Bioelectromagnetics 36:1^9 (2015)
Pulsed Magnetic Field Accelerate Proliferation and Migration of Cardiac Microvascular Endothelial Cells Fei Li,1 YuanYuan,1 Ying Guo,1 Nan Liu,1 Da Jing,2 HaichangWang,1 and Wenyi Guo1* 1
Department of Cardiology, Xijing Hospital, Fourth Military Medical University, Xi’an, China 2 Faculty of Biomedical Engineering, Fourth Military Medical University, Xi’an, China
Heart failure is a disease with multifactorial causes. Recently it was established that reduction in vascular density promoted the progression from adaptive cardiac hypertrophy to heart failure, therefore, therapeutic angiogenesis may be a promising method for treating heart failure. Cardiac microvascular endothelial cells (CMECs) play a major role in cardiac angiogenesis. In the present study, we investigated the direct and indirect effect of pulsed magnetic field (PMF) on the proliferation and migration of CMECs. CMECs were isolated from adult Sprague–Dawley (SD) rat hearts. We found PMF with a frequency of 15 Hz and an intensity of 1.8 mT accelerated the proliferation and migration of CMECs and cardiac myocytes (CMs). Moreover, CMECs treated with PMF released 1.5-fold higher vascular endothelial growth factor (VEGF) and 2-fold higher fibroblast growth factor-2 (FGF-2) when compared with PMF-free cells. In addition, CMs treated with PMF released twofold higher FGF-2 compared with PMF-free cells, but there was no change in VEGF levels. Those results suggested PMF has both a direct autocrine mitogenic and an indirect paracrine effect on CMECs proliferation and migration, and the effect of PMF on intercellular communication between CMECs and CMs was partially dependent on FGF-2, but independent on VEGF. Bioelectromagnetics 36:1–9, 2015. © 2014 Wiley Periodicals, Inc. Key words: pulsed magnetic field; cardiac microvascular endothelial cells; cardiac myocytes; angiogenesis
INTRODUCTION Heart failure, a leading cause of mortality worldwide, is a final common consequence of various heart diseases [Neubauer, 2007; Krum and Teerlink, 2011]. Pathological cardiac hypertrophy is a cause of failing myocardium. It was previously reported that physiological hypertrophy was associated with increased numbers of myocardial capillaries, whereas pathological hypertrophy was correlated with a reduction in capillary density [Rakusan et al., 1992]. Recently, convincing evidence has shown that an imbalance between myocyte growth and coronary angiogenesis plays a crucial role in the processes of pathological cardiac hypertrophy [Shiojima et al., 2005]. Disruption of coordinated tissue growth and angiogenesis in the heart contributes to the progression from adaptive cardiac hypertrophy to heart failure, while, supplementation of angiogenic factors during progression from adaptive to maladaptive cardiac hypertrophy preserves cardiac function [Walsh and Shiojima, 2007; Serpi et al., 2011; Higashikuni et al., 2012]. Thus, promoting angiogenesis in hypertrophic myocardium becomes a new therapeutic target of heart failure. 2014 Wiley Periodicals, Inc.
It was shown that PMF is a safe and effective means of treating nonunion bone fractures [Bassett et al., 1974a,b]. Tepper et al. [2004] attributed the beneficial role of PMF to promoting angiogenesis by the direct effect on endothelial cells through the release of fibroblast growth factor b-2 (FGF-2). Recently, Hopper et al. [2009] found conditioned media from osteoblast cells stimulated with PMF increased endoGrant sponsors: National Natural Science Foundation of China; grant number: 30600580; Foundation of Xijing Hospital; grant number: XJZT13M17. Fei Li and Yuan Yuan contributed equally to this work. *Correspondence to: Wenyi Guo, Department of Cardiology, Xijing Hospital, Fourth Military Medical University, Xi’an, China. E-mail: [email protected] Received for review 20 November 2012; Accepted 17 July 2014 DOI: 10.1002/bem.21875 Published online 22 October 2014 in Wiley Online Library (wileyonlinelibrary.com).
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Li et al.
