Rac1 mediates intestinal epithelial cell apoptosis via JNK
Abstract
Apoptosis plays a key role in the maintenance of a constant cell number and a low incidence of cancer in the mucosa of the intestine. Although the small GTPase Rac1 has been established as an important regulator of migration of intestinal epithelial cells, whether Rac1 is also involved in apoptosis is unclear. The present study tested the hypothesis that Rac1 mediates TNF-α-induced apoptosis in IEC-6 cells. Rac1 is activated during TNF-α-induced apoptosis as judged by the level of GTP-Rac1, the level of microsomal membrane-associated Rac1, and lamellipodia formation. Although expression of constitutively active Rac1 does not increase apoptosis in the basal condition, inhibition of Rac1 either by NSC-23766 (Rac1 inhibitor) or expression of dominant negative Rac1 protects cells from TNF-α-induced apoptosis by inhibiting caspase-3, -8, and -9 activities. Inhibition of Rac1 before the administration of apoptotic stimuli significantly prevents TNF-α-induced activation of JNK1/2, the key proapoptotic regulator in IEC-6 cells. Inhibition of Rac1 does not modulate TNF-α-induced ERK1/2 and Akt activation. Inhibition of ERK1/2 and Akt activity by U-0126 and LY-294002, respectively, increased TNF-α-induced apoptosis. However, inhibition of Rac1 significantly decreased apoptosis in the presence of ERK1/2 and Akt inhibitors, similar to the effect observed with NSC-23766 alone in response to TNF-α. Thus, Rac1 inhibition protects cells independently of ERK1/2 and Akt activation during TNF-α-induced apoptosis. Although p38 MAPK is activated in response to TNF-α, inhibition of p38 MAPK did not decrease apoptosis. Rac1 inhibition did not alter p38 MAPK activity. Thus, these results indicate that Rac1 mediates apoptosis via JNK and plays a key role in proapoptotic pathways in intestinal epithelial cells.
apoptosis is a normal physiological form of cell death that plays a key role in both embryonic development and the maintenance of adult tissues. In adults, apoptosis is responsible for balancing cell proliferation and maintaining constant cell numbers in tissues undergoing cell turnover (25). Among all tissues, mammalian small intestinal epithelial cells have one of the most rapid turnover rates, a possible explanation for the low incidence of cancer in this tissue (17, 19). Therefore, an analysis of signal transduction cascades in apoptosis may help in the understanding of mechanisms responsible for the rapid turnover of small intestinal epithelial cells. Recent evidence implicates the small GTPase Rac1 in the regulation of apoptotic signaling pathways (1, 10, 24, 39, 40, 44, 52, 53).
Rac1 belongs to the Rho family of small GTPases and plays diverse roles in cellular function including ROS production, membrane ruffling, lamellipodia formation, the activity of transcriptional factors, cell cycle control, and the integrity of cell-cell adhesions (7, 23, 28, 29, 38, 45). Lores et al. (27) reported the first evidence showing that there is a high degree of thymus atrophy in mice expressing activated mutants of Rac protein, leading them to propose that Rac-dependent pathways might play an important role in apoptosis of T lymphocytes. This study resulted in many subsequent investigations of the role of Rac1 in apoptosis. It appears that whether Rac1 enhances or inhibits apoptosis is highly dependent on cellular context and/or the inducers of apoptosis (30, 53). The role of Rac1 in apoptosis of intestinal epithelial cells is unclear.
We have examined the function of polyamines (putrescine, spermine, and spermidine) in intestinal epithelial cells using a nontransformed line derived from rat crypt cells, the IEC-6 cell line (34). Depletion of polyamines with α-difluoromethylornithine (DFMO), which inhibits ornithine decarboxylase, the first rate-limiting enzyme in polyamine synthesis, causes a significant increase in the resistance of cells to apoptosis (2, 37). We have also found that the activity of Rac1 is significantly inhibited in polyamine-depleted cells (36, 47). Given this background, the present study tested the hypothesis that Rac1 plays a role in TNF-α-induced apoptosis in intestinal epithelial cells. We examined whether Rac1 is activated during apoptosis and whether inactivation of Rac1 either by treatment with the specific chemical inhibitor NSC-23766 or the expression of a dominant negative (DN)-Rac1 mutant prevents apoptosis. We tested whether inactivation of Rac1 prevents apoptosis by inhibiting the activation of JNK, which is required for TNF-α-induced apoptosis (2, 3). Furthermore, we determined the role of TNF-α-induced ERK1/2, p38 MAPK, and Akt activation in Rac1-mediated apoptosis. Taken together, our results demonstrate that Rac1 mediates TNF-α-induced apoptosis via JNK irrespective of ERK1/2 and Akt activation and plays a key role in the proapoptotic pathway in intestinal epithelial cells.
MATERIALS AND METHODS
Reagents.
