![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
|
|
|
|
|
||
Vol. 20, Issue 24, 5236-5249, December 15, 2009
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Shedding
,||
*Department of Biochemistry and Biophysics and Lineberger Comprehensive Cancer Center, University of North Carolina School of Medicine, Chapel Hill, NC 27599; Departments of Pediatrics and Cell Biology, Vanderbilt University, Nashville, TN 37232; and Department of Biochemistry and Molecular Genetics, Ehime University Graduate School of Medicine, Shitsukawa, Toon, Ehime 791-0295, Japan
Submitted January 2, 2009;
Revised September 28, 2009;
Accepted October 14, 2009
Monitoring Editor: J. Silvio Gutkind
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
(TGF-) shedding by reactive oxygen species (ROS) through the ATP-dependent activation of the P2Y family of GPCRs. We report that ATP stimulates TGF- proteolysis with concomitant EGFR activation and that this process requires TACE/ADAM17 activity in both murine fibroblasts and CHO cells. ATP-induced TGF- shedding required calcium and was independent of Src family kinases and PKC and MAPK signaling. Moreover, ATP-induced TGF- shedding was completely inhibited by scavengers of ROS, whereas calcium-stimulated shedding was partially inhibited by ROS scavenging. Hydrogen peroxide restored TGF- shedding after calcium chelation. Importantly, we also found that ATP-induced shedding was independent of the cytoplasmic NADPH oxidase complex. Instead, mitochondrial ROS production increased in response to ATP and mitochondrial oxidative complex activity was required to activate TACE-dependent shedding. These results reveal an essential role for mitochondrial ROS in regulating GPCR-induced growth factor shedding. |
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
). The discovery of the essential role of EGFR in propagating signals generated by G-protein–coupled receptor (GPCR) agonists indicates that EGFR may function as a central signal integration point for stimuli impacting the cell surface (Blobel, 2005; Ohtsu et al., 2006). A wide variety of GPCR agonists, including lysophosphatidic acid, phenylephrine, and carbachol, can harness EGFR to promote ERK activation, leading to physiological and pathophysiological responses (Yan et al., 2002; Gschwind et al., 2003; Schafer et al., 2004b; Zhang et al., 2004). GPCR cross-talk was previously thought to consist of entirely intracellular signaling pathways that led to EGFR phosphorylation, independent of ligand binding and dimerization. However, elucidation of a rapid, metalloprotease-dependent growth factor cleavage step leading to EGFR activation (Prenzel et al., 1999) revealed the importance of regulated proteolysis in GPCR-EGFR transactivation. More recent studies have shown the in vivo importance of this phenomenon (Zhang et al., 2004; Lautrette et al., 2005).
The seven members of the EGF family of growth factors, amphiregulin (AR), betacellulin, EGF, epigen, epiregulin, heparin-binding EGF (HB-EGF), and transforming growth factor alpha (TGF-), are all initially synthesized as type-I transmembrane precursors, containing the growth factor moiety in the ectodomain (Lee et al., 2003). Several studies have demonstrated the biological activity of the noncleavable, membrane-anchored precursor molecules (Brachmann et al., 1989; Wong et al., 1989), but the metalloprotease dependence of transactivation, the loss of EGFR signaling in cells treated with metalloprotease inhibitors (Dong et al., 1999; Borrell-Pages et al., 2003), and the convergence of phenotypes of growth factor and protease knockout models indicate that proteolytic cleavage of the growth factors is an important and regulatable step in most contexts (Peschon et al., 1998; Iwamoto et al., 2003; Jackson et al., 2003; Sternlicht et al., 2005).
A variety of in vitro and in vivo evidence points to the ADAM family member tumor necrosis factor alpha–converting enzyme (TACE/ADAM17) as the critical convertase for TGF-, AR, and HB-EGF. ADAMs (a disintegrin and metalloprotease), along with matrix metalloproteases, belong to the metzincin family of zinc-dependent proteases (Blobel, 2005). When mice lacking active TACE were compared with TGF-, AR, and HB-EGF knockouts, they were found to share epithelial defects with homozygous TGF- null mice (Peschon et al., 1998), loss of mammary gland branching as in mice lacking AR (Sternlicht et al., 2005), and heart and lung defects with HB-EGF–/– animals (Iwamoto et al., 2003; Jackson et al., 2003). Fibroblasts derived from the TACE-deficient mice were impaired in shedding of TGF-, HB-EGF, and AR, but shedding could be rescued by transfection of wild-type TACE into the cells (Sunnarborg et al., 2002). TACE also cleaved each growth factor in vitro at the physiologically relevant site (Sunnarborg et al., 2002; Hinkle et al., 2004). In cell culture, knockdown of TACE expression can also have an inhibitory effect on growth factor–dependent transactivation by lysophosphatidic acid, angiotensin II, and epoxyeicosatrienoic acid stimulation (Schafer et al., 2004b; Mifune et al., 2005; Chen et al., 2007). The overlapping phenotypes of mice lacking these growth factors and those lacking TACE/ADAM17, along with the in vitro results, support a critical role for the soluble forms of the growth factors and highlight the importance of their proteolysis as a regulatory event.
Despite its presence at a critical signaling juncture, the regulation of ADAM metalloprotease activity is still not fully understood. ADAMs are type I transmembrane proteins that possess an archetypal organization, including the metalloprotease and disintegrin domains, along with a cytoplasmic domain often rich in SH3-binding sites that could potentially regulate ADAM inside-out signaling (Seals and Courtneidge, 2003; Blobel, 2005). Several signaling components have been implicated in GPCR-initiated TACE activation. Src-like nonreceptor tyrosine kinases have long been accepted as intermediates in EGFR transactivation and have been found in association with several ADAMs leading to phosphorylation (Pierce et al., 2001; Seals and Courtneidge, 2003; Luttrell and Luttrell, 2004; Zhang et al., 2006). Elevation of intracellular calcium was also found to stimulate the release of ErbB ligands in an ADAM-dependent manner (Mifune et al., 2005; Sanderson et al., 2005; Horiuchi et al., 2007), whereas protein kinase C (PKC) has long been suspected to play a role in ADAM activation because of the ability of phorbol 12-myristate 13-acetate (PMA) to trigger PKC signaling and to stimulate ectodomain shedding. Mitogen-activated protein kinase (MAPK) proteins have also been implicated both prior and subsequent to EGFR activation (Prenzel et al., 1999; Umata et al., 2001; Gschwind et al., 2003; Fischer et al., 2004). Lastly, reactive oxygen species (ROS) can function as second messengers in response to G-protein signaling (Zhang et al., 2001; Cross and Templeton, 2006). ROS have been shown to be necessary for angiotensin II- and endothelin-1–mediated TACE activation and HB-EGF shedding, possibly through direct modification of a cysteine that coordinates the binding of the inhibitory prodomain (Zhang et al., 2001; Mifune et al., 2005; Chen et al., 2006). The wide variety of possible regulatory mechanisms suggests substrate- or cell type-specific pathways dependent on the stimulant or intended signal effect.
In this study we characterize a ROS-mediated pathway wherein ATP initiates P2Y signaling, stimulating TACE-dependent TGF- proteolysis and EGFR phosphorylation. TGF- shedding is independent of Src, PKC, and MAPK signaling. We report for the first time the requirement for mitochondrially derived ROS, whose production is stimulated by ATP, for regulation of GPCR-induced growth factor shedding. Collectively, these results offer a model that can be utilized to further our understanding of the regulatory mechanisms of TACE-dependent cell surface proteolysis.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
(MF9) was purchased from Neomarkers (Fremont, CA) and anti-TACE from Exalpha Biologicals (Shirley, MA). Horseradish peroxidase–conjugated secondary antibodies and ATP
S were from Roche Molecular Biochemicals (Indianapolis, IN). Adenosine and uridine triphosphate were purchased from GE Healthcare (Waukesha, WI). EGF was from Harlan Bioproducts (Indianapolis, IN) and TAPI-2 from Peptides International (Louisville, KY). N-Acetyl-L-cysteine, rotenone (NAC), myxothiazol, and AG1478 were from Sigma (St. Louis, MO) and all other chemicals were from Calbiochem (La Jolla, CA).
Cell Lines, Transfections, and Stable Clones
EC-4 (Tace+/+) and EC-2 (Tace
Zn/Zn) fibroblasts, provided by Amgen (Thousand Oaks, CA; Reddy et al., 2000), and their derivatives were maintained in DMEM/F-12, 1% FBS with antibiotic. Wild-type and M2 Chinese hamster ovary (CHO) cells and their derivatives were maintained in DMEM, 10% FBS, 1x nonessential amino acids and antibiotic. HL-60 cells were maintained in RPMI media, 10% FBS with antibiotic. Plasmid transfections were performed using LipofectAMINE (Invitrogen, Carlsbad, CA). The human EGFR cDNA (Saxon and Lee, 1999) was subcloned into modified pCEP4 (Invitrogen) and stably expressed in all indicated cell lines after selection with 800 µg/ml hygromycin B (Roche). Alkaline phosphatase–tagged TGF- (Tokumaru et al., 2000) was kindly sent by Dr. Carl Blobel (Hospital for Special Surgery, New York, NY). Stable expression of AP-TGF- in wild-type CHO cells was achieved by selecting clones in 500 µg/ml G418 (Sigma). For stable expression in M2 cells that had already been subjected to G418 selection, AP-TGF- was subcloned into pcDNA 3.1 hygromycin B vector (Invitrogen) using a PCR-generated fragment (HindIII-capped forward PCR primer 5'-AAGCTTGTTCTAGCGGCACCGC-3'; KpnI-capped reverse PCR primer 5'-GGTACCCGTTCTTACAGCAAAAGGC-3') and the TOPO cloning kit (Invitrogen). Stable clones were selected in 800 µg/ml hygromycin B. For stable expression of pcDNA3 mouse TACE, transfected EC-2 and M2 CHO cells were selected in 100 µg/ml Zeocin (Reddy et al., 2000).
RNA Interference
Small interfering RNA (siRNA) duplexes against mouse TACE (5'GGGAUUCCUUUCAGCAUUCUUGUCCA-3') from Invitrogen were transiently transfected into EC-4 EGFR cells by adding 6 µl RNAiMAX (Invitrogen) to 100 nM siRNA per 35-mm plate according to the manufacturer's instructions. A fluorescent-tagged scrambled sequence was used as a control. Cells were harvested 48 h after transfection as described below.
