- Email: [email protected]
Low voltage pulses can induce apoptosis
Available online at www.sciencedirect.com
Cancer Letters 269 (2008) 93–100 www.elsevier.com/locate/canlet
Low voltage pulses can induce apoptosis Noriaki Matsuki a,b,*, Takuji Ishikawa a, Yousuke Imai a, Takami Yamaguchi a a
Department of Bioengineering and Robotics, Tohoku University, 6-6-01, Aoba, Aramaki, Aoba-ku, 980-8579 Sendai, Japan b New Industry Creation Hatchery Center, Tohoku University, 6-6-10, Aoba, Aramaki, Aoba-ku, 980-8579 Sendai, Japan Received 21 January 2008; received in revised form 14 April 2008; accepted 15 April 2008
Abstract Electroporation is used for gene transfection, drug delivery, and cell fusion. While studies have shown that high voltage electroporation induces apoptosis in vitro, a strong electric field can lower cell survival rates. As there are no published reports which have examined apoptotic properties associate with low voltage electric charges, we demonstrated for the first time that consecutive low voltage pulses with a voltage lower than the membrane breakdown threshold of human cells can increase the membrane potential to the threshold required to induce electroporation. This led to apoptosis through caspase pathways. Moreover, necrotic cell damage was less than that caused by high voltage pulses. Therefore, low voltage electroporation can be a suitable anticancer method. Ó 2008 Elsevier Ireland Ltd. All rights reserved. Keywords: Apoptosis; Caspase; Electroporation; Low voltage pulse
1. Introduction Electroporation has been commonly used in a variety of in vitro [1–3] and in vivo [4–7] biotechnical applications. The applied electric field generates large depolarizing and hyperpolarizing transmembrane potentials across the cell. Electrical membrane breakdown occurs when the membrane potential reaches a critical value, resulting in the formation of pores. The number and size of the pores are determined by the pulse voltage, pulse
*
Corresponding author. Address: Department of Bioengineering and Robotics, Tohoku University, 6-6-01, Aoba, Aramaki, Aoba-ku, 980-8579 Sendai, Japan. Tel.: +81 22 714 8514; fax: +81 22 795 6959. E-mail address: [email protected] (N. Matsuki).
duration, waveform, pH, and the ionic strength of the suspending medium [2,7–12]. If the electric field is relatively weak, the pores shrink and disappear once the pulse is removed, and the electrical membrane breakdown can thus be reversed. Conversely, when a strong electric field is applied, the pores continue to exist long after the pulse, leading to irreversible damage of the cell membrane [13– 16]. During the effective pore open time, solutes added to the extracellular medium that are normally unable to permeate the membrane will be able to enter cell via the pores [16]. Cell death occurs by either necrosis or apoptosis [17]. Necrotic cells are characterized by irreversible plasma membrane rupturing as a result of severe cell damage and dysfunction, which in turn induces inflammatory reactions. In contrast, the membrane integrity of an apoptotic cell is maintained, and the
0304-3835/$ - see front matter Ó 2008 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.canlet.2008.04.019
94
N. Matsuki et al. / Cancer Letters 269 (2008) 93–100
cell is removed by phagocytosis without inducing an inflammatory response. Therefore, apoptosis is considered to be the most suitable method for novel targeted anticancer therapy [18]. Apoptosis is characterized by morphological and biochemical alterations of a cell including the externalization of phosphatidylserine on the plasma membrane, membrane blebbing, chromatin condensation, DNA fragmentation, and the degradation of intracellular proteins such as poly-ADP-ribose polymerase (PARP), lamine, and others. It involves the concerted action of many enzymatic steps including caspase activation [17,19]. Apoptosis signals can arise from the plasma membrane [17] or intracellular structures [20,21] such as the nucleus [17], mitochondria [22], and endoplasmic reticulum [23]. These signals activate pathways that converge to activate caspases [24,25]. In the past, several studies have reported that high electric voltage electroporation (>4.5 kV/ cm) can induce mild apoptosis as well as severe necrosis [26,27]. However, such a strong electric field reduces cell survival through severe cell explosion and causes undesirable side effects such as burn due to Joule heating and severe inflammatory reaction due to necrosis. On the other hand, electroporation with relatively low voltage electric field (0.2 kV/cm) demonstrated successful gene transfection in vivo [28]. Therefore, apoptosis induced by a mild electric field would be a favorable alternative to high voltage electroporation, and it would also be suitable for targeted anticancer treatments. To date, no published studies have examined the apoptotic capabilities of low voltage electric fields. We thus investigate whether low voltage and long duration pulses can induce apoptosis and examine the role of caspase pathways in the response. 2. Materials and methods 2.1. Cell culture The SCC 9 (human squamous cell carcinoma of the tongue) cell line was obtained from and cultured as recommended by the American Type Culture Collection (Rockville, MD, USA). Cells were cultured in RPMI 1640 (Irvine Scientific, Santa Ana, CA, USA) media containing 10% heat-incubated fetal bovine serum (FBS; Gibco, Grand Island, NY, USA), 50 U/ml penicillin (Gibco), and 50 lg/ml streptomycin (Gibco) at 37 °C in a 95% humidified, 5% carbon dioxide atmosphere.
2.2. Electroporation Cells were collected by trypsin treatment and centrifuged for 5 min at 300g, washed with phosphate buffer saline (PBS), and resuspended in PBS or RPMI 1640 at a concentration of 2 106 cells/ml. The cell suspensions (400 ll) were transferred to parallel aluminium-plated electroporation/fusion chambers with 2 mm gaps. Low voltage long duration (7.5 V/mm, 100 ms) and high voltage short duration (1.5 kV/mm, 0.05 ms) square wave pulses were applied using a Gene Pulser II (Bio-Rad, Hercules, CA, USA). 2.3. Detection of membrane breakdown (7-AAD uptake) Membrane breakdown was assessed by staining the cells with 10 ll 7-amino-actinomycin D (7-AAD; Becton–Dickinson, McKinley, MN, USA) and flow cytometric analysis. Cells were washed, resuspended at a concentration of 2 106 cells/ml in PBS, and stained with 10 ll 7-AAD. Following exposure to electrical pulses, the cells were immediately analyzed by flow cytometry according to the manufacturer’s protocol. 2.4. Viability assay For the viability analysis, the cells were washed and resuspended at a concentration of 2 106 cells/ml with PBS. After 2 h of exposure to electric pulses, the cells were stained with 10 ll 7-AAD and analyzed by flow cytometry. 2.5. Apoptosis assay A flow cytometry apoptosis detection kit (Becton– Dickinson) was used to identify apoptotic and necrotic cells according to the manufacturer’s protocol. Briefly, cells (3 105 cells/ml) were double-stained with 10 ll fluorescein isothiocyanate-labeled-Annexin V (R&D Systems, Minneapolis, MN, USA) to detect phosphatidylserine expression during early apoptotic phases, and 10 ll 7AAD to exclude late apoptotic and necrotic cells. Samples were analyzed using an EPICS XL-MCL cytometer (Beckman Coulter, Fullerton, CA, USA). The number of positive cells was determined in the annexin-positive fraction of the cell suspension by using an arbitrary threshold setting, which allows no more than 5% positive counts in the negative control. Cells positive for annexin only were defined as apoptotic, whereas those positive for both annexin and 7-AAD were defined as necrotic/late apoptotic. 2.6. CPP 32 activity (caspase-3-like enzyme activity) Caspase activity was measured as described previously [24]. Briefly, cells were washed twice with PBS, and the pellets were lysed in PBS containing 0.2% Triton X-100
N. Matsuki et al. / Cancer Letters 269 (2008) 93–100
on ice for 10 min and centrifuged at 12,000 rpm for 5 min. The cell extracts were incubated with 10 lM Ac-DEVDMCA (Peptide Institute, Osaka, Japan) as a fluorescent substrate in the incubation buffer [50 lM Tris–HCl (pH 7.5), 1 mM DDT] for 30 min at 37 °C. The fluorescence intensity of the substrate cleaved by caspase-3 was measured at 380 nm for excitation and 460 nm for emission by a spectrofluorometer.
