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Biological studies with tin oxide materials
Biological studies with tin oxide materials
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Jia-Bo Lyau, Hsiang-Chiu Wu, Hsin Chen Department of Electrical Engineering, National Tsing Hua University, Hsinchu, Taiwan Chapter outline 21.1 Introduction 599 21.2 Electrochemical characterization of ITO 600 21.3 Designing transparent microfluidic systems above ITO microelectrodes 602 21.4 A transparent microlab for studying electrophysiology of biological cells 605 21.5 Biological experiments 607 21.5.1 Recording heartbeat signals of zebrafish 607 21.5.2 Biocompatibility test 608 21.5.3 Cell-trapping capability test 609
21.6 Summary 610 References 611
21.1 Introduction The rapidly advancing photolithography and micromachining technologies not only enable us to fabricate miniaturized transistors and sensors for engineering applications, but also underpin the development of innovative devices, such as the labon-a-chip or the micro-total analysis system (μTAS), for biomedical applications. These miniaturized microlabs require much less testing samples and allow different experimental conditions to be carried out in parallel and automatically. Therefore, experiments based on microlabs are more efficient and favorable than conventional experimental approaches. The indium tin oxide (ITO) exhibits great transmittance for visible light, low resistivity (1.408 × 10−3–1.956 × 10−3 Ω cm), and stable electrochemical property [1]. In addition, ITO films have good adhesion to either glass or polyethylene terephthalate PET substrates. These features make ITO a good candidate for forming microelectrodes and microwires in a lab-on-chip device. In many biological experiments, it is crucial to detect the intrinsic charges of specific biomolecules [2, 3], or the electrophysiological signals generated by cells (e.g., neurons) before and after a certain drug or a specific electrical stimulation is applied to these cells. In these experiments, monitoring cell morphology or biomedical reactions in real time is very helpful for experimenters to ensure an experiment is well controlled, as well as to correlate the measured electrophysiological signals to the states of biology essays easily. Driven Tin Oxide Materials: Synthesis, Properties, and Applications. https://doi.org/10.1016/B978-0-12-815924-8.00021-9 © 2020 Elsevier Inc. All rights reserved.
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by this demand, ITO microelectrodes have been widely employed in lab-on-chip systems to facilitate microchip electrophoresis [4], electrochemiluminescence [5, 6], and cancer-cell detection [7]. In addition to the applications above, electrophysiological studies further demand methodologies for immobilizing cells on ITO electrodes or for guiding the growth of cells above ITO electrodes. One possibility is using micro- or nano-imprint technology to coat materials with good cell affinity on specific locations, so as to promote the growth and adhesion of cells to these locations [8, 9]. However, this method is unable to control the number of cells falling on the promoted positions precisely. In case too many cells aggregate at a single electrode, the aggregation would impede cells from distributing evenly on all electrodes [10]. Instead, using the polydimethylsiloxane (PDMS) to fabricate microstructures for confining or guiding the movements of cells is a popular alternative proposed in Refs. [11–15]. Although the fabrication process of the PDMS is simple and low cost, the PDMS exhibits nonideal properties such as pairing, sagging, or shrinking. These properties make it difficult to fabricate complex structures with a high geometrical resolution for guiding or trapping cells smaller than 10 μm in diameter. In addition, the PDMS elastomers may absorb hydrophobic molecules to alter buffer concentrations and subsequently affect experimental outcomes [16]. As the limitations above are unacceptable, the SU-8 photoresist becomes a good alternative for fabricating transparent, multilayer microstructures with high geometrical resolution. This chapter introduces the possibility of integrating ITO microelectrodes with microfluidic systems to form a transparent microlab. The microlab contains microfluidic channels to guide biological cells to stay right above ITO microelectrodes, so that both morphology and electrophysiology of the cells can be studied simultaneously. The content is organized as follows. Following the introduction to potential biological applications and advantages for ITO electrodes, Section 21.2 investigates whether the electrochemical properties of ITO microelectrodes are suitable for electrophysiological experiments. Section 21.3 then proposes a process for fabricating transparent, multilayer, microfluidic channels with SU-8 above an ITO microelectrode array. Afterward, Section 21.4 presents the fabricated microlab in detail, and Section 21.5 presents the feasibility of using the microlab for different biological experiments. The biocompatibility of ITO materials is also examined. Finally, the experimental results are discussed and summarized in Section 21.6.
21.2 Electrochemical characterization of ITO How well ITO electrodes are able to record or to elicit electrophysiological activities of biological cells depend very much on the electrochemical impedance and the charge-injection capacity of the electrode-electrolyte interface of ITO electrodes. The electrochemical impedance spectrum of the interface between ITO microelectrodes and phosphate-buffered saline (PBS) is thus measured with a LCR meter. The charge-injection capacity of ITO microelectrodes is also examined by cyclic voltammetry. These experimental results are presented and discussed below.
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The experimental setup for measuring the electrochemical impedance spectrum is shown in Fig. 21.1A. A 120-nm-thick ITO on a glass substrate is first patterned by photolithography to form circular microelectrodes with a diameter of 1.6 μm. A glass O-ring is then attached to the substrate to form a reservoir for containing the buffer solution above ITO microelectrodes. Subsequently, the electrochemical impedance of the ITO microelectrode is measured by an impedance analyzer (HP 4284A). One end of the analyzer is connected to an ITO microelectrode, and the other end to a silver/silver-chloride electrode immersed in the buffer. The measured impedance spectrum is shown in Fig. 21.1B and compared to that of gold microelectrodes reported in Ref. [18]. The ITO microelectrode exhibits 5–20 times higher impedance than gold microelectrodes over the bandwidth (0.1– 10 kHz) of electrophysiological signals of biological cells [19]. Large electrode impedance could impede the recording of biopotential signals because the signals could be attenuated due to the voltage division between the electrode impedance of ITO and the input impedance of a recording amplifier. Nevertheless, the maximum electrode impedance is 112 kΩ/mm2 for ITO and the input impedance of a recording amplifier is normally greater than 10 MΩ [20]. As long as the area of an ITO microelectrode is large enough to result in an electrode impedance below 1 MΩ, recording biopotential signals through ITO microelectrode is feasible.
Fig. 21.1 (A) The setup for measuring electrode impedance spectrum. (B) the measured electrode impedance spectra of ITO and gold microelectrodes [17].