Harvesting, Immunoprecipitation, and Western Blot Analysis
Confluent cells were washed twice in serum-free medium (SFM) and then starved for 4 h at 37°C in SFM. After a 5-min stimulation, conditioned medium was collected, and cells were washed in phosphate-buffered saline (PBS) and lysed in 1% Triton X-100, 50 mM Tris, pH 7.4, 150 mM NaCl with 10 µg/ml leupeptin, 20 µg/ml aprotinin, 1 mM phenylmethylsulfonyl fluoride, 2 µM sodium orthovanadate, 10 mM sodium fluoride, and 5 mM sodium molybdate (lysis buffer). Protein concentration was determined with a BCA assay kit (Pierce, Rockford, IL). For some experiments media were concentrated using Sep-Pak C-18 reverse-phase cartridges (Waters, Milford, MA; Sandgren et al., 1990), and TGF- in both lysates and media was measured with a specific radioimmunoassay (RIA; Russell et al., 1993). For immunoprecipitations, equal amounts of protein were incubated for 60 min with primary antibody at 4°C followed by protein G agarose (Invitrogen) at 4°C for 1 h. Beads were washed in 20 mM Tris, pH 7.5, 0.1 M NaCl, 0.1 mM EDTA, and 0.1% Nonidet P-40, separated by SDS-PAGE, and transferred to Immobilon polyvinylidene difluoride (Millipore, Billerica, MA). Membranes were blocked in Tris-buffered saline, 0.1% Tween 20, and 3% BSA or 5% nonfat dry milk. Bands were visualized using Pierce SuperSignal West Pico chemiluminescent system for autoradiography.
Expression of P2Y Receptors by RT-PCR
RNA was extracted from wild-type EC-4 or CHO cells grown in 10-cm dishes using RNeasy Mini Kit (Qiagen, Valencia, CA). RNA was isolated from fresh jejunum tissue with Trizol (Invitrogen) according to the manufacturer's protocol (Troyer et al., 2001). Total RNA was used to prepare cDNA with MMTV Reverse Transcriptase (Invitrogen). PCR amplification used primers for mouse P2Y2 or P2Y4, corresponding to analogous regions to yield the same size products and species-specific primers for β-actin as a loading control. Primers for P2Y2, 5'-ACGGTGCTCTGCAAGCTGGTGC-3'; 5'-GTAGAGGGTGCGCGTGACGTGG-3'; P2Y4, 5'-TACTACTATGCTGCCAGAAACCAC-3'; 5'-AGCAAAGACAGTCAGCACCACAGC-3'; mouse β-actin, 5'-GACAACGGATCCGGCATGTG C-3'; 5'-GCGGCAGTGGCC ATCTCCTGC-3'; and hamster β-actin, 5'-ACTCCTACGTGGGTGACGAG-3'; 5'-AAGGAAGGCTGGAAAAGAGC-3'.
Inositol Phosphate Incorporation
EC-4 cells were seeded into 24-well plates and cultured overnight. The next day, [3H]myo-inositol was added at 0.4 µCi/well and incubated overnight. Cells were stimulated for 15 min with vehicle, 100 µM ATP or 100 µM UTP as indicated, and [3H]inositol incorporation was assayed as previously described (Hoffmann et al., 1999).
Alkaline Phosphatase Assay
Cells were plated in 24-well dishes at a density of 2.5 x 105 per well for 18 h. Cells were washed twice in SFM and starved in fresh SFM for 4 h at 37°C. Vehicle (DMSO or water) or inhibitors were added directly to cells for 30 min. After inhibition, media were changed to fresh SFM containing inhibitor and agonist for 5 min. Media were then collected and centrifuged for 30 min at 14,000 x g at 4°C. Media aliquots were mixed with an equal volume of 2x AP buffer (100 mM glycine, pH 10.4, 1 mM MgCl2, 1 mM ZnCl2, and 5.4 mM p-nitrophenylphosphate; Sahin et al., 2006) and allowed to develop at room temperature. Absorbance at 405 nm was measured to quantify alkaline phosphatase (AP) activity. To test for possible interference in alkaline phosphatase activity by signaling inhibitors, conditioned media from 150-mm dishes of vehicle or 100 µM ATP-stimulated CHO-APT cells were divided after harvesting from cells for treatment with the appropriate concentration of each reagent.
Exosome Purification
Exosomes were purified from conditioned media by differential centrifugation, as previously described (Thery et al., 2006). In short, 8 x 107 cells were serum-starved for 4 h and then pretreated with vehicle or 20 µM TAPI-2 for 30 min. In fresh SFM, cells were stimulated with vehicle or 100 µM ATP for 5 min, and the media were collected and pooled. To remove live, whole cells the media were subjected to centrifugation at 300 x g for 10 min. An aliquot of supernatant along with the entire pellet, which was collected in lysis buffer, were stored at –80°C. The remaining supernatant was centrifuged again at 2000 x g for 10 min to separate dead cells and 10,000 x g for 30 min to remove cell debris. Aliquots were again collected for analysis. Finally, exosomes were pelleted by centrifugation at 100,000 x g for 60 min. The lysed pellets were prepared for Western blot analysis, and the media aliquots were split and prepared for Western blot analysis and AP assays.
Cell Viability
An equal number of cells was plated in duplicate, 24 h before treatment for each condition. Cells were then serum-starved for 4 h before treatment with the appropriate concentration of reagent for 30 min. After that, cells were washed twice in PBS and then trypsinized and collected. Viable cells were counted on a hemacytometer, and total cells per plate were compared with vehicle-treated cells to determine the effect of each reagent on cell number.
Measurement of Membrane NADPH Activity
CHO or HL-60 cells (1.5 x 108) were stimulated with vehicle, 100 µM ATP, or 20 µM PMA for 5 min at 37°C. Media were removed, and cells washed twice in ice-cold PBS. Intact cells were collected from the plates in PBS. Plasma membranes were isolated using the Membrane Protein Extraction Kit as directed by manufacturer (BioVision, Mountain View, CA). Protein concentrations were determined, and 2 µg of protein was used to measure SOD inhibitable superoxide production by cytochrome C reduction in vitro (Teufelhofer et al., 2003; Dikalov et al., 2007). Briefly, 100 µM cytochrome C and 100 µM NADPH with or without 100 U/ml SOD were prepared in HBSS and mixed with plasma membrane proteins in an equal volume in a 96-well plate and incubated at 37°C for the length of the assay. Triplicate samples were assayed. Absorbance at 540, 550, and 560 nm was measured in a SpectraMAX platereader (Molecular Devices, Sunnyvale, CA). The difference in absorbance of 550 nm and the average of 540 and 560 nm was calculated for each well. The average background absorbance (reaction solution without protein) was then subtracted and the difference in SOD-treated and untreated samples was determined. This value was then converted to picomoles of reduced cytochrome C using the extinction coefficient 21.1 mM–1 cm–1.
Measurement of Mitochondrial ROS
Mouse cells were plated in 24-well dishes at a density of 2.5 x 105/well and cultured for 18 h. Cells were washed twice in SFM and starved in fresh SFM for 4 h at 37°C. Cells were loaded with 250 nM CM-H2XRos MitoTracker Red (MTR; Molecular Probes, Eugene, OR) for 15 min at 37°C away from direct light. Media were aspirated, and cells were washed twice in phenol-free, SFM supplemented with 2 mM L-glutamine and left in washing media. Cells were then stimulated for 5 min and excited at 544 nm, and emission was detected at 612 nm on the fluorescent platereader SpectraMax Gemini XS (Molecular Devices). Data are presented as a box-and-whisker plot to exhibit the full range of data points.
Statistical Analysis
The nonparametric Wilcoxon rank-sum test was used for two-group comparisons. p < 0.05 was considered statistically significant. Statistical analyses were performed using Prism 5.0 from GraphPad Software (San Diego, CA) and SAS statistical software (version 9.1, SAS Institute, Cary, NC).
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
as a model system. Stimulation of EC-4–transformed fibroblasts that stably express EGFR (EC-4 EGFR; Figure 1A, bottom panel) with the purinergic receptor agonist ATP led to the phosphorylation of EGFR, as did treatment with exogenous EGF (Figure 1A). ATP transactivation was also able to stimulate phosphorylation of Shc, an adaptor molecule for EGFR (Figure 1B). ATP-induced Shc phosphorylation was dependent on EGFR activity as the EGFR-kinase inhibitor AG1478 was able to block both ATP- and EGF-stimulated Shc activation. Therefore, these cells possess the endogenous components to accomplish GPCR-induced EGFR transactivation and downstream signaling.
|
(Sunnarborg et al., 2002) and fibroblasts often express P2Y receptors, we asked if ATP stimulation would lead to rapid release of soluble TGF-. ATP stimulates P2Y2 purinergic receptors with an EC50 of 
25 µM and the maximal response at 100 µM (Soltoff et al., 1998). Using a highly specific, TGF- RIA (Russell et al., 1993), we detected a greater than twofold increase in endogenous TGF- shedding 5 min after addition of 100 µM ATP (Figure 1C). The effect of ATP was not dependent on hydrolysis, as indicated by comparable results with the noncleavable ATP analog, ATPS. These results indicate that GPCR stimulation via purinergic receptors leads to rapid release of soluble TGF-, and EGFR transactivation.
We also wanted to identify the purinergic receptor subtype responsible for this shedding event. Purinergic receptors are classified on the basis of their agonist profile. ATP can stimulate members of the P2X and P2Y family of receptors. UTP, in contrast, stimulates only P2Y receptors, and murine P2Y2 and P2Y4 receptors are equipotently activated by ATP and UTP (Abbracchio et al., 2006). In mouse EC-4 cells, as shown in Figure 1C, UTP stimulates TGF- shedding at levels similar to that of ATP, indicating use of P2Y receptors. ATP and UTP also stimulated comparable [3H]inositol incorporation in these cells (Supplemental Figure S1). Semiquantitative reverse transcriptase-PCR on RNA from EC-4 mouse fibroblasts revealed expression of only the P2Y2 receptor subtype. P2Y4 receptor was not visible at this low cycle number but was visible in the positive control jejunum tissue sample (Figure 1D). These results, together with the ability of ATP and UTP to stimulate GPCR signaling and TGF- shedding, identify P2Y2 as the critical receptor.
