Chemico-Biological Interactions 221 (2014) 24–34

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Spiruchostatin A and B, novel histone deacetylase inhibitors, induce apoptosis through reactive oxygen species-mitochondria pathway in human lymphoma U937 cells Mati Ur Rehman a, Paras Jawaid b, Yoko Yoshihisa a, Peng Li b, Qing Li Zhao b, Koichi Narita c, Tadashi Katoh c, Takashi Kondo b,⇑, Tadamichi Shimizu a a

Department of Dermatology, Graduate School of Medicine and Pharmaceutical Science, University of Toyama, 2630 Sugitani, Toyama 930-0194, Japan Department of Radiological Sciences, Graduate School of Medicine and Pharmaceutical Science, University of Toyama, 2630 Sugitani, Toyama 930-0194, Japan c Laboratory of Synthetic and Medicinal Chemistry, Faculty of Chemical Pharmaceutical Science, Tohoku Pharmaceutical University, Aoba-ku, Sendai 981-8558, Japan b

a r t i c l e

i n f o

Article history: Received 14 May 2014 Received in revised form 30 June 2014 Accepted 11 July 2014 Available online 29 July 2014 Keywords: Spiruchostatin A Spiruchostatin B HDAC inhibitors Apoptosis ROS

a b s t r a c t Spiruchostatin A (SP-A) and spiruchostatin B (SP-B) are the potent histone deacetylase inhibitors (HDACi), that has the potential for chemotherapy of leukemia but the exact mechanism of these compounds remains unclear. In the present study, the role of reactive oxygen species (ROS) production and the mechanism involved in the apoptosis was investigated in human lymphoma U937 cell. When the U937 cells were treated with SP-A and SP-B for 24 h at different concentrations, evidence of apoptotic features, including increase in DNA fragmentation and changes in nuclear morphology, were obtained. SP-B showed maximum potency to induce apoptosis, while SP-A was less potent. Apoptosis was also determined by increase in the fraction of sub-G1 cells and Annexin V-FITC staining cells. SP-A and SP-B induced apoptosis was accompanied by significant increase in the formation of intracellular reactive oxygen species (ROS). Pre-treatment with N-acetyl-L-cysteine (NAC), significantly inhibited the SP-A and SPB mediated apoptosis, suggesting a vital role of ROS involved in the lethality of both agents. Moreover, SPA and SP-B treatment resulted in the loss of mitochondrial membrane potential (MMP), and Fas, caspase8 and caspase-3 activation. In addition Bid activation and the release of cytochrome-c to the cytosol was also observed. In this study, we suggest that a marked induction of intracellular ROS mediated mitochondrial pathway and the Fas plays a role in the SP-A and SP-B induced apoptosis. Taken together, our data provides further insights of the mechanism of action of SP-A and SP-B and their potential application as novel chemotherapeutic agents. Ó 2014 Elsevier Ireland Ltd. All rights reserved.

1. Introduction Acetylation and deacetylation of histones play a crucial role to control transcriptional activity of different genes through affecting the interaction between DNA and histones [1,2]. Histones acetylation and deacetylation is regulated by a balance between two Abbreviations: SP-A, Spiruchostatin A; SP-B, Spiruchostatin B; ROS, reactive oxygen species; HDAC, histone deacetylase; MMP, mitochondrial membrane potential; NAC, N-acetyl-L-cysteine; FITC, fluorescein isothiocyanate; HATs, histone acetyltransferases; HDACi, histone deacetylase inhibitors; CTCL, cutaneous T-cell lymphoma; SAHA, suberoylanilide hydroxamic acid; SCLC, small cell lung cancer; DCFH-DA, dichlorofluorescein diacetate; DMSO, dimethyl sulfoxide; DTT, dithiothreitol; HE, hydroethidine; pNA, chromophore p-nitroanilide; TCA, trichloroacetic acid; TMRM, tetramethylrhodamine methyl ester. ⇑ Corresponding author. Tel.: +81 76 434 7267; fax: +81 76 434 5190. E-mail address: [email protected] (T. Kondo). http://dx.doi.org/10.1016/j.cbi.2014.07.004 0009-2797/Ó 2014 Elsevier Ireland Ltd. All rights reserved.

