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Sonomagnetic Stimulation of Live Cells: Electrophysiologic, Biochemical and Behavioral Responses
ARTICLE IN PRESS Ultrasound in Med. & Biol., Vol. 00, No. 00, pp. 114, 2019 Copyright © 2019 World Federation for Ultrasound in Medicine & Biology. All rights reserved. Printed in the USA. All rights reserved. 0301-5629/$ - see front matter
https://doi.org/10.1016/j.ultrasmedbio.2019.07.009
Original Contribution SONOMAGNETIC STIMULATION OF LIVE CELLS: ELECTROPHYSIOLOGIC, BIOCHEMICAL AND BEHAVIORAL RESPONSES AGEDPY T AXIN
HU,*,y YANCHENG WANG,*,y XIN CHEN,*,y and SIPING CHEN*,yTAGEDEN
* School of Biomedical Engineering, Health Science Center, Shenzhen University, Shenzhen, P.R. China; and y National-Regional Key Technology Engineering Laboratory for Medical Ultrasound, Shenzhen, P.R. China (Received 19 November 2018; revised 1 July 2019; in final from 6 July 2019)
Abstract—Various physical methods have been developed to modulate the electrophysiologic properties of cells and their biochemical signaling pathways. In this study, we propose a sonomagnetic method using pulsed ultrasound (1.1 MHz frequency, 1.1 or 2.2 MPa pressure, 50 cycles per pulse and 500 Hz pulse repetition frequency) and a static magnetic field (680 mT) to stimulate live cells. We found that sonomagnetic stimulation promoted the cell and mitochondrial membrane potentials to more hyperpolarized states. The degree of cell membrane hyperpolarization was cell-type dependent. Furthermore, we found that the intracellular concentrations of Ca2+ ions, reactive oxygen species and nitric oxide were substantially increased after sonomagnetic stimulation, and a small decrease in intracellular pH was also observed. Lastly, we found that the daily sonomagnetic stimulation for 3 d inhibited the proliferation rate of neuro-2a cancer cells by 48.64%. Our work demonstrates that sonomagnetic stimulation can effectively perturb cell signaling and drive cancer cells into relatively quiescent states. (E-mail: [email protected]) © 2019 World Federation for Ultrasound in Medicine & Biology. All rights reserved. Key Words: Ultrasound, Magnetic field, Membrane potential, Biochemical signal.
triphosphate for metabolic activities. By contrast, it has been proposed that mitochondria with high DCm initiate programmed cell death (Heiskanen et al. 1999; Zorova et al. 2018). Therefore, the modulation of cellular and organelle membrane potentials is important for controlling cell functions. Considerable research has revealed the complex ways in which the cell and organelle membrane potentials affect cell behavior. Mechanistic studies have shown that the bioelectrical signals of membrane potentials are first transduced into biochemical signals to trigger subsequent intracellular signaling cascades. For example, changes in Vmem can be transduced into the cell via one of two pathways: (i) activation of voltage-gated ion channels and (ii) activation of voltage-sensitive phosphatase. In the first pathway, a hyperpolarized Vmem results in the influx of divalent Ca2+, which rapidly triggers the production of signaling molecules, such as calmodulin and nitric oxide (NO) to modulate cell behavior (Krasznai et al. 2000; Gilbert et al. 2017). In the second pathway, after membrane potential alternation, the activated phosphatase removes phosphate from the inositol rings of phosphatidylinositol messengers (e.g., phosphatidylinositol (3,4,5)-trisphosphate (PI[3,4,5]P3), and phosphatidylinositol 4,5-bisphosphate
INTRODUCTION A eukaryotic cell manages its autonomous life-sustaining processes via exquisite physical and chemical signals. One important cell signaling mechanism is the electrical modulation of voltage gradients generated across the cell and organelle membranes (McCaig et al. 2005). For example, the balance of ion currents produced by transmembrane channels and pumps results in a resting cell membrane potential (Vmem) ranging from 10 to 90 mV, depending on the specific cell type. An increase of this membrane potential (a hyperpolarized Vmem) can turn the cell into a relatively quiescent state with a slower proliferation rate, whereas a decrease of this membrane potential (a depolarized Vmem) makes the cell mitotically active and proliferate more quickly (Blackiston et al. 2009; Yang and Brackenbury 2013). In addition, mitochondria, the energy “power-plant” of cells, also have a transmembrane potential (DCm), ranging from 130 to 180 mV, which has an important role in various cell functions. Mitochondria with low values of DCm cannot generate sufficient adenosine Address correspondence to: Xin Chen, PhD, 3688 Nanhai Avenue, Shenzhen, Guangdong, P.R. China. E-mail: [email protected]
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(PI[4,5]P2)) to regulate cell motility and shape (Iwasaki et al. 2008). Mitochondrial DCm is also transduced into chemical signals. For example, a high DCm can accelerate the mitochondrial respiratory chain in an exponential manner to produce more reactive oxygen species (ROS). The oxidative stress induced by harmful levels of ROS (e.g., superoxide [O2 ] and hydroxyl radicals [OH ]) opens membrane permeability transition pores and releases cytochrome c to trigger apoptosis (Lee et al. 2001; Sena and Chandel 2012). Because the cell and organelle membrane potentials are important modulators of intracellular biochemical signals, various electrical stimulation methods have been developed to modulate membrane potentials. In in vitro studies, chemicals or drugs have been directly added to the extracellular medium (e.g., potassium ions for cell membrane depolarization and oligomycin for mitochondria hyperpolarization) (Giovannini et al. 2002; Sundelacruz et al. 2015). In in vivo studies, direct current stimulation has proven to be an effective approach; however, this method needs invasive implantation of electrodes (Perlmutter and Mink 2006). It has been shown that magnetic stimulation can affect cell membrane potentials, and transcranial magnetic stimulation (TMS) has been approved by the US Food and Drug Administration (FDA). The technical limitation of magnetic stimulation is its spatial resolution of several centimeters, which is too large for precise regional stimulation (Hallett 2007). Recently, ultrasound has been shown to be able to stimulate neuronal circuits (Legon et al. 2014). However, the underlying mechanism of ultrasound stimulation is still largely unknown. In this work, we propose a sonomagnetic method to evoke electrophysiologic, biochemical and behavioral responses of live cells. This sonomagnetic method is developed based on the following two effects: (i) the Debye effect, which shows that ions with different masses and charges in a physiologic solution would vibrate and separate from each other when driven by ultrasound energy (Debye 1933) and (ii) the Hall effect, which employs a static magnetic field to generate Lorentz forces in opposite directions to further separate the moving ions of different charges and to produce an electrical current (Wright and Van Der Beken 1972). Note that the feasibility of employing sonomagnetic stimulation for generating Lorentz currents in physiologic solution has been proven. Montalibet et al. (2001) measured a 320 nA Lorentz current evoked by 2 MPa ultrasound (500 kHz frequency) combined with a 350 mT strength magnetic field in NaCl solution with a conductivity of 1.6 S/m. Using a 420 kPa ultrasound (1.1 MHz frequency) and a 2 T strength magnetic field, Rekhi and Arbabian (2017) measured a 3.1 mA Lorentz current in
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1 M KCl solution (11.7 S/m conductivity). The sonomagnetically-stimulated Lorentz current was also confirmed in issue phantoms and ex vivo tissue samples (Grasland-Mongrain et al. 2013; Kunyansky et al. 2017). However, the question of whether the sonomagnetic method could be used for electrical stimulation of the electrophysiologic state of live cells and the production of biochemical signaling species has not been answered. Therefore, we established a sonomagnetic experiment platform for live cell stimulation. Cell and mitochondrial membrane potentials were fluorescently measured to examine the electrophysiologic responses of cells. Biochemical signaling species, including Ca2+, NO, H+ and ROS, were also fluorescently measured to examine the cells’ biochemical responses. Finally, the behavioral responses of the cells after sonomagnetic stimulation were monitored. Our work aims to provide experimental evidence for the feasibility and effectiveness of sonomagnetic stimulation of live cells. THEORY As demonstrated in Figure 1a, the propagation of ultrasound (longitudinal pressure wave along the z-axis) in an electrolyte solution results in the oscillation of the fluid and ions. When a static magnetic field is applied in the direction (y-axis) perpendicular to the direction of ultrasound propagation, Lorentz forces will be generated on the oscillating ions. The magnitude of the Lorentz force of an ion with charge q is given by FLorentz ¼ qvB
ð1Þ
where v is the velocity of the ion and B is the magnetic field strength. Driven by the generated Lorentz forces, positive and negative ions will move in opposite directions to produce an electrical current. Assuming the positive and negative ions have charges of q and q, respectively, the electrical current density, j, can be calculated as: j ¼ ðnþ mþ þ nmÞFLorentz
ð2Þ
where n+ and n are the ion concentrations, and m+ and m are the ion mobilities. Using eqn (1) and eqn (2), j can be written as: j ¼ qðnþ mþ þ nmÞvB
ð3Þ
Because the electrical conductivity of the electrolyte solution is given by: s ¼ qðnþ mþ þ nmÞ
ð4Þ
j can be expressed as: j ¼ svB
ð5Þ
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Fig. 1. Experimental setup and Lorentz current detection. (a) Graphical illustration of sonomagnetic stimulation theory. (b) Generation of the ultrasonic and magnetic fields for live cell stimulation. (c) The ultrasound pressure map measured by a needle hydrophone. (d) The measured magnetic field strength (left) and the calculated field strength gradients along the x-axis (middle) and the y-axis (right). (e) Immersible chamber with live cells grown on its bottom. (f) Immersible chamber with a 0.7 mm width copper wire placed on its bottom and the Lorentz current detection system. (g) Lorentz current elicited in the copper wire by 1.1 MPa pressure ultrasound (20 cycles per pulse and 500 Hz pulse repetition frequency) and 680 mT magnetic field strength. (h) The spectrum of the Lorentz current generated by sonomagnetic stimulation in the copper wire. DPO = oscilloscope; f = frequency; FLorentz = Lorentz Force; NdFeB = Neodymium Iron Boron; p(t,z) = ultrasound pressure; PXPA6-40 dB = 40 dB differential amplifier; q = electrical charge; t = time; u = electrical voltage; v(t,z) = ion oscillation speed.
