: evidence for plasminogen and urokinase receptors on the same cell type,” Journal of Cell Biology, vol. 103, no. 6 I, pp. 2411–2420, 1986. View at Scopus
  16. S. W. Hall, J. E. Humphries, and S. L. Gonias, “Inhibition of cell surface receptor-bound plasmin by α2-antiplasmin and α2-macroglobulin,” The Journal of Biological Chemistry, vol. 266, no. 19, pp. 12329–12336, 1991. View at Scopus
  17. Q. Jiang, L. Taupenot, S. K. Mahata et al., “Proteolytic cleavage of chromogranin A (CgA) by plasmin: selective liberation of a specific bioactive CgA fragment that regulates catecholamine release,” The Journal of Biological Chemistry, vol. 276, no. 27, pp. 25022–25029, 2001. View at Publisher · View at Google Scholar · View at Scopus
  18. N. M. Andronicos, E. I. Chen, N. Baik et al., “Proteomics-based discovery of a novel, structurally unique, and developmentally regulated plasminogen receptor, Plg-RKT, a major regulator of cell surface plasminogen activation,” Blood, vol. 115, no. 7, pp. 1319–1330, 2010. View at Publisher · View at Google Scholar · View at Scopus
  19. R. B. Kelly, “Pathways of protein secretion in eukaryotes,” Science, vol. 230, no. 4721, pp. 25–32, 1985. View at Scopus
  20. H. P. Moore, “Factors controlling packaging of peptide hormones into secretory granules,” Annals of the New York Academy of Sciences, vol. 493, pp. 50–61, 1987. View at Scopus
  21. H. Winkler and E. Westhead, “The molecular organization of adrenal chromaffin granules,” Neuroscience, vol. 5, no. 11, pp. 1803–1823, 1980. View at Publisher · View at Google Scholar · View at Scopus
  22. R. J. Parmer, X. P. Xi, H. J. Wu, L. J. Helman, and L. N. Petz, “Secretory protein traffic. Chromogranin A contains a dominant targeting signal for the regulated pathway,” Journal of Clinical Investigation, vol. 92, no. 2, pp. 1042–1054, 1993. View at Scopus
  23. E. S. Schweitzer and R. B. Kelly, “Selective packaging of human growth hormone into synaptic vesicles in a rat neuronal (PC12) cell line,” Journal of Cell Biology, vol. 101, no. 2, pp. 667–676, 1985. View at Scopus
  24. L. A. Miles, N. M. Andronicos, N. Baik, and R. J. Parmer, “Cell-surface actin binds plasminogen and modulates neurotransmitter release from catecholaminergic cells,” Journal of Neuroscience, vol. 26, no. 50, pp. 13017–13024, 2006. View at Publisher · View at Google Scholar · View at Scopus
  25. H. Bai, N. Baik, W. B. Kiosses, S. Krajewski, L. A. Miles, and R. J. Parmer, “The novel plasminogen receptor, plasminogen receptorKT (Plg-RKT), regulates catecholamine release,” The Journal of Biological Chemistry, vol. 286, pp. 33125–33133, 2011. View at Publisher · View at Google Scholar
  26. K. Fujita, P. Lazarovici, and G. Guroff, “Regulation of the differentiation of PC12 pheochromocytoma cells,” Environmental Health Perspectives, vol. 80, pp. 127–142, 1989. View at Scopus
  27. J. E. Lochner, M. Kingma, S. Kuhn, C. D. Meliza, B. Cutler, and B. A. Scalettar, “Real-time imaging of the axonal transport of granules containing a tissue plasminogen activator/green fluorescent protein hybrid,” Molecular Biology of the Cell, vol. 9, no. 9, pp. 2463–2476, 1998. View at Scopus
  28. J. R. Abney, C. D. Meliza, B. Cutler, M. Kingma, J. E. Lochner, and B. A. Scalettar, “Real-time imaging of the dynamics of secretory granules in growth cones,” Biophysical Journal, vol. 77, no. 5, pp. 2887–2895, 1999. View at Scopus
  29. J. W. Taraska, D. Perrais, M. Ohara-Imaizumi, S. Nagamatsu, and W. Almers, “Secretory granules are recaptured largely intact after stimulated exocytosis in cultured endocrine cells,” Proceedings of the National Academy of Sciences of the United States of America, vol. 100, no. 4, pp. 2070–2075, 2003. View at Publisher · View at Google Scholar · View at Scopus
  30. T. Tsuboi and M. Fukuda, “Synaptotagmin VII modulates the kinetics of dense-core vesicle exocytosis in PC12 cells,” Genes to Cells, vol. 12, no. 4, pp. 511–519, 2007. View at Publisher · View at Google Scholar · View at Scopus
  31. F. Felmy, “Modulation of cargo release from dense core granules by size and actin network,” Traffic, vol. 8, no. 8, pp. 983–997, 2007. View at Publisher · View at Google Scholar · View at Scopus