specific enzymes, histone acetyltransferases (HATs) and histone deacetylase (HDAC) [3,4]. Aberrant regulation of HATs and HDAC activity has been reported to play an important role in uncontrolled cancer growth [5,6]. Increased HDAC activity was reported in several human cancer cell lines and therefore HDAC inhibition is considered as the novel strategy in the cancer treatment [7,8]. Recently, histone deacetylase inhibitors (HDACi) have gained much attention and emerged as promising anti-cancer agents [9], because of their diverse mode of action which include disruption of co-repressor complexes, ROS induction, death receptors up-regulation, generation of lipid second messengers (e.g. ceramide), interference with chaperone protein function [10,11]. Until now, several HDACi have been identified and on the basis of their structural features they were divided into different classes [12]. FK228 (romidepsin) and suberoylanilide hydroxamic acid (SAHA) has

M.U. Rehman et al. / Chemico-Biological Interactions 221 (2014) 24–34

already been approved by US Food and Drug administration (FDA), for the treatment of cutaneous T cell lymphoma (CTCL) [13], and others such as valproic acid and MS-275 are still in clinical trials [13,14]. In spite of the fact that HDAC inhibitors are promising anti-cancer agents, the molecular mechanism involved in the cell death after treatment with HDAC inhibitors remain unclear, but the induction of oxidative stress seems to be common theme involved in the cell death caused by HDAC inhibitors [15]. In solid tumor and leukemia cells, HDAC inhibitors have been reported to induce apoptosis through ROS generation [16]. Recently, Spiruchostatin A and B, the structurally similar 15-membered bicyclic depsipeptides, were successfully isolated from a culture broth of Pseudomonas sp. [17]. Both agents were found to possess potent HDAC inhibitory activity and have the potential for chemotherapy of leukemia [18,19]. Previously, the HDAC inhibitory activity of these agents were tested against the panel of 39 cancer cell line, in which SP-B was found to be the most potent inhibitor of HDAC1 and it showed the most potent inhibitory activity as compared to other HDACi including FK228 and SP-A [19]. Like other HDACi, SP-A and SP-B also characterized by the biological effects which include growth inhibition, cell cycle arrest and induction of p21 expression [3,20]. The purpose of this study is to determine the possible apoptosis inducing activity of SP-A and SP-B, and the detail molecular mechanism involved. To date, no report has regarded the effects of these compounds on ROS production and apoptosis in human lymphoma U937 cells. Therefore, this study was undertaken to further clarify the molecular mechanism involved in the apoptosis induced by SPA and SP-B in human lymphoma U937 cells.

2. Materials and methods

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2.3. Cell cycle analysis For flow cytometry, cells were fixed with 70% ice cold ethanol at least for 2 h or stored overnight at 20 °C, and subsequently treated with 0.25 mg/ml RNase A (Nacalai Tesque, Kyoto, Japan) and 50 lg/ml PI to obtain the distribution of PI-based cell-cycle phases. The samples were finally run on an Epics XL flow cytometer (Beckman Coulter, Fullerton, CA) [21]. 2.4. DNA fragmentation assay For the detection of apoptosis the percentage of DNA fragmentation was assessed up till 24 h post treatment using the method of Sellins and Cohen [22], with minor modifications. Briefly, approximately 3  106 cells were lysed using 200 ll of lysis buffer (10 mM Tris, 1 mM EDTA and 0.2% Triton X-100, pH 7.5) and centrifuged at 13,000g for 10 min. Subsequently, each DNA sample in the supernatant and the resulting pellet was precipitated in the 25% trichloroacetic acid (TCA) at 4 °C overnight and quantified using a diphenylamine reagent after hydrolysis in 5% TCA at 90 °C for 20 min. The percentage of fragmented DNA in each sample was calculated as the amount of DNA in the supernatant divided by total DNA for that sample (supernatant plus pellet). 2.5. Morphological detection of apoptosis The morphological changes in the cells were examined by Giemsa staining. To identify the apoptotic cells after treatment with SP-A and SP-B, cells were harvested after 24 h of incubation at 37 °C, were washed with PBS and collected by centrifugation. Then the cells were fixed with methanol and acetic acid (3:1) and spread on the glass slides. After drying, staining was performed with 5% Giemsa solution (pH 6.8) for 5 min [23].

