Electrochemicolor Imaging using an LSI-Based Device for Multiplexed

Subscriber access provided by READING UNIV

Article

Electrochemicolor Imaging using an LSIBased Device for Multiplexed Cell Assays Yusuke Kanno, Kosuke Ino, Hiroya Abe, Chika Sakamoto, Takehiro Onodera, Kumi Y Inoue, Atsushi Suda, Ryota Kunikata, Masahki Matsudaira, Hitoshi Shiku, and Tomokazu Matsue Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.7b03042 • Publication Date (Web): 01 Nov 2017 Downloaded from http://pubs.acs.org on November 3, 2017

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

Analytical Chemistry is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 22

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Analytical Chemistry

Electrochemicolor Imaging using an LSI-Based Device for Multiplexed Cell Assays Yusuke Kanno,a Kosuke Ino,*b Hiroya Abe,a Chika Sakamoto,a Takehiro Onodera,b Kumi Y. Inoue,a Atsushi Suda,c Ryota Kunikata,c Masahki Matsudaira,d Hitoshi Shiku,b Tomokazu Matsue*a a Graduate

School

of

Environmental

Studies,

Tohoku

University,

6-6-11-604

Aramaki-aza Aoba, Aoba-ku, Sendai 980-8579, Japan. b Graduate

School of Engineering, Tohoku University, 6-6-11-406 Aramaki-aza Aoba,

Aoba-ku, Sendai 980-8579, Japan. cJapan

Aviation Electronics Industry, Ltd., 1-1 Musashino 3-chome, Akishima-shi,

Tokyo 196-8555, Japan. dMicro

System Integration Center, Tohoku University, 519-1176 Aramaki-aza Aoba,

Aoba-ku, Sendai 980-0845, Japan.

1

ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Abstract: Multiplexed bioimaging systems have triggered the development of effective assays, contributing new biological information. Although electrochemical imaging is beneficial for quantitative analysis in real time, monitoring multiple cell functions is difficult. We have developed a novel electrochemical imaging system, herein, using a large-scale integration (LSI)-based amperometric device for detecting multiple biomolecules simultaneously. This system is designated as an electrochemicolor imaging system in which the current signals from two different types of biomolecules are depicted as a multicolor electrochemical image. The mode-selectable function of the 400-electrode device enables the imaging system and two different potentials can be independently applied to the selected electrodes. The imaging system is successfully applied for detecting multiple cell functions of the embryonic stem (ES) cell and the rat pheochromocytoma (PC12) cell aggregates. To the best of our knowledge, this is the first time that a real-time electrochemical mapping technique for multiple electroactive species, simultaneously, has been reported. The imaging system is a promising bioanalytical method for exploring complex biological phenomena. Keywords: Bioimaging Analytical electrochemistry Amperometry LSI-based chip device Multi detection system

2


Page 2 of 22

Page 3 of 22

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


INTRODUCTION Bioimaging is a powerful analytical method for bioanalysis to acquire visual and mapping information from single cells to three-dimensional cultured cells. Among the various bioimaging tools, electrochemical sensors have been determined beneficial for rapid and quantitative cell analysis.1 Along with the developments in micro/nano fabrication processes, several types of electrochemical devices have been developed as dynamic bioimaging tools by incorporating large electrode sensors.2,3 In particular, a large-scale integration (LSI) chip including a charge-coupled device (CCD) and a complementary metal-oxide semiconductor (CMOS) sensor has been developed for real-time bioimaging.4-13 Electrochemical devices are generally divided into two types: potentiometric and amperometric. Potentiometric devices have been widely used for monitoring signal transduction mainly based on the changes in the charges on electrode surfaces. These devices are used for monitoring ion concentrations such as the pH potentials including the action potentials of neuron cells

15

14

and membrane

in the biological field. For

selective assays, the sensors have been modified with ionophores for the detection of specific ions and ligand molecules such as antibodies for the detection of specific molecules.4,16,17 Amperometric devices have been applied for the selective and quantitative detection of electroactive molecules 20,21.

5-10,12,13,18,19

such as neurotransmitters in real time

The enzymatic activities in single cells and cell aggregates have also been

evaluated by amperometric devices. We have used amperometric devices for the selective imaging of alkaline phosphatase (ALP) activity in embryonic stem (ES) cells based on the oxidation of enzymatic products and have elucidated the differentiation levels of ES cells.22,23 Amperometric devices can be used to determine the local O2 concentrations. Cell respiration activity is monitored by the changes in the local O2 concentration in real time.24-26 The primary advantage of amperometric detection is that the species of interest can be selectively determined by applying a suitable potential. However, it is not easy for an integrated electrochemical device with several sensor points to detect multiple species simultaneously. Recently, Bellin and coworkers used square wave voltammetry (SWV) on a CMOS-based electrode array for the imaging of multiple metabolites in biofilms.13 However, the time resolution in SWV is insufficient for following rapid events. Therefore, it is important to develop an electrochemical device/system, which enables both rapid and multi-species imaging for monitoring fast, transient biological phenomena such as exocytosis events. 3