The paradigm for GPCR-induced EGFR transactivation requires a metalloprotease-dependent growth factor cleavage step (Prenzel et al., 1999). We verified the dependence on metalloprotease activity in this model system by treating cells with the hydroxamate compound TAPI-2. In the presence of this inhibitor, ATP transactivation of EGFR was abolished, whereas EGF still activated the receptor (Figure 2A). Therefore, purinergic stimulation of EGFR requires metalloprotease activity.
|
). In our system, ATP stimulated TGF- release, and as proTGF- is preferentially cleaved by TACE over ADAM10 (Le Gall et al., 2009), we hypothesized that TACE was responsible for the ATP-induced TGF- release and EGFR transactivation that we have observed. To test this hypothesis, we performed siRNA experiments to knock down endogenous TACE in EC-4 EGFR cells. In control transfected cells, both EGF and ATP stimulated phosphorylation of EGFR, whereas in the absence of TACE, ATP-dependent phosphorylation of EGFR was significantly decreased (Figure 2B). These results were confirmed in TACE-deficient EC-2 mouse fibroblasts, which are the complementary cells to wild-type EC-4 cells (Reddy et al., 2000), stably expressing EGFR. Although EGF activated its receptor in wild-type and TACE-deficient cells, ATP was unable to stimulate transactivation of EGFR in the EC-2 cells compared with the wild-type EC-4 cells (Figure 2C). Similar to our previous observations for constitutive shedding (Sunnarborg et al., 2002), induction of TGF- release by treatment with 100 µM ATP was abolished in the absence of TACE (Figure 2D). RIA measurements of unstimulated cell lysate indicated comparable TGF- levels in EC-4 and EC-2 cells. We were also able to restore ATP-induced shedding in EC-2 cells by stably expressing mouse TACE (Figure 2E). Thus, we conclude that TACE is the crucial protease responsible for ATP-induced EGFR transactivation and TGF- shedding in mouse fibroblasts, though TGF- may not be exclusively responsible for ATP-induced EGFR activation in this system.
Further investigation into the regulation of TGF- shedding required an assay system with high-throughput capabilities. Alkaline phosphatase-tagged EGF-like growth factors are an established method of monitoring growth factor cleavage by quantifying AP activity in conditioned media (Tokumaru et al., 2000; Sahin et al., 2004; Tanaka et al., 2004; Horiuchi et al., 2007). To facilitate transfection and culturing, we shifted our cell model to CHO cells, which have been used previously to study the regulation of growth factor shedding (Teixido et al., 1990; Gechtman et al., 1999). Although lacking endogenous EGF-family growth factors, CHO cells express TACE (Borroto et al., 2003; Li and Fan, 2004) and have been reported to respond to purinergic stimulation (Iredale and Hill, 1993). RT-PCR of CHO RNA revealed strong expression of P2Y2, although P2Y4 was undetectable even with elevated cycle numbers (Figure 3A and data not shown). Thus, CHO cells should provide a suitable model for further study of GPCR-induced TGF- shedding.
|
(AP-TGF-) or empty vector (Tokumaru et al., 2000). We next determined the effect of ATP on AP-TGF- shedding. Analogous to our results in mouse cells, a 5-min stimulation with 100 µM ATP caused a significant increase of AP activity in the media compared with mock stimulation (Figure 3B). Likewise, 100 µM UTP was able to stimulate a strong TGF- shedding response in CHO cells. Metalloprotease dependence was verified for ATP- and UTP-induced shedding by pretreatment of the cells with TAPI-2. When these cells were stimulated with water, ATP, or UTP, shedding was almost completely abrogated (Figure 3B). We next confirmed that TACE was responsible for AP-TGF- shedding in CHO cells by using CHO M2 cells that carry inactivating mutations in each endogenous TACE allele (Arribas and Massagué, 1995; Li and Fan, 2004). After ATP treatment, induction of AP activity was negligible in M2-AP-TGF- cells stably expressing AP-TGF-, compared with vehicle-treated cells (Figure 3C). When TACE was stably expressed in these cells, constitutive shedding remained low, whereas ATP-induced shedding was partially recovered. These results confirm the presence of a P2Y2-TACE-TGF- pathway in CHO cells.
We wanted to confirm that the TGF- and AP activity we detected in conditioned media resulted from proteolytic cleavage and not a protease-independent secretory event such as exosome release (Stoeck et al., 2006). We therefore isolated exosomes from ATP-stimulated CHO AP-TGF-–conditioned medium and examined TGF- levels in this fraction. In ATP-treated cells, AP-TGF- was detected in supernatants of conditioned media after depletion of each membrane fraction (Supplemental Figure S2A, top panel), including exosomes. The AP activity in conditioned media nonetheless remained strong in the membrane-depleted fractions (Supplemental Figure S2B). AP-TGF- was also detected in each of the ATP-stimulated membrane fractions (Supplemental Figure S2A, middle panel) including in exosomes as identified by the marker, flotillin (Supplemental Figure S2A, bottom panel). TAPI-2 treatment was able to block release of AP-TGF- in each case. A small loss of signal at each stage was observed, which likely corresponds to the membrane bound AP-TGF- removed from the media by removal of the membrane fractions. These results demonstrate that after ATP stimulation the majority of TGF- released from cells is cleaved and not secreted in membranes.
To determine potential regulatory steps of TACE-dependent ATP-induced TGF- shedding, we used chemical inhibitors to ask which pathways contributed to the release of TGF- after ATP stimulation in CHO AP-TGF- cells. Calcium signaling has been implicated in shedding and EGFR transactivation (Mifune et al., 2005; Horiuchi et al., 2007) and is also a common signaling component of ATP-induced purinergic pathways (Abbracchio et al., 2006); therefore, we wanted to test the effect of calcium on TGF- shedding in our cell model. Treatment with the calcium specific chelator BAPTA-AM significantly inhibited ATP-dependent TGF- shedding (Figure 4). We therefore tested whether calcium influx stimulated shedding with a nontoxic concentration of calcium ionophore A23187 (1 µM, Supplemental Table S1), which led to a significant increase in shedding over control levels, comparable to ATP (Figure 5A). A23187-induced shedding was TACE-dependent, as shedding was not significantly induced in the TACE-deficient M2 AP-TGF- CHO cells, while being restored by expression of mouse TACE in these cells (Figure 5B). Similar to ATP, A23187-induced shedding was partially inhibited by BAPTA-AM (Figure 5A). One explanation for the partial inhibition by BAPTA-AM is the possibility that a local influx of extracellular calcium may avoid BAPTA-AM chelation, thus allowing for ATP- or A23187-dependent shedding. To test for a role for extracellular calcium, we used the nonspecific chelator EGTA in aqueous solution to sequester extracellular calcium before stimulation. EGTA strongly inhibited TGF- shedding by both ATP and A23187 (Figure 5C). Despite a high affinity for calcium, EGTA can also chelate other metal ions such as zinc that could interfere with TACE activity. To rule out this possibility, we used hydrogen peroxide as a calcium-independent stimulator of TGF- shedding (Shao and Nadel, 2005). Hydrogen peroxide stimulated TGF- shedding, but was not sensitive to EGTA pretreatment (Figure 5D). These results demonstrate that extracellular calcium is an important signaling component in TACE-mediated ATP-induced TGF- shedding and indicate that hydrogen peroxide can stimulate TGF- shedding independently of calcium.
|
|
; Izumi et al., 1998; Pierce et al., 2001; McCole et al., 2002; Nagano et al., 2004; Stoeck et al., 2006). Involvement of PKC proteins can be tested with the inhibitor bisindolylmaleimide-1 (BIM-1), whereas Src family members can be probed with the inhibitor PP2 and the nonfunctional structural homolog PP3. As shown in Figure 4, PP2 had an insignificant effect with results indistinguishable from PP3, indicating no specific contribution of Src and related nonreceptor tyrosine kinase pathways. Treatment with the broad spectrum PKC inhibitor BIM-1 also did not significantly inhibit ATP-stimulated TGF- shedding. MAPK pathways have also been associated with growth factor shedding after stress or other stimuli (Fan and Derynck, 1999; Gechtman et al., 1999; Umata et al., 2001; Fischer et al., 2004; Zhuang and Schnellmann, 2004). However, we observed no inhibition of TGF- release when p38 MAPK signaling (SB202190; SB203580) or ERK activity (PD98059) was blocked with inhibitors for each pathway (Figure 4). These results were confirmed by RIA analysis in mouse EC-4 cells (data not shown). To rule out toxicity of inhibitors as a trivial explanation for our results, we tested each inhibitor's effect on cell viability at the concentration used in these experiments. We found that cell numbers remained the same as control-treated cells for all reagents (Supplemental Table S1). Moreover, these signaling inhibitors did not interfere with the ability of the TGF- antibody to measure TGF- by RIA (Supplemental Figure S3A), nor did they interfere with the ability to detect AP activity (Supplemental Figure S3B), though BAPTA-AM increased the level of AP activity in some experiments. Therefore, we have established that these reagents have no effect on cell viability or assay function, yet despite the demonstrated importance of these pathways, they are not required for ATP-stimulated TGF- shedding.