Fig.1. A:Schematic representation of the PMF stimulationwaveform, which consisted of a pulsed burst repeated at 15 Hz. B: Picture of coils and matched perspex holder, which fits inside the coil inincubator.Thermometer wasput by thesideof 24-wellcellculture dishonthestand.
thelial cell proliferation, which suggested PMF promoted angiogenesis by manipulation of the interaction between osteoblast and endothelial cells. Given this data, it may be reasonable for PMF to become a potentially therapeutic mean for pathological myocardial hypertrophy and ischemic myocardial disease. However, one problem that cannot be ignored is the endothelial cells heterogeneity. Cardiac microvascular endothelial cells (CMECs) are a protagonist in myocardial angiogenesis and have very different structure and functions from human umbilical vein endothelial cells (HUVEC). Actually, CMECs are not the only protagonist; abundant evidence has shown cardiac myocytes (CMs) also play a crucial role in myocardial angiogenesis [Levy et al., 1995]. Recently, we demonstrated that pulse-burst magnetic field (PMF) could augment angiogenesis, with associated improvement in ventricular function and reduced infarct size in Sprague–Dawley (SD) rats [Yuan et al., 2010]. To our knowledge, this is the first study to suggest the potential application of PMF in therapeutic myocardial angiogenesis. To better explore the role and mechanism of PMF on myocardial angiogenesis, we investigated the effects of PMF on proliferation, migration of CMECs and the influence of PMF on interaction between cardiomyocytes and CMECs. MATERIALS AND METHODS
with 1.0 mm diameter coil with 197 turns in total, winding a cylindrical Plexiglas frame [17 cm in diameter and 20 cm in length (Fig. 1)]. The PMF waveform produced by the generator consisted of a pulsed burst (burst width, 4.4 ms; pulse width, 0.2 ms; pulse wait, 0.02 ms; burst wait, 62 ms; pulse rise and fall times, 0.3 and 2.0 ms) repeated at 15 Hz. Similar PMF waveforms have been widely employed and proven effective in the prevention and treatment of a range of clinical diseases by many studies, such as osteoporosis and cardiovascular diseases [Luo et al., 2007; Jing et al., 2010, 2011]. The cell culture dish was placed in the center of the solenoid coil to ensure that cells were confined to the center of the electromagnetic fields where the flux density was considered uniform. A resistor with 1 V was placed in series with the coils to record the voltage drop across the resistor with an oscilloscope (Agilent Technologies, Santa Clara, CA). The peak-to-peak magnetic field of the coils was determined to be 0– 18 mT. The background electromagnetic field was determined using a gaussmeter (Lakeshore, Westerville, OH) after the pulse generator was switched off. In this study, all solenoid coils were placed in an incubator, and a Yuyue thermometer (Yancheng, Jiangsu, China) was placed in each coil to detect the temperature in both experimental and control solenoid coils. The measured temperature in experimental (15 Hz, 1.8 mT PMF) and control (0 Hz, 0 mT PMF) solenoid coils in incubator was 38.0 0.2 8C and 37.0 0.2 8C, respectively.
PMF Equipment The PMF generator system consisted of a signal generator and cylindrical solenoid coil. The solenoid coil was constituted by enameled coated copper wire
Cell Culture Cardiac microvascular endothelial cells were isolated from adult SD rat hearts as previously described [Nishida et al., 1993]. Briefly, hearts were
Bioelectromagnetics
Enhancement of Cardiac Microvascular Cells by PMF
removed from adult male SD rats (80–100 g) under sterile conditions. After removing the atria, visible connective tissue, right ventricle and valvular tissue, the left ventricle was immersed in 75% ethanol for 10 s to devitalize epicardial and endocardial endothelial cells. After mincing and digesting the tissue [0.02% collagenase type II (Invitrogen, Carlsbad, CA) for 10 min and 0.025% trypsin (Invitrogen) for 10 min at 37 8C in a shaking bath], the solution was filtered through a 100 mm nylon mesh to remove undigested tissue. The dissociated cells were resuspended in Dulbecco’s modified Eagle’s medium (DMEM, Invitrogen) supplemented with 10% fetal bovine serum (FBS, Invitrogen) and then were seeded on cell culture dishes. Cultured CMECs were tested for their purity using morphological and functional characteristics: (1) the phenotypic profile showed the CMECs displayed a uniform “cobblestone” morphology; (2) positive immunofluorescence assay of Von Willebrand factor (vWF; Abcam, Cambridge, England) and factor VIII (Abcam) were demonstrated positive; (3) uptake of acetylated low-density lipoprotein. SD rat neonatal ventricular myocytes were isolated as described previously [Li et al., 2008] and cultured in DMEM supplemented with 10% FBS, which was changed to serum-free medium after 48 h. Cells were cultured under serum-free condition for 24 h before being used. The investigation conformed to the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH Publication No. 85–23, revised 1996, Bethesda, MD) and it was approved by our Institutional Ethics Review Committee. Cell Viability and Apoptosis The 3rd passage CMECs and primary CMs were used to measure cell viability and apoptosis. Confluent cells were cultured under serum free condition for 24 h before being used. After exposure to PMF for 24 h, cell viability and apoptosis were evaluated. Cell viability was evaluated by trypan blue staining and the fraction of blue cells was quantified by light microscopy. In each dish, at least 200 cells were counted in 10 random fields. To assess the rate of cell apoptosis, apoptosis was quantified by annexin-V-FITC and propidium iodide (PI) double staining using an Annexin-V/FITC kit (Antigene, Wuhan, China). Cells were collected according to the manufacturer’s instructions 48 h after transfection, washed with cold PBS, suspended in binding buffer, and then the cells were incubated 30 min in the dark at 4 8C with Annexin V-FITC and PI in phosphate buffer and analyzed on the flow cytometer (BD Bioscience, Franklin Lakes, NJ) 1 h after staining.