The ion oscillation speed, v, can be calculated from the classical ultrasound propagation equation: r
@vðt; zÞ @Pðt; zÞ þ ¼0 @t @z
ð6Þ
where r is the mass density of the solution and P(t, z) is the ultrasound pressure, which varies in time, t, and with distance, z. Therefore, the ion oscillation speed, v(t, z), can be calculated by integrating the above equation with respect to time: Zt 1 @Pðt; zÞ vðt; zÞ ¼ dt ð7Þ r @z 1
Substituting eqn (7) into eqn (5), we get: sB jðt; zÞ ¼ r
Zt 1
@Pðt; zÞ dt @z
ð8Þ
According to eqn (8), the generated electrical density can be modulated by three parameters: the solution conductivity, s, the magnetic field strength, B, and the ultrasound pressure, P. For example, when s and B are constant, increasing the ultrasound pressure increases the generated electrical current density, j. Comparing the parameters used in this study (frequency of 1.1 MHz, conductivity of 1.6 S/m and magnetic field strength of
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680 mT) with those employed by Rekhi and Arbabian (2017), it can be inferred that the amplitude of the generated Lorentz current in the extracellular medium by an ultrasound wave of 2.2 MPa pressure is in the range of 0.75 mA. It is worth noting that the conductivity of the lipid membrane (5 £ 107 S/m) is much lower than that of physiologic solutions (0.21.6 S/m) (Pucihar et al. 2001). Therefore, voltage gradients are generated at cell and organelle membrane boundaries (Kunyansky et al. 2017). MATERIALS AND METHODS Experimental setup To produce ultrasound waves, an electrical signal was first edited in a function generator (DG1022, Agilent Technologies, Santa Clara, CA, USA), then amplified by a 50 dB broadband power amplifier (2100L, Electronics & Innovation, Rochester, NY, USA), and finally transformed into acoustic waves by an annular high-intensity focused transducer (H102, Sonic Concepts, Bothell, WA, USA) (Fig. 1b). The outer diameter of the transducer is 64 mm, and its inner diameter is 20 mm. The focal depth of the transducer is 51.74 mm. The cell samples were placed 61 cm from the transducer (9.26 mm from the transducer focus), where the ultrasound pressure distribution was relatively homogenous. The ultrasound pressure map generated in the cell sample region was measured using a needle hydrophone (HNR-0500, Onda Corp, Sunnyvale, CA, USA) (Fig. 1c). Pulsed ultrasound waves (1.1 MHz center frequency, 50 cycles per pulse and 500 Hz pulse repetition frequency) with two peak negative pressure settings (1.1 and 2.2 MPa) were used for electrochemical stimulation of live cells. To generate a static magnetic field, two Neodymium Iron Boron (NdFeB) magnets (cubic shape, 5 £ 5 £ 2.5 cm) were used. The gap between these two magnets was 13 mm. The measured magnetic field strength in the live cell region (indicated by the dashed circle in left image of Fig. 1d) was 680 mT. The magnetic field gradients calculated along the x-axis and the y-axis were also given in Figure 1d (the middle and the right image). Immersible cell chamber It is reported that when the cells were grown in culture wells for in vitro ultrasound experiments, standing waves would be generated in the culture medium to affect the cell stimulation results (Hensel et al. 2011). To minimize the influence of standing waves, Hensel et al. (2011) suggested that the air-medium interface in the wells should be avoided, and the wells should be submersed into the water tank to receive ultrasound
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exposure. Therefore, in this study, we designed an immersible cell chamber for sonomagnetic stimulation. The rectangular chamber (61 mm [L] £ 12.6 mm [W] £ 5 mm [H]) had a circular window area (8.6 mm diameter) for cell culture. In particular, both sides of the circular window were sealed by an 170 mm thick cover glass. Live cells could be seeded into the chamber through an injection channel of 1.5 mm diameter. During the experiment, the chamber was filled with degassed Hank’s balanced salt solution (1672547, Invitrogen, Carlsbad, CA, USA) with 10 mM N-2-hydroxyethylpiperazine-N-2-ethane sulfonic acid (HEPES) (15630080, Invitrogen). In this way, no air would be left in the chamber, and no air-water interface was formed. Furthermore, the injection channel of the chamber could be sealed by sterilized polystyrene membranes. The chamber could be submersed in the degassed water to further minimize ultrasound reflection and standing wave formation. Lorentz current confirmation To confirm the generation of Lorentz current in our experimental setup, a 0.7 mm width copper wire was inserted into the immersible chamber (Fig. 1f). The chamber was placed between the two magnets to receive the ultrasound exposure. The electrical signal generated at the two ends of the copper wire was amplified by a 40 dB differential amplifier (PXPA6-40, Pengxiang, Changsha, Hunan, China) and recorded by an oscilloscope (DPO 5054, Tektronix, Beaverton, OR, USA). As shown in Figure 1g, the averaged amplitude of the electrical signal elicited in copper wire by the 1.1 MPa pressure ultrasound wave (20 cycles per pulse and 500 Hz pulse repetition frequency) and 680 mT strength magnetic field was 24.3 mV. Note that, when the magnets were removed, the electrical signal disappeared. As shown in Figure 1h, the frequency of the Lorentz current was the same as the ultrasound wave, which was consistent with eqn (8) given in the Theory section. Cell culture In this study, we employed mouse neuroblastoma cell line (neuro-2a or N2a, ATCC CCL-131, Manassas, VA, USA) to study the cellular effects induced by sonomagnetic stimulation. The N2a cells were cultured in minimum essential medium (10370070, Invitrogen) supplemented with 10% fetal bovine serum (10099141, Invitrogen). When the cells reached a confluency of 70%, they were detached from the bottom of a 25 cm2 flask by trypsin- ethylenediaminetetraacetic acid (25200072, Invitrogen). Then, 5 £ 103 cells were seeded onto the glass bottom of the immersible cell chamber. After a 24 h incubation time at 37˚C and 5% CO2, the cells were fully attached to the glass bottom
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and could be used for the sonomagnetic stimulation experiments. In addition to N2a cells, we employed another six types of cells to examine the question of whether the cell membrane potential alternation induced by sonomagnetic stimulation was a cell-type dependent phenomenon. These six types of cells were categorized according to their morphology: (i) cells of neuroblast-like morphology, including mouse dendritic cell of DC2.4 and rat pheochromocytoma cell of PC12; (ii) cells of endothelial-like morphology, including mouse endothelioma cell of BEND3 and human cervical cancer cell of HeLa; and (iii) cells of fibroblast-like morphology, including human lung cell of MRC5 and mouse embryo cell of 3T3. Four types of cells (PC12, BEND3, HeLa and 3T3) were grown in Dulbecco’s modified Eagle medium (11965092, Invitrogen). The DC2.4 cells were grown in Roswell Park Memorial Institute 1640 medium (11875093, Invitrogen). MRC5 cells were grown in minimum essential medium. Note that, all of the media were supplemented with 10% fetal bovine serum for cell culture. To avoid the influence of phenol red on the fluorescence measurement, the cells were temporarily cultured in Hanks’ balanced salt solution (14025092, Invitrogen) supplemented with 20 mM HEPES during the examination of bioeffects induced by sonomagnetic stimulation. Membrane potential labeling To monitor changes in Vmem, a voltage-sensitive fluorescent probe, Bis-(1,3-Dibutylbarbituric Acid)Trimethine Oxonol (DiBAC4(3)) (B438, Invitrogen), was loaded into live cells at a final concentration of 200 nM. In general, the green fluorescence intensity of DiBAC4(3) increases with membrane depolarization and decreases with membrane hyperpolarization. To monitor changes in the mitochondrial DCm, a red fluorescent tetramethylrhodamine, ethyl ester (TMRE) probe (ENZ52309, Enzo Life Sciences, Farmingdale, NY, USA) was used to stain live cells at a final concentration of 100 nM. We note that TMRE loaded at this high concentration works in the quenching mode, which means mitochondrial hyperpolarization results in a relative decrease in fluorescence intensity of the dye and vice versa (Perry et al. 2011). Labeling of biochemical signaling species The fluorescent probe Fluo-4 AM (F14201, Invitrogen), at a final concentration of 2 mM, was used to monitor the cytoplasmic Ca2+ concentration. The intracellular NO level was monitored using the probe 4-Amino-5-Methylamino-2’,7’-Difluorofluorescein (DAF-FM) diacetate (D23844, Invitrogen) at a final concentration of 5 mM. The cytoplasmic pH was monitored using the probe pHrodo Red AM (P35372, Invitrogen) at a final concentration of
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5 mM. The intracellular ROS level was assessed using the probe 20 ,70 -Dichlorodihydrofluorescein diacetate (H2DCFDA) (D399, Invitrogen) at a final concentration of 3 mM. Fluorescent labeling of live cells was carried out according to the product protocols. Monitoring of cell proliferation To allow the cells to proliferate for 3 d without the contact-induced inhibition of division, N2a cells were seeded into the immersible chamber at relatively low density (1 £ 103 cells per chamber). The cells were then divided into four groups (a control group, an ultrasoundexposure group, a magnetic field group and a sonomagnetic stimulation group) and received daily physical treatment, respectively. To monitor the proliferation status of cells in different groups, the cells in the center of the immersible chamber were imaged before each treatment using an inverted microscope (Axio Observer A1, Carl Zeiss Meditec AG, Oberkochen, Germany) and a 63 £ objective. The number of the cells in the imaging view (135 £ 100 mm) was counted manually using the ImageJ software (v.1.52a, National Institutes of Health, Bethesda, MD, USA) and its plug-in (Cell Counter). Quantitative calculation Fluorescently labeled cells in the central area (diameter of 4 mm) of the immersible chamber were imaged using the inverted microscope (Axio Observer A1, Carl Zeiss Meditec AG). Microscope images were then imported into the image processing and analysis software ImageJ. The fluorescence intensity of each cell was calculated by manually outlining the cell boundary (white dashed line, Fig. 2a) with the aid of the Find Edges and Measure functions of ImageJ. The background fluorescence intensity was also calculated in the rectangular region of interest (10 mm [L] £ 12.6 mm [W]; yellow dashed line, Fig. 2a). We note that the background fluorescence intensity was subtracted from the cellular fluorescence intensity. To determine the statistical significance of the fluorescent intensities of the different experimental groups, the data were analyzed using GraphPad Prism software (v.5, GraphPad Software, San Diego, CA, USA) using the Friedman test (i.e., a nonparametric repeated measures analysis of variance). RESULTS Cell membrane hyperpolarization To calibrate the fluorescence responses of DiBAC4(3) probe in our system, we modulated the plasma membrane of N2a cells by increasing the extracellular K+ concentration, which linearly depolarizes the membrane potential (Yamada et al. 2001). We found that the green fluorescence intensity of DiBAC4(3)
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Fig. 2. Cell membrane hyperpolarization induced by sonomagnetic stimulation. (a) Microscope images showing the fluorescence intensity increase of DiBAC4(3)-loaded cells after extracellular K+ stimulation. (b) Quantitative calibration of the DiBAC4(3) fluorescence intensity measured under K+ stimulation at different concentrations. (c) Microscope images showing the fluorescence intensity decrease of DiBAC4(3)-loaded cells after sonomagnetic stimulation. (d) Quantitative analysis of the DiBAC4(3) fluorescence intensities measured after sham exposure (Ctrl), magnetic stimulation (M), ultrasonic stimulation (US) and sonomagnetic stimulation (US + M). (e) Quantitative analysis of the DiBAC4(3) fluorescence intensities measured after US and US + M at different pressures. (f) Quantitative analysis of the DiBAC4(3) fluorescence intensities measured after sonomagnetic stimulation for different exposure times. Data (mean with SEM) were calculated from three independent experiments (n = 150). The image contrast was increased by 10%. **p < 0.01; ***p < 0.001. Ctrl = sham exposure; DiBAC4(3) = Bis-(1,3-Dibutylbarbituric Acid)Trimethine Oxonol; F = Fluorescence intensity; K+ = potassium; M = magnetic stimulation; M + US = sonomagnetic stimulation; SEM = standard error of the mean; US = ultrasonic stimulation.
increased with the concentration of extracellular K+ (Fig. 2a). Quantitative analysis further showed the relationship between the concentration of K+ and DiBAC4(3) fluorescence intensity was almost linear, demonstrating the effectiveness of this voltage-sensitive probe in our experiments (Fig. 2b). We found that sonomagnetic stimulation resulted in a decrease in fluorescence intensity of the membraneloaded DiBAC4(3) probe, which was indicative of hyperpolarization of the membrane potential (Fig. 2c). Moreover, sonomagnetic stimulation at higher ultrasound pressure (2.2 MPa) induced a greater decrease of
DiBAC4(3) fluorescence intensity than that at lower ultrasound pressure (1.1 MPa). We also studied the changes in cell membrane potential induced by different physical stimuli, including magnetic, ultrasonic and sonomagnetic stimulation. As shown in Figure 2d, magnetic stimulation had little effect on the cell membrane potential. Ultrasonic stimulation induced a small-scale hyperpolarization of the cell membrane (4.17%). By contrast, sonomagnetic stimulation resulted in significant hyperpolarization of the cell membrane (47.89%), confirming the effectiveness of sonomagnetic stimulation in modulating the membrane potential.
