Activation of Transforming Growth Factor-?1/p38/Smad3 Signaling in Stromal Cells Requires Reactive Oxygen Species–Mediated MMP-2 Activity Du
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《干细胞学杂志》
a Department of Pediatrics,
b Department of Microbiology, Immunology, and Cell Biology, and
c Mary Babb Randolph Cancer Center, Robert C. Byrd Health Sciences Center, West Virginia University, School of Medicine, Morgantown, West Virginia, USA
Key Words. Transforming growth factor-?1 ? Smad3 ? Reactive oxygen species ? MMP-2 ? Chemotherapy ? Microenvironment
Correspondence: Laura F. Gibson, Ph.D., P.O. Box 9214, Department of Pediatrics, School of Medicine, WV University, Morgantown, West Virginia, 26506, USA. Telephone: 304-293-5820; Fax: 304-293-4341; e-mail: lgibson@hsc.wvu.edu
ABSTRACT
The bone marrow microenvironment serves as the primary site of normal postnatal hematopoiesis and supports hematopoietic recovery after myelosuppressive chemotherapy or irradiation-induced injury of the immune system . Hematopoietic reconstitution requires efficient migration of transplanted stem/progenitor cells to the bone marrow and relocation to stromal cell niches in this microenvironment . The effects of preparative regimens on the marrow microenvironment remain an area requiring further investigation. The assumption that aggressive chemotherapy spares the bone marrow microenvironment grows increasingly more suspect because dose escalation of chemotherapy reveals unexpected problems with hematopoietic recovery . The dilemma remains maintaining efficacy of tumor eradication while reducing damage to the microenvironment.
Of the signaling molecules in the bone marrow microenvironment that may be involved in chemotherapy-induced bone marrow damage, transforming growth factor (TGF)-?1 is specifically noteworthy. TGF-?1 regulates a variety of biological responses, including angiogenesis, chemotaxis, cell-cycle progression, differentiation, and apoptosis of target cells in a context- and cell-specific manner . TGF-?1 is also involved in regulating extracellular matrix (ECM) remodeling, collagen gene expression, and degradation of matrix proteins during the processes of tissue injury and repair . Upregulated expression or activation of TGF-?1 at sites of injury is associated with proliferation of fibroblasts, progressive fibrosis, and subsequent organ dysfunction in diverse systems, including kidney, liver, and lung . In contrast to its promotion of mesenchymal cell proliferation and survival, TGF-?1 is a potent inhibitor of hematopoietic stem cell proliferation .
TGF-?1 is initially synthesized as a large precursor that is processed to a mature protein during secretion. After secretion, mature TGF-?1 (25 kD) noncovalently associates with its N-terminal pro-peptide, the 75-kD latency-associated protein (LAP) . The TGF-?1–LAP complex predominantly binds to a latent TGF-?1–binding protein (LTBP), which mediates deposition of the latent complex (230 and 195 kD) to the ECM . Release of mature TGF-?1 from the latent complex can be accomplished by different mechanisms, such as proteolytic cleavage of LAP by plasmin , deglycosylation of LAP , or interaction with thrombospondin-1 , platelet , or integrin 4, ?6 . After TGF-?1 ligand binding, TGF-?1 receptor II recruits and activates TGF-?1 receptor I, which in turn phosphorylates and activates the R-Smads, including Smad2 or Smad3 . Phosphorylated R-Smads homodimerize, form a transcriptional complex with Smad4, and translocate into the nucleus to regulate target gene expression .
Data presented in the current study suggest that the TGF-?1/p38/Smad3 signaling cascades are activated through reactive oxygen species (ROS)–mediated matrix metalloproteinase-2 (MMP-2) activity in bone marrow stromal cells during etoposide chemotherapy. Increased availability of active TGF-?1 has the potential to alter stromal cell function through regulation of diverse Smad-driven gene expression in stromal cells. Moreover, release of TGF-?1 from ECM during chemotherapy may also directly regulate growth and proliferation of transplanted hematopoietic stem cells. This in vitro model provides a setting in which we can further delineate the effects of chemotherapy on marrow stromal cells and evaluate the role of TGF-?1 in influencing hematopoietic recovery after transplantation.
MATERIALS AND METHODS
Chemotherapy Activates Smad3 Through Phosphorylation at Serines 433/435 in Human Bone Marrow–Derived Stromal Cells
To investigate the phosphorylation of Smad3 in stromal cells after chemotherapeutic stimulation, stromal cells were treated either for different times or with various concentrations of etoposide. Exposure of stromal cells to etoposide resulted in phosphorylation of Smad3 at serines 433/435 in a time-dependent (Fig. 1A) and dose-dependent (Fig. 1B) manner. Etoposide induced a rapid elevation of phospho-Smad3 signal as early as 30 minutes, which was sustained for approximately 6–7 hours (Fig. 1A). After the transient increase, phospho-Smad3 levels diminished for up to 24 hours in the presence of chemotherapy. Etoposide, melphalan, vincristine, daunorubicin, doxorubicin, 4-hydroperocyclophosphomide, and Ara-C induced Smad3 phosphorylation in stromal cells to varying degrees (Fig. 1C). Treatment of stromal cells with recombinant TGF-?1 served as a positive control and induced the most pronounced phosphorylation of Smad3. Total Smad3 protein remained unchanged and served as the lane loading control.
Figure 1. Chemotherapy activates Smad3 through phosphorylation at serines 433/435 in human bone marrow–derived stromal cells. Western blot analyses of HS-27A stromal cells treated with (A) 100 μM etoposide for the indicated time points, (B) various concentrations of etoposide for 1 hour, or (C) 3 ng/ml transforming growth factor -?1 (TGF-?1), 100 μM etoposide (VP-16), 200 μg/ml melphalan (Mel), 20 μg/ml vincristine (VCR), 100 μg/ml daunorubicin (DNR), 100 μM doxorubicin (DOX), 100 μg/ml 4-hydroperoxylcyclophosphamide (4-HC), or 100 μg/ml Ara-C for 1 hour. Membranes were probed with anti-phospho-Smad3 (P-Smad3, ser433/435) and then stripped and reprobed with total Smad3 (T-Smad3)–specific antibodies.
Chemotherapy-Induced Smad3 Phosphorylation Is Mediated by TGF-?1
To investigate the potential involvement of TGF-?1 in phosphorylation of Smad3 in bone marrow stromal cells during chemotherapy, HS-27A stromal cells were exposed to etoposide for different times or at various concentrations. Quantitative analysis of TGF-?1 by ELISA was performed using the cell supernatants after treatment. To better distinguish the free (active) and latent (total) TGF-?1 released, nonacidification and acidification of the stromal supernatants were simultaneously used before assay as described. Exposure of stromal cells to chemotherapy resulted in elevated TGF-?1 release from stromal cells both in a time-and dose-dependent manner (Fig. 2A). Chemotherapy rapidly induced the release of active and latent forms of TGF-?1 from stromal cell ECM. In our stromal cell model, active TGF-?1 constituted approximately 5%–9% of the total TGF-?1 pool during each treatment phase. Activation of TGF-?1 preceded phosphorylation of Smad3 with initial increases rapidly after etoposide treatment of 15 minutes and further elevation at 1 hour. Activation of TGF-?1 in stromal cells was transient, as longer than 1 hour exposure of stromal cells to chemotherapy correlated with gradual regression of active and total TGF-?1 to the baseline level.
Figure 2. Chemotherapy-induced Smad3 phosphorylation is mediated by TGF-?1. (A): Quantitative analysis of the release of active and total TGF-?1 from HS-27A stromal cell supernatants exposed to eto-poside at 0–100 μM for 1 hour (left graph) or at 100 μM for 0–6 hours (right graph). Bars marked with * or # indicate significant differences compared with untreated controls (p < .05). (B): Immunoprecipitation of human TGF-?1 from HS-27A and P148 supernatants after exposure of stromal cells to the indicated concentrations of etoposide for 1 hour. Supernatants were immunoprecipitated with anti–TGF-?1 and run under reducing conditions. Western blots were probed with anti–TGF-?1 antibody. Recombinant human TGF-?1 served as the molecular-size control. (C): Immunoprecipitation of TGF-?1 from HS-27A cell supernatants preincubated with 3 μg of anti–TGF-?1, anti–LAP/TGF-?1, or anti–LTBP-1/TGF-?1 antibodies followed by exposure to 100 μM etoposide for 1 hour. Samples were run under both reducing and nonreducing conditions, and Western blots were probed with anti-human TGF-?1 antibody. Abbreviations: Ig, immunoglobulin; LAP, latency-associated protein; LTBP, latent TGF-?1–binding protein; TGF-?1, transforming growth factor beta-1.
IP of TGF-?1 from the stromal cell supernatants indicated that baseline TGF-?1 in untreated stromal cell supernatants was negligible, with increased TGF-?1 immunoprecipitated from etoposide-treated stromal cell supernatants in a dose-dependent fashion (Fig. 2B).