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Gold microelectrode
Current density (µA/mm2)
Current density (µA/mm2)
1.5 1.0 0.5 0 –0.5
(A)
ITO microelectrode
1.0 0.5 0 –0.5
–0.2
0
0.2
0.4
Potential (Volts vs. Ag/AgCl)
0.6
(B)
–0.6
–0.2
0
0.4
0.8
Potential (Volts vs. Ag/AgCl)
Fig. 21.2 The cyclic voltammetry measurement of (A) gold microelectrodes. (B) ITO microelectrodes.
High electrode impedance is also unfavorable for delivering electrical stimulation currents because high-voltage compliance is required for a faradic current to flow through an electrode with high impedance. For example, if the impedance of an ITO microelectrode is 1 MΩ, a faradic current of 10 μA already results in a voltage drop of 10 V across the electrode-electrolyte interface. To avoid unfeasibly high-voltage compliance, stimulation current can be delivered by capacitive coupling, instead, through the double-layer capacitor at the electrode-electrolyte interface [21]. The ability of a microelectrode to inject charge through capacitive coupling is usually characterized by cyclic voltammetry. In our cyclic voltammetry experiment, a source meter (Keithley 2602) is used to replace the impedance analyzer. One end of the source meter is connected to an ITO electrode, and the other to a silver/silver-chloride electrode immersed in the buffer. The source meter generates a voltage difference across the electrode-electrolyte interface of the ITO electrode and measures the corresponding current. The voltage is swept within the dynamic range of [−0.8 0.8] V without inducing Faradic current, and the voltage changes slowly at a rate of 100 mV/s. The measured cyclic voltammetry curves for a gold and an ITO microelectrode are shown in Fig. 21.2 for comparison. The charge-injection capacity of a microelectrode is proportional to the area within the cyclic voltammetry curve of that microelectrode. Fig. 21.2 indicates that the ITO microelectrode has smaller charge-injection capacity than the gold microelectrode. Nevertheless, the difference in charge-injection capacity is smaller than that in electrode impedance. Inducing stimulation currents by capacitive coupling rather than Faradic mechanism is more feasible for ITO microelectrodes. Certainly, it is also possible to improve the charge-injection capacity of ITO microelectrodes by surface modification [22, 23].
21.3 Designing transparent microfluidic systems above ITO microelectrodes As ITO microelectrodes are employed to record or to elicit electrophysiological activities of biological cells (e.g., neurons), a microstructure able to guide biological cells to stay right on top of ITO microelectrodes is crucial for enhancing the signal-to-noise
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ratio of recorded signals, as well as for maximizing stimulation efficacy. The transparency of ITO further facilitates studying the morphology and the electrophysiology of cells simultaneously. To retain the advantage of transparency, this section introduces the feasibility of using the photoresist SU-8 to build a multilayer microfluidic system above ITO microelectrode arrays. A microlab integrating transparent microfluidic system with ITO microelectrodes has been proposed in Ref. [17]. Fig. 21.3A shows a modified structure for increasing the likelihood to trap cells in the micro-wells above ITO microelectrodes. The microfluidic system consists of three layers, the cell-guiding layer, the cell- trapping layer, and the cell-culturing layer. Fig. 21.3B shows the detailed structure around ITO microelectrodes in the cell-culturing layer, and Fig. 21.4 shows a clearer structure of the proposed device with the three layers assembled. The cell-guiding layer contains three circular holes. The large one (2.5 mm in diameter) connected with microfluidic channels is called the cell reservoir, and the other two are called medium reservoirs. As a physiological buffer containing cells is dripped into the cell reservoir, the cells flow along the microchannels and fall into the micro-well array, as the red arrows indicated in Fig. 21.5. Each micro-well penetrates through the cell-trapping layer and aligns with one ITO microelectrode on the glass substrate (Fig. 21.5B). As shown in Fig. 21.3B, yellow pillars are used to support the cell-trapping layer above, so as
Fig. 21.3 The multilayer microfluidic system made of SU-8. (A) The structure consists of the cell-guiding layer (top), the cell-trapping layer (middle), and the cell-culturing layer (bottom) with an ITO microelectrode array on the glass substrate. (B) The magnified cell-culturing region at the center of the cell-culturing layer.
Fig. 21.4 The assembled multilayer microfluidic system with ITO microelectrodes.
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Cell
Cell
PLL solution
Cell guiding layer
Cell trapping layer
Fluid out Cell
(A)
(B) Fig. 21.5 The cross-sectional view of the microfluidic system for (A) illustrating how cells are trapped above microelectrodes (red arrows) and how culture medium is supplied to maintain cell health (green arrows). (B) illustrating how cells flow from cell reservoir (red arrows) into the cell-trapping holes (purple arrow).
to create a cell-culture region of 10 μm high. This 10-μm high-culturing region allows trapped cells, especially neurons, to grow synaptic connections between each other. In addition, culture media can be supplied into this region, through the two medium reservoirs (Fig. 21.4) to maintain the health of cultured cells. For example, coating poly-l-lysine (PLL) on microelectrodes promotes the adhesion of cells to microelectrodes. The PLL can be supplied to the culture region by flowing from one medium reservoir to the other, as the green arrows indicated in Fig. 21.5A. The cell-trapping layer is designed to be 50 μm thick for separating cell-guiding channels from the cell- culturing layer effectively. Sufficient separation is essential for preventing the culture medium (e.g., PLL) from flowing into the cell-guiding channels in the top layer. This avoids cells from being adhered to the guiding channels and subsequently blocking the flow of other cells. To keep cells staying on top of ITO microelectrodes, cell-trapping fences are further built around each microelectrode, as shown by Fig. 21.3B. Compared to other lab-on-a-chip systems reported in literatures [8–10], the proposed system has the following advantages. First, integrating transparent microfluidic
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system with ITO microelectrodes enables studying electrophysiology and morphology of cells simultaneously. Secondly, the customized, solid microfluidic structure helps to guide and to trap cells more precisely and more reliably above ITO electrodes, as compared to simply coating PLL on electrodes by micro- or nano-imprint technologies [8–10]. Thirdly, the multilayer structure allows media for cell adhesion or cell nutrition to be supplied into the bottom layer directly, avoiding the contamination to the fluids or channels in other layers (e.g., the cell-guiding layer).