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Endothelial Cells Growth Curve To plot growth curve, 3rd passage CMECs were used; 104/ml CMECs were cultivated in a 24 well 15.6 mm plate (Greiner Bio-One, Frickenhausen, Germany). After exposure to PMF for 2 h/day, cell numbers were counted from 1st to 6th day using a counting chamber (Bürker-Türk, Lauda-Königshofen, Germany). Measurement of Cell Proliferation by CCK-8 The influence of PMF on the CMECs proliferation was measured by Cell Counting Kit-8 (CCK-8; Beyotime, Shanghai, China) according to the manufacturer’s instructions. Briefly, CMECs were cultured in 96-well plates and exposed to PMF 2 h/day for 5 days. After treatment, 10 ml of CCK-8 reagent was added to each well. The absorbance measured at 450 nm after incubation at 37 8C for 1 h. All experiments were repeated three times and eight wells were used per experiment. Scratch Wound-Healing Test Confluent endothelial cells were exposed to PMF for 2 h/day for 2 days. The cells were wounded by scraping with a 200 ml pipette tip, denuding a strip of the monolayer 0.3 mm in diameter. The variation in the wound diameter within three experiments was
Pulsed Magnetic Field Accelerate Proliferation and Migration of Cardiac Microvascular Endothelial Cells Fei Li,1 YuanYuan,1 Ying Guo,1 Nan Liu,1 Da Jing,2 HaichangWang,1 and Wenyi Guo1* 1
Department of Cardiology, Xijing Hospital, Fourth Military Medical University, Xi’an, China 2 Faculty of Biomedical Engineering, Fourth Military Medical University, Xi’an, China
Heart failure is a disease with multifactorial causes. Recently it was established that reduction in vascular density promoted the progression from adaptive cardiac hypertrophy to heart failure, therefore, therapeutic angiogenesis may be a promising method for treating heart failure. Cardiac microvascular endothelial cells (CMECs) play a major role in cardiac angiogenesis. In the present study, we investigated the direct and indirect effect of pulsed magnetic field (PMF) on the proliferation and migration of CMECs. CMECs were isolated from adult Sprague–Dawley (SD) rat hearts. We found PMF with a frequency of 15 Hz and an intensity of 1.8 mT accelerated the proliferation and migration of CMECs and cardiac myocytes (CMs). Moreover, CMECs treated with PMF released 1.5-fold higher vascular endothelial growth factor (VEGF) and 2-fold higher fibroblast growth factor-2 (FGF-2) when compared with PMF-free cells. In addition, CMs treated with PMF released twofold higher FGF-2 compared with PMF-free cells, but there was no change in VEGF levels. Those results suggested PMF has both a direct autocrine mitogenic and an indirect paracrine effect on CMECs proliferation and migration, and the effect of PMF on intercellular communication between CMECs and CMs was partially dependent on FGF-2, but independent on VEGF. Bioelectromagnetics 36:1–9, 2015. © 2014 Wiley Periodicals, Inc. Key words: pulsed magnetic field; cardiac microvascular endothelial cells; cardiac myocytes; angiogenesis
INTRODUCTION Heart failure, a leading cause of mortality worldwide, is a final common consequence of various heart diseases [Neubauer, 2007; Krum and Teerlink, 2011]. Pathological cardiac hypertrophy is a cause of failing myocardium. It was previously reported that physiological hypertrophy was associated with increased numbers of myocardial capillaries, whereas pathological hypertrophy was correlated with a reduction in capillary density [Rakusan et al., 1992]. Recently, convincing evidence has shown that an imbalance between myocyte growth and coronary angiogenesis plays a crucial role in the processes of pathological cardiac hypertrophy [Shiojima et al., 2005]. Disruption of coordinated tissue growth and angiogenesis in the heart contributes to the progression from adaptive cardiac hypertrophy to heart failure, while, supplementation of angiogenic factors during progression from adaptive to maladaptive cardiac hypertrophy preserves cardiac function [Walsh and Shiojima, 2007; Serpi et al., 2011; Higashikuni et al., 2012]. Thus, promoting angiogenesis in hypertrophic myocardium becomes a new therapeutic target of heart failure. 2014 Wiley Periodicals, Inc.