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 4 of 22

In this study, we develop a novel electrochemical imaging system called the electrochemicolor imaging system for the simultaneous analysis of multiple biomolecules, beyond the conventional electrochemical imaging systems. Two different potentials, as a checkered potential pattern, are applied to each electrode to acquire the oxidation and reduction currents from two different species derived from the cells, simultaneously. The two electrochemical responses consisting of reduction or oxidation currents are image-processed to acquire an electrochemical image consisting of two signal scales for the oxidation and reduction currents, providing an electrochemicolor image. In a conventional amperometric method using an electrode array device, only a single type of target molecule is monitored because the same potential is applied to all the electrodes (Fig. S1). Therefore, the electrochemical image using the previous method consists of a single signal scale and the resulting images are similar to single signal scale black and white images (Fig. S2). In contrast, two types of target molecules are simultaneously monitored in real time using the proposed method and the electrochemical image consists of two signal scales. Therefore, the image is similar to a color image with multi-signal scales (Fig. S2). Therefore, we designate the proposed method as electrochemicolor imaging. We have presented a very preliminary concept in the conference proceedings.27 This technique of applying two different potentials (V1 and V2 modes) as a checkered potential pattern was possible using an addressable device such as the “2G Bio-LSI” depicted in our previous report.10 In this study, this imaging system was applied for monitoring the ALP and respiration activities of stem cells. Moreover, dopamine release and the respiration activities of neuron-like cells were also imaged simultaneously. This electrochemicolor imaging system can enable the exploration

of

complex

biological

phenomena

including

brain

functions

and

sophisticated bioassays. To the best of our knowledge, this is the first time that a real-time electrochemical mapping technique for multiple electroactive species, simultaneously, has been reported.

4


Page 5 of 22

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


EXPERIMENTAL SECTION Device fabrication. The detailed fabrication process for the device has been described in our previous paper.10,28 Briefly, 400 Au or Pt working electrodes were prepared. SU-8 microwells (40-µm diameter, 5-µm deep) were then fabricated on the electrodes such that disk electrodes were formed on the bottom of the microwells. The sensor electrodes are arranged at a pitch of 250 µm. The general outline of the Bio-LSI device is shown in Fig. S3.

Electrochemicolor imaging. The electrochemicolor imaging process is depicted in Fig. 1. In the illustration, redox compounds (Redox A and Redox B) are placed on the device (Fig. 1A). As shown in Fig. 1B, a potential for the oxidation of Redox A and another for the reduction of Redox B are applied to the designated sensors, respectively. For electrochemicolor imaging, a checkered potential pattern is adapted (Fig. 1B). As shown in Fig. 1B, V1 and V2 are simultaneously applied on two electrode sets. The electronic circuits have been described in our previous paper (Fig. S3). As a result, two electrochemical images with a single signal scale are acquired for the oxidation of Redox A and the reduction of Redox B, respectively (Fig. 1C). These electrochemical images consist of 200 electrochemical signals and 200 blanks; hence a checkered image appears. The blanks are filled using a mathematical approach based on a biharmonic spline in LabVIEW to complete the electrochemical images consisting of 400 electrochemical signals (Fig. 1C). The two images are merged to create an electrochemical image consisting of two signal scales, designated as an electrochemicolor image (Fig. 1C). In all the experiments, an Ag/AgCl (sat. KCl) reference (RE-1CP, BAS Inc., Japan) and Pt counter electrodes were inserted into a solution on the LSI device. Amperometry was performed to obtain an electrochemical image every 200 ms. The biosamples were observed under a stereomicroscope during electrochemical imaging.

Electrochemicolor imaging of the enzyme membranes. The enzymatic reaction of glucose oxidase (GOx) consumes O2 in the presence of glucose.29 The consumption of O2 monitored from the reduction current of O2 at -0.5 V 30

was used to estimate the GOx activity. The ALP activity was measured by detecting

p-aminophenol (PAP), which is converted from p-aminophenyl phosphate (PAPP) by ALP (Fig. 2B).23,31 PAP is oxidized to QI p-quinone imine (QI) at 0.4 V. An enzyme membrane was prepared by mixing 24 mg of bovine serum albumin, 5