2.1. Chemicals

2.6. Detection of apoptosis using Annexin V-FITC/PI staining

Spiruchostatin A and B were kindly provided by Prof. Tadashi Katoh (Faculty of Pharmaceutical Sciences, Tohoku Pharmaceutical University, Sendai, Japan). Both agents were dissolved in dimethyl sulfoxide (DMSO) and stored at 20 °C. The agents were diluted with culture media to prepare serial concentrations just before the use. The Annexin V-FITC kit was from Immunotech (Marseille, France). RNase A was purchased from Nacalai Tesque (Kyoto, Japan). DCFH-DA and HE was from molecular probes (Eugene). The FLICE/caspase-8 colorimetric protease assay kit was obtained from MBL (Nagoya, Japan). Antibodies against caspase-3, Bid, Bcl2, Ac-H3, Ac-H4 and secondary horseradish peroxide (HRP)-conjugated anti-rabbit or anti-mouse IgG were obtained from Cell Signaling technology (Danvers M.A.). Anti-b-actin was from Sigma Aldrich (Saint Louis) and anti-cytochrome-c pAb, anti Bax and Fas were purchased from Santa Cruz Biotechnology Inc (CA). Anti caspase-8 was from R&D systems (Minneapolis, USA). All other reagents were of analytical grade.

To determine early apoptosis and secondary necrosis, phosphatidylserine (PS) externalization of apoptosis was determined by analysis of propidium iodide (PI) and fluorescein isothiocyanate (FITC)-labeled Annexin V (Immunotech, Marseille, France) using Flow cytometry (Epics XL, Beckman-Coulter, Miami, FL) [24] according to the instructions of the manufacturer. Briefly, cells were treated with SP-A and SP-B at different concentrations for 24 h at 37 °C, cells were collected, washed with cold PBS at 4 °C and centrifuged at 1200 rpm for 3 min. The resulting pellet was mixed with the binding buffer of the Annexin V-FITC kit. FITClabeled Annexin V (5 ll) and PI (5 ll) were added to the 490 ll suspension and mixed gently. After incubation at 4 °C for 20 min in the dark, the cells were analyzed with a flow cytometry.

2.2. Cell culture A human myelomonocytic lymphoma cell lines, U937 and Molt4 were obtained from Human Sciences Research Resource Bank (Japan Human Sciences Foundation, Tokyo, Japan). HL-60 cells were kindly provided as a gift by Emeritus Prof. Yoshisada Fujiwara, Kobe University. The cells were grown in RPMI 1640 culture medium supplemented with 10% heat-inactivated fetal bovine serum (FBS) at 37 °C in humidified air with 5% CO2.

2.7. Assessment of intracellular reactive oxygen species (ROS) Evaluation of intracellular H2O2 was performed by flow cytometry using the fluorescence generate in the cells loaded with the peroxide-sensitive fluorescent probe, a H2O2 sensitive dye, dichlorofluorescein diacetate (DCFH-DA) (Molecular probes, Eugene, OR). DCFH-DA rapidly diffuses in to the cytosol of cells where it is hydrolyzed to the non-fluorescent, oxidation sensitive DCFH. In the presence of cytosolic peroxide, DCFH is rapidly oxidized to the non-diffusible, fluorescent DCF. Briefly, after treatment with SP-A and SP-B for different time periods, the cells were collected and washed with PBS, and were stained with 10 lM of DCFH-DA at 37 °C for 30 min. The fraction of DCF fluorescence positive cells was measured by flow cytometry as the proportion of cells containing intracellular H2O2 [25].

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Hydroethidine (HE, Molecular Probes, Eugene) was used to determine superoxide generation using the method of Gorman et al [26].