95
10
5
2.7. Statistical analysis 0
Statistical analysis was performed using StatView 5.0 software (SAS Institute Inc., Cary, NC, USA). Comparisons of means were conducted using analysis of variance (ANOVA) with P < 0.05 considered to be statistically significant.
To determine the membrane breakdown threshold voltage, cells in PBS containing 7-AAD were exposed to a square wave with a 100 ms duration, single electric pulses of different voltages (0–50 V), and to a single square wave, 3000 V, 0.05 ms duration pulse, then they were analyzed using flow cytometry. The cells exhibited 5.55 ± 1.51%, 5.58 ± 1.42%, 5.85 ± 1.55%, 6.90 ± 1.71%, and 10.30 ± 1.31% 7-AAD uptake for control, 20, 30, 50, and 3000 V treatments, respectively. Therefore, the membrane breakdown threshold of SCC 9 cells was determined to be between 30 and 50 V (Fig. 1a). To determine whether consecutive electric pulses with a lower voltage than the membrane breakdown threshold could disrupt the structural integrity of the membrane, cell suspensions containing 7-AAD were exposed to 15 V, 100 ms duration, square wave electric pulses with an interval of 100 ms and immediately analyzed using flow cytometry. The control cells exhibited 5.55 ± 1.51% 7-AAD uptake, whereas treated cells showed 5.53 ± 1.46% uptake for a single pulse (P < 0.05 vs. control and single pulse), 6.90 ± 1.71% for 10 pulses (P < 0.05 vs. control, single and 10 pulses), 14.30 ± 3.09% for 100 pulses, and 19.52 ± 4.26% for 200 pulses (Fig. 1b). The 3000 V square wave, 0.05 ms duration, 5-pulse treatment resulted in 48.47 ± 14.30% 7-AAD-uptake (data not shown). Therefore, the 7-AAD-uptake of the cells increase as the pulse frequency is increased. This indicates that pulses with a lower voltage than the membrane breakdown threshold are able to break the membrane as the pulse frequency increases. 3.2. Viability It can take seconds, minutes, or hours for a membrane to recover (reseal) following electroporation [13–15]. Therefore, to examine cell viability, cells were stained with
30 V
50 V
3000 V
25
7 - AAD uptake (%)
3.1. Membrane breakdown
20 V
Pulse voltage (V)
20 15 10 5 0 control
1
10
100
200
Pulse frequency (Times) Fig. 1. The 7-AAD uptake of cells after the application of electric pulses. (a) Cells cultured in the presence of 7-AAD were exposed to single electric pulses of different voltages and analyzed using flow cytometry. (b) Cells cultured in the presence of 7-AAD were exposed to consecutive 15 V, 100 ms pulses of different frequencies and analyzed using flow cytometry. Data are shown as means ± SEM of four separate experiments.
100
Viability ( %)
3. Results
control
75
50
25
0 control
15 V
3000 V
Electric pulse application Fig. 2. Cell viability after the application of electric pulses. Cells were exposed to 200 low (15 V, 100 ms) or five high (3000 V, 0.05 ms) voltage electric pulses. The 7-AAD was added 2 h after the administration of the electric pulses. Viability was analyzed using flow cytometry. Data are shown as means ± SEM of four separate experiments.
N. Matsuki et al. / Cancer Letters 269 (2008) 93–100
10
4
10
3
10
2
10
1
10
0
control
15 V
104
103
7-AAD
7-AAD
96
102
101
100 10
0
10
1
10
2
10
3
10
4
100
101
10
3
10
2
10
1
10
0
102
103
104
10 3
10 4
Annexin
3000 V
cisplatin
104
103
7-AAD
4
102
101
100 10
0
10
1
10
2
10
3
10
4
10 0
Annexin
10 1
10 2
Annexin
15
Apoptosis rate ( %)
10
10
5
0 control
15 V
3000 V
cisplatin
Electric pulse / Cisplatin application 25
Necrosis rate ( %)
7-AAD
Annexin
20 15 10 5 0
control
15 V
3000 V
cisplatin
Electric pulse / Cisplatin application
7-AAD 2 h following the application of electric pulses and analyzed by flow cytometry. The control cells showed 91.8 ± 1.6% viability. Cells exposed to 15 V pulses showed 82.3 ± 4.5% viability, whereas 65.1 ± 3.0% of those treated with 3000 V survived (P < 0.01 vs. control and P < 0.02 vs. 15 V; Fig. 2). These data indicate that high voltage pulses severely damage cells. 3.3. Induction of apoptosis and cell death To determine whether low voltage pulses can induce apoptosis, cells were exposed to 200 15 V, 100 ms, square wave pulses and analyzed 48 h later using flow cytometry with an apoptosis detection kit. Cells treated with cisplatin (3.0 lg/ml) for 48 h were used as positive controls for apoptosis. Fig. 3a shows the flow cytometric plots of annexin and 7-AAD staining. Cells exposed to electrical pulses shifted to the right and upward compared to control cells. These results indicate that electrical pulses can induce apoptosis and necrosis. The apoptotic rates of the cells are shown in Fig. 3b. On average, apoptosis occurred in 3.0 ± 0.9% of the control cells, 6.0 ± 0.8% of the cells exposed to 15 V (P = 0.0514 vs. control), 13.2 ± 1.7% of the cells exposed to 3000 V (P < 0.01 vs. control and 15 V), and 6.7 ± 1.5% of the cisplatin-treated cells (P < 0.05 vs. 3000 V). Fig. 3c shows necrosis rates of the cells exposed to electrical pulses. On average, cell death occurred in 3.7 ± 1.2% of the control cells, 6.2 ± 1.4% of cells exposed to 15 V, 16.9 ± 5.2% of cells exposed to 3000 V (P < 0.05 vs. control), and 8.9 ± 0.8% of cisplatin-treated cells (P < 0.02 vs. control). Together, these data indicate that high voltage pulses were capable of inducing significant cell damage by necrosis as well as apoptosis. In contrast, low voltage pulses resulted in fewer damaged (necrotic) cells while still inducing apoptosis. 3.4. CPP 32 (caspase-3-like enzyme) activity To determine whether caspase pathways were involved in apoptosis induced by electric pulses, cells were exposed to 200 square wave pulses at 15 V for 100 ms each. CPP32 activity was measured 24 h following the application of electric pulses using synthetic peptide Ac-DEVD-MCA as a substrate. Cells treated with cisplatin (3.0 lg/ml) for 24 h were used as positive controls for activated caspase.