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Consistent with eqn (8), according to which the electrical current density generated by sonomagnetic stimulation is positively correlated with ultrasound pressure, we found that ultrasound at higher pressure (2.2 MPa) evoked a significant higher degree of membrane hyperpolarization (57.92%) (Fig. 2e). Moreover, we further studied the cumulative effect of the sonomagnetic stimulation on membrane hyperpolarization from 010 min. We found that longer exposure time resulted in a greater hyperpolarization of cell membrane potential (Fig. 2f).
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Cell-type-dependent membrane hyperpolarization The membrane electrophysiology of different cell types varies greatly (e.g., resting membrane potential and ion channel type). Even within a single cell, the distribution of ion channels on cell membrane is heterogeneous (Schram 2002; Nusser 2009). Therefore, it is of great interest to explore membrane potential responses of different types of cells after sonomagnetic stimulation. In this work, which focused on stimulating the resting membrane potential, we used non-excitable cells.
Fig. 3. Cell-type-dependent cell membrane hyperpolarization. Sonomagnetic stimulation resulted in the decrease in DiBAC4(3) fluorescence intensity of neuroblast-like cells (a), endothelial-like cells (b) and fibroblast-like cells (c). (d, e) Quantitative analysis showing cell-type-dependent membrane potential responses observed under ultrasound pressures of 1.1 and 2.2 MPa. Data (mean with SEM) were calculated from three independent experiments (n = 150). The image contrast was increased by 10%. *p < 0.05. 3T3 = mouse embryo cells; BEND3 = mouse endothelioma cells; Ctrl = sham exposure; DC2.4 = mouse dendritic cells; DiBAC4(3) = Bis-(1,3-Dibutylbarbituric Acid) Trimethine Oxonol; Elike = endothelial-like; F = Fluorescence intensity; F-like = fibroblast-like; HELA = human cervical cancer cells; M = magnetic stimulation; MRC5 = human lung cells; M + US = sonomagnetic stimulation; N2a = neuro-2a; Nlike = neuroblast-like; PC12 = rat pheochromocytoma cells; SEM = standard error of the mean; US = ultrasonic stimulation.
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Fig. 4. Mitochondrial membrane hyperpolarization. (ac) Fluorescence images of TMRE-loaded cells after sham-exposure, sonomagnetic stimulation at 1.1 MPa ultrasound pressure and sonomagnetic stimulation at 2.2 MPa ultrasound pressure, respectively. (d) Quantitative analysis of the TMRE fluorescence intensities measured after sham exposure (Ctrl), magnetic stimulation (M), ultrasonic stimulation (US) and sonomagnetic stimulation (US + M). Data (mean with SEM) were calculated from three independent experiments (n = 150). The image contrast was increased by 10%. ***p < 0.001. Ctrl = sham exposure; F = Fluorescence intensity; M = magnetic stimulation; M + US = sonomagnetic stimulation; SEM = standard error of the mean; TMRE = tetramethylrhodamine, ethyl ester; US = ultrasonic stimulation.
The membrane potential responses of neuroblast-like cells (DC2.4, N2a and PC12), endothelial-like cells (BEND3 and HeLa) and fibroblast-like cells (MRC5 and 3T3) are shown in Figure 3ac, respectively. We observed a decrease in fluorescence intensity and membrane hyperpolarization in these cells, which have different morphologies. In particular, the neuroblast-like cells exhibited a significantly higher degree of membrane hyperpolarization than that of the endothelial-like cells when stimulated sonomagnetically at 1.1 MPa ultrasound pressure (Fig. 3d). At higher ultrasound pressure (2.2 MPa), the response of DC2.4 cells was significantly greater than that of the other types of cells. Mitochondrial membrane hyperpolarization The mitochondrial DCm plays a vital role in driving protons, ions and proteins to maintain healthy mitochondrial functions (Zorova et al. 2018). To explore whether sonomagnetic stimulation alters the DCm of live cells, we measured the mitochondrial DCm using the voltageresponsive probe TMRE. In Figure 4a, the red, granular
fluorescence indicated a normal mitochondrial potential. After sonomagnetic stimulation, the fluorescence intensity of the probe decreased, indicating a hyperpolarized state of DCm (Fig. 4b, 4c). Quantitative analysis (Fig. 4d) shows that ultrasonic stimulation induced a slight decrease of TMRE fluorescence, whereas sonomagnetic stimulation resulted in a substantial decrease of fluorescence. In line with the response of the cell membrane potential, sonomagnetic stimulation with higher ultrasound pressure (2.2 Pa) induced a greater mitochondrial membrane hyperpolarization. Increase in Ca2+ and NO concentration Ca2+ serves as a second messenger for cell signaling, and its intracellular concentration is affected by the cell membrane potential (Krasznai et al. 2000). Therefore, we examined the Ca2+ response of N2a cells after sonomagnetic stimulation. As shown in Figure 5a, before external stimulation, the green fluorescence of Fluo-4 was weak because the concentration of normal cytoplasmic Ca2+ was relatively low (150250 nM). After
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sonomagnetic stimulation of 1.1 MPa ultrasound pressure, the fluorescence intensity of Fluo-4 increased, indicating the elevation of cytoplasmic Ca2+ concentration. We note that at higher ultrasound pressure (2.2 MPa), the intracellular Ca2+ concentration further increased. This increase in intracellular Ca2+ concentration after sonomagnetic stimulation is quantified in Figure 5d. It is worth noting that confined Ca2+ regions in the form of bright spots were observed under sonomagnetic stimulation at both 1.1 and 2.2 MPa (indicated by yellow arrows in Fig. 5b, 5c). Because the intracellular production of NO is closely correlated with cytoplasmic Ca2+ signaling (Li et al. 2003), we investigated the influence of sonomagnetic stimulation on NO messenger production. As shown in Figure 6ac, the intracellular NO concentration of N2a cells was monitored with the green fluorescence probe DAF-FM diacetate. An increase in fluorescence intensity was observed after sonomagnetic stimulation. In particular, in both of the sonomagnetic groups (Fig. 6b, 6c; 1.1
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and 2.2 MPa, respectively), the localized NO concentration increased, as was evident from the bright spots indicated by the yellow arrows. Quantitative analysis in Figure 6d confirmed that sonomagnetic stimulation significantly elevated the intracellular production of NO. Intracellular acidification The intracellular pH plays an important role in modulating cell function and changes depending on the cell membrane potential (Schackmann et al. 1981). Because we confirmed that sonomagnetic stimulation can induce cell membrane hyperpolarization, we examined whether the intracellular pH was also affected by external physical stimulation. As shown in Figure 7ac, the increased fluorescence intensity of the pHrodo probe indicates that the intracellular pH of the exposed N2a cells decreased (i.e., the concentration of cytoplasmic H+ increased). Quantitative measurements showed that the pH shift was relatively small (i.e., fluorescence intensity
Fig. 5. Increase in intracellular Ca2+ concentration. (ac) Fluorescence images of Fluo-4-loaded cells after sham-exposure, sonomagnetic stimulation at 1.1 MPa ultrasound pressure and sonomagnetic stimulation at 2.2 MPa ultrasound pressure, respectively. (d) Quantitative analysis of the Fluo-4 fluorescence intensities measured after sonomagnetic stimulation at different ultrasound pressures (1.1 and 2.2 MPa). Data (mean with SEM) were calculated from three independent experiments (n = 150). The image contrast was increased by 10%. ***p < 0.001. Ca2+ = calcium; Ctrl = sham exposure; F = Fluorescence intensity; M = magnetic stimulation; M + US = sonomagnetic stimulation; SEM = standard error of the mean; US = ultrasonic stimulation.
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Fig. 6. Increase in intracellular NO concentration. (ac) Fluorescence images of DAF-FM-loaded cells after shamexposure, sonomagnetic stimulation at 1.1 MPa ultrasound pressure and sonomagnetic stimulation at 2.2 MPa ultrasound pressure, respectively. (d) Quantitative analysis of the DAF-FM fluorescence intensities measured after sonomagnetic stimulation at different ultrasound pressures (1.1 and 2.2 MPa). Data (mean with SEM) were calculated from three independent experiments (n = 150). The image contrast was increased by 10%. ***p < 0.001. Ctrl = sham exposure; DAFFM = to 4-Amino-5-Methylamino-20 ,70 -Difluorofluorescein; F = Fluorescence intensity; M = magnetic stimulation; M + US = sonomagnetic stimulation; NO = nitric oxide; SEM = standard error of the mean; US = ultrasonic stimulation.
increase of approximately 35% and pH decrease of approximately 0.3) (Fig. 7d). Increase in intracellular ROS level Hyperpolarization of the mitochondrial membrane induces chemical production of intracellular ROS. Having confirmed DCm hyperpolarization, we then investigated the intracellular level of ROS after sonomagnetic stimulation. The ROS levels of N2a cells were monitored with a green fluorescent H2DCF-DA probe. As shown in Figure 8ac, the stimulated cells exhibited increased fluorescence intensities and cytoplasmic ROS levels. Quantitative analysis in Figure 8d confirmed that ROS elevation in the sonomagnetic group was significantly higher than that of the sham-exposed control group.
proliferation (Roderick and Cook 2008; Blackiston et al. 2009). Therefore, we examined whether daily sonomagnetic stimulation (1.1 MHz frequency, 2.2 MPa ultrasound pressure, 680 mT-strength magnetic field and 10 min exposure time) affects the growth of N2a cancer cells. As shown in Figure 9a, the cells were imaged at 0, 24, 48 and 72 h after the first sonomagnetic stimulation. We found that the cells in the sonomagnetic group grew more slowly than those in sham-exposed group. Quantitative analysis confirmed this observation by showing that the proliferation rate of cells in the sonomagnetic group was inhibited by 48.64% at 72 h after the first stimulation (Fig. 9d).
