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Institute of Human Nutrition [D. X.], Herbert Irving Comprehensive Cancer Center [J-W. S., A. D., J-T. Y., I. B. W.], Department of Anatomy and Cell Biology [G. G. G., A. F. P.], College of Physicians and Surgeons, Columbia University, New York, New York 10032; American Health Foundation, Valhalla, New York 10595 [J. T. P.]; and E. Wolfson Medical Center, Holon, 58100 Israel [H. S.]
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ABSTRACT |
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 Top ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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MTs are major cellular structural components. They play important roles in the development and maintenance of cell shape, cell replication and division, cell signaling, and cellular movement. MTs are composed of a backbone of 
- and ß-tubulin heterodimers and smaller amounts of MAPs. They are in an unstable steady state of a highly dynamic process of polymerization and depolymerization, which can be influenced by numerous factors (3)
. Certain cancer chemotherapy agents exert their antitumor actions by disrupting the dynamics of MT assembly, thus perturbing the formation and function of the mitotic spindle apparatus and arresting cells in mitosis. These agents are called MIAs. For example, paclitaxel (Taxol) binds to and stabilizes MTs, whereas Colcemid and Vinca alkaloids bind to and depolymerize MTs (4)
. An in-depth understanding of the molecular mechanisms by which these and other MIAs act may enhance their use in both cancer prevention and chemotherapy.
Because we found that a fraction of the SAMC-treated cells are specifically arrested in mitosis, in the present study we examined the effects of this compound on MTs. We found that SAMC can act directly to cause MT depolymerization and thereby disrupt the MT cytoskeletal network in interphase cells and also interfere with spindle formation in mitotic cells. We also obtained evidence that the activation of JNK1 and caspase-3 play important roles in SAMC-induced apoptosis. These novel effects of SAMC provide new insights into the molecular mechanisms responsible for the antitumor effects of this compound and may encourage further application of this and related compounds to cancer treatment and prevention.
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MATERIALS AND METHODS |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Chemicals and Antibodies.
The selective JNK1 inhibitor SP, p38 inhibitor SB, MEK1/2 inhibitor PD, and general caspase inhibitor VAD were purchased from Calbiochem (La Jolla, CA). The fluorogenic tetrapeptide substrates of caspase-1 (Z-YVAD-AFC), caspase-6 (Ac-VEID-AMC), and caspase-9 (Ac-LEHD-AFC) were purchased from Calbiochem. The fluorogenic substrates of caspase-3 (Ac-DEVD-AFC) and caspase-8 (Ac-IETD-AFC) were purchased from PharMingen (San Diego, CA). Primary antibodies used in Western blotting for ERK1/2, phospho-ERK1/2, p38, and phospho-p38 were obtained from Cell Signaling Technology (Beverly, MA), and the antibody to actin was obtained from Sigma-Aldrich Biotechnology (St. Louis, MO). Paclitaxel (Taxol) and cytochalasin D were obtained from Sigma, and Colcemid (N-methyl-N-deacetyl-colchicine) was obtained from Roche (Indianapolis, IN).
Cell Lines and Culture Conditions.
The human colon cancer SW480 and mouse fibroblast NIH3T3 cell lines were obtained from the American Type Culture Collection (Manassas, VA). The cultures of WT MEF cells (JNK+/+), MEF JNK1-/- cells, and MEF JNK2-/- cells were established from mutant mice with the respective genotypes (5)
and were generously provided by Dr. Zigang Dong (University of Minnesota, Austin, MN). SW480 cells and MEF cells were grown in DMEM with 10% fetal bovine serum, and NIH3T3 cells were grown in DMEM with 10% calf serum (Life Technologies, Inc., Grand Island, NY). Cells were incubated in a 100% humidified incubator at 37°C with 5% CO2. All of the biochemical studies were performed with exponentially growing cells at 50% confluence.
Cell Cycle Analysis.
PI staining was used to analyze DNA content, as reported previously (6)
. After the desired time of exposure to the indicated compounds, both adherent and floating cells were harvested, fixed with 70% ethanol, incubated with PI (0.05 mg/ml) and 1 µg/ml RNase A (Sigma) at room temperature in the dark for 30 min, and analyzed by flow cytometry, using a FACS Calibur instrument with a CellQuest software (Becton Dickinson, San Jose, CA). The apoptotic cells were considered to constitute the sub-G1 population, and the percentage of nonapoptotic cells in each phase of cell cycle was determined. All of the experiments were performed at least twice and yielded similar results.
Apoptosis Assay.
