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Investigation of Hypoxia-Induced Myocardial Injury Dynamics in a Tissue Interface Mimicking Microfluidic Device Li Ren,†,§ Wenming Liu,†,§ Yaolei Wang,†,§ Jian-Chun Wang,† Qin Tu,† Juan Xu,† Rui Liu,† Shao-Fei Shen,† and Jinyi Wang*,†,‡ †

Colleges of Veterinary Medicine and Science and ‡Shaanxi Key Laboratory of Molecular Biology for Agriculture, Northwest A&F University, Yangling, Shaanxi 712100, People’s Republic of China S Supporting Information *

ABSTRACT: Myocardial infarction is a major cause of morbidity and mortality worldwide. However, the methodological development of a spatiotemporally controllable investigation of the damage events in myocardial infarction remains challengeable. In the present study, we describe a micropillar array-aided tissue interface mimicking microfluidic device for the dynamic study of hypoxia-induced myocardial injury in a microenvironment-controllable manner. The mass distribution in the device was visually characterized, calculated, and systematically evaluated using the micropillar-assisted biomimetic interface, physiologically relevant flows, and multitype transportation. The fluidic microenvironment in the specifically functional chamber for cell positioning and analysis was successfully constructed with high fluidic relevance to the myocardial tissue. We also performed a microenvironment-controlled microfluidic cultivation of myocardial cells with high viability and regular structure integration. Using the well-established culture device with a tissue-mimicking microenvironment, a further on-chip investigation of hypoxia-induced myocardial injury was carried out and the varying apoptotic responses of myocardial cells were temporally monitored and measured. The results show that the hypoxia directionally resulted in observable cell shrinkage, disintegration of the cytoskeleton, loss of mitochondrial membrane potential, and obvious activation of caspase-3, which indicates its significant apoptosis effect on myocardial cells. We believe this microfluidic device can be suitable for temporal investigations of cell activities and responses in myocardial infarction. It is also potentially valuable to the microcontrol development of tissue-simulated studies of multiple clinical organ/tissue disease dynamics.

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concerns.7,8 However, either the real-time detection of the fickle process in vivo or the reliable spatiotemporal construction of extracellular signal dynamics in vitro has been one of the long-standing challenges in cardiac science. Accordingly, the methodological development of an effective investigation of specific myocardial infarction events (e.g., myocardial hypoxia) remains largely out of reach. Recently, microengineering technologies have presented a series of striking capabilities in conducting spatial and temporal manipulations of both mammalian cells and their microenvironments with high microscale resolution.9−11 For example, microfabrication and micropattern methods have been employed to perform controllable orientation and organization of myocardial cells.12,13 Microfluidics, a microfabrication technique stemming from the field of microelectromechanical systems, is a promising platform for cell biology because of its excellent performance in the precise control of fluid perfusion and biological sample localization as well as in keeping the organized cells in a tissue-relevant

yocardial infarction, a common presentation of coronary artery disease, arises from the life-threatening interruption of blood supply to a part of the heart.1 The resulting ischemia and ensuing oxygen shortage, if left untreated for a sufficient period of time, can cause damage or even death of the cardiac muscle tissue.2 Although therapeutic options such as reperfusion have greatly improved this situation over the past 25 years, more than 7 million people worldwide are estimated to have myocardial infarction each year.3 Cardiomyocyte injury still occurs even if reperfusion therapy is carried out as soon as possible after the onset of ischemia/hypoxia.4 Therefore, the meticulous exploration of the pathogenesis (e.g., formation and rupture of a vulnerable atherosclerotic plaque) and the development (e.g., hypoxia-managed myocardial injury and apoptosis) of the myocardial infarction process has been the subject of much interest and extensive research.3,5 On the basis of the conventional in vivo (e.g., mice model) and in vitro (e.g., plate-based cell culture assay) models, the study on myocardial infarction has largely enhanced the histopathological identification of tissue composition and its spatial organization to date.6 The in vivo model characteristically presents high tissue relevance, and the in vitro model shows a cost-effective operation and avoids ethical or legal © 2012 American Chemical Society

Received: September 6, 2012 Accepted: December 3, 2012 Published: December 4, 2012 235

