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CHEMOTHERAPY, ANTIBIOTICS, AND GENE THERAPY
Departments of Biopharmaceutics (N.K., H.K., T.K., M.O.) and Clinical Pharmaceutics (T.M.), Graduate School of Pharmaceutical Sciences, and Department of Cell Pathology (T.N., M.T.), Graduate School of Medical Sciences, Kumamoto University, Kumamoto, Japan; and Pharmaceutical Research Center, Nipro Corporation, Shiga, Japan (N.K., K.N., K.U., J.-i.Y., N.Y., H.Y., T.K., M.S.)
Received for publication
September 26, 2007
Accepted
January 23, 2008.
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). Therefore, induction of apoptosis in neoplastic cells is a very effective therapy for tumor eradication (Meng et al., 2006). However, this type of chemotherapy often has negative side effects, such as transient cell cycle arrest, senescence, and autophagy. Drug delivery systems that facilitate selective apoptosis of neoplastic cells have been suggested as a way of overcoming this problem (Kaufmann and Gores, 2000; Kondo et al., 2005).
NO is a unique diffusible molecular messenger that occupies a central role in mammalian pathophysiology (Brune et al., 1998). Its multiple actions include vascular smooth muscle relaxation (Moncada et al., 1986; Ignarro, 1989), inhibition of platelet aggregation (Azuma et al., 1986), effects on neurotransmission (Garthwaite, 1991), and regulation of immune function (Marletta et al., 1988). Alternatively, under some circumstances, NO is cytotoxic (Laval and Wink, 1994). NO causes cellular iron losses and inhibits DNA synthesis, mitochondrial respiration, and aconitase activity in L10 hepatoma cells (Hibbs et al., 1988). In addition, NO reacts with superoxide anion (which is produced by activated macrophages and other cells), to form peroxynitrite. This by-product of NO is a potent chemical oxidant, which alters protein function and damages DNA (Beckman and Crow, 1993). These effects are part of the nonspecific host defense, which facilitates killing of tumor cells and intracellular pathogens. In addition, the cytotoxicity arising from long-lasting NO generation has been attributed to induction of apoptosis (Brune et al., 1998).
In recent studies, the cytotoxic activity of NO has been studied to assess its therapeutic potential in cancer treatment. NO-donating nonsteroidal anti-inflammatory drugs (NSAIDs), especially NO-aspirin (NO-ASA), have been investigated as promising chemopreventive agents (Williams et al., 2001; Kashfi et al., 2002; Fabbri et al., 2005). NO-ASA consists of traditional ASA to which an NO-releasing moiety is bound via a spacer. This agent induces oxidative stress by increasing intracellular peroxide and 
, thereby inducing apoptosis via activation of the intrinsic apoptosis pathway (Gao et al., 2005). JS-K is a prodrug designed to release NO after reacting with glutathione transferase, which induces double-stranded DNA breaks, activates DNA damage response pathways, and induces apoptosis in human multiple myeloma cells both in vitro and in vivo (Kiziltepe et al., 2007).
Human serum albumin (HSA) is an abundant circulating protein, and the nitrosated form serves as a reservoir of NO (Stamler et al., 1992). Therefore, NO-HSA is an NO donor that is currently being investigated for its potential therapeutic applications. For example, administration of NO-HSA to animals with ischemia-reperfusion injury minimizes the tissue damage that occurs after reperfusion (Semsroth et al., 2005). In a balloon-injured rabbit femoral artery model, locally delivered NO-HSA preferentially binds to sites of vessel injury and inhibits both platelet accumulation and the subsequent development of neointimal hyperplasia (Marks et al., 1995). NO-HSA also shows potent antibacterial activity and inhibits the proliferation of cultured human vascular smooth muscle cells (Ishima et al., 2007). However, there are no reports describing the effects of NO-HSA on cancer.
Accordingly, the present study evaluated the antitumor activity of NO bound to HSA (NO-HSA) via an S-nitrosothiol linkage using iminothiolane as a spacer. The molecular events related to induction of apoptosis by NO-HSA were studied in vitro, and the antitumor activity of NO-HSA was studied in vivo using a murine model of C26 colon carcinoma.