Recent studies of ADAM regulation, and specifically TACE, have focused on the ability of ROS to act as signaling intermediates. Our results indicate that hydrogen peroxide could stimulate TGF- shedding (Figure 5D), consistent with studies of TACE activation and shedding of both HB-EGF and TGF- (Zhang et al., 2001; Frank and Eguchi, 2003; Shao and Nadel, 2005). Cell-derived ROS have also been found as signaling intermediates downstream of the angiotensin II receptor and P2Y purinergic receptors (Sauer et al., 2001; Mifune et al., 2005; Pines et al., 2005); thus we hypothesized that cellular ROS may also play a role in ATP-induced TGF- shedding. We tested this by treating CHO AP-TGF- cells with the ROS scavenger NAC. Shedding of TGF- was dramatically reduced in the presence of NAC, pointing to a significant role for ROS in the regulation of TGF- shedding (Figures 4 and 6A). The importance of cell-derived ROS was confirmed by testing endogenous TGF- shedding in mouse fibroblasts, where NAC also significantly inhibited ATP-stimulated shedding (Supplemental Figure S4A). To determine if ROS-induced TGF- shedding was metalloprotease-dependent, we used the inhibitor TAPI-2, which completely blocked hydrogen peroxide–induced TGF- shedding (Figure 6B). The importance of TACE as a specific metalloprotease was demonstrated by the lack of shedding from M2-AP-TGF- cells and the restoration of hydrogen peroxide–induced shedding with the expression of mouse TACE (Figure 6C). Thus, ROS signaling appears to be a key intermediate in TACE-mediated TGF- shedding that is part of a conserved pathway following P2Y receptor activation.
|
). In most nonphagocytic cells the NADPH oxidase produces the majority of cytoplasmic superoxide, which is then rapidly dismutated into hydrogen peroxide (Brandes and Kreuzer, 2005). Several reports present evidence of a role for the NADPH oxidase in UV-induced (Singh et al., 2009) and GPCR-induced transactivation of EGFR (Ushio-Fukai et al., 1996; Lemjabbar et al., 2003; Zhang et al., 2004; Shao and Nadel, 2005; Stoeck et al., 2006). To investigate a role for the NADPH oxidase in TGF- shedding, we tested the effect of apocynin, which blocks formation of the complete NADPH oxidase complex, thus blocking ROS production by this source. CHO AP-TGF- cells preincubated with 1 mM apocynin and then stimulated with water or ATP showed no effects on TGF- shedding (Figure 7A). This might be expected since CHO cells have been reported to lack p22phox, a key component of the NADPH oxidase complex (Takeya et al., 2003). To further confirm the inability of CHO cells to produce NADPH oxidase-dependent ROS, we measured the production of superoxide from isolated membrane proteins of stimulated cells. CHO cells and HL-60 cells, which are human promyelocytic cells that can be stimulated by PMA to form an active NADPH oxidase complex, were stimulated with water, 100 µM ATP, or 20 µM PMA for 5 min. Membrane proteins were isolated from these cells and used to measure superoxide production through cytochrome C reduction. In CHO cells, neither ATP nor PMA treatment produced any superoxide, confirming that these cells lack the endogenous machinery to form an active NADPH oxidase complex (Figure 7B). In contrast, in control HL-60 cells, neither water nor ATP was able to stimulate superoxide production, while PMA-treated cells showed a steady increase over time (Figure 7C). Therefore the ROS involved in ATP-stimulated TGF- shedding in CHO cells must come from another source. We also tested the involvement of the NADPH oxidase complex on endogenous shedding from EC-4 mouse fibroblasts. In these cells, apocynin also had no effect on ATP-stimulated TGF- shedding at any concentration tested (Supplemental Figure S4B), indicating an NADPH-oxidase independent mechanism of ROS production following P2Y receptor stimulation.
|
shedding, we examined two independent inhibitors of the electron transport chain: rotenone, which inhibits transfer of electrons from iron-sulfur centers to ubiquinone in complex I, and myxothiazol, which inhibits transfer of electrons between cytochrome b and cytochrome c1 in complex III. Treatment with myxothiazol resulted in a consistent, significant decrease in shedding compared with ATP alone (Figure 8A). The complex I inhibitor rotenone blocked the majority of ATP-stimulated TGF- shedding (Figure 8B). These inhibitors were individually unable to inhibit shedding to the same extent as the ROS scavenger NAC. However, when used in tandem, rotenone and myxothiazol reduced shedding as effectively as NAC (Figure 8C). Confirming the relevance of this pathway in endogenous ATP–stimulated shedding of TGF-, rotenone and myxothiazol in combination also inhibited shedding to the same level as NAC in mouse EC-4 cells (Supplemental Figure S4A). None of these inhibitors had any effect on cell viability or AP enzyme activity (Supplemental Table 1 and Supplemental Figure 3B), ruling out trivial effects on the assay system. These results strongly suggest that ROS produced in combination from mitochondrial complex I and complex III are an essential component of the regulatory pathway for GPCR-induced TGF- shedding.
|
shedding.
Increases in calcium concentration due to ATP activation of purinergic receptors occur rapidly through G-protein coupling to calcium channels (Abbracchio et al., 2006). Calcium is a known effector of mitochondrial function, including increasing production of ROS (Brookes et al., 2004). This suggests that calcium signaling could be an early step in ATP activation of TACE-dependent shedding and could occur upstream of mitochondrial ROS production. This is supported by our results that show that calcium chelation by EGTA was unable to inhibit hydrogen peroxide–dependent TGF- shedding (Figure 5D). To test whether calcium-induced TGF- shedding was dependent on mitochondrial ROS we examined ROS inhibitor effects on A23187
[GenBank]
-stimulated shedding. The ROS scavenger NAC partially inhibited this shedding event (Figure 8E). Mitochondrial inhibitors myxothiazol and rotenone individually and together also partially inhibited A23187
[GenBank]
-induced shedding as did treatment with both inhibitors. These results further support our hypothesis that ATP-induced calcium mobilization contributes to increases in mitochondrial ROS and activation of TACE-dependent TGF- shedding.
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
; Ohtsu et al., 2006). This study examines a ROS-dependent transactivation pathway from the P2Y family of GPCRs that utilizes TACE/ADAM17 metalloprotease activity to cleave TGF-, followed by activation of EGFR. Our goal was to begin to elucidate the signaling events leading to TACE-mediated growth factor cleavage after stimulation with GPCR agonists. We report here the first observation of mitochondrially produced ROS as a critical step in regulated growth factor proteolysis.
Extracellular nucleotides such as ATP are released from many cell types in response to stress and mechanical or biochemical stimulation (Abbracchio et al., 2006) and can function as a paracrine signal for the P2Y family of purinergic receptors. In this study we identified a P2Y-dependent pathway leading to TGF- shedding and transactivation of EGFR. Prominent expression of P2Y2 in mouse cells, along with the lack of detectable P2Y4 expression in CHO cells, suggests a favored role for the P2Y2 subtype in this transactivation pathway. P2Y2 receptors have been previously linked to EGFR transactivation in PC12 cells and rat fibroblasts (Soltoff, 1998; Soltoff et al., 1998), consistent with the ability of these receptors to stimulate proliferation of cells such as epidermal keratinocytes (Greig et al., 2003) and smooth muscle cells (Wilden et al., 1998). P2Y2 receptors are also important for regulating ion channel function in epithelial cells (Abbracchio et al., 2006). Activation of P2Y2 can promote mucin release in airway cells, an event that has also been tied to TACE-dependent TGF- shedding and EGFR transactivation (Yerxa et al., 2002; Shao et al., 2003). Extracellular release of ATP is also a hallmark of epithelial wound healing. In vitro wounding experiments utilize EGFR transactivation by purinergic receptors to initiate migration and healing of epithelial cells (Boucher et al., 2007; Yin et al., 2007). Thus, elucidation of the mechanisms responsible for regulating EGFR transactivation pathways could have important physiological and therapeutic implications.
A recent study showed that ATP stimulation through P2Y2 receptors led to both ADAM10 and TACE metalloprotease activity toward amyloid precursor protein in astrocytoma cells (Camden et al., 2005). Here, we show that ATP stimulated endogenous TACE-dependent TGF- shedding in mouse fibroblasts and AP-TGF- shedding in CHO cells, consistent with previous evidence for TACE as the major constitutive and PMA-stimulated TGF- sheddase (Peschon et al., 1998; Hinkle et al., 2004; Sahin et al., 2004; Horiuchi et al., 2007). A variety of studies have implicated roles for other ADAM proteases in TGF- processing under specific conditions (Merlos-Suarez et al., 2001; Hinkle et al., 2003; Schafer et al., 2004b; Horiuchi et al., 2007), including for ADAM10 in the absence of TACE (Le Gall et al., 2009). Although ADAM10 activity may explain the minor residual shedding in both TACE-deficient EC-2 mouse and CHO cells, the significant loss of TGF- release and of EGFR transactivation associated with TACE deficiency, or knockdown by siRNA, together with restoration of shedding by TACE in EC-2 and CHO M2 cells, indicates that TACE is the predominant TGF- sheddase in this model.
There is a high degree of correlation between ATP-dependent TGF- shedding and EGFR transactivation in the mouse fibroblast system that we tested. However, this does not exclude other EGFR ligands from involvement in ATP-induced receptor activation in this system. Other transactivation models have shown TACE-dependent cleavage of AR and HB-EGF to be important for EGFR phosphorylation as well (Hart et al., 2004; Schafer et al., 2004a). It is therefore possible that in the system we tested, additional EGF family members are also shed in response to ATP stimulation and this may contribute to the transactivation that we observed.
In this study, we undertook a further examination of the signaling pathways leading to TACE activity. ROS have recently gained attention as second messengers in GPCR signaling pathways (Forman et al., 2002). P2Y family receptors have been specifically linked to ROS production in eosinophils and prostate tumor cells (Ferrari et al., 2000; Sauer et al., 2001). Cellular ROS have also been implicated in EGFR transactivation initiated via other GPCRs (Wang et al., 2000; Zhang et al., 2004; Stoeck et al., 2006). We used hydrogen peroxide and the ROS scavenger NAC to establish a critical role for ROS in P2Y receptor-dependent TGF- shedding. These observations are also consistent with other reports that exogenous hydrogen peroxide can stimulate the shedding of TGF- and other EGF family members such as HB-EGF, AR, and betacellulin (Lemjabbar et al., 2003; Fischer et al., 2004; Kim et al., 2005; Shao and Nadel, 2005; Stoeck et al., 2006).
The cytoplasmic NADPH oxidase complex has been implicated as the main source of cellular ROS in a number of EGFR transactivation pathways. A role for NADPH oxidase-derived ROS has been reported for GPCR agonists ATP, angiotensin II, phenylephrine, endothelin-1, and tobacco smoke (Ushio-Fukai et al., 2001; Lemjabbar et al., 2003; Schafer et al., 2004a; Zhang et al., 2004; Chen et al., 2006; Stoeck et al., 2006; Boots et al., 2009), for UV-induced signaling (Singh et al., 2009), and for the global activator of shedding, PMA (Shao and Nadel, 2005). In addition to inhibitor studies, a role for NADPH oxidase-derived ROS in EGFR transactivation was identified using induced phosphorylation of p47 (Pietri et al., 2005), a key subunit of NADPH oxidase, and also the more stringent knockdown of the NADPH oxidase subunits Duox1 or p47phox (Shao and Nadel, 2005; Chen et al., 2006). ATP signaling through purinergic receptors has also been demonstrated to activate Duox1/Nox in airway epithelial cells (Wesley et al., 2007; Boots et al., 2009). We investigated the involvement of the NADPH oxidase complex in our system using the inhibitor apocynin, which directly blocks complex formation. We observed that at apocynin concentrations ranging from 30 µM up to 1 mM, ATP-induced TGF- shedding was unaffected. Furthermore, we found no evidence of an active NADPH oxidase complex in CHO cells, supporting previous findings in the literature (Takeya et al., 2003). These results rule out the NADPH oxidase complex as a component of this pathway.