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Figure 1. Structural and functional design of the microfluidic device. (A) Optical image of the actual device. The dotted circles mark the inlets and outlets of the device (1, the central inlet; 2, the central outlet; 3, the side inlet; and 4, the side outlet). (B) Design and spatial arrangement of the microfluidic components. The micropillar arrays were specifically set in this study for the interface reconstitution between the blood vessel and myocardial tissue. The inserted images are the sectional (left) and planar (right) presentation of the micropillar arrays. (C) Schematic diagrams of the microfluidic device for studying controllable myocardial hypoxia and for the myocardial fluidic microenvironment mimicking.

context.14−17 Combined with the feasible sequential manipulation in a microfluidic device, these advantages effectively extend microfluidic application to cardiac research, ranging from mechanical activity18,19 to electrophysiology,18,20,21 cell metabolism and communication,21,22 and cytotoxicity.23 Lin et al. measured the isometric contraction force in a single cardiomyocyte by clamping the myocardial cell using freestanding microfabricated clamps and measuring the deflections of the microbeams.20 Zhao et al. cultured cardiomyocytes in a micropillar-substrate, and the cellular contractile forces were recorded in situ by observing the deformation of the micropillars.12 Parker’s group reported the design of a “heart on a chip” that exploits muscular thin film technology, biohybrid constructs of an engineered, anisotropic ventricular myocardium on an elastomeric thin film, to measure contractility.18,24,25 These studies greatly improved the microscale evaluation of myocardial tissue/cell properties, especially the studies on the physiological functions of myocardial cells. Nevertheless, the reproduction of the mechanical and functional features of a complex myocardial microenvironment for the study of tissue-simulated cardiac diseases is still being explored, and it requires a systematic consideration of the multifactor microenvironment construction.26 To the best of our knowledge, the pathological dynamics investigation of particular myocardial infarction processes through microfluidics has been less advanced. Here, we describe a study of hypoxia-induced myocardial injury dynamics in a tissue interface mimicking microfluidic device. In the device, we designed and fabricated two sets of interdigitated−castellated micropillars that decouple a central cell culture chamber from the two side perfusion channels. Using these microstructures, the capillary blood flow− endothelial cell−myocardial tissue structure was reconstructed in the device. Furthermore, the physiological fluidic micro-

environment and the hypoperfusion/hypoxia condition during myocardial infarction were mimicked based on the micropillaraided interface. Hereafter, the hypoxia-induced myocardial injury dynamics, namely, cell viability, cytoskeleton, membrane potential of mitochondria, and caspase-3 activation, were recorded and quantified at a single-cell resolution in a precisely controlled device.

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EXPERIMENTAL SECTION Materials and Reagents. RTV 615 polydimethylsiloxane (PDMS) prepolymer and curing agent were purchased from Momentive Performance Materials (Waterford, NY). The surface-oxidized silicon wafers were obtained from Shanghai Xiangjing Electronic Technology Ltd. (Shanghai, China). The AZ 50XT photoresist and developer were bought from AZ Electronic Materials (Somerville, NJ). The SU-8 2005 photoresist and developer were purchased from Microchem (Newton, MA). Collagen-I, acridine orange (AO), propidium iodide (PI), Hoechst 33258, and carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP) were bought from Sigma-Aldrich (MO). The cell culture medium, fetal bovine serum (FBS), trypsin, and TRITC-phalloidin were obtained from Gibco Invitrogen Corporation (CA). The DEVD-NucView 488 Caspase-3 assay kit and the JC-1 mitochondrial membrane potential detection kit were purchased from Biotium, Inc. (Hayward, CA). All solvents and other chemicals were purchased from local commercial suppliers and were of analytical reagent grade, unless otherwise stated. All solutions were prepared using ultrapurified water supplied by a Milli-Q system (Millipore). Device Design and Fabrication. The microfluidic device used in this study was fabricated using soft lithography with PDMS.27,28 We designed patterns of the device using the AutoCAD software. Generally, the microfluidic device is 236