  32. R. Aoki, T. Kitaguchi, M. Oya et al., “Duration of fusion pore opening and the amount of hormone released are regulated by myosin II during kiss-and-run exocytosis,” Biochemical Journal, vol. 429, no. 3, pp. 497–504, 2010. View at Publisher · View at Google Scholar · View at Scopus
  33. T. Tsuboi, T. Kitaguchi, S. Karasawa, M. Fukuda, and A. Miyawaki, “Age-dependent preferential dense-core vesicle exocytosis in neuroendocrine cells revealed by newly developed monomeric fluorescent timer protein,” Molecular Biology of the Cell, vol. 21, no. 1, pp. 87–94, 2010. View at Publisher · View at Google Scholar · View at Scopus
  34. R. G. Macfarlane and R. Biggs, “Observations on fibrinolysis; spontaneous activity associated with surgical operations, trauma, & c,” The Lancet, vol. 2, no. 6433, pp. 862–864, 1946. View at Scopus
  35. R. Biggs, R. G. Macfarlane, and J. Pilling, “Observations on fibrinolysis; experimental activity produced by exercise and adrenaline,” The Lancet, vol. 1, no. 6448, pp. 402–405, 1947. View at Scopus
  36. P. Fantl and S. E. Simon, “Fibrinolysis following electrically-induced convulsions,” Australian Journal of Experimental Biology & Medical Science, vol. 26, no. 6, pp. 521–529, 1948.
  37. S. A. Schneck and K. N. von Kaulla, “Fibrinolysis and the nervous system,” Neurology, vol. 11, pp. 959–969, 1961. View at Scopus
  38. T. Peng, X. Jiang, Y. F. Wang et al., “Sympathectomy decreases and adrenergic stimulation increases the release of tissue plasminogen activator (t-PA) from blood vessels: functional evidence for a neurologic regulation of plasmin production within vessel walls and other tissue matrices,” Journal of Neuroscience Research, vol. 57, no. 5, pp. 680–692, 1999.
  39. X. Jiang, Y. Wang, A. R. Hand et al., “Storage and release of tissue plasminogen activator by sympathetic axons in resistance vessel walls,” Microvascular Research, vol. 64, no. 3, pp. 438–447, 2002. View at Publisher · View at Google Scholar · View at Scopus
  40. S. J. An, C. P. Grabner, and D. Zenisek, “Real-time visualization of complexin during single exocytic events,” Nature Neuroscience, vol. 13, no. 5, pp. 577–583, 2010. View at Publisher · View at Google Scholar · View at Scopus
  41. S. Ishigami, M. Sandkvist, F. Tsui, E. Moore, T. A. Coleman, and D. A. Lawrence, “Identification of a novel targeting sequence for regulated secretion in the serine protease inhibitor neuroserpin,” Biochemical Journal, vol. 402, no. 1, pp. 25–34, 2007. View at Publisher · View at Google Scholar · View at Scopus
  42. A. Estreicher, A. Wohlwend, D. Belin, W. D. Schleuning, and J. D. Vassalli, “Characterization of the cellular binding site for the urokinase-type plasminogen activator,” The Journal of Biological Chemistry, vol. 264, no. 2, pp. 1180–1189, 1989. View at Scopus
  43. H. Solberg, D. Lober, J. Eriksen et al., “Identification and characterization of the murine cell surface receptor for the urokinase-type plasminogen activator,” European Journal of Biochemistry, vol. 205, no. 2, pp. 451–458, 1992. View at Scopus
  44. B. Fowler, N. Mackman, R. J. Parmer, and L. A. Miles, “Binding of human single chain urokinase to chinese hamster ovary cells and cloning of hamster u-PAR,” Thrombosis and Haemostasis, vol. 80, no. 1, pp. 148–154, 1998. View at Scopus
  45. S. A. Rabbani, N. Rajwans, A. Achbarou, K. K. Murthy, and D. Goltzman, “Isolation and acharacterization of multiple isoforms of the rat urokinase receptor in osteoclasts,” FEBS Letters, vol. 338, no. 1, pp. 69–74, 1994. View at Publisher · View at Google Scholar
  46. R. Farias-Eisner, L. Vician, A. Silver, S. Reddy, S. A. Rabbani, and H. R. Herschman, “The urokinase plasminogen activator receptor (UPAR) is preferentially induced by nerve growth factor in PC12 pheochromocytoma cells and is required for NGF-driven differentiation,” Journal of Neuroscience, vol. 20, no. 1, pp. 230–239, 2000. View at Scopus
  47. R. Farias-Eisner, L. Vician, S. Reddy et al., “Expression of the urokinase plasminogen activator receptor is transiently required during, “priming” of PC12 cells in nerve growth factor-directed cellular differentiation,” Journal of Neuroscience Research, vol. 63, no. 4, pp. 341–346, 2001.