It was shown that PMF is a safe and effective means of treating nonunion bone fractures [Bassett et al., 1974a,b]. Tepper et al. [2004] attributed the beneficial role of PMF to promoting angiogenesis by the direct effect on endothelial cells through the release of fibroblast growth factor b-2 (FGF-2). Recently, Hopper et al. [2009] found conditioned media from osteoblast cells stimulated with PMF increased endoGrant sponsors: National Natural Science Foundation of China; grant number: 30600580; Foundation of Xijing Hospital; grant number: XJZT13M17. Fei Li and Yuan Yuan contributed equally to this work. *Correspondence to: Wenyi Guo, Department of Cardiology, Xijing Hospital, Fourth Military Medical University, Xi’an, China. E-mail: [email protected] Received for review 20 November 2012; Accepted 17 July 2014 DOI: 10.1002/bem.21875 Published online 22 October 2014 in Wiley Online Library (wileyonlinelibrary.com).
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Li et al.
Fig.1. A:Schematic representation of the PMF stimulationwaveform, which consisted of a pulsed burst repeated at 15 Hz. B: Picture of coils and matched perspex holder, which fits inside the coil inincubator.Thermometer wasput by thesideof 24-wellcellculture dishonthestand.
thelial cell proliferation, which suggested PMF promoted angiogenesis by manipulation of the interaction between osteoblast and endothelial cells. Given this data, it may be reasonable for PMF to become a potentially therapeutic mean for pathological myocardial hypertrophy and ischemic myocardial disease. However, one problem that cannot be ignored is the endothelial cells heterogeneity. Cardiac microvascular endothelial cells (CMECs) are a protagonist in myocardial angiogenesis and have very different structure and functions from human umbilical vein endothelial cells (HUVEC). Actually, CMECs are not the only protagonist; abundant evidence has shown cardiac myocytes (CMs) also play a crucial role in myocardial angiogenesis [Levy et al., 1995]. Recently, we demonstrated that pulse-burst magnetic field (PMF) could augment angiogenesis, with associated improvement in ventricular function and reduced infarct size in Sprague–Dawley (SD) rats [Yuan et al., 2010]. To our knowledge, this is the first study to suggest the potential application of PMF in therapeutic myocardial angiogenesis. To better explore the role and mechanism of PMF on myocardial angiogenesis, we investigated the effects of PMF on proliferation, migration of CMECs and the influence of PMF on interaction between cardiomyocytes and CMECs. MATERIALS AND METHODS
with 1.0 mm diameter coil with 197 turns in total, winding a cylindrical Plexiglas frame [17 cm in diameter and 20 cm in length (Fig. 1)]. The PMF waveform produced by the generator consisted of a pulsed burst (burst width, 4.4 ms; pulse width, 0.2 ms; pulse wait, 0.02 ms; burst wait, 62 ms; pulse rise and fall times, 0.3 and 2.0 ms) repeated at 15 Hz. Similar PMF waveforms have been widely employed and proven effective in the prevention and treatment of a range of clinical diseases by many studies, such as osteoporosis and cardiovascular diseases [Luo et al., 2007; Jing et al., 2010, 2011]. The cell culture dish was placed in the center of the solenoid coil to ensure that cells were confined to the center of the electromagnetic fields where the flux density was considered uniform. A resistor with 1 V was placed in series with the coils to record the voltage drop across the resistor with an oscilloscope (Agilent Technologies, Santa Clara, CA). The peak-to-peak magnetic field of the coils was determined to be 0– 18 mT. The background electromagnetic field was determined using a gaussmeter (Lakeshore, Westerville, OH) after the pulse generator was switched off. In this study, all solenoid coils were placed in an incubator, and a Yuyue thermometer (Yancheng, Jiangsu, China) was placed in each coil to detect the temperature in both experimental and control solenoid coils. The measured temperature in experimental (15 Hz, 1.8 mT PMF) and control (0 Hz, 0 mT PMF) solenoid coils in incubator was 38.0 0.2 8C and 37.0 0.2 8C, respectively.