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atmosphere of 5% CO2 at 37 °C. The cells were normally passaged at a ratio of 1:2 every 3 days to maintain them in the exponential growth phase. When the cells reached confluence, they were harvested through trypsinization with 0.25% trypsin in phosphate buffered solution (PBS, 0.01 M, pH 7.4) at 37 °C. Trypsinization was stopped by adding freshly supplemented DMEM. The cell suspension was centrifuged at 1000 rpm for 3 min. The cells were then resuspended in supplemented DMEM for further use. Cell Seeding and Hypoxia Treatment. The microfluidic device was first sterilized with UV light for 2 h and then coated with collagen-I (200 μg/mL) for another 2 h.30 After rinsing thrice with DMEM, the cells were introduced from the central inlet into the cell culture chambers by flowing the suspension (4 μL/min) for 3 min at a cell density of 7.0 × 106 cells/mL. The device was then placed in a humidified atmosphere with 5% CO2 at 37 °C for 1 h to enable the cells to attach. The supplemented DMEM medium was loaded from the two adjacent side inlets into the device to supply the culture nutrients of the cells. For the hypoxia treatment of the cultured heart myocardium H9c2 cells, FCCP solution (50 μM in FBS-free DMEM) was pumped into one of the adjacent side channels at 1 μL/min. Fresh supplemented DMEM medium was simultaneously introduced into the other side channel at the same flow rate. The treating flow was maintained for at least 2 h.4 Cell Staining. Cell viability assessment was performed using the AO/PI staining protocol.28 After removing the growth medium and washing with PBS, the AO/PI staining solution (10 μg/mL each in PBS) was introduced from the central inlet into the cell culture chamber at 2 μL/min, and the staining process was performed for 10 min at room temperature. Then, PBS was introduced for 10 min as a final rinse. For a clear visualization of the cytoskeleton, the actin filament staining of H9c2 cells was also performed.31 Briefly, the cells were fixed using 4% paraformaldehyde for 10 min at room temperature after washing thrice with PBS. The cultures were permeabilized with PBS containing 0.2% Triton X-100 for 30 min. Then, the cultures were incubated at 37 °C for 20 min with TRITC-phalloidin (100 nM in PBS). The cultures were also incubated for 10 min in PBS containing Hoechst dye (H33258 fluorochrome, 0.5 μg/mL) for nuclear staining. Mitochondrial Membrane Potential and Caspase-3 Activity of H9c2 Cells. The mitochondrial membrane potential and caspase-3 activity of H9c2 cells were analyzed in the current study to evaluate the hypoxic injury dynamics of myocardial cells. Briefly, the solution of 5,5′,6,6′-tetrachloro1,1′,3,3′-tetraethylbenzimidazolylcarbocyanine iodide (JC-1, Biotium; 5 μM) was loaded into the cell culture chamber at a flow rate of 2 μL/min and incubated for 15 min at room temperature before PBS rinsing and hypoxia treatment. The analysis of caspase-3 activation of H9c2 cells was performed using NucView 488 caspase-3 substrate (1 μM, a cell membrane-permeable fluorogenic caspase substrate) with the same procedure as above, except the 30 min incubation time. All steps were performed in a biologically safe cabinet to maintain the sterility of all reagents. Caution was used in handling all human biological material. Microscopy and Image Analysis. Bright-field and fluorescence images were timely acquired using an inverted microscope (Olympus, CKX41) equipped with a CCD camera (Olympus, DP72) and a mercury lamp (Olympus, U-RFLT50). Software Image-Pro Plus 6.0 (Media Cyternetics) and SPSS