Cell cultureware was purchased from Corning Glassworks (Corning, NY). Media and other cell culture reagents were obtained from Invitrogen (Carlsbad, CA). Dialyzed FBS, cycloheximide (CHX), and secondary antibodies conjugated to horseradish peroxidase were purchased from Sigma (St. Louis, MO). FuGENE 6 transfection reagent and the Cell Death Detection ELISA Plus kit were purchased from Roche Diagnostic (Indianapolis, IN). The ECL Western blot detection system was purchased from DuPont-New England Nuclear (Boston, MA). Caspase-3, -8, and -9 substrates were purchased from BioMol Research Laboratories (Plymouth Meeting, PA). Mouse anti-phospho-JNK1/2, rabbit anti-phospho-ERK1/2, rabbit anti-JNK1/2, rabbit anti-phospho-Akt, rabbit anti-Akt, rabbit anti-phospho-p38, rabbit anti-p38, and rabbit anti-caspase-3 antibodies were purchased from Cell Signaling (Beverly, MA). Mouse anti-Rac1 antibody was obtained from Upstate Biotechnology (Lake Placid, NY). Mouse anti-ERK1/2 was obtained from Zymed Laboratories (San Francisco, CA). Mouse anti-actin antibody was purchased from Santa Cruz Biotechnology (Santa Cruz, CA). TNF-α was obtained from Pharmingen (San Diego, CA). Rhodamine-phalloidin was purchased from Molecular Probes (Eugene, OR). NSC-23766 (Rac1 inhibitor), LY-294002 [phosphatidylinositol 3-kinase (PI3K)-Akt inhibitor], SP-600125 (JNK1/2 inhibitor), and SB-203580 (p38 MAPK inhibitor) were purchased from Calbiochem (La Jolla, CA). U-0126 (MEK inhibitor) was purchased from Promega (Madison, WI). The IEC-6 cell line (CRL 1592) was obtained from the American Type Culture Collection (Manassas, VA) at passage 13. The cell line was derived from the normal rat intestine and developed and characterized by Quaroni et al. (34). IEC-6 cells originate from intestinal crypt cells as judged by morphological and immunological criteria. They are nontumorigenic and retain the undifferentiated character of epithelial stem cells. Tests for mycoplasma were always negative. All chemicals were of the highest purity commercially available.
Cell culture.
IEC-6 cell stock was maintained in T-150 flasks in a humidified, 37°C incubator in an atmosphere of 90% air-10% CO2. The medium consisted of DMEM with 5% heat-inactivated FBS, 10 μg/ml insulin, and 50 μg/ml gentamicin sulfate. The stock was passaged weekly and fed three times per week, and passages 15–22 were used. During the experimental setup, cells were trypsinized with 0.05% trypsin and 0.53 mM EDTA and counted using a Beckman Coulter counter (model Z1).
Transfection.
IEC-6 cells were transfected with pMX-internal ribosome entry site (IRES)-green fluorescent protein (GFP)-V12-Rac1 [constitutively active (CA)], pMX-IRES-GFP-N17-Rac1 (DN), and pMX-IRES-GFP (vector). Stable clones were isolated and characterized as previously described (36).
Apoptosis studies.
Cells were plated (day 0) in T-75 flasks at a density of 6.25 × 104 cells/cm2 in DMEM-dialyzed FBS with triplicate samples for each group. Cells were fed on day 2. On day 3, the cell culture medium was removed and replaced with serum-free medium. On day 4, TNF-α (20 ng/ml) with CHX (25 μg/ml) was added to the serum-free medium for 3 h. After various treatments, images were photographed with a charge-coupled device camera using NIH Image software (version 1.58).
Caspase activity assay.
The protocol used for the caspase assay was similar to that described previously (2). In brief, cells were harvested and washed with cold Dulbecco’s phosphate-buffered saline (DPBS) buffer. The cell pellet was resuspended in ice-cold lysis buffer. After a 15-min incubation on ice, the lysate was centrifuged at 10,000 g at 4°C for 10 min. The supernatant was used for measurements of caspase-3, -8, and -9 activities. Each reaction (100 μl) contained 10 μl of cytosolic protein, 70 μl of assay buffer, and 20 μl of 2 mM Ac-DEVD-p-nitroanilide (pNA) for caspase-3, Ac-IETD-pNA for caspase-8, or Ac-LEHD-pNA for caspase-9 dissolved in assay buffer. The enzymatic reaction was carried out in 96-well plates at 37°C. Absorbance was read at 405 nm in a microplate reader. Protein was determined using the BCA method (Pierce, Rockford, IL), and caspase activity was expressed as picomoles of pNA released per milligram of protein per minute.
Quantitative DNA fragmentation ELISA.
The protocol for quantitative DNA fragmentation ELISA was similar to that described previously (2). In brief, cells were harvested, lysed in lysis buffer for 30 min, and centrifuged at 200 g for 5 min to pellet nuclei. An aliquot of the supernatant was incubated with immunoreagents (anti-histone-biotin plus anti-DNA peroxidase-conjugated antibody) in 96-well streptavidin-coated plates on a shaker for 2 h. After samples were washed with incubation buffer, 100 μl of substrate buffer were added to each well, and samples were incubated for an additional 5–10 min. Absorbance was read at 405 nm in a microplate reader. Triplicates of the samples were used to quantify protein by the BCA method from Pierce. DNA fragmentation was expressed as absorbance units per milligram of protein per minute.
Cell migration assay.
Cells were grown to confluence for 4 days in 35-mm plates, serum starved for 24 h, and then treated with or without 30 μM Rac1 inhibitor NSC-23766 3 h before the monolayer was wounded. Wounding was done by scratching with a gel-loading microtip as described previously (47). Immediately thereafter, scratches were photographed by a charge-coupled device camera using NIH Image software (version 1.58). The same fields were also photographed after 7 h to monitor the migration of cells.
Western blot analysis.
Cells were first washed with ice-cold DPBS and lysed for 10 min in ice-cold extraction buffer. Lysates were centrifuged at 10,000 rpm for 10 min at 4°C. Supernatants from 10 to 50 μg protein were TCA precipitated, eluted in 1× SDS sample buffer for 5 min, and separated by 10–15% SDS-PAGE. Proteins were transferred to Immunobilon-P membranes (Millipore, Bedford, MA) and probed with the indicated antibodies overnight at 4°C in TBS buffer containing 0.1% Tween 20 and 5% BSA or nonfat dry milk (blotting grade, Bio-Rad); all antibodies were diluted 1:1,000 with the exception of anti-caspase-3 at 1:1,500 and anti-actin at 1:5,000. Membranes were subsequently incubated with secondary antibody conjugated to horseradish peroxidase at room temperature for 1 h, and immunocomplexes were visualized by the ECL detection system. Quantitative analysis of the Western blots (densitometry) was carried out using NIH Image software. All data are expressed as means ± SE. All experiments were repeated three times.