The percentage of cells undergoing apoptosis was determined by Annexin V-PE assay, as described previously (6)
. Briefly, after the cells were treated with either PBS (control) or SAMC (300 µM) for the desired time, both adherent and floating cells were harvested and double stained with Annexin V-PE (stained apoptotic cells) and 7-AAD (stained dead cells; PharMingen). The percentage of apoptotic cells was then determined by flow cytometric analysis. All of the experiments were performed three times, independently, and the results are expressed as mean ± SD.
Transfection and Flow Cytometric Analyses.
SW480 cells were transfected with either the control vector pcDNA3 or pcDNA3-DN-JNK1, by a liposome-mediated method using the Lipofectin Reagent (Invitrogen Life Technologies, Inc., Carlsbad, CA). The pcDNA3-DN-JNK1 plasmid encodes a FLAG epitope-tagged, kinase-inactive dn JNK1 and was generated from a pCMV-DN-JNK1 plasmid by subcloning and neomycin selection (7)
. The pCMV-DN-JNK1 was kindly provided by Dr. Audrey Minden (Ref. 8
; Columbia University, New York, NY). A CMV promoter-driven farnesylated-EGFP construct, pEGFPF, was cotransfected as a selectable marker for transfected cells (9)
. The pEGFPF plasmid was kindly provided by Dr. Wei Jiang (The Salk Institute, La Jolla, CA). After a 24-h transfection, the cells were treated with either PBS (control) or SAMC (300 µM) for the indicated times (18, 24 or 48 h). Both adherent and floating cells were harvested, double stained with Annexin V-PE and 7-AAD, and analyzed by flow cytometry. The early apoptotic transfected cells were determined as the 7-AAD-negative, GFP-positive and Annexin V-PE-positive cells, and the percentage in live transfected cells (7-AAD-negative, GFP-positive cells) was calculated. Aliquots of the cells were also stained with PI after treatment with SAMC, and the cell cycle distribution among transfected cells was analyzed by DNA flow cytometry.
Protein Extraction and Western Blotting.
The methods for protein extraction and Western blot analysis have been described previously (6)
. Briefly, after treatment with the indicated compounds for the desired time, both adherent and floating cells were collected and cell lysates were prepared. Protein concentrations were determined using the Bio-Rad Protein assay (Bio-Rad, Richmond, CA). Protein samples, 50 µg each, were subjected to SDS-PAGE, transferred to nitrocellulose membranes (Millipore, Bedford, MA), and blocked with 5% milk protein. Membranes were then incubated with the indicated primary antibody for 1 h or overnight, depending on the antibody, and were incubated with the corresponding horseradish peroxidase-conjugated secondary antibody for 1 h. Protein-antibody complexes were detected by an enhanced chemiluminescence system (Amersham, Piscataway, NJ). Immunoblotting for actin was performed to verify equivalent amounts of loaded protein. Results were analyzed on a Macintosh computer using the public domain NIH Image program (developed at the United States NIH and available on the Internet at http://rsb.info.nih.gov/nih-image/).
Indirect Immunofluorescence Microscopy.
The immunostaining method was essentially as described by Yoon et al. (10)
, with some modifications. Briefly, cells were grown on glass coverslips in 6-well plates (35-mm diameter). After exposure to PBS (control), 150 µM SAMC, or SAC for the indicated time period, the cells were washed once in DMEM at 37°C and were fixed with methanol at -20°C for 10 min. Fixed cells were washed three times with TBS for 10 min and were incubated with TBS, 10% normal goat serum, (Vector Labs, Burlingame, CA) plus the proper concentration of the primary antibody, as described below, for 1 h at 37°C in a 100% humidified chamber. After three washes with TBS, the cells were incubated with TBS and 10% normal goat serum plus the appropriate secondary antibody, as described below, for 45 min at 37°C in a 100% humidified chamber. After incubation, the cells were washed three times with TBS and refixed with 4% formaldehyde for 10 min. Coverslips were rinsed with tap water, mounted with Histomount (Vector Labs), and sealed to the microscope slide with clear nail polish. The slides were observed with a Nikon Optiphot microscope, and images were captured with a MicroMax cooled CCD (charge-coupled device) camera (Kodak KAF 1400 chip; Princeton Scientific Instruments, Monmouth Junction, NJ) using Metamorph software (Universal Imaging, Downingtown, PA).
Cells were stained for MTs with a rat monoclonal antibody against tyrosinated -tubulin (YL1/2; 1:10 dilution; Ref. 11
) and visualized with a goat antirat IgG-FITC conjugate (1:100; Chemicon International Inc., Temecula, CA). Cells were stained for centrosomes with a rabbit antipericentrin antibody (1:200; Babco, Richmond, CA) or Golgi complex with a rabbit anti-Giatin antibody (1:200) and were visualized with a goat antirabbit IgG-rhodamine conjugate (1:200), and were also stained with 4',6-diamidino-2-phenylindole (DAPI; 1:1000) to visualize DNA.