21.4 A transparent microlab for studying electrophysiology of biological cells Fig. 21.6 shows the process flow for fabricating the proposed transparent microfluidic system with ITO microelectrodes. The fabrication process consists of five photolithography steps (masks). The positive photoresist, AZ9260, is employed as a sacrificial layer, defining the cell-trapping structure and supporting pillars in the cell-culturing layer. The negative photoresist, SU-8, is the main material for building the whole microfluidic system. After the bottom layer AZ9260 is patterned and developed, two layers of SU-8 are grown and patterned by photolithography. Afterwards, all unwanted SU-8 and the sacrificial layer of AZ9260 are developed to form the microfluidic system. Figs. 21.7 and 21.8 illustrate the photolithography processes more clearly. The first mask defines the locations and patterns of alignment marks made of Au and Ti on the ITO glass.
SU-8 coating
Aliment mark lift-off process (Mask 1)
SU-8 soft bake
AZ9260 patterning (Mask 3)
SU-8 patterning (Mask 4)
Mask 5
ITO electrode etching (Mask 2)
AZ9260 coating
SU-8 post bake AZ9260 development SU-8 development
Fig. 21.6 The process steps for fabricating a transparent, multilayer microfluidic system above ITO microelectrodes.
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UV
(B) (A)
(C)
UV
(D)
Fig. 21.7 The photolithography process for coating and patterning the sacrificial layer made of AZ9260.
Fig. 21.8 The photolithography process for coating and patterning multiple layers of SU-8.
The second mask defines the patterns of microelectrodes, wire connections, and wire-bonding pads made of the ITO layer. As shown in Fig. 21.7A–C, the unwanted ITO is removed by wet etching. Afterwards, the photoresist AZ9260 is coated above the ITO microelectrode, and the third mask is employed to define the structure of cell- trapping fences and pillars in the culture region (Fig. 21.7D). The AZ9260 exposed to UV light is developed as shown in Fig. 21.8A. A 50-μm-thick SU-8 layer is then coated and patterned by the forth mask, defining the cell-trapping wells and medium reservoirs (Fig. 21.8B). The SU-8 also fills into the holes of the AZ9260 to form the pillars and cell-trapping fences. Subsequently, another 50-μm-thick layer of SU-8 is coated and patterned by the fifth mask, defining the cell reservoir and cell-guiding channels on the top layer (Fig. 21.8C). The uncured SU-8 (the region not exposed to UV light) and remaining AZ9260 are removed by the development process for 20 min (Fig. 21.8D). The fabricated microfluidic sample is heated to 300°C and then immersed in cold water. During the cooling process, the microfluidic system can be easily peeled off from the glass substrate due to the difference in thermal expansion coefficients b etween
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Fig. 21.9 The scanning-electron-microscopy (SEM) images of (A) the top view and (B) the bottom view of the fabricated multilayer microfluidic system.
glass and SU-8. Fig. 21.9 shows the SEM images of a fabricated microfluidic system. The cell-guiding layer in Fig. 21.9A has five cell-guiding channels, each of which has five branches leading to the cell-trapping wells. The bottom view of the cell-culturing layer in Fig. 21.9B further reveals that the height of the culture region is actually 7.51 μm in this sample. The height is smaller than our intended design (10 μm) because repetitive baking of the multilayer structure could induce shrinkage of AZ9260. The other cause would be process variations dependent on the rotational speed for coating AZ9260. Nevertheless, a lower culture region actually helps to trap cells in the wells reliably. The fabricated microfluidic system integrated with ITO microelectrodes on the glass substrate forms a transparent microlab for studying electrophysiology and morphology of biological cells simultaneously.
21.5 Biological experiments The biocompatibility of SU-8 is not well studied and not as good as the PDMS, a material proved to be stable, biocompatible, and thus widely used for fabricating microfluidic systems. However, the PDMS exhibits nonideal characteristics, which make it difficult to fabricate sophisticate structures with PDMS with a resolution below a few micrometers [24]. This section first examines the capability of ITO microelectrodes to record biopotential signals. The biocompatibility of SU-8 will then be investigated by culturing hippocampal neurons dissociated from the brain of rat fetus at embryonic day 18. The embryonic neurons are particularly sensitive to the culturing environment and the toxicity of materials. Finally, the ability of the proposed microfluidic system to trap miniature cells such as neurons is tested.
21.5.1 Recording heartbeat signals of zebrafish Fig. 21.10A shows the experimental setup for recording the heartbeat signals from a dissected heart of zebrafish. Adult, wide-type zebrafish with a size of 2–4 cm was
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Fig. 21.10 Experimental results with zebrafish heart [17]. (A) The experimental setup (B) the measured electrocardiogram of zebrafish.
obtained from the Zebrafish Core Facility at Academia Sinica in Taiwan. Dissection of the heart from zebrafish followed the protocol in Ref. [25], and the dissected hear was immersed in phosphate-buffered saline (PBS; pH = 7.4) within the reservoir above ITO microelectrodes. All experimental protocols had been reviewed and approved by the Institutional Animal Care and Use Committee in the National Tsing Hua University, Taiwan. The heartbeat signals of the dissected heart were measured by an electrocardiogram (ECG) recorder (EZ-BIO-01-S1-E, EZ instrument, Taiwan) through the two ITO microelectrodes, denoted as the positive (+) and negative (−) terminals in Fig. 21.10A. The reference terminal of the ECG recorder was connected to the reference electrode in the medium reservoir. The measured ECG signals are shown in Fig. 21.10B. The average heart rate of the zebrafish is 113 beats/min, agreeing with the fact that the heart rate of the zebrafish ranges from 80 to 180 beats/min [26]. This confirms that heartbeat signals can be recorded through ITO microelectrodes reliably. Moreover, Fig. 21.10B shows that the QRS waves of heartbeat signals are also recorded clearly to facilitate further analysis.