To further examine the source of ROS for regulation of P2Y-dependent TGF- shedding, we used specific inhibitors of complex I and complex III of the mitochondrial electron transport chain to gauge the contributions of mitochondrial ROS. The inhibitors myxothiazol and rotenone partially decreased shedding when used individually, but when used together inhibited fully, as did the general ROS scavenger NAC, indicating that the functional ROS generated in this pathway can be entirely attributed to mitochondrial sources. In mouse cells, NAC and the combination of rotenone and myxothiazol also inhibited ATP-stimulated shedding to the same extent, indicating the requirement for mitochondrial ROS in this cell type as well. Mitochondrial ROS have been implicated in aging and cell fate, cell responses to hypoxia and many aspects of cardiovascular biology (Zhang and Gutterman, 2007). Hydrogen peroxide–induced EGFR activation can depend on mitochondrial ROS (Chen et al., 2004). Induction of mitochondrial ROS as a downstream signal of EGFR, estrogen, and NMDA receptors reveals a role for mitochondrial ROS as signaling messengers for important cellular receptors (Krieg et al., 2004; Felty et al., 2005; Hao et al., 2006; Lee et al., 2006; Duan et al., 2007). Here, we present evidence for mitochondrial ROS as a signaling intermediate for regulation of P2Y-induced, TACE-dependent growth factor shedding. This novel observation of mitochondrial ROS in regulating shedding reveals another possible mechanism that cells may use to specify certain pathways for responding to various stimuli.
ROS-mediated TACE-dependent shedding may also involve other signaling molecules that have been implicated in EGFR transactivation. Increases in intracellular calcium concentrations are a well-characterized result of GPCR activation (Werry et al., 2003), and this extracellular calcium influx can subsequently stimulate ROS-induced signaling (Sauer et al., 2001; Ushio-Fukai et al., 2001; Mifune et al., 2005; Pines et al., 2005; Watanabe et al., 2006). In our CHO AP-TGF- cells, TACE-dependent TGF- shedding was stimulated by the ionophore A23187, and inhibited by calcium chelation with BAPTA-AM and EGTA. Calcium chelation had little to no effect on hydrogen peroxide–stimulated shedding, suggesting that the calcium signal preceded ROS production in our system. This was further supported by the blocking of A23187-induced TGF- shedding with the mitochondrial inhibitors used to block ATP-induced shedding. In contrast to ATP, A23187-induced shedding was only partially inhibited by the ROS scavenger NAC and the mitochondrial inhibitors myxothiazol and rotenone. This may be explained by the previous observation in CHO-BQ1 cells that A23187 induced higher levels of intracellular calcium than ATP treatment (Marcet et al., 2003). The calcium ionophore-induced residual shedding seen in M2 cells and in NAC-treated wild-type CHO cells may indicate a ROS- and TACE- independent shedding event involving ADAM10 such as that shown in mouse Tace–/– fibroblasts (Horiuchi et al., 2007; Le Gall et al., 2009). Calcium signaling is a well-known effector of mitochondrial function and mitochondrial ROS production (Brookes et al., 2004). Although the mechanism is still under investigation, the observations here demonstrate a role for calcium signaling upstream of required mitochondrial ROS.
PKC has been widely implicated in regulation of ectodomain shedding based on the well-established ability of phorbol esters to stimulate TACE-dependent TGF- shedding (Fan et al., 2003; Hinkle et al., 2004; Sahin et al., 2004; Horiuchi et al., 2007). Previous reports implicated PKC isoforms in TACE phosphorylation and activation (Alfa Cisse et al., 2008; Reddy et al., 2008). In our study, treatment of cells with the broad spectrum PKC inhibitor BIM-1 (GF109203X) did not affect ATP-stimulated TGF- release. Src family kinases have also been implicated in several GPCR-mediated transactivation cascades upstream of growth factor shedding, including the direct association of Src with TACE, that led to TACE phosphorylation and increased shedding of amphiregulin (Zhang et al., 2006) or with P2Y2 receptors via SH3 motifs (Liu et al., 2004). There is also evidence suggesting ROS may modulate Src function in transactivation schemes (Ushio-Fukai et al., 2001; Zhuang and Schnellmann, 2004). In contrast, Camden et al. (2005) demonstrated Src-independent activation of TACE and ADAM10 after ATP stimulation. We also show that ATP-dependent activation of TGF- shedding is independent of the PP2-sensitive Src family members Src, Fyn, Hck, and Lck. Likewise ERK/MAPK pathways have been implicated in activation of TACE-dependent growth factor shedding by serum, stress, VEGF-A, ATP, and other stimuli (Fan and Derynck, 1999; Gechtman et al., 1999; Umata et al., 2001; Gschwind et al., 2003; Fischer et al., 2004; Zhuang and Schnellmann, 2004; Swendeman et al., 2008; Yin and Yu, 2009) possibly by phosphorylation of the cytoplasmic domain of TACE (Soond et al., 2005). We found no evidence for ERK/MAPK signals playing a regulatory role in TGF- shedding in our model. This further underscores the selective use of available regulatory pathways for particular substrates or physiological events.
The mechanism of TACE activation remains unclear. However, the large number of agonists and substrates utilizing ROS strongly suggest ROS are common regulators of TACE activity. ROS may cause direct modifications of TACE in response to stimulation. The prodomain of TACE can bind to a cysteine in the metalloprotease active site to inhibit cleavage (Seals and Courtneidge, 2003; Li et al., 2008). ROS have been shown to oxidize the cysteine thiol group thereby relieving the inhibition of the metalloprotease activity (Zhang et al., 2001). TACE also contains an extracellular cysteine-rich domain including two cysteine residues previously shown to be important in TACE functionality (Li and Fan, 2004) and specifically for hydrogen peroxide–induced, TACE-dependent L-selectin shedding in neutrophils (Wang et al., 2009). These could be potential targets for ROS-dependent modifications. ROS may also indirectly activate TACE by alteration and modification of effector proteins. Demonstrated phosphorylation of TACE by interaction with ERK (Soond et al., 2005), one of several PKC isoforms (Roghani et al., 1999; Alfa Cisse et al., 2008; Reddy et al., 2008), or with phosphoinositide-dependent kinase-1 (Zhang et al., 2006) could all be regulated by ROS modifications of these kinases or by the negative regulation of phosphatases. Modifications by ROS on scaffold proteins that are involved in substrate selection or presentation is another potential regulatory mechanism. The noncatalytic Eve-1 protein is thought to regulate EGF-like growth factor shedding in response to PMA (Tanaka et al., 2004), whereas NRDc enhances shedding of the TACE substrates HB-EGF (Nishi et al., 2006) and TNF- (Hiraoka et al., 2008). Most recently a tripartite complex between DLG1/SAP97, TACE and proTGF- was identified, where both knockdown and overexpression of DLG1 led to inhibition of PMA-induced TGF- shedding (Surena et al., 2009). Lastly, TACE activity regulated by localization in membrane subdomains (Tellier et al., 2006) could be affected by ROS modifications of lipids and membrane components (Zmijewski et al., 2005). By altering the membrane and creating or abolishing subdomains, protein localization complexes and enzyme–substrate interactions could all be influenced. Thus, the ability of ROS to freely diffuse throughout the cell and their high reactivity with proteins and membranes allow many different possible regulatory mechanisms.
In summary, we present evidence for the first time of mitochondrially derived ROS regulation of ATP-stimulated, TACE-dependent shedding of TGF-. Because TGF- signals act exclusively through EGFR, this may represent one mechanism to maintain the specificity of biochemical signaling pathways. A recent report examining the existence of multiple transactivating mechanisms in a single cell highlights the need for specific pathways for different transactivation schemes (Rodland et al., 2008). The discovery of mitochondrial ROS as a novel, key regulatory intermediate in TACE-dependent shedding of TGF- reveals an additional mechanism for regulating the complexity of signaling through common cellular pathways.
|
ACKNOWLEDGMENTS |
|---|
|
Footnotes |
|---|
||Present address: The Office of the Vice President for Research, University of Georgia, Athens, GA 30602. 
Address correspondence to: Susan Wohler Sunnarborg (susan_sunnarborg{at}med.unc.edu).
Abbreviations used: ADAM, a disintegrin and metalloprotease; AR, amphiregulin; BIM-I, bisindolylmaleimide I; DPI, diphenyleneiodonium; EGF, epidermal growth factor; EGFR, epidermal growth factor receptor; GPCR, G-protein–coupled receptor; HB-EGF, heparin-binding growth factor; MAPK, mitogen-activated protein kinase; NAC, N-acetyl-L-cysteine; PKC, protein kinase C; PMA, phorbol 12-myristate 13-acetate; RIA, radioimmunoassay; ROS, reactive oxygen species; TACE, tumor necrosis factor-–converting enzyme; TGF-, transforming growth factor-.
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Alfa Cisse, M., Louis, K., Braun, U., Mari, B., Leitges, M., Slack, B. E., Fisher, A., Auberger, P., Checler, F., and Vincent, B. (2008). Isoform-specific contribution of protein kinase C to prion processing. Mol. Cell Neurosci 39, 400–410.[CrossRef][Medline]
Arribas, J., and Massagué, J. (1995). Transforming growth factor- and β-amyloid precursor protein share a secretory mechanism. J. Cell Biol 128, 433–441.
Aslan, M., and Freeman, B. A. (2002). Oxidases and oxygenases in regulation of vascular nitric oxide signaling and inflammatory responses. Immunol. Res 26, 107–118.[CrossRef][Medline]
Blobel, C. P. (2005). ADAMs: key components in EGFR signalling and development. Nat. Rev. Mol. Cell Biol 6, 32–43.[CrossRef][Medline]
Boots, A. W., Hristova, M., Kasahara, D. I., Haenen, G. R., Bast, A., and van der Vliet, A. (2009). ATP-mediated activation of the NADPH oxidase DUOX1 mediates airway epithelial responses to bacterial stimuli. J. Biol. Chem 284, 17858–17867.