CPP 32 activity ( Fluorescence Intensity)
N. Matsuki et al. / Cancer Letters 269 (2008) 93–100
97
800
600
400
200
0 control
15 V
3000 V
cisplatin
Electric pulse / Cisplatin application Fig. 4. CPP32 activity after the administration of electric pulses. CPP32 activity was measured using the synthetic peptide AcDEVD-MCA as a substrate 24 h after the application of electric pulses. Data are shown as fluorescence intensity (FI) and means ± SEM of four separate experiments, each performed in duplicate.
Fig. 4 illustrates the CPP32 activities after the application of electric pulses. The control cells showed an average of 161.6 ± 31.9 fluorescence intensity (FI). Cells exposed to electrical pulses showed a significant increase of caspase activities. Specifically, 15 V produced 305 ± 25.8 FI (P < 0.01 vs. control), 3000 V induced 806.8 ± 35.4 FI (P < 0.01 vs. control and 15 V), and cisplatin-treated cells exhibited 604.6 ± 78.8 FI (P < 0.01 vs. control and 15 V; P < 0.02 vs. 3000 V). This indicates that caspase pathways play a significant role in electroporation-induced apoptosis.
4. Discussion In the past, several publications reported that high voltage electroporation could induce apoptosis in vitro [26,27]. However, applying such strong electric fields tend to severely diminish cell survival rates and induce undesired side effects as a result of Joule heating. We have demonstrated that mild electric fields are capable of inducing apoptosis while minimizing the undesirable side effects that accompany necrosis. Therefore, this shows great potential as an anticancer treatment method.
3 Fig. 3. Induction of apoptosis and cell death following the administration of electric pulses. Cells were exposed to 200 low (15 V, 100 ms) or five high (3000 V, 0.05 ms) voltage electric pulses and analyzed using a flow cytometry with annexin and 7-AAD staining carried out 48 h later. Cells treated with cisplatin (3.0 lg/ml) for 48 h were used as positive controls for apoptosis. (a) Flow cytometry plot of annexin and 7-AAD staining. Annexin () and 7-AAD () cells were considered intact; annexin (+) and 7-AAD () cells were considered apoptotic; 7-AAD (+) cells were considered necrotic. (b) Apoptosis rate. (c) Necrosis rate. Data are shown as means ± SEM of four separate experiments.
98
N. Matsuki et al. / Cancer Letters 269 (2008) 93–100
Given the results of the 7-AAD uptake experiments (Fig. 1a and b), the membrane breakdown threshold of SCC 9 cells was expected to be between 30 and 50 V. However, this is dependent on the membrane composition and the pulse duration as well as other stresses [2,14,16]. Therefore, we chose 15 V, less than a half of the expected membrane breakdown threshold of SCC 9 cells, for the low voltage electrical stimulation. The transmembrane potential (Vm) is theoretically given as follows: V m ¼ 1:5E a cos h where E is the applied electric field strength (7.5 V/mm), a is the cell radius (10 lm), and h is the position angle relative to the electrical field (cos h 5 1) (Fig. 5) [11]. According to this formula, Vm is expected to be approximately 0.11 V. This is much less than the threshold at which typical mammalian cells experience membrane breakdown (electroporation), which is reported to be between 0.2 and 1.5 V [14]. In our experiment using the single pulse application, no membrane breakdown could be detected at this voltage. However, it is possible that micropores smaller than the 7-AAD molecules were generated. This would still block 7-AAD from entering the cell and they would thereby remain undetected [29]. Our data indicated that there is an increase in the membrane permeability as the pulse frequency increases. Ryttsen et al. [10] reported that the transmembrane potential increased during the first 50 ms, reached the maximum state where it stabilized for up to 1 ms after the application of the elec-
Cell
Vm a Θ
π
0
E Fig. 5. Schematic of a spherical cell with radius a in an electric field. The transmembrane potential Vm is theoretically given as 1.5 E a cos h. The electric field E is oriented such that the depolarized pole is at h = 0 and the hyperpolarized pole is at h = p.
trical pulse. Gowrishankar et al. reported that longer pulse can induce more electroporation [30]. Since we have applied 100-ms pulses with an interval of 100 ms, it is possible that the transmembrane potential could accumulate with each pulse, eventually reaching the threshold and resulting in electroporation. The net percentage (sample-control value) of electroporated cells, as defined by the 7-AAD uptake immediately following the application of electric pulses, was 14% for cells exposed to 15 V and 43% for cells exposed to 3000 V. Similarly, the net percentage of apoptosis/necrosis as defined by the percentage of cells stained with annexin/7AAD 2 days following the application of electric pulses, was higher in cells exposed to high voltage pulses than those exposed to low voltage pulses (10.2% and 13.2% vs. 3% and 2.5% for high and low voltages, respectively). Therefore, apoptosis was induced in 21.4% of the electroporated cells using low voltage pulses and 23.7% of the electroporated cells using high voltage pulses. Interestingly, the ratio between apoptosis and necrosis in the electroporated cells was higher in the low voltage group (1.2 vs. 0.77 for low and high voltages, respectively). Therefore, we have demonstrated that low voltage pulses were capable of inducing less necrosis (i.e., fewer harmful side effects), and more apoptosis than high voltage treatments. Caspase activity, a marker for apoptosis via the caspase pathways, was observed to increase with increasing voltage. These results are in accordance with apoptosis levels detected by annexin. Hofmann et al. [26] also reported the occurrence of caspase activation occurred in high voltage-induced apoptosis. To date, the mechanisms and pathways of apoptosis in electroporated cells remain unclear. Several reports [26,27] have suggested that electroporation with high voltage electrical pulse charges induces apoptosis by changing the membrane integrity via the influx of extracellular substances including ions such as Ca++, Na+, K+, and Cl [26]. This is thought to break the membranes of organelles such as mitochondria and the endoplasmic reticulum, and activate intracellular signaling [31]. Interestingly, very high voltage (MeV/m), ultrashort (nanosecond) pulses can also induce apoptosis without membrane breakdown [32–36] and bring DNA fragmentation [37]. However, the formation of undetectable micropores may take place. Apoptosis induced by electrical charges could be based on changing the membrane integrity, the pore forma-
N. Matsuki et al. / Cancer Letters 269 (2008) 93–100
tion, and the activation of caspase enzymes. Prausnitz et al. [12] reported that transport caused by electroporation occurs predominantly by electrophoresis and electroosmosis during a pulse, and by diffusion after pulse. Therefore, it is possible that ions such as Ca++ could pass through micropores and activate caspase pathways. In summary, consecutive pulses of a voltage less than the membrane breakdown threshold can increase the membrane potential to the threshold, disrupt the membrane integrity, cause membrane permeabilization through pores, lead to the influx of extracellular substances including ions, activate caspase pathways, and induce apoptosis. Furthermore, low voltage electroporation can induce apoptosis while mitigating cellular damage that typically accompanies high voltage electroporation such as that caused by necrosis. Therefore, low voltage electroporation can represent a useful anticancer method targeting apoptosis. Further studies are required to define the exact threshold of membrane breakdown and resolve the apoptosis signaling mechanisms involved in electroporation. Acknowledgments This study was supported by the Japanese Society for the Promotion of Science and the Austrian Academic Exchange Service. References [1] T.Y. Tsong, Electroporation of cell membranes, Biophys. J. 60 (1991) 297–306. [2] J.C. Weaver, Y.A. Chizmadzhev, Theory of electroporation: a review, Bioelectrochem. Bioenerg. 41 (1996) 135–160. [3] J.C. Weaver, Electroporation of cell and tissues, IEE. Trans. Plasm. Sci. 28 (2000) 24–33. [4] B. Rubinsky, Irreversible electroporation in medicine, Technol. Cancer. Res. Treat. 6 (2007) 255–259. [5] J.P. Cantella, M.M. Black, M.D. Bonnichsen, Tissue electroporation: quantification and analysis of heterogenous transport in multicellular environments, Biophys. J. 86 (2004) 3260–3268. [6] G. Sersa, D. Miklavcic, M. Cemazar, Electrochemotherapy in treatment of tumours, Eur. J. Surg. Oncol. 34 (2008) 232–240. [7] J.A. Kim, K. Cho, M.S. Shin, A novel electroporation method using a capillary and wire-type electrode, Biosens. Bioelectron. 23 (2008) 1353–1360. [8] J. Deng, H.K. Schoenbach, S. Buescher, The effects of intense submicrosecond electrical pulses on cells, Biophys. J. 84 (2003) 2709–2714. [9] J.P. Cantella, F.J. Karr, A.J. Petros, Quantitative study of electroporation-mediated molecular uptake and cell viability, Biophys. J. 80 (2001) 755–764.