Because the anti–TGF-?1 antibody we used for IP of TGF-?1 may recognize both active and latent forms of TGF-?1, we performed additional IP experiments with antibodies recognizing the free and total TGF-?1 (i.e., anti–TGF-?1), the small latent complex (i.e., anti–LAP/TGF-?1), or the large latent complex (anti–LTBP-1/TGF-?1) to further address this issue. As shown in Figure 2C, under nonreducing electrophoretic conditions, the major forms of TGF-?1 activated via etopo-side treatment are the 230- and 195-kD large latency complexes (i.e., LTBP-1/LAP/TGF-?1) as immunoprecipitated by anti–TGF-?1, anti–LAP/TGF-?1, and anti–LTBP-1/TGF-?1 antibodies. In addition, a 100-kD band, which represents the small latency complex (i.e., LAP/TGF-?1), and a 75-kD LAP band were also detected in etoposide-treated samples. When the same samples were electrophoresed under reducing conditions, the high-molecular-weight large and small latency complexes were partially dissociated, and two bands of molecular size of 195 kD (LTBP-1/LAP- TGF-?1) and 25 kD (TGF-?1) were observed.
Disruption of the Availability of TGF-?1 Blocks the Signal Transduction Initiated by Chemotherapy
To better understand the role of TGF-?1 in mediating chemotherapy-triggered signals during bone marrow damage, marrow-derived stromal cells were treated with chemotherapeutic agents in the presence or absence of anti–TGF-?1 neutralizing antibody. Etoposide, melphalan, and 4-HC promoted phosphorylation of Smad3 when cells were pretreated with the isotype control antibody, whereas phosphorylation of Smad3 was diminished in the presence of TGF-?1 neutralizing antibody (Fig. 3A).
Figure 3. Disruption of the availability of transforming growth factor-?1 (TGF-?1) blocks the signal transduction initiated by chemotherapy. (A): Western blot analysis of HS-27A stromal cells treated with 100 μM etoposide, 200 μg/ml melphalan, or 100 μg/ml 4-hydroperoxylcyclophosphamide (4-HC) for 1 hour. (B): Enzyme-linked immunosorbent assay of released total TGF-?1 from HS-27A and P148 stromal cells transfected with the indicated concentrations of TGF-?1 knockdown small interfering RNA (siRNA) or control double-stranded RNA (dsRNA) for 48 hours followed by exposure to 100 μM etoposide for 1 hour (upper panel). Bars marked with * or # indicate significant differences compared with control dsRNA transfections (p < .05). Western blot analysis of Smad-3 phosphorylation using the same TGF-?1 siRNA transfection cell lysates is shown in the lower panel.
This prompted us to more specifically test whether chemotherapy-induced effects on marrow stromal cells could be disrupted through downregulation of TGF-?1 expression. Human HS-27A and P148 stromal cells were transiently transfected with TGF-?1 knockdown siRNA before exposure to etoposide. TGF-?1 targeting siRNA transfection diminished the amount of total TGF-?1 release induced by etoposide treatment in a concentration-dependent manner compared with control dsRNA transfection (Fig. 3B, upper panel). Complete loss of TGF-?1 release occurred when stromal cells were exposed to 150 nM siRNA in the presence of chemotherapy. Consistent with the diminished availability of TGF-?1 presented in the supernatants, phosphorylation of Smad3 was also reduced after transfection of stromal cells with various concentrations of TGF-?1 siRNA during chemotherapy (Fig. 3B, lower panel).
Chemotherapy-Induced MMP-2 Activity Is Required for Activation of Latent TGF-?1
Gelatin zymography revealed that MMP-2 activity was elevated in S10 stromal cell supernatants after etoposide exposure as early as 5 minutes and increased further at 30 to 60 minutes (Fig. 4A). Inhibition of MMP-2 activity by OA-Hy diminished phosphorylation of Smad3 after etoposide treatment of stromal cells (Fig. 4B). To determine whether MMP-2 was required for bone marrow stromal cell activation of TGF-?1, we established stromal cells from MMP-2–/– knockout mice. Etoposide, Melphalan, or 4-HC exposure induced Smad3 phosphorylation in murine MMP-2+/+ stromal cells, whereas phospho-Smad3 signals were less pronounced in MMP-2–/– stromal cells (Fig. 4C). Addition of active MMP-2 partially restored treatment-induced phospho-Smad3 signals in MMP-2–/– cells and further increased phosphorylation of Smad3 in MMP-2+/+ stromal cells treated with etoposide (Fig. 4D).
Figure 4. Chemotherapy-induced MMP-2 activity is required for activation of latent TGF-1. (A): Gelatin zymography analysis of supernatants from S-10 stromal cells treated with 100 μM etoposide for 0 to 6 hours. (B): Western blot analysis of HS-27A and Ped604 stromal cells treated with 1 μM MMP-2 inhibitor OA-Hy for 30 minutes before exposure to 100 μM etoposide for 1 hour. (C): Western blot analysis of murine C57BL/6 MMP-2 knockout stromal cells (MMP-2–/–, KO) or wild-type stromal cells (MMP-2+/+, WT) treated with 100 μM etoposide, 200 μg/ml melphalan, or 100 μg/ml 4-HC for 1 hour. (D): Western blot analysis of MMP-2+/+ and MMP-2–/– stromal cells treated with 100 μM etoposide in the presence or absence of 250 ng/ml recombinant human active MMP-2 for 1 hour. Untreated controls were not exposed to either recombinant MMP-2 or etoposide. (E): Gelatinolytic analysis of supernatants and cell lysates from S-10 parental, vector, or MMP-2 transfected S-10 stromal cells treated with 100 μM etoposide for 1 hour and Western blot analysis of Smad3 phosphorylation and NPT II expression using the same transfection samples. Abbreviations: 4-HC, 4-hydroperoxycyclophosphamide; KO, knockout; MMP-2, matrix metalloproteinase-2; NPT II, neomycin phosphotransferase II; OA-Hy, cis-9-octadecenoyl-N-hydroxylamide; TGF, transforming growth factor; WT, wild-type.
To further investigate the role of MMP-2 in mediating activation of TGF-?1 in marrow stromal cells during chemotherapy, S-10 murine stromal cells transfected with a human MMP-2 construct or vector control were established. Stromal cell clones with comparable expression of the neomycin resistance gene NPT II were selected for further experiments. As shown in Figure 4E, although no substantial differences were observed between S-10 parental and SV-2 vector–transfected cells in terms of activation of Smad3 and MMP-2 during treatment, overexpression of MMP-2 in SM-8 stromal cells increased baseline and etoposide-induced Smad3 phosphorylation.
Activation of Latent MMP-2 by Chemotherapy Requires the Generation of Reactive Oxygen Species
Because MMP-2 exists largely as a latent form in stromal cell matrix, we next sought to explore the mechanism underlying the activation of pro–MMP-2 during etoposide chemotherapy. ROS was generated after etoposide treatment and was required for conversion of pro–MMP-2 to its active form. Etoposide rapidly induced production of intracellular ROS in HS-27A stromal cells as early as 5 minutes after etoposide exposure, which preceded activation of MMP-2 and TGF-?1 (Fig. 5A, upper panel).
Figure 5. Activation of latent MMP-2 by chemotherapy requires the generation of ROS. (A): Flow cytometric analysis of intracellular ROS in HS-27A stromal cells treated with 100 μM etoposide for 0–4 hours (upper panel) or HS-27A stromal cells treated with the indicated chemotherapeutic agents identical to those shown in Figure 1C for 1 hour (lower panel). Untreated control stromal cells in the lower panel are indicated by the solid histogram. Ara-C–treated stromal cell ROS overlays the untreated control histogram. (B): Western blot analysis of HS-27A or P148 stromal cells pretreated with 20 mM NAC overnight before exposure to 100 μM etoposide (VP-16) for 1 hour. (C): Western blot analysis of HS-27A stromal cells treated with recombinant pro–MMP-2 that was activated in vitro. Cells were exposed to 0–500 ng/ml activated MMP-2 for 1 hour. (D): Western blot analysis of MMP-2+/+ and MMP-2–/– stromal cells treated with 10 μM H2O2 in the presence or absence of 250 ng/ml pro–MMP-2 for 1 hour. Abbreviations: 4-HC, 4-hydroperoxylcyclophosphamide; DNR, daunorubicin; DOX, doxorubicin; KO, knockout; Mel, melphalan; MMP-2, matrix metalloproteinase-2; NAC, N-acetyl-cysteine; ROS, reactive oxygen species; VCR, vincristine; WT, wild-type.
Comparable to the activation of MMP-2 and TGF-?1, ROS generation is also a transient event during chemotherapy in our stromal cell model. The mean fluorescence intensity emitted by oxidized 2',7-dichlorodihydrofluorescein diacetate in etoposide-treated stromal cells was increased greater than twofold to threefold in all lines evaluated compared with untreated controls. Uniquely, Ara-C did not stimulate stromal cell production of H2O2 during short-term (1-hour) chemotherapy (Fig. 5A, lower panel). Reduction of intracellular ROS accumulation with the hydroxyl radical scavenger NAC reduced phospho-Smad3 in stromal cells treated with etoposide (Fig. 5B).