DISCUSSION Inhibition of proliferation of N2a cells We found that sonomagnetic stimulation resulted in cell membrane hyperpolarization and intracellular NO production. In the literature, these two cellular responses are functionally correlated with the inhibition of cell
Modulation of cellular electrical and chemical signals by ultrasound is attractive for technical reasons because ultrasound can be non-invasively focused into deep tissues with good spatial resolution. When using ultrasound alone, high acoustic energy levels are
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Fig. 7. Intracellular acidification. (ac) Fluorescence images of pHrodo-loaded cells after sham-exposure, sonomagnetic stimulation at 1.1 MPa ultrasound pressure and sonomagnetic stimulation at 2.2 MPa ultrasound pressure, respectively. (d) Quantitative analysis of the pHrodo fluorescence intensities measured after sonomagnetic stimulation at different ultrasound pressures (1.1 and 2.2 MPa). Data (mean with SEM) were calculated from three independent experiments (n = 150). The image contrast was increased by 10%. ***p < 0.001. Ctrl = sham exposure; F = Fluorescence intensity; M = magnetic stimulation; M + US = sonomagnetic stimulation; SEM = standard error of the mean; US = ultrasonic stimulation.
required to evoke sonoelectrical and sonochemical effects (Honda et al. 2004). To decrease the acoustic energy to safe levels, ultrasound contrast agents (e.g., microbubble) and ultrasound-sensitive chemicals (e.g., 5-aminolevulinic acid) can be used (Qin et al. 2014; Sun et al. 2015). In this work, we proposed a new method using ultrasound and a static magnetic field to evoke electrophysiologic and biochemical responses of live cells. Our results demonstrated that the sonomagnetic stimulation method (1.1 MHz frequency, 1.1 or 2.2 MPa acoustic pressure, 50 cycles per pulse, 500 Hz pulse repetition frequency and 680 mT strength magnetic field) induced cell and mitochondrial membrane hyperpolarization and also increased the production of intracellular biochemical signaling species (Ca2+, NO, H+ and ROS). We note that, according to eqn (8), the same evoked cellular effects could be achieved using lower ultrasound energy levels if we increase the magnetic field strength accordingly. In 2017, Rekhi and Arbabian (2017) used a 2 T strength magnetic field for generating Lorentz current. If we increased the magnetic field strength from 680 mT to 2 T, the ultrasound pressure could be
theoretically decreased from 2.20.748 MPa. In this way, the calculated spatial-peak time-averaged intensity of the ultrasound wave is 463 mW/cm2, which is smaller than the FDA-approved safety level of 720 mW/cm2 for diagnostic ultrasound equipment (Nelson et al. 2009). The electrophysiologic and biochemical responses induced in this study can affect other cell behavior (e.g., proliferation). For example, it has been reported that hyperpolarization of the cell membrane turns the cell into a quiescent state with slow proliferation rate, whereas depolarization drives cells into an active state whereby they grow more quickly (Levin 2007). A low concentration of cytoplasmic NO promotes cell proliferation, whereas a higher concentration of NO induces cell cycle arrest and inhibits cell growth (Napoli et al. 2013). Moreover, when the intracellular pH is decreased, the proliferation rate of cancer cells has been shown to be inhibited (Che et al. 2008). Although the roles of mitochondrial DCm, Ca2+ and ROS concentrations in cell proliferation are still unclear, these signals are highly correlated with cell cycle progression and proliferation rate (Pinto et al. 2015; Diebold and Chandel 2016). With
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Fig. 8. Increase in intracellular ROS level. (ac) Fluorescence images of H2DCF-DA-loaded cells after sham-exposure, sonomagnetic stimulation at 1.1 MPa ultrasound pressure and sonomagnetic stimulation at 2.2 MPa ultrasound pressure, respectively. (d) Quantitative analysis of the H2DCF-DA fluorescence intensities measured after sonomagnetic stimulation at different ultrasound pressures (1.1 and 2.2 MPa). Data (mean with SEM) were calculated from three independent experiments (n = 150). The image contrast was increased by 10%. ***p < 0.001. Ctrl = sham exposure; F = Fluorescence intensity; H2DCF-DA = 20 ,70 -Dichlorodihydrofluorescein; diacetate F = Fluorescence intensity; M = magnetic stimulation; M + US = sonomagnetic stimulation; ROS = reactive oxygen species; SEM = standard error of the mean; US = ultrasonic stimulation.
our results demonstrating that sonomagnetic stimulation can induce cell membrane hyperpolarization and modulate the production of signaling molecules (NO and H+), we further examined the proliferation of the stimulated cells. We found that the proliferation of these cells was inhibited, demonstrating the potential of the sonomagnetic stimulation method in controlling cancer cell proliferation. In future work, two critical questions associated with sonomagnetic stimulation need further investigation. The first question is how the scale of cell membrane hyperpolarization is linked to the cell type. In the present study, we found that cells of different type responded to sonomagnetic stimulation differently. Such cell-type dependence has been previously reported in a sonoporation study (Shi et al. 2017). Exploration of this question is of particular interest because this cell-type-dependent behavior could be used for cell diagnosis and imaging (Kunyansky et al. 2017). The second question is how to precisely control the cellular responses of live cells evoked by sonomagnetic stimulation. It is known that even for the same
physical stimulation method, different parameters can lead to different biologic responses. For example, highfrequency (>5 Hz) TMS excites neurons, whereas at low frequency (0.21 Hz), it inhibits the electrical activity of neurons (Hallett 2007). Therefore, the ultrasound parameters of sonomagnetic stimulation (e.g., frequency, pulse duration and pressure) need to be better controlled to achieve precise modulation of electrophysiologic, biochemical and behavioral responses. The proposed sonomagnetic stimulation method holds great promise to non-invasively modulate the in vivo signaling of cells at a spatial resolution of approximately 12 mm depending on the focusing ability of the ultrasound wave. However, the clinical application of this method was limited by the technical difficulty in the generation of large-scale (e.g., 45 £ 45 £ 45 cm) homogenous magnetic field with high strength. In this study, the length of the gap between the two NdFeB magnets where the cell samples were placed was 13 mm. In the work carried out by Rekhi and Arbabian (2017), the length of the gap between magnets was 5 mm. Note
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Fig. 9. Inhibition of proliferation of N2a cells. (a) Images of N2a cells at 0, 24, 48 and 72 h after the first sonomagnetic stimulation. (b) Quantitative analysis of the proliferation rate of N2a cells after sham exposure (Ctrl), magnetic stimulation (M), ultrasonic stimulation (US) and sonomagnetic stimulation (US + M). For each experiment, five imaging areas were selected for cell number counting. Data (mean with SEM) were calculated from three independent experiments (n = 150). Ctrl = sham exposure; F = Fluorescence intensity; M = magnetic stimulation; M + US = sonomagnetic stimulation; N2a = Neuro-2a; SEM = standard error of the mean; US = ultrasonic stimulation.
that the increase in the gap length will decrease magnetic field strength. Therefore, it is not practical to use permanent magnets for sonomagnetic stimulation of cells at significant tissue depths. We suggest that electromagnets made from coils of superconducting wire that have been implemented for magnetic resonance imaging can be employed for the clinical investigation of sonomagnetic stimulation (Shen et al. 2017).
can be technically enhanced by increasing the magnetic field strength.
CONCLUSION
REFERENCES
In this study, we demonstrated the feasibility and effectiveness of sonomagnetic stimulation in evoking electrophysiologic, biochemical and behavioral responses of live cells. Our results suggest that this method could potentially be applied to modulate cell and mitochondrial membrane potentials, to trigger the production of intracellular signaling species and to inhibit the growth of cancer cells. The advantages of this physical stimulation method are that its spatial energy distribution can be effectively confined inside the ultrasound focal region and its stimulation effects
Blackiston DJ, McLaughlin KA, Levin M. Bioelectric controls of cell proliferation ion channels, membrane voltage and the cell cycle. Cell Cycle 2009;8:3527–3536. Che X-F, Akiyama S-I, Tomoda A. Suppression of the proliferation of cancer cell lines, KB-3-1 and K562 cells preceded by a decrease in intracellular pH caused by phenoxazine derivatives. Oncol Rep 2008;19:1253–1258. Debye P. A method for the determination of the mass of electrolytic ions. J Chem Phys 1933;1:13–16. Diebold L, Chandel NS. Mitochondrial ROS regulation of proliferating cells. Free Radic Biol Med 2016;100:86–93. Gilbert G, Courtois A, Dubois M, Cussac LA, Ducret T, Lory P, Marthan R, Savineau JP, Quignard JF. T-type voltage gated calcium
Acknowledgments—This work was funded by the National Natural Science Foundation of China (Grant No. 61427806), the Shenzhen Science and Technology Planning Project (Grant No. JCYJ20160520170055193), the Medical Project of Guangdong Province (A2019542) and the Natural Science Foundation of Shenzhen University (Grant No.000218). Conflict of interest—The authors declare no other potential conflicts of interest.
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channels are involved in endothelium-dependent relaxation of mice pulmonary artery. Biochem Pharmacol 2017;138:61–72. Giovannini C, Matarrese P, Scazzocchio B, Sanchez M, Masella R, Malorni W. Mitochondria hyperpolarization is an early event in oxidized low-density lipoprotein-induced apoptosis in Caco-2 intestinal cells. FEBS Letters 2002;523:200–206. Grasland-Mongrain P, Mari JM, Chapelon JY, Lafon C. Lorentz force electrical impedance tomography. Irbm 2013;34:357–360. Hallett M. Transcranial magnetic stimulation: A primer. Neuron 2007;55:187–199. Heiskanen KM, Bhat MB, Wang H-W, Ma J, Nieminen A-L. Mitochondrial depolarization accompanies cytochrome c release during apoptosis in PC6 Cells. J Biol Chem 1999;274:5654–5658. Hensel K, Mienkina MP, Schmitz G. Analysis of ultrasound fields in cell culture wells for in vitro ultrasound therapy experiments. Ultrasound Med Biol 2011;37:2105–2115. Honda H, Kondo T, Zhao QL, Feril LB, Jr., Kitagawa H. Role of intracellular calcium ions and reactive oxygen species in apoptosis induced by ultrasound. Ultrasound Med Biol 2004;30:683–692. Iwasaki H, Murata Y, Kim Y, Hossain MI, Worby CA, Dixon JE, McCormack T, Sasaki T, Okamura Y. A voltage-sensing phosphatase, Ci-VSP, which shares sequence identity with PTEN, dephosphorylates phosphatidylinositol 4,5-bisphosphate. Proc Natl Acad Sci USA 2008;105:7970–7975. Krasznai Z, Marian T, Izumi H, Damjanovich S, Balkay L, Tron L, Morisawa M. Membrane hyperpolarization removes inactivation of Ca2+ channels, leading to Ca2+ influx and subsequent initiation of sperm motility in the common carp. Proc Natl Acad Sci USA 2000;97:2052–2057. Kunyansky L, Ingram CP, Witte RS. Rotational magneto-acoustoelectric tomography (MAET): theory and experimental validation. Phys Med Biol 2017;62:3025–3050. Lee I, Bender E, Arnold S, Kadenbach B. New control of mitochondrial membrane potential and ROS formationa hypothesis. Biol Chem 2001;382:1629–1636. Legon W, Sato TF, Opitz A, Mueller J, Barbour A, Williams A, Tyler WJ. Transcranial focused ultrasound modulates the activity of primary somatosensory cortex in humans. Nat Neurosci 2014;17: 322–329. Levin M. Large-scale biophysics: Ion flows and regeneration. Trends Cell Biol 2007;17:261–270. Li N, Sul J-Y, Haydon PG. A calcium-induced calcium influx factor, nitric oxide, modulates the refilling of calcium stores in astrocytes. J Neurosci 2003;23:10302–10310. McCaig CD, Rajnicek AM, Song B, Zhao M. Controlling cell behavior electrically: Current views and future potential. Physiol Rev 2005;85:943–978. Montalibet A, Jossinet J, Matias A, Cathignol D. Electric current generated by ultrasonically induced Lorentz force in biological media. Med Biol Eng Comput 2001;39:15–20. Napoli C, Paolisso G, Casamassimi A, Al-Omran M, Barbieri M, Sommese L, Infante T, Ignarro LJ. Effects of nitric oxide on cell proliferation: Novel insights. J Am Coll Cardiol 2013;62:89–95. Nelson TR, Fowlkes JB, Abramowicz JS, Church CC. Ultrasound biosafety considerations for the practicing sonographer and sonologist. J Ultrasound Med 2009;28:139–150.