99
[10] F. Ryttsen, C. Farre, C. Brennan, Characterization of single cell electroporation by using patch–clamp and fluorescence microscopy, Biophys. J. 79 (2000) 1993–2001. [11] K. DeBruin, W. Krassowska, Modeling electroporation in a single cell. I. Effects of field strength and rest potential, Biophys. J. 77 (1999) 1213–1224. [12] M.R. Prausnitz, J.D. Corbett, J.A. Gimm, Millisecond measurement of transport during and after an electroporation pulse, Biophys. J. 68 (1995) 1864–1870. [13] G. Saulis, Pore disappearance in a cell after electroporation: theoretical simulation and comparison with experiments, Biophys. J. 73 (1997) 1299–1309. [14] R.T. Gowrishankar, U. Pliquett, R. Lee, Dynamics of membrane sealing in transient electropermeabilization of skeletal muscle membranes, Ann. NY Acad. Sci. 888 (1999) 196–210. [15] F. Hilpert, A. Heiser, W. Wieckhorst, The impact of electrical charge on the viability and physiology of dendritic cells, Scand. J. Immunol. 62 (2005) 399–406. [16] M. Tarek, Membrane electroporation: a molecular dynamics simulation, Biophys. J. 88 (2005) 4045–4053. [17] M.O. Hengarther, The biochemistry of apoptosis, Nature 407 (2000) 770–776. [18] A. Russo, M. Terrasi, V. Agnese, Apoptosis: a relevant tool for anticancer therapy, Ann. Oncol. 17 (2006) vii115–123. [19] J. Liu, A. Lin, Role of JNK activation in apoptosis: a double-edged sword, Cell Res. 15 (2005) 36–42. [20] M.H. Beere, The stress of dying: the role of heat shock proteins in the regulation of apoptosis, J. Cell Sci. 117 (2004) 2641–2651. [21] M.H. Beere, Death versus survival: functional interaction between the apoptotic and stress-inducible heat shock protein pathways, J. Clin. Invest. 115 (2005) 2633–2639. [22] K. Maaser, P.A. Sutter, H. Scheruebl, Mechanisms of mitochondrial apoptosis induced by peripheral benzodiazepine receptor ligands in human colorectal cancer cells, Biochem. Biophys. Res. Commun. 332 (2005) 646–652. [23] J.S. Kim, M.K. Park, N. Kim, Doxorubicin prevents endoplasmic reticulum stress-induced apoptosis, Biochem. Biophys. Res. Commun. 339 (2005) 463–468. [24] Y. Fukamachi, Y. Karasaki, T. Sugiura, Zinc suppresses apoptosis of U937 cells induced by hydrogen peroxide through an increase of the Bcl-2/Bax ratio, Biochem. Biophys. Res. Commun. 246 (1998) 364–369. [25] R.C. Taylor, S.P. Cullen, S.J. Martin, Apoptosis: controlled demolition at the cellular level, Nature Rev. Mol. Cell Biol. 9 (2008) 231–241. [26] F. Hofmann, H. Ohnimus, C. Scheller, Electric field pulses can induce apoptosis, J. Membr. Biol. 169 (1999) 103–109. [27] J. Pinero, M. Lopez-Baena, T. Ortiz, Apoptotic and necrotic cell death are both induced by electroporation in HL60 human promyeloid leukemia cells, Apoptosis 2 (1997) 330– 336. [28] D. Machado-Aranda, Y. Adir, J.L. Yong, Gene transfer of Na+, K+-ATPase b1 subunit using electroporation increases lung liquid clearance, Am. J. Respir. Crit. Care Med. 171 (2005) 204–211. [29] P.T. Vernier, Y. Sun, Y. Marcu, Calcium bursts induced by nanosecond electric pulses, Biochem. Biophys. Res. Commun. 310 (2003) 286–295. [30] T.R. Gowrishankar, J.C. Weaver, Electrical behavior and pore accumulation in a multicellular model for conventional
100
N. Matsuki et al. / Cancer Letters 269 (2008) 93–100
and supra-electroporation, Biochem. Biophys. Res. Commun. 349 (2006) 643–653. [31] F. Bovanovic, D.M. Bootman, J.M. Berridge, Elementary [Ca++]i signals generated by electroporation functionally mimic those evoked by hormonal stimulation, FASEB J. 13 (1999) 365–376. [32] J.S. Beebe, M.P. Fox, J.L. Rec, Nanosecond, high-intensity pulsed electric fields induce apoptosis in human cells, FASEB J. 17 (2003) 1493–1495. [33] J.S. Beebe, F.P. Blackmore, J. White, Nanosecond pulsed electric fields modulate cell function through intracellular signal transduction mechanisms, Physiol. Meas. 25 (2004) 1077–1093.
[34] P.T. Verner, Y. Sun, L. Marc, Nanoelectropulse-induced phosphatidylserine translocation, Biophys. J. 86 (2004) 4040–4048. [35] P.T. Vennier, Y. Sun, L. Marc, Nanosecond pulsed electric fields perturb membrane phospholipids in T lymphoblast, FEBS Lett. 572 (2004) 103–108. [36] A.J. White, F.P. Blackmore, H.K. Schoenbach, Stimulation of capacitative calcium entry in HL-60 cells by nanosecond pulsed electric field, J. Biol. Chem. 279 (2004) 22964–22972. [37] Y. Sun, P.T. Vernier, M. Behrend, Fluorescence microscopy imaging of electroperturbation in mammalian cells, J. Biomed. Optics 11 (2006) 024010.