3. Results

2.8. Measurement of mitochondrial membrane potential (MMP)

The chemical structures of the HDAC inhibitors used in this study are shown in (Fig. 1). U937 cells were treated in a concentrationdependent manner for 24 h, followed by the colorimetric DNA fragmentation assay, a hallmark of apoptosis. As illustrated in Fig. 2 A, the results showed significant increase in the DNA fragmentation (%) with SP-A 30 nM and SP-B at 10 and 30 nM concentration as compared to control, respectively. While, 10 nM of SP-A does not induced any cell death in U937 cells. Further time-dependent analysis considering these two doses showed no DNA fragmentation at initial 6 and 12 h. However, DNA fragmentation was apparent at 18 h and increased at 24 h with SP-B (10 and 30 nM) and SP-A (30 nM) treatment (Fig. 2B and C). Thus, 10 and 30 nM of SP-A and SP-B were often used in this study. DNA fragmentation analysis was also done using human promyelocytic leukemia cells (HL-60) and Molt-4 cells. Both SP-A and SP-B induced apoptosis in a similar manner with those observed in U937 cells. Taken together these findings indicate that SP-A and SP-B induce apoptosis in human leukemia cells in a concentration- and time-dependent manner (Fig. 2D and E). Increased apoptosis was observed in U937 cells, as evidenced by DNA fragmentation analysis. Human lymphoma U937 cells were most susceptible to SP-A and SP-B treatment as compared to Molt-4 and HL-60 cells. Based on the finding U937 cells were selected to evaluate the detail mechanism involved in the apoptosis induced by SP-A and SP-B. Further, to examined the HDAC inhibition we analyzed the effects of SP-A and SP-B on acetylation of histones using Western blot, time dependent increase in the acetylated histone H3 and H4 levels were seen with SP-A and SP-B 30 nM (Fig. 2F). The cell morphology indicate that cells treated with SP-A and SP-B for 24 h showed typical signs of apoptosis, i.e. cytoplasmic aggregation, nuclear condensation, and fragmentation (Fig. 3A). Even at higher concentration of 100 nM, no signs of necrosis were observed (data not shown). Annexin V-FITC and PI double staining has been used extensively to distinguish between cells in early and late apoptosis. In this study we measured early and late apoptosis induced by SP-A (3–30 nM) and SP-B (3–30 nM) for 24 h in U937 cells. Flow cytometry using Annexin V-FITC and PI staining showed that treatment with SP-A (30 nM) and SP-B (10, 30 nM) caused significant apoptosis in U937 cells compared to control in a concentration dependent manner (p < 0.01). Interestingly, SP-B 10 nM induced significantly higher apoptosis than that of SP-A 10 nM (Fig. 3B and C).

To measure changes in MMP, U937 cells were harvested at different time periods and stained with 10 nM tetramethylrhodamine methyl ester (TMRM) (Molecular Probes) for 15 min at 37 °C in 1 ml of PBS, followed by the immediate flow cytometry of red TMRM fluorescence (excitation at 488 nm; emission at 575 nm). 2.9. Measurement of caspase-8 activity To measure caspase-8 activity, a FLICE/caspase-8 colorimetric protease assay kit (MBL, Nagoya, Japan) was used according to manufacturer’s instructions. The assay is based on the spectrophotometric detection of chromophore p-nitroanilide (pNA), a substrate of activated caspase-8. The pNA absorbance was quantified using a spectrophotometer at a wavelength of 400 nm (Beckman Instruments Inc., Fullerton, CA) [27]. 2.10. Western blot analysis for proteins Cells were collected and washed with cold PBS. They were lysed at a density of 2.5  106 cells/70 ll of RIPA buffer (50 mM Tris–HCl, 150 mM NaCl, 1% Nonidet P-40 (v/v), 1% sodium deoxycholate, 0.05% SDS, 1 lg of each aprotinin, pepstatin and leupeptin and 1 mm phenylmethyl sulfonyl fluoride) for 20 min. Following brief sonification, the lysates were centrifuged at 12,000g for 10 min at 4 °C, and the protein content in the supernatant was measured using the Bio- Rad protein assay kit (Bio-Rad, Hercules, CA). Protein lysates were denatured at 96 °C for 5 min after mixing with 2 ll SDS-loading buffer, applied on an SDS polyacrylamide gel for electrophoresis, and transferred to nitrocellulose membrane. Western blot analysis was performed to detect caspase-3, caspase-8, Bid, Bax, Bcl-2, Fas, Ac-H3 and Ac-H4 expression using specific antibodies. Blots were then probed with either secondary horseradish peroxide (HRP)-conjugated anti-rabbit or anti-mouse IgG antibodies obtained from Cell Signaling. Band signals were visualized on a luminescent image analyzer (LAS 4000, Fujifilm Co., Tokyo, Japan) by using chemiluminescence ECL detection reagents (Amersham Biosciences, Buckinghamshire, UK). For the preparation of cytosolic extracts, 5  107 cells were harvested and washed with 10 ml ice-cold PBS then centrifuged at 600g for 5 min and the supernatant was removed. Cells were resuspended in 300 ll of 1 Cytosol Extraction Buffer Mix containing DTT and a Protease inhibitor according to the manufacturer‘s instruction and then incubated on ice for 10 min. Cells were homogenized in an ice-cold dounce tissue grinder and then centrifuged at 3000 rpm for 10 min at 4 °C to remove nuclei and debris. The supernatant was collected and centrifuged again at 13,000 rpm for 30 min at 4 °C. The resulting supernatant was used as the soluble cytosolic fraction. Protein contents in cytosolic fraction were determined as described above. Following SDS–PAGE, Western blotting was performed to detect cytochrome-c release into cytosol using anti-cytochrome-c pAb (Santa Cruz Biotechnology Inc., Santa Cruz, CA) and anti-b-actin mAb was used as a control (Sigma).