21.5.2 Biocompatibility test Rat hippocampal neurons were dissociated from the brain of rat fetus at embryonic day 18 in minimum essential medium. All experimental protocols complied with the provisions of the Animal Care and Use Committee of the National Tsing Hua University, Taiwan. The preparation and culturing procedures of hippocampal neurons were detailed in Ref. [17]. The hippocampal neurons were cultured on a cover glass and the proposed SU-8 microfluidic system for comparison. After 14 days of culturing, fluorescence immunohistochemical stain was used to examine the cytotoxicity. Fig. 21.11 shows the fluorescent images of cultured neurons (A) on the cover glass and (B) on the SU-8 microfluidic system. The blue circles are the cell nucleus of hippocampal neurons, while the green color reveals the cell morphology. Neurons in both cultures exhibit well-retained nucleus and skeleton.
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Fig. 21.11 The fluorescent images of rat hippocampal neurons on (A) the glass substrate and (B) the SU-8 substrate at DIV14 [17].
Fig. 21.12 Average numbers of neurons on the glass coverslip (control) and the SU-8 substrate. p = 0.0249 [17].
This result indicates that neurons on SU-8 substrate exhibit comparable health conditions to neurons on the glass substrate. The fluorescent images are further examined under a microscope. Three regions with an area of 1.5 mm2 are randomly selected from each sample. The number of nucleus in each selected region is counted. Fig. 21.12 shows and compares the average numbers of nucleus found on the SU-8 and the glass substrates. Taking the experiment with the cover glass as a control group, the statistical p-value for the two different groups is only 0.0249. This proves that the biocompatibility of the fabricated SU-8 microfluidic system is comparable to that of the glass substrate.
21.5.3 Cell-trapping capability test The cell-trapping capability of the proposed microfluidic system is further tested with microbeads and hippocampal neurons. Microbeads with a diameter of 10 μm (Micro
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Fig. 21.13 The snapshot of the experiment with microbeads (diameter = 10 μm), which are trapped in the microholes of the microfluidic system.
particles, No. 72986, SIGMA) were mixed with PBS with a concentration of 1.05 g/ cm3. As the PBS with microbeads were dripped into the cell reservoir, the flow in the microfluidic system was monitored and video recorded by a microscope. A snapshot is shown in Fig. 21.13, revealing that the microbeads are guided to enter cell-trapping wells and confined above ITO microelectrodes. The culture medium containing hippocampal neurons is further dripped into the cell reservoir of another microfluidic system. The neurons are also guided into the cell-trapping wells reliably. However, as shown by the snapshots in Fig. 21.14, the red circles highlight the neuron which enters a cell-trapping well for less than 1 s. This nonideal phenomenon comes from the fact that, as fluid continues to flow from the cell reservoir to the culture region, the fluidic pressure forces cells to deform themselves and subsequently to squeeze themselves out of the cell-trapping fences. The cell-trapping structure shown in this chapter should thus be improved further for trapping neurons for a long term. Nevertheless, the experiment with microbeads demonstrates that the proposed structure is able to trap cells (e.g., red blood cells in (JMM)) whose diameter is much larger than 10 μm.
21.6 Summary A transparent microlab consisting of ITO microelectrodes and wires on a glass substrate provides a great opportunity to study both electrophysiology and morphology of cells simultaneously. Although ITO microelectrodes have greater impedance than microelectrodes made of inert metals, the electrode impedance can be reduced by either increasing the physical electrode area or modifying the electrode surface through electrochemistry. The experiment with the zebrafish further demonstrates that ITO microelectrodes are able to record biopotential signals reliably. To facilitate long-term
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Fig. 21.14 The snapshots of the experiment with dissected neurons. The time difference between successive frames is 1–2 s.
study with biological cells, a transparent microfluidic structure can be built with SU-8 for guiding biological cells to stay above ITO microelectrodes. Our experimental results demonstrate the feasibility of using photolithography to construct a multilayer SU-8 microfluidic system, which is not only biocompatible but also capable of trapping biological cells with sophisticated structures. The slight imperfection is that neurons could escape trapping wells when the fluidic pressure is too strong. This could be solved by controlling the flow speed of the solution containing biological cells. Therefore, integrating a transparent microfluidic system with ITO microelectrodes is believed to be a useful platform for drug-screening and neuroscience research.
References [1] Zhang J, Au KH, Zhu ZQ, O’shea S. Sol–gel preparation of poly (ethylene glycol) doped indium tin oxide thin films for sensing applications. Opt Mater 2004;26(1):47–55. [2] Gao Z, Ting BP. A DNA biosensor based on a morpholino oligomer coated indium-tin oxide electrode and a cationic redox polymer. Analyst 2009;134(5):952–7.
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[3] Armistead PM, Thorp HH. Oxidation kinetics of guanine in DNA molecules adsorbed onto indium tin oxide electrodes. Anal Chem 2001;73(3):558–64. [4] Zhu G, Song Q, Liu W, Yan X, Xiao J, Chen C. A gold nanoparticle-modified indium tin oxide microelectrode for in-channel amperometric detection in dual-channel microchip electrophoresis. Anal Methods 2017;9(29):4319–26. [5] Zhao X, You T, Qiu H, Yan J, Yang X, Wang E. Electrochemiluminescence detection with integrated indium tin oxide electrode on electrophoretic microchip for direct bioanalysis of lincomycin in the urine. J Chromatogr B 2004;810(1):137–42. [6] Qiu H, Yan J, Sun X, Liu J, Cao W, Yang X, et al. Microchip capillary electrophoresis with an integrated indium tin oxide electrode-based electrochemiluminescence detector. Anal Chem 2003;75(20):5435–40. [7] Sun D, Lu J, Wang X, Zhang Y, Chen Z. Voltammetric aptamer based detection of HepG2 tumor cells by using an indium tin oxide electrode array and multifunctional nanoprobes. Microchim Acta 2017;184(9):3487–96. [8] Jungblut M, Knoll W, Thielemann C, Pottek M. Triangular neuronal networks on microelectrode arrays: an approach to improve the properties of low-density networks for extracellular recording. Biomed Microdevices 2009;11(6):1269. [9] Scholl M, Sprössler C, Denyer M, Krause M, Nakajima K, Maelicke A, et al. Ordered networks of rat hippocampal neurons attached to silicon oxide surfaces. J Neurosci Methods 2000;104(1):65–75. [10] Jun SB, Hynd MR, Dowell-Mesfin N, Smith KL, Turner JN, Shain W, et al. Low-density neuronal networks cultured using patterned poly-l-lysine on microelectrode arrays. J Neurosci Methods 2007;160(2):317–26. [11] Nam Y, Musick K, Wheeler BC. Application of a PDMS microstencil as a replaceable insulator toward a single-use planar microelectrode array. Biomed Microdevices 2006;8(4):375–81. [12] Kim SM, Lee SH, Suh KY. Cell research with physically modified microfluidic channels: a review. Lab Chip 2008;8(7):1015–23. [13] Rettig JR, Folch A. Large-scale single-cell trapping and imaging using microwell arrays. Anal Chem 2005;77(17):5628–34. [14] Di Carlo D, Aghdam N, Lee LP. Single-cell enzyme concentrations, kinetics, and inhibition analysis using high-density hydrodynamic cell isolation arrays. Anal Chem 2006;78(14):4925–30. [15] Yang M, Li CW, Yang J. Cell docking and on-chip monitoring of cellular reactions with a controlled concentration gradient on a microfluidic device. Anal Chem 2002;74(16):3991–4001. [16] Toepke MW, Beebe DJ. PDMS absorption of small molecules and consequences in microfluidic applications. Lab Chip 2006;6(12):1484–6. [17] Wu HC, Lyau JB, Lin MH, Chuang YJ, Chen H. Multilayer microfluidic systems with indium-tin-oxide microelectrodes for studying biological cells. J Micromech Microeng 2017;27(7):075013. [18] Chen YC, Hsu HL, Lee YT, Su HC, Yen SJ, Chen CH, et al. An active, flexible carbon nanotube microelectrode array for recording electrocorticograms. J Neural Eng 2011;8(3):034001. [19] Webster JG. Medical instrumentation: Application and design. 2nd ed. Boston, MA: Houghton Mifflin; 1992. [20] Yoo HJ, Hoof CV. Bio-medical CMOS ICs (integrated circuits and systems). New York: Springer; 2010. [21] He B. Neural engineering. New York: Springer; 2013.