Rac1 activation assay.
The biological activity of Rac1 protein was analyzed using pulldown assays performed as described previously (36, 47). In brief, GST-p21-activated kinase fusion protein (GST-PAK; Rac1-binding domain of human PAK residues 51–135) was prepared by lysing bacteria transformed with the GST-PAK plasmid construct in bacteria lysis buffer. The cell lysate was sonicated and clarified by centrifugation at 10,000 g for 15 min. The fusion protein was recovered by the addition of glutathione-agarose beads to the supernatant. The beads were washed three times in pulldown assay buffer and resuspended before the addition of cell lysates (150 μg). After being tumbled for 45 min at 4°C, beads were washed with pulldown assay buffer three times, and the amount of Rac1 protein bound to GST-PAK protein was analyzed by Western blot analysis. Proteins (20 μg) from each sample were resolved using SDS-PAGE to determine the level of total Rac1 protein.
Cell fractionation.
The protocol used for the cell fractionation assay was similar to that described previously (47). Cells were grown for 4 days and serum starved for 24 h before being harvested. Cell monolayers were washed twice with DPBS, harvested in ice-cold cell lysis buffer [containing (in mM) 50 HEPES (pH 7.5), 50 NaCl, 1 MgCl2, 2 EDTA, and protease inhibitors], homogenized with a Dounce homogenizer using a type B pestle, and centrifuged at 1,500 g for 10 min. The resulting supernatant was centrifuged at 15,000 g for 10 min. The pellet was resuspended in cell lysis buffer. Equal amounts of protein were used for the detection of Rac1 using Western blot analysis.
Fluorescence detection of actin filaments.
Cells were plated in 35-mm dishes, fixed with 3.7% formaldehyde, washed with PBS, permeabilized with 0.1% Triton X-100 for 5 min, blocked with 3% BSA for 20 min, and rinsed with 0.1% BSA for 20 min. Monolayers were stained with rhodamine-conjugated phalloidin for 1 h. Images were observed using a Nikon Diaphot inverted microscope with appropriate filters and processed with NIH Image.
Statistics.
Data are expressed as means ± SE. All experiments were repeated three times (n = 3). Western blots are representative of three experiments. ANOVA with appropriate post hoc testing was used to determine the significance of the differences between the means of multiple treatments, and Student's t-test was performed for measuring the significance of the differences between the means of two treatments. P < 0.05 was regarded as statistically significant.
RESULTS
TNF-α-mediated apoptotic signaling induces Rac1 activation.
Since GTP-bound Rac1, membrane-associated Rac1, and the formation of lamellipodia reflect the activation of Rac1, we analyzed these events in response to TNF-α. The level of GTP-bound Rac1 (active Rac1) was determined using a GST-PAK pulldown assay as previously described (36). As shown in Fig. 1A, TNF-α increased GTP-Rac1 levels transiently within 1 min (Fig. 1A, lane 2), which peaked between 10 and 30 min (Fig. 1A, lanes 4 and 5). To determine the levels of membrane-bound Rac1, a microsomal membrane fraction (heavy membrane) and whole cell extract were analyzed from untreated cells and those treated with TNF-α (Fig. 1B). TNF-α significantly increased membrane-associated Rac1 levels within 30 min and decreased them within 60 min without changes in the total amount of Rac1 protein. When Rac1 is activated, it interacts with downstream effectors and mediates cellular effects, such as lamellipodia formation. As shown in Fig. 1C, cells exposed to TNF-α for a short period of time (30 min) exhibited extensive lamellipodia accompanied by intense stress fiber formation. At a longer incubation time (60 min), these peripheral structures and stress fibers were gradually decreased. The concomitant activation of Rac1 during apoptosis suggests that Rac1 may modulate apoptotic signaling in these cells.
Fig. 1.Effect of TNF-α-cycloheximide (CHX) on the level of GTP-Rac1, membrane-associated Rac1, and lamelliodia formation. IEC-6 cells were grown in DMEM-5% dialyzed FBS (dFBS) and serum deprived for 24 h before treatment. A: cells were exposed to TNF-α (20 ng/ml) + CHX (25 μg/ml) for 0, 1, 5, 10, 30, 60, and 90 min. Cell extracts were prepared and subjected to a GST-p21-activated kinase (PAK) fusion protein pulldown assay as described in materials and methods, Rac1 activation assay. Equal amounts of protein were applied to each lane and subjected to electrophoresis by 12% SDS-PAGE. GTP-Rac1 and total Rac1 protein (∼21 kDa) were detected by anti-Rac1 antibody. Representative Western blots and values of densitometry readings are shown. Values are means ± SE for 3 independent experiments. B: cells were treated with TNF-α (20 ng/ml) + CHX (25 μg/ml) for 0, 30, and 60 min. Total cell extracts and membrane fractions were prepared, applied to each lane (10 μg for whole cell extracts and 30 μg for the membrane fraction), and subjected to electrophoresis by 12% SDS-PAGE. Rac1 was identified by anti-Rac1 antibody. Representative Western blots and values of densitometry readings are shown. Values are means ± SE for 3 independent experiments. C: cells were treated with TNF-α (20 ng/ml) + CHX (25 μg/ml) for 0, 30, and 60 min, washed with DPBS, fixed, and stained with rhodamine-conjugated phalloidin for the visualization of F-actin. Images are representative of 3 observations. Arrows indicate lamellipodia.
Inhibition of Rac1 significantly decreases TNF-α-induced apoptosis.