composed of four parallel functional units (Figure 1). Each unit has a cell culture chamber (550 μm wide and 30 μm high), two side channels (adjacent to the cell culture chamber; 200 μm wide and 30 μm high), and two groups of optimized micropillar arrays (20 μm in diameter, 4 μm in space, and 5 μm in height) each located between the cell culture chamber and the side channels (for the optimization and analysis of the micropillar design in the device, see the Supporting Information). Two types of inlets and outlets were set up including a central inlet, four side inlets, four central outlets, and eight side outlets. Thus, four microfluidic units shared the same central inlet, and each two adjacent units shared the same side inlet. Next, the patterns of the device were printed at 20 000 dots per inch on glass substrates (MicroCAD Photomask Ltd., Suzhou, China) and used as a photomask. Then, the mold was produced by a photolithographic process to create the microchamber, microchannels, and micropillar arrays. A 5 μm-thick negative photoresist (SU-8 2005) was spin-coated onto a silicon wafer to prepare the mold for the micropillar arrays. After ultraviolet (UV) light exposure, the microfluidic components on the wafer were developed using a SU-8 developer. The mold for the fabrication of the other microfluidic components was made by introducing a 30 μm-thick positive photoresist (AZ 50XT) pattern on the same silicon wafer. Before fabricating the microfluidic device, the mold was exposed to trimethylchlorosilane vapor for 3 min. A well-mixed PDMS prepolymer (RTV 615 A and B in 10:1 ratio) was poured onto the mold and placed in a Petri dish to yield a 3 mm-thick layer. The thick layer was cured in an 80 °C oven for 60 min. After incubation, the PDMS layer was peeled off the mold, and holes were introduced for sample access and waste exclusion. The PDMS layer was then trimmed, cleaned, and placed on top of a glass slide (3000 rpm, 45 s, ramp 15 s) coated with PDMS prepolymer (GE RTV 615 A and B in 15:1 ratio) that had been cured for 10 min in the oven (80 °C). The microfluidic device was ready for use after baking at 80 °C for 48 h. On-Chip Mass Transportation and Distribution. In this study, on-chip mass transportation and distribution were comprehensively evaluated based on the physiologically relevant microflow to mimic and reconstitute the fluidic microenvironment of myocardial tissue in the microfluidic device. The channels and the chamber in the device were treated with collagen-I (200 μg/mL, 2 h) similar to the microfluidic cell culture pretreatment to improve their hydrophilic condition and the biologically relevant microenvironment.29,30 Fluorescein (100 μM fluorescein in NaHCO3 buffer, pH 8.3) was used in the present study as a model cue. It was introduced from one inlet into the side channel using a syringe pump (LSP01-1A, Longer Pump) to visually characterize the solute distribution dynamics in the device. Fresh NaHCO3 buffer was simultaneously injected into the other side channel. Various flow rates ranging from 0.2 μL/min to 20 μL/ min were applied here. Each perfusion lasted for at least 15 min, and each experiment was repeated at least three times. Finite element analysis was conducted using the ESI-CFD software (V2010.0, ESI CFD Inc., Huntsville, AL) to evaluate further the flow profile and solute distribution in the device. Cell Culture. Rat heart myocardium H9c2 cells were obtained from the Chinese Academy of Sciences (Shanghai, China). The cells were cultured using Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% FBS, 100 U/ mL penicillin, and 100 μg/mL streptomycin in a humidified 237

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Figure 2. Mass transportation and distribution in the opening condition in the device using fluorescence visualization and a series of biologically relevant flows (0.1−10 μL/min in the side channels). (A) Observable mass distribution at different flow rates. The central inlet and outlet are opened. (B) Quantitative concentration distribution at the same flow rate. (C) Quantitative concentration distribution at different flow rates. (D) Simulated concentration distribution in the device at 0.5 μL/min in the two side channels. (E) Simulated flow condition in the device at 0.5 μL/min in the two side channels.

(Figure 1C) that is capable of maintaining the rapid mass transportation and substantially weakening the negative shear influence of blood flow to the parenchymal cells.32 On the basis of the biological truth that the velocity of blood in capillaries (typically from 0.15 to 8.6 mm/s) directly relates to the volume of blood flow and thus characterizes the degree of local tissue perfusion,34 we selected a series of flow rates ranging from 0.1 to 10 μL/min in the side channel, corresponding to the velocity of 0.28−27.8 mm/s used for mass transportation evaluation in the device. Two parallel flows with changeable ratios of velocity between each other were introduced into the two adjacent side channels. Aided by the fluorescence visualization, the temporal imaging smoothly presented the concentration dynamics of the solutes.35 For convenient monitoring of the mass transportation and distribution, the fluorescein solution was only loaded into one side channel, and fluorescein-free solution was loaded into the other side channel during the study of mass transportation and distribution (Figure 2A). For the ratio of 1:1 (i.e., the same flow rate was used for the two parallel flows similar to the physiological blood flows in capillary networks penetrating through the local area of the

12.0 (SPSS Inc.) were used to perform image analysis and statistical data analysis, respectively. The results, including the error bars in the graphs, were given as the mean ± standard deviation.