Cancer Letters 269 (2008) 93–100 www.elsevier.com/locate/canlet
Low voltage pulses can induce apoptosis Noriaki Matsuki a,b,*, Takuji Ishikawa a, Yousuke Imai a, Takami Yamaguchi a a
Department of Bioengineering and Robotics, Tohoku University, 6-6-01, Aoba, Aramaki, Aoba-ku, 980-8579 Sendai, Japan b New Industry Creation Hatchery Center, Tohoku University, 6-6-10, Aoba, Aramaki, Aoba-ku, 980-8579 Sendai, Japan Received 21 January 2008; received in revised form 14 April 2008; accepted 15 April 2008
Abstract Electroporation is used for gene transfection, drug delivery, and cell fusion. While studies have shown that high voltage electroporation induces apoptosis in vitro, a strong electric field can lower cell survival rates. As there are no published reports which have examined apoptotic properties associate with low voltage electric charges, we demonstrated for the first time that consecutive low voltage pulses with a voltage lower than the membrane breakdown threshold of human cells can increase the membrane potential to the threshold required to induce electroporation. This led to apoptosis through caspase pathways. Moreover, necrotic cell damage was less than that caused by high voltage pulses. Therefore, low voltage electroporation can be a suitable anticancer method. Ó 2008 Elsevier Ireland Ltd. All rights reserved. Keywords: Apoptosis; Caspase; Electroporation; Low voltage pulse
1. Introduction Electroporation has been commonly used in a variety of in vitro [1–3] and in vivo [4–7] biotechnical applications. The applied electric field generates large depolarizing and hyperpolarizing transmembrane potentials across the cell. Electrical membrane breakdown occurs when the membrane potential reaches a critical value, resulting in the formation of pores. The number and size of the pores are determined by the pulse voltage, pulse
*
Corresponding author. Address: Department of Bioengineering and Robotics, Tohoku University, 6-6-01, Aoba, Aramaki, Aoba-ku, 980-8579 Sendai, Japan. Tel.: +81 22 714 8514; fax: +81 22 795 6959. E-mail address: [email protected] (N. Matsuki).
duration, waveform, pH, and the ionic strength of the suspending medium [2,7–12]. If the electric field is relatively weak, the pores shrink and disappear once the pulse is removed, and the electrical membrane breakdown can thus be reversed. Conversely, when a strong electric field is applied, the pores continue to exist long after the pulse, leading to irreversible damage of the cell membrane [13– 16]. During the effective pore open time, solutes added to the extracellular medium that are normally unable to permeate the membrane will be able to enter cell via the pores [16]. Cell death occurs by either necrosis or apoptosis [17]. Necrotic cells are characterized by irreversible plasma membrane rupturing as a result of severe cell damage and dysfunction, which in turn induces inflammatory reactions. In contrast, the membrane integrity of an apoptotic cell is maintained, and the
0304-3835/$ - see front matter Ó 2008 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.canlet.2008.04.019
94
N. Matsuki et al. / Cancer Letters 269 (2008) 93–100
cell is removed by phagocytosis without inducing an inflammatory response. Therefore, apoptosis is considered to be the most suitable method for novel targeted anticancer therapy [18]. Apoptosis is characterized by morphological and biochemical alterations of a cell including the externalization of phosphatidylserine on the plasma membrane, membrane blebbing, chromatin condensation, DNA fragmentation, and the degradation of intracellular proteins such as poly-ADP-ribose polymerase (PARP), lamine, and others. It involves the concerted action of many enzymatic steps including caspase activation [17,19]. Apoptosis signals can arise from the plasma membrane [17] or intracellular structures [20,21] such as the nucleus [17], mitochondria [22], and endoplasmic reticulum [23]. These signals activate pathways that converge to activate caspases [24,25]. In the past, several studies have reported that high electric voltage electroporation (>4.5 kV/ cm) can induce mild apoptosis as well as severe necrosis [26,27]. However, such a strong electric field reduces cell survival through severe cell explosion and causes undesirable side effects such as burn due to Joule heating and severe inflammatory reaction due to necrosis. On the other hand, electroporation with relatively low voltage electric field (0.2 kV/cm) demonstrated successful gene transfection in vivo [28]. Therefore, apoptosis induced by a mild electric field would be a favorable alternative to high voltage electroporation, and it would also be suitable for targeted anticancer treatments. To date, no published studies have examined the apoptotic capabilities of low voltage electric fields. We thus investigate whether low voltage and long duration pulses can induce apoptosis and examine the role of caspase pathways in the response. 2. Materials and methods 2.1. Cell culture The SCC 9 (human squamous cell carcinoma of the tongue) cell line was obtained from and cultured as recommended by the American Type Culture Collection (Rockville, MD, USA). Cells were cultured in RPMI 1640 (Irvine Scientific, Santa Ana, CA, USA) media containing 10% heat-incubated fetal bovine serum (FBS; Gibco, Grand Island, NY, USA), 50 U/ml penicillin (Gibco), and 50 lg/ml streptomycin (Gibco) at 37 °C in a 95% humidified, 5% carbon dioxide atmosphere.
2.2. Electroporation Cells were collected by trypsin treatment and centrifuged for 5 min at 300g, washed with phosphate buffer saline (PBS), and resuspended in PBS or RPMI 1640 at a concentration of 2 106 cells/ml. The cell suspensions (400 ll) were transferred to parallel aluminium-plated electroporation/fusion chambers with 2 mm gaps. Low voltage long duration (7.5 V/mm, 100 ms) and high voltage short duration (1.5 kV/mm, 0.05 ms) square wave pulses were applied using a Gene Pulser II (Bio-Rad, Hercules, CA, USA). 2.3. Detection of membrane breakdown (7-AAD uptake) Membrane breakdown was assessed by staining the cells with 10 ll 7-amino-actinomycin D (7-AAD; Becton–Dickinson, McKinley, MN, USA) and flow cytometric analysis. Cells were washed, resuspended at a concentration of 2 106 cells/ml in PBS, and stained with 10 ll 7-AAD. Following exposure to electrical pulses, the cells were immediately analyzed by flow cytometry according to the manufacturer’s protocol. 2.4. Viability assay For the viability analysis, the cells were washed and resuspended at a concentration of 2 106 cells/ml with PBS. After 2 h of exposure to electric pulses, the cells were stained with 10 ll 7-AAD and analyzed by flow cytometry. 2.5. Apoptosis assay A flow cytometry apoptosis detection kit (Becton– Dickinson) was used to identify apoptotic and necrotic cells according to the manufacturer’s protocol. Briefly, cells (3 105 cells/ml) were double-stained with 10 ll fluorescein isothiocyanate-labeled-Annexin V (R&D Systems, Minneapolis, MN, USA) to detect phosphatidylserine expression during early apoptotic phases, and 10 ll 7AAD to exclude late apoptotic and necrotic cells. Samples were analyzed using an EPICS XL-MCL cytometer (Beckman Coulter, Fullerton, CA, USA). The number of positive cells was determined in the annexin-positive fraction of the cell suspension by using an arbitrary threshold setting, which allows no more than 5% positive counts in the negative control. Cells positive for annexin only were defined as apoptotic, whereas those positive for both annexin and 7-AAD were defined as necrotic/late apoptotic. 2.6. CPP 32 activity (caspase-3-like enzyme activity) Caspase activity was measured as described previously [24]. Briefly, cells were washed twice with PBS, and the pellets were lysed in PBS containing 0.2% Triton X-100
N. Matsuki et al. / Cancer Letters 269 (2008) 93–100
on ice for 10 min and centrifuged at 12,000 rpm for 5 min. The cell extracts were incubated with 10 lM Ac-DEVDMCA (Peptide Institute, Osaka, Japan) as a fluorescent substrate in the incubation buffer [50 lM Tris–HCl (pH 7.5), 1 mM DDT] for 30 min at 37 °C. The fluorescence intensity of the substrate cleaved by caspase-3 was measured at 380 nm for excitation and 460 nm for emission by a spectrofluorometer.