We examined the role of Rac1 in TNF-α-induced apoptosis by inhibiting its activation. We studied the effects of Rac1 inhibition on apoptosis by using a specific Rac1 inhibitor, NSC-23766. Gao et al. (14) recently identified this compound by structure-based virtual screening from the National Cancer Institute database. NSC-23766 fits the surface groove of Rac1 that is critical for Rac1-guanine nucleotide exchange factor (GEF) interaction. NSC-23766 has been tested in fibroblasts, human cancer cells (prostate cancer and breast cancer), myelin-forming cells, vascular smooth muscle cells, lung epithelial cells, and hematopoietic stem cells (6, 13, 14, 31, 41, 49). In these cell lines, NSC-23766 effectively blocks Rac1 activation and abolishes Rac1-mediated cellular events. However, its effectiveness in IEC-6 cells has not been tested. Therefore, we studied the effect of NSC-23766 on Rac1-mediated cellular events by examining wound healing, Rac1 activation, and lamelipodia formation. The results depicted in Fig. 2 demonstrate that pretreatment of the cells with 30 μM NSC-23766 for 3 h significantly inhibited migration (55%) compared with untreated cells. NSC-23766 significantly abolished the formation of lamellipodia compared with that in untreated cells exposed to TNF-α at different time points (Fig. 2C). In addition, NSC-23766 effectively inhibited both basal as well as TNF-α-induced Rac1 activation (Fig. 2D). The results shown in Fig. 2 clearly show that NSC-23766 effectively blocked Rac1-mediated cellular responses and confirmed the effectiveness of this compound for our study.
Fig. 2.Effect of a Rac1 inhibitor on cell migration, formation of lammellipodia, and Rac1 activity in response to TNF-α-CHX. IEC-6 cells were grown to confluence for 3 days in DMEM-5% dFBS and serum deprived for 24 h before treatment with 30 μM NSC-23766 (Rac1 inhibitor) for 3 h. A: after treatment, confluent monolayers of the control and NSC-23766-treated groups were wounded with a gel-loading tip, marked to localize the wound site, washed, and incubated with serum-free medium ± 30 μM NSC-23766. Plates were photographed immediately to record the wound width (0 h) and again at the marked wound location after 7 h of incubation. Images are representative of 3 observations. B: quantitative analysis of migration showing wound width covered compared with initial wound size (0 h) using NIH Image software analysis. Values are means ± SE. *P < 0.05 compared with the control group. C: control and NSC-23766-treated groups were exposed to TNF-α (20 ng/ml) + CHX (25 μg/ml) for 0 and 30 min, washed with DPBS, fixed, and stained with rhodamine-conjugated phalloidin for the visualization of F-actin. Images are representative of 3 observations. Arrows indicate lamellipodia. D: control and NSC-23766-treated groups were exposed to TNF-α (20 ng/ml) + CHX (25 μg/ml) for 0 and 15 min. Cell extracts were prepared and subjected to a GST-PAK fusion protein pulldown assay as described in materials and methods, Rac1 activation assay. Active (GTP-Rac1) and total Rac1 were detected by a Rac1 antibody. Representative Western blots and values of densitometry readings are shown. Values are means ± SE for 3 independent experiments.
Using our previously established model for TNF-α-induced apoptosis (2), we examined the effect of the inhibition of Rac1 on apoptosis. TNF-α caused detachment of cells, as evident by the appearance of refractory cells (arrows in Fig. 3Ab). Pretreatment of the cells with 30 μM NSC-23766 for 3 h followed by TNF-α exposure almost completely prevented the detachment, and cultures retained the morphological features of untreated monolayers (Fig. 3A, a and d). We further analyzed the effect of NSC-23766 on DNA fragmentation and capase-3 activation. TNF-α treatment significantly increased DNA fragmentation (4-fold) compared with untreated cells. Inhibition of Rac1 by NSC-23766 decreased TNF-α-induced DNA fragmentation by 50% (Fig. 3B). Caspase-3 activation as determined by a casapse-3 activity assay followed a pattern similar to DNA fragmentation (Fig. 3C). Because caspase-3 activation is the result of cleavage of procaspase-3, we determined the effect of NSC-23766 on procaspase-3 cleavage by Western blot analysis. Rac1 inhibition significantly decreased the formation of the caspase-3 active fragment (∼17 kDa) compared with the control group in response to TNF-α (Fig. 3D). Furthermore, NSC-23766 alone had no effect on either DNA fragmentation or capase-3 activation in the basal condition. These results suggest that inhibition of Rac1 protects intestinal epithelial cells from apoptosis by inhibiting the activation of caspase-3 and subsequently decreasing DNA fragmentation.
Fig. 3.Effect of a Rac1 inhibitor on apoptosis. IEC-6 cells were grown in DMEM-5% dFBS to confluence for 3 days. After being serum deprived for 24 h, cells were pretreated with 30 μM NSC-23766 and then incubated with TNF-α (20 ng/ml) + CHX (25 μg/ml) for 3 h in the presence of NSC-23766. A: apoptosis was measured by morphological analysis. a, Control; b, cells exposed to TNF-α-CHX for 3 h; c, cells exposed to NSC-23766 alone; d, cells pretreated with NSC-23766 and then exposed to TNF-α-CHX. B: apoptotic-induced DNA fragmentation was measured using a colorimetric ELISA kit as described in materials and methods. Values are means ± SE in optical density units (OD) read at 405 nm per milligram of protein per minute. *P < 0.05 compared with the respective control group treated with TNF-α-CHX. C: cell extracts from the above experiments were incubated with DEVD-p-nitroanilide (pNA), a colorimetric substrate of caspase-3, at 37°C for 4 h. Absorbance was read at 405 nm, and enzyme activity was calculated as picomoles of pNA released per minute per milligram of protein. Values are means ± SE. *P < 0.05 compared with the respective control group treated with TNF-α-CHX. D: cell extracts from the above experiments were collected in lysis buffer, applied to each lane (50 μg), and subjected to electrophoresis by 15% SDS-PAGE. The activated fragments of casapase-3 (∼17 kDa) were identified by caspase-3 antibody. Actin (∼42 kDa) immunoblotting was performed as an internal control for equal loading. Blots shown are representative of 3 observations.