3.1. Induction of apoptosis by SP-A and SP-B

2.11. Statistical analysis All values are expressed as the means ± S.D. of the respective test or control group. Statistical significance between the control group and test groups was evaluated by either Student’s t-test or one-way ANOVA. Values of p < 0.05 were considered to be significant. All experiments were performed in triplicate.

Fig. 1. Chemical structure of the Spiruchostatin A and Spiruchostatin B.

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Fig. 2. DNA fragmentation induced by SP-A and SP-B. (A) U937 cells were treated with SP-A and SP-B at different concentrations (3–100 nM) for 24 h followed by DNA fragmentation assay. (B) Time-dependent induction of DNA fragmentation by SP-A 10 and 30 nM in U937 cells. (C) Time-dependent induction of DNA fragmentation by SP-B 10 and 30 nM in U937 cells. (D) Molt-4 cells were treated with SP-A and SP-B 10 and 30 nM for 12 and 24 h followed by DNA fragmentation assay. (E) HL-60 cells were treated with SP-A and SP-B 10 and 30 nM for 12 and 24 h followed by DNA fragmentation assay. (F) SP-A and SP-B 30 nM caused a time-dependent increase in acetylated histone H3 and acetylated histone H4 levels. Where indicated, values for the treated cells are significantly different to the untreated cells at the same time point at ⁄p < 0.05; ⁄⁄p < 0.01. The results are presented as the mean ± S.D. (n = 3).

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Fig. 3. Assessment of apoptosis. (A) Morphological features of apoptosis after treatment of U937 cells with 10 and 30 nM SP-A and SP-B for 24 h. Signs of apoptosis were detected by Giemsa staining and then examined under microscope at 400 magnification. (B) Representative flow cytometric histograms of Annexin V-FITC/PI staining are shown (C) U937 cells were treated with 3, 6, 10 and 30 nM SP-A and SP-B. The cells were stained with Annexin V-FITC and PI for flow cytometry after 24 h. Where indicated, values for the treated cells are significantly different to the untreated cells at the same time point at ⁄p < 0.05; ⁄⁄p < 0.01.

3.2. Effects of SP-A and SP-B on cell cycle distribution

3.3. SP-A and SP-B induced ROS generation in U937 cells

Cell cycle distributions were examined in SP-A and SP-B treated U937 cells. U937 cells were treated with or without SP-A and SP-B (10 and 30 nM) for 18 and 24 h. As shown in Table 1, after 24 h incubation, the percentage of cells in sub-G1 population increased from 2.3 ± 1.2% to 27.0 ± 6.4% with SP-A 30 nM, from 3.0 ± 0.2% to 25.1 ± 1.9% with SP-B 10 nM and 36.7 ± 0.8% with SP-B 30 nM as compared to control. This increase in the percentage of sub-G1 population was determined by a decrease of cells in the G1 and G2/M phases, which was caused by apoptosis.