To confirm the role of ROS in the activation of MMP-2, pro–MMP-2 was activated in vitro by hydrogen peroxide. As shown in Figure 5C, treatment of HS-27A stromal cells with in vitro–activated MMP-2 induced phosphorylation of Smad3 in a dose-dependent manner. Because inhibition of extracellular MMP-2 activity and reduction of intracellular ROS both disrupted etoposide-induced Smad3 phosphorylation, we sought to determine which one was the initiating factor in modulating TGF-?1/Smad3 signaling. Hydrogen peroxide–induced phosphorylation of Smad3 only occurred in MMP-2+/+ but not MMP-2–/– cells, whereas in the presence of pro–MMP-2, oxidative stress led to phosphorylation of Smad3 in MMP-2–/– cells (Fig. 5D).
P38 Mediates Etoposide-Induced Smad3 Phosphorylation in Bone Marrow Stromal Cells
Smad3 was not directly phosphorylated by TGF-?1 receptor I in a classic fashion but appeared to be regulated by p38 MAPK in this specific setting. As indicated in Figure 6A, all the chemotherapeutic drugs evaluated in this study activated p38 kinase. Chemotherapy induced phosphorylation and activation of both Erk1/2 and p38 kinases in HS-27A cells (Fig. 6B); however, inhibition of Erk1/2 MAPK with U0126 did not result in diminished phosphorylation of Smad3. In contrast, interruption of p38 kinase activity with SB220025 blocked etoposide-triggered Smad3 phosphorylation. JNK/SAPK was not involved in chemotherapy-induced activation of TGF-?1 signaling in bone marrow stromal cells.
Figure 6. P38, but not Erk1/2 or JNK kinase, is involved in mediating etoposide (VP-16)–induced Smad3 phosphorylation. (A): Western blot analysis of HS-27A stromal cells treated with TGF-?1 or the same chemotherapeutic agents shown in Figures 1C and 5A (lower panel). (B): Western blot analysis of HS-27 stromal cells pretreated with vehicle, 10 μM Erk1/2 kinase inhibitor U0126, 20 μM p38 kinase inhibitor SB220025, or 5 μM JNK/SAPK inhibitor SP600125 followed by etoposide exposure for 1 hour. Blot was stripped and reprobed with antibodies specific for the proteins indicated. Abbreviations: 4-HC, 4-hydroperoxylcyclophosphamide; DMSO, dimethyl sulfoxide; DNR, daunorubicin; DOX, doxorubicin; Mel, melphalan; TGF-?1, transforming growth factor-beta 1; VCR, vincristine.
Etoposide Treatment Results in Redistribution of Phosphorylated Smad3 Protein in Human Stromal Cells
Changes in cellular distribution of Smad3 protein after etoposide-induced phosphorylation were evaluated (Fig. 7). The phospho-Smad3 signal was negligible in untreated stromal cells, with only the PI-counterstained cell nuclei clearly detected. Cytoplasmic Smad3 was rapidly phosphorylated in response to etoposide stimulation as early as 30 minutes. Longer exposure of stromal cells to etoposide induced a gradual redistribution and accumulation of Smad3 protein in nucleus. After approximately 4 hours of etoposide treatment, most phospho-Smad3 had translocated into the stromal cell nuclei.
Figure 7. Etoposide treatment results in phosphorylation and subsequent nuclear localization of Smad3 protein in human stromal cells. Double-channel confocal microscopy analysis of HS-27A stromal cells treated with 100 μM etoposide for up to 6 hours. Cells were double-stained with 5 μg/100 μl propidium iodide (PI) (rhodamine, red signal) and 3 μg/100 μl of antiphospho-Smad3 antibody followed by staining with fluorescein isothiocyanate (FITC)–conjugated anti-rabbit IgG secondary antibody (green signal). Panels on the right represent merged images of cytosolic and nuclear phospho-smad3 staining. Original magnifications x200.
Recombinant TGF-?1 Activates Smad3 and Impairs Stromal Cell Support of Pro-B Cell Adhesion and Proliferation
To characterize the response of stromal cells to TGF-?1 exposure, human primary P156 stromal cells were treated with rh-TGF-?1 (Fig. 8A). TGF-?1 rapidly induced phosphorylation of Smad3 in P156 stromal cells in a time-dependent manner with elevated Smad3 phosphorylation as early as 30 minutes and decreased phospho-Smad3 signals thereafter (Fig. 8A, upper panel). In contrast to the Smad3 activation pattern induced by chemotherapy, rh-TGF-?1 treatment resulted in the most pronounced Smad3 phosphorylation at 1 and 5 ng/ml of TGF-?1; however, 10 and 20 ng/ml resulted in diminished phospho-Smad3 signals (Fig. 8A, lower panel).
Figure 8. Recombinant transforming growth factor (TGF)-?1 activates Smad3 and impairs stromal cells support of pro-B cell adhesion and proliferation. (A): Western blot analysis of Smad3 phosphorylation of HS-27A stromal cells treated with 3 ng/ml rh-TGF-?1 for the indication time points or the indicated concentrations of rh-TGF-?1 for 24 hours. (B): Adhesion of C1.92 pro-B cells labeled with PKH-26 and cocultured on P156 and S-10 stromal cells pretreated with the indicated concentrations of rh-TGF-?1 for 72 hours. Bars marked with * or # indicate significant differences compared with untreated controls (p < .05). (C): 3H-thymidine incorporation of C1.92 pro-B cells cocultured on P156 and S-10 stromal cells pretreated with the indicated concentrations of rh-TGF-?1 for 72 hours. Bars marked with * or # indicate significant differences compared with untreated controls (p < .05).
To explore the functional consequences of activation of TGF-?1/Smad3 signaling during bone marrow damage, bone marrow stromal cells were treated with rh-TGF-?1 followed by coculture with C1.92 hematopoietic stem cells. Stromal cells pretreated with TGF-?1 diminished the ability to support C1.92 cell adhesion to the stromal cell layer (Fig. 8B). In addition to the diminished adhesion of the pro-B cells, C1.92 cells cocultured on TGF-?1–pretreated human or murine stromal cells had lower cell proliferation (Fig. 8C) compared with those on control stromal cells.
DISCUSSION
This work was supported by NIH grant R01 HL056888 (to L.F.G.) and NIEHS Training grant ES010953 (to S.C.). The authors would like to acknowledge Dr. Michael Reiss (The Cancer Institute of New Jersey, New Brunswick, NJ), who generously provided phospho-Smad2 antibody used in experiments that preceded those shown in the current report as well as anti-phospho-Smad3 antibody before commercial availability.
REFERENCES
Gordon MY, Goldman JM, Gordon-Smith EC. Spatial and functional relationships between human hemopoietic and marrow stromal cells in vitro. STEM CELLS 1983;1:429–439.
Harvey K, Dzierzak E. Cell-cell contact and anatomical compatibility in stromal cell-mediated HSC support during development. STEM CELLS 2004;22:253–258.
Bianco P, Riminucci M, Gronthos S et al. Bone marrow stromal stem cells: nature, biology, and potential applications. STEM CELLS 2001;19:180–192.
Bacigalupo A. Hematopoietic stem cell transplants after reduced intensity conditioning regimen (RI-HSCT): report of a workshop of the European group for Blood and Marrow Transplantation (EBMT). Bone Marrow Transplant 2000;25:803–805.
de LM, Anagnostopoulos A, Munsell M et al. Nonablative versus reduced-intensity conditioning regimens in the treatment of acute myeloid leukemia and high-risk myelodysplastic syndrome: dose is relevant for long-term disease control after allogeneic hematopoietic stem cell transplantation. Blood 2004;104:865–872.
Attisano L, Wrana JL. Signal transduction by the TGF-? superfamily. Science 2002;296:1646–1647.
Derynck R, Zhang YE. Smad-dependent and Smad-independent pathways in TGF-? family signalling. Nature 2003;425:577–584.
Leask A, Abraham DJ. TGF-? signaling and the fibrotic response. FASEB J 2004;18:816–827.
Schnaper HW, Hayashida T, Poncelet AC. It’s a Smad world: regulation of TGF-? signaling in the kidney. J Am Soc Nephrol 2002;13:1126–1128.
Kanzler S, Lohse AW, Keil A et al. TGF-beta1 in liver fibrosis: an inducible transgenic mouse model to study liver fibrogenesis. Am J Physiol 1999;276:G1059–G1068.
Daniels CE, Wilkes MC, Edens M et al. Imatinib mesylate inhibits the profibrogenic activity of TGF-? and prevents bleomycin-mediated lung fibrosis. J Clin Invest 2004;114:1308–1316.
Fortunel N, Hatzfeld J, Kisselev S et al. Release from quiescence of primitive human hematopoietic stem/progenitor cells by blocking their cell-surface TGF-? type II receptor in a short-term in vitro assay. STEM CELLS 2000;18:102–111.