Expression of DN-Rac1 prevents TNF-α-induced apoptosis.
We used CA-recombinant Rac1 and DN-recombinant Rac1 to validate our results from the Rac1 inhibitor study. IEC-6 cells were transfected with CA-Rac1, DN-Rac1, and vector DNA, and stably transfected clones were selected by limiting dilution (36). Selected clones transfected with CA-Rac1, DN-Rac1, and vector were characterized by determining the levels of GTP-Rac1. Cells expressing DN-Rac1 had significantly less GTP-Rac1 compared with vector- and CA-Rac1-expressing cells (Fig. 4A). As expected, CA-Rac1-expressing cells showed significantly higher levels of GTP-Rac1. In addition, recombinant Rac1-expressing cells had relatively more Rac1 protein compared with empty vector-transfected cells, indicating robust expression of these proteins. The above results confirmed the suitability of these clones for use in this study. Based on our inhibitor study, we predicted that DN-Rac1 expression should protect cells from apoptosis. CA-Rac1-transfected cells and vector-transfected cells showed relatively more refractory cells, indicating detachment (arrows in Fig. 4B, b and d) in response to the apoptotic stimulus, whereas untreated cells maintained a normal firmly adherent monolayer (Fig. 4B, a and c). TNF-α failed to induce detachment and morphological changes in DN-Rac1-transfected cells (Fig. 4Bf). These observations were further supported by DNA fragmentation (Fig. 4C). TNF-α induced a 3.5-fold increase in DNA fragmentation in vector-transfected cells compared with untreated cells. In CA-Rac1-transfected cells, TNF-α further increased DNA fragmentation, which was significantly higher compared with vector-transfected cells treated with TNF-α. As expected, expression of DN-Rac1 significantly decreased DNA fragmentation compared with both vector- and CA-Rac1-transfected cells.
Fig. 4.Effect of expression of stable constitutively active (CA)-Rac1 and dominant negative (DN)-Rac1 on apoptosis. A: IEC-6 cells transfected with empty vector (V), CA-Rac1, and DN-Rac1 plasmids were grown as described in materials and methods, Cell culture. Cell extracts were prepared and subjected to a GST-PAK fusion protein pulldown assay as described in materials and methods, Rac1 activation assay. Active (GTP-Rac1) and total Rac1 were detected by a Rac1 antibody. Representative Western blots and values of densitometry readings are shown. Values are means ± SE for 3 independent experiments. B: cells transfected with vector, CA-Rac1, and DN-Rac1 were grown to confluence for 3 days in DMEM-5% dFBS. After being serum deprived for 24 h, cells were incubated with TNF-α (20 ng/ml) + CHX (25 μg/ml) for 3 h. Apoptosis was measured by morphological analysis. a, Vector-transfected cell; b, vector-transfected cell exposed to TNF-α-CHX for 3 h; c, CA-Rac1-transfected cell; d, CA-Rac1-transfected cell exposed to TNF-α-CHX; e, DN-Rac1-transfected cell; f, DN-Rac1-transfected cell exposed to TNF-α-CHX. Arrows indicate apoptotic cells. C: cells transfected with vector, CA-Rac1, and DN-Rac1 were grown and exposed to apoptotic stimuli as described in B. DNA fragmentation was measured by using a colorimetric ELISA kit as described in materials and methods. Values are means ± SE. *P < 0.05 compared with TNF-α-CHX-treated cells transfected with vector.
Activation of caspase-8 and -9 is indicative of the involvement of death domain receptors and the mitochondrial pathway, respectively. It is also believed that the mitochondrial pathway is required for caspase-8 and -9 activation (8). Caspase-8 and-9 activation was significantly prevented in DN-Rac1-transfected cells compared with cells transfected with vector and CA-Rac1 (Fig. 5, A and B). CA-Rac1 expression significantly increased the activities of both caspase-8 and -9 compared with vector-transfected cells in response to TNF-α. The activation of caspase-8 and -9 leads to activation of caspase-3 (2). DN-Rac1 significantly prevented the activation of caspase-3 compared with the levels seen in cells transfected with vector and CA-Rac1 (Fig. 5C).
Fig. 5.Effect of expression of stable CA-Rac1 and DN-Rac1 on the activities of caspase-8, -9, and -3 after TNF-α-CHX treatment. Cells transfected with vector, CA-Rac1, and DN-Rac1 were grown to confluence for 3 days in DMEM-5% dFBS. After being serum deprived for 24 h, cells were incubated with TNF-α (20 ng/ml) + CHX (25 μg/ml) for 3 h. Apoptosis-induced activation of caspase-8 (A), -9 (B), and -3 (C) was determined by using specific substrates for the caspases. Values are means ± SE. *P < 0.05 compared with TNF-α-CHX-treated cells transfected with vector.
Inhibition of Rac1 prevents TNF-α-induced JNK activation.