Induction of ROS has been implicated in the lethality of several HDACs inhibitors, however a detailed analysis of its role in SP-A and SP-B induced apoptosis has not been studied yet. Exposure of U937 cells with SP-A and SP-B 30 nM does not induce any profound change in ROS production at initial 3 and 6 h but a peak in the ROS generation was observed at 14 h with both the agents, more profound increased in the ROS production was observed with SP-B 30 nM as evidenced by the oxidation-sensitive dye dichlorofluorescein diacetate (DCFH-DA) (Fig. 4A–C). To confirm the role

Table 1 Cell cycle analysis by flow-cytometry. Data are mean ± S.D.; n = 3. 18 h

* **

24 h

SubG1 (%)

G0/G1 (%)

S (%)

G2/M (%)

SubG1 (%)

G0/G1 (%)

S (%)

G2/M (%)

SP-A Control 10 nM 30 nM

2.5 ± 0.7 2.0 ± 0.3 7.9 ± 0.2

37.6 ± 2.5 36.4 ± 4.9 14.8 ± 1.9

33.5 ± 0.8 33.3 ± 1.0 44.7 ± 1.6

19.7 ± 0.8 21.0 ± 4.8 24.0 ± 1.1

2.3 ± 1.2 2.9 ± 1.4 27.0 ± 6.4**

36.5 ± 3 35.0 ± 4.7 18.2 ± 2.3

34.5 ± 1.3 34.4 ± 3.4 37.0 ± 3.9

23.5 ± 0.6 23.8 ± 2.0 16.7 ± 2.9

SP-B Control 10 nM 30 nM

2.1 ± 0.8 5.3 ± 1.7* 12.3 ± 2.7**

39.0 ± 3.1 22.3 ± 5.1 13.1 ± 2.3

33.5 ± 0.8 39.4 ± 6.0 46.8 ± 5.6

19.7 ± 0.8 25.7 ± 1.4 17.4 ± 3.1

3.0 ± 0.2 25.1 ± 1.9** 36.7 ± 0.8**

36.3 + 3.3 25.3 ± 4.6 19.2 ± 2.3

34.5 ± 1.3 35.8 ± 4.1 37.3 ± 5.0

23.0 ± 1.0 17.0 ± 3.0 10.0 ± 1.0

p < 0.05 vs control. p < 0.01 vs control.

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Fig. 4. Effects of SP-A and SP-B on generation of intracellular ROS in U937 cells. (A and B) U937 cells were treated with 30 nM SP-A and SP-B for 14 h. SP-A and SP-B treated U937 cells exhibited an increase in DCF fluorescence compared with the control cells. One representative photomicrograph from three independent experiments is shown. (C) Induction of intracellular ROS generation by SP-A and SP-B. U937 cells were treated with SP-A and SP-B 30 nM for 3, 6 and 14 h. The cells were subjected to 10 lM DCFH-DA staining, were incubated at 37 °C for 30 min before measuring of the intracellular ROS by flow cytometry. Where indicated, values for the treated cells are significantly different to the untreated cells at the same time point at ⁄⁄p < 0.01. The results are expressed as means ± S.D. (n = 3). (D and E) U937 cells were pre-treated with or without 5 mM NAC for 1 h and stimulated with SP-A and SP-B 30 nM for 24 h followed by measurement of apoptosis with DNA fragmentation assay and Annexin V-FITC/PI staining, ⁄⁄ p < 0.01 as compared to treatment alone. The results are expressed as means ± S.D. (n = 3).

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of ROS production in the apoptosis induced by SP-A and SP-B, cells were pre-treated with 5 mM NAC (an antioxidant, N-acetyl-L-cysteine) for 1 h. NAC pre-treatment significantly inhibited the apoptosis induced by SP-A and SP-B (p < 0.01) (Fig. 4D and E), suggesting a crucial role of ROS generation in the apoptosis induction by these agents.

3.4. Measurement of mitochondrial membrane potential To assess the effects of SP-A and SP-B on the mitochondrial apoptotic pathway, mitochondrial membrane potential of U937 cells treated with SP-A (10, 30 nM) and SP-B (10, 30 nM) was evaluated. A significant time and concentration-dependent MMP loss was observed (Fig 5C and D).