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Expression and Purification of Recombinant HSA. rHSA was produced using a yeast expression system as described previously (Matsushita et al., 2004). In brief, for constructing the HSA expression vector pPIC9-HSA, native HSA coding region was incorporated into the methanol-inducible pPIC9 vector (Invitrogen, Carlsbad, CA). The resulting vector was introduced into the yeast species Pichia pastoris (strain GS115) to express rHSA. Secreted rHSA was isolated from the growth medium by a combination of precipitation with 60% (w/v) (NH4)2SO4 and purification on a Blue Sepharose CL-6B column (GE Healthcare, Little Chalfont, Buckinghamshire, UK) followed by Phenyl HP column (GE Healthcare). Isolated protein was defatted by using the charcoal procedure described by Chen (1967), deionized, freeze-dried, and then stored at –20°C until use. The resulting rHSA (treated with dithiothreitol) exhibited a single band on SDS-PAGE. Density analysis of protein bands stained with Coomassie Brilliant Blue showed that its purity was more than 97%.
Synthesis of NO-HSA. Terminal sulfhydryl groups were added to the HSA molecule by incubating 0.15 mM rHSA with 3 mM Traut's reagent in 100 mM potassium phosphate buffer containing 0.5 mM DTPA, pH 7.8, for 1 h at room temperature. The resultant modified rHSA then was S-nitrosylated by 3-h incubation with 15 mM isopentyl nitrite at room temperature (Fig. 1). The resulting NO-HSA was concentrated, exchanged with saline using a PelliconXL filtration device (Millipore Corporation, Billerica, MA), and the final concentration adjusted to 2 mM NO-HSA. The sample was stored at –80°C until use.
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Cellular Experiments with C26 Cells. C26 cells, which were donated by the Institute of Development, Aging and Cancer, at Tohoku University (Sendai, Miyagi, Japan), were cultured at 37°C in RPMI 1640 medium containing 10% fetal calf serum, 100 U/ml penicillin, and 10 µg/ml streptomycin (medium A). Trypsin (0.25%) EDTA solution was used to detach the cells from the culture flask for plating and passing the cells. All cell culture experiments were performed at 37°C in a humidified atmosphere of 5% CO2 in air.
For detection of reactive oxygen species (ROS), C26 cells (1.0 x 104 cells/well) were cultured in 96-well plates in medium A for 12 h, they were washed twice with PBS, and then they were incubated for an additional 30 min in HBSS containing 5 µM 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H2DCFDA) (Invitrogen, Carlsbad, CA). After washing twice with HBSS, the cells were cultured in HBSS containing 5% fetal calf serum for 15 min followed by the addition of PBS, 50 µM HSA, or 50 µM NO-HSA. After incubation, fluorescence was measured using a plate reader (excitation wavelength, 485 nm; emission wavelength, 535 nm). The change in fluorescence was calculated by subtracting the fluorescence at 0 h from the fluorescence measured at the indicated times. The fluorescence intensities of cells incubated with PBS, 50 µM HSA, and 50 µM NO-HSA at 0 h were 201.3, 166.1, and 181.3, respectively.
Changes in the mitochondrial membrane potential of C26 cells were monitored using flow cytometric analysis with rhodamine 123 staining. C26 cells (1.0 x 106 cells/well) were cultured in six-well plates for 12 h, washed twice with PBS, and incubated with PBS and either 100 µM HSA or various concentrations of NO-HSA in medium A for 24 h. The cells were then trypsinized, washed twice with PBS, and incubated for 15 min with 5 µg/ml rhodamine 123. The mean fluorescence intensity of rhodamine 123 in the cells was measured using a flow cytometer (FACSCalibur; BD Biosciences, Franklin Lakes, NJ).