Based on our previous study, which showed that TNF-α-induced JNK activation stimulates caspase-9 activation and, thereby, apoptosis in IEC-6 cells (2), it is highly possible that inhibition of caspase-3 and -9 by Rac1 inhibition is via altering the activation of JNK. Therefore, we determined the effect of Rac1 inhibition on JNK activation. During TNF-α-induced apoptosis, the phosphorylation of JNK1/2 increased in a time-dependent manner and then plateaued (Fig. 6A). Inhibition of Rac1 by NSC-23766 prevented TNF-α-induced JNK1/2 activation, as evident by the decreased phosphorylation of JNK1/2 (Fig. 6B). Interestingly, DN-Rac1 expression almost completely prevented TNF-α-induced JNK2 phosphorylation and significantly decreased JNK1 phosphorylation (Fig. 6C). JNK1/2 inhibition by a specific JNK inhibitor, SP-600125, or Rac1 inhibition by NSC-23766 significantly prevented TNF-α-induced DNA fragmentation compared with the control group (Fig. 6D). Taken together, these data clearly indicate that the inhibition of Rac1 protects cell from apoptosis via inhibiting TNF-α-induced JNK activation.
Fig. 6.Effect of inactivation of Rac1 on phosphorylation of JNK1/2. A: cells were grown for 3 days to confluence in DMEM-5% dFBS. After being serum deprived for 24 h, cells were incubated with TNF-α (20 ng/ml) + CHX (25 μg/ml) for 0, 5, 10, 30, 60, 180, and 300 min. Whole cell extracts were collected in lysis buffer, applied to each lane (30 μg), and subjected to electrophoresis by 12% SDS-PAGE. Phospho (p)-JNK1/2 (∼46/54 kDa) was identified by a specific antibody that recognizes p-JNK1/2 at Thr183/Tyr204. Total (T)-JNK1/2 (∼46/54 kDa) was determined by a JNK1/2 antibody. Blots shown are representative of 3 observations. B: cells were grown and serum deprived as described in A. After being preincubated with NSC-23766 (30 μM) for 3 h, cells were exposed to TNF-α (20 ng/ml) + CHX (25 μg/ml) for 3 h along with NSC-23766. p-JNK1/2 and T-JNK1/2 were identified by Western blot analysis as described in A. Blots shown are representative of 3 observations. C: cells transfected with vector, CA-Rac1, and DN-Rac1 were exposed to TNF-α (20 ng/ml) + CHX (25 μg/ml) for 3 h. p-JNK1/2 and T-JNK1/2 (∼46/54 kDa) were identified by Western blot analysis as described in A. Blots shown are representative of 3 observations. D: cells were exposed to TNF-α-CHX for 3 h in the absence and presence of NSC-23766 (30 μM) or SP-600125 (25 μM). Apoptosis was measured using a colorimetric ELISA kit as described in materials and methods. Values are means ± SE. *P < 0.05 compared with control cells exposed to TNF-α-CHX.
Inhibition of Rac1 does not alter TNF-α-induced ERK1/2 and Akt activation.
Previous studies (3, 4, 54) have shown that the MEK1/2-ERK1/2 and PI3K-Akt pathways are the two major prosurvival pathways in IEC-6 cells. Therefore, we examined whether inhibition of Rac1 attenuates apoptosis by modulating TNF-α-induced ERK1/2 and Akt activity. Inhibition of Rac1 by NSC-23766 had no effect on basal as well as TNF-α-induced ERK1/2 and Akt activation (Fig. 7A). In addition, TNF-α-induced activation of ERK1/2 or Akt was similar in CA-Rac1-, DN-Rac1-, and vector-transfected cells (Fig. 7B). Furthermore, prevention of TNF-α-induced ERK1/2 and Akt activation by U-0126 and LY-294002, respectively, significantly increased apoptosis as judged by DNA fragmentation. However, Rac1 inhibition by NSC-23766 significantly prevented TNF-α-induced DNA fragmentation in the presence and absence of these inhibitors. The inhibition of JNK1/2 by SP-600125 protected cells in a manner similar to NSC-23766 treatment in the presence and absence of U-0126 or LY-294002 (Fig. 8, A and B). These results indicate that inhibition of Rac1 mediates protection against TNF-α-induced apoptosis by inhibiting JNK1/2 independently of ERK1/2 and Akt activation.
Fig. 7.Effect of inactivation of Rac1 on phosphorylation of ERK1/2 and Akt. IEC-6 cells were grown to confluence for 3 days in DMEM-5% dFBS and serum deprived for 24 h. A: after being preincubated with NSC-23766 (30 μM) 3 h, cells were incubated with TNF-α (20 ng/ml) + CHX (25 μg/ml) for 3 h. Whole cell extracts were collected in lysis buffer, applied to each lane (15 μg), and subjected to electrophoresis by 12% SDS-PAGE. p-ERK1/2 (∼42/44 kDa) and p-Akt (∼60 kDa) were identified by a specific antibody that recognizes p-ERK1/2 (Thr202/Tyr204) and a specific antibody that recognizes p-Akt (Ser473). T-ERK1/2 (∼42/44 kDa) and T-Akt (∼60 kDa) were determined by the respective specific antibodies. Blots shown are representative of 3 observations. B: cells transfected with vector, CA-Rac1, and DN-Rac1 were exposed to TNF-α (20 ng/ml) + CHX (25 μg/ml) for 3 h. p-ERK1/2, p-Akt, T-ERK1/2, and T-Akt were identified by Western blot analysis as described in A. Blots shown are representative of 3 observations.
Fig. 8.Effect of Rac1 inhibition on apoptosis in the absense of ERK1/2 or Akt activation. IEC-6 cells were grown to confluence for 3 days in DMEM-5% dFBS and serum deprived for 24 h. A: cells were exposed to TNF-α-CHX for 3 h in the absence and presence of NSC-23766 (30 μM), SP-600125 (25 μM), U-0126 (10 μM), NSC-23766 + U-0126, and SP-600125 + U-0126. Apoptosis was measured using a colorimetric ELISA kit as described in materials and methods. Values are means ± SE. *P < 0.05 compared with control cells exposed to TNF-α-CHX. B: cells were exposed to TNF-α-CHX for 3 h in the absence and presence of NSC-23766 (30 μM), SP-600125 (25 μM), LY-294002 (10 μM), NSC-23766 + LY-294002, and SP-600125 + LY-294002. Apoptosis was measured using a colorimetric ELISA kit as described in materials and methods. Values are means ± SE. *P < 0.05 compared with control cells exposed to TNF-α-CHX.
p38 MAPK does not modulate TNF-α-induced apoptosis.