95
10
5
2.7. Statistical analysis 0
Statistical analysis was performed using StatView 5.0 software (SAS Institute Inc., Cary, NC, USA). Comparisons of means were conducted using analysis of variance (ANOVA) with P < 0.05 considered to be statistically significant.
To determine the membrane breakdown threshold voltage, cells in PBS containing 7-AAD were exposed to a square wave with a 100 ms duration, single electric pulses of different voltages (0–50 V), and to a single square wave, 3000 V, 0.05 ms duration pulse, then they were analyzed using flow cytometry. The cells exhibited 5.55 ± 1.51%, 5.58 ± 1.42%, 5.85 ± 1.55%, 6.90 ± 1.71%, and 10.30 ± 1.31% 7-AAD uptake for control, 20, 30, 50, and 3000 V treatments, respectively. Therefore, the membrane breakdown threshold of SCC 9 cells was determined to be between 30 and 50 V (Fig. 1a). To determine whether consecutive electric pulses with a lower voltage than the membrane breakdown threshold could disrupt the structural integrity of the membrane, cell suspensions containing 7-AAD were exposed to 15 V, 100 ms duration, square wave electric pulses with an interval of 100 ms and immediately analyzed using flow cytometry. The control cells exhibited 5.55 ± 1.51% 7-AAD uptake, whereas treated cells showed 5.53 ± 1.46% uptake for a single pulse (P < 0.05 vs. control and single pulse), 6.90 ± 1.71% for 10 pulses (P < 0.05 vs. control, single and 10 pulses), 14.30 ± 3.09% for 100 pulses, and 19.52 ± 4.26% for 200 pulses (Fig. 1b). The 3000 V square wave, 0.05 ms duration, 5-pulse treatment resulted in 48.47 ± 14.30% 7-AAD-uptake (data not shown). Therefore, the 7-AAD-uptake of the cells increase as the pulse frequency is increased. This indicates that pulses with a lower voltage than the membrane breakdown threshold are able to break the membrane as the pulse frequency increases. 3.2. Viability It can take seconds, minutes, or hours for a membrane to recover (reseal) following electroporation [13–15]. Therefore, to examine cell viability, cells were stained with
30 V
50 V
3000 V
25
7 - AAD uptake (%)
3.1. Membrane breakdown
20 V
Pulse voltage (V)
20 15 10 5 0 control
1
10
100
200
Pulse frequency (Times) Fig. 1. The 7-AAD uptake of cells after the application of electric pulses. (a) Cells cultured in the presence of 7-AAD were exposed to single electric pulses of different voltages and analyzed using flow cytometry. (b) Cells cultured in the presence of 7-AAD were exposed to consecutive 15 V, 100 ms pulses of different frequencies and analyzed using flow cytometry. Data are shown as means ± SEM of four separate experiments.
100
Viability ( %)
3. Results
control
75
50
25
0 control
15 V
3000 V
Electric pulse application Fig. 2. Cell viability after the application of electric pulses. Cells were exposed to 200 low (15 V, 100 ms) or five high (3000 V, 0.05 ms) voltage electric pulses. The 7-AAD was added 2 h after the administration of the electric pulses. Viability was analyzed using flow cytometry. Data are shown as means ± SEM of four separate experiments.
N. Matsuki et al. / Cancer Letters 269 (2008) 93–100
10
4
10
3
10
2
10
1
10
0
control
15 V
104
103
7-AAD
7-AAD
96
102
101
100 10
0
10
1
10
2
10
3
10
4
100
101
10
3
10
2
10
1
10
0
102
103
104
10 3
10 4
Annexin
3000 V
cisplatin
104
103
7-AAD
4
102
101
100 10
0
10
1
10
2
10
3
10
4
10 0
Annexin
10 1
10 2
Annexin
15
Apoptosis rate ( %)
10
10
5
0 control
15 V
3000 V
cisplatin
Electric pulse / Cisplatin application 25
Necrosis rate ( %)
7-AAD
Annexin
20 15 10 5 0
control
15 V
3000 V
cisplatin
Electric pulse / Cisplatin application
7-AAD 2 h following the application of electric pulses and analyzed by flow cytometry. The control cells showed 91.8 ± 1.6% viability. Cells exposed to 15 V pulses showed 82.3 ± 4.5% viability, whereas 65.1 ± 3.0% of those treated with 3000 V survived (P < 0.01 vs. control and P < 0.02 vs. 15 V; Fig. 2). These data indicate that high voltage pulses severely damage cells. 3.3. Induction of apoptosis and cell death To determine whether low voltage pulses can induce apoptosis, cells were exposed to 200 15 V, 100 ms, square wave pulses and analyzed 48 h later using flow cytometry with an apoptosis detection kit. Cells treated with cisplatin (3.0 lg/ml) for 48 h were used as positive controls for apoptosis. Fig. 3a shows the flow cytometric plots of annexin and 7-AAD staining. Cells exposed to electrical pulses shifted to the right and upward compared to control cells. These results indicate that electrical pulses can induce apoptosis and necrosis. The apoptotic rates of the cells are shown in Fig. 3b. On average, apoptosis occurred in 3.0 ± 0.9% of the control cells, 6.0 ± 0.8% of the cells exposed to 15 V (P = 0.0514 vs. control), 13.2 ± 1.7% of the cells exposed to 3000 V (P < 0.01 vs. control and 15 V), and 6.7 ± 1.5% of the cisplatin-treated cells (P < 0.05 vs. 3000 V). Fig. 3c shows necrosis rates of the cells exposed to electrical pulses. On average, cell death occurred in 3.7 ± 1.2% of the control cells, 6.2 ± 1.4% of cells exposed to 15 V, 16.9 ± 5.2% of cells exposed to 3000 V (P < 0.05 vs. control), and 8.9 ± 0.8% of cisplatin-treated cells (P < 0.02 vs. control). Together, these data indicate that high voltage pulses were capable of inducing significant cell damage by necrosis as well as apoptosis. In contrast, low voltage pulses resulted in fewer damaged (necrotic) cells while still inducing apoptosis. 3.4. CPP 32 (caspase-3-like enzyme) activity To determine whether caspase pathways were involved in apoptosis induced by electric pulses, cells were exposed to 200 square wave pulses at 15 V for 100 ms each. CPP32 activity was measured 24 h following the application of electric pulses using synthetic peptide Ac-DEVD-MCA as a substrate. Cells treated with cisplatin (3.0 lg/ml) for 24 h were used as positive controls for activated caspase.