For the determination of caspase-3 activity, cells were cultured to confluence in six-well plates, washed twice with PBS, and incubated with medium A containing 100 µM HSA or various concentrations of NO-HSA. Cells were incubated for 24 h, trypsinized, and washed with 0.2 ml of ice-cold PBS. The cell pellet was resuspended in 15 µl of cell lysis buffer, it was lysed by freeze-thawing, and then it was incubated on ice for 15 min. The cell lysates were centrifuged at 15,000 rpm for 20 min at 4°C, and the supernatant fraction was collected (cell extract). The caspase-3 activity in the cell extract was assessed using the colorimetric CaspASE Assay System (Promega, Madison, WI), according to the manufacturer's instructions.
For the detection of DNA degradation (DNA ladder), C26 cells (1.0 x 106 cell/well) were cultured in six-well plates. Cells were cultured for 12 h, they were washed twice with PBS, and then they were incubated with PBS and either 100 µM HSA or various concentrations of NO-HSA for 24 h. The cells then were trypsinized, collected, and centrifuged at 4000 rpm for 10 min. After removing the supernatant, the cell pellet was resuspended in 0.2 ml of PBS, and then it was centrifuged at 4000 rpm for 10 min. The supernatant was again removed, and the remaining pellet was incubated in 20 µl of 10 mM Tris-HCl buffer, pH 7.8, containing 2 mM EDTA and 0.5% SDS (cell lysis buffer) for 10 min at 4°C, followed by centrifugation at 15,000 rpm for 5 min. The resulting supernatant (cell extract) was collected and incubated with 1 µl of RNase A (10 µg/ml) for 30 min at 50°C. One microliter of proteinase K (10 µg/ml) was added to the cell extract, followed by a 1-h incubation at 50°C. The resulting DNA extract was electrophoresed in a 2.0% agarose gel, followed by staining of the gel with ethidium bromide and visualization of the DNA bands using ultraviolet illumination.
The cell viability assay was performed using WST-8, which is based on the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium assay. C26 cells were plated in 96-well plates at 1.0 x 104 cells/well, and they were cultured for 32 h in medium A. Then, the cells were washed twice with PBS and incubated in a total volume of 0.2 ml of medium A containing various concentrations of HSA or NO-HSA. After incubating the cultures for various lengths of time, 5 µl of WST-8 solution was added to each well, and the cells were incubated for an additional 2 h at 37°C. The number of surviving cells was determined by measuring the absorbance at 450 nm. Cell viability was calculated as the percentage of the control value (without HSA or NO-HSA) (Ishiyama et al., 1996).
Animal Experiments. Five-week-old male BALB/c AnNCrlCrlj mice (17–20 g) were purchased from Charles River Italica (Calco, Italy). The mice were housed in a 12-h light/dark cycle in a humidity-controlled room. Mice were acclimated for at least 5 days before being used in experiments.
For tumor induction, mice were inoculated with C26 cells (1.0 x 106 cells/mouse) by a subcutaneous injection into the dorsal skin. Three days after inoculation, C26 carcinoma-bearing mice were randomly divided into three groups: control, HSA, and NO-HSA. The mice received a daily i.v. injection of saline, HSA (10 µmol/kg), or NO-HSA (10 µmol/kg) for 10 days from day 3 to day 12 after inoculation. Tumor volume was calculated using the formula 0.4 (a x b2), where a is the largest diameter and b is the smallest diameter of the tumor (Shimizu et al., 2005), and volume was monitored from day 7 to day 17 after inoculation. Variance in each group was evaluated using the Bartlett test, and differences in mean tumor volume were evaluated using the Tukey-Kramer test.
Some animals in each of the three treatment groups were used for immunohistochemical analysis and serum biochemistry. When the mice received five times per day from day 3 to day 7 after inoculation (the tumors in each group reached approximately 5 mm in diameter), blood samples were collected from the abdominal vena cava under diethyl ether anesthesia approximately 2 h after the daily injection, and then the mice were sacrificed.