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RESULTS AND DISCUSSION Mass Transportation and Distribution in the Device. The extracellular microenvironment plays a critical role in determining cell function and fate.32 The cardiovascular system delivers the necessary substances such as oxygen and other nutrients to myocardial cells and passes metabolic wastes away through extracellular microenvironment by fluid movement to maintain the normal survival, proliferation, and metabolism of myocardial cells.33 A well-prepared attempt at a systematical investigation of the controllable flow and mass distribution was performed in the present study using the designed device (Figure 1A) to reconstitute the fluidic microenvironment of myocardial tissue. Micropillar arrays (Figure 1B) were specifically set in the device between the side channel and the cell culture chamber for the effective mimicking of the capillary−myocardial tissue interface 238

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Figure 3. On-chip cultivation and structural identification of myocardial cells. (A) Optical image of the cultured H9c2 cells in the chamber. (B) Fluorescence image of H9c2 cells using AO/PI double staining for cell viability assessment (green for live cells and red for dead cells). (C) Fluorescence image of actin filament (red) and nuclear (blue) staining. The dotted line corresponds to Figure 4E. (D) Enlarged image of actin filament (red) and nuclear (blue) staining.

parenchymal organs),36 the result shows that a concentration gradient was produced in the cell culture chamber (Figure 2B). Moreover, both the increasing concentration and the broadening gradient were positively related to the increasing flow. However, the variable concentrations in the chamber during the perfusion at those flow rates were completely lower than that in the fluorescein-passed side channel, indicating that it is difficult to realize the total mass transportation from the side channel to the chamber. The theoretical calculations also indicate that the data on the concentration gradient were similar to the actual experimental results (Figure 2A,D). In this study, the intentional opening of the central inlet and outlet was performed to improve further the tissue simulation in the mass distribution test above. This consideration is based on the fact that interstitial flow characteristics and diffusive/convective transport powered by both hydrostatic pressure and osmotic pressure guarantee the physiological function of myocardial tissue.37 The previous microfluidic approaches, which commonly generate a gradient through diffusion at an absolutely closed state,38 did not involve all these cues, particularly the interstitial flow, which is present in all vascularized tissues in vivo as an important microcirculatory component to complete a rapid and sufficient mass transportation for cells. In the present study, the relatively open condition in the cell chamber facilitated the slow flow formation and permitted the diffusive passing. For example, while the flow rate in the side channel is 0.5 μL/min (i.e., 1.39 mm/s), the average velocity of flow within the micropillar arrays and in the chamber is 13.4 and 7.7 μm/s, respectively (Figure 2E). These flow results using the open condition (especially for the flows with lower flow rates such as 0.5 and 0.1 μL/min) are close to the well-known fluidic condition in the extracellular microenvironment in vivo including the myocardial microenvironment (typically 1 μm/ s).37 Compared with the closed state (i.e., the central inlet and outlet were blocked), the open condition caused a faster and

higher concentration gradient formation in the cell chamber (Figure 2B and the Supporting Information, Figures S2−S5), which implies more sufficient nutrient and signal distribution for myocardial cells. These results suggest that the physiologically fluidic microenvironment of the myocardial tissue accompanied by multitype mass transportations can be mimicked in the micropillar array-containing device. To evaluate further the mass transportation and distribution under different flow conditions, two flows with different flow rates (i.e., the ratio ≠ 1) were used for the fluidic microenvironment-controllable investigation of fluidic dynamics and mass distribution in myocardial infarction-induced ischemia. The use of two different flow rates is based on the fact that the inhibition or blockade of blood flow in the upstream of the capillary can result in a significant change in the mass transportation in the downstream condition around the myocardial cells.3 For convenient manipulation and relevance to myocardial infarction, the flow rate in one side channel was maintained constantly, hypothetically corresponding to the normal blood flow. The flow rate in the other side channel was decreased, hypothetically corresponding to the pathological blood flow. In the following text, the two side channels were, respectively, designated as normal channel (N channel) and pathological channel (P channel). The results show that the fluorescence-visualized concentration distribution in the chamber shifted back toward the P channel (Figure 2C and Supporting Information, Figures S6 and S7), which is mainly caused by the increase in fluidic pressure difference between the two side channels. This result implies that the amount of growth/interactive cues around myocardial cells adjacent to the pathological capillary declines, and the insufficient blood flow possibly leads to the subsequent pathological changes in this zone. We also tried to close the P channel to explore further the quantitative dynamics of mass distribution in the blood-blocked condition during myocardial infarction. As the result shows 239