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[22] Zhang J, Oyama M. Gold nanoparticle-attached ITO as a biocompatible matrix for myoglobin immobilization: direct electrochemistry and catalysis to hydrogen peroxide. J Electroanal Chem 2005;577(2):273–9. [23] Chang G, Oyama M, Hirao K. In situ chemical reductive growth of platinum nanoparticles on indium tin oxide surfaces and their electrochemical applications. J Phys Chem B 2006;110(4):1860–5. [24] Xia Y, Whitesides GM. Soft lithography. Angew Chem Int Ed 1998;37(5):550–75. [25] Hecker L, Khait L, Sessions SK, Birla RK. Functional evaluation of isolated zebrafish hearts. Zebrafish 2008;5(4):319–22. [26] Huang WC, Hsieh YS, Chen IH, Wang CH, Chang HW, Yang CC, et al. Combined use of MS-222 (tricaine) and isoflurane extends anesthesia time and minimizes cardiac rhythm side effects in adult zebrafish. Zebrafish 2010;7(3):297–304.
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Jia-Bo Lyau, Hsiang-Chiu Wu, Hsin Chen Department of Electrical Engineering, National Tsing Hua University, Hsinchu, Taiwan Chapter outline 21.1 Introduction 599 21.2 Electrochemical characterization of ITO 600 21.3 Designing transparent microfluidic systems above ITO microelectrodes 602 21.4 A transparent microlab for studying electrophysiology of biological cells 605 21.5 Biological experiments 607 21.5.1 Recording heartbeat signals of zebrafish 607 21.5.2 Biocompatibility test 608 21.5.3 Cell-trapping capability test 609
21.6 Summary 610 References 611
21.1 Introduction The rapidly advancing photolithography and micromachining technologies not only enable us to fabricate miniaturized transistors and sensors for engineering applications, but also underpin the development of innovative devices, such as the labon-a-chip or the micro-total analysis system (μTAS), for biomedical applications. These miniaturized microlabs require much less testing samples and allow different experimental conditions to be carried out in parallel and automatically. Therefore, experiments based on microlabs are more efficient and favorable than conventional experimental approaches. The indium tin oxide (ITO) exhibits great transmittance for visible light, low resistivity (1.408 × 10−3–1.956 × 10−3 Ω cm), and stable electrochemical property [1]. In addition, ITO films have good adhesion to either glass or polyethylene terephthalate PET substrates. These features make ITO a good candidate for forming microelectrodes and microwires in a lab-on-chip device. In many biological experiments, it is crucial to detect the intrinsic charges of specific biomolecules [2, 3], or the electrophysiological signals generated by cells (e.g., neurons) before and after a certain drug or a specific electrical stimulation is applied to these cells. In these experiments, monitoring cell morphology or biomedical reactions in real time is very helpful for experimenters to ensure an experiment is well controlled, as well as to correlate the measured electrophysiological signals to the states of biology essays easily. Driven Tin Oxide Materials: Synthesis, Properties, and Applications. https://doi.org/10.1016/B978-0-12-815924-8.00021-9 © 2020 Elsevier Inc. All rights reserved.
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by this demand, ITO microelectrodes have been widely employed in lab-on-chip systems to facilitate microchip electrophoresis [4], electrochemiluminescence [5, 6], and cancer-cell detection [7]. In addition to the applications above, electrophysiological studies further demand methodologies for immobilizing cells on ITO electrodes or for guiding the growth of cells above ITO electrodes. One possibility is using micro- or nano-imprint technology to coat materials with good cell affinity on specific locations, so as to promote the growth and adhesion of cells to these locations [8, 9]. However, this method is unable to control the number of cells falling on the promoted positions precisely. In case too many cells aggregate at a single electrode, the aggregation would impede cells from distributing evenly on all electrodes [10]. Instead, using the polydimethylsiloxane (PDMS) to fabricate microstructures for confining or guiding the movements of cells is a popular alternative proposed in Refs. [11–15]. Although the fabrication process of the PDMS is simple and low cost, the PDMS exhibits nonideal properties such as pairing, sagging, or shrinking. These properties make it difficult to fabricate complex structures with a high geometrical resolution for guiding or trapping cells smaller than 10 μm in diameter. In addition, the PDMS elastomers may absorb hydrophobic molecules to alter buffer concentrations and subsequently affect experimental outcomes [16]. As the limitations above are unacceptable, the SU-8 photoresist becomes a good alternative for fabricating transparent, multilayer microstructures with high geometrical resolution. This chapter introduces the possibility of integrating ITO microelectrodes with microfluidic systems to form a transparent microlab. The microlab contains microfluidic channels to guide biological cells to stay right above ITO microelectrodes, so that both morphology and electrophysiology of the cells can be studied simultaneously. The content is organized as follows. Following the introduction to potential biological applications and advantages for ITO electrodes, Section 21.2 investigates whether the electrochemical properties of ITO microelectrodes are suitable for electrophysiological experiments. Section 21.3 then proposes a process for fabricating transparent, multilayer, microfluidic channels with SU-8 above an ITO microelectrode array. Afterward, Section 21.4 presents the fabricated microlab in detail, and Section 21.5 presents the feasibility of using the microlab for different biological experiments. The biocompatibility of ITO materials is also examined. Finally, the experimental results are discussed and summarized in Section 21.6.