Recent studies have shown that TNF-α-induced p38 MAPK activation requires Rac1 in human dermal microvascular endothelial cells (33) and that p38 MAPK is a proapoptotic molecule in myocytes (26). We determined whether TNF-α-induced Rac1 activation mediates apoptotic signaling via p38 MAPK in intestinal epithelial cells. As shown in Fig. 9A, TNF-α dramatically increased the phosphorylation of p38 MAPK, resulting in its activation, and Rac1 inhibition did not alter the response (Fig. 9A). In addition, treatment of cells with a p38 MAPK-specific inhibitor, SB-203580, did not attenuate TNF-α-induced DNA fragmentation compared with the control group (Fig. 9B). These data suggest that TNF-α-induced p38 MAPK activation does not mediate apoptosis and that Rac1-mediated effects on apoptosis are independent of p38 MAPK activation.
Fig. 9.Effect of Rac1 inhibition on phosphorylation of p38 MAPK and effect of a p38 MAPK inhibitor on TNF-α-induced DNA fragmentation. IEC-6 cells were grown to confluence for 3 days in DMEM-5% dFBS and serum deprived for 24 h. A: after being preincubated with NSC-23766 (30 μM) 3 h, cells were incubated with TNF-α (20 ng/ml) + CHX (25 μg/ml) for 3 h. Whole cell extracts were collected in lysis buffer, applied to each lane (20 μg), and subjected to electrophoresis by SDS-PAGE. p-p38 MAPK (∼43 kDa) was identified using a phospho-specific antibody [1:1,000 in Tris-buffered saline (TBS) solution with 5% BSA]. A specific antibody (1:1,000 in TBS solution with 5% BSA) was used for the measurement of T-p38 (∼43 kDa). Blots shown are representative of 3 observations. B: cells were exposed to TNF-α (20 ng/ml) + CHX (25 μg/ml) for 3 h in the absence and presence of NSC-23766 (30 μM), SB-203580 (10 μM), or NSC-23766 + SB-203580. Apoptosis-induced DNA fragmentation was measured using a colorimetric ELISA kit as described in materials and methods. Values are means ± SE. *P < 0.05 compared with control cells exposed to TNF-α-CHX.
DISCUSSION
Previous observations from our laboratory have shown that polyamine depletion inhibits apoptosis in cultured small intestinal epithelial cells (IEC-6 cells) by preventing the activation of JNK (2). It also significantly inhibits the activation of Rac1, a key regulator of cell migration (36), but whether Rac1 plays a role in apoptosis is unclear. In view of the above results, we hypothesized that Rac1 is involved in apoptosis and mediates JNK activation.
Switching between an inactive GDP-bound form and an active GTP-bound form, Rac1 plays a central role in signaling pathways related to cytoskeletal organization, cell migration, proliferation, and apoptosis (1, 5, 10, 18, 35, 46). In the resting stage, GDP-bound Rac1 resides in the cytosol as a complex with a Rho guanine nucleotide dissociation inhibitor (RhoGDI). After stimulation, Rac1 is released from RhoGDI and translocates to the membrane, where GTP is exchanged for GDP by GEFs such as Tiam1 (14).
Esteve et al. (12) showed that expression of DN-Rac1 protected the U937 leukemic cell line from TNF-α-induced apoptosis; however, Deshpande et al. (9) reported that DN-Rac1 increased the susceptibility of endothelial cells to TNF-α-induced apoptosis. These controversial results suggest that the role of Rac1 in TNF-α-induced apoptosis is highly dependent on cellular context. However, none of these studies examined the effect of TNF-α on Rac1 activity. Geijsen et al. (15) reported that TNF-α is not able to induce Rac activation in human neutrophils, suggesting that TNF-α does not activate Rac1 in all cell lines. Therefore, in this study, we first determined Rac1 activity during TNF-α-induced apoptosis in IEC-6 cells. We have shown that TNF-α alone does not induce apoptosis and that it requires the inhibition of protein synthesis (2). We used CHX with TNF-α to suppress the synthesis of short-lived members of the inhibitors of apoptosis group of proteins. Rac1 regulates the formation of lamellipodia in a large number of cell lines including IEC-6 cells (36). Wojciak-Stothard et al. (50) reported that TNF-α activated Rac1 as assessed by actin cytoskeleton changes. They demonstrated that quiescent human endothelial cells incubated with TNF-α for 5–30 min increased the formation of lamellipodia. With respect to IEC-6 cells, the levels of GTP-Rac1 began to increase within 1 min and achieved a peak at about 30 min in response to TNF-α (Fig. 1A). The increased level of membrane-associated Rac1 (Fig. 1B) following incubation with TNF-α for a short time (30 min) indicated that it does translocate from the cytosol to the membrane system for its activation. Exposure to TNF-α for 30 min led to increased formation of lamellipodia and stress fibers (Fig. 1C). Taken together, these results indicate that Rac1 is activated during TNF-α-induced apoptosis in IEC-6 cells.