Borrell-Pages, M., Rojo, F., Albanell, J., Baselga, J., and Arribas, J. (2003). TACE is required for the activation of the EGFR by TGF-alpha in tumors. EMBO J 22, 1114–1124.[CrossRef][Medline]
Borroto, A., Ruiz-Paz, S., de la Torre, T. V., Borrell-Pages, M., Merlos-Suarez, A., Pandiella, A., Blobel, C. P., Baselga, J., and Arribas, J. (2003). Impaired trafficking and activation of tumor necrosis factor-alpha-converting enzyme in cell mutants defective in protein ectodomain shedding. J. Biol. Chem 278, 25933–25939.
Boucher, I., Yang, L., Mayo, C., Klepeis, V., and Trinkaus-Randall, V. (2007). Injury and nucleotides induce phosphorylation of epidermal growth factor receptor: MMP and HB-EGF dependent pathway. Exp. Eye Res 85, 130–141.[CrossRef][Medline]
Brachmann, R., Lindquist, P. B., Nagashima, M., Kohr, W., Lipari, T., Napier, M., and Derynck, R. (1989). Transmembrane TGF-alpha precursors activate EGF/TGF-alpha receptors. Cell 56, 691–700.[CrossRef][Medline]
Brandes, R. P., and Kreuzer, J. (2005). Vascular NADPH oxidases: molecular mechanisms of activation. Cardiovasc. Res 65, 16–27.
Brookes, P. S., Yoon, Y., Robotham, J. L., Anders, M. W., and Sheu, S. S. (2004). Calcium, ATP, and ROS: a mitochondrial love-hate triangle. Am. J. Physiol. Cell Physiol 287, C817–C833.
Camden, J. M., Schrader, A. M., Camden, R. E., Gonzalez, F. A., Erb, L., Seye, C. I., and Weisman, G. A. (2005). P2Y2 nucleotide receptors enhance -secretase-dependent amyloid precursor protein processing. J. Biol. Chem 280, 18696–18702.
Chen, C. H. et al. (2006). Reactive oxygen species generation is involved in epidermal growth factor receptor transactivation through the transient oxidization of Src homology 2-containing tyrosine phosphatase in endothelin-1 signaling pathway in rat cardiac fibroblasts. Mol. Pharmacol 69, 1347–1355.
Chen, J., Chen, J. K., Falck, J. R., Anjaiah, S., Capdevila, J. H., and Harris, R. C. (2007). Mitogenic activity and signaling mechanism of 2-(14,15-epoxyeicosatrienoyl)glycerol, a novel cytochrome p450 arachidonate metabolite. Mol. Cell. Biol 27, 3023–3034.
Chen, K., Thomas, S. R., Albano, A., Murphy, M. P., and Keaney, J. F., Jr. (2004). Mitochondrial function is required for hydrogen peroxide-induced growth factor receptor transactivation and downstream signaling. J. Biol. Chem 279, 35079–35086.
Cross, J. V., and Templeton, D. J. (2006). Regulation of signal transduction through protein cysteine oxidation. Antioxid. Redox Signal 8, 1819–1827.[CrossRef][Medline]
Dikalov, S., Griendling, K. K., and Harrison, D. G. (2007). Measurement of reactive oxygen species in cardiovascular studies. Hypertension 49, 717–727.
Dong, J., Opresko, L. K., Dempsey, P. J., Lauffenburger, D. A., Coffey, R. J., and Wiley, H. S. (1999). Metalloprotease-mediated ligand release regulates autocrine signaling through the epidermal growth factor receptor. Proc. Natl. Acad. Sci. USA 96, 6235–6240.
Duan, Y., Gross, R. A., and Sheu, S. S. (2007). Ca2+-dependent generation of mitochondrial reactive oxygen species serves as a signal for poly(ADP-ribose) polymerase-1 activation during glutamate excitotoxicity. J. Physiol 585, 741–758.
Fan, H., and Derynck, R. (1999). Ectodomain shedding of TGF-alpha and other transmembrane proteins is induced by receptor tyrosine kinase activation and MAP kinase signaling cascades. EMBO J 18, 6962–6972.[CrossRef][Medline]
Fan, H., Turck, C. W., and Derynck, R. (2003). Characterization of growth factor-induced serine phosphorylation of tumor necrosis factor- converting enzyme and of an alternatively translated polypeptide. J. Biol. Chem 278, 18617–18627.
Felty, Q., Xiong, W. C., Sun, D., Sarkar, S., Singh, K. P., Parkash, J., and Roy, D. (2005). Estrogen-induced mitochondrial reactive oxygen species as signal-transducing messengers. Biochemistry 44, 6900–6909.[CrossRef][Medline]
Ferrari, D., Idzko, M., Dichmann, S., Purlis, D., Virchow, C., Norgauer, J., Chiozzi, P., Di Virgilio, F., and Luttmann, W. (2000). P2 purinergic receptors of human eosinophils: characterization and coupling to oxygen radical production. FEBS Lett 486, 217–224.[CrossRef][Medline]
Fischer, O. M., Hart, S., Gschwind, A., Prenzel, N., and Ullrich, A. (2004). Oxidative and osmotic stress signaling in tumor cells is mediated by ADAM proteases and heparin-binding epidermal growth factor. Mol. Cell. Biol 24, 5172–5183.
Forman, H. J., Torres, M., and Fukuto, J. (2002). Redox signaling. Mol. Cell Biochem. 234– 235, 49–62.[CrossRef]
Frank, G. D., and Eguchi, S. (2003). Activation of tyrosine kinases by reactive oxygen species in vascular smooth muscle cells: significance and involvement of EGF receptor transactivation by angiotensin II. Antioxid. Redox Signal 5, 771–780.[CrossRef][Medline]
Gechtman, Z., Alonso, J. L., Raab, G., Ingber, D. E., and Klagsbrun, M. (1999). The shedding of membrane-anchored heparin-binding epidermal-like growth factor is regulated by the Raf/mitogen-activated protein kinase cascade and by cell adhesion and spreading. J. Biol. Chem 274, 28828–28835.
Greig, A. V., James, S. E., McGrouther, D. A., Terenghi, G., and Burnstock, G. (2003). Purinergic receptor expression in the regeneration epidermis in a rat model of normal and delayed wound healing. Exp. Dermatol 12, 860–871.[CrossRef][Medline]
Gschwind, A., Hart, S., Fischer, O. M., and Ullrich, A. (2003). TACE cleavage of proamphiregulin regulates GPCR-induced proliferation and motility of cancer cells. EMBO J 22, 2411–2421.[CrossRef][Medline]
Hao, L., Nishimura, T., Wo, H., and Fernandez-Patron, C. (2006). Vascular responses to alpha1-adrenergic receptors in small rat mesenteric arteries depend on mitochondrial reactive oxygen species. Arterioscler. Thromb. Vasc. Biol 26, 819–825.
Hart, S., Fischer, O. M., and Ullrich, A. (2004). Cannabinoids induce cancer cell proliferation via tumor necrosis factor alpha-converting enzyme (TACE/ADAM17)-mediated transactivation of the epidermal growth factor receptor. Cancer Res 64, 1943–1950.
Hinkle, C. L., Mohan, M. J., Lin, P., Yeung, N., Rasmussen, F., Milla, M. E., and Moss, M. L. (2003). Multiple metalloproteinases process protransforming growth factor- (proTGF-). Biochemistry 42, 2127–2136.[CrossRef][Medline]
Hinkle, C. L., Sunnarborg, S. W., Loiselle, D., Parker, C. E., Stevenson, M., Russell, W. E., and Lee, D. C. (2004). Selective roles for tumor necrosis factor -converting enzyme/ADAM17 in the shedding of the epidermal growth factor receptor ligand family: the juxtamembrane stalk determines cleavage efficiency. J. Biol. Chem 279, 24179–24188.
Hiraoka, Y., Yoshida, K., Ohno, M., Matsuoka, T., Kita, T., and Nishi, E. (2008). Ectodomain shedding of TNF-alpha is enhanced by nardilysin via activation of ADAM proteases. Biochem. Biophys. Res. Commun 370, 154–158.[CrossRef][Medline]
Hoffmann, C., Moro, S., Nicholas, R. A., Harden, T. K., and Jacobson, K. A. (1999). The role of amino acids in extracellular loops of the human P2Y1 receptor in surface expression and activation processes. J. Biol. Chem 274, 14639–14647.
Holbro, T., and Hynes, N. E. (2004). ErbB receptors: directing key signaling networks throughout life. Annu. Rev. Pharmacol. Toxicol 44, 195–217.[CrossRef][Medline]
Horiuchi, K., Le Gall, S., Schulte, M., Yamaguchi, T., Reiss, K., Murphy, G., Toyama, Y., Hartmann, D., Saftig, P., and Blobel, C. P. (2007). Substrate selectivity of epidermal growth factor-receptor ligand sheddases and their regulation by phorbol esters and calcium influx. Mol. Biol. Cell 18, 176–188.
Iredale, P. A., and Hill, S. J. (1993). Increases in intracellular calcium via activation of an endogenous P2-purinoceptor in cultured CHO-K1 cells. Br. J. Pharmacol 110, 1305–1310.[Medline]
Iwamoto, R. et al. (2003). Heparin-binding EGF-like growth factor and ErbB signaling is essential for heart function. Proc. Natl. Acad. Sci. USA 100, 3221–3226.
Izumi, Y. et al. (1998). A metalloprotease-disintegrin, MDC9/meltrin-/ADAM9 and PKC are involved in TPA-induced ectodomain shedding of membrane-anchored heparin-binding EGF-like growth factor. EMBO J 17, 7260–7272.[CrossRef][Medline]
Jackson, L. F., Qiu, T. H., Sunnarborg, S. W., Chang, A., Zhang, C., Patterson, C., and Lee, D. C. (2003). Defective valvulogenesis in HB-EGF and TACE-null mice is associated with aberrant BMP signaling. EMBO J 22, 2704–2716.[CrossRef][Medline]
Kim, J., Adam, R. M., and Freeman, M. R. (2005). Trafficking of nuclear heparin-binding epidermal growth factor-like growth factor into an epidermal growth factor receptor-dependent autocrine loop in response to oxidative stress. Cancer Res 65, 8242–8249.