21.2 Electrochemical characterization of ITO How well ITO electrodes are able to record or to elicit electrophysiological activities of biological cells depend very much on the electrochemical impedance and the charge-injection capacity of the electrode-electrolyte interface of ITO electrodes. The electrochemical impedance spectrum of the interface between ITO microelectrodes and phosphate-buffered saline (PBS) is thus measured with a LCR meter. The charge-injection capacity of ITO microelectrodes is also examined by cyclic voltammetry. These experimental results are presented and discussed below.
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The experimental setup for measuring the electrochemical impedance spectrum is shown in Fig. 21.1A. A 120-nm-thick ITO on a glass substrate is first patterned by photolithography to form circular microelectrodes with a diameter of 1.6 μm. A glass O-ring is then attached to the substrate to form a reservoir for containing the buffer solution above ITO microelectrodes. Subsequently, the electrochemical impedance of the ITO microelectrode is measured by an impedance analyzer (HP 4284A). One end of the analyzer is connected to an ITO microelectrode, and the other end to a silver/silver-chloride electrode immersed in the buffer. The measured impedance spectrum is shown in Fig. 21.1B and compared to that of gold microelectrodes reported in Ref. [18]. The ITO microelectrode exhibits 5–20 times higher impedance than gold microelectrodes over the bandwidth (0.1– 10 kHz) of electrophysiological signals of biological cells [19]. Large electrode impedance could impede the recording of biopotential signals because the signals could be attenuated due to the voltage division between the electrode impedance of ITO and the input impedance of a recording amplifier. Nevertheless, the maximum electrode impedance is 112 kΩ/mm2 for ITO and the input impedance of a recording amplifier is normally greater than 10 MΩ [20]. As long as the area of an ITO microelectrode is large enough to result in an electrode impedance below 1 MΩ, recording biopotential signals through ITO microelectrode is feasible.
Fig. 21.1 (A) The setup for measuring electrode impedance spectrum. (B) the measured electrode impedance spectra of ITO and gold microelectrodes [17].
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Gold microelectrode
Current density (µA/mm2)
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1.5 1.0 0.5 0 –0.5
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Fig. 21.2 The cyclic voltammetry measurement of (A) gold microelectrodes. (B) ITO microelectrodes.
High electrode impedance is also unfavorable for delivering electrical stimulation currents because high-voltage compliance is required for a faradic current to flow through an electrode with high impedance. For example, if the impedance of an ITO microelectrode is 1 MΩ, a faradic current of 10 μA already results in a voltage drop of 10 V across the electrode-electrolyte interface. To avoid unfeasibly high-voltage compliance, stimulation current can be delivered by capacitive coupling, instead, through the double-layer capacitor at the electrode-electrolyte interface [21]. The ability of a microelectrode to inject charge through capacitive coupling is usually characterized by cyclic voltammetry. In our cyclic voltammetry experiment, a source meter (Keithley 2602) is used to replace the impedance analyzer. One end of the source meter is connected to an ITO electrode, and the other to a silver/silver-chloride electrode immersed in the buffer. The source meter generates a voltage difference across the electrode-electrolyte interface of the ITO electrode and measures the corresponding current. The voltage is swept within the dynamic range of [−0.8 0.8] V without inducing Faradic current, and the voltage changes slowly at a rate of 100 mV/s. The measured cyclic voltammetry curves for a gold and an ITO microelectrode are shown in Fig. 21.2 for comparison. The charge-injection capacity of a microelectrode is proportional to the area within the cyclic voltammetry curve of that microelectrode. Fig. 21.2 indicates that the ITO microelectrode has smaller charge-injection capacity than the gold microelectrode. Nevertheless, the difference in charge-injection capacity is smaller than that in electrode impedance. Inducing stimulation currents by capacitive coupling rather than Faradic mechanism is more feasible for ITO microelectrodes. Certainly, it is also possible to improve the charge-injection capacity of ITO microelectrodes by surface modification [22, 23].
21.3 Designing transparent microfluidic systems above ITO microelectrodes As ITO microelectrodes are employed to record or to elicit electrophysiological activities of biological cells (e.g., neurons), a microstructure able to guide biological cells to stay right on top of ITO microelectrodes is crucial for enhancing the signal-to-noise
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ratio of recorded signals, as well as for maximizing stimulation efficacy. The transparency of ITO further facilitates studying the morphology and the electrophysiology of cells simultaneously. To retain the advantage of transparency, this section introduces the feasibility of using the photoresist SU-8 to build a multilayer microfluidic system above ITO microelectrode arrays. A microlab integrating transparent microfluidic system with ITO microelectrodes has been proposed in Ref. [17]. Fig. 21.3A shows a modified structure for increasing the likelihood to trap cells in the micro-wells above ITO microelectrodes. The microfluidic system consists of three layers, the cell-guiding layer, the cell- trapping layer, and the cell-culturing layer. Fig. 21.3B shows the detailed structure around ITO microelectrodes in the cell-culturing layer, and Fig. 21.4 shows a clearer structure of the proposed device with the three layers assembled. The cell-guiding layer contains three circular holes. The large one (2.5 mm in diameter) connected with microfluidic channels is called the cell reservoir, and the other two are called medium reservoirs. As a physiological buffer containing cells is dripped into the cell reservoir, the cells flow along the microchannels and fall into the micro-well array, as the red arrows indicated in Fig. 21.5. Each micro-well penetrates through the cell-trapping layer and aligns with one ITO microelectrode on the glass substrate (Fig. 21.5B). As shown in Fig. 21.3B, yellow pillars are used to support the cell-trapping layer above, so as
Fig. 21.3 The multilayer microfluidic system made of SU-8. (A) The structure consists of the cell-guiding layer (top), the cell-trapping layer (middle), and the cell-culturing layer (bottom) with an ITO microelectrode array on the glass substrate. (B) The magnified cell-culturing region at the center of the cell-culturing layer.