Volume 00, Number 00, 2019 Nusser Z. Variability in the subcellular distribution of ion channels increases neuronal diversity. Trends Neurosci 2009;32:267–274. Perlmutter JS, Mink JW. Deep brain stimulation. Annu Rev Neurosci 2006;29:229–257. Perry SW, Norman JP, Barbieri J, Brown EB, Gelbard HA. Mitochondrial membrane potential probes and the proton gradient: A practical usage guide. Biotechniques 2011;50:98–115. Pinto MC, Kihara AH, Goulart VA, Tonelli FM, Gomes KN, Ulrich H, Resende RR. Calcium signaling and cell proliferation. Cell Signal 2015;27:2139–2149. Pucihar G, Kotnik T, Kanduser M, Miklavcic D. The influence of medium conductivity on electropermeabilization and survival of cells in vitro. Bioelectrochemistry 2001;54:107–115. Qin P, Xu L, Hu Y, Zhong W, Cai P, Du L, Jin L, Yu AC. Sonoporationinduced depolarization of plasma membrane potential: Analysis of heterogeneous impact. Ultrasound Med Biol 2014;40:979–989. Rekhi AS, Arbabian A. Remote sub-wavelength focusing of ultrasonically activated Lorentz current. Appl Phys Lett 2017;110 164104. Roderick HL, Cook SJ. Ca2+ signalling checkpoints in cancer: Remodelling Ca2+ for cancer cell proliferation and survival. Nat Rev Cancer 2008;8:361–375. Schackmann RW, Christen R, Shapiro BM. Membrane potential depolarization and increased intracellular pH accompany the acrosome reaction of sea urchin sperm. Proc Nat Acad Sci USA 1981;78:6066–6070. Schram G. Differential distribution of cardiac ion channel expression as a basis for regional specialization in electrical function. Circ Res 2002;90:939–950. Sena LA, Chandel NS. Physiological roles of mitochondrial reactive oxygen species. Mol Cell 2012;48:158–167. Shen B, Geng J, Li C, Zhang X, Fu L, Zhang H, Ma J, Coombs TA. Optimization study on the magnetic field of superconducting Halbach Array magnet. Physica C Supercond 2017;538:46–51. Shi D, Guo L, Duan S, Shang M, Meng D, Cheng L, Li J. Influence of tumor cell lines derived from different tissue on sonoporation efficiency under ultrasound microbubble treatment. Ultrason Sonochem 2017;38:598–603. Sun X, Xu H, Shen J, Guo S, Shi S, Dan J, Tian F, Tian Y, Tian Y. Real-time detection of intracellular reactive oxygen species and mitochondrial membrane potential in THP-1 macrophages during ultrasonic irradiation for optimal sonodynamic therapy. Ultrason Sonochem 2015;22:7–14. Sundelacruz S, Levin M, Kaplan DL. Comparison of the depolarization response of human mesenchymal stem cells from different donors. Sci Rep 2015;5:18279. Wright JJ, Van Der Beken S. The Hall effect in a flowing electrolyte. Am J Phys 1972;40:245–247. Yamada A, Gaja N, Ohya S, Muraki K, Narita H, Ohwada T, Imaizumi Y. Usefulness and limitation of DiBAC4(3), a voltage-sensitive fluorescent dye, for the measurement of membrane potentials regulated by recombinant large conductance Ca2+-activated K+ channels in HEK293 cells. Jpn J Pharmacol 2001;86:342–350. Yang M, Brackenbury WJ. Membrane potential and cancer progression. Front Physiol 2013;4:185. Zorova LD, Popkov VA, Plotnikov EY, Silachev DN, Pevzner IB, Jankauskas SS, Babenko VA, Zorov SD, Balakireva AV, Juhaszova M, Sollott SJ, Zorov DB. Mitochondrial membrane potential. Anal Biochem 2018;552:50–59.
https://doi.org/10.1016/j.ultrasmedbio.2019.07.009
Original Contribution SONOMAGNETIC STIMULATION OF LIVE CELLS: ELECTROPHYSIOLOGIC, BIOCHEMICAL AND BEHAVIORAL RESPONSES AGEDPY T AXIN
HU,*,y YANCHENG WANG,*,y XIN CHEN,*,y and SIPING CHEN*,yTAGEDEN
* School of Biomedical Engineering, Health Science Center, Shenzhen University, Shenzhen, P.R. China; and y National-Regional Key Technology Engineering Laboratory for Medical Ultrasound, Shenzhen, P.R. China (Received 19 November 2018; revised 1 July 2019; in final from 6 July 2019)
Abstract—Various physical methods have been developed to modulate the electrophysiologic properties of cells and their biochemical signaling pathways. In this study, we propose a sonomagnetic method using pulsed ultrasound (1.1 MHz frequency, 1.1 or 2.2 MPa pressure, 50 cycles per pulse and 500 Hz pulse repetition frequency) and a static magnetic field (680 mT) to stimulate live cells. We found that sonomagnetic stimulation promoted the cell and mitochondrial membrane potentials to more hyperpolarized states. The degree of cell membrane hyperpolarization was cell-type dependent. Furthermore, we found that the intracellular concentrations of Ca2+ ions, reactive oxygen species and nitric oxide were substantially increased after sonomagnetic stimulation, and a small decrease in intracellular pH was also observed. Lastly, we found that the daily sonomagnetic stimulation for 3 d inhibited the proliferation rate of neuro-2a cancer cells by 48.64%. Our work demonstrates that sonomagnetic stimulation can effectively perturb cell signaling and drive cancer cells into relatively quiescent states. (E-mail: [email protected]) © 2019 World Federation for Ultrasound in Medicine & Biology. All rights reserved. Key Words: Ultrasound, Magnetic field, Membrane potential, Biochemical signal.
triphosphate for metabolic activities. By contrast, it has been proposed that mitochondria with high DCm initiate programmed cell death (Heiskanen et al. 1999; Zorova et al. 2018). Therefore, the modulation of cellular and organelle membrane potentials is important for controlling cell functions. Considerable research has revealed the complex ways in which the cell and organelle membrane potentials affect cell behavior. Mechanistic studies have shown that the bioelectrical signals of membrane potentials are first transduced into biochemical signals to trigger subsequent intracellular signaling cascades. For example, changes in Vmem can be transduced into the cell via one of two pathways: (i) activation of voltage-gated ion channels and (ii) activation of voltage-sensitive phosphatase. In the first pathway, a hyperpolarized Vmem results in the influx of divalent Ca2+, which rapidly triggers the production of signaling molecules, such as calmodulin and nitric oxide (NO) to modulate cell behavior (Krasznai et al. 2000; Gilbert et al. 2017). In the second pathway, after membrane potential alternation, the activated phosphatase removes phosphate from the inositol rings of phosphatidylinositol messengers (e.g., phosphatidylinositol (3,4,5)-trisphosphate (PI[3,4,5]P3), and phosphatidylinositol 4,5-bisphosphate
INTRODUCTION A eukaryotic cell manages its autonomous life-sustaining processes via exquisite physical and chemical signals. One important cell signaling mechanism is the electrical modulation of voltage gradients generated across the cell and organelle membranes (McCaig et al. 2005). For example, the balance of ion currents produced by transmembrane channels and pumps results in a resting cell membrane potential (Vmem) ranging from 10 to 90 mV, depending on the specific cell type. An increase of this membrane potential (a hyperpolarized Vmem) can turn the cell into a relatively quiescent state with a slower proliferation rate, whereas a decrease of this membrane potential (a depolarized Vmem) makes the cell mitotically active and proliferate more quickly (Blackiston et al. 2009; Yang and Brackenbury 2013). In addition, mitochondria, the energy “power-plant” of cells, also have a transmembrane potential (DCm), ranging from 130 to 180 mV, which has an important role in various cell functions. Mitochondria with low values of DCm cannot generate sufficient adenosine Address correspondence to: Xin Chen, PhD, 3688 Nanhai Avenue, Shenzhen, Guangdong, P.R. China. E-mail: [email protected]
1
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(PI[4,5]P2)) to regulate cell motility and shape (Iwasaki et al. 2008). Mitochondrial DCm is also transduced into chemical signals. For example, a high DCm can accelerate the mitochondrial respiratory chain in an exponential manner to produce more reactive oxygen species (ROS). The oxidative stress induced by harmful levels of ROS (e.g., superoxide [O2 ] and hydroxyl radicals [OH ]) opens membrane permeability transition pores and releases cytochrome c to trigger apoptosis (Lee et al. 2001; Sena and Chandel 2012). Because the cell and organelle membrane potentials are important modulators of intracellular biochemical signals, various electrical stimulation methods have been developed to modulate membrane potentials. In in vitro studies, chemicals or drugs have been directly added to the extracellular medium (e.g., potassium ions for cell membrane depolarization and oligomycin for mitochondria hyperpolarization) (Giovannini et al. 2002; Sundelacruz et al. 2015). In in vivo studies, direct current stimulation has proven to be an effective approach; however, this method needs invasive implantation of electrodes (Perlmutter and Mink 2006). It has been shown that magnetic stimulation can affect cell membrane potentials, and transcranial magnetic stimulation (TMS) has been approved by the US Food and Drug Administration (FDA). The technical limitation of magnetic stimulation is its spatial resolution of several centimeters, which is too large for precise regional stimulation (Hallett 2007). Recently, ultrasound has been shown to be able to stimulate neuronal circuits (Legon et al. 2014). However, the underlying mechanism of ultrasound stimulation is still largely unknown. In this work, we propose a sonomagnetic method to evoke electrophysiologic, biochemical and behavioral responses of live cells. This sonomagnetic method is developed based on the following two effects: (i) the Debye effect, which shows that ions with different masses and charges in a physiologic solution would vibrate and separate from each other when driven by ultrasound energy (Debye 1933) and (ii) the Hall effect, which employs a static magnetic field to generate Lorentz forces in opposite directions to further separate the moving ions of different charges and to produce an electrical current (Wright and Van Der Beken 1972). Note that the feasibility of employing sonomagnetic stimulation for generating Lorentz currents in physiologic solution has been proven. Montalibet et al. (2001) measured a 320 nA Lorentz current evoked by 2 MPa ultrasound (500 kHz frequency) combined with a 350 mT strength magnetic field in NaCl solution with a conductivity of 1.6 S/m. Using a 420 kPa ultrasound (1.1 MHz frequency) and a 2 T strength magnetic field, Rekhi and Arbabian (2017) measured a 3.1 mA Lorentz current in
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1 M KCl solution (11.7 S/m conductivity). The sonomagnetically-stimulated Lorentz current was also confirmed in issue phantoms and ex vivo tissue samples (Grasland-Mongrain et al. 2013; Kunyansky et al. 2017). However, the question of whether the sonomagnetic method could be used for electrical stimulation of the electrophysiologic state of live cells and the production of biochemical signaling species has not been answered. Therefore, we established a sonomagnetic experiment platform for live cell stimulation. Cell and mitochondrial membrane potentials were fluorescently measured to examine the electrophysiologic responses of cells. Biochemical signaling species, including Ca2+, NO, H+ and ROS, were also fluorescently measured to examine the cells’ biochemical responses. Finally, the behavioral responses of the cells after sonomagnetic stimulation were monitored. Our work aims to provide experimental evidence for the feasibility and effectiveness of sonomagnetic stimulation of live cells. THEORY As demonstrated in Figure 1a, the propagation of ultrasound (longitudinal pressure wave along the z-axis) in an electrolyte solution results in the oscillation of the fluid and ions. When a static magnetic field is applied in the direction (y-axis) perpendicular to the direction of ultrasound propagation, Lorentz forces will be generated on the oscillating ions. The magnitude of the Lorentz force of an ion with charge q is given by FLorentz ¼ qvB
ð1Þ
where v is the velocity of the ion and B is the magnetic field strength. Driven by the generated Lorentz forces, positive and negative ions will move in opposite directions to produce an electrical current. Assuming the positive and negative ions have charges of q and q, respectively, the electrical current density, j, can be calculated as: j ¼ ðnþ mþ þ nmÞFLorentz
ð2Þ
where n+ and n are the ion concentrations, and m+ and m are the ion mobilities. Using eqn (1) and eqn (2), j can be written as: j ¼ qðnþ mþ þ nmÞvB
ð3Þ
Because the electrical conductivity of the electrolyte solution is given by: s ¼ qðnþ mþ þ nmÞ
ð4Þ
j can be expressed as: j ¼ svB
ð5Þ
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Fig. 1. Experimental setup and Lorentz current detection. (a) Graphical illustration of sonomagnetic stimulation theory. (b) Generation of the ultrasonic and magnetic fields for live cell stimulation. (c) The ultrasound pressure map measured by a needle hydrophone. (d) The measured magnetic field strength (left) and the calculated field strength gradients along the x-axis (middle) and the y-axis (right). (e) Immersible chamber with live cells grown on its bottom. (f) Immersible chamber with a 0.7 mm width copper wire placed on its bottom and the Lorentz current detection system. (g) Lorentz current elicited in the copper wire by 1.1 MPa pressure ultrasound (20 cycles per pulse and 500 Hz pulse repetition frequency) and 680 mT magnetic field strength. (h) The spectrum of the Lorentz current generated by sonomagnetic stimulation in the copper wire. DPO = oscilloscope; f = frequency; FLorentz = Lorentz Force; NdFeB = Neodymium Iron Boron; p(t,z) = ultrasound pressure; PXPA6-40 dB = 40 dB differential amplifier; q = electrical charge; t = time; u = electrical voltage; v(t,z) = ion oscillation speed.