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Figure 4. Microfluidic hypoxia-induced myocardial cell injury. (A) Fluorescence image of cytoskeletal presentation after 2-h hypoxia treatment. The dotted line corresponds to part D. (B) Changes in myocardial cell size during hypoxia treatment. (C) Cytoskeletal organization after hypoxia treatment in different hypoxia zones. (D) Fluorescence-based fluctuation analysis for cell morphology and cytoskeletal integration after the controlled hypoxia treatment. (E) Fluorescence-based fluctuation analysis of the common cultured myocardial cells (Figure 3C) as a blank control for the evaluation of cell morphology and cytoskeletal integration.

condition for 1 h for firm adherence. The cell culture medium was then transported from the side channels into the cell chamber. The medium flow rate of 0.5 μL/min (i.e., 1.39 mm/ s) in the side channels was applied in this test, specifically referring to the actual blood flow state in the capillary around the myocardial tissue (commonly 1 mm/s).39 The nutrient distribution in the chamber during the perfusion cultivation was visualized dynamically (Supporting Information, Figure S9). The liquid environment in the chamber presented an approximate 40% original concentration of solute after the mass transportation. The concentration profile in the chamber appeared as two similar narrow gradients, in which the maximum concentration appeared near the side channel. Thus, a uniform and continual medium supply can be produced and maintained to ensure cell growth and function. During the medium perfusion-based culture, myocardial cells presented complete spreading and adherence as well as proliferation (Figure 3 and the Supporting Information, Figure S10). The viability of the myocardial cells is measured by AO/ PI double staining (Figure 3B) and is as high as 99.6%. One of the primary structural proteins of cardiac muscle (actin) was routinely selected and visualized by using TRITC-phalloidin

(Supporting Information, Figures S6 and S7), nearly no visual mass transmission occurred from the P channel into the chamber. Thus, although the low interstitial flow transports the supplies, the myocardial cells in the pathological zone would hardly survive because of the inadequate feeding and communication. Overall, the results suggest that the physiological and pathological fluidic dynamics of extracellular microenvironments in the myocardial tissue can be constructed using the micropillar array-aided device in a precisely controllable and highly tissue-mimicking manner. On-Chip Culture of Myocardial Cells in the Established Microenvironment. For the precise control and quantified investigation of the myocardial infarction event in the microfluidic device, we prepared an on-chip myocardial cell culture using the well-mimicked physiological fluidic microenvironment above. Cardiomyocytes, like all tissues in the body, rely on ample blood supply to deliver oxygen and nutrients and to remove wastes such as carbon dioxide. The coronary arteries and their branches fulfill this function.33,39 In the present study, rat heart myocardium H9c2 cells were seeded from the central inlet into the cell culture chamber (Supporting Information, Figure S8) and were left at a static 240

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Figure 5. Quantitative investigation of hypoxia-induced mitochondrial membrane potential dynamics. (A) Fluorescence image of JC-1 aggregates in mitochondria before hypoxia treatment. (B) Fluorescence image of JC-1 aggregates in mitochondria after 15-min hypoxia treatment. (C) Fluorescence image of JC-1 monomers in cytoplasm before hypoxia treatment. (D) Fluorescence image of JC-1 monomers in cytoplasm after 15-min hypoxia treatment. (E) Ratio of JC-1 aggregates to its monomers before hypoxia treatment using fluorescence visualization for mitochondrial membrane potential assessment in the hypoxia treatment. The result corresponds to the dotted lines in parts A and C. (F) Ratio of red fluorescence intensity over green fluorescence intensity after 15-min hypoxia treatment. The result corresponds to the dotted lines in parts B and D.