Kim SJ, Letterio J. Transforming growth factor-? signaling in normal and malignant hematopoiesis. Leukemia 2003;17:1731–1737.
Gleizes PE, Munger JS, Nunes I et al. TGF-? latency: biological significance and mechanisms of activation. STEM CELLS 1997;15:190–197.
Chu TM, Kawinski E. Plasmin, substilisin-like endoproteases, tissue plasminogen activator, and urokinase plasminogen activator are involved in activation of latent TGF-?1 in human seminal plasma. Biochem Biophys Res Commun 1998;253:128–134.
Mitchell EJ, O’Connor-McCourt MD. A transforming growth factor beta (TGF-?) receptor from human placenta exhibits a greater affinity for TGF-?2 than for TGF-?1. Biochemistry 1991;30:4350–4356.
Yevdokimova N, Wahab NA, Mason RM. Thrombospondin-1 is the key activator of TGF-?1 in human mesangial cells exposed to high glucose. J Am Soc Nephrol 2001;12:703–712.
Blakytny R, Ludlow A, Martin GE et al. Latent TGF-?1 activation by platelets. J Cell Physiol 2004;199:67–76.
Annes JP, Chen Y, Munger JS et al. Integrin alphaV?6-mediated activation of latent TGF-? requires the latent TGF-? binding protein-1. J Cell Biol 2004;165:723–734.
Shi Y, Massague J. Mechanisms of TGF-? signaling from cell membrane to the nucleus. Cell 2003;113:685–700.
Roecklein BA, Torok-Storb B. Functionally distinct human marrow stromal cell lines immortalized by transduction with the human papilloma virus E6/E7 genes. Blood 1995;85:997–1005.
Gibson LF, Fortney J, Landreth KS et al. Disruption of bone marrow stromal cell function by etoposide. Biol Blood Marrow Transplant 1997;3:122–132.
Corry DB, Kiss A, Song LZ et al. Overlapping and independent contributions of MMP2 and MMP9 to lung allergic inflammatory cell egression through decreased CC chemokines. FASEB J 2004;18:995–997.
Gibson LF, Piktel D, Landreth KS. Insulin-like growth factor-1 potentiates expansion of interleukin-7-dependent pro-B cells. Blood 1993;82:3005–3011.
Hande KR. The importance of drug scheduling in cancer chemotherapy: etoposide as an example. STEM CELLS 1996;14:18–24.
Wang L, Fortney JE, Gibson LF. Stromal cell protection of B-lineage acute lymphoblastic leukemic cells during chemotherapy requires active Akt. Leuk Res 2004;28:733–742.
Wang L, Chen L, Benincosa J et al. VEGF-induced phosphorylation of Bcl-2 influences B lineage leukemic cell response to apoptotic stimuli. Leukemia 2005;19:344–353.
Fischer OM, Giordano S, Comoglio PM et al. Reactive oxygen species mediate Met receptor transactivation by G protein-coupled receptors and the epidermal growth factor receptor in human carcinoma cells. J Biol Chem 2004;279:28970–28978.
Thomas ED Sr. Stem cell transplantation: past, present and future. STEM CELLS 1994;12:539–544.
Clift RA, Appelbaum FR, Thomas ED. Treatment of chronic myeloid leukemia by marrow transplantation. Blood 1993;82:1954–1956.
Imrie K, Dicke KA, Keating A. Autologous bone marrow transplantation for acute myeloid leukemia. STEM CELLS 1996;14:69–78.
Gorin NC. Autologous stem cell transplantation in acute lymphocytic leukemia. STEM CELLS 2002;20:3–10.
Socie G, Salooja N, Cohen A et al. Nonmalignant late effects after allogeneic stem cell transplantation. Blood 2003;101:3373–3385.
Pedersen-Bjergaard J, Andersen MK, Christiansen DH. Therapy-related acute myeloid leukemia and myelodysplasia after high-dose chemotherapy and autologous stem cell transplantation. Blood 2000;95:3273–3279.
Guillaume T, Rubinstein DB, Symann M. Immune reconstitution and immunotherapy after autologous hematopoietic stem cell transplantation. Blood 1998;92:1471–1490.
Thiele J, Kvasnicka HM, Beelen DW et al. Megakaryopoiesis and myelofibrosis in chronic myeloid leukemia after allogeneic bone marrow transplantation: an immunohistochemical study of 127 patients. Mod Pathol 2001;14:129–138.
Thiele J, Kvasnicka HM, Beelen DW et al. Relevance and dynamics of myelofibrosis regarding hematopoietic reconstitution after allogeneic bone marrow transplantation in chronic myelogenous leukemia: a single center experience on 160 patients. Bone Marrow Transplant 2000;26:275–281.
Hall BM, Fortney JE, Gibson LF. Alteration of nuclear factor-B (NF-B) expression in bone marrow stromal cells treated with etoposide. Biochem Pharmacol 2001;61:1243–1252.
Sontag E. Protein phosphatase 2A: the Trojan Horse of cellular signaling. Cell Signal 2001;13:7–16.
Yu Q, Stamenkovic I. Cell surface-localized matrix metalloproteinase-9 proteolytically activates TGF-? and promotes tumor invasion and angiogenesis. Genes Dev 2000;14:163–176.
Maeda S, Dean DD, Gomez R et al. The first stage of transforming growth factor beta1 activation is release of the large latent complex from the extra-cellular matrix of growth plate chondrocytes by matrix vesicle stromelysin-1 (MMP-3). Calcif Tissue Int 2002;70:54–65.
Goodsell DS. The molecular perspective: matrix metalloproteinase 2. STEM CELLS 2000;18:73–75.
Kunugi S, Fukuda Y, Ishizaki M et al. Role of MMP-2 in alveolar epithelial cell repair after bleomycin administration in rabbits. Lab Invest 2001;81:1309–1318.
Miralles F, Battelino T, Czernichow P et al. TGF-? plays a key role in morphogenesis of the pancreatic islets of Langerhans by controlling the activity of the matrix metalloproteinase MMP-2. J Cell Biol 1998;143:827–836.
Stawowy P, Margeta C, Kallisch H et al. Regulation of matrix metalloproteinase MT1-MMP/MMP-2 in cardiac fibroblasts by TGF-?1 involves furin-convertase. Cardiovasc Res 2004;63:87–97.
Kim ES, Kim MS, Moon A. TGF-?-induced upregulation of MMP-2 and MMP-9 depends on p38 MAPK, but not ERK signaling in MCF10A human breast epithelial cells. Int J Oncol 2004;25:1375–1382.
Kim HS, Shang T, Chen Z et al. TGF-beta1 stimulates production of gelatinase (MMP-9), collagenases (MMP-1, -13) and stromelysins (MMP-3, -10, -11) by human corneal epithelial cells. Exp Eye Res 2004;79:263–274.
Sauer H, Wartenberg M, Hescheler J. Reactive oxygen species as intracellular messengers during cell growth and differentiation. Cell Physiol Biochem 2001;11:173–186.
Thannickal VJ, Fanburg BL. Reactive oxygen species in cell signaling. Am J Physiol Lung Cell Mol Physiol 2000;279:L1005–L1028.
Pociask DA, Sime PJ, Brody AR. Asbestos-derived reactive oxygen species activate TGF-?1. Lab Invest 2004;84:1013–1023.
Rajagopalan S, Meng XP, Ramasamy S et al. Reactive oxygen species produced by macrophage-derived foam cells regulate the activity of vascular matrix metalloproteinases in vitro. Implications for atherosclerotic plaque stability. J Clin Invest 1996;98:2572–2579.
Deem TL, Cook-Mills JM. Vascular cell adhesion molecule 1 (VCAM-1) activation of endothelial cell matrix metalloproteinases: role of reactive oxygen species. Blood 2004;104:2385–2393.
Furukawa F, Matsuzaki K, Mori S et al. p38 MAPK mediates fibrogenic signal through Smad3 phosphorylation in rat myofibroblasts. Hepatology 2003;38:879–889.
Leivonen SK, Chantry A, Hakkinen L et al. Smad3 mediates transforming growth factor-?-induced collagenase-3 (matrix metalloproteinase-13) expression in human gingival fibroblasts: evidence for cross-talk between Smad3 and p38 signaling pathways. J Biol Chem 2002;277:46338–46346.
Mori S, Matsuzaki K, Yoshida K et al. TGF-beta and HGF transmit the signals through JNK-dependent Smad2/3 phosphorylation at the linker regions. Oncogene 2004;23:7416–7429.
Wright N, de Lera TL, Garcia-Moruja C et al. Transforming growth factor-beta1 down-regulates expression of chemokine stromal cell-derived factor-1: functional consequences in cell migration and adhesion. Blood 2003;102:1978–1984.