PMF Equipment The PMF generator system consisted of a signal generator and cylindrical solenoid coil. The solenoid coil was constituted by enameled coated copper wire
Cell Culture Cardiac microvascular endothelial cells were isolated from adult SD rat hearts as previously described [Nishida et al., 1993]. Briefly, hearts were
Bioelectromagnetics
Enhancement of Cardiac Microvascular Cells by PMF
removed from adult male SD rats (80–100 g) under sterile conditions. After removing the atria, visible connective tissue, right ventricle and valvular tissue, the left ventricle was immersed in 75% ethanol for 10 s to devitalize epicardial and endocardial endothelial cells. After mincing and digesting the tissue [0.02% collagenase type II (Invitrogen, Carlsbad, CA) for 10 min and 0.025% trypsin (Invitrogen) for 10 min at 37 8C in a shaking bath], the solution was filtered through a 100 mm nylon mesh to remove undigested tissue. The dissociated cells were resuspended in Dulbecco’s modified Eagle’s medium (DMEM, Invitrogen) supplemented with 10% fetal bovine serum (FBS, Invitrogen) and then were seeded on cell culture dishes. Cultured CMECs were tested for their purity using morphological and functional characteristics: (1) the phenotypic profile showed the CMECs displayed a uniform “cobblestone” morphology; (2) positive immunofluorescence assay of Von Willebrand factor (vWF; Abcam, Cambridge, England) and factor VIII (Abcam) were demonstrated positive; (3) uptake of acetylated low-density lipoprotein. SD rat neonatal ventricular myocytes were isolated as described previously [Li et al., 2008] and cultured in DMEM supplemented with 10% FBS, which was changed to serum-free medium after 48 h. Cells were cultured under serum-free condition for 24 h before being used. The investigation conformed to the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH Publication No. 85–23, revised 1996, Bethesda, MD) and it was approved by our Institutional Ethics Review Committee. Cell Viability and Apoptosis The 3rd passage CMECs and primary CMs were used to measure cell viability and apoptosis. Confluent cells were cultured under serum free condition for 24 h before being used. After exposure to PMF for 24 h, cell viability and apoptosis were evaluated. Cell viability was evaluated by trypan blue staining and the fraction of blue cells was quantified by light microscopy. In each dish, at least 200 cells were counted in 10 random fields. To assess the rate of cell apoptosis, apoptosis was quantified by annexin-V-FITC and propidium iodide (PI) double staining using an Annexin-V/FITC kit (Antigene, Wuhan, China). Cells were collected according to the manufacturer’s instructions 48 h after transfection, washed with cold PBS, suspended in binding buffer, and then the cells were incubated 30 min in the dark at 4 8C with Annexin V-FITC and PI in phosphate buffer and analyzed on the flow cytometer (BD Bioscience, Franklin Lakes, NJ) 1 h after staining.
3
Endothelial Cells Growth Curve To plot growth curve, 3rd passage CMECs were used; 104/ml CMECs were cultivated in a 24 well 15.6 mm plate (Greiner Bio-One, Frickenhausen, Germany). After exposure to PMF for 2 h/day, cell numbers were counted from 1st to 6th day using a counting chamber (Bürker-Türk, Lauda-Königshofen, Germany). Measurement of Cell Proliferation by CCK-8 The influence of PMF on the CMECs proliferation was measured by Cell Counting Kit-8 (CCK-8; Beyotime, Shanghai, China) according to the manufacturer’s instructions. Briefly, CMECs were cultured in 96-well plates and exposed to PMF 2 h/day for 5 days. After treatment, 10 ml of CCK-8 reagent was added to each well. The absorbance measured at 450 nm after incubation at 37 8C for 1 h. All experiments were repeated three times and eight wells were used per experiment. Scratch Wound-Healing Test Confluent endothelial cells were exposed to PMF for 2 h/day for 2 days. The cells were wounded by scraping with a 200 ml pipette tip, denuding a strip of the monolayer 0.3 mm in diameter. The variation in the wound diameter within three experiments was
