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. 2019 Nov 4;13(6):064104. doi: 10.1063/1.5123316

A pumpless solution exchange system for nanopore sensors

Tetsuya Yamada 1, Koki Kamiya 1, Toshihisa Osaki 1, Shoji Takeuchi 1,2,1,2,a)
PMCID: PMC6828575  PMID: 31700563

Abstract

This paper proposes a nanopore-based sensor exploiting the solution exchange of a droplet-based lipid bilayer driven by a superabsorbent polymer. Biological nanopores are candidates for use in portable sensors because of their potential to recognize and detect single molecules. One of the current challenges in the development of portable nanopore sensors is the inability to achieve continuous detection. To achieve continuous detection, we have exploited the suction force of a superabsorbent polymer to drive the continuous microfluidic flow required to wash the analyte out of the droplet. The superabsorbent polymer drives the microfluidic flow without electricity, and the developed solution exchange system remains compact. To demonstrate solution exchange in the droplet containing the lipid bilayer, the concentration of heptakis(6-O-sulfo)-β-cyclodextrin was monitored in a time-dependent manner using α-hemolysin nanopores. A reduction in the concentration, attributable to solution exchange, was successfully observed. We believe that the proposed system will increase the portability and usability of nanopore sensors.

I. INTRODUCTION

A number of studies have been carried out to develop biological nanopore sensors because of their selectivity and sensitivity, which allow the recognition and detection of single molecules.1–3 To date, various applications for nanopore sensors have been reported, including mass spectral analysis of polyethylene glycol,4 cocaine sensors,5 DNA/RNA sequencers,6 and pesticide vapor detectors.7 The size of a nanopore is in the nanometer scale, and therefore, the nanopore sensors have a potential to become a portable sensor device.

The study of nanopore sensors has been facilitated by microfabrication technology, resulting in improved reproducibility of nanopore preparation.8 Nanopores have been prepared using the droplet contact method.9 This method is simple and reproducible as a lipid bilayer is obtained by simply pipetting two aqueous droplets into a lipid-dispersed oil, in which a lipid bilayer is self-assembled between the two droplets surrounded by two lipid monolayers. Nanopores are incorporated into the bilayer spontaneously. The simplicity of the droplet contact method enables the utilization of the nanopore sensors in various fields.10

One of the current challenges in the development of portable nanopore sensors is the inability to achieve continuous detection. To achieve continuous detection, a compact solution exchange system is required, which can exchange the analytes for the next measurement. In the previous studies, the solution exchange of droplets was carried out by introducing microfluidic channels.11–13 However, this system is complex and cumbersome, as it uses two syringe pumps that hinder the portability of the nanopore sensors.

In this study, we have developed a pump-free solution exchange system for continuous detection [Fig. 1(a)]. The superabsorbent polymer, poly(acrylic acid), has a high water absorption capacity and can generate continuous flow.14–16 We have developed a device equipped with a reservoir and this superabsorbent polymer [Fig. 1(b)]. The solution in the detector is well connected to the reservoir and superabsorbent polymer by microchannels. The flow rate can be controlled by varying the microchannel dimensions. To demonstrate the solution exchange, we conducted time-dependent monitoring of the reduction in the blockade signal caused by heptakis(6-O-sulfo)-β-cyclodextrin (s7βCD) using α-hemolysin (αHL) nanopores, as schematically illustrated in Fig. 1(c).

FIG. 1.

FIG. 1.

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(a) Conceptual diagram of solution exchange driven by a superabsorbent polymer. The solution in a droplet is exchanged through the two microfluidic channels by the suction force of the superabsorbent polymer. (b) Photograph of the developed solution exchange device. (c) Schematic of the blockade signal of αHL nanopores attributed to the insertion of s7βCD. The flow is driven by the suction force of the superabsorbent polymer.

II. EXPERIMENTAL

A. Materials and reagents

n-Decane, calcein, and wild-type αHL were obtained from Sigma–Aldrich (St. Louis, MO, USA). 1,2-Diphytanoyl-sn-glycero-3-phosphocholine (DPhPC) was purchased from Avanti Polar Lipids (Alabaster, AL, USA). The poly(acrylic acid) superabsorbent polymer, AQUALIC CA, was obtained from Nippon Shokubai (Osaka, Japan). Heptakis(6-O-sulfo)-β-cyclodextrin heptasodium salt was purchased from Toronto Research Chemicals (Toronto, Canada). Chloroform, KCl, K2HPO4, and KH2PO4 were purchased from Wako Pure Chemical Industries, Ltd. (Tokyo, Japan). All aqueous solutions were prepared with ultrapure water from a Milli-Q system (Millipore, MA, USA). The buffer solution of 1.0M KCl was adjusted to pH 7.6 with 10 mM potassium phosphate. Poly(methyl methacrylate) (PMMA) substrates of 0.075, 1, and 4-mm thickness were purchased from Mitsubishi Chemical (Tokyo, Japan). The Super X adhesive was purchased from Cemedine Co., Ltd. (Japan). A superhydrophilic coating fluid, WG-R1, was purchased from Marusyosangyo Co., Ltd. (Tochigi, Japan).