  48. L. Marek, V. Levresse, C. Amura et al., “Multiple signaling conduits regulate global differentiation-specific gene expression in PC12 cells,” Journal of Cellular Physiology, vol. 201, no. 3, pp. 459–469, 2004. View at Publisher · View at Google Scholar · View at Scopus
  49. F. Su, K. R. Kozak, H. Herschman, S. T. Reddy, and R. Farias-Eisner, “Characterization of the rat urokinase plasminogen activator receptor promoter in PC12 cells,” Journal of Neuroscience Research, vol. 85, no. 9, pp. 1952–1958, 2007. View at Publisher · View at Google Scholar · View at Scopus
  50. T. J. Maures, L. Chen, and C. Carter-Su, “Nucleocytoplasmic shuttling of the adapter protein SH2B1β (SH2-Bβ) is required for nerve growth factor (NGF)-dependent neurite outgrowth and enhancement of expression of a subset of NGF-responsive genes,” Molecular Endocrinology, vol. 23, no. 7, pp. 1077–1091, 2009. View at Publisher · View at Google Scholar · View at Scopus
  51. L. Chen, T. J. Maures, H. Jin et al., “SH2B1β (SH2-Bβ) enhances expression of a subset of nerve growth factor-regulated genes important for neuronal differentiation including genes encoding urokinase plasminogen activator receptor and matrix metalloproteinase 3/10,” Molecular Endocrinology, vol. 22, no. 2, pp. 454–476, 2008. View at Publisher · View at Google Scholar · View at Scopus
  52. F. J. Castellino and S. G. McCance, “The kringle domains of human plasminogen,” CIBA Foundation Symposia, no. 212, pp. 46–60, 1997. View at Scopus
  53. J. Hoover-Plow, O. Skomorovska-Prokvolit, and S. Welsh, “Selective behaviors altered in plasminogen-deficient mice are reconstituted with intracerebroventricular injection of plasminogen,” Brain Research, vol. 898, no. 2, pp. 256–264, 2001. View at Publisher · View at Google Scholar · View at Scopus
  54. B. Colombo, R. Longhi, C. Marinzi et al., “Cleavage of chromogranin A N-terminal domain by plasmin provides a new mechanism for regulating cell adhesion,” The Journal of Biological Chemistry, vol. 277, no. 48, pp. 45911–45919, 2002. View at Publisher · View at Google Scholar · View at Scopus
  55. Q. Jiang, S. Yasothornsrikul, L. Taupenot, L. A. Miles, and R. J. Parmer, “The local chromaffin cell plasminogen/plasmin system and the regulation of catecholamine secretion,” Annals of the New York Academy of Sciences, vol. 971, pp. 445–449, 2002. View at Scopus
  56. P. T. Pang, H. K. Teng, E. Zaitsev et al., “Cleavage of proBDNF by tPA/plasmin is essential for long-term hippocampal plasticity,” Science, vol. 306, no. 5695, pp. 487–491, 2004. View at Publisher · View at Google Scholar · View at Scopus
  57. N. Wang, L. Zhang, L. Miles, and J. Hoover-Plow, “Plasminogen regulates pro-opiomelanocortin processing,” Journal of Thrombosis and Haemostasis, vol. 2, no. 5, pp. 785–796, 2004. View at Publisher · View at Google Scholar · View at Scopus
  58. H. Winkler, “The composition of adrenal chromaffin granules: an assessment of controversial results,” Neuroscience, vol. 1, no. 2, pp. 65–80, 1976. View at Publisher · View at Google Scholar · View at Scopus
  59. D. T. O'Connor, R. J. Parmer, and L. J. Deftos, “Chromogranin A: studies in the endocrine system,” Transactions of the Association of American Physicians, vol. 97, pp. 242–250, 1984. View at Scopus
  60. K. Tatemoto, S. Efendic, and V. Mutt, “Pancreastatin, a novel pancreatic peptide that inhibits insulin secretion,” Nature, vol. 324, no. 6096, pp. 476–478, 1986. View at Scopus
  61. J. C. Hutton, H. W. Davidson, and M. Peshavaria, “Proteolytic processing of chromogranin A in purified insulin granules. Formation of a 20 kDa N-terminal fragment (betagranin) by the concerted action of a Ca2+-dependent endopeptidase and carboxypeptidase H (EC 3.4.17.10),” Biochemical Journal, vol. 244, no. 2, pp. 457–464, 1987. View at Scopus