Fig. 21.4 The assembled multilayer microfluidic system with ITO microelectrodes.
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Cell
Cell
PLL solution
Cell guiding layer
Cell trapping layer
Fluid out Cell
(A)
(B) Fig. 21.5 The cross-sectional view of the microfluidic system for (A) illustrating how cells are trapped above microelectrodes (red arrows) and how culture medium is supplied to maintain cell health (green arrows). (B) illustrating how cells flow from cell reservoir (red arrows) into the cell-trapping holes (purple arrow).
to create a cell-culture region of 10 μm high. This 10-μm high-culturing region allows trapped cells, especially neurons, to grow synaptic connections between each other. In addition, culture media can be supplied into this region, through the two medium reservoirs (Fig. 21.4) to maintain the health of cultured cells. For example, coating poly-l-lysine (PLL) on microelectrodes promotes the adhesion of cells to microelectrodes. The PLL can be supplied to the culture region by flowing from one medium reservoir to the other, as the green arrows indicated in Fig. 21.5A. The cell-trapping layer is designed to be 50 μm thick for separating cell-guiding channels from the cell- culturing layer effectively. Sufficient separation is essential for preventing the culture medium (e.g., PLL) from flowing into the cell-guiding channels in the top layer. This avoids cells from being adhered to the guiding channels and subsequently blocking the flow of other cells. To keep cells staying on top of ITO microelectrodes, cell-trapping fences are further built around each microelectrode, as shown by Fig. 21.3B. Compared to other lab-on-a-chip systems reported in literatures [8–10], the proposed system has the following advantages. First, integrating transparent microfluidic
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system with ITO microelectrodes enables studying electrophysiology and morphology of cells simultaneously. Secondly, the customized, solid microfluidic structure helps to guide and to trap cells more precisely and more reliably above ITO electrodes, as compared to simply coating PLL on electrodes by micro- or nano-imprint technologies [8–10]. Thirdly, the multilayer structure allows media for cell adhesion or cell nutrition to be supplied into the bottom layer directly, avoiding the contamination to the fluids or channels in other layers (e.g., the cell-guiding layer).
21.4 A transparent microlab for studying electrophysiology of biological cells Fig. 21.6 shows the process flow for fabricating the proposed transparent microfluidic system with ITO microelectrodes. The fabrication process consists of five photolithography steps (masks). The positive photoresist, AZ9260, is employed as a sacrificial layer, defining the cell-trapping structure and supporting pillars in the cell-culturing layer. The negative photoresist, SU-8, is the main material for building the whole microfluidic system. After the bottom layer AZ9260 is patterned and developed, two layers of SU-8 are grown and patterned by photolithography. Afterwards, all unwanted SU-8 and the sacrificial layer of AZ9260 are developed to form the microfluidic system. Figs. 21.7 and 21.8 illustrate the photolithography processes more clearly. The first mask defines the locations and patterns of alignment marks made of Au and Ti on the ITO glass.
SU-8 coating
Aliment mark lift-off process (Mask 1)
SU-8 soft bake
AZ9260 patterning (Mask 3)
SU-8 patterning (Mask 4)
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ITO electrode etching (Mask 2)
AZ9260 coating
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Fig. 21.6 The process steps for fabricating a transparent, multilayer microfluidic system above ITO microelectrodes.
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UV
(B) (A)
(C)
UV
(D)
Fig. 21.7 The photolithography process for coating and patterning the sacrificial layer made of AZ9260.
Fig. 21.8 The photolithography process for coating and patterning multiple layers of SU-8.
The second mask defines the patterns of microelectrodes, wire connections, and wire-bonding pads made of the ITO layer. As shown in Fig. 21.7A–C, the unwanted ITO is removed by wet etching. Afterwards, the photoresist AZ9260 is coated above the ITO microelectrode, and the third mask is employed to define the structure of cell- trapping fences and pillars in the culture region (Fig. 21.7D). The AZ9260 exposed to UV light is developed as shown in Fig. 21.8A. A 50-μm-thick SU-8 layer is then coated and patterned by the forth mask, defining the cell-trapping wells and medium reservoirs (Fig. 21.8B). The SU-8 also fills into the holes of the AZ9260 to form the pillars and cell-trapping fences. Subsequently, another 50-μm-thick layer of SU-8 is coated and patterned by the fifth mask, defining the cell reservoir and cell-guiding channels on the top layer (Fig. 21.8C). The uncured SU-8 (the region not exposed to UV light) and remaining AZ9260 are removed by the development process for 20 min (Fig. 21.8D). The fabricated microfluidic sample is heated to 300°C and then immersed in cold water. During the cooling process, the microfluidic system can be easily peeled off from the glass substrate due to the difference in thermal expansion coefficients b etween
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Fig. 21.9 The scanning-electron-microscopy (SEM) images of (A) the top view and (B) the bottom view of the fabricated multilayer microfluidic system.
glass and SU-8. Fig. 21.9 shows the SEM images of a fabricated microfluidic system. The cell-guiding layer in Fig. 21.9A has five cell-guiding channels, each of which has five branches leading to the cell-trapping wells. The bottom view of the cell-culturing layer in Fig. 21.9B further reveals that the height of the culture region is actually 7.51 μm in this sample. The height is smaller than our intended design (10 μm) because repetitive baking of the multilayer structure could induce shrinkage of AZ9260. The other cause would be process variations dependent on the rotational speed for coating AZ9260. Nevertheless, a lower culture region actually helps to trap cells in the wells reliably. The fabricated microfluidic system integrated with ITO microelectrodes on the glass substrate forms a transparent microlab for studying electrophysiology and morphology of biological cells simultaneously.