B. Device design and fabrication

The dependence of the rates of inflow and outflow against the microchannel dimensions (width, depth, and length) is important for maintaining the volume of the droplet forming a lipid bilayer. To estimate the outflow caused by the hydrostatic pressure, we fabricated devices comprising a reservoir well and an observation well. The two wells were connected by a microchannel. We added 3 mL water in the reservoir and monitored the increase in the water level in the observation well by a microscope to estimate the inflow. The decrease in the water level in the observation well was also recorded. In this case, 45 μL of water and 100 mg of superabsorbent polymer were added in the observer well and the reservoir well, respectively. The experimental setup is schematically illustrated in Fig. S1 of the supplementary material.

The design of the device is shown in Fig. S2 of the supplementary material. Both the diameter and depth of the two wells are 4.0 mm. The depth, width, and length of the inflow microchannel were 0.2, 0.2, and 15 mm, respectively. The microchannel depth, width, and length for the outflow were 0.1, 0.2, and 20 mm, respectively.

The device was made from three PMMA substrates. Two microchannels for inflow and outflow were manufactured on the 1-mm thick bottom substrate via micromachining (MM-100, Modia system, Japan). A pair of holes (diameter: 1.1 mm) was also drilled on the bottom substrate for the placement of Ag wires. The absorbent well and reservoir were manufactured on the 4-mm thick substrates (Fig. S2 in the supplementary material). These three substrates were bonded by thermocompression for 5 min at 95 °C, with an applied force of 5 × 105 Nm−2 and then cooled to room temperature gradually. Subsequently, Ag wires were inserted into the device from the bottom surface of the wells and coated with Ag/AgCl paste to allow electrical measurements. A parylene film with 11 micropores (diameter: 100 μm) was sandwiched between two 75-μm thick PMMA films with an adhesive to act as a separator. The parylene film was fabricated with some modification of the method provided in a previous study to obtain a pore-size diameter of 100 μm and a film thickness of 5 μm.17 This separator was inserted between the double wells and bonded by an adhesive. The absorbent well and the microchannels were coated with a superhydrophilic fluid. The wettability of the surface allows good accessibility of the solution coming from the microchannel to the polymer.

C. Observing solution exchange using ink

To demonstrate the solution exchange driven by a superabsorbent polymer, blue ink (2 μL) was added to the two microchannels. A lipid bilayer was formed by the droplet contact method. First, both wells of the device were filled with 3 μL of n-decane containing a DPhPC (20 mg/mL); then, two droplets of ink (20 μL) were injected into each well. This procedure encapsulated the droplets within the oil and produced a lipid bilayer between the two droplets. Subsequently, 1.6 mL of buffer solution was added to the reservoir, and 70 mg of the superabsorbent polymer was added to the absorbent well. This polymer was used to drive the continuous flow. After the blue ink in the well had been washed out, 100 μL red ink was added to the reservoir to introduce red ink to the well.

D. Observing solution exchange using fluorescence

A solution of calcein (322 nM), a fluorescent dye, was used to examine the duration of the solution exchange. Calcein (32 μL) in the detector well was washed out by the buffer solution, and the images were obtained with an inverted microscope (IX71-DP72 system, Olympus, Japan. Objective Lens: UPLANFLN 10×, filter: NIBA Ex 470–490 nm, Em 510–550 nm). The light source was a Hg Lamp (OLYMPUS, U-RFL-T). The concentration of calcein was estimated from the fluorescence intensity using ImageJ (NIH, USA). We had initially obtained the calibration curve (fluorescence intensity vs calcein concentration). To drive the continuous flow, 3 mL buffer solution was added to the reservoir, and 100 mg superabsorbent polymer was added to the absorbent well.

E. Ion current measurements under solution exchange conditions

We verified the function of this system by monitoring the concentration of s7βCD based on the blockade rate of αHL nanopores. The calibration curve was constructed by investigating the blockade rate per αHL vs. the concentration of s7βCD. To form a lipid bilayer, 3 μL DPhPC (20 mg/mL) was added to both the wells. Then, 20 μL buffer solution containing s7βCD was added to the well connecting the two microchannels, while 20 μL buffer containing αHL (10 nM) was added to the other well. The ion current passing through the lipid bilayer was recorded using a wired custom-built amplifier. The current was monitored with a sampling frequency of 5 kHz, with a 1 kHz Bessel low-pass filter at a hold voltage of +60 mV. The gain was 1 mV/pA. We used a simple Faraday cage made of aluminum foil, and this covered the chamber device.

Under this condition, the αHL nanopores were spontaneously formed, generating stepwise ion current signals, and the number of the nanopores changed over time. The blockade rate per αHL nanopore was estimated from the slope of the plot of the blockade rate vs. the number of αHLs in the lipid bilayer.

To demonstrate solution exchange, 3.0 mL buffer solution was added to the reservoir, and 100 mg of the superabsorbent polymer was added to the absorbent well. Then, 3 μL DPhPC (20 mg/mL) was added to both wells. Subsequently, 20 μL buffer solution was added to the well connecting the two microchannels. In addition, a buffer solution containing αHL (1 nM) was added to the opposite well. After the formation of the nanopores, 1 μL s7βCD (2 mM) was repeatedly added, and the blockade rate was analyzed by appropriate software (Clampfit, Molecular Devices, LLC, CA, USA). The concentration of s7βCD around αHL nanopores was estimated from the calibration curve. The representative analysis shows the time-dependent change in the concentration of s7βCD (the third addition of the s7βCD).

III. RESULTS AND DISCUSSION

Both inflow and outflow occur in the droplet through the microchannels. The inflow is driven by hydrostatic pressure, while the outflow is driven by the suction force of the superabsorbent polