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1 Department of Pediatrics, Division of Hematology/Oncology, Stanford University School of Medicine, Stanford, CA and
2 Department of Biological Sciences, Stanford University, Stanford, CA
Requests for reprints: Louie Naumovski, Department of Pediatrics, Division of Hematology/Oncology, Stanford University School of Medicine, CCSR Rm. 1215, Stanford, CA 94305-5149. Phone: (650) 723-5113; Fax: (650) 736-0195. E-mail: naumovsk{at}stanford.edu
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Abstract |
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, and the ER stressors brefeldin A, tunicamycin, and thapsigargin. Analysis of the caspase proteolytic cascade reveals that ARMER inhibits proteolysis of the caspase-9-specific fluorogenic substrate LEHD-AFC as well as endogenous substrates downstream of caspase-9; however, it does not inhibit cytochrome c release or cleavage of caspase-9 itself. Apoptotic stimuli cause endogenous levels of ARMER protein and RNA to decrease, leading to cell death; however, sustaining ARMER protein levels through exogenous expression inhibits apoptosis. These data suggest that ARMER is a novel ER integral membrane protein which protects cells by inhibiting caspase-9 activity and reveal a possible role for ARMER in cell survival.
Key Words: apoptosis • endoplasmic reticulum • KIAA0069 • ARL6ip
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Introduction |
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TopNotesAbstractIntroductionResultsDiscussionMaterials and MethodsReferences |
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A central mechanism in the apoptotic pathway is the activation of caspases, a family of cysteine aspartic acid-specific proteases (5). Caspases are synthesized as zymogens which require specific cleavage for activation. Caspases involved in apoptosis can be divided into two groups: initiator caspases (caspase-8, -9, and -12); and effector caspases (caspase-3, -6, and -7) (6). Through binding specific co-factors recruited by apoptotic stimuli, initiator caspases become cleaved and thus activated, and in turn activate downstream effector caspases, which cleave an array of cellular proteins, resulting in the morphological and biochemical hallmarks of apoptosis (6). The three main apoptotic pathways identified thus far (the mitochondrial, ER stress, and death receptor pathways) are activated by caspase-9, -12, and -8, respectively (7). The mitochondrial pathway may also serve as a point of convergence for the other two pathways, since death receptor activation and ER stress can both lead to cytochrome c release from the mitochondria and subsequent caspase activation (8).
Once released, cytochrome c initiates formation of a high-molecular-weight caspase-activating complex, the apoptosome, composed of cytochrome c, an adaptor protein Apaf-1, and caspase-9. Caspase-9 is cleaved at the apoptosome and recruits and cleaves caspase-3, the primary executioner caspase, which then cleaves many cellular proteins, including poly-ADP ribose polymerase (PARP) and vimentin, a major component of intermediate filaments (7, 9). Mice deficient in caspase-9 exhibit expansion and protrusion of brain tissue caused by defective apoptosis in neural progenitor cells. In addition, caspase-9-/- thymocytes are resistant to apoptotic stimuli which signal through the mitochondrial pathway, such as dexamethasone- and 
irradiation-induced death (10). Proteins which can modulate caspase-9 activity, such as inhibitor of apoptosis proteins (IAPs), can inhibit apoptosis induced by a variety of stimuli, such as tumor necrosis factor (TNF), staurosporine, taxol, and growth factor withdrawal (11). Thus, caspase-9 and proteins that can modulate caspase-9 activity may play central biological roles in the pathogenesis of diseases such as stroke, neurodegenerative diseases, and cancer.
In this study, we report on a novel protein, apoptotic regulator in the membrane of the endoplasmic reticulum (ARMER). We show by immunofluorescence that ARMER is localized in the ER and by membrane-partitioning assays that it fractionates with membrane proteins. Overexpression of ARMER under control of a ponasterone-inducible promoter protects HT1080 fibrosarcoma cells from apoptosis induced by all stimuli tested, including serum deprivation, doxorubicin, UV irradiation, TNF-, brefeldin A, tunicamycin, and thapsigargin. Analysis of the caspase proteolytic cascade reveals that ARMER inhibits proteolysis of substrates downstream of caspase-9; however, it does not affect cytochrome c release or cleavage of caspase-9 itself. Apoptotic stimuli cause endogenous levels of ARMER protein and RNA to decrease, leading to cell death; however, sustaining ARMER protein levels through exogenous expression inhibits apoptosis. Thus, our results suggest that ARMER may be a novel modulator of caspase-9 activity and have possible implications in cancer and other diseases involving aberrant apoptosis.
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Results |
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Full-length human and mouse sequences show extensive homology with four conservative amino acid substitutions and one less conservative change (F v. A) (Fig. 4). An almost full-length rat sequence diverges by one amino acid from the mouse (not shown). Partial ESTs derived from zebrafish, Xenopus laevis, and Brugia malayi were also found to have extensive homology. Significant but more distant homology was also found in a Drosophila melanogaster EST (Fig. 4).
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ARMER Is an Integral Membrane Protein Co-Localizing With the ER Marker Calreticulin
We used biochemical methods to determine if ARMER was an ER integral membrane protein as suggested from computer analysis of the protein sequence. Fractionation of post-nuclear homogenates (H) of F-ARMER-expressing cells into membrane (M) and soluble fractions (S) showed that the vast majority of F-ARMER was present in the membrane fraction (Fig. 5A, lanes labeled fractionation). Triton X-114 partitioning studies revealed that ARMER is a hydrophobic protein because it was found in the Triton X-114 phase (L) (Fig. 5A, lanes labeled TX-114). Finally, carbonate extraction of membrane fractions demonstrated that F-ARMER was retained in the membrane (P) (Fig. 5A, lanes labeled sodium carbonate), consistent with an integral membrane protein (17). In each case, F-ARMER fractionated with calnexin, a protein known to be in the ER membrane, and not calreticulin, a protein known to be in the ER lumen.
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ARMER Protects Against Apoptosis Induced by Multiple Stimuli
Since our findings strongly suggested that ARMER localizes to the ER, we hypothesized that ARMER may also be able to protect cells against apoptosis induced by ER stressors. Brefeldin A, an inhibitor of ER-Golgi transport, tunicamycin, a specific inhibitor of N-glycosylation in the ER, and thapsigargin, an inhibitor of the ER Ca2+-ATPase, are all ER stressors as evidenced by induction of grp78 and grp94 (18). To test the ability of ARMER to inhibit apoptosis from ER stressors, we grew pIND F-ARMER and pIND LacZ cells in the presence or absence of ponA induction, then treated them with the ER stressors brefeldin A, tunicamycin, or thapsigargin, as well as the chemotherapy drug doxorubicin, TNF- plus cycloheximide, and serum starvation. Doxorubicin is an anthracycline antibiotic used to treat a variety of malignancies through many proposed but uncertain mechanisms, including DNA synthesis inhibition and inhibition of DNA repair (19). TNF- alone does not kill HT1080 cells (20); however, TNF- plus cycloheximide causes apoptosis by binding to cell surface TNF receptors and activating caspase-8 (21). Serum starvation deprives cells of the necessary growth factors required to sustain cell cycle progression and viability. Cells were harvested between 1 and 4 days after apoptosis induction and analyzed for sub-G0 DNA content. F-ARMER expression significantly protected cells against each apoptotic agent (Fig. 6A). LacZ expression did not have any significant effect on cell viability (data not shown), indicating that apoptotic protection was a specific effect of F-ARMER expression.
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To further determine at which point ARMER inhibited the apoptotic pathway, we studied the caspase cascade. Various apoptotic inducers have been suggested to converge on mitochondrial cytochrome c release, which leads to caspase-9 activation. Having shown that cytochrome c release was unaffected by ARMER, we next measured amounts of caspase-9 cleavage by Western blotting using antibodies directed to cleaved caspase-9. pIND F-ARMER and pIND LacZ cells were grown in the absence or presence of ponA, then deprived of serum or treated with brefeldin A or doxorubicin. One to 3 days after induction of apoptosis, cells were collected for quantitative Western blotting. In addition to measuring cleaved caspase-9, we also monitored downstream events, such as cleavage of vimentin, caspase-3, and PARP. In healthy, untreated cells, cleaved caspases were not detectable (Fig. 7, Day 0). On treatment with apoptotic stimuli, cleaved caspase-9 was detectable to the same extent in both F-ARMER-expressing (ponA +) and non-expressing cells (ponA -) (Fig. 7). However, cleavage of substrates downstream of caspase-9 was inhibited by F-ARMER. Cleavage of caspase-3, a direct target of activated caspase-9, was inhibited when F-ARMER was expressed (Fig. 7). In turn, cleavage of caspase-3 substrates, including PARP and vimentin, was also inhibited in F-ARMER-expressing cells. Vimentin was recently identified as a substrate that can be cleaved by both caspase-3, yielding a Mr 47,000 fragment, and also by caspase-9, yielding a Mr 28,000 fragment (7). F-ARMER inhibited cleavage of vimentin by both caspase-3 and caspase-9 (Fig. 7). LacZ expression did not affect cleavage of any of these caspase substrates (data not shown). These data demonstrate that ARMER inhibits the caspase proteolytic cascade downstream of caspase-9 cleavage.
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24,000) slightly lower than F-ARMER, corresponding to endogenous ARMER (Fig. 9A). Using this antibody, we determined by co-immunofluorescence that endogenous ARMER co-localized with calreticulin, indicating that ARMER resides in the ER (data not shown). We also used this antibody to monitor the expression of ARMER in response to apoptotic stimuli. On treatment with brefeldin A or thapsigargin, cells exhibited a decrease in endogenous ARMER protein (Fig. 9B). Northern blots revealed that endogenous ARMER RNA expression is also decreased in cells treated with apoptotic stress. In cells inducibly expressing F-ARMER, however, F-ARMER protein and RNA expression persisted (Fig. 9, B and C), suggesting that F-ARMER expression may be maintaining viability in the absence of endogenous ARMER. Note that F-ARMER RNA migrates at a lower molecular weight than endogenous ARMER RNA because the pIND F-ARMER plasmid contains a truncated version of the endogenous RNA lacking a portion of the 3' untranslated region.
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6-fold compared to cells grown in the absence of ponA (Fig. 10A). pIND ARMER and pIND LacZ cells were grown in the absence or presence of ponA, then deprived of serum or treated with tunicamycin or doxorubicin, and harvested for FACS analysis for sub-G0 DNA content. Cells overexpressing ARMER exhibited less apoptosis than cells not overexpressing ARMER (Fig. 10B). These results indicate that the Flag epitope tag used in experiments described in this study does not affect the biological function of ARMER.
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Discussion |
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TopNotesAbstractIntroductionResultsDiscussionMaterials and MethodsReferences |
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Our study shows that ARMER inhibits apoptosis, possibly by modulating the protease activity of cleaved caspase-9. Western blot analysis showed that caspase-9 is cleaved in response to serum starvation, brefeldin A, and doxorubicin, supporting the theory that various apoptotic pathways may converge at a point upstream of apoptosome formation. Although ARMER did not affect release of cytochrome c into the cytosol or cleavage of caspase-9, events downstream of caspase-9 cleavage were inhibited in cells overexpressing ARMER. These events include the cleavage of vimentin and of the primary executioner caspase, caspase-3, as well as its substrate, PARP. In addition, cleavage of the exogenous caspase-9-specific fluorogenic substrate LEHD-AFC in extracts prepared from ARMER-expressing cells was also inhibited. These results suggest that ARMER, directly or indirectly, inhibits the proteolytic capacity of cleaved caspase-9.
We tested the ability of microsomes containing F-ARMER to inhibit the cleavage activity of recombinant human active caspase-9 in a fluorogenic assay using LEHD-AFC as substrate. F-ARMER was unable to inhibit recombinant caspase-9 activity in this context (data not shown), suggesting that inhibition of caspase-9 by F-ARMER may be indirect and may require other molecular components in the cell.
Our results support the contention that cleaved caspase-9 is not necessarily proteolytically active (7). For example, IAPs can physically interact with cleaved caspase-3, -7, and -9, and block their activity (25). IAPs inhibit the activities of caspases either by directly binding to and blocking the catalytic sites of cleaved caspases or functioning as ubiquitin ligases to promote the proteolysis of caspases once bound to them (25). IAPs contain homologous domains called baculoviral IAP repeat (BIR) domains that are necessary to inhibit caspase activity. The third BIR domain of XIAP specifically inhibits caspase-9 while the second BIR domain along with some flanking sequences was sufficient to inhibit caspase-3 and -7 (11). Expression of XIAP, as well as its family members c-IAP-1, c-IAP-2, and survivin were examined in our laboratory, and levels were found to be similar in cells overexpressing F-ARMER compared to uninduced cells (data not shown). Thus, the mechanism by which ARMER protects cells from apoptosis does not appear to be through up-regulating protein expression of IAP family members. However, it remains possible that ARMER may enhance IAP function, perhaps by facilitating IAP binding and inhibition of their caspase substrates, or may function in a manner similar to the IAPs.
ER stress stimulates cleavage of caspase-12 in the mouse as detected by a specific monoclonal antibody directed against the mouse protein (18). Interestingly, the human orthologue of mouse caspase-12 remains elusive and has not been clearly defined (26). Fischer et al. (27) recently described a potential human homologue (68% identical to mouse caspase-12), but the gene contains severe mutations causing truncations in several splice variants and destruction of the SHG box, a crucial element for caspase enzymatic activity, precluding its function as an active caspase. Neither monoclonal nor polyclonal antibodies directed against the human caspase-12 have been described in the literature or are commercially available. We have attempted to use a number of different antibodies raised against mouse caspase-12 to show cleavage of the putative human caspase-12 but have been unsuccessful in detecting any cleavage products after ER stress. Therefore, analysis of caspase-12 cleavage in our experimental system will not be possible until the human gene is identified and antibody reagents are available.
Endogenous ARMER was detected by Western blotting in numerous cell lines (data not shown). By co-immunofluorescence, endogenous ARMER was determined to co-localize with the ER protein, calreticulin, indicating that endogenous ARMER localizes to the ER, as would be predicted based on its KKNE ER retrieval sequence. The finding that endogenous ARMER and overexpressed F-ARMER both co-localize with calreticulin suggests that the Flag epitope tag does not affect the localization of ARMER. The epitope tag also did not affect the function of ARMER since we demonstrated that regulated overexpression of native ARMER also protected cells against serum starvation, tunicamycin, and doxorubicin.
Our studies showing that overexpression of epitope-tagged or native ARMER inhibits apoptosis suggests but does not prove that endogenous ARMER has the same function. However, our observation that endogenous ARMER decreases when cells are challenged with apoptotic stimuli suggests that ARMER is involved in apoptosis. The first possibility is that apoptotic stimuli down-regulate ARMER, and that lack of anti-apoptotic ARMER function then contributes to cell death. However, in cells expressing F-ARMER from an exogenous promoter, viability is maintained since F-ARMER levels do not drop. The second possibility is that ARMER expression decreases as a consequence of cell death. Our results do not support this latter possibility since endogenous ARMER still decreases in F-ARMER-expressing cells that are resistant to cell death.
The high conservation of ARMER across species suggests that ARMER function will be amenable to genetic analysis. Recent studies have identified a mouse homologue of ARMER (91% identity to human ARMER), which was isolated in a yeast two-hybrid screen using ARL-6, a small ADP-ribosylation-like factor that may be involved in intracellular protein transport, as bait (28). The mouse homologue of ARMER was also recently identified as a gene down-regulated during differentiation of myeloid progenitor cells (29). Neither of these studies provided biological significance; therefore, the relevance of these findings is currently unknown.
In summary, our studies describe a novel ER membrane protein, ARMER, which protects human HT1080 fibrosarcoma cells from apoptosis induced by various agents. ARMER inhibits the ability of cleaved caspase-9 to cleave its substrates, both endogenous and exogenous, suggesting that ARMER may, directly or indirectly, inhibit the activity of cleaved caspase-9. The mechanism by which ARMER inhibits cleaved caspase-9 activity is the subject of current investigation in our laboratory. Although ARMER inhibits events downstream of caspase-9 cleavage, that may not be its only mechanism of action in inhibiting apoptosis. We have not definitively identified the mechanism of ARMER action; however, we note that the mechanism of action of another anti-apoptotic protein, Bcl-2, remains controversial even after more than 15 years of intense study (30).
Proteins with anti-apoptotic activity, such as ARMER, may contribute to the etiology of cancer or our inability to treat the disease with cytotoxic therapy. Since ARMER has anti-apoptotic activity, we would expect to find ARMER up-regulated in malignancies either at presentation or after relapse. Indeed, the gene for ARMER (KIAA0069) is more highly expressed in colon adenocarcinomas than in adenomas or normal colon tissues (31) and is highly expressed in highly proliferating breast tumors (32). The significance of these findings, suggesting a potential role for ARMER in cancer, is being investigated. Further characterization of ARMER will provide valuable insight with potential implications in malignant, neurodegenerative, and ischemic disorders.
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Materials and Methods |
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To establish inducible cell lines, HT1080 cells were co-transfected with pIND F-ARMER or pIND ARMER and pVgRXR receptor plasmid using calcium phosphate precipitation of DNA. The media were changed the next day. The following day, transfected cells were serially diluted into 10-cm tissue culture plates and selected with G418 (1.0 mg/ml) and Zeocin (0.5 mg/ml). Colonies that formed after 2 weeks were transferred into individual wells of a 24-well plate for further expansion (33). Some cells from each well were transferred to another 24-well assay plate and incubated for 48 h in media containing 2.5 µM ponA, an ecdysone analogue. Cells were solubilized with SDS loading buffer and expression of F-ARMER or native ARMER was detected by Western blotting using an anti-Flag antibody or an anti-ARMER antibody. Purity was assessed by immunofluorescence. As a control, a cell line inducibly expressing ß-galactosidase was constructed in a similar manner using the pIND LacZ and pVgRXR plasmids (Invitrogen). Colonies expressing ß-galactosidase were screened using X-gal as a substrate.
Survival Assays
Cells were grown in the presence of ponasterone (2.5 µM) or ethanol alone for 3 days and then treated with apoptotic stimuli as specified in the text. Survival was assayed at the times indicated by combining media and cells harvested using trypsin/EDTA. Cell pellets were washed with PBS with 5 mM EDTA and fixed with 70% ethanol. Cell pellets were treated with RNaseA (0.2 mg/ml for 30 min) and stained with propidium iodide (40 µg/ml) before analysis on a Becton Dickinson FACScan (Becton Dickinson, Franklin Lakes, NJ). Ten thousand cells were assayed for sub-G0 DNA. Hoechst staining of treated cells and immunofluorescence analysis revealed condensed and fragmented nuclei consistent with apoptosis (not shown).
For statistical analysis, percentages of non-apoptotic cells were compared between cells expressing and not expressing F-ARMER using Student's independent two-tailed t test to determine statistical significance. Each experiment was performed in triplicate.
Homogenization, Triton X-114, and Sodium Carbonate Extraction
Cells were collected and resuspended in 1.5 ml of homogenizing buffer (HB: 0.25 M sucrose, 10 mM Tris-HCl (pH 7.4), 1 mM DTT, 1 mM EDTA, and protease inhibitors). A Dounce homogenizer with a tightly fitting pestle was used for homogenization, and cell disruption was evaluated by phase contrast microscopy. The homogenate was separated into membrane fraction (M) and supernatant (S) by centrifuging at 100,000 x g for 1 h. The membrane fraction (120 µg of protein) was suspended in 1 ml of ice-cold Tris-buffered saline. Pre-condensed Triton X-114 (0.2 ml) was added and the suspension was incubated on ice for 30 min with occasional mixing. The suspension was centrifuged at 16,000 x g for 15 min at 4°C, and the pellet (P) was collected. The supernatant was transferred to a fresh tube and incubated in a 37°C water bath until the solution became cloudy. It was centrifuged at 16,000 x g for 10 min at room temperature. The upper aqueous phase (U) and the lower detergent phase (L) were collected and the proteins were precipitated with TCA (final concentration 10%). After incubation on ice for 30 min, the precipitated proteins were pelleted by centrifuging at 10,000 x g for 5 min. The pellets were washed twice with acetone and dissolved in protein loading buffer.
The integral membrane and non-integral membrane proteins in the membrane fraction were separated with sodium carbonate (17). Specifically, 60 µg of membrane proteins were diluted 50-fold in 100 mM sodium carbonate (pH 11.5) and incubated on ice for 30 min or longer. The suspension was centrifuged at 240,000 x gmax for 1 h at 4°C. The pellet (P) was directly dissolved in 1x protein loading buffer. The supernatant (S) was precipitated with TCA as described above.
Immunofluorescence Localization of Protein Expression
Cells were plated in chamber slides and grown in the presence or absence of ponA (2.5 µM) for 3 days. Cells were fixed in 2% paraformaldehyde in PBS for 15 min and permeabilized for 15 min with 0.5% Triton X-100 in PBS. After washing with PBS, slides were blocked with 5% BSA in PBS for 30 min at room temperature. F-ARMER was visualized with mouse anti-Flag monoclonal antibody, and endogenous ARMER was visualized with mouse anti-ARMER monoclonal antibody followed by goat anti-mouse FITC (green fluorescence). In co-immunofluorescence experiments, calreticulin was visualized with rabbit anti-calreticulin polyclonal antibodies followed by goat anti-rabbit Alexa 594 (red fluorescence). Coverslips were mounted to slides with an aqueous anti-fade mounting solution. A Nikon ECLIPSE E800 microscope with a VFM epi-fluorescence attachment was used to visualize cells. Images were captured using a Spot RT Slider camera and Spot RT v3.0 software.
Preparation of Cytosolic Fractions
For cytochrome c releasing assays, cells were homogenized as described above. After removing nuclei by centrifuging at 800 x g for 8 min, the homogenate was separated into the mitochondria-containing membrane fraction and cytosolic fraction by centrifuging at 100,000 x g for 1 h. Cytosolic fractions were analyzed by Western blotting for cytochrome c and cleaved caspase-3. Purity of fractions was confirmed by Western blotting for cytochrome c oxidase II (data not shown).
Quantitative Western Blotting
Cells were lysed in triple-detergent lysis buffer [50 mM Tris-Cl (pH 8.0), 150 mM NaCl, 0.1% SDS, 0.5% deoxycholic acid, 1.0% NP40, and a cocktail of protease inhibitors] on ice for 10 min. After centrifugation at 15,000 x g for 5 min, supernatants were quantitated and equal quantities of protein were resolved on the appropriate percentage SDS-polyacrylamide gels. Gels were transferred to polyvinylidene difluoride membrane using a Bio-Rad Semi-dry Transfer Cell (Bio-Rad, Hercules, CA). Western blotting was performed using primary and horseradish peroxidase-conjugated secondary antibodies specified in the text. Anti-Flag and -vimentin antibodies were purchased from Sigma Chemical Co., St. Louis, MO. Anti-cytochrome c antibody was purchased from Santa Cruz Biotechnology, Santa Cruz, CA. Antibodies to caspases and PARP specifically recognized the cleaved forms (Cell Signaling Technologies, Beverly, MA). Anti-ARMER antibodies were generated against a KLH-coupled peptide containing the first 20 amino acids of ARMER by Antibody Solutions (Palo Alto, CA). Protein bands were detected using SuperSignal Chemiluminescent Substrate (Pierce, Rockford, IL). All membranes were blotted with an anti-tubulin antibody (Sigma) to control for loading and transfer. Bands were imaged and quantitated in the linear range and normalized to tubulin using the Bio-Rad Fluor-S MultiImager.
Quantitation of Caspase-9 Enzymatic Activity
Caspase-9 activity was assayed using the Caspase-9/Mch6 Fluorometric Protease Assay Kit (MBL, Japan). Cells were harvested, rinsed in cold PBS, and lysed, and supernatants were quantitated. One hundred fifty micrograms cell lysate were analyzed according to the manufacturer's protocol. Reactions were incubated in a reaction mixture containing LEHD-AFC (0.1 mM) at 37°C for 2 h, and fluorescence levels were determined at 2-min intervals at an excitation of 400 nm and emission of 512 nm using a fluorescence plate reader. The Vmax was obtained using 30 data points in the linear range of the reaction.
Northern Blotting
Total RNA was isolated using TRIzol reagent (Life Technologies, Inc., Carlsbad, CA) and dissolved in diethyl pyrocarbonate-treated H2O. Using standard techniques, 15 µg total RNA were resolved on a 1% formaldehyde-agarose gel, transferred to nylon membrane, and hybridized with a 32P-labeled ARMER cDNA probe using Rapid-hyb buffer (Amersham, Piscataway, NJ). The probe was prepared by isolating and purifying a BglII fragment containing ARMER from the pIND ARMER plasmid. The probe was labeled with random primers extended by Klenow fragment in the presence of deoxynucleotide triphosphates and [32P]dCTP using Ready-To-Go DNA Laddering Beads (Amersham). Blots were imaged and in the linear range of X-ray film and reprobed with 18S cDNA as a control.
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Received November 15, 2002; revised March 20, 2003; accepted April 3, 2003.
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: effects on proliferation of normal and transformed cells in vitro. Science, 230: 943–945, 1985.This article has been cited by other articles:
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Y. Takahashi, L. Li, M. Kamiryo, T. Asteriou, A. Moustakas, H. Yamashita, and P. Heldin Hyaluronan Fragments Induce Endothelial Cell Differentiation in a CD44- and CXCL1/GRO1-dependent Manner J. Biol. Chem., June 24, 2005; 280(25): 24195 - 24204. [Abstract] [Full Text] [PDF]
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, pIND LacZ; 
, pIND F-ARMER clone 1; 
, pIND F-ARMER clone 2. B. ARMER expression protects serum-deprived cells over time. Clone 1 was grown in the absence or presence of ponA for 3 days and treated with serum deprivation. Cells were harvested at 0, 2, 4, and 6 days. Points, means of one representative of three independent experiments, each done in triplicate at each time point; bars, SD.