  62. S. Aardal and K. B. Helle, “The vasoinhibitory activity of bovine chromogranin A fragment (vasostatin) and its independence of extracellular calcium in isolated segments of human blood vessels,” Regulatory Peptides, vol. 41, no. 1, pp. 9–18, 1992. View at Publisher · View at Google Scholar · View at Scopus
  63. B. M. Drees, J. Rouse, J. Johnson, and J. W. Hamilton, “Bovine parathyroid glands secrete a 26-kDa N-terminal fragment of chromogranin-A which inhibits parathyroid cell secretion,” Endocrinology, vol. 129, no. 6, pp. 3381–3387, 1991. View at Scopus
  64. B. H. Fasciotto, C. A. Trauss, G. H. Greeley, and D. V. Cohn, “Parastatin (porcine chromogranin A347-419), a novel chromogranin A-derived peptide, inhibits parathyroid cell secretion,” Endocrinology, vol. 133, no. 2, pp. 461–466, 1993. View at Publisher · View at Google Scholar · View at Scopus
  65. J. P. Simon, M. F. Bader, and D. Aunis, “Secretion from chromaffin cells is controlled by chromogranin A-derived peptides,” Proceedings of the National Academy of Sciences of the United States of America, vol. 85, no. 5, pp. 1712–1716, 1988. View at Scopus
  66. E. Galindo, A. Rill, M. F. Bader, and D. Aunis, “Chromostatin, a 20-amino acid peptide derived from chromogranin A, inhibits chromaffin cell secretion,” Proceedings of the National Academy of Sciences of the United States of America, vol. 88, no. 4, pp. 1426–1430, 1991. View at Scopus
  67. S. K. Mahata, D. T. O'Connor, M. Mahata et al., “Novel autocrine feedback control of catecholamine release: a discrete chromogranin A fragment is a noncompetitive nicotinic cholinergic antagonist,” Journal of Clinical Investigation, vol. 100, no. 6, pp. 1623–1633, 1997. View at Scopus
  68. C. Bordier, “Phase separation of integral membrane proteins in Triton X-114 solution,” The Journal of Biological Chemistry, vol. 256, no. 4, pp. 1604–1607, 1981. View at Scopus
  69. L. A. Miles, C. M. Dahlberg, J. Plescia, J. Felez, K. Kato, and E. F. Plow, “Role of cell-surface lysines in plasminogen binding to cells: identification of α-enolase as a candidate plasminogen receptor,” Biochemistry, vol. 30, no. 6, pp. 1682–1691, 1991. View at Scopus
  70. G. Kassam, K. S. Choi, J. Ghuman et al., “The role of annexin II tetramer in the activation of plasminogen,” The Journal of Biological Chemistry, vol. 273, no. 8, pp. 4790–4799, 1998. View at Publisher · View at Google Scholar · View at Scopus
  71. G. Kassam, B. H. Le, K. S. Choi et al., “The p11 subunit of the annexin II tetramer plays a key role in the stimulation of t-PA-dependent plasminogen activation,” Biochemistry, vol. 37, no. 48, pp. 16958–16966, 1998. View at Publisher · View at Google Scholar · View at Scopus
  72. T. Herren, T. A. Burke, R. Das, and E. F. Plow, “Identification of histone H2B as a regulated plasminogen receptor,” Biochemistry, vol. 45, no. 31, pp. 9463–9474, 2006. View at Publisher · View at Google Scholar · View at Scopus
  73. S. B. Hawley, T. A. Tamura, and L. A. Miles, “Purification, cloning, and characterization of a profibrinolytic plasminogen-binding protein, TIP49a,” The Journal of Biological Chemistry, vol. 276, no. 1, pp. 179–186, 2001. View at Publisher · View at Google Scholar · View at Scopus
  74. R. J. Parmer and L. A. Miles, “Targeting of tissue plasminogen activator to the regulated pathway of secretion,” Trends in Cardiovascular Medicine, vol. 8, no. 7, pp. 306–312, 1998. View at Publisher · View at Google Scholar · View at Scopus
  75. A. P. Sappino, R. Madani, J. Huarte et al., “Extracellular proteolysis in the adult murine brain,” Journal of Clinical Investigation, vol. 92, no. 2, pp. 679–685, 1993. View at Scopus
  76. G. C. Friedman and N. W. Seeds, “Tissue plasminogen activator mRNA expression in granule neurons coincides with their migration in the developing cerebellum,” Journal of Comparative Neurology, vol. 360, no. 4, pp. 658–670, 1995. View at Publisher · View at Google Scholar · View at Scopus
  77. L. Zhang, D. Seiffert, B. J. Fowler et al., “Plasminogen has a broad extrahepatic distribution,” Thrombosis and Haemostasis, vol. 87, no. 3, pp. 493–501, 2002. View at Scopus
  78. Z. Qian, M. E. Gilbert, M. A. Colicos, E. R. Kandel, and D. Kuhl, “Tissue-plasminogen activator is induced as an immediate-early gene during seizure, kindling and long-term potentiation,” Nature, vol. 361, no. 6411, pp. 453–457, 1993. View at Publisher · View at Google Scholar · View at Scopus
  79. S. E. Tsirka, A. D. Rogove, T. H. Bugge, J. L. Degen, and S. Strickland, “An extracellular proteolytic cascade promotes neuronal degeneration in the mouse hippocampus,” Journal of Neuroscience, vol. 17, no. 2, pp. 543–552, 1997. View at Scopus
  80. D. Baranes, D. Lederfein, V. Y. Huang, M. Chen, C. H. Bailey, and E. R. Kandel, “Tissue plasminogen activator contributes to the late phase of LTP and to synaptic growth in the hippocampal mossy fiber pathway,” Neuron, vol. 21, no. 4, pp. 813–825, 1998. View at Publisher · View at Google Scholar · View at Scopus
  81. F. J. Sallés and S. Strickland, “Localization and regulation of the tissue plasminogen activator-plasmin system in the hippocampus,” Journal of Neuroscience, vol. 22, no. 6, pp. 2125–2134, 2002. View at Scopus
  82. Z. Hao, C. Guo, X. Jiang et al., “New transgenic evidence for a system of sympathetic axons able to express tissue plasminogen activator (t-PA) within arterial/arteriolar walls,” Blood, vol. 108, no. 1, pp. 200–202, 2006. View at Publisher · View at Google Scholar · View at Scopus
  83. A. T. Jacovina, F. Zhong, E. Khazanova, E. Lev, A. B. Deora, and K. A. Hajjar, “Neuritogenesis and the nerve growth factor-induced differentiation of PC-12 cells requires annexin II-mediated plasmin generation,” The Journal of Biological Chemistry, vol. 276, no. 52, pp. 49350–49358, 2001. View at Publisher · View at Google Scholar · View at Scopus
  84. A. Gutiérrez-Fernández, N. A. Gingles, H. Bai, F. J. Castellino, R. J. Parmer, and L. A. Miles, “Plasminogen enhances neuritogenesis on laminin-1,” Journal of Neuroscience, vol. 29, no. 40, pp. 12393–12400, 2009. View at Publisher · View at Google Scholar · View at Scopus
  85. S. E. Tsirka, A. Gualandris, D. G. Amaral, and S. Strickland, “Excitotoxin-induced neuronal degeneration and seizure are mediated by tissue plasminogen activator,” Nature, vol. 377, no. 6547, pp. 340–344, 1995. View at Scopus
  86. S. E. Tsirka, A. D. Rogove, and S. Strickland, “Neuronal cell death and tPA,” Nature, vol. 384, no. 6605, pp. 123–124, 1996. View at Scopus
  87. P. Carmeliet, L. Schoonjans, L. Kieckens et al., “Physiological consequences of loss of plasminogen activator gene function in mice,” Nature, vol. 368, no. 6470, pp. 419–424, 1994. View at Publisher · View at Google Scholar · View at Scopus
  88. U. Frey, M. Müller, and D. Kuhl, “A different form of long-lasting potentiation revealed in tissue plasminogen activator mutant mice,” Journal of Neuroscience, vol. 16, no. 6, pp. 2057–2063, 1996. View at Scopus
  89. Y. Y. Huang, M. E. Bach, H. P. Lipp et al., “Mice lacking the gene encoding tissue-type plasminogen activator show a selective interference with late-phase long-term potentiation in both Schaffer collateral and mossy fiber pathways,” Proceedings of the National Academy of Sciences of the United States of America, vol. 93, no. 16, pp. 8699–8704, 1996. View at Publisher · View at Google Scholar · View at Scopus
  90. N. W. Seeds, M. E. Basham, and J. E. Ferguson, “Absence of tissue plasminogen activator gene or activity impairs mouse cerebellar motor learning,” Journal of Neuroscience, vol. 23, no. 19, pp. 7368–7375, 2003. View at Scopus