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 6 of 22

155 µL PBS, 5.3 µL 25% glutaraldehyde, 2.7 mg of GOx (210 U/mg, Wako Pure Chemical Industries, Ltd.), and/or 1 µL ALP (6.05×104 U/mL, calf intestine, Oriental Yeast Co., Ltd., Japan).22 As a control, membranes without enzymes were also prepared. The enzyme membranes were immersed in PBS containing 20 mM glucose + 4.7 mM PAPP (LKT Laboratories, USA), PBS containing 4.7 mM PAPP, or PBS on the device. The Au electrodes were stepped to -0.50 and 0.40 V for the reduction of O2 and the oxidation of PAP, respectively. The O2 reduction current at every sensor point in the membrane area was calculated by subtracting the reduction current of O2 without the membranes as the background current, from that with the membranes. Among the subtracted currents, the maximum value, ∆current, was used for the discussion of the GOx activity. The background current was estimated from the baseline of the reduction currents in a row of the image. For comparison of the ALP activities, the maximum oxidation current of PAP in the membrane area was selected for discussing the ALP activities.

Cell culture. Mouse ES cells (129/SvEv; DS Pharma, Japan) were cultured in a stem medium (DSRK100, DS Pharma, Japan) with 1000 U/mL of mouse leukemia inhibitory factor (mLIF) (Millipore, USA), 1% PS, and 0.1 mM 2-mercaptoethanol on a flask coated with 0.1% gelatin (ES-006-B, Millipore, USA). The medium was utilized to keep the cells undifferentiated. To form embryoid bodies (EBs) using the hanging drop method,32 20 µL droplets of the cell suspension containing 1000 cells were hung from a dish cover. To prepare undifferentiated or differentiated EBs, the cells were cultured for four or eight days in the droplets of the medium or in α-MEM (Gibco, USA) supplemented with 10% FBS, 1% PS, 0.1 mM MEM Non-Essential Amino Acids Solution, 0.1 mM 2-mercaptoethanol, 1 mM sodium pyruvate, and 2 mM L-glutamine for cell differentiation. PC12 cells were three-dimensionally cultured using the hanging-drop method to form cell aggregates. In brief, PC12 cells were suspended in an RPMI-1640 medium (Gibco) supplemented with 10% FBS and 1% PS. Then, 20 µL droplets containing 2000 cells were hung from a dish cover for eight days to form PC12 cell aggregates. For preparing fixed cell aggregates, the cell aggregates were washed with PBS (Catalog no. 041-20211, Wako Pure Chemical Industries, Ltd., Japan) and then, immersed in 4% paraformaldehyde

(Wako

Pure

Chemical

Industries,

Ltd.)

for

1

h.

Before

electrochemicolor imaging, the fixed cell aggregates were washed with PBS. The cells and cell aggregates were cultured at 37 °C in a humidified 6


Page 7 of 22

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


atmosphere of air containing 5% CO2.

Electrochemicolor imaging of the respiration and ALP activities of the EBs. Au electrodes were used in the experiments. The O2 reduction and PAP oxidation currents are acquired using the same schemes mentioned above. The ∆current for the O2 reduction was calculated as mentioned above. EBs were introduced into a 4.7 mM PAPP solution containing Tris and 2 mM MgCl2 (pH 9.5) on the device using a micropipette. The electrodes were stepped to -0.50 and 0.30 V for the reduction of O2 and the oxidation of PAP, respectively.

Electrochemicolor imaging of the respiration activity and dopamine release of the PC12 cell aggregates. Pt electrodes were used in the experiments. PC12 cell aggregates were introduced into an embryo respiration assay medium-2 (ERAM-2: pH 7.3, [K+] = 4.7 mM, [Ca2+] = 2.0 mM: Research Institute for the Functional Peptides, Japan) on the device using a micropipette. ERAM-2 containing 100 mM K+ was prepared as the stimulation solution to induce exocytotic dopamine release from the PC12 cells.33 Dopamine released from the PC12 cells was oxidized at 0.60 V.20 O2 was reduced at -0.50 V to monitor the cell respiration activity.34

∆Currents of the O2 reduction, with and without the stimulation of the PC12 cell aggregates, were calculated by subtracting the reduction current immediately before the introduction of the aggregates from that at approximately 120 s after the introduction. ∆Peak currents for the dopamine oxidation were calculated by subtracting the peak oxidation current from the background current. The peak oxidation current was obtained by averaging the currents for 1.8 s after reaching a peak. The background current was obtained by averaging the currents for 1.8 s before introducing the aggregates.