Tumors were dissected, they were fixed immediately with 2% periodate/lysine/paraform-aldehyde fixative at 4°C for 5 h, and then they were washed with a graded series of sucrose solutions in PBS (10, 15, and 20%). After immersion in 20% sucrose in PBS to inhibit ice crystal formation, the tissues were embedded in O.C.T. compound (Sakura Fine Technochemical, Tokyo, Japan), they were frozen in liquid nitrogen, and then they were stored at –80°C. Five-micrometer tumor sections were prepared using a cryostatic microtome (HM500M; Microm, Walldorf, Germany), and they were mounted on poly-L-lysine-coated slides. The slides were stained using the TUNEL method and an in situ apoptosis detection kit (Takara-Bio Co. Ltd., Shiga, Japan). The slides were washed three times with 0.01 M phosphate buffer, pH 7.4, containing 0.9% NaCl, followed by application of methanol containing 0.3% H2O2 to inactivate endogenous peroxidase and incubation at room temperature for 30 min. The slides were washed 3 times with 0.01 M phosphate-buffered saline, and then they were incubated in 100 ml of permeabilization buffer on ice for 5 min. The slides were washed three times with 0.01 M phosphate-buffered saline, and then they were incubated with 50 ml of freshly prepared terminal deoxynucleotidyl transferase reaction mixture (5 ml of terminal deoxynucleotidyl transferase enzyme + 45 ml of Labeling Safe buffer) at 37°C for 60 min. After the slides were washed three times with 0.01 M PBS, they were incubated in 70 ml of anti-fluorescein isothiocyanate-horseradish peroxidase conjugate antibody at 37°C for 30 min. After the slides were washed three times with 0.01 M PBS, they were incubated in 3,3-diaminobenzidine-H2O2 reaction buffer at room temperature for 10 min. After the slides were washed three times with distilled water, they were stained with 3% methyl green for 10 min, dehydrated, penetrated, and sealed (Gavrieli et al., 1992). Each slide then was visualized under a light microscope (Olympus, Tokyo, Japan), at a magnification of 400x.
Serum was separated by centrifugation. Routine clinical laboratory techniques were used to determine the concentrations of total protein, serum creatinine (Cr), blood urea nitrogen (BUN), alanine aminotransferase (ALT), aspartate aminotransferase (AST), and alkaline phosphatase (ALP) in serum. Variance in each group was evaluated using the Bartlett test, and differences were evaluated using the Tukey-Kramer test.
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, and peroxynitrite, causes apoptosis. Moreover, production of ROS also plays a major role in NO-ASA-induced apoptosis. To examine whether NO-HSA promoted ROS production in C26 cells, a fluorescent probe (CM-H2DCFDA), which undergoes conversion to 2',7'-dichlorofluorescein in the presence of intracellular ROS, was used. Addition of NO-HSA to C26 cells increased the amount of ROS compared with treatment with vehicle or HSA (Fig. 2). In addition, the ROS in the C26 cell culture medium increased with time after addition of NO-HSA. This result suggests that NO-HSA promotes a signal cascade leading to apoptosis by increasing intracellular production of ROS.
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Caspase-3 is a cell death protease that is involved in the downstream execution phase of apoptosis. During this phase of apoptosis, cells undergo morphological changes, such as DNA fragmentation, chromatin condensation, and formation of apoptotic bodies. Compared with the effect of vehicle, cells treated with 25 or 50 µM NO-HSA had slightly increased caspase-3 activity, and cells treated with 100 µM NO-HSA showed a larger increase in caspase-3 activity (Fig. 3B). Cells treated with HSA had the same level of caspase-3 activity as cells treated with PBS.
To further confirm that NO-HSA induced apoptosis in C26 cells, DNA fragmentation, which is a typical morphological change in the execution phase of apoptosis, was examined. DNA fragmentation was observed in C26 cells treated with 100 µM NO-HSA, but not in C26 cells treated with 25 or 50 µM NO-HSA (Fig. 3C). Neither vehicle nor 100 µM HSA induced DNA fragmentation in C26 cells (Fig. 3C). These findings suggest that NO-HSA induces apoptosis by increasing ROS production, activating caspase-3, and hyperpolarizing the mitochondrial membrane potential.