The ion oscillation speed, v, can be calculated from the classical ultrasound propagation equation: r
@vðt; zÞ @Pðt; zÞ þ ¼0 @t @z
ð6Þ
where r is the mass density of the solution and P(t, z) is the ultrasound pressure, which varies in time, t, and with distance, z. Therefore, the ion oscillation speed, v(t, z), can be calculated by integrating the above equation with respect to time: Zt 1 @Pðt; zÞ vðt; zÞ ¼ dt ð7Þ r @z 1
Substituting eqn (7) into eqn (5), we get: sB jðt; zÞ ¼ r
Zt 1
@Pðt; zÞ dt @z
ð8Þ
According to eqn (8), the generated electrical density can be modulated by three parameters: the solution conductivity, s, the magnetic field strength, B, and the ultrasound pressure, P. For example, when s and B are constant, increasing the ultrasound pressure increases the generated electrical current density, j. Comparing the parameters used in this study (frequency of 1.1 MHz, conductivity of 1.6 S/m and magnetic field strength of
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680 mT) with those employed by Rekhi and Arbabian (2017), it can be inferred that the amplitude of the generated Lorentz current in the extracellular medium by an ultrasound wave of 2.2 MPa pressure is in the range of 0.75 mA. It is worth noting that the conductivity of the lipid membrane (5 £ 107 S/m) is much lower than that of physiologic solutions (0.21.6 S/m) (Pucihar et al. 2001). Therefore, voltage gradients are generated at cell and organelle membrane boundaries (Kunyansky et al. 2017). MATERIALS AND METHODS Experimental setup To produce ultrasound waves, an electrical signal was first edited in a function generator (DG1022, Agilent Technologies, Santa Clara, CA, USA), then amplified by a 50 dB broadband power amplifier (2100L, Electronics & Innovation, Rochester, NY, USA), and finally transformed into acoustic waves by an annular high-intensity focused transducer (H102, Sonic Concepts, Bothell, WA, USA) (Fig. 1b). The outer diameter of the transducer is 64 mm, and its inner diameter is 20 mm. The focal depth of the transducer is 51.74 mm. The cell samples were placed 61 cm from the transducer (9.26 mm from the transducer focus), where the ultrasound pressure distribution was relatively homogenous. The ultrasound pressure map generated in the cell sample region was measured using a needle hydrophone (HNR-0500, Onda Corp, Sunnyvale, CA, USA) (Fig. 1c). Pulsed ultrasound waves (1.1 MHz center frequency, 50 cycles per pulse and 500 Hz pulse repetition frequency) with two peak negative pressure settings (1.1 and 2.2 MPa) were used for electrochemical stimulation of live cells. To generate a static magnetic field, two Neodymium Iron Boron (NdFeB) magnets (cubic shape, 5 £ 5 £ 2.5 cm) were used. The gap between these two magnets was 13 mm. The measured magnetic field strength in the live cell region (indicated by the dashed circle in left image of Fig. 1d) was 680 mT. The magnetic field gradients calculated along the x-axis and the y-axis were also given in Figure 1d (the middle and the right image). Immersible cell chamber It is reported that when the cells were grown in culture wells for in vitro ultrasound experiments, standing waves would be generated in the culture medium to affect the cell stimulation results (Hensel et al. 2011). To minimize the influence of standing waves, Hensel et al. (2011) suggested that the air-medium interface in the wells should be avoided, and the wells should be submersed into the water tank to receive ultrasound
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exposure. Therefore, in this study, we designed an immersible cell chamber for sonomagnetic stimulation. The rectangular chamber (61 mm [L] £ 12.6 mm [W] £ 5 mm [H]) had a circular window area (8.6 mm diameter) for cell culture. In particular, both sides of the circular window were sealed by an 170 mm thick cover glass. Live cells could be seeded into the chamber through an injection channel of 1.5 mm diameter. During the experiment, the chamber was filled with degassed Hank’s balanced salt solution (1672547, Invitrogen, Carlsbad, CA, USA) with 10 mM N-2-hydroxyethylpiperazine-N-2-ethane sulfonic acid (HEPES) (15630080, Invitrogen). In this way, no air would be left in the chamber, and no air-water interface was formed. Furthermore, the injection channel of the chamber could be sealed by sterilized polystyrene membranes. The chamber could be submersed in the degassed water to further minimize ultrasound reflection and standing wave formation. Lorentz current confirmation To confirm the generation of Lorentz current in our experimental setup, a 0.7 mm width copper wire was inserted into the immersible chamber (Fig. 1f). The chamber was placed between the two magnets to receive the ultrasound exposure. The electrical signal generated at the two ends of the copper wire was amplified by a 40 dB differential amplifier (PXPA6-40, Pengxiang, Changsha, Hunan, China) and recorded by an oscilloscope (DPO 5054, Tektronix, Beaverton, OR, USA). As shown in Figure 1g, the averaged amplitude of the electrical signal elicited in copper wire by the 1.1 MPa pressure ultrasound wave (20 cycles per pulse and 500 Hz pulse repetition frequency) and 680 mT strength magnetic field was 24.3 mV. Note that, when the magnets were removed, the electrical signal disappeared. As shown in Figure 1h, the frequency of the Lorentz current was the same as the ultrasound wave, which was consistent with eqn (8) given in the Theory section. Cell culture In this study, we employed mouse neuroblastoma cell line (neuro-2a or N2a, ATCC CCL-131, Manassas, VA, USA) to study the cellular effects induced by sonomagnetic stimulation. The N2a cells were cultured in minimum essential medium (10370070, Invitrogen) supplemented with 10% fetal bovine serum (10099141, Invitrogen). When the cells reached a confluency of 70%, they were detached from the bottom of a 25 cm2 flask by trypsin- ethylenediaminetetraacetic acid (25200072, Invitrogen). Then, 5 £ 103 cells were seeded onto the glass bottom of the immersible cell chamber. After a 24 h incubation time at 37˚C and 5% CO2, the cells were fully attached to the glass bottom
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and could be used for the sonomagnetic stimulation experiments. In addition to N2a cells, we employed another six types of cells to examine the question of whether the cell membrane potential alternation induced by sonomagnetic stimulation was a cell-type dependent phenomenon. These six types of cells were categorized according to their morphology: (i) cells of neuroblast-like morphology, including mouse dendritic cell of DC2.4 and rat pheochromocytoma cell of PC12; (ii) cells of endothelial-like morphology, including mouse endothelioma cell of BEND3 and human cervical cancer cell of HeLa; and (iii) cells of fibroblast-like morphology, including human lung cell of MRC5 and mouse embryo cell of 3T3. Four types of cells (PC12, BEND3, HeLa and 3T3) were grown in Dulbecco’s modified Eagle medium (11965092, Invitrogen). The DC2.4 cells were grown in Roswell Park Memorial Institute 1640 medium (11875093, Invitrogen). MRC5 cells were grown in minimum essential medium. Note that, all of the media were supplemented with 10% fetal bovine serum for cell culture. To avoid the influence of phenol red on the fluorescence measurement, the cells were temporarily cultured in Hanks’ balanced salt solution (14025092, Invitrogen) supplemented with 20 mM HEPES during the examination of bioeffects induced by sonomagnetic stimulation. Membrane potential labeling To monitor changes in Vmem, a voltage-sensitive fluorescent probe, Bis-(1,3-Dibutylbarbituric Acid)Trimethine Oxonol (DiBAC4(3)) (B438, Invitrogen), was loaded into live cells at a final concentration of 200 nM. In general, the green fluorescence intensity of DiBAC4(3) increases with membrane depolarization and decreases with membrane hyperpolarization. To monitor changes in the mitochondrial DCm, a red fluorescent tetramethylrhodamine, ethyl ester (TMRE) probe (ENZ52309, Enzo Life Sciences, Farmingdale, NY, USA) was used to stain live cells at a final concentration of 100 nM. We note that TMRE loaded at this high concentration works in the quenching mode, which means mitochondrial hyperpolarization results in a relative decrease in fluorescence intensity of the dye and vice versa (Perry et al. 2011). Labeling of biochemical signaling species The fluorescent probe Fluo-4 AM (F14201, Invitrogen), at a final concentration of 2 mM, was used to monitor the cytoplasmic Ca2+ concentration. The intracellular NO level was monitored using the probe 4-Amino-5-Methylamino-2’,7’-Difluorofluorescein (DAF-FM) diacetate (D23844, Invitrogen) at a final concentration of 5 mM. The cytoplasmic pH was monitored using the probe pHrodo Red AM (P35372, Invitrogen) at a final concentration of
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5 mM. The intracellular ROS level was assessed using the probe 20 ,70 -Dichlorodihydrofluorescein diacetate (H2DCFDA) (D399, Invitrogen) at a final concentration of 3 mM. Fluorescent labeling of live cells was carried out according to the product protocols. Monitoring of cell proliferation To allow the cells to proliferate for 3 d without the contact-induced inhibition of division, N2a cells were seeded into the immersible chamber at relatively low density (1 £ 103 cells per chamber). The cells were then divided into four groups (a control group, an ultrasoundexposure group, a magnetic field group and a sonomagnetic stimulation group) and received daily physical treatment, respectively. To monitor the proliferation status of cells in different groups, the cells in the center of the immersible chamber were imaged before each treatment using an inverted microscope (Axio Observer A1, Carl Zeiss Meditec AG, Oberkochen, Germany) and a 63 £ objective. The number of the cells in the imaging view (135 £ 100 mm) was counted manually using the ImageJ software (v.1.52a, National Institutes of Health, Bethesda, MD, USA) and its plug-in (Cell Counter). Quantitative calculation Fluorescently labeled cells in the central area (diameter of 4 mm) of the immersible chamber were imaged using the inverted microscope (Axio Observer A1, Carl Zeiss Meditec AG). Microscope images were then imported into the image processing and analysis software ImageJ. The fluorescence intensity of each cell was calculated by manually outlining the cell boundary (white dashed line, Fig. 2a) with the aid of the Find Edges and Measure functions of ImageJ. The background fluorescence intensity was also calculated in the rectangular region of interest (10 mm [L] £ 12.6 mm [W]; yellow dashed line, Fig. 2a). We note that the background fluorescence intensity was subtracted from the cellular fluorescence intensity. To determine the statistical significance of the fluorescent intensities of the different experimental groups, the data were analyzed using GraphPad Prism software (v.5, GraphPad Software, San Diego, CA, USA) using the Friedman test (i.e., a nonparametric repeated measures analysis of variance). RESULTS Cell membrane hyperpolarization To calibrate the fluorescence responses of DiBAC4(3) probe in our system, we modulated the plasma membrane of N2a cells by increasing the extracellular K+ concentration, which linearly depolarizes the membrane potential (Yamada et al. 2001). We found that the green fluorescence intensity of DiBAC4(3)