3.5. Effects of SP-A and SP-B on activation of extrinsic pathway of apoptosis To determine the involvement of extrinsic pathway of apoptosis, effects of SP-A and SP-B on Fas and caspase 8 were investigated. The Fas receptor is a death receptor on the surface of cells that leads to one of the apoptotic pathways, extrinsic pathway, through DISC assembly and subsequent caspase-8 activation. SP-A and SP-B induced a clear time-dependent increase in the protein expression of Fas at 30 nM concentration (Fig. 6A). Simultaneously, increased in the cleaved caspase-8 protein expression following treatment with SP-A (30 nM) and SP-B (10, 30 nM) for 24 h was observed. Moreover, increased caspase-8 activity was also observed with

SP-A (30 nM) and SP-B (10, 30 nM) as compared to control (Fig. 6B and C). 3.6. SP-A and SP-B induced caspase activation Caspases play a key role in the apoptosis signaling pathways. To investigate the role of caspase-3 in the apoptosis induced by SP-A and SP-B. Cells were treated with SP-A and SP-B, activation of caspase cascade was determined by Western blot using caspase 3 specific antibody. An increased in the cleaved form of caspase-3 was observed following treatment with SP-A (30 nM) and SP-B (10, 30 nM) for 18 and 24 h (Fig 7). 3.7. Changes in the apoptosis-related proteins Bcl-2 family proteins with anti- or pro-apoptotic functions cause the release of cytochrome-c from the mitochondrial membrane, leading to the activation of caspase cascade. In this study, Western blot analysis revealed the activation of Bid following treatment with SP-A 30 nM and SP-B (10, 30 nM) for 18 and 24 h in the U937 cells. The expression of pro-apoptotic Bax and an anti-apoptotic Bcl-2 remained unchanged (Fig 8A). In addition, cytochrome-c release from mitochondria to cytosol was induced by SP-A (30 nM) and SP-B (10, 30 nM) at 24 h (Fig 8B). 4. Discussion SP-A and SP-B have been reported for the potent HDAC inhibitory activity [18,19], however the molecular mechanism involved

Fig. 5. Effects of SP-A and SP-B on mitochondrial membrane potential. (A and B) Representative flow cytometric histograms of SP-A and SP-B induced MMP loss. (C) Timedependent effects of SP-A 10 and 30 nM on the mitochondrial membrane potential of U937 cells. (D) Time-dependent effects of SP-B 10 and 30 nM on the mitochondrial membrane potential of U937 cells. The loss of MMP was observed in the cells treated with SP-A and SP-B, as determined by flow cytometry using TMRM staining. The results are expressed as means ± S.D. (n = 3).

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Fig. 6. Effects of SP-A and SP-B on Fas and caspase-8 activation. (A) Time-dependent changes of Fas expression after treatment with SP-A and SP-B 30 nM in U937 cells. (B) Concentration-dependent changes in the protein expression of caspase-8 after treatment with SP-A for 24 h. The caspase-8 activity in U937 cells induced by SP-A was also measured with FLICE/caspase-8 colorimetric protease kit (C) Concentration-dependent changes in the protein expression of caspase-8 after treatment with SP-B for 24 h. The caspase-8 activity in U937 cells induced by SP-B was also measured with FLICE/caspase-8 colorimetric protease kit. Where indicated, values for the treated cells are significantly different to the untreated cells at the same time point at ⁄p < 0.05, ⁄⁄p < 0.01. The results are presented as the mean ± S.D. (n = 3), as determined by one way ANOVA.

in the lethality of these agents needs further investigation. This is the first report that evaluated the apoptosis activity of these agents in U937 cells. Both SP-A and SP-B share many structural activities with FK228, which is already approved for the treatment of CTCL. Both compounds possess intermolecular disulfide bond that is in FK228 reduced intra-cellularly to provide the zinc-binding group and ROS may be generated in the redox-cycling reaction and take part in apoptosis [18,28]. Recently, several HDAC inhibitors have been reported to induce apoptosis through ROS generation including FK228, SAHA, trichostatin A, sodium butyrate, MS275 and LAQ-824. [15,29,30]. Suberoylanilide hydroxamic acid (SAHA) and FK228 were reported to induce a cell death pathway by cleavage of Bid [31,32]. A recent study showed that FK228 induce apoptosis via ROS generation, and FK228-induced apoptosis was attenuated by pre-incubation with NAC (N-acetyl cysteine), suggesting that ROS production is likely involved in the mode of action of FK228 in solid cancer cell lines, caused mitochondrial damage and apoptosis [28] In consistent, we have demonstrated that in human lymphoma U937 cells, a marked increase in the ROS production was observed with SP-A and SP-B. Pre-incubation with NAC significantly blocked the generation of ROS and apoptosis induced by SP-A and SP-B, supporting the notion that ROS production is involved in the death mechanism. In term of differences between SP-A and SP-B, we did not