To determine the effect of NO-HSA on cell growth, the viability of C26 cells was examined after treatment with HSA or various concentrations of NO-HSA. NO-HSA inhibited growth of C26 cells in a concentration-dependent manner; cell growth was suppressed by 71, 80, and 85% after a 48-h incubation with 25, 50, and 100 µM NO-HSA, respectively (Fig. 4A). The viability of C26 cells incubated with 50 µM NO-HSA significantly decreased with increasing incubation times (Fig. 4B). NO-HSA inhibited growth to a greater extent than did HSA, which had only a weak inhibitory effect. These results suggest that NO-HSA inhibits cell growth of C26 cells by inducing apoptosis.
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NO-HSA Exerts Antitumor Effect via the Apoptotic Pathway in Vivo. To investigate the antitumor effect of NO-HSA in vivo, C26 tumor-bearing mice received i.v. injections with saline, HSA, or NO-HSA. Mean tumor area increased with time in the saline-treated group. In the HSA-treated group, tumor growth was suppressed, compared with that in the control group, but the difference was not statistically significant. In contrast, tumor growth was significantly inhibited by administration of NO-HSA (Fig. 5).
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To clarify whether the suppression of tumor growth by NO-HSA is mediated via apoptosis, tumor tissues from C26 tumor-bearing mice receiving injections with NO-HSA were examined using immunohistochemistry. In NO-HSA-treated mice, there were more TUNEL-positive cells than in the saline- and HSA-treated animals. In addition, the tumor tissue architecture was less defined in animals treated with NO-HSA than in the other groups, suggesting that NO-HSA induced apoptosis in C26 tumor cells and thus exerted an antitumor effect in vivo (Fig. 6).
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). However, the in vivo half-life of NO (
0.1 s) is often too short to capitalize on its potential biological actions. The half-life of NO can be prolonged by adding S-nitrosothiol moieties with cysteine residues of proteins. For example, nitrosated HSA seems to act as a reservoir of NO in vivo (Stamler et al., 1992). Simon et al. (1996) incubated bovine serum albumin (BSA) with 200-fold excess concentration of NaNO2 under acidic condition to synthesize polynitrosylated BSA that is highly modified at the thiol group of cysteine, hydroxyl group of tyrosine and amines (38 mol NO/mol BSA). The polynitrosylated BSA has been shown to exhibit anti-platelet activity. However, polynitrosylated S-NO-BSA, an NO-BSA conjugate prepared with the same method except that the BSA has been reduced with dithiothreitol and it contains 19 mol of "S-NO" per mol of BSA, was a significantly more potent platelet inhibitor than polynitrosylated BSA described above. These findings show that nitrosylated BSA behaves as an NO donor; in particular, the poly(S-nitroso) derivative could be by far the most potent compound. One molecule of HSA contains 35 cysteine residues, 34 of which form 17 nonreactive disulfide bonds, and one of which (Cys-34) forms a reactive free thiol (Peters, 1985). Thus, the number of NO molecules that can be bound to HSA is limited because only one free cysteine per HSA molecule is available for conjugation. Ewing et al. increased the number of free sulfhydryl groups on BSA by reduction with dithiothreitol and thiolation with N-acetylhomocysteine, thereby preparing polynitrosated BSA (12–15 mol NO/mol BSA) (Ewing et al., 1997). Marks et al. (1995) produced polynitrosated BSA (5.9 mol NO/mol BSA) by adding free sulfhydryl groups to the molecule and by treating the BSA with N-acetylhomocysteine thiolactone. However, the polynitrosated BSAs prepared in these studies formed aggregates as a result of intermolecular disulfide formation. Aggregate formation results in molecular heterogeneity, which limits the therapeutic application of S-nitroso residues. In the present study, iminothiolane, which reacts with primary amines to introduce sulfhydryl groups while maintaining charge properties similar to the original amino groups, was selected as the thiolation reagent. Iminothiolane was used to produce polynitrosated HSA (NO-HSA) (6.6 mol NO/mol HSA), which did not form aggregates after nonreducing SDS-PAGE or native-PAGE (data not shown). Moreover, the far-UV CD spectra of NO-HSA were nearly identical to those of HSA (data not shown). Therefore, NO-HSA is expected to be clinically applicable as a biocompatible pharmacological agent, although further study is required to clarify other potential issues, including the antigenicity of this protein.