Dittel BN, McCarthy JB, Wayner EA et al. Regulation of human B-cell precursor adhesion to bone marrow stromal cells by cytokines that exert opposing effects on the expression of vascular cell adhesion molecule-1 (VCAM-1). Blood 1993;81:2272–2282.(Lin Wanga, Suzanne Clutte)
b Department of Microbiology, Immunology, and Cell Biology, and
c Mary Babb Randolph Cancer Center, Robert C. Byrd Health Sciences Center, West Virginia University, School of Medicine, Morgantown, West Virginia, USA
Key Words. Transforming growth factor-?1 ? Smad3 ? Reactive oxygen species ? MMP-2 ? Chemotherapy ? Microenvironment
Correspondence: Laura F. Gibson, Ph.D., P.O. Box 9214, Department of Pediatrics, School of Medicine, WV University, Morgantown, West Virginia, 26506, USA. Telephone: 304-293-5820; Fax: 304-293-4341; e-mail: lgibson@hsc.wvu.edu
ABSTRACT
The bone marrow microenvironment serves as the primary site of normal postnatal hematopoiesis and supports hematopoietic recovery after myelosuppressive chemotherapy or irradiation-induced injury of the immune system . Hematopoietic reconstitution requires efficient migration of transplanted stem/progenitor cells to the bone marrow and relocation to stromal cell niches in this microenvironment . The effects of preparative regimens on the marrow microenvironment remain an area requiring further investigation. The assumption that aggressive chemotherapy spares the bone marrow microenvironment grows increasingly more suspect because dose escalation of chemotherapy reveals unexpected problems with hematopoietic recovery . The dilemma remains maintaining efficacy of tumor eradication while reducing damage to the microenvironment.
Of the signaling molecules in the bone marrow microenvironment that may be involved in chemotherapy-induced bone marrow damage, transforming growth factor (TGF)-?1 is specifically noteworthy. TGF-?1 regulates a variety of biological responses, including angiogenesis, chemotaxis, cell-cycle progression, differentiation, and apoptosis of target cells in a context- and cell-specific manner . TGF-?1 is also involved in regulating extracellular matrix (ECM) remodeling, collagen gene expression, and degradation of matrix proteins during the processes of tissue injury and repair . Upregulated expression or activation of TGF-?1 at sites of injury is associated with proliferation of fibroblasts, progressive fibrosis, and subsequent organ dysfunction in diverse systems, including kidney, liver, and lung . In contrast to its promotion of mesenchymal cell proliferation and survival, TGF-?1 is a potent inhibitor of hematopoietic stem cell proliferation .
TGF-?1 is initially synthesized as a large precursor that is processed to a mature protein during secretion. After secretion, mature TGF-?1 (25 kD) noncovalently associates with its N-terminal pro-peptide, the 75-kD latency-associated protein (LAP) . The TGF-?1–LAP complex predominantly binds to a latent TGF-?1–binding protein (LTBP), which mediates deposition of the latent complex (230 and 195 kD) to the ECM . Release of mature TGF-?1 from the latent complex can be accomplished by different mechanisms, such as proteolytic cleavage of LAP by plasmin , deglycosylation of LAP , or interaction with thrombospondin-1 , platelet , or integrin 4, ?6 . After TGF-?1 ligand binding, TGF-?1 receptor II recruits and activates TGF-?1 receptor I, which in turn phosphorylates and activates the R-Smads, including Smad2 or Smad3 . Phosphorylated R-Smads homodimerize, form a transcriptional complex with Smad4, and translocate into the nucleus to regulate target gene expression .
Data presented in the current study suggest that the TGF-?1/p38/Smad3 signaling cascades are activated through reactive oxygen species (ROS)–mediated matrix metalloproteinase-2 (MMP-2) activity in bone marrow stromal cells during etoposide chemotherapy. Increased availability of active TGF-?1 has the potential to alter stromal cell function through regulation of diverse Smad-driven gene expression in stromal cells. Moreover, release of TGF-?1 from ECM during chemotherapy may also directly regulate growth and proliferation of transplanted hematopoietic stem cells. This in vitro model provides a setting in which we can further delineate the effects of chemotherapy on marrow stromal cells and evaluate the role of TGF-?1 in influencing hematopoietic recovery after transplantation.
MATERIALS AND METHODS
Chemotherapy Activates Smad3 Through Phosphorylation at Serines 433/435 in Human Bone Marrow–Derived Stromal Cells
To investigate the phosphorylation of Smad3 in stromal cells after chemotherapeutic stimulation, stromal cells were treated either for different times or with various concentrations of etoposide. Exposure of stromal cells to etoposide resulted in phosphorylation of Smad3 at serines 433/435 in a time-dependent (Fig. 1A) and dose-dependent (Fig. 1B) manner. Etoposide induced a rapid elevation of phospho-Smad3 signal as early as 30 minutes, which was sustained for approximately 6–7 hours (Fig. 1A). After the transient increase, phospho-Smad3 levels diminished for up to 24 hours in the presence of chemotherapy. Etoposide, melphalan, vincristine, daunorubicin, doxorubicin, 4-hydroperocyclophosphomide, and Ara-C induced Smad3 phosphorylation in stromal cells to varying degrees (Fig. 1C). Treatment of stromal cells with recombinant TGF-?1 served as a positive control and induced the most pronounced phosphorylation of Smad3. Total Smad3 protein remained unchanged and served as the lane loading control.
Figure 1. Chemotherapy activates Smad3 through phosphorylation at serines 433/435 in human bone marrow–derived stromal cells. Western blot analyses of HS-27A stromal cells treated with (A) 100 μM etoposide for the indicated time points, (B) various concentrations of etoposide for 1 hour, or (C) 3 ng/ml transforming growth factor -?1 (TGF-?1), 100 μM etoposide (VP-16), 200 μg/ml melphalan (Mel), 20 μg/ml vincristine (VCR), 100 μg/ml daunorubicin (DNR), 100 μM doxorubicin (DOX), 100 μg/ml 4-hydroperoxylcyclophosphamide (4-HC), or 100 μg/ml Ara-C for 1 hour. Membranes were probed with anti-phospho-Smad3 (P-Smad3, ser433/435) and then stripped and reprobed with total Smad3 (T-Smad3)–specific antibodies.
Chemotherapy-Induced Smad3 Phosphorylation Is Mediated by TGF-?1
To investigate the potential involvement of TGF-?1 in phosphorylation of Smad3 in bone marrow stromal cells during chemotherapy, HS-27A stromal cells were exposed to etoposide for different times or at various concentrations. Quantitative analysis of TGF-?1 by ELISA was performed using the cell supernatants after treatment. To better distinguish the free (active) and latent (total) TGF-?1 released, nonacidification and acidification of the stromal supernatants were simultaneously used before assay as described. Exposure of stromal cells to chemotherapy resulted in elevated TGF-?1 release from stromal cells both in a time-and dose-dependent manner (Fig. 2A). Chemotherapy rapidly induced the release of active and latent forms of TGF-?1 from stromal cell ECM. In our stromal cell model, active TGF-?1 constituted approximately 5%–9% of the total TGF-?1 pool during each treatment phase. Activation of TGF-?1 preceded phosphorylation of Smad3 with initial increases rapidly after etoposide treatment of 15 minutes and further elevation at 1 hour. Activation of TGF-?1 in stromal cells was transient, as longer than 1 hour exposure of stromal cells to chemotherapy correlated with gradual regression of active and total TGF-?1 to the baseline level.
Figure 2. Chemotherapy-induced Smad3 phosphorylation is mediated by TGF-?1. (A): Quantitative analysis of the release of active and total TGF-?1 from HS-27A stromal cell supernatants exposed to eto-poside at 0–100 μM for 1 hour (left graph) or at 100 μM for 0–6 hours (right graph). Bars marked with * or # indicate significant differences compared with untreated controls (p < .05). (B): Immunoprecipitation of human TGF-?1 from HS-27A and P148 supernatants after exposure of stromal cells to the indicated concentrations of etoposide for 1 hour. Supernatants were immunoprecipitated with anti–TGF-?1 and run under reducing conditions. Western blots were probed with anti–TGF-?1 antibody. Recombinant human TGF-?1 served as the molecular-size control. (C): Immunoprecipitation of TGF-?1 from HS-27A cell supernatants preincubated with 3 μg of anti–TGF-?1, anti–LAP/TGF-?1, or anti–LTBP-1/TGF-?1 antibodies followed by exposure to 100 μM etoposide for 1 hour. Samples were run under both reducing and nonreducing conditions, and Western blots were probed with anti-human TGF-?1 antibody. Abbreviations: Ig, immunoglobulin; LAP, latency-associated protein; LTBP, latent TGF-?1–binding protein; TGF-?1, transforming growth factor beta-1.
IP of TGF-?1 from the stromal cell supernatants indicated that baseline TGF-?1 in untreated stromal cell supernatants was negligible, with increased TGF-?1 immunoprecipitated from etoposide-treated stromal cell supernatants in a dose-dependent fashion (Fig. 2B).