CPP 32 activity ( Fluorescence Intensity)
N. Matsuki et al. / Cancer Letters 269 (2008) 93–100
97
800
600
400
200
0 control
15 V
3000 V
cisplatin
Electric pulse / Cisplatin application Fig. 4. CPP32 activity after the administration of electric pulses. CPP32 activity was measured using the synthetic peptide AcDEVD-MCA as a substrate 24 h after the application of electric pulses. Data are shown as fluorescence intensity (FI) and means ± SEM of four separate experiments, each performed in duplicate.
Fig. 4 illustrates the CPP32 activities after the application of electric pulses. The control cells showed an average of 161.6 ± 31.9 fluorescence intensity (FI). Cells exposed to electrical pulses showed a significant increase of caspase activities. Specifically, 15 V produced 305 ± 25.8 FI (P < 0.01 vs. control), 3000 V induced 806.8 ± 35.4 FI (P < 0.01 vs. control and 15 V), and cisplatin-treated cells exhibited 604.6 ± 78.8 FI (P < 0.01 vs. control and 15 V; P < 0.02 vs. 3000 V). This indicates that caspase pathways play a significant role in electroporation-induced apoptosis.
4. Discussion In the past, several publications reported that high voltage electroporation could induce apoptosis in vitro [26,27]. However, applying such strong electric fields tend to severely diminish cell survival rates and induce undesired side effects as a result of Joule heating. We have demonstrated that mild electric fields are capable of inducing apoptosis while minimizing the undesirable side effects that accompany necrosis. Therefore, this shows great potential as an anticancer treatment method.
3 Fig. 3. Induction of apoptosis and cell death following the administration of electric pulses. Cells were exposed to 200 low (15 V, 100 ms) or five high (3000 V, 0.05 ms) voltage electric pulses and analyzed using a flow cytometry with annexin and 7-AAD staining carried out 48 h later. Cells treated with cisplatin (3.0 lg/ml) for 48 h were used as positive controls for apoptosis. (a) Flow cytometry plot of annexin and 7-AAD staining. Annexin () and 7-AAD () cells were considered intact; annexin (+) and 7-AAD () cells were considered apoptotic; 7-AAD (+) cells were considered necrotic. (b) Apoptosis rate. (c) Necrosis rate. Data are shown as means ± SEM of four separate experiments.
98
N. Matsuki et al. / Cancer Letters 269 (2008) 93–100
Given the results of the 7-AAD uptake experiments (Fig. 1a and b), the membrane breakdown threshold of SCC 9 cells was expected to be between 30 and 50 V. However, this is dependent on the membrane composition and the pulse duration as well as other stresses [2,14,16]. Therefore, we chose 15 V, less than a half of the expected membrane breakdown threshold of SCC 9 cells, for the low voltage electrical stimulation. The transmembrane potential (Vm) is theoretically given as follows: V m ¼ 1:5E a cos h where E is the applied electric field strength (7.5 V/mm), a is the cell radius (10 lm), and h is the position angle relative to the electrical field (cos h 5 1) (Fig. 5) [11]. According to this formula, Vm is expected to be approximately 0.11 V. This is much less than the threshold at which typical mammalian cells experience membrane breakdown (electroporation), which is reported to be between 0.2 and 1.5 V [14]. In our experiment using the single pulse application, no membrane breakdown could be detected at this voltage. However, it is possible that micropores smaller than the 7-AAD molecules were generated. This would still block 7-AAD from entering the cell and they would thereby remain undetected [29]. Our data indicated that there is an increase in the membrane permeability as the pulse frequency increases. Ryttsen et al. [10] reported that the transmembrane potential increased during the first 50 ms, reached the maximum state where it stabilized for up to 1 ms after the application of the elec-
Cell
Vm a Θ
π
0
E Fig. 5. Schematic of a spherical cell with radius a in an electric field. The transmembrane potential Vm is theoretically given as 1.5 E a cos h. The electric field E is oriented such that the depolarized pole is at h = 0 and the hyperpolarized pole is at h = p.
trical pulse. Gowrishankar et al. reported that longer pulse can induce more electroporation [30]. Since we have applied 100-ms pulses with an interval of 100 ms, it is possible that the transmembrane potential could accumulate with each pulse, eventually reaching the threshold and resulting in electroporation. The net percentage (sample-control value) of electroporated cells, as defined by the 7-AAD uptake immediately following the application of electric pulses, was 14% for cells exposed to 15 V and 43% for cells exposed to 3000 V. Similarly, the net percentage of apoptosis/necrosis as defined by the percentage of cells stained with annexin/7AAD 2 days following the application of electric pulses, was higher in cells exposed to high voltage pulses than those exposed to low voltage pulses (10.2% and 13.2% vs. 3% and 2.5% for high and low voltages, respectively). Therefore, apoptosis was induced in 21.4% of the electroporated cells using low voltage pulses and 23.7% of the electroporated cells using high voltage pulses. Interestingly, the ratio between apoptosis and necrosis in the electroporated cells was higher in the low voltage group (1.2 vs. 0.77 for low and high voltages, respectively). Therefore, we have demonstrated that low voltage pulses were capable of inducing less necrosis (i.e., fewer harmful side effects), and more apoptosis than high voltage treatments. Caspase activity, a marker for apoptosis via the caspase pathways, was observed to increase with increasing voltage. These results are in accordance with apoptosis levels detected by annexin. Hofmann et al. [26] also reported the occurrence of caspase activation occurred in high voltage-induced apoptosis. To date, the mechanisms and pathways of apoptosis in electroporated cells remain unclear. Several reports [26,27] have suggested that electroporation with high voltage electrical pulse charges induces apoptosis by changing the membrane integrity via the influx of extracellular substances including ions such as Ca++, Na+, K+, and Cl [26]. This is thought to break the membranes of organelles such as mitochondria and the endoplasmic reticulum, and activate intracellular signaling [31]. Interestingly, very high voltage (MeV/m), ultrashort (nanosecond) pulses can also induce apoptosis without membrane breakdown [32–36] and bring DNA fragmentation [37]. However, the formation of undetectable micropores may take place. Apoptosis induced by electrical charges could be based on changing the membrane integrity, the pore forma-
N. Matsuki et al. / Cancer Letters 269 (2008) 93–100
tion, and the activation of caspase enzymes. Prausnitz et al. [12] reported that transport caused by electroporation occurs predominantly by electrophoresis and electroosmosis during a pulse, and by diffusion after pulse. Therefore, it is possible that ions such as Ca++ could pass through micropores and activate caspase pathways. In summary, consecutive pulses of a voltage less than the membrane breakdown threshold can increase the membrane potential to the threshold, disrupt the membrane integrity, cause membrane permeabilization through pores, lead to the influx of extracellular substances including ions, activate caspase pathways, and induce apoptosis. Furthermore, low voltage electroporation can induce apoptosis while mitigating cellular damage that typically accompanies high voltage electroporation such as that caused by necrosis. Therefore, low voltage electroporation can represent a useful anticancer method targeting apoptosis. Further studies are required to define the exact threshold of membrane breakdown and resolve the apoptosis signaling mechanisms involved in electroporation. Acknowledgments This study was supported by the Japanese Society for the Promotion of Science and the Austrian Academic Exchange Service. References [1] T.Y. Tsong, Electroporation of cell membranes, Biophys. J. 60 (1991) 297–306. [2] J.C. Weaver, Y.A. Chizmadzhev, Theory of electroporation: a review, Bioelectrochem. Bioenerg. 41 (1996) 135–160. [3] J.C. Weaver, Electroporation of cell and tissues, IEE. Trans. Plasm. Sci. 28 (2000) 24–33. [4] B. Rubinsky, Irreversible electroporation in medicine, Technol. Cancer. Res. Treat. 6 (2007) 255–259. [5] J.P. Cantella, M.M. Black, M.D. Bonnichsen, Tissue electroporation: quantification and analysis of heterogenous transport in multicellular environments, Biophys. J. 86 (2004) 3260–3268. [6] G. Sersa, D. Miklavcic, M. Cemazar, Electrochemotherapy in treatment of tumours, Eur. J. Surg. Oncol. 34 (2008) 232–240. [7] J.A. Kim, K. Cho, M.S. Shin, A novel electroporation method using a capillary and wire-type electrode, Biosens. Bioelectron. 23 (2008) 1353–1360. [8] J. Deng, H.K. Schoenbach, S. Buescher, The effects of intense submicrosecond electrical pulses on cells, Biophys. J. 84 (2003) 2709–2714. [9] J.P. Cantella, F.J. Karr, A.J. Petros, Quantitative study of electroporation-mediated molecular uptake and cell viability, Biophys. J. 80 (2001) 755–764.