directly construct a hypoxia microenvironment by nonsupply of oxygen to myocardial cells. Under the condition of decreased supply of oxygen, there is not enough or no oxygen to involve the respiratory chain, and glycolysis becomes the main cellular metabolism and energy production pattern.43,44 As a commonly used chemical hypoxia reagent, FCCP can uncouple the mitochondrial respiratory chain, so oxygen could not involve the respiratory chain; the oxidative phosphorylation is inhibited and the energy production is dependent on glycolysis.43 This is very similar with the real hypoxia condition and prevails during myocardial disease.43,44 Therefore, FCCP (50 μM) was chosen in this study to produce in situ a spatially controllable hypoxia condition. Initially, the myocardial cells were routinely cultivated with high viability in the cell chamber for 12 h by nutrient feeding perfusion (0.5 μL/min) through the two side channels. Then, the medium in the P channel was replaced by the FCCPcontaining medium without FBS to create a less nourishing condition that closely simulates a serious myocardial ischemia/ hypoxia in myocardial infarction. According to the mass distribution assay (Figure 2A), there was an invisible hypoxia gradient with high concentration adjacent to the P channel. The first observable cell response from the imaging results was cell shrinkage (Figure 4A,B), which was characterized as a sizedecreasing gradient, i.e., the higher the concentration of the FCCP, the smaller is the cell size. These results positively correlate with the experimental results using the conventional plate-based culture method (Supporting Information, Figures S11−S13). Further, cell shrinkage became more serious along with the 2 h of hypoxia treatment and broader in the lower

staining to visualize intracellular architecture after the culture. The fluorescence images (Figure 3C,D) show that actin proteins seem to self-assemble into long polymers to build networks and bundles of filaments, which have been demonstrated to provide cellular mechanical integrity and contribute to intracellular transport.40 The cell nucleus generally appeared in a normal oval shape. These results demonstrate that the microfluidic device can complete a tissue interface-mimicked myocardial cell culture in a well-controlled microenvironment, which can be useful in a wide range of culture-based myocardial cell assays. Hypoxia-Induced Myocardial Injury Dynamics and Quantitative Assay. Myocardial ischemia has been demonstrated to always result in myocardial hypoxia, especially in myocardial infarction.41 In the present study, the hypoxiainduced myocardial injury dynamics was studied in the established tissue-simulated microfluidic myocardial device to investigate the response of myocardial cells in a hypoxia condition. PDMS, an optically transparent elastomer, has been widely used in biological microfluidics because of its easy fabrication, low cost, practical scalability, optical transparency, and gas permeability. Despite the many advantages of PDMSbased microfluidics, the gas permeability of PDMS hinders the creation of the hypoxia condition, particularly the hypoxia efficiency, if only the manipulation of flow blockage in the P channel is performed. Recently, two hypoxia models in vitro were used for the study of myocardial injury. One is the airproof capsule-based physical hypoxia model.42 The other one is oxygen consumption blocking reagent (e.g., FCCP) -assisted chemical hypoxia model.43 The physical hypoxia model can 241

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Figure 6. Caspase-3 activation of myocardial cells in a controlled hypoxia condition. (A) Fluorescence image of the caspase-3+ cells in the chamber before hypoxia treatment. (B) Fluorescence image of the caspase-3+ cells in the chamber after 2-h hypoxia treatment. The arrows with numbers correspond to part C. (C) Dynamic fluorescence intensity of the caspase-3+ cells during the hypoxia-induced caspase-3 activation. (D) The percentage of the caspase-3+ cells at different hypoxia treating times (i.e., 0, 15, 30, 60, 90, or 120 min) and at different regions of the chamber.

depolarization and membrane potential alteration, which appeared in the early stages of apoptosis. Generally, JC-1 exists in two forms, namely, aggregate and monomer.47,48 After uptake, JC-1 aggregates in the healthy mitochondria with high membrane potential and emits red fluorescence at 590 nm. On the contrary, JC-1 is monomeric in the membrane-damaged mitochondria with low membrane potential, sequentially leaking into the cytoplasm. More importantly, these monomers emit green fluorescence at 527 nm, and the variability can indirectly show the dynamics of the membrane potential.47,48 For a more accurate assessment, the fluorescence ratio between the two types of JC-1 forms, as a substitute to the simple value of fluorescence intensity, was used to reflect the quantitative change in the membrane potential.48 The result shows that the ratio significantly decreased after the introduction of FCCP (Figure 5). The gradual fading of visible red was observed from the high FCCP zone to the low FCCP zone. Comparatively, the green fluorescence in the chamber was maintained. The results suggest that hypoxia treatment induced the decrease in the membrane potential of the mitochondria in the myocardial