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Fig. 2. Cell membrane hyperpolarization induced by sonomagnetic stimulation. (a) Microscope images showing the fluorescence intensity increase of DiBAC4(3)-loaded cells after extracellular K+ stimulation. (b) Quantitative calibration of the DiBAC4(3) fluorescence intensity measured under K+ stimulation at different concentrations. (c) Microscope images showing the fluorescence intensity decrease of DiBAC4(3)-loaded cells after sonomagnetic stimulation. (d) Quantitative analysis of the DiBAC4(3) fluorescence intensities measured after sham exposure (Ctrl), magnetic stimulation (M), ultrasonic stimulation (US) and sonomagnetic stimulation (US + M). (e) Quantitative analysis of the DiBAC4(3) fluorescence intensities measured after US and US + M at different pressures. (f) Quantitative analysis of the DiBAC4(3) fluorescence intensities measured after sonomagnetic stimulation for different exposure times. Data (mean with SEM) were calculated from three independent experiments (n = 150). The image contrast was increased by 10%. **p < 0.01; ***p < 0.001. Ctrl = sham exposure; DiBAC4(3) = Bis-(1,3-Dibutylbarbituric Acid)Trimethine Oxonol; F = Fluorescence intensity; K+ = potassium; M = magnetic stimulation; M + US = sonomagnetic stimulation; SEM = standard error of the mean; US = ultrasonic stimulation.
increased with the concentration of extracellular K+ (Fig. 2a). Quantitative analysis further showed the relationship between the concentration of K+ and DiBAC4(3) fluorescence intensity was almost linear, demonstrating the effectiveness of this voltage-sensitive probe in our experiments (Fig. 2b). We found that sonomagnetic stimulation resulted in a decrease in fluorescence intensity of the membraneloaded DiBAC4(3) probe, which was indicative of hyperpolarization of the membrane potential (Fig. 2c). Moreover, sonomagnetic stimulation at higher ultrasound pressure (2.2 MPa) induced a greater decrease of
DiBAC4(3) fluorescence intensity than that at lower ultrasound pressure (1.1 MPa). We also studied the changes in cell membrane potential induced by different physical stimuli, including magnetic, ultrasonic and sonomagnetic stimulation. As shown in Figure 2d, magnetic stimulation had little effect on the cell membrane potential. Ultrasonic stimulation induced a small-scale hyperpolarization of the cell membrane (4.17%). By contrast, sonomagnetic stimulation resulted in significant hyperpolarization of the cell membrane (47.89%), confirming the effectiveness of sonomagnetic stimulation in modulating the membrane potential.
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Consistent with eqn (8), according to which the electrical current density generated by sonomagnetic stimulation is positively correlated with ultrasound pressure, we found that ultrasound at higher pressure (2.2 MPa) evoked a significant higher degree of membrane hyperpolarization (57.92%) (Fig. 2e). Moreover, we further studied the cumulative effect of the sonomagnetic stimulation on membrane hyperpolarization from 010 min. We found that longer exposure time resulted in a greater hyperpolarization of cell membrane potential (Fig. 2f).
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Cell-type-dependent membrane hyperpolarization The membrane electrophysiology of different cell types varies greatly (e.g., resting membrane potential and ion channel type). Even within a single cell, the distribution of ion channels on cell membrane is heterogeneous (Schram 2002; Nusser 2009). Therefore, it is of great interest to explore membrane potential responses of different types of cells after sonomagnetic stimulation. In this work, which focused on stimulating the resting membrane potential, we used non-excitable cells.
Fig. 3. Cell-type-dependent cell membrane hyperpolarization. Sonomagnetic stimulation resulted in the decrease in DiBAC4(3) fluorescence intensity of neuroblast-like cells (a), endothelial-like cells (b) and fibroblast-like cells (c). (d, e) Quantitative analysis showing cell-type-dependent membrane potential responses observed under ultrasound pressures of 1.1 and 2.2 MPa. Data (mean with SEM) were calculated from three independent experiments (n = 150). The image contrast was increased by 10%. *p < 0.05. 3T3 = mouse embryo cells; BEND3 = mouse endothelioma cells; Ctrl = sham exposure; DC2.4 = mouse dendritic cells; DiBAC4(3) = Bis-(1,3-Dibutylbarbituric Acid) Trimethine Oxonol; Elike = endothelial-like; F = Fluorescence intensity; F-like = fibroblast-like; HELA = human cervical cancer cells; M = magnetic stimulation; MRC5 = human lung cells; M + US = sonomagnetic stimulation; N2a = neuro-2a; Nlike = neuroblast-like; PC12 = rat pheochromocytoma cells; SEM = standard error of the mean; US = ultrasonic stimulation.
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Fig. 4. Mitochondrial membrane hyperpolarization. (ac) Fluorescence images of TMRE-loaded cells after sham-exposure, sonomagnetic stimulation at 1.1 MPa ultrasound pressure and sonomagnetic stimulation at 2.2 MPa ultrasound pressure, respectively. (d) Quantitative analysis of the TMRE fluorescence intensities measured after sham exposure (Ctrl), magnetic stimulation (M), ultrasonic stimulation (US) and sonomagnetic stimulation (US + M). Data (mean with SEM) were calculated from three independent experiments (n = 150). The image contrast was increased by 10%. ***p < 0.001. Ctrl = sham exposure; F = Fluorescence intensity; M = magnetic stimulation; M + US = sonomagnetic stimulation; SEM = standard error of the mean; TMRE = tetramethylrhodamine, ethyl ester; US = ultrasonic stimulation.
The membrane potential responses of neuroblast-like cells (DC2.4, N2a and PC12), endothelial-like cells (BEND3 and HeLa) and fibroblast-like cells (MRC5 and 3T3) are shown in Figure 3ac, respectively. We observed a decrease in fluorescence intensity and membrane hyperpolarization in these cells, which have different morphologies. In particular, the neuroblast-like cells exhibited a significantly higher degree of membrane hyperpolarization than that of the endothelial-like cells when stimulated sonomagnetically at 1.1 MPa ultrasound pressure (Fig. 3d). At higher ultrasound pressure (2.2 MPa), the response of DC2.4 cells was significantly greater than that of the other types of cells. Mitochondrial membrane hyperpolarization The mitochondrial DCm plays a vital role in driving protons, ions and proteins to maintain healthy mitochondrial functions (Zorova et al. 2018). To explore whether sonomagnetic stimulation alters the DCm of live cells, we measured the mitochondrial DCm using the voltageresponsive probe TMRE. In Figure 4a, the red, granular
fluorescence indicated a normal mitochondrial potential. After sonomagnetic stimulation, the fluorescence intensity of the probe decreased, indicating a hyperpolarized state of DCm (Fig. 4b, 4c). Quantitative analysis (Fig. 4d) shows that ultrasonic stimulation induced a slight decrease of TMRE fluorescence, whereas sonomagnetic stimulation resulted in a substantial decrease of fluorescence. In line with the response of the cell membrane potential, sonomagnetic stimulation with higher ultrasound pressure (2.2 Pa) induced a greater mitochondrial membrane hyperpolarization. Increase in Ca2+ and NO concentration Ca2+ serves as a second messenger for cell signaling, and its intracellular concentration is affected by the cell membrane potential (Krasznai et al. 2000). Therefore, we examined the Ca2+ response of N2a cells after sonomagnetic stimulation. As shown in Figure 5a, before external stimulation, the green fluorescence of Fluo-4 was weak because the concentration of normal cytoplasmic Ca2+ was relatively low (150250 nM). After
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sonomagnetic stimulation of 1.1 MPa ultrasound pressure, the fluorescence intensity of Fluo-4 increased, indicating the elevation of cytoplasmic Ca2+ concentration. We note that at higher ultrasound pressure (2.2 MPa), the intracellular Ca2+ concentration further increased. This increase in intracellular Ca2+ concentration after sonomagnetic stimulation is quantified in Figure 5d. It is worth noting that confined Ca2+ regions in the form of bright spots were observed under sonomagnetic stimulation at both 1.1 and 2.2 MPa (indicated by yellow arrows in Fig. 5b, 5c). Because the intracellular production of NO is closely correlated with cytoplasmic Ca2+ signaling (Li et al. 2003), we investigated the influence of sonomagnetic stimulation on NO messenger production. As shown in Figure 6ac, the intracellular NO concentration of N2a cells was monitored with the green fluorescence probe DAF-FM diacetate. An increase in fluorescence intensity was observed after sonomagnetic stimulation. In particular, in both of the sonomagnetic groups (Fig. 6b, 6c; 1.1
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and 2.2 MPa, respectively), the localized NO concentration increased, as was evident from the bright spots indicated by the yellow arrows. Quantitative analysis in Figure 6d confirmed that sonomagnetic stimulation significantly elevated the intracellular production of NO. Intracellular acidification The intracellular pH plays an important role in modulating cell function and changes depending on the cell membrane potential (Schackmann et al. 1981). Because we confirmed that sonomagnetic stimulation can induce cell membrane hyperpolarization, we examined whether the intracellular pH was also affected by external physical stimulation. As shown in Figure 7ac, the increased fluorescence intensity of the pHrodo probe indicates that the intracellular pH of the exposed N2a cells decreased (i.e., the concentration of cytoplasmic H+ increased). Quantitative measurements showed that the pH shift was relatively small (i.e., fluorescence intensity
Fig. 5. Increase in intracellular Ca2+ concentration. (ac) Fluorescence images of Fluo-4-loaded cells after sham-exposure, sonomagnetic stimulation at 1.1 MPa ultrasound pressure and sonomagnetic stimulation at 2.2 MPa ultrasound pressure, respectively. (d) Quantitative analysis of the Fluo-4 fluorescence intensities measured after sonomagnetic stimulation at different ultrasound pressures (1.1 and 2.2 MPa). Data (mean with SEM) were calculated from three independent experiments (n = 150). The image contrast was increased by 10%. ***p < 0.001. Ca2+ = calcium; Ctrl = sham exposure; F = Fluorescence intensity; M = magnetic stimulation; M + US = sonomagnetic stimulation; SEM = standard error of the mean; US = ultrasonic stimulation.