7



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

RESULTS AND DISCUSSION Electrochemicolor imaging of the enzyme membranes. To validate the concept, GOx and the ALP activities in the membranes were imaged simultaneously. The detection scheme is shown in Figs. 2A and 2 B. Enzyme membranes consisting of GOx, ALP, GOx + ALP, and without an enzyme were introduced into solutions containing glucose as the enzymatic substrate for GOx, PAPP as the enzymatic substrate for ALP, or with no enzymatic substrate. Figures 2C–2E present the optical and electrochemical images of the enzyme membranes on the device. The red and green scales in the electrochemical images indicate the reduction currents of O2 and the oxidation currents of PAP, respectively. PAP is an enzymatic product of the PAPP hydrolysis by ALP. PAP is oxidized at 0.4 V to QI. The results are summarized in Fig. 2F, indicating a decrease in the O2 reduction currents and in Fig. 2G for the PAP oxidation currents. In the image based on O2 reduction, the reduction currents are slightly lower in the area of the membrane with no enzyme than in those without the membranes (Figs. 2C–2F), indicating that the membranes themselves block the O2 supply to the electrodes from the bulk. In contrast, the reduction currents significantly decrease in the area under the membranes containing GOx in the glucose solution (Figs. 2C and 2F). As the GOx-catalyzed oxidation of glucose consumes oxygen, the differences in the current clearly demonstrate that the present system successfully affords images for the GOx activity. The images based on the oxidation currents of PAP show increased responses in the area under the membranes containing ALP in the PAPP solution (Figs. 2C, 2D, and 2G). The electrochemical response is directly related to the ALP activity of the membrane; therefore, the system provides the ALP activity images. When the membrane containing GOx and ALP was placed in the solution containing glucose and PAPP, a yellow color, consisting of a mixture of red and green colors, appeared in the electrochemicolor image (Fig. 2C). The above results indicate that the activities of GOx and ALP in the same membrane can be successfully detected simultaneously by the proposed electrochemicolor imaging system.

Electrochemicolor imaging of the respiration and ALP activities in EBs. Electrochemicolor imaging was applied for the evaluation of ES cells. ES cells were three-dimensionally cultured to form EBs, which were introduced into a solution containing PAPP, on the device, to detect their respiration and ALP activities. The detection scheme is shown in Figs. 3A and 3B. Figures 3C–3F display the 8


Page 8 of 22

Page 9 of 22

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


electrochemicolor images of the EBs. The results are summarized in Fig. 3G for the O2 consumption and Fig. 3H for the PAP generation. The proposed imaging system was used to characterize EBs cultured under different conditions. Optical images of the undifferentiated EBs show continuous proliferation from Day 4–Day 8 (Figs. 3C and 3E). On the other hand, an obvious change in the size of the differentiated EBs was not found during this period because cell differentiation suppresses cell growth (Figs. 3D and 3F). The image based on the O2 reduction currents indicates the respiration activity of the EBs. The consumption of O2 by respiration decreases the O2 reduction currents in the areas under the EBs. Although the sizes of the undifferentiated EBs on Day 8 were obviously large (diameter, above 500 µm), the respiratory activities of the EBs were similar to those of the undifferentiated EBs on Day 4 and the differentiated EBs on Days 4 and 8 (Fig. 3G). The above findings indicate that the cells inside the undifferentiated EBs on Day 8 would die due to the depletion of oxygen and a cavity might form inside the EBs.35 This electrochemical imaging shows a marked difference in the ALP activities between the undifferentiated and differentiated EBs because undifferentiated ES cells have a considerable amount of ALP.36 As the ALP reaction occurs mainly on the surface of the EBs,37 the ALP activities in the undifferentiated EBs on Day 8 are more than that on Day 4 (Fig. 3H). Electrochemicolor images from the undifferentiated EBs clearly depict a yellow color, indicating that the undifferentiated EBs show both ALP and respiration activities (Figs. 3C and 3E). This system provides information on both the activities of the EBs simultaneously; therefore, an easy assessment of the undifferentiation state of the EBs can be made. The blocking effects of oxygen diffusion from the bulk to the sensors is described in Supporting Information, although the culture condition of the living EBs in Fig. 3 were not the same. As shown in Fig. S4, the oxygen reduction currents changed only slightly near the fixed EBs. This observation indicates that the blocking effect is minimal, compared to the effect of respiratory activity of the cell aggregates. However, for precise evaluation, control experiments in the same condition should have been performed.

Electrochemicolor imaging of the respiration activity and dopamine release of the PC12 cell aggregates. Electrochemicolor imaging was applied for monitoring the respiration activity and dopamine release of the PC12 cell aggregates, simultaneously. The detection scheme is shown in Figs. 4A and 4B. A set of sensor electrodes was used for the detection 9