Because the anti–TGF-?1 antibody we used for IP of TGF-?1 may recognize both active and latent forms of TGF-?1, we performed additional IP experiments with antibodies recognizing the free and total TGF-?1 (i.e., anti–TGF-?1), the small latent complex (i.e., anti–LAP/TGF-?1), or the large latent complex (anti–LTBP-1/TGF-?1) to further address this issue. As shown in Figure 2C, under nonreducing electrophoretic conditions, the major forms of TGF-?1 activated via etopo-side treatment are the 230- and 195-kD large latency complexes (i.e., LTBP-1/LAP/TGF-?1) as immunoprecipitated by anti–TGF-?1, anti–LAP/TGF-?1, and anti–LTBP-1/TGF-?1 antibodies. In addition, a 100-kD band, which represents the small latency complex (i.e., LAP/TGF-?1), and a 75-kD LAP band were also detected in etoposide-treated samples. When the same samples were electrophoresed under reducing conditions, the high-molecular-weight large and small latency complexes were partially dissociated, and two bands of molecular size of 195 kD (LTBP-1/LAP- TGF-?1) and 25 kD (TGF-?1) were observed.
Disruption of the Availability of TGF-?1 Blocks the Signal Transduction Initiated by Chemotherapy
To better understand the role of TGF-?1 in mediating chemotherapy-triggered signals during bone marrow damage, marrow-derived stromal cells were treated with chemotherapeutic agents in the presence or absence of anti–TGF-?1 neutralizing antibody. Etoposide, melphalan, and 4-HC promoted phosphorylation of Smad3 when cells were pretreated with the isotype control antibody, whereas phosphorylation of Smad3 was diminished in the presence of TGF-?1 neutralizing antibody (Fig. 3A).
Figure 3. Disruption of the availability of transforming growth factor-?1 (TGF-?1) blocks the signal transduction initiated by chemotherapy. (A): Western blot analysis of HS-27A stromal cells treated with 100 μM etoposide, 200 μg/ml melphalan, or 100 μg/ml 4-hydroperoxylcyclophosphamide (4-HC) for 1 hour. (B): Enzyme-linked immunosorbent assay of released total TGF-?1 from HS-27A and P148 stromal cells transfected with the indicated concentrations of TGF-?1 knockdown small interfering RNA (siRNA) or control double-stranded RNA (dsRNA) for 48 hours followed by exposure to 100 μM etoposide for 1 hour (upper panel). Bars marked with * or # indicate significant differences compared with control dsRNA transfections (p < .05). Western blot analysis of Smad-3 phosphorylation using the same TGF-?1 siRNA transfection cell lysates is shown in the lower panel.
This prompted us to more specifically test whether chemotherapy-induced effects on marrow stromal cells could be disrupted through downregulation of TGF-?1 expression. Human HS-27A and P148 stromal cells were transiently transfected with TGF-?1 knockdown siRNA before exposure to etoposide. TGF-?1 targeting siRNA transfection diminished the amount of total TGF-?1 release induced by etoposide treatment in a concentration-dependent manner compared with control dsRNA transfection (Fig. 3B, upper panel). Complete loss of TGF-?1 release occurred when stromal cells were exposed to 150 nM siRNA in the presence of chemotherapy. Consistent with the diminished availability of TGF-?1 presented in the supernatants, phosphorylation of Smad3 was also reduced after transfection of stromal cells with various concentrations of TGF-?1 siRNA during chemotherapy (Fig. 3B, lower panel).
Chemotherapy-Induced MMP-2 Activity Is Required for Activation of Latent TGF-?1
Gelatin zymography revealed that MMP-2 activity was elevated in S10 stromal cell supernatants after etoposide exposure as early as 5 minutes and increased further at 30 to 60 minutes (Fig. 4A). Inhibition of MMP-2 activity by OA-Hy diminished phosphorylation of Smad3 after etoposide treatment of stromal cells (Fig. 4B). To determine whether MMP-2 was required for bone marrow stromal cell activation of TGF-?1, we established stromal cells from MMP-2–/– knockout mice. Etoposide, Melphalan, or 4-HC exposure induced Smad3 phosphorylation in murine MMP-2+/+ stromal cells, whereas phospho-Smad3 signals were less pronounced in MMP-2–/– stromal cells (Fig. 4C). Addition of active MMP-2 partially restored treatment-induced phospho-Smad3 signals in MMP-2–/– cells and further increased phosphorylation of Smad3 in MMP-2+/+ stromal cells treated with etoposide (Fig. 4D).
Figure 4. Chemotherapy-induced MMP-2 activity is required for activation of latent TGF-1. (A): Gelatin zymography analysis of supernatants from S-10 stromal cells treated with 100 μM etoposide for 0 to 6 hours. (B): Western blot analysis of HS-27A and Ped604 stromal cells treated with 1 μM MMP-2 inhibitor OA-Hy for 30 minutes before exposure to 100 μM etoposide for 1 hour. (C): Western blot analysis of murine C57BL/6 MMP-2 knockout stromal cells (MMP-2–/–, KO) or wild-type stromal cells (MMP-2+/+, WT) treated with 100 μM etoposide, 200 μg/ml melphalan, or 100 μg/ml 4-HC for 1 hour. (D): Western blot analysis of MMP-2+/+ and MMP-2–/– stromal cells treated with 100 μM etoposide in the presence or absence of 250 ng/ml recombinant human active MMP-2 for 1 hour. Untreated controls were not exposed to either recombinant MMP-2 or etoposide. (E): Gelatinolytic analysis of supernatants and cell lysates from S-10 parental, vector, or MMP-2 transfected S-10 stromal cells treated with 100 μM etoposide for 1 hour and Western blot analysis of Smad3 phosphorylation and NPT II expression using the same transfection samples. Abbreviations: 4-HC, 4-hydroperoxycyclophosphamide; KO, knockout; MMP-2, matrix metalloproteinase-2; NPT II, neomycin phosphotransferase II; OA-Hy, cis-9-octadecenoyl-N-hydroxylamide; TGF, transforming growth factor; WT, wild-type.
To further investigate the role of MMP-2 in mediating activation of TGF-?1 in marrow stromal cells during chemotherapy, S-10 murine stromal cells transfected with a human MMP-2 construct or vector control were established. Stromal cell clones with comparable expression of the neomycin resistance gene NPT II were selected for further experiments. As shown in Figure 4E, although no substantial differences were observed between S-10 parental and SV-2 vector–transfected cells in terms of activation of Smad3 and MMP-2 during treatment, overexpression of MMP-2 in SM-8 stromal cells increased baseline and etoposide-induced Smad3 phosphorylation.
Activation of Latent MMP-2 by Chemotherapy Requires the Generation of Reactive Oxygen Species
Because MMP-2 exists largely as a latent form in stromal cell matrix, we next sought to explore the mechanism underlying the activation of pro–MMP-2 during etoposide chemotherapy. ROS was generated after etoposide treatment and was required for conversion of pro–MMP-2 to its active form. Etoposide rapidly induced production of intracellular ROS in HS-27A stromal cells as early as 5 minutes after etoposide exposure, which preceded activation of MMP-2 and TGF-?1 (Fig. 5A, upper panel).
Figure 5. Activation of latent MMP-2 by chemotherapy requires the generation of ROS. (A): Flow cytometric analysis of intracellular ROS in HS-27A stromal cells treated with 100 μM etoposide for 0–4 hours (upper panel) or HS-27A stromal cells treated with the indicated chemotherapeutic agents identical to those shown in Figure 1C for 1 hour (lower panel). Untreated control stromal cells in the lower panel are indicated by the solid histogram. Ara-C–treated stromal cell ROS overlays the untreated control histogram. (B): Western blot analysis of HS-27A or P148 stromal cells pretreated with 20 mM NAC overnight before exposure to 100 μM etoposide (VP-16) for 1 hour. (C): Western blot analysis of HS-27A stromal cells treated with recombinant pro–MMP-2 that was activated in vitro. Cells were exposed to 0–500 ng/ml activated MMP-2 for 1 hour. (D): Western blot analysis of MMP-2+/+ and MMP-2–/– stromal cells treated with 10 μM H2O2 in the presence or absence of 250 ng/ml pro–MMP-2 for 1 hour. Abbreviations: 4-HC, 4-hydroperoxylcyclophosphamide; DNR, daunorubicin; DOX, doxorubicin; KO, knockout; Mel, melphalan; MMP-2, matrix metalloproteinase-2; NAC, N-acetyl-cysteine; ROS, reactive oxygen species; VCR, vincristine; WT, wild-type.
Comparable to the activation of MMP-2 and TGF-?1, ROS generation is also a transient event during chemotherapy in our stromal cell model. The mean fluorescence intensity emitted by oxidized 2',7-dichlorodihydrofluorescein diacetate in etoposide-treated stromal cells was increased greater than twofold to threefold in all lines evaluated compared with untreated controls. Uniquely, Ara-C did not stimulate stromal cell production of H2O2 during short-term (1-hour) chemotherapy (Fig. 5A, lower panel). Reduction of intracellular ROS accumulation with the hydroxyl radical scavenger NAC reduced phospho-Smad3 in stromal cells treated with etoposide (Fig. 5B).