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Fig. 6. Increase in intracellular NO concentration. (ac) Fluorescence images of DAF-FM-loaded cells after shamexposure, sonomagnetic stimulation at 1.1 MPa ultrasound pressure and sonomagnetic stimulation at 2.2 MPa ultrasound pressure, respectively. (d) Quantitative analysis of the DAF-FM fluorescence intensities measured after sonomagnetic stimulation at different ultrasound pressures (1.1 and 2.2 MPa). Data (mean with SEM) were calculated from three independent experiments (n = 150). The image contrast was increased by 10%. ***p < 0.001. Ctrl = sham exposure; DAFFM = to 4-Amino-5-Methylamino-20 ,70 -Difluorofluorescein; F = Fluorescence intensity; M = magnetic stimulation; M + US = sonomagnetic stimulation; NO = nitric oxide; SEM = standard error of the mean; US = ultrasonic stimulation.
increase of approximately 35% and pH decrease of approximately 0.3) (Fig. 7d). Increase in intracellular ROS level Hyperpolarization of the mitochondrial membrane induces chemical production of intracellular ROS. Having confirmed DCm hyperpolarization, we then investigated the intracellular level of ROS after sonomagnetic stimulation. The ROS levels of N2a cells were monitored with a green fluorescent H2DCF-DA probe. As shown in Figure 8ac, the stimulated cells exhibited increased fluorescence intensities and cytoplasmic ROS levels. Quantitative analysis in Figure 8d confirmed that ROS elevation in the sonomagnetic group was significantly higher than that of the sham-exposed control group.
proliferation (Roderick and Cook 2008; Blackiston et al. 2009). Therefore, we examined whether daily sonomagnetic stimulation (1.1 MHz frequency, 2.2 MPa ultrasound pressure, 680 mT-strength magnetic field and 10 min exposure time) affects the growth of N2a cancer cells. As shown in Figure 9a, the cells were imaged at 0, 24, 48 and 72 h after the first sonomagnetic stimulation. We found that the cells in the sonomagnetic group grew more slowly than those in sham-exposed group. Quantitative analysis confirmed this observation by showing that the proliferation rate of cells in the sonomagnetic group was inhibited by 48.64% at 72 h after the first stimulation (Fig. 9d).
DISCUSSION Inhibition of proliferation of N2a cells We found that sonomagnetic stimulation resulted in cell membrane hyperpolarization and intracellular NO production. In the literature, these two cellular responses are functionally correlated with the inhibition of cell
Modulation of cellular electrical and chemical signals by ultrasound is attractive for technical reasons because ultrasound can be non-invasively focused into deep tissues with good spatial resolution. When using ultrasound alone, high acoustic energy levels are
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Fig. 7. Intracellular acidification. (ac) Fluorescence images of pHrodo-loaded cells after sham-exposure, sonomagnetic stimulation at 1.1 MPa ultrasound pressure and sonomagnetic stimulation at 2.2 MPa ultrasound pressure, respectively. (d) Quantitative analysis of the pHrodo fluorescence intensities measured after sonomagnetic stimulation at different ultrasound pressures (1.1 and 2.2 MPa). Data (mean with SEM) were calculated from three independent experiments (n = 150). The image contrast was increased by 10%. ***p < 0.001. Ctrl = sham exposure; F = Fluorescence intensity; M = magnetic stimulation; M + US = sonomagnetic stimulation; SEM = standard error of the mean; US = ultrasonic stimulation.
required to evoke sonoelectrical and sonochemical effects (Honda et al. 2004). To decrease the acoustic energy to safe levels, ultrasound contrast agents (e.g., microbubble) and ultrasound-sensitive chemicals (e.g., 5-aminolevulinic acid) can be used (Qin et al. 2014; Sun et al. 2015). In this work, we proposed a new method using ultrasound and a static magnetic field to evoke electrophysiologic and biochemical responses of live cells. Our results demonstrated that the sonomagnetic stimulation method (1.1 MHz frequency, 1.1 or 2.2 MPa acoustic pressure, 50 cycles per pulse, 500 Hz pulse repetition frequency and 680 mT strength magnetic field) induced cell and mitochondrial membrane hyperpolarization and also increased the production of intracellular biochemical signaling species (Ca2+, NO, H+ and ROS). We note that, according to eqn (8), the same evoked cellular effects could be achieved using lower ultrasound energy levels if we increase the magnetic field strength accordingly. In 2017, Rekhi and Arbabian (2017) used a 2 T strength magnetic field for generating Lorentz current. If we increased the magnetic field strength from 680 mT to 2 T, the ultrasound pressure could be
theoretically decreased from 2.20.748 MPa. In this way, the calculated spatial-peak time-averaged intensity of the ultrasound wave is 463 mW/cm2, which is smaller than the FDA-approved safety level of 720 mW/cm2 for diagnostic ultrasound equipment (Nelson et al. 2009). The electrophysiologic and biochemical responses induced in this study can affect other cell behavior (e.g., proliferation). For example, it has been reported that hyperpolarization of the cell membrane turns the cell into a quiescent state with slow proliferation rate, whereas depolarization drives cells into an active state whereby they grow more quickly (Levin 2007). A low concentration of cytoplasmic NO promotes cell proliferation, whereas a higher concentration of NO induces cell cycle arrest and inhibits cell growth (Napoli et al. 2013). Moreover, when the intracellular pH is decreased, the proliferation rate of cancer cells has been shown to be inhibited (Che et al. 2008). Although the roles of mitochondrial DCm, Ca2+ and ROS concentrations in cell proliferation are still unclear, these signals are highly correlated with cell cycle progression and proliferation rate (Pinto et al. 2015; Diebold and Chandel 2016). With
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Fig. 8. Increase in intracellular ROS level. (ac) Fluorescence images of H2DCF-DA-loaded cells after sham-exposure, sonomagnetic stimulation at 1.1 MPa ultrasound pressure and sonomagnetic stimulation at 2.2 MPa ultrasound pressure, respectively. (d) Quantitative analysis of the H2DCF-DA fluorescence intensities measured after sonomagnetic stimulation at different ultrasound pressures (1.1 and 2.2 MPa). Data (mean with SEM) were calculated from three independent experiments (n = 150). The image contrast was increased by 10%. ***p < 0.001. Ctrl = sham exposure; F = Fluorescence intensity; H2DCF-DA = 20 ,70 -Dichlorodihydrofluorescein; diacetate F = Fluorescence intensity; M = magnetic stimulation; M + US = sonomagnetic stimulation; ROS = reactive oxygen species; SEM = standard error of the mean; US = ultrasonic stimulation.
our results demonstrating that sonomagnetic stimulation can induce cell membrane hyperpolarization and modulate the production of signaling molecules (NO and H+), we further examined the proliferation of the stimulated cells. We found that the proliferation of these cells was inhibited, demonstrating the potential of the sonomagnetic stimulation method in controlling cancer cell proliferation. In future work, two critical questions associated with sonomagnetic stimulation need further investigation. The first question is how the scale of cell membrane hyperpolarization is linked to the cell type. In the present study, we found that cells of different type responded to sonomagnetic stimulation differently. Such cell-type dependence has been previously reported in a sonoporation study (Shi et al. 2017). Exploration of this question is of particular interest because this cell-type-dependent behavior could be used for cell diagnosis and imaging (Kunyansky et al. 2017). The second question is how to precisely control the cellular responses of live cells evoked by sonomagnetic stimulation. It is known that even for the same
physical stimulation method, different parameters can lead to different biologic responses. For example, highfrequency (>5 Hz) TMS excites neurons, whereas at low frequency (0.21 Hz), it inhibits the electrical activity of neurons (Hallett 2007). Therefore, the ultrasound parameters of sonomagnetic stimulation (e.g., frequency, pulse duration and pressure) need to be better controlled to achieve precise modulation of electrophysiologic, biochemical and behavioral responses. The proposed sonomagnetic stimulation method holds great promise to non-invasively modulate the in vivo signaling of cells at a spatial resolution of approximately 12 mm depending on the focusing ability of the ultrasound wave. However, the clinical application of this method was limited by the technical difficulty in the generation of large-scale (e.g., 45 £ 45 £ 45 cm) homogenous magnetic field with high strength. In this study, the length of the gap between the two NdFeB magnets where the cell samples were placed was 13 mm. In the work carried out by Rekhi and Arbabian (2017), the length of the gap between magnets was 5 mm. Note
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Fig. 9. Inhibition of proliferation of N2a cells. (a) Images of N2a cells at 0, 24, 48 and 72 h after the first sonomagnetic stimulation. (b) Quantitative analysis of the proliferation rate of N2a cells after sham exposure (Ctrl), magnetic stimulation (M), ultrasonic stimulation (US) and sonomagnetic stimulation (US + M). For each experiment, five imaging areas were selected for cell number counting. Data (mean with SEM) were calculated from three independent experiments (n = 150). Ctrl = sham exposure; F = Fluorescence intensity; M = magnetic stimulation; M + US = sonomagnetic stimulation; N2a = Neuro-2a; SEM = standard error of the mean; US = ultrasonic stimulation.
that the increase in the gap length will decrease magnetic field strength. Therefore, it is not practical to use permanent magnets for sonomagnetic stimulation of cells at significant tissue depths. We suggest that electromagnets made from coils of superconducting wire that have been implemented for magnetic resonance imaging can be employed for the clinical investigation of sonomagnetic stimulation (Shen et al. 2017).
can be technically enhanced by increasing the magnetic field strength.
CONCLUSION
REFERENCES
In this study, we demonstrated the feasibility and effectiveness of sonomagnetic stimulation in evoking electrophysiologic, biochemical and behavioral responses of live cells. Our results suggest that this method could potentially be applied to modulate cell and mitochondrial membrane potentials, to trigger the production of intracellular signaling species and to inhibit the growth of cancer cells. The advantages of this physical stimulation method are that its spatial energy distribution can be effectively confined inside the ultrasound focal region and its stimulation effects
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Acknowledgments—This work was funded by the National Natural Science Foundation of China (Grant No. 61427806), the Shenzhen Science and Technology Planning Project (Grant No. JCYJ20160520170055193), the Medical Project of Guangdong Province (A2019542) and the Natural Science Foundation of Shenzhen University (Grant No.000218). Conflict of interest—The authors declare no other potential conflicts of interest.
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