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

of O2 as described above and another set was used to detect dopamine from the oxidation currents.29 A solution containing 100 mM K+ was utilized to stimulate the cells for releasing dopamine. PC12 cell aggregates were introduced into the solution on the device to monitor their activities in real time. The electrochemicolor image in Fig. 4C indicates that both the activities were successfully imaged, when the cells were stimulated. Although two of the three cell aggregates exhibited both the activities, the other cell did not show any activity (Fig. 4C), indicating that a large number of cells in the cell aggregate died during the 8-day inoculation. Unstimulated cell aggregates, as controls, exhibited only respiration activities because dopamine was not released (Fig. 4D). In the fixed and unstimulated cell aggregates, both the activities were not monitored (Fig. 4E). Figure 4F shows the time-course analysis of the dopamine release from living cell aggregates, when stimulated. The oxidation currents of dopamine rapidly increased, after the transfer of the aggregates into a solution containing 100 mM K+, indicating that a considerable quantity of dopamine was rapidly released by the K+ stimulation. Then, the currents gradually decreased due to the depletion of the dopamine release (Fig. 4G). The peak and the oxidation currents correspond to our previous report.20 The influence of stimulation on the respiration activity was investigated by electrochemicolor imaging. As indexes of O2 consumption, the ∆currents of the O2 reduction were calculated by subtracting the reduction currents immediately before introducing the aggregates from those at approximately 120 s after the introduction. The ∆currents with the stimulation were compared to those without the stimulation. The O2 consumption of the cell aggregates with and without stimulation did not demonstrate significant differences (Fig. 4H), indicating that the respiration activity did not depend on dopamine release, under the present conditions. Figure 4I shows the relationship among the dopamine release, O2 consumption, and the sizes of the aggregates. The respiration activity of the cell aggregates increases as the size of aggregates increase, indicating that the respiration activity significantly depends on the cell number. On the other hand, no distinct correlation between the dopamine release and size was found in Fig. 4I. The above results demonstrate that electrochemicolor imaging is beneficial for monitoring the metabolism and exocytotic events of the cells, simultaneously. Figure 4H shows that the respiration activity does not depend on the dopamine release of the PC12 cell aggregates, under the present conditions. Zhsanov et al. reported that there was no O2 gradient in an extracellular medium containing differentiated PC12 cells with high K+ stimulation using a fluorescence-based sensing 10


Page 10 of 22

Page 11 of 22

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


technique, which agrees well with our result.38 However, research has revealed that O2 consumption decreases upon dopamine release on the surface of a slice of rat caudate nucleus, using fast-scan cyclic voltammetry (FSCV).39 In electrochemical imaging with an electrode array, the electrochemical signals depend upon the position and a size of each cell aggregate.40 Therefore, more precise analysis regarding the relationship between the respiration activity and dopamine release is needed by a correction of the electrochemical signals, considering the position and size of each cell aggregate or by calculating the gradient of the oxygen concentration.41 If the time resolution is improved significantly, single events of dopamine release can be monitored. In addition, electrochemicolor imaging can be applied for evaluation of 2D signal propagation in neuron networks. The maximum imaging rate is 3.5 ms/image in the present read-out system, although the noise is high at that rate. The read-out system including data acquisition devices is currently being modified to improve the time resolution, so that the maximum theoretical rate of 0.17 ms/image can be reached. In the near future, the fast imaging technique will be utilized for evaluation of single events of dopamine release. However, the size of the sensor electrodes should be reduced to separate clearly two events. The present device is more useful to investigate cellular activity for ensemble cell populations than single cells. Although the pitch is sufficient for evaluation of cell aggregates, it should be shortened for single-cell analysis. To achieve this, working electrodes connected from the aluminum pads at a pitch of 250 µm on the LSI wafer should be redesigned, and/or the wafer should be redesigned to pack sensors at a higher density. The concept for electrochemicolor imaging is not limited to potential-based selective detection. If the sensors are modified locally with enzyme or polymer, enzyme or polymer-based selective assay can be performed. For example, when glucose oxidase is modified locally at sensors in checked patterns, glucose and oxygen consumption of cell aggregates can be monitored simultaneously using the modified sensors and bare sensors. Also, amperometry can be combined with potentiometry in electrochemicolor imaging. For example, two sets of sensors can be utilized for the evaluation of action potentials and dopamine release to achieve 2D imaging of signal propagation in neuron networks. Thus, the concept of electrochemicolor imaging is useful for multiple analysis.

CONCLUSION As an overview, we have innovated a novel electrochemical imaging system using an LSI-based amperometric device to simultaneously monitor multiple 11



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

biomolecules in real time. This system, called the electrochemicolor imaging system, simultaneously detects the current signals from two different biomolecules, enabling multicolor electrochemical imaging with two signal scales. The electrochemicolor imaging was successfully applied for the real-time monitoring of the respiration activity and the undifferentiation marker enzyme of EBs, and the respiration activity and dopamine release of the PC12 cell aggregates. The results demonstrate that the new imaging system provides a promising bioanalytical method for effective cell analysis and correlation search of different biological phenomena in real time.