detect any clear differences between these two structurally similar compounds but both the agents can induce apoptosis associated with increase in p21 [20]. Therefore, it might be possible that these agents can induce apoptosis via a ROS production and p21 increase as FK228 does, or indirectly by the activation of NAD (P) H oxidase and xanthine oxidase [33,34]. However, further future investigations are necessary to rule out the source of this ROS generation. In this study we mainly detected changes in ROS level by DCFHDA probe, which is more sensitive towards hydrogen peroxide [35]. Unfortunately, we did not detect any significant changes with other probes like HE (data not shown). The mechanism involved in the HDAC inhibitors induced cell death is still unclear. Although, oxidative stress has been identified as a mechanism involved in the lethality of HDAC inhibitors but the manner by which HDACi induce oxidative stress is poorly understood. Two distinct mechanisms are apparently involved; one involves the mitochondrial injury, while other implicates the modulation of anti-oxidant levels. HDACi induces expression of Bid, which binds to and disrupts the mitochondrial membrane and resulted in increased ROS and apoptosis [31,36]. Previously, in osteosarcoma and chronic lymphocytic leukemia cells, depsipeptide has been reported to induce apoptosis selectively via a caspase-8 (extrinsic) pathway [37,38]. Depsipeptide induced apoptosis via activation of mitochondrial pathway, independent

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Fig. 7. Effects of SP-A and SP-B on activation of caspase-3. Concentration-dependent changes in the expression of caspase 3 after 18 and 24 h incubation with SP-A and SP-B. Date is representative of three independent experiments.

Fig. 8. Effects of SP-A and SP-B on apoptosis related proteins and release of cytochrome-c. (A) U937 cells were treated with SP-A and SP-B in a dose-dependent manner for 18 and 24 h. The cell lysates were harvested for Western blot analysis. Changes in expressions of Bax, Bcl-2, and Bid induced by SP-A and SP-B. (B) Release of cytochrome-c (Cytc) in cytosolic fraction after treatment with SP-A and SP-B for 24 h. One representative picture from three independent experiments is shown.

of caspase activation in acute leukemia cells [32] and in the small cell lung cancer (SCLC) induced caspase dependent apoptosis via mitochondrial pathway without the involvement of death receptor

pathway [39]. In contrast to these reports, findings with SP-A and SP-B demonstrate the involvement of both the extrinsic and intrinsic pathway of apoptosis. The discrepancy in between the current

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and previous report can be resolved by accounting the fact that the induction of different cell pathways by different HDACi may be dependent on the different targets and the cell type used. In this study, treatment with SP-A and SP-B significantly caused MMP loss in a time-dependent manner with a concurrent release of cytochrome-c into the cytosol. These findings suggest the involvement of mitochondria in SP-A and SP-B induced cell death. The involvement of the extrinsic pathway is suggested by the increased Fas and caspase-8 activation with SP-A and SP-B treatment. The extrinsic pathway is due to the direct interaction of cell surface receptors, such as Fas, with caspase-8, that in turn activates downstream effector caspases. Bcl-2 family proteins are involved in pro- or anti-apoptotic processes by interacting with the mitochondria [40]. HDACi, alter the factors that are involved in the regulation of intrinsic pathway. Bid activation, can initiate the intrinsic pathway [9]. In consistent, our findings also revealed activation of Bid with both SP-A and SP-B treatment. Bid can be cleaved by caspase-8 and the cleaved Bid as the carboxyl-terminal fragment translocates to the mitochondria to induce the release of cytochrome-c [41,42]. These two apoptotic pathways may be interconnected by the caspase-8-mediated cleavage of Bid, which triggers the activation of the mitochondrial pathway. In conclusion, the current study is the first to demonstrate that SP-A and SP-B induces apoptosis in human lymphoma U937 cells. SP-A and SP-B treatment resulted in increased accumulation of ROS and pre-treatment with NAC significantly prevented the apoptosis induced by these agents suggesting the involvement of ROSdependent pathways in the cell death. Additionally, FAS activation may also play an important role in the SP-A and SP-B induced apoptosis. Our findings may help the potential application of these agents as future chemo-therapeutics agents. Conflict of Interest The author has no conflict of interest. Transparency Document The Transparency document associated with this article can be found in the online version.

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