99
[10] F. Ryttsen, C. Farre, C. Brennan, Characterization of single cell electroporation by using patch–clamp and fluorescence microscopy, Biophys. J. 79 (2000) 1993–2001. [11] K. DeBruin, W. Krassowska, Modeling electroporation in a single cell. I. Effects of field strength and rest potential, Biophys. J. 77 (1999) 1213–1224. [12] M.R. Prausnitz, J.D. Corbett, J.A. Gimm, Millisecond measurement of transport during and after an electroporation pulse, Biophys. J. 68 (1995) 1864–1870. [13] G. Saulis, Pore disappearance in a cell after electroporation: theoretical simulation and comparison with experiments, Biophys. J. 73 (1997) 1299–1309. [14] R.T. Gowrishankar, U. Pliquett, R. Lee, Dynamics of membrane sealing in transient electropermeabilization of skeletal muscle membranes, Ann. NY Acad. Sci. 888 (1999) 196–210. [15] F. Hilpert, A. Heiser, W. Wieckhorst, The impact of electrical charge on the viability and physiology of dendritic cells, Scand. J. Immunol. 62 (2005) 399–406. [16] M. Tarek, Membrane electroporation: a molecular dynamics simulation, Biophys. J. 88 (2005) 4045–4053. [17] M.O. Hengarther, The biochemistry of apoptosis, Nature 407 (2000) 770–776. [18] A. Russo, M. Terrasi, V. Agnese, Apoptosis: a relevant tool for anticancer therapy, Ann. Oncol. 17 (2006) vii115–123. [19] J. Liu, A. Lin, Role of JNK activation in apoptosis: a double-edged sword, Cell Res. 15 (2005) 36–42. [20] M.H. Beere, The stress of dying: the role of heat shock proteins in the regulation of apoptosis, J. Cell Sci. 117 (2004) 2641–2651. [21] M.H. Beere, Death versus survival: functional interaction between the apoptotic and stress-inducible heat shock protein pathways, J. Clin. Invest. 115 (2005) 2633–2639. [22] K. Maaser, P.A. Sutter, H. Scheruebl, Mechanisms of mitochondrial apoptosis induced by peripheral benzodiazepine receptor ligands in human colorectal cancer cells, Biochem. Biophys. Res. Commun. 332 (2005) 646–652. [23] J.S. Kim, M.K. Park, N. Kim, Doxorubicin prevents endoplasmic reticulum stress-induced apoptosis, Biochem. Biophys. Res. Commun. 339 (2005) 463–468. [24] Y. Fukamachi, Y. Karasaki, T. Sugiura, Zinc suppresses apoptosis of U937 cells induced by hydrogen peroxide through an increase of the Bcl-2/Bax ratio, Biochem. Biophys. Res. Commun. 246 (1998) 364–369. [25] R.C. Taylor, S.P. Cullen, S.J. Martin, Apoptosis: controlled demolition at the cellular level, Nature Rev. Mol. Cell Biol. 9 (2008) 231–241. [26] F. Hofmann, H. Ohnimus, C. Scheller, Electric field pulses can induce apoptosis, J. Membr. Biol. 169 (1999) 103–109. [27] J. Pinero, M. Lopez-Baena, T. Ortiz, Apoptotic and necrotic cell death are both induced by electroporation in HL60 human promyeloid leukemia cells, Apoptosis 2 (1997) 330– 336. [28] D. Machado-Aranda, Y. Adir, J.L. Yong, Gene transfer of Na+, K+-ATPase b1 subunit using electroporation increases lung liquid clearance, Am. J. Respir. Crit. Care Med. 171 (2005) 204–211. [29] P.T. Vernier, Y. Sun, Y. Marcu, Calcium bursts induced by nanosecond electric pulses, Biochem. Biophys. Res. Commun. 310 (2003) 286–295. [30] T.R. Gowrishankar, J.C. Weaver, Electrical behavior and pore accumulation in a multicellular model for conventional
100
N. Matsuki et al. / Cancer Letters 269 (2008) 93–100
and supra-electroporation, Biochem. Biophys. Res. Commun. 349 (2006) 643–653. [31] F. Bovanovic, D.M. Bootman, J.M. Berridge, Elementary [Ca++]i signals generated by electroporation functionally mimic those evoked by hormonal stimulation, FASEB J. 13 (1999) 365–376. [32] J.S. Beebe, M.P. Fox, J.L. Rec, Nanosecond, high-intensity pulsed electric fields induce apoptosis in human cells, FASEB J. 17 (2003) 1493–1495. [33] J.S. Beebe, F.P. Blackmore, J. White, Nanosecond pulsed electric fields modulate cell function through intracellular signal transduction mechanisms, Physiol. Meas. 25 (2004) 1077–1093.
[34] P.T. Verner, Y. Sun, L. Marc, Nanoelectropulse-induced phosphatidylserine translocation, Biophys. J. 86 (2004) 4040–4048. [35] P.T. Vennier, Y. Sun, L. Marc, Nanosecond pulsed electric fields perturb membrane phospholipids in T lymphoblast, FEBS Lett. 572 (2004) 103–108. [36] A.J. White, F.P. Blackmore, H.K. Schoenbach, Stimulation of capacitative calcium entry in HL-60 cells by nanosecond pulsed electric field, J. Biol. Chem. 279 (2004) 22964–22972. [37] Y. Sun, P.T. Vernier, M. Behrend, Fluorescence microscopy imaging of electroperturbation in mammalian cells, J. Biomed. Optics 11 (2006) 024010.