NO-NSAIDs have been extensively investigated as therapeutic agents for cancer due to their ability to release NO, thereby promoting apoptosis. NO-NSAIDs are categorized as organic nitrate esters, which are readily reduced to organic nitrite esters by cytosolic enzymes. Subsequently, glutathione reacts with organic nitrite esters to form GSNO, indicating that NO-NSAIDs release NO via S-nitrosothiol (Wong and Fukuto, 1999). Alternatively, the transfer of NO from NO-HSA to the cytosol could be inferred from a study by Ramachandran et al. (2001). They reported that NO is released from extracellular S-nitrosothiols by a cell surface enzyme (protein disulfide isomerase) and that it accumulates in the cell membrane where it reacts with O2 to produce N2O3, which is then available for nitrosation reactions with intracellular thiols at the membrane-cytosol interface (Ramachardran et al., 2001). Therefore, it is possible that NO-HSA also releases NO by the intracellular formation of S-nitrosothiol, suggesting that the species of NO released within the cell by S-nitrosothiols, as well as the reactive substances (such as ROS) derived from the released NO, would not differ significantly between NO-NSAIDs and NO-HSA. In support of this hypothesis, NO-HSA caused depolarization of the mitochondrial membrane potential, activation of caspase-3 and DNA fragmentation in the present study, consistent with the effects of NO-NSAIDs. Additional studies are needed to determine the details of the molecular events and the systematic pathways affected by NO-HSA, but the mechanism of action should be similar to that of NO-NSAIDs. In a recent study, Gao et al. (2005) elucidated the detailed mechanism of apoptosis induced by NO-ASA. Intracellular accumulation of ROS is a key proximal event in NO-ASA-induced apoptosis, and it correlates with the effect on tumor cell growth (Gao et al., 2005). In the present study, NO-HSA induced accumulation of ROS in tumor cells, suggesting that increased ROS production may be an important proximal event leading to induction of apoptosis.
The results of the in vivo study showed that NO-HSA significantly suppressed tumor growth by inducing apoptosis, without adverse changes in serum biochemical parameters in treated mice. In a recent study, Trachootham et al. (2006), using immortalized cell lines and their oncogenic progeny transfected with H-RasV12, demonstrated that cancer cells typically produce more ROS than normal cells. Moreover, the pro-oxidant status of cancer cells increases their susceptibility to treatment with agents that cause oxidative stress, as demonstrated in a study using β-phenylethyl isothiocyanate (Trachootham et al., 2006). In addition, Feng et al. (2007) reported that cyaniding-3-rutinoside selectively induces accumulation of peroxides in HL-60 human leukemic cells, but not in normal peripheral blood mononuclear cells (Feng et al., 2007). Schumacker (2006) has proposed that ROS toxicity induced by certain chemotherapeutic agents may be an effective means of selectively eradicating malignant cells. In the present study, we presumed that although NO reacts with superoxide anion to form peroxynitrite (a potent oxidant and nitrating agent), these highly reactive oxidant species are probably produced at higher levels in C26 cells compared with normal cells. Therefore, NO-HSA may show selective cytotoxicity for tumor cells and not affect normal cells. These findings strongly suggest that NO-HSA is a promising therapeutic anticancer agent, given the unusual redox conditions typical of malignant cells.