Krieg, T., Cui, L., Qin, Q., Cohen, M. V., and Downey, J. M. (2004). Mitochondrial ROS generation following acetylcholine-induced EGF receptor transactivation requires metalloproteinase cleavage of proHB-EGF. J. Mol. Cell Cardiol 36, 435–443.[CrossRef][Medline]
Lautrette, A., Li, S., Alili, R., Sunnarborg, S. W., Burtin, M., Lee, D. C., Friedlander, G., and Terzi, F. (2005). Angiotensin II and EGF receptor cross-talk in chronic kidney diseases: a new therapeutic approach. Nat. Med 11, 867–874.[CrossRef][Medline]
Le Gall, S. M., Bobe, P., Reiss, K., Horiuchi, K., Niu, X. D., Lundell, D., Gibb, D. R., Conrad, D., Saftig, P., and Blobel, C. P. (2009). ADAMs 10 and 17 represent differentially regulated components of a general shedding machinery for membrane proteins such as transforming growth factor , L-selectin, and tumor necrosis factor . Mol. Biol. Cell 20, 1785–1794.
Lee, D. C., Hinkle, C. L., Jackson, L. F., Li, S., and Sunnarborg, S. W. (2003). EGF family ligands. In: The Cytokine Handbook, A. W. Thomson and M. T. Lotze, London: Elsevier Science, 959–987.
Lee, S. B., Bae, I. H., Bae, Y. S., and Um, H. D. (2006). Link between mitochondria and NADPH oxidase 1 isozyme for the sustained production of reactive oxygen species and cell death. J. Biol. Chem 281, 36228–36235.
Lemjabbar, H., Li, D., Gallup, M., Sidhu, S., Drori, E., and Basbaum, C. (2003). Tobacco smoke-induced lung cell proliferation mediated by tumor necrosis factor alpha-converting enzyme and amphiregulin. J. Biol. Chem 278, 26202–26207.
Li, X., and Fan, H. (2004). Loss of ectodomain shedding due to mutations in the metalloprotease and cysteine-rich/disintegrin domains of the tumor necrosis factor- converting enzyme (TACE). J. Biol. Chem 279, 27365–27375.
Li, X., Yan, Y., Huang, W., Yang, Y., Wang, H., and Chang, L. (2008). The regulation of TACE catalytic function by its prodomain. Mol. Biol. Rep 36, 641–651.[Medline]
Liu, J., Liao, Z., Camden, J., Griffin, K. D., Garrad, R. C., Santiago-Perez, L. I., Gonzalez, F. A., Seye, C. I., Weisman, G. A., and Erb, L. (2004). Src homology 3 binding sites in the P2Y2 nucleotide receptor interact with Src and regulate activities of Src, proline-rich tyrosine kinase 2, and growth factor receptors. J. Biol. Chem 279, 8212–8218.
Luttrell, D. K., and Luttrell, L. M. (2004). Not so strange bedfellows: G-protein-coupled receptors and Src family kinases. Oncogene 23, 7969–7978.[CrossRef][Medline]
Marcet, B., Chappe, V., Delmas, P., Gola, M., and Verrier, B. (2003). Negative regulation of CFTR activity by extracellular ATP involves P2Y2 receptors in CFTR-expressing CHO cells. J. Membr. Biol 194, 21–32.[CrossRef][Medline]
McCole, D. F., Keely, S. J., Coffey, R. J., and Barrett, K. E. (2002). Transactivation of the epidermal growth factor receptor in colonic epithelial cells by carbachol requires extracellular release of transforming growth factor-. J. Biol. Chem 277, 42603–42612.
Merlos-Suarez, A., Ruiz-Paz, S., Baselga, J., and Arribas, J. (2001). Metalloprotease-dependent protransforming growth factor- ectodomain shedding in the absence of tumor necrosis factor--converting enzyme. J. Biol. Chem 276, 48510–48517.
Mifune, M. et al. (2005). G protein coupling and second messenger generation are indispensable for a metalloprotease-dependent HB-EGF shedding through angiotensin II type-1 receptor. J. Biol. Chem 280, 26592–26599.
Nagano, O., Murakami, D., Hartmann, D., De Strooper, B., Saftig, P., Iwatsubo, T., Nakajima, M., Shinohara, M., and Saya, H. (2004). Cell-matrix interaction via CD44 is independently regulated by different metalloproteinases activated in response to extracellular Ca(2+) influx and PKC activation. J. Cell Biol 165, 893–902.
Nishi, E., Hiraoka, Y., Yoshida, K., Okawa, K., and Kita, T. (2006). Nardilysin enhances ectodomain shedding of heparin-binding epidermal growth factor-like growth factor through activation of tumor necrosis factor--converting enzyme. J. Biol. Chem 281, 31164–31172.
Ohtsu, H., Dempsey, P. J., and Eguchi, S. (2006). ADAMs as mediators of EGF receptor transactivation by G protein-coupled receptors. Am. J. Physiol. Cell Physiol 291, C1–C10.
Peschon, J. J. et al. (1998). An essential role for ectodomain shedding in mammalian development. Science 282, 1281–1284.
Pierce, K. L., Tohgo, A., Ahn, S., Field, M. E., Luttrell, L. M., and Lefkowitz, R. J. (2001). Epidermal growth factor (EGF) receptor-dependent ERK activation by G protein-coupled receptors: a co-culture system for identifying intermediates upstream and downstream of heparin-binding EGF shedding. J. Biol. Chem 276, 23155–23160.
Pietri, M., Schneider, B., Mouillet-Richard, S., Ermonval, M., Mutel, V., Launay, J. M., and Kellermann, O. (2005). Reactive oxygen species-dependent TNF- converting enzyme activation through stimulation of 5-HT2B and -1D autoreceptors in neuronal cells. FASEB J 19, 1078–1087.
Pines, A., Perrone, L., Bivi, N., Romanello, M., Damante, G., Gulisano, M., Kelley, M. R., Quadrifoglio, F., and Tell, G. (2005). Activation of APE1/Ref-1 is dependent on reactive oxygen species generated after purinergic receptor stimulation by ATP. Nucleic Acids Res 33, 4379–4394.
Prenzel, N., Zwick, E., Daub, H., Leserer, M., Abraham, R., Wallasch, C., and Ullrich, A. (1999). EGF receptor transactivation by G-protein-coupled receptors requires metalloproteinase cleavage of proHB-EGF. Nature 402, 884–888.[Medline]
Reddy, A. B., Ramana, K. V., Srivastava, S., Bhatnagar, A., and Srivastava, S. K. (2008). Aldose reductase regulates high glucose-induced ectodomain shedding of TNF- via PKC- and TACE in vascular smooth muscle cells. Endocrinology 150, 63–74.[CrossRef][Medline]
Reddy, P., Slack, J. L., Davis, R., Cerretti, D. P., Kozlosky, C. J., Blanton, R. A., Shows, D., Peschon, J. J., and Black, R. A. (2000). Functional analysis of the domain structure of tumor necrosis factor- converting enzyme. J. Biol. Chem 275, 14608–14614.
Rodland, K. D., Bollinger, N., Ippolito, D., Opresko, L. K., Coffey, R. J., Zangar, R., and Wiley, H. S. (2008). Multiple mechanisms are responsible for transactivation of the epidermal growth factor receptor in mammary epithelial cells. J. Biol. Chem 283, 31477–31487.
Roghani, M. et al. (1999). Metalloprotease-disintegrin MDC9, intracellular maturation and catalytic activity. J. Biol. Chem 274, 3531–3540.
Russell, W. E., Dempsey, P. J., Sitaric, S., Peck, A. J., and Coffey, R. J., Jr. (1993). Transforming growth factor- (TGF-) concentrations increase in regenerating rat liver: evidence for a delayed accumulation of mature TGF . Endocrinology 133, 1731–1738.
Sahin, U., Weskamp, G., Kelly, K., Zhou, H. M., Higashiyama, S., Peschon, J., Hartmann, D., Saftig, P., and Blobel, C. P. (2004). Distinct roles for ADAM10 and ADAM17 in ectodomain shedding of six EGFR ligands. J. Cell Biol 164, 769–779.
Sahin, U., Weskamp, G., Zheng, Y., Chesneau, V., Horiuchi, K., and Blobel, C. P. (2006). A sensitive method to monitor ectodomain shedding of ligands of the epidermal growth factor receptor. Methods Mol. Biol 327, 99–113.[Medline]
Sanderson, M. P., Erickson, S. N., Gough, P. J., Garton, K. J., Wille, P. T., Raines, E. W., Dunbar, A. J., and Dempsey, P. J. (2005). ADAM10 mediates ectodomain shedding of the betacellulin precursor activated by p-aminophenylmercuric acetate and extracellular calcium influx. J. Biol. Chem 280, 1826–1837.
Sandgren, E. P., Luetteke, N. C., Palmiter, R. D., Brinster, R. L., and Lee, D. C. (1990). Overexpression of TGF- in transgenic mice: induction of epithelial hyperplasia, pancreatic metaplasia, and carcinoma of the breast. Cell 61, 1121–1135.[CrossRef][Medline]
Sauer, H., Klimm, B., Hescheler, J., and Wartenberg, M. (2001). Activation of p90RSK and growth stimulation of multicellular tumor spheroids are dependent on reactive oxygen species generated after purinergic receptor stimulation by ATP. FASEB J 15, 2539–2541.
Saxon, M. L., and Lee, D. C. (1999). Mutagenesis reveals a role for epidermal growth factor receptor extracellular subdomain IV in ligand binding. J. Biol. Chem 274, 28356–28362.
Schafer, B., Gschwind, A., and Ullrich, A. (2004a). Multiple G-protein-coupled receptor signals converge on the epidermal growth factor receptor to promote migration and invasion. Oncogene 23, 991–999.[CrossRef][Medline]
Schafer, B., Marg, B., Gschwind, A., and Ullrich, A. (2004b). Distinct ADAM metalloproteinases regulate G protein coupled receptor-induced cell proliferation and survival. J. Biol. Chem 279, 47929–47938.
Seals, D. F., and Courtneidge, S. A. (2003). The ADAMs family of metalloproteases: multidomain proteins with multiple functions. Genes Dev 17, 7–30.
Shao, M. X., and Nadel, J. A. (2005). Dual oxidase 1-dependent MUC5AC mucin expression in cultured human airway epithelial cells. Proc. Natl. Acad. Sci. USA 102, 767–772.
Shao, M. X., Ueki, I. F., and Nadel, J. A. (2003). Tumor necrosis factor -converting enzyme mediates MUC5AC mucin expression in cultured human airway epithelial cells. Proc. Natl. Acad. Sci. USA 100, 11618–11623.