FCCP treated zone (Figure 4B). The cell morphological change implies that sustained hypoxia can induce an aggressive injury in myocardial cells. The injured cells become spatially independent, and they have no contact with each other, especially in the high FCCP gradient area (Figure 4). Alternatively, the results from the stain-based evaluation indicate that the actin filaments in the injured cells underwent a disintegration process (Figure 4C,D). The long actin bundles, which were stained a deep red, were damaged and disassembled, leading to the destruction of the cytoskeletal structure of myocardial cells by hypoxia.45 For a direct and reasonable confirmation of the hypoxia injury and the possible starting of apoptosis in myocardial cells, we measured the membrane potential of the mitochondria, which are the primary energy-generating systems in most eukaryotic cells and the key sites of oxidative phosphorylation and ATP generation.46 Mitochondria also participate in various cellular events such as intermediary metabolism, calcium signaling, and apoptosis.46 A common cationic dye (JC-1) was used in this test for the detection of mitochondrial 242

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cells, indicating the progress of mitochondrial dysfunction and the start of the cell death process.47 Caspases are a family of cysteine proteases that play essential roles in apoptosis; when activated, they trigger the apoptotic process of cells.49 In the final phase of this study, the activation signal of caspase-3 was monitored using the NucView 488 Caspase-3 substrate.50 The substrate consists of a fluorogenic DNA dye and a DEVD substrate moiety specific for caspase-3, which are both nonfluorescent and nonfunctional as a DNA dye. After cleavage by caspase-3, a high-affinity DNA dye was released and stained the nucleus bright green. During microfluidic hypoxia dynamics, the caspase-3-positive cells (caspase-3+ cells) were quantified, and the results (Figure 6) show that the percentage of the caspase-3+ cells increased during the hypoxia treatment. In detail, the percentage of the caspase-3+ cells near to the P channel (higher hypoxia) was 18.0 ± 5.5% after 30 min of incubation and was 33.0 ± 2.6% after 60 min of incubation. Conversely, the percentage of the caspase-3+ cells near to the N channel (lower hypoxia) was only near 2.0% after incubation of 30 and 60 min. This increase became more serious with the continuous hypoxia treatment. Therefore, the controlled hypoxia caused the gradient distribution of the caspase-3+ cells in the chamber. Many caspase-3-negative cells remained in the normoxic zone of the chamber during the hypoxia treatment. Therefore, the regional hypoxia gradient in the myocardial tissue due to the myocardial ischemia results in significant apoptotic responses from the myocardial cells in both the morphology and the organelle and functional protein operation. These responses lead to cell death and myocardial infarction at the local district of the heart.

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CONCLUSIONS

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ASSOCIATED CONTENT

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AUTHOR INFORMATION

Corresponding Author

*Phone: + 86-29-870 825 20. Fax: + 86-29-870 825 20. E-mail: [email protected]. Author Contributions §

These three authors contributed equally to this work.

Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS

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REFERENCES

This study was supported by the National Natural Science Foundation of China (Grants 21175107, 20975082, and 207750 59), the Ministry of Education of the People’s Republic of China (Grant NCET-08-602 0464), the Scientific Research Foundation for the Returned Overseas Chinese Scholars, the State Education Ministry, and the Northwest A&F University.

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In this study, we presented a micropillar array-aided tissue interface mimicking microfluidic device and an experimental demonstration of its application in studying hypoxia-induced myocardial cell injury with precise microfluidic control. The results show that the physiological and pathological fluidic conditions of the extracellular microenvironments in the myocardial tissue can be successfully reconstructed, monitored, and measured. Using the well-established microenvironment in the microfluidic device, the myocardial cells can be sufficiently fed and cultured with high viability and a well-organized structure. More importantly, we demonstrated that the study of the hypoxia-induced myocardial injury dynamics could be successfully performed at a single cell resolution in the precisely controlled device. We consider this approach will facilitate the microfluidic device development in a wide range of myocardial infarction-related applications such as myocardial metabolism assessment, cardiac marker isolation and analysis, and electrical signal monitoring. The microenvironment-controlled achievement is potentially useful in the tissue-mimicking exploration of various cell investigations in many biological events such as liver toxicity and failure, inflammation, and myocardial infarction.

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