Next, we established the proapoptotic role of Rac1 in TNF-α-induced apoptosis by using a Rac1-specific inhibitor and expression of DN-Rac1. DN-Rac1 (threonine to asparagine substitution at codon 17) lacks the ability to bind GTP and downstream effectors and inhibits Rac1 activity by competing with GEFs for endogenous Rac1. Inactivation of Rac1, either by treatment with Rac1 inhibitor or expression of the DN-Rac1 mutant, protected IEC-6 cells from apoptosis. Furthermore, expression of the CA-Rac1 mutant enhanced apoptosis, as indicated by the morphology, DNA fragmentation, and caspase-3 activity results (Figs. 3–5). These findings are consistent with results from others who have demonstrated that Rac1 is a proapoptotic molecule in response to different types of apoptotic stimuli in a variety of cell types. For example, overexpression of wide-type Rac1 protein accelerates apoptosis in chondrocytes during bone development (48), and the expression of DN-Rac1 protects H-Ras-transformed human breast epithelial cells from capsaicin-induced apoptosis (22). Moreover, in human hepatoma cells, overexpression of DN-Rac1 prevents growth factor deprivation-induced apoptosis (21). Eom et al. (11) also reported that overexpression of DN-Rac1 prevents UV-induced apoptosis in Rat-2 fibroblasts, and, recently, Ito et al. (20) demonstrated that β-adrenergic receptor stimulation induces apoptosis in rat ventricular myocytes in a Rac1-dependent manner. It is important to note that the CA-Rac1 mutant had no effect on apoptosis, caspase activation, and DNA fragmentation in the absence of apoptotic stimuli compared with DN-Rac1- and vector-transfected cells, suggesting that Rac1 by itself does not induce apoptosis. However, the rapid activation of Rac1 by TNF-α and enhancement of apoptosis in CA-Rac1-transfected cells indicate that Rac1 is a key modulator of receptor-mediated proapoptotic signaling pathways in intestinal epithelial cells.
As a “molecular switch,” Rac1 has been implicated in the regulation of several signaling pathways related to apoptosis, including MAPK pathways (ERK1/2, JNK1/2, and p38 MAPK) and the PI3K-Akt pathway (30, 32, 52). The role of Rac1 in these signaling cascades is highly dependent on the cellular context and apoptotic stimulus. JNK is a key regulator of many cellular events, including epithelial cell shape changes during dorsal closure, proliferation, and apoptosis (25, 51). Our previous studies (2, 3) have indicated that JNK1/2 mediates TNF-α-induced apoptosis by promoting cytochrome c release, subsequently activating caspase-9, and sustained activation of ERK1/2 inhibits JNK activity in IEC-6 cells. In fact, whether Rac1 mediates JNK activation in response to apoptotic inducers including TNF-α is still debatable. Minden et al. (28) reported that Rac1 mediates JNK activity in response to EGF but not to TNF-α; Su et al. (43) also reported that Cdc42 rather than Rac1 mediates JNK activation during Fas ligand-related apoptosis in human leukemia HL-60 cells. Our current study indicated that inhibition of Rac1 was associated with a strong inhibition of the activation of JNK1/2 (Fig. 6, B and C) and nearly prevented the activation of caspase-9, whose activity is regulated by JNK1/2 during TNF-α-induced apoptosis in IEC-6 cells (Fig. 5B). In addition, TNF-α-induced ERK1/2, Akt, and p38 MAPK activities are independent of Rac1. Moreover, in the absence of TNF-α-induced ERK1/2 and Akt activation, the Rac1 inhibitor effectively prevented apoptosis similar to the effect of the JNK1/2 inhibitor. This indicates that Rac1 inhibition mediated the protection against TNF-α-induced apoptosis by inhibiting JNK1/2 independently of ERK1/2 and Akt (Figs. 7–9). These findings are consistent with the results of several studies (5, 16, 18, 42, 46) showing that Rac1 mediates the JNK apoptotic pathway in oligodendrocytes, thyroid cells, COS-1 cells, Jurkat cells, and human breast cancer cells.
In summary, our present work showed that TNF-α induced lamellipodia formation, increased levels of membrane-associated Rac1, and increased GTP-Rac1 levels, indicating that Rac1 is activated during the induction of apoptosis by TNF-α. Inactivation of Rac1 by treatment with either a specific inhibitor, NSC-23766, or expression of DN-Rac1 prevented TNF-α-induced apoptosis by blocking the activation of JNK. Inhibition of Rac1 protected cells from apoptosis in the absence of ERK1/2 or Akt activation, and Rac1 did not modulate the TNF-α-induced activation of p38 MAPK. Based on our present findings and earlier observations (summarized in Fig. 10), we conclude that Rac1 is an upstream positive modulator of TNF-α-induced JNK activation and plays an important role in the biological regulation of apoptosis in normal small intestinal epithelial cells. Future efforts will focus on elucidating the molecular mechanisms through which Rac1 regulates JNK-mediated apoptosis.
Fig. 10.Schematic representation of the Rac1-mediated proapoptotic signaling pathway and prosurvival signaling pathways in IEC-6 cells. Rac1, ERK1/2, Akt, JNK1/2, and p38 are activated during TNF-α-induced apoptosis. Among them, the small GTPase Rac1 mediates the activation of JNK1/2. Activated JNK1/2 activates caspase-9 via promoting cytochrome c release (2), leading to activation of caspase-3, DNA fragmentation, and apoptosis. CA-MEK1/2 inhibits the activation of JNK1/2 (3). Akt inhibition by LY-294002 increases the activity of caspase-9 (4). The mechanisms through which Rac1 regulates JNK activation and apoptosis warrant further investigation.
GRANTS
This study was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-16505 and the Thomas A. Gerwin Endowment.
FOOTNOTES
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
We sincerely acknowledge Mary Jane Viar for critically reading the manuscript and sincerely thank Mary Jane Viar, Dr. Rajivkumar J. Vaidya, Dr. Sujoy Bhattacharya, Dr. Wenlin Deng, Dr. Huazhang Guo, and Rebecca L. West for technical support.
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