ASSOCIATED CONTENT Supporting Information The Supporting Information is available free of charge on the ACS Publications website. Figures S1-S4 describe the conventional electrochemical imaging, the concept of the electrochemicolor imaging, the general outline of the Bio-LSI device and the electrochemical imaging of the fixed EBs, respectively.

AUTHOR INFORMATION Corresponding Author *E-mail: [email protected] and [email protected] Phone: +81-22-795-5872 and +81-22-795-7209 Notes The authors declare no competing financial interest.

ACKNOWLEDGMENTS This work was supported in part by the Grant-in-Aid for Scientific Research (A) (No. 16H02280), Grant-in-Aid for Scientific Research (B) (No. 15H035420), Grant-in-Aid for Young Scientists (A) (No. 15H05415), and by the Grant-in-Aid for JSPS Fellows from the Japan Society for the Promotion of Science (JSPS). This work was also supported by the Special Coordination Funds for Promoting Science and Technology, Creation of Innovation Centers for Advanced Interdisciplinary Research Areas Program from the Japan Science and Technology Agency, the Asahi Glass Foundation, and by the Grant-in-Aid of the Tohoku University Institute for International Advanced Research and Education.

12


Page 12 of 22

Page 13 of 22

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


REFERENCES (1) Wang, J.; Trouillon, R.; Dunevall, J.; Ewing, A. G. Anal. Chem. 2014, 2014 86, 4515-4520. (2) Wolfrum, B.; Katelhon, E.; Yakushenko, A.; Krause, K. J.; Adly, N.; Huske, M.; Rinklin, P. Accounts Chem. Res. 2016, 2016 49, 2031-2040. (3) Miyamoto, K.; Yoshinobu, T. Sens. Mater. 2016, 2016 28, 1091-1104. (4) Hattori, T.; Satou, H.; Tokunaga, K.; Kato, R.; Sawada, K. Sens. Mater. 2015, 2015 27, 1023-1034. (5) Schienle, M.; Paulus, C.; Frey, A.; Hofmann, F.; Holzapfl, B.; Schindler-Bauer, P.; Thewes, R. IEEE J. Solid-State Circuit 2004, 2004 39, 2438-2445. (6) Ghindilis, A. L.; Smith, M. W.; Schwarzkopf, K. R.; Roth, K. M.; Peyvan, K.; Munro, S. B.; Lodes, M. J.; Stover, A. G.; Bernards, K.; Dill, K.; McShea, A. Biosens. Bioelectron. 2007, 2007 22, 1853-1860. (7) Kim, B. N.; Herbst, A. D.; Kim, S. J.; Minch, B. A.; Lindau, M. Biosens. Bioelectron. 2013, 2013 41, 736-744. (8) Kuno, T.; Niitsu, K.; Nakazato, K. Jpn. J. Appl. Phys. 2014, 2014 53. (9) Wydallis, J. B.; Feeny, R. M.; Wilson, W.; Kern, T.; Chen, T.; Tobet, S.; Reynolds, M. M.; Henry, C. S. Lab Chip 2015, 2015 15, 4075-4082. (10) Inoue, K. Y.; Matsudaira, M.; Nakano, M.; Ino, K.; Sakamoto, C.; Kanno, Y.; Kubo, R.; Kunikata, R.; Kira, A.; Suda, A.; Tsurumi, R.; Shioya, T.; Yoshida, S.; Muroyama, M.; Ishikawa, T.; Shiku, H.; Satoh, S.; Esashi, M.; Matsue, T. Lab Chip 2015, 2015 15, 848-856. (11) Laborde, C.; Pittino, F.; Verhoeven, H. A.; Lemay, S. G.; Selmi, L.; Jongsma, M. A.; Widdershoven, F. P. Nat. Nanotechnol. 2015, 2015 10, 791-795. (12) Bellin, D. L.; Sakhtah, H.; Rosenstein, J. K.; Levine, P. M.; Thimot, J.; Emmett, K.; Dietrich, L. E. P.; Shepard, K. L. Nat. Commun. 2014, 2014 5, 3256 (13) Bellin, D. L.; Sakhtah, H.; Zhang, Y. H.; Price-Whelan, A.; Dietrich, L. E. P.; Shepard, K. L. Nat. Commun. 2016, 2016 7, 10535. (14) Rothberg, J. M.; Hinz, W.; Rearick, T. M.; Schultz, J.; Mileski, W.; Davey, M.; Leamon, J. H.; Johnson, K.; Milgrew, M. J.; Edwards, M.; Hoon, J.; Simons, J. F.; Marran, D.; Myers, J. W.; Davidson, J. F.; Branting, A.; Nobile, J. R.; Puc, B. P.; Light, D.; Clark, T. A.; Huber, M.; Branciforte, J. T.; Stoner, I. B.; Cawley, S. E.; Lyons, M.; Fu, Y. T.; Homer, N.; Sedova, M.; Miao, X.; Reed, B.; Sabina, J.; Feierstein, E.; Schorn, M.; Alanjary, M.; Dimalanta, E.; Dressman, D.; Kasinskas, R.; Sokolsky, T.; Fidanza, J. A.; Namsaraev, E.; McKernan, K. J.; Williams, A.; Roth, G. T.; Bustillo, J. Nature 2011, 2011 475, 348-352. (15) Hai, A.; Shappir, J.; Spira, M. E. Nat. Methods 2010, 2010 7, 200-U250. (16) Maruyama, Y.; Terao, S.; Sawada, K. Biosens. Bioelectron. 2009, 2009 24, 3108-3112. 13