Tubulin Turbidity Assay.
In vitro MT and tubulin assembly were monitored by the turbidity assay. Fifty µl of 5 mg/ml pure tubulin (Cytoskeleton, Denver, CO) was assembled to a steady state in G-PEM buffer [100 mM PIPES (pH 6.9), 1 mM EGTA, 1 mM MgCl2, and 1 mM GTP] plus 10% glycerol in 96-well plate (0.33 cm2/well), by incubation at 37°C for 20–30 min. This is referred to as the "polymer stock." The tested compounds were prepared in G-PEM buffer at 3.5 times the indicated final concentrations and prewarmed to 37°C. Twenty µl of the test-compound solution was pipetted into the polymer stock wells and the plate was incubated at 37°C for an additional 20–30 min. The effects on polymerization/depolymerization of the tested compounds were quantitated by measuring the increase/decrease of the absorbance at 340 nm (A340) over time, using a Spectramax O.D. Reader and SOFTmax PRO 3.0 software (Molecular Devices, Sunnyvale, CA) in a kinetic model (one reading/min). Similar assays were also performed with MAP-rich tubulin (Cytoskeleton) instead of pure tubulin at 1 mg/ml in G-PEM buffer, without glycerol. In all of the assays the absorbance at 340 nm of the tested compound itself was subtracted from the total absorbance. Results are presented as a percentage of absorbance, with 100% representing the A340 value at 20 or 30 min, when the tubulin polymerization reached a steady state.
JNK Activity Assay.
A nonradioactive kinase assay was used according to the instruction manual of Cell Signaling Technology, Inc. Briefly, after the cells were treated with PBS (negative control) or SAMC at the indicated concentrations and times, the media were removed, and the cells were rinsed once with ice-cold PBS and scraped off the plate. Cells were also exposed to a germicidal UV lamp (254 nm, 30 W, 70-cm distance between plates and the UV lamp) in a cell culture hood for 30 min as a positive control for activating JNK. The UV dose was 
40 J/m2 (12)
. Cell pellets were then suspended in 500 µl of M2 lysis buffer [20 mM Tris (pH 7.6) containing 0.5% NP40, 250 mM NaCl, 3 mM EDTA, 3 mM EGTA, 1 mM DTT, 1 µg/ml leupeptin, 0.02 M ß-glycerophosphate, 1 mM sodium orthovanadate and 1 mM phenylmethylsulfonyl fluoride]. JNK1 was immunoprecipitated by incubation of 200 µg of this cell protein extract with 2 µg (20 µl) of c-Jun fusion protein beads (Cell Signaling), with gentle rocking overnight at 4°C. The precipitates were washed twice with M2 buffer and twice with kinase buffer [25 mM Tris (pH 7.5) containing 10 mM MgCl2, 5 mM ß-glycerophosphate, 2 mM DTT, and 0.1 mM Na3VO4]. Pellets were suspended in 50 µl of kinase buffer supplemented with 100 µM ATP and were incubated at 30°C for 30 min. The kinase reaction was terminated by adding 25 µl of 3x SDS sample buffer, and the reaction mixtures were boiled for 5 min. Samples were analyzed by Western blotting using a phospho-c-Jun (Ser63) antibody (1:1000 dilution; Cell Signaling).
Caspase Activity Assay.
After treatment with SAMC or SAC (300 µM) for the indicated times, the cells were washed twice with ice-cold PBS and harvested and extracts prepared as described previously (2)
. Caspase activity was determined using a fluorometric assay. Briefly, 10 µg of total protein were incubated with 2 µg of the fluorogenic peptide substrates Z-YVAD-AFC, Ac-DEVD-AFC, Ac-VEID-AMC, Ac-IETD-AFC, or Ac-LEHD-AFC, the fluorogenic substrate for caspase 1, -3, -6, -8-, or -9, respectively, in a 200-µl caspase buffer at 37°C for 3 h. The release of AFC or AMC was measured with a Gemini Fluoro/Luminometer (Molecular Devices) at an excitation wavelength of 400 or 360 nm and an emission wavelength of 505 or 460 nm, respectively. Results are presented as the fold increase of caspase activity, as compared with that of the untreated control cells.
Statistical Analysis.
Data are expressed as mean ± SD. Comparisons between control cells and drug-treated cells were made using Student’s t test. A difference between groups with P < 0.05 were considered statistically significant.
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RESULTS |
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-tubulin antibody and a goat antirat IgG-FITC conjugate indicated that treatment of NIH3T3 cells with 150 µM SAMC (the IC50 concentration for growth inhibition) for 24 h caused extensive MT depolymerization and disruption of the MT network in interphase cells (Fig. 1B)
. In addition, treatment with SAMC interfered with the normal spindle formation in M-phase cells. There was a marked increase of mitotic cells with monopolar spindles, which contained a ring of condensed chromosomes and a monopolar spindle pole in the center with astral MTs irradiating from it (Figs. 1E
and 2F
). Multipolar spindles were also observed in some of the SAMC-treated mitotic cells, together with abnormal chromosome alignments (Fig. 1H)
. Bipolar spindles, which were present in control cells (Fig. 1, D and G)
, were hardly seen in SAMC-treated cells. As expected from our previous studies (2)
, SAC (150 µM) did not show any effect on MTs or mitotic spindles in these cells (Fig. 1, C, F, and I)
. A time course study indicated that the depolymerization effect of SAMC on MTs occurred as early as 10 min after treatment (Fig. 2B)
. Using antibodies to pericentrin or Giatin, we observed additional effects of SAMC on cells. Treatment of interphase cells with SAMC caused fragmentation of the centrosome (Fig. 2, B and C)
. It also disrupted the distribution of the Golgi complex (Fig. 2E)
, when compared with the control cells in which the Golgi stacks were gathered juxtanuclearly (Fig. 2D)
. As expected (14)
, the Golgi elements were dispersed throughout the cytoplasm in untreated mitotic cells, and this was also seen in SAMC-treated mitotic cells (Fig. 2F)
. Studies with SW480 cells indicated that SAMC produced effects on MT depolymerization in interphase cells and abnormal spindle formation in mitotic cells, similar to those described above with NIH3T3 cells (data not shown).
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. The addition of 300 µM SAMC caused about a 50% decrease in tubulin polymerization (Fig. 3B
, curve 2), and an even greater effect was seen with 1 mM SAMC (Fig. 3B
, curve 4). We were surprised to find that in these in vitro assays with pure tubulin, SAC had an effect on MT depolymerization similar to that obtained with SAMC (Fig. 3B
, curves 1 and 3), although in living cells, SAC did not inhibit cell proliferation (2)
and had no detectable effect on MT depolymerization (Fig. 1)
. Because of this apparent discrepancy, we conducted in vitro turbidity assays with MAP-rich tubulin, to more closely resemble the situation in intact cells. In this system, SAC still induced MT depolymerization (Fig. 3C
, curves 1 and 2), but the effect was much weaker than that obtained with an equivalent concentration of SAMC (Fig. 3C
, curve 3 and 4). These findings suggest that the MAPs partially protect MT from the depolymerizing effect of SAC, although other factors probably also play a role in explaining why SAC lacks cytotoxicity in cultured cells (see "Discussion").
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, ß-ME (1 mM) itself had no significant effect on tubulin polymerization (curve 1). SAMC still caused a decrease in tubulin polymerization in the presence of 1 mM ß-ME, but this effect was markedly reduced. Tubulin polymerization was decreased by 49% after the addition of 300 µM SAMC alone (Fig. 3B
, curve 2), but was decreased by only 12% after the addition of 300 µM SAMC and 1 mM ß-ME (Fig. 3D
, curve 2). One mM SAMC decreased tubulin polymerization by 74% (Fig. 3B
, curve 4), whereas it decreased tubulin polymerization by only 35% when added together with 1 mM ß-ME (Fig. 3D
, curve 3). These data suggest that there is competitive binding between SAMC and ß-ME to thiol (-SH) groups in tubulin, and provide evidence that thiol groups of amino acids in tubulin (i.e., cysteine residues) may be the binding sites for SAMC (see "Discussion").
SAMC Induces JNK1 Activation.
Activation of JNK1 plays an important role in mediating the apoptosis induced by various toxic agents, including the MT-stabilizing agent Taxol and the MT-depolymerizing agent vinblastine (12)
. To investigate the signaling pathways involved in SAMC-induced apoptosis, we measured stress-activated protein kinase (SAPK)/JNK activity in extracts of SAMC-treated SW480 cells using a nonradioactive in vitro kinase assay. The cells were also treated with 40 J/m2 UV as a positive control (Fig. 4)
. Treatment of cells with 300 µM SAMC induced a rapid (within 15 min after treatment) and strong (3.5-fold increase) activation of JNK, which persisted for up to 12 h, with fluctuations in intensity during this interval including a decrease at 6 h (Fig. 4A)
. Additional studies, including the transient decrease at 4–6 h (data not shown), confirmed these results.
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. We found that pretreatment of cells with 100 µM SP also caused about a 50% inhibition of SAMC (300 µM)-induced apoptosis when assayed at 24 h after the addition of SAMC (Fig. 5, A and B)
. It is of interest that this effect was highly time dependent because pretreatment with 100 µM SP caused only 20% inhibition at 48 h, and no inhibition of apoptosis occurred at 72 h (Fig. 5B)
. By contrast, during this time course within the nonapoptotic population, pretreatment of cells with 100 µM SP did not inhibit the G2-M arrest induced by SAMC and, in fact, enhanced it (Fig. 5C)
. The fact that SP did not inhibit the accumulation of SAMC-treated cells in G2-M is consistent with our evidence that SAMC induces cells to arrest in M phase by directly binding to tubulin and causing MT depolymerization (Fig. 3)
. Therefore, the latter effects are independent of JNK1 activation.
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, expression of dn-JNK1 in these cells inhibited SAMC-induced apoptosis, and this inhibition was highly time-dependent. At 18 h, SAMC treatment caused about a 2-fold increase in apoptosis in the vector-transfected cells, and at this early time point, the apoptosis induced by SAMC was almost completely abolished by the expression of dn-JNK1. However, at the 24 and 48 h time points, the expression of dn-JNK1 inhibited SAMC-induced apoptosis by only 50 and 25%, respectively. These results are consistent with the data obtained in Fig. 5B
with the SP JNK inhibitor, thus providing further evidence that JNK1 plays a more important role in mediating the early phase than the late phase of apoptosis induced by SAMC. The dn-JNK1-transfected cells were also stained with PI and analyzed by flow cytometry for cell cycle distribution. Expression of dn-JNK1 did not inhibit the G2-M arrest induced by SAMC (data not shown).
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, at both the 18 and 24 h time points, the extent of apoptosis induced by SAMC in the JNK1-/- cells was significantly less than that in the WT MEF cells. However, there was no significant difference between these two cell lines with respect to the extent of apoptosis induced at 48 h. Thus, a lack of JNK1 activity makes these cells relatively resistant to SAMC-induced apoptosis at 18 and 24 h, but not at 48 h. By contrast, SAMC-induced apoptosis was not reduced in the JNK2-/- cells at 18, 24, or 48 h, and was actually enhanced, especially at 48 h, when compared with the WT MEF cells (Fig. 6B)
. DNA flow cytometry indicated that all three types of MEF cells displayed the same extent of arrest in G2-M after treatment with SAMC (data not shown).
Taken together with the data in Fig. 5
, these findings suggest that SAMC induces apoptosis in SW480 cells through two different pathways: one occurs early after treatment with SAMC (within 24 h) and is mediated at least in part by JNK1 signaling, and the other occurs later (at 48–72 h) and is presumably a consequence of abnormal spindle assembly and M phase arrest. It is of interest that JNK1 and JNK2 appear to play different roles in this process (see Discussion).
Activation of ERK1/2 and p38 Are Not Required for SAMC-Induced Apoptosis.
To investigate the possible roles of other MAPKs in SAMC-induced apoptosis, we measured the protein levels of ERK1/2 and p38 and the extent of their phosphorylation in SAMC-treated SW480 cells. Treatment of cells with SAMC (300 µM) caused a rapid (within 15–30 min) increase in the levels of both phospho-ERK1/2 and phospho-p38 without changing their total protein levels (Fig. 7A)
. To assess the functional significance of the activation of these two signaling kinases, we used specific inhibitors of MEK1/2 (PD) and p38 (SB; Ref. 21
). MEK1/2 are upstream kinases of ERK1/2; therefore, the inhibition of MEK1/2 should result in the inhibition of the activation of ERK1/2. Cells were pretreated for 4 h with increasing concentrations (10–100 µM) of either PD or SB, and then were exposed to SAMC (300 µM) for 24 h. Pretreatment with 50 or 100 µM PD markedly inhibited SAMC-induced ERK1/2 phosphorylation, and pretreatment with 20–100 µM SB inhibited the formation of phospho-p38 (Fig. 7B)
. However, when we pretreated the cells with PD or SB at the concentrations that inhibited ERK1/2 or p38 phosphorylation, neither compound inhibited SAMC-induced apoptosis at 24 h (Fig. 7C)
or at 48 h (data not shown). These results suggest that, in contrast to JNK1 activation (Figs. 4
and 5
), ERK1/2 and p38 activation do not play critical roles in SAMC-induced apoptosis.
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, when SW480 cells were treated with SAMC (300 µM), caspase-3-like activity increased within 24 h and continued to rise until 70 h. However, during this time course, there was no significant increase in the activities of caspase-1, -6, -8, or -9 (Fig. 8A)
. Western blot analysis confirmed the activation of caspase-3 by SAMC and indicated that this was associated with the cleavage of poly(ADP-ribose) polymerase (PARP). SAC (300 µM) exhibited no significant effect on caspase activity using both methods (data not shown).
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, VAD attenuated SAMC-induced apoptosis by about 50% at both 24 and 48 h. In the SAMC-treated cells, pretreatment with VAD reduced the percentage of sub-G1 cells from 30 to 15% at 24 h, and from 44 to 22% at 48 h. This inhibition of apoptosis by VAD was similar at the 50- and 100-µM doses (Fig. 8B)
. These results demonstrate that both the early and late phases of SAMC-induced apoptosis in SW480 colon cancer cells are caspase dependent, although it is not apparent why the VAD compound did not produce 100% inhibition of apoptosis. As expected, this compound did not inhibit SAMC-induced arrest of the cell cycle in G2-M (data not shown). |
DISCUSSION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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MIAs can be subdivided into classes based on their binding sites or domains on tubulin: the colchicine-binding site, the Vinca alkaloid-binding site, the paclitaxel-binding site, tubulin sulfhydryl groups, and uncharacterized binding sites (17)
. Our results indicate that SAMC interacts directly with tubulin and that MAPs may assist in this interaction (Fig. 3)
. Furthermore, the sulfhydryl reactive agent ß-ME inhibited the ability of SAMC to cause MT depolymerization in in vitro turbidity assays (Fig. 3D)
, suggesting that one or more sulfhydryl (-SH) groups in tubulin form a covalent bond with SAMC. It is known that the thiol groups on cysteine residues in tubulin are highly reactive (17)
. There are 12 cysteine residues in -tubulin and 8 in ß-tubulin. These cysteine residues are sensitive reporters for determining the conformation of tubulin, and thiol-disulfide exchanges between them may be a key regulator of ligand interactions and MT assembly (27)
. Although not fully understood, oxidation or covalent interactions of certain key thiol groups (i.e., Cys239 or Cys354) can cause structural modifications of tubulin, inhibit MT assembly, and also interfere with the formation of the mitotic spindle (17
, 28, 29, 30)
. Additional studies are required to demonstrate the direct formation of disulfide bonds between SAMC and one or more specific cysteine residues in tubulin.
In addition to these in vitro and in vivo effects on MT depolymerization, our study revealed other cellular effects of SAMC. SAMC induced centrosome fragmentation within 10 min after exposure of cells to this compound (Fig. 2B)
. The centrosome serves as the organization center for MTs. This fragmentation may account for the formation of multipolar spindles during mitosis in SAMC-treated cells (Fig. 1H)
. A recent report (31)
indicates that monopolar spindles are induced by both MT-stabilizing and -depolymerizing agents, but that multipolar spindles are induced only by MT-stabilizing drugs (such as Taxol, epothilone B, and discodermolide), but not by MT-depolymerizing agents (such as colchicine, nocodazole, and vinblastine). Chen and Horwitz (31)
suggested that the aberrant mitosis in cells with multipolar spindles may explain the higher sensitivity of cells to MT-stabilizing agents. However, our study indicates that even though SAMC is a MT-depolymerizing agent, it induced multipolar spindles, perhaps because of the unique mechanism by which it disrupts MT structure. It is well known that a close relation exists between the Golgi elements and MTs (14)
. We found that SAMC induced the break-up of Golgi stacks and caused their dispersal within the cytoplasm of interphase cells, an effect similar to that produced by nocodazole (14)
. Because an important function of the Golgi complex is the processing, sorting, and transport of membrane and luminal proteins after their synthesis, the disruption of the Golgi apparatus by SAMC may contribute to the cytotoxicity of this compound.
Our findings may also be relevant to the antitumor effects of other garlic-derived compounds. Allicin (diallyl thiosulfinate), the major ingredient of crushed garlic, has been shown to inhibit the proliferation of human breast, endometrial, and colon cancer cells (32) . However, allicin is rapidly metabolized, both in vitro and in vivo, and could not be detected in human blood or urine even after a large oral dose (33) . When allicin was incubated with cysteine at a physiological temperature and pH, under conditions mimicking those in the intestinal tract, it was found that allicin reacted with cysteine in less than one min and yielded two moles of SAMC per mol of allicin (34) . Other garlic-derived compounds, including diallyl trisulfide, diallyl disulfide, and ajoene, were also transformed to SAMC in this in vitro model system (34) . Furthermore, allicin can also react rapidly with glutathione- or thiol-containing proteins to produce SAMC (35 , 36) . These studies suggest that after the consumption of garlic, SAMC can be a major metabolic product in the intestinal tract and could, therefore, reach a high local concentration in the intestinal mucosa, thus enhancing its effects on colon cancer prevention. A recent study found that an aqueous garlic extract inhibited cell growth and markedly arrested the cell cycle at G2-M in HT-29 human colon cancer cells. Thus, 89% of the treated cells were in G2-M versus 12% in the control cells (37) . Allicin was also found to arrest MCF-7 human breast cancer cells in G2-M (32) . Thus, it seems likely that the cellular and molecular effects of SAMC that we observed in the present study are relevant to the antitumor effects of garlic and other garlic derivatives. At the same time, our findings do not exclude the possibility that garlic and specific garlic derivatives exert antitumor effects through additional mechanisms, for example, effects on drug metabolizing enzymes (1 , 38) , alteration of protein phosphorylation status (39) , and so forth.
In contrast to the effects of SAMC, our studies with SAC indicated that this compound does not have any detectable effects on growth, cell cycle progression, or MT polymerization in intact SW480 or NIH3T3 cells, at concentrations of 150–300 µM. These negative findings are consistent with previous reports indicating that similar concentrations of SAC failed to show growth-inhibitory effects in human colon, lung, skin, and prostate cancer cell lines (25
, 40)
. Previous studies showed that SAC did have antiproliferative effects on neuroblastoma (41)
and melanoma (42)
cells, but in these studies the doses were extremely high, i.e., in the range of 1–10 mM. A pharmacokinetic study (43)
of SAC in rats, mice, and dogs indicated that after oral administration SAC was rapidly and efficiently absorbed in the gastrointestinal tract. In rats, the bioavailability was 98%. The maximum plasma concentration appeared within 0.5–1 h, and was 50, 112, and 227 µM after oral administration of 12.5, 25, or 50 mg/kg SAC, respectively. Unfortunately, no pharmacokinetic data are available for SAMC. It is, however, encouraging that the plasma levels of SAC in the above-cited study (43)
were in the range of the SAMC concentrations that we used in our cell culture studies, i.e., 150–300 µM. Two in vivo studies showed that oral administration of SAMC (200 mg/kg) decreased acetaminophen-induced liver injury in mice, thus indicating the bioavailability of orally administrated SAMC (44
, 45)
. It is curious that, although in our studies, SAC did not have any observable toxicity to intact cells, like SAMC, it did interfere with tubulin polymerization in in vitro turbidity assays (Fig. 3)
, presumably because it can also interact with sulfhydryl groups in purified tubulin. Presumably the lack of a cytotoxic effect of SAC in intact cells reflects the limited uptake and/or rapid metabolism of this compound by cells, but the precise mechanism remains to be determined.
Our studies of the signal transduction pathways involved in SAMC-induced apoptosis demonstrate that treatment of cells with SAMC leads to rapid, marked, and persistent activation of the JNK1 pathway (Fig. 4A)
. Presumably this contributes to the apoptotic effect of SAMC, because activation of JNK1 is known to play a role in the induction of apoptosis by other cytotoxic agents (12)
. Indeed, our studies with the JNK inhibitor SP, a dn-JNK1, and with JNK1-/- cells provide evidence that this pathway plays an important role in the early phase of apoptosis induced by SAMC, and that this effect is independent of the G2-M cell cycle arrest induced by this compound (Figs. 5
and 6
). A hypothetical scheme based on our results is shown in Fig. 9
. SAMC interacts directly with tubulin and induces MT depolymerization, shortly after it enters the cells, i.e., within 10 min. This leads, through unknown mechanisms, to rapid activation of the JNK1 pathway within 15 min; this, in turn, leads to activation of caspase-3, PARP cleavage, and other events that mediate an early phase of apoptosis, occurring during the first 24 h, which we designate "Phase 1." During this period, SAMC-induced MT depolymerization also interferes with assembly of the mitotic spindle, thus arresting cells in mitosis, within the first 12–24 h. Presumably, this arrest triggers spindle checkpoint and other responses, which contribute to a late phase of apoptosis, termed "Phase 2," which occurs at about 24–48 h after treatment with SAMC. We found that SAMC also causes activation of the ERK1/2 and p38 kinases (Fig. 7A)
, but the functional significance of these effects with respect to cytotoxicity are not known.
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. Although similar studies have not been previously reported for other MT-depolymerizing agents, it seems likely that this model may apply to diverse MIAs.
Our studies with JNK1-/- and JNK2-/- cells demonstrated that a lack of JNK1 activity inhibits SAMC-induced apoptosis, whereas a lack of JNK2 activity actually enhances SAMC-induced apoptosis (Fig. 6B)
, indicating different roles for JNK1 and JNK2 in these cellular events. Our findings are consistent with previous evidence that JNK1 is specifically involved in mediating the apoptotic response in a variety of human cancer cells treated with UV radiation or MIAs (12
, 48)
, whereas JNK2 plays a more specific role in cell survival (49)
. In a DMBA/TPA two-stage skin tumor model, JNK1-/- mice were more susceptible to TPA-induced skin tumor formation and growth than were WT mice (50)
, whereas skin tumorigenesis was suppressed in JNK2-/- mice (51)
. These findings suggest that JNK1 normally suppresses DMBA/TPA-induced skin tumor development, whereas JNK2 enhances it. A cDNA microarray analysis revealed different patterns of gene expression in JNK1-/-, JNK2-/-, and WT MEFs after TPA treatment (5)
. In JNK1-/- cells, TPA up-regulated 16 genes, including the genes encoding well-known antiapoptotic proteins, such as the A20 zinc finger protein, GST5-5, and the c-akt oncoprotein. In TPA-treated JNK2-/- cells, most of the genes that were highly expressed were related to tumor suppression and induction of cell differentiation, apoptosis, and cell growth arrest (5)
. These findings appear to explain the results obtained with respect to the mouse skin tumor model (50
, 51)
and are also consistent with our results.
It appears that, in NIH3T3 cells, one-third of the total cellular content of MAPK is associated with the MT cytoskeleton (52) . In addition, mixed linkage kinase 2 (MLK2), a MAPK kinase kinase, interacts with two KIF3 kinesin motor proteins and colocalizes with dually phosphorylated JNK1/2 in punctuate structures distributed along MTs (53) . These studies indicate that several signaling molecules localize on MTs, and suggest that MTs may act as a scaffold to bring together related signaling components. This association may explain the above-described correlations between activation of MAPK activities and disruptions in MT structure caused by various drugs. Studies are in progress to examine these effects with respect to the action of SAMC. Additional studies are also required to elucidate how the arrest of cells in mitosis induced by SAMC and other MIAs leads to what we have termed the Phase 2 apoptotic response. Inactivation of the antiapoptotic protein Bcl-2 by phosphorylation has been implicated in the apoptosis induced by paclitaxel (47) , but this mechanism is controversial (54) . Answers to the above questions will increase our understanding of the molecular mechanisms by which MIAs exert antitumor effects and will also help in the identification and/or synthesis of novel anticancer compounds.
In summary, the present studies provide the first evidence that the garlic derivative SAMC can inhibit the growth of cancer cells by directly binding to sulfhydryl residues in tubulin and thereby disrupt MT structures in the cytoplasm of interphase cells and the spindle apparatus of mitotic cells. We have also obtained evidence that the apoptosis induced by this compound is related, at least in part, to activation of the JNK1 pathway, but additional signaling mechanisms remain to be elucidated. Our findings may encourage the development of derivatives of SAMC that are more potent and suggest that SAMC or related compounds might provide a novel approach to cancer chemoprevention and/or cancer therapy.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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1 Supported by funding from the National Foundation for Cancer Research and the T. J. Martell Foundation (to I. B. W.), the NIH Grant GM62939 (to G. G. G.), and the NIH Grant CA89815 (to J. T. P.). 
2 To whom requests for reprints should be addressed, at the Herbert Irving Comprehensive Cancer Center, College of Physicians and Surgeons, Columbia University, Columbia-Presbyterian Medical Center, 701 West 168th Street, Suite 1509, New York, NY 10032. Phone: (212) 305-6921; Fax: (212) 305-6889; E-mail: ibw1{at}columbia.edu
3 The abbreviations used are: SAMC, S-allylmercaptocysteine; dn, dominant negative; EGFP, enhanced green fluorescent protein; ERK, extracellular signal-regulated kinase; JNK, c-Jun NH2-terminal kinase; MAPK, mitogen-activated protein kinase; MEF, mouse embryonic fibroblast; MT, microtubule; MAP, MT-associated protein; MIA, MT-interfering agent; SAC, S-allylcysteine; ß-ME or BME, ß-mercaptoethanol; TPA, 12-O-tetradecanoylphorbol-13-acetate; WT, wild type; SB, SB203580; SP, SP600125; PD, PD98059; VAD, z-VAD-fmk; PI, propidium iodide; CMV, cytomegalovirus; TBS, Tris-buffered saline; DMBA, 7,12-dimethylbenz(a)anthracene; 7-AAD, 7-amino-actinomycin; AFC, 7-amino-4-trifluoromethylcoumarin; AMC, 7-amino-4-methylcoumarin; G-PEM, buffer composed of GTP, PIPES, EGTA and MgCI2.
4 D. Xiao and I. B. Weinstein, unpublished data.
Received 2/13/03. Revised 7/16/03. Accepted 7/22/03.
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