To confirm the role of ROS in the activation of MMP-2, pro–MMP-2 was activated in vitro by hydrogen peroxide. As shown in Figure 5C, treatment of HS-27A stromal cells with in vitro–activated MMP-2 induced phosphorylation of Smad3 in a dose-dependent manner. Because inhibition of extracellular MMP-2 activity and reduction of intracellular ROS both disrupted etoposide-induced Smad3 phosphorylation, we sought to determine which one was the initiating factor in modulating TGF-?1/Smad3 signaling. Hydrogen peroxide–induced phosphorylation of Smad3 only occurred in MMP-2+/+ but not MMP-2–/– cells, whereas in the presence of pro–MMP-2, oxidative stress led to phosphorylation of Smad3 in MMP-2–/– cells (Fig. 5D).
P38 Mediates Etoposide-Induced Smad3 Phosphorylation in Bone Marrow Stromal Cells
Smad3 was not directly phosphorylated by TGF-?1 receptor I in a classic fashion but appeared to be regulated by p38 MAPK in this specific setting. As indicated in Figure 6A, all the chemotherapeutic drugs evaluated in this study activated p38 kinase. Chemotherapy induced phosphorylation and activation of both Erk1/2 and p38 kinases in HS-27A cells (Fig. 6B); however, inhibition of Erk1/2 MAPK with U0126 did not result in diminished phosphorylation of Smad3. In contrast, interruption of p38 kinase activity with SB220025 blocked etoposide-triggered Smad3 phosphorylation. JNK/SAPK was not involved in chemotherapy-induced activation of TGF-?1 signaling in bone marrow stromal cells.
Figure 6. P38, but not Erk1/2 or JNK kinase, is involved in mediating etoposide (VP-16)–induced Smad3 phosphorylation. (A): Western blot analysis of HS-27A stromal cells treated with TGF-?1 or the same chemotherapeutic agents shown in Figures 1C and 5A (lower panel). (B): Western blot analysis of HS-27 stromal cells pretreated with vehicle, 10 μM Erk1/2 kinase inhibitor U0126, 20 μM p38 kinase inhibitor SB220025, or 5 μM JNK/SAPK inhibitor SP600125 followed by etoposide exposure for 1 hour. Blot was stripped and reprobed with antibodies specific for the proteins indicated. Abbreviations: 4-HC, 4-hydroperoxylcyclophosphamide; DMSO, dimethyl sulfoxide; DNR, daunorubicin; DOX, doxorubicin; Mel, melphalan; TGF-?1, transforming growth factor-beta 1; VCR, vincristine.
Etoposide Treatment Results in Redistribution of Phosphorylated Smad3 Protein in Human Stromal Cells
Changes in cellular distribution of Smad3 protein after etoposide-induced phosphorylation were evaluated (Fig. 7). The phospho-Smad3 signal was negligible in untreated stromal cells, with only the PI-counterstained cell nuclei clearly detected. Cytoplasmic Smad3 was rapidly phosphorylated in response to etoposide stimulation as early as 30 minutes. Longer exposure of stromal cells to etoposide induced a gradual redistribution and accumulation of Smad3 protein in nucleus. After approximately 4 hours of etoposide treatment, most phospho-Smad3 had translocated into the stromal cell nuclei.
Figure 7. Etoposide treatment results in phosphorylation and subsequent nuclear localization of Smad3 protein in human stromal cells. Double-channel confocal microscopy analysis of HS-27A stromal cells treated with 100 μM etoposide for up to 6 hours. Cells were double-stained with 5 μg/100 μl propidium iodide (PI) (rhodamine, red signal) and 3 μg/100 μl of antiphospho-Smad3 antibody followed by staining with fluorescein isothiocyanate (FITC)–conjugated anti-rabbit IgG secondary antibody (green signal). Panels on the right represent merged images of cytosolic and nuclear phospho-smad3 staining. Original magnifications x200.
Recombinant TGF-?1 Activates Smad3 and Impairs Stromal Cell Support of Pro-B Cell Adhesion and Proliferation
To characterize the response of stromal cells to TGF-?1 exposure, human primary P156 stromal cells were treated with rh-TGF-?1 (Fig. 8A). TGF-?1 rapidly induced phosphorylation of Smad3 in P156 stromal cells in a time-dependent manner with elevated Smad3 phosphorylation as early as 30 minutes and decreased phospho-Smad3 signals thereafter (Fig. 8A, upper panel). In contrast to the Smad3 activation pattern induced by chemotherapy, rh-TGF-?1 treatment resulted in the most pronounced Smad3 phosphorylation at 1 and 5 ng/ml of TGF-?1; however, 10 and 20 ng/ml resulted in diminished phospho-Smad3 signals (Fig. 8A, lower panel).
Figure 8. Recombinant transforming growth factor (TGF)-?1 activates Smad3 and impairs stromal cells support of pro-B cell adhesion and proliferation. (A): Western blot analysis of Smad3 phosphorylation of HS-27A stromal cells treated with 3 ng/ml rh-TGF-?1 for the indication time points or the indicated concentrations of rh-TGF-?1 for 24 hours. (B): Adhesion of C1.92 pro-B cells labeled with PKH-26 and cocultured on P156 and S-10 stromal cells pretreated with the indicated concentrations of rh-TGF-?1 for 72 hours. Bars marked with * or # indicate significant differences compared with untreated controls (p < .05). (C): 3H-thymidine incorporation of C1.92 pro-B cells cocultured on P156 and S-10 stromal cells pretreated with the indicated concentrations of rh-TGF-?1 for 72 hours. Bars marked with * or # indicate significant differences compared with untreated controls (p < .05).
To explore the functional consequences of activation of TGF-?1/Smad3 signaling during bone marrow damage, bone marrow stromal cells were treated with rh-TGF-?1 followed by coculture with C1.92 hematopoietic stem cells. Stromal cells pretreated with TGF-?1 diminished the ability to support C1.92 cell adhesion to the stromal cell layer (Fig. 8B). In addition to the diminished adhesion of the pro-B cells, C1.92 cells cocultured on TGF-?1–pretreated human or murine stromal cells had lower cell proliferation (Fig. 8C) compared with those on control stromal cells.
DISCUSSION
This work was supported by NIH grant R01 HL056888 (to L.F.G.) and NIEHS Training grant ES010953 (to S.C.). The authors would like to acknowledge Dr. Michael Reiss (The Cancer Institute of New Jersey, New Brunswick, NJ), who generously provided phospho-Smad2 antibody used in experiments that preceded those shown in the current report as well as anti-phospho-Smad3 antibody before commercial availability.
REFERENCES
Gordon MY, Goldman JM, Gordon-Smith EC. Spatial and functional relationships between human hemopoietic and marrow stromal cells in vitro. STEM CELLS 1983;1:429–439.
Harvey K, Dzierzak E. Cell-cell contact and anatomical compatibility in stromal cell-mediated HSC support during development. STEM CELLS 2004;22:253–258.
Bianco P, Riminucci M, Gronthos S et al. Bone marrow stromal stem cells: nature, biology, and potential applications. STEM CELLS 2001;19:180–192.
Bacigalupo A. Hematopoietic stem cell transplants after reduced intensity conditioning regimen (RI-HSCT): report of a workshop of the European group for Blood and Marrow Transplantation (EBMT). Bone Marrow Transplant 2000;25:803–805.
de LM, Anagnostopoulos A, Munsell M et al. Nonablative versus reduced-intensity conditioning regimens in the treatment of acute myeloid leukemia and high-risk myelodysplastic syndrome: dose is relevant for long-term disease control after allogeneic hematopoietic stem cell transplantation. Blood 2004;104:865–872.
Attisano L, Wrana JL. Signal transduction by the TGF-? superfamily. Science 2002;296:1646–1647.
Derynck R, Zhang YE. Smad-dependent and Smad-independent pathways in TGF-? family signalling. Nature 2003;425:577–584.
Leask A, Abraham DJ. TGF-? signaling and the fibrotic response. FASEB J 2004;18:816–827.
Schnaper HW, Hayashida T, Poncelet AC. It’s a Smad world: regulation of TGF-? signaling in the kidney. J Am Soc Nephrol 2002;13:1126–1128.
Kanzler S, Lohse AW, Keil A et al. TGF-beta1 in liver fibrosis: an inducible transgenic mouse model to study liver fibrogenesis. Am J Physiol 1999;276:G1059–G1068.
Daniels CE, Wilkes MC, Edens M et al. Imatinib mesylate inhibits the profibrogenic activity of TGF-? and prevents bleomycin-mediated lung fibrosis. J Clin Invest 2004;114:1308–1316.
Fortunel N, Hatzfeld J, Kisselev S et al. Release from quiescence of primitive human hematopoietic stem/progenitor cells by blocking their cell-surface TGF-? type II receptor in a short-term in vitro assay. STEM CELLS 2000;18:102–111.