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(17) Klein, M.; Kates, R.; Chucholowski, N.; Schmitt, M.; Lyden, C. Sens. Actuator

B-Chem. 1995, 1995 27, 474-476. (18) Mathwig, K.; Zafarani, H. R.; Speck, J. M.; Sarkar, S.; Lang, H.; Lemay, S. G.; Rassaei, L.; Schmidt, O. G. J. Phys. Chem. C 2016, 2016 120, 23262-23267. (19) Yao, J.; Liu, X. A.; Gillis, K. D. Anal. Methods 2015, 2015 7, 5760-5766. (20) Abe, H.; Ino, K.; Li, C. Z.; Kanno, Y.; Inoue, K. Y.; Suda, A.; Kunikata, R.; Matsudaira, M.; Takahashi, Y.; Shiku, H.; Matsue, T. Anal. Chem. 2015, 2015 87, 6364-6370. (21) Sansuk, S.; Bitziou, E.; Joseph, M. B.; Covington, J. A.; Boutelle, M. G.; Unwin, P. R.; Macpherson, J. V. Anal. Chem. 2013, 2013 85, 163-169. (22) Ino, K.; Kanno, Y.; Nishijo, T.; Komaki, H.; Yamada, Y.; Yoshida, S.; Takahashi, Y.; Shiku, H.; Matsue, T. Anal. Chem. 2014 2014, 14 86, 4016-4023. (23) Ino, K.; Nishijo, T.; Arai, T.; Kanno, Y.; Takahashi, Y.; Shiku, H.; Matsue, T. Angew.

Chem. Int. Ed. 2012, 2012 51, 6648-6652. (24) Yasukawa, T.; Kondo, Y.; Uchida, I.; Matsue, T. Chem. Lett. 1998, 1998 27, 767-768. (25) Takii, Y.; Takoh, K.; Nishizawa, M.; Matsue, T. Electrochim. Acta 2003, 2003 48, 3381-3385. (26) Shiku, H.; Shiraishi, T.; Ohya, H.; Matsue, T.; Abe, H.; Hoshi, H.; Kobayashi, M.

Anal. Chem. 2001, 2001 73, 3751-3758. (27) Kanno, Y.; Ino, K.; Abe, H.; Inoue, K. Y.; Matsudaira, M.; Suda, A.; Kunikata, R.; Shiku, H.; Matsue, T. Proceedings of MicroTAS 2016, Dublin, Ireland 2016, 2016 104-105. (28) Kanno, Y.; Ino, K.; Sakamoto, C.; Inoue, K. Y.; Matsudaira, M.; Suda, A.; Kunikata, R.; Ishikawa, T.; Abe, H.; Shiku, H.; Matsue, T. Biosens. Bioelectron. 2016, 2016 77, 709-714. (29) Heller, A.; Feldman, B. Chem. Rev. 2008, 2008 108, 2482-2505. (30) Shen, Y.; Trauble, M.; Wittstock, G. Anal. Chem. 2008, 2008 80, 750-759. (31) Tang, H. T.; Lunte, C. E.; Halsall, H. B.; Heineman, W. R. Anal. Chim. Acta 1988, 1988

214, 187-195. (32) Kurosawa, H.; Imamura, T.; Koike, M.; Sasaki, K.; Amano, Y. J. Biosci. Bioeng. 2003, 2003 96, 409-411. (33) Chen, T. K.; Luo, G.; Ewing, A. G. Anal. Chem. 1994, 1994 66, 3031-3035. (34) Obregon, R.; Horiguchi, Y.; Arai, T.; Abe, S.; Zhou, Y.; RyosukeTakahashi; Hisada, A.; Ino, K.; Shiku, H.; Matsue, T. Talanta 2012, 2012 94, 30-35. (35) Van Winkle, A. P.; Gates, I. D.; Kallos, M. S. Cells Tissues Organs 2012, 2012 196, 34-47. (36) Sen, M.; Ino, K.; Inoue, K. Y.; Arai, T.; Nishijo, T.; Suda, A.; Kunikata, R.; Shiku, H.; Matsue, T. Biosens. Bioelectron. 2013, 2013 48, 12-18. (37) Arai, T.; Nishijo, T.; Matsumae, Y.; Zhou, Y.; Ino, K.; Shiku, H.; Matsue, T. Anal.