21.5 Biological experiments The biocompatibility of SU-8 is not well studied and not as good as the PDMS, a material proved to be stable, biocompatible, and thus widely used for fabricating microfluidic systems. However, the PDMS exhibits nonideal characteristics, which make it difficult to fabricate sophisticate structures with PDMS with a resolution below a few micrometers [24]. This section first examines the capability of ITO microelectrodes to record biopotential signals. The biocompatibility of SU-8 will then be investigated by culturing hippocampal neurons dissociated from the brain of rat fetus at embryonic day 18. The embryonic neurons are particularly sensitive to the culturing environment and the toxicity of materials. Finally, the ability of the proposed microfluidic system to trap miniature cells such as neurons is tested.
21.5.1 Recording heartbeat signals of zebrafish Fig. 21.10A shows the experimental setup for recording the heartbeat signals from a dissected heart of zebrafish. Adult, wide-type zebrafish with a size of 2–4 cm was
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Fig. 21.10 Experimental results with zebrafish heart [17]. (A) The experimental setup (B) the measured electrocardiogram of zebrafish.
obtained from the Zebrafish Core Facility at Academia Sinica in Taiwan. Dissection of the heart from zebrafish followed the protocol in Ref. [25], and the dissected hear was immersed in phosphate-buffered saline (PBS; pH = 7.4) within the reservoir above ITO microelectrodes. All experimental protocols had been reviewed and approved by the Institutional Animal Care and Use Committee in the National Tsing Hua University, Taiwan. The heartbeat signals of the dissected heart were measured by an electrocardiogram (ECG) recorder (EZ-BIO-01-S1-E, EZ instrument, Taiwan) through the two ITO microelectrodes, denoted as the positive (+) and negative (−) terminals in Fig. 21.10A. The reference terminal of the ECG recorder was connected to the reference electrode in the medium reservoir. The measured ECG signals are shown in Fig. 21.10B. The average heart rate of the zebrafish is 113 beats/min, agreeing with the fact that the heart rate of the zebrafish ranges from 80 to 180 beats/min [26]. This confirms that heartbeat signals can be recorded through ITO microelectrodes reliably. Moreover, Fig. 21.10B shows that the QRS waves of heartbeat signals are also recorded clearly to facilitate further analysis.
21.5.2 Biocompatibility test Rat hippocampal neurons were dissociated from the brain of rat fetus at embryonic day 18 in minimum essential medium. All experimental protocols complied with the provisions of the Animal Care and Use Committee of the National Tsing Hua University, Taiwan. The preparation and culturing procedures of hippocampal neurons were detailed in Ref. [17]. The hippocampal neurons were cultured on a cover glass and the proposed SU-8 microfluidic system for comparison. After 14 days of culturing, fluorescence immunohistochemical stain was used to examine the cytotoxicity. Fig. 21.11 shows the fluorescent images of cultured neurons (A) on the cover glass and (B) on the SU-8 microfluidic system. The blue circles are the cell nucleus of hippocampal neurons, while the green color reveals the cell morphology. Neurons in both cultures exhibit well-retained nucleus and skeleton.
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Fig. 21.11 The fluorescent images of rat hippocampal neurons on (A) the glass substrate and (B) the SU-8 substrate at DIV14 [17].
Fig. 21.12 Average numbers of neurons on the glass coverslip (control) and the SU-8 substrate. p = 0.0249 [17].
This result indicates that neurons on SU-8 substrate exhibit comparable health conditions to neurons on the glass substrate. The fluorescent images are further examined under a microscope. Three regions with an area of 1.5 mm2 are randomly selected from each sample. The number of nucleus in each selected region is counted. Fig. 21.12 shows and compares the average numbers of nucleus found on the SU-8 and the glass substrates. Taking the experiment with the cover glass as a control group, the statistical p-value for the two different groups is only 0.0249. This proves that the biocompatibility of the fabricated SU-8 microfluidic system is comparable to that of the glass substrate.
21.5.3 Cell-trapping capability test The cell-trapping capability of the proposed microfluidic system is further tested with microbeads and hippocampal neurons. Microbeads with a diameter of 10 μm (Micro
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Fig. 21.13 The snapshot of the experiment with microbeads (diameter = 10 μm), which are trapped in the microholes of the microfluidic system.
particles, No. 72986, SIGMA) were mixed with PBS with a concentration of 1.05 g/ cm3. As the PBS with microbeads were dripped into the cell reservoir, the flow in the microfluidic system was monitored and video recorded by a microscope. A snapshot is shown in Fig. 21.13, revealing that the microbeads are guided to enter cell-trapping wells and confined above ITO microelectrodes. The culture medium containing hippocampal neurons is further dripped into the cell reservoir of another microfluidic system. The neurons are also guided into the cell-trapping wells reliably. However, as shown by the snapshots in Fig. 21.14, the red circles highlight the neuron which enters a cell-trapping well for less than 1 s. This nonideal phenomenon comes from the fact that, as fluid continues to flow from the cell reservoir to the culture region, the fluidic pressure forces cells to deform themselves and subsequently to squeeze themselves out of the cell-trapping fences. The cell-trapping structure shown in this chapter should thus be improved further for trapping neurons for a long term. Nevertheless, the experiment with microbeads demonstrates that the proposed structure is able to trap cells (e.g., red blood cells in (JMM)) whose diameter is much larger than 10 μm.
21.6 Summary A transparent microlab consisting of ITO microelectrodes and wires on a glass substrate provides a great opportunity to study both electrophysiology and morphology of cells simultaneously. Although ITO microelectrodes have greater impedance than microelectrodes made of inert metals, the electrode impedance can be reduced by either increasing the physical electrode area or modifying the electrode surface through electrochemistry. The experiment with the zebrafish further demonstrates that ITO microelectrodes are able to record biopotential signals reliably. To facilitate long-term
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Fig. 21.14 The snapshots of the experiment with dissected neurons. The time difference between successive frames is 1–2 s.
study with biological cells, a transparent microfluidic structure can be built with SU-8 for guiding biological cells to stay above ITO microelectrodes. Our experimental results demonstrate the feasibility of using photolithography to construct a multilayer SU-8 microfluidic system, which is not only biocompatible but also capable of trapping biological cells with sophisticated structures. The slight imperfection is that neurons could escape trapping wells when the fluidic pressure is too strong. This could be solved by controlling the flow speed of the solution containing biological cells. Therefore, integrating a transparent microfluidic system with ITO microelectrodes is believed to be a useful platform for drug-screening and neuroscience research.
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