Singh, B., Schneider, M., Knyazev, P., and Ullrich, A. (2009). UV-induced EGFR signal transactivation is dependent on proligand shedding by activated metalloproteases in skin cancer cell lines. Int. J. Cancer 124, 531–539.[CrossRef][Medline]
Soltoff, S. P. (1998). Related adhesion focal tyrosine kinase and the epidermal growth factor receptor mediate the stimulation of mitogen-activated protein kinase by the G-protein-coupled P2Y2 receptor. Phorbol ester or [Ca2+]i elevation can substitute for receptor activation. J. Biol. Chem 273, 23110–23117.[Medline]
Soltoff, S. P., Avraham, H., Avraham, S., and Cantley, L. C. (1998). Activation of P2Y2 receptors by UTP and ATP stimulates mitogen-activated kinase activity through a pathway that involves related adhesion focal tyrosine kinase and protein kinase C. J. Biol. Chem 273, 2653–2660.
Soond, S. M., Everson, B., Riches, D. W., and Murphy, G. (2005). ERK-mediated phosphorylation of Thr735 in TNF-converting enzyme and its potential role in TACE protein trafficking. J. Cell Sci 118, 2371–2380.
Sternlicht, M. D., Sunnarborg, S. W., Kouros-Mehr, H., Yu, Y., Lee, D. C., and Werb, Z. (2005). Mammary ductal morphogenesis requires paracrine activation of stromal EGFR via ADAM17-dependent shedding of epithelial amphiregulin. Development 132, 3923–3933.
Stoeck, A., Keller, S., Riedle, S., Sanderson, M. P., Runz, S., Le Naour, F., Gutwein, P., Ludwig, A., Rubinstein, E., and Altevogt, P. (2006). A role for exosomes in the constitutive and stimulus-induced ectodomain cleavage of L1 and CD44. Biochem. J 393, 609–618.[CrossRef][Medline]
Sunnarborg, S. W. et al. (2002). Tumor necrosis factor- converting enzyme (TACE) regulates epidermal growth factor receptor ligand availability. J. Biol. Chem 277, 12838–12845.
Surena, A. L., de Faria, G. P., Studler, J. M., Peiretti, F., Pidoux, M., Camonis, J., Chneiweiss, H., Formstecher, E., and Junier, M. P. (2009). DLG1/SAP97 modulates transforming growth factor alpha bioavailability. Biochim. Biophys. Acta 1793, 264–272.[Medline]
Swendeman, S., Mendelson, K., Weskamp, G., Horiuchi, K., Deutsch, U., Scherle, P., Hooper, A., Rafii, S., and Blobel, C. P. (2008). VEGF-A stimulates ADAM17-dependent shedding of VEGFR2 and crosstalk between VEGFR2 and ERK signaling. Circ. Res 103, 916–918.
Takeya, R., Ueno, N., Kami, K., Taura, M., Kohjima, M., Izaki, T., Nunoi, H., and Sumimoto, H. (2003). Novel human homologues of p47phox and p67phox participate in activation of superoxide-producing NADPH oxidases. J. Biol. Chem 278, 25234–25246.
Tanaka, M., Nanba, D., Mori, S., Shiba, F., Ishiguro, H., Yoshino, K., Matsuura, N., and Higashiyama, S. (2004). ADAM binding protein Eve-1 is required for ectodomain shedding of epidermal growth factor receptor ligands. J. Biol. Chem 279, 41950–41959.
Teixidó, J., Wong, S. T., Lee, D. C., and Massagué, J. (1990). Generation of transforming growth factor-alpha from the cell surface by an O-glycosylation-independent multistep process. J. Biol. Chem 265, 6410–6415.
Tellier, E., Canault, M., Rebsomen, L., Bonardo, B., Juhan-Vague, I., Nalbone, G., and Peiretti, F. (2006). The shedding activity of ADAM17 is sequestered in lipid rafts. Exp. Cell Res 312, 3969–3980.[CrossRef][Medline]
Teufelhofer, O., Weiss, R. M., Parzefall, W., Schulte-Hermann, R., Micksche, M., Berger, W., and Elbling, L. (2003). Promyelocytic HL60 cells express NADPH oxidase and are excellent targets in a rapid spectrophotometric microplate assay for extracellular superoxide. Toxicol. Sci 76, 376–383.
Thery, C., Amigorena, S., Raposo, G., and Clayton, A. (2006). Isolation and characterization of exosomes from cell culture supernatants and biological fluids. Curr. Protoc. Cell. Biol Chapter 3, Unit 3 22.
Tokumaru, S. et al. (2000). Ectodomain shedding of epidermal growth factor receptor ligands is required for keratinocyte migration in cutaneous wound healing. J. Cell Biol 151, 209–220.
Troyer, K. L., Luetteke, N. C., Saxon, M. L., Qiu, T. H., Xian, C. J., and Lee, D. C. (2001). Growth retardation, duodenal lesions, and aberrant ileum architecture in triple null mice lacking EGF, amphiregulin, and TGF-alpha. Gastroenterology 121, 68–78.[CrossRef][Medline]
Umata, T. et al. (2001). A dual signaling cascade that regulates the ectodomain shedding of heparin-binding epidermal growth factor-like growth factor. J. Biol. Chem 276, 30475–30482.
Ushio-Fukai, M., Griendling, K. K., Becker, P. L., Hilenski, L., Halleran, S., and Alexander, R. W. (2001). Epidermal growth factor receptor transactivation by angiotensin II requires reactive oxygen species in vascular smooth muscle cells. Arterioscler. Thromb. Vasc. Biol 21, 489–495.
Ushio-Fukai, M., Zafari, A. M., Fukui, T., Ishizaka, N., and Griendling, K. K. (1996). p22phox is a critical component of the superoxide-generating NADH/NADPH oxidase system and regulates angiotensin II-induced hypertrophy in vascular smooth muscle cells. J. Biol. Chem 271, 23317–23321.
Wang, D., Yu, X., Cohen, R. A., and Brecher, P. (2000). Distinct effects of N-acetylcysteine and nitric oxide on angiotensin II-induced epidermal growth factor receptor phosphorylation and intracellular Ca(2+) levels. J. Biol. Chem 275, 12223–12230.
Wang, Y., Herrera, A. H., Li, Y., Belani, K. K., and Walcheck, B. (2009). Regulation of mature ADAM17 by redox agents for L-selectin shedding. J. Immunol 182, 2449–2457.
Watanabe, N. et al. (2006). Activation of mitogen-activated protein kinases by lysophosphatidylcholine-induced mitochondrial reactive oxygen species generation in endothelial cells. Am. J. Pathol 168, 1737–1748.
Werry, T. D., Wilkinson, G. F., and Willars, G. B. (2003). Mechanisms of cross-talk between G-protein-coupled receptors resulting in enhanced release of intracellular Ca2+. Biochem. J 374, 281–296.[CrossRef][Medline]
Wesley, U. V., Bove, P. F., Hristova, M., McCarthy, S., and van der Vliet, A. (2007). Airway epithelial cell migration and wound repair by ATP-mediated activation of dual oxidase 1. J. Biol. Chem 282, 3213–3220.
Wilden, P. A., Agazie, Y. M., Kaufman, R., and Halenda, S. P. (1998). ATP-stimulated smooth muscle cell proliferation requires independent ERK and PI3K signaling pathways. Am. J. Physiol 275, H1209–H1215.[Medline]
Wong, S. T., Winchell, L. F., McCune, B. K., Earp, H. S., Teixidó, J., Massagué, J., Herman, B., and Lee, D. C. (1989). The TGF- precursor expressed on the cell surface binds to the EGF receptor on adjacent cells, leading to signal transduction. Cell 56, 495–506.[CrossRef][Medline]
Yan, Y., Shirakabe, K., and Werb, Z. (2002). The metalloprotease Kuzbanian (ADAM10) mediates the transactivation of EGF receptor by G protein-coupled receptors. J. Cell Biol 158, 221–226.
Yerxa, B. R. et al. (2002). Pharmacology of INS37217 [P(1)-(uridine 5')-P(4)- (2'-deoxycytidine 5')tetraphosphate, tetrasodium salt], a next-generation P2Y(2) receptor agonist for the treatment of cystic fibrosis. J. Pharmacol. Exp. Ther 302, 871–880.
Yin, J., Xu, K., Zhang, J., Kumar, A., and Yu, F. S. (2007). Wound-induced ATP release and EGF receptor activation in epithelial cells. J. Cell Sci 120, 815–825.
Yin, J., and Yu, F. S. (2009). ERK1/2 mediate wounding- and G-protein-coupled receptor ligands-induced EGFR activation via regulating ADAM17 and HB-EGF shedding. Invest. Ophthalmol. Vis. Sci 50, 132–139.
Zhang, D. X., and Gutterman, D. D. (2007). Mitochondrial reactive oxygen species-mediated signaling in endothelial cells. Am. J. Physiol. Heart Circ. Physiol 292, H2023–H2031.
Zhang, H., Chalothorn, D., Jackson, L. F., Lee, D. C., and Faber, J. E. (2004). Transactivation of epidermal growth factor receptor mediates catecholamine-induced growth of vascular smooth muscle. Circ. Res 95, 989–997.
Zhang, Q., Thomas, S. M., Lui, V. W., Xi, S., Siegfried, J. M., Fan, H., Smithgall, T. E., Mills, G. B., and Grandis, J. R. (2006). Phosphorylation of TNF- converting enzyme by gastrin-releasing peptide induces amphiregulin release and EGF receptor activation. Proc. Natl. Acad. Sci. USA 103, 6901–6906.
Zhang, Z., Oliver, P., Lancaster, J. J., Schwarzenberger, P. O., Joshi, M. S., Cork, J., and Kolls, J. K. (2001). Reactive oxygen species mediate tumor necrosis factor -converting, enzyme-dependent ectodomain shedding induced by phorbol myristate acetate. FASEB J 15, 303–305.
Zhuang, S., and Schnellmann, R. G. (2004). H2O2-induced transactivation of EGF receptor requires Src and mediates ERK1/2, but not Akt, activation in renal cells. Am. J. Physiol. Renal Physiol 286, F858–F865.
Zmijewski, J. W., Landar, A., Watanabe, N., Dickinson, D. A., Noguchi, N., and Darley-Usmar, V. M. (2005). Cell signalling by oxidized lipids and the role of reactive oxygen species in the endothelium. Biochem. Soc. Trans 33, 1385–1389.[CrossRef][Medline]
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