Chem. 2013, 2013 85, 9647-9654. 14


Page 14 of 22

Page 15 of 22

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(38) Zhdanov, A. V.; Ward, M. W.; Prehn, J. H.; Papkovsky, D. B. J. Biol. Chem. 2008, 2008 283, 5650-5661. (39) Kennedy, R. T.; Jones, S. R.; Wightman, R. M. Neuroscience 1992, 1992 47, 603-612. (40) Kanno, Y.; Ino, K.; Inoue, K. Y.; Suda, A.; Kunikata, R.; Matsudaira, M.; Shiku, H.; Matsue, T. Anal. Sci. 2015, 2015 31, 715-719. (41) Shiku, H.; Arai, T.; Zhou, Y.; Aoki, N.; Nishijo, T.; Horiguchi, Y.; Ino, K.; Matsue, T.

Mol. Biosyst. 2013, 2013 9, 2701-2711.

15



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure Figure 1. Electrochemicolor imaging scheme. (A) Illustration of the samples on the device. Redox A and Redox B are reductant and oxidant, respectively. (B) Applied potential map of the 400 electrochemical sensors using the V1 and V2 modes. The red and green boxes indicate the potentials for the oxidation of Redox A and the reduction of Redox B, respectively. (C) Imaging process: Two electrochemical images are constructed using the electrochemical signals from the sensor electrodes in the V1 and V2 modes, respectively. The white color indicates the blanks in the current signals. These images show the oxidation currents of Redox A and the reduction currents of Redox B, respectively. Further, the blanks are filled using a mathematical approach. Finally, these images are merged to construct an electrochemicolor image consisting of two signal scales.

16


Page 16 of 22

Page 17 of 22

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure Figure 2. Electrochemicolor imaging of the GOx and ALP activities in the membranes. (A, B) Detection schemes for the GOx and ALP activities, respectively. The blue and orange colors indicate the O2 and PAP concentrations, respectively. (C-E) Optical and electrochemical images of the membranes on the device. Enzymatic substrates (C: glucose and PAPP, D: PAPP, E: no substrate) were added. These electrochemical images consist of signals at -0.5 and 0.4 V for depicting the reduction currents of O2 and the oxidation currents of PAP, respectively. The reduction currents of O2 (F) and the oxidation currents of PAP (G) on the membranes are plotted onto graphs, respectively. These images were acquired after 220 s of the potential steps. Au sensor electrodes were utilized for detection. 17



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure Figure 3. Electrochemicolor imaging of the respiration and ALP activities of EBs. (A, B) Detection schemes for the respiration and ALP activities, respectively. The blue and orange colors indicate the O2 and PAP concentrations, respectively. (C-F) Optical and electrochemical images of the undifferentiated (C, E) and differentiated EBs (D, F). 18


Page 18 of 22

Page 19 of 22

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


These EBs were cultured for four (C, D) or eight (E, F) days. The electrochemical images consist of signals at -0.5 and 0.3 V for depicting the reduction currents of O2 and the oxidation currents of PAP, respectively. These images were acquired after 180 s of the potential steps. (G) ∆Currents of the O2 reduction currents in the EBs. (H) Oxidation currents of PAP in the EBs. Au sensor electrodes were utilized for detection.

19



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure Figure 4. Electrochemicolor imaging of the respiration activity and dopamine release of the PC12 cell aggregates. (A, B) Detection schemes for the respiration activity and 20


Page 20 of 22

Page 21 of 22

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


dopamine release of the PC12 cell aggregates, respectively. The blue and orange colors indicate the O2 and dopamine concentrations, respectively. (C-E) Optical and electrochemical images of the aggregates on the device. The electrochemical images consist of signals at -0.5 and 0.6 V for depicting the reduction currents of O2 and the oxidation currents of dopamine, respectively. The cells were stimulated (C) or unstimulated (D). (E) Fixed and unstimulated PC12 cell aggregates. (F) Time-course electrochemicolor images after stimulation. (G) Amperogram for the oxidation currents of dopamine from the sensor indicated by an arrow in (F). (H) ∆Currents for the O2 reduction currents in the stimulated or unstimulated cell aggregates. (I) Relationship among the dopamine- oxidation ∆peak current, ∆current of the O2 reduction currents, and the sizes of the cell aggregates. Pt sensor electrodes were utilized for detection.

21



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

For TOC only.

22


Page 22 of 22

Electrochemicolor Imaging using an LSI-Based Device for Multiplexed

Recommend Documents