Kim SJ, Letterio J. Transforming growth factor-? signaling in normal and malignant hematopoiesis. Leukemia 2003;17:1731–1737.
Gleizes PE, Munger JS, Nunes I et al. TGF-? latency: biological significance and mechanisms of activation. STEM CELLS 1997;15:190–197.
Chu TM, Kawinski E. Plasmin, substilisin-like endoproteases, tissue plasminogen activator, and urokinase plasminogen activator are involved in activation of latent TGF-?1 in human seminal plasma. Biochem Biophys Res Commun 1998;253:128–134.
Mitchell EJ, O’Connor-McCourt MD. A transforming growth factor beta (TGF-?) receptor from human placenta exhibits a greater affinity for TGF-?2 than for TGF-?1. Biochemistry 1991;30:4350–4356.
Yevdokimova N, Wahab NA, Mason RM. Thrombospondin-1 is the key activator of TGF-?1 in human mesangial cells exposed to high glucose. J Am Soc Nephrol 2001;12:703–712.
Blakytny R, Ludlow A, Martin GE et al. Latent TGF-?1 activation by platelets. J Cell Physiol 2004;199:67–76.
Annes JP, Chen Y, Munger JS et al. Integrin alphaV?6-mediated activation of latent TGF-? requires the latent TGF-? binding protein-1. J Cell Biol 2004;165:723–734.
Shi Y, Massague J. Mechanisms of TGF-? signaling from cell membrane to the nucleus. Cell 2003;113:685–700.
Roecklein BA, Torok-Storb B. Functionally distinct human marrow stromal cell lines immortalized by transduction with the human papilloma virus E6/E7 genes. Blood 1995;85:997–1005.
Gibson LF, Fortney J, Landreth KS et al. Disruption of bone marrow stromal cell function by etoposide. Biol Blood Marrow Transplant 1997;3:122–132.
Corry DB, Kiss A, Song LZ et al. Overlapping and independent contributions of MMP2 and MMP9 to lung allergic inflammatory cell egression through decreased CC chemokines. FASEB J 2004;18:995–997.
Gibson LF, Piktel D, Landreth KS. Insulin-like growth factor-1 potentiates expansion of interleukin-7-dependent pro-B cells. Blood 1993;82:3005–3011.
Hande KR. The importance of drug scheduling in cancer chemotherapy: etoposide as an example. STEM CELLS 1996;14:18–24.
Wang L, Fortney JE, Gibson LF. Stromal cell protection of B-lineage acute lymphoblastic leukemic cells during chemotherapy requires active Akt. Leuk Res 2004;28:733–742.
Wang L, Chen L, Benincosa J et al. VEGF-induced phosphorylation of Bcl-2 influences B lineage leukemic cell response to apoptotic stimuli. Leukemia 2005;19:344–353.
Fischer OM, Giordano S, Comoglio PM et al. Reactive oxygen species mediate Met receptor transactivation by G protein-coupled receptors and the epidermal growth factor receptor in human carcinoma cells. J Biol Chem 2004;279:28970–28978.
Thomas ED Sr. Stem cell transplantation: past, present and future. STEM CELLS 1994;12:539–544.
Clift RA, Appelbaum FR, Thomas ED. Treatment of chronic myeloid leukemia by marrow transplantation. Blood 1993;82:1954–1956.
Imrie K, Dicke KA, Keating A. Autologous bone marrow transplantation for acute myeloid leukemia. STEM CELLS 1996;14:69–78.
Gorin NC. Autologous stem cell transplantation in acute lymphocytic leukemia. STEM CELLS 2002;20:3–10.
Socie G, Salooja N, Cohen A et al. Nonmalignant late effects after allogeneic stem cell transplantation. Blood 2003;101:3373–3385.
Pedersen-Bjergaard J, Andersen MK, Christiansen DH. Therapy-related acute myeloid leukemia and myelodysplasia after high-dose chemotherapy and autologous stem cell transplantation. Blood 2000;95:3273–3279.
Guillaume T, Rubinstein DB, Symann M. Immune reconstitution and immunotherapy after autologous hematopoietic stem cell transplantation. Blood 1998;92:1471–1490.
Thiele J, Kvasnicka HM, Beelen DW et al. Megakaryopoiesis and myelofibrosis in chronic myeloid leukemia after allogeneic bone marrow transplantation: an immunohistochemical study of 127 patients. Mod Pathol 2001;14:129–138.
Thiele J, Kvasnicka HM, Beelen DW et al. Relevance and dynamics of myelofibrosis regarding hematopoietic reconstitution after allogeneic bone marrow transplantation in chronic myelogenous leukemia: a single center experience on 160 patients. Bone Marrow Transplant 2000;26:275–281.
Hall BM, Fortney JE, Gibson LF. Alteration of nuclear factor-B (NF-B) expression in bone marrow stromal cells treated with etoposide. Biochem Pharmacol 2001;61:1243–1252.
Sontag E. Protein phosphatase 2A: the Trojan Horse of cellular signaling. Cell Signal 2001;13:7–16.
Yu Q, Stamenkovic I. Cell surface-localized matrix metalloproteinase-9 proteolytically activates TGF-? and promotes tumor invasion and angiogenesis. Genes Dev 2000;14:163–176.
Maeda S, Dean DD, Gomez R et al. The first stage of transforming growth factor beta1 activation is release of the large latent complex from the extra-cellular matrix of growth plate chondrocytes by matrix vesicle stromelysin-1 (MMP-3). Calcif Tissue Int 2002;70:54–65.
Goodsell DS. The molecular perspective: matrix metalloproteinase 2. STEM CELLS 2000;18:73–75.
Kunugi S, Fukuda Y, Ishizaki M et al. Role of MMP-2 in alveolar epithelial cell repair after bleomycin administration in rabbits. Lab Invest 2001;81:1309–1318.
Miralles F, Battelino T, Czernichow P et al. TGF-? plays a key role in morphogenesis of the pancreatic islets of Langerhans by controlling the activity of the matrix metalloproteinase MMP-2. J Cell Biol 1998;143:827–836.
Stawowy P, Margeta C, Kallisch H et al. Regulation of matrix metalloproteinase MT1-MMP/MMP-2 in cardiac fibroblasts by TGF-?1 involves furin-convertase. Cardiovasc Res 2004;63:87–97.
Kim ES, Kim MS, Moon A. TGF-?-induced upregulation of MMP-2 and MMP-9 depends on p38 MAPK, but not ERK signaling in MCF10A human breast epithelial cells. Int J Oncol 2004;25:1375–1382.
Kim HS, Shang T, Chen Z et al. TGF-beta1 stimulates production of gelatinase (MMP-9), collagenases (MMP-1, -13) and stromelysins (MMP-3, -10, -11) by human corneal epithelial cells. Exp Eye Res 2004;79:263–274.
Sauer H, Wartenberg M, Hescheler J. Reactive oxygen species as intracellular messengers during cell growth and differentiation. Cell Physiol Biochem 2001;11:173–186.
Thannickal VJ, Fanburg BL. Reactive oxygen species in cell signaling. Am J Physiol Lung Cell Mol Physiol 2000;279:L1005–L1028.
Pociask DA, Sime PJ, Brody AR. Asbestos-derived reactive oxygen species activate TGF-?1. Lab Invest 2004;84:1013–1023.
Rajagopalan S, Meng XP, Ramasamy S et al. Reactive oxygen species produced by macrophage-derived foam cells regulate the activity of vascular matrix metalloproteinases in vitro. Implications for atherosclerotic plaque stability. J Clin Invest 1996;98:2572–2579.
Deem TL, Cook-Mills JM. Vascular cell adhesion molecule 1 (VCAM-1) activation of endothelial cell matrix metalloproteinases: role of reactive oxygen species. Blood 2004;104:2385–2393.
Furukawa F, Matsuzaki K, Mori S et al. p38 MAPK mediates fibrogenic signal through Smad3 phosphorylation in rat myofibroblasts. Hepatology 2003;38:879–889.
Leivonen SK, Chantry A, Hakkinen L et al. Smad3 mediates transforming growth factor-?-induced collagenase-3 (matrix metalloproteinase-13) expression in human gingival fibroblasts: evidence for cross-talk between Smad3 and p38 signaling pathways. J Biol Chem 2002;277:46338–46346.
Mori S, Matsuzaki K, Yoshida K et al. TGF-beta and HGF transmit the signals through JNK-dependent Smad2/3 phosphorylation at the linker regions. Oncogene 2004;23:7416–7429.
Wright N, de Lera TL, Garcia-Moruja C et al. Transforming growth factor-beta1 down-regulates expression of chemokine stromal cell-derived factor-1: functional consequences in cell migration and adhesion. Blood 2003;102:1978–1984.
Dittel BN, McCarthy JB, Wayner EA et al. Regulation of human B-cell precursor adhesion to bone marrow stromal cells by cytokines that exert opposing effects on the expression of vascular cell adhesion molecule-1 (VCAM-1). Blood 1993;81:2272–2282.(Lin Wanga, Suzanne Clutte)