Antiapoptotic effects of NO have been observed in a variety of cells, including T cells, hepatocytes, endothelial cells, neurons, ovarian follicle cells, eosinophils, thymocytes, and embryonic kidney cells (Liu and Stamler, 1999). In a recent study using U937 human promonocytic cells, NO-R410C (a genetic variant of HSA) had antiapoptotic activity (Ishima et al., 2007). Whether NO ultimately inhibits or promotes apoptosis probably depends on the cell, the signal, the source, the molecule, the amount, and the presence or absence of coreactants. For example, the amount of NO bound to the carrier molecule seems to account for the discrepant results between previous investigations and the present study. The NO content of NO-R410C (in the previous study) and NO-HSA (in this study) were 1.3 and 6.6 mol NO/mol HSA, respectively, and the S-nitroso moiety concentrations in vitro were 26 to 130 and 165 to 660 µM, respectively. Mohr et al. (1997) reported that 10 to 100 µM GSNO inhibits the activation of caspase-3 induced by actinomycin D in U937 cells. In contrast, apoptosis (characterized by DNA fragmentation and morphological changes) was observed in U937 cells treated with GSNO at concentrations in excess of 250 µM (Messmer et al., 1996). Based on the results of the present study and those of previously published investigations, the critical NO threshold concentration between promotion and inhibition of apoptosis seems to be 100 to 200 µM.
Matsumura et al. (1987) examined the accumulation of differently sized proteins within tumor tissues of tumor-bearing mice. Macromolecules containing HSA tended to accumulate in tumor tissues, apparently due to hypervascularization and enhanced vascular permeability (even to macromolecules) of the tumors, with little export of macromolecules from the tumor tissue via blood or lymphatic vessels (Matsumura et al., 1987). Therefore, NO-HSA may be a useful agent for targeting chemotherapeutics to tumor tissue. However, the short half-life of NO has been one of the greatest obstacles to therapeutic application of NO donors. Consequently, the pharmacokinetic properties of NO-HSA in mice were measured to determine the biological fate of NO. The apparent half-life of S-nitroso moieties in NO-HSA was estimated to be 18.9 min (data not shown), which is similar to that of NO-R410C, but much longer than that of the low-molecular-weight NO donor GSNO (4.2 min) (Ishima et al., 2007). In the present study, the difference in NO half-life between NO-HSA and GSNO seemed to be due to reduced renal excretion of HSA compared with glutathione due to its larger molecular size, suggesting that HSA may be a useful NO carrier in vivo.
In summary, NO-HSA was synthesized by inducing S-nitrosothiol linkages using iminothiolane as a spacer. NO-HSA generated ROS in C26 cells, and it induced intrinsic apoptotic events, such as depolarization of mitochondrial membrane potentials, activation of caspase-3, and induction of DNA fragmentation. Moreover, NO-HSA inhibited proliferation of tumor cells in vitro in a concentration-dependent manner. In the in vivo experiments, NO-HSA also strongly inhibited tumor growth by inducing apoptosis, with no side effects. The results of the present study suggest that NO-HSA has promise as a new generation anticancer agent with few side effects.
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Footnotes |
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ABBREVIATIONS: NSAID, nonsteroidal anti-inflammatory drug; ASA, aspirin; HSA, human serum albumin; DTPA, diethylenetriaminepentaacetic acid; HBSS, Hanks' balanced salt solution; rHSA, recombinant human serum albumin; PAGE, polyacrylamide gel electrophoresis; C26, murine colon 26 carcinoma; ROS, reactive oxygen species; PBS, phosphate-buffered saline; CM-H2DCFDA, 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate, acetyl ester; TUNEL, terminal deoxynucleotidyl transferase dUTP nick-end labeling; Cr, serum creatinine; BUN, blood urea nitrogen; ALT, alanine aminotransferase; AST, aspartate aminotransferase; ALP, alkaline phosphatase; BSA, bovine serum albumin; GSNO, S-nitrosoglutathione; R410C, genetic variant of human serum albumin mutated at position 410.
Address correspondence to: Dr. Masaki Otagiri, Department of Biopharmaceutics, Graduate school of Pharmaceutical Sciences, Kumamoto University, 5-1 Oe-honmachi, Kumamoto 862-0973, Japan. E-mail: otagirim{at}gpo.kumamoto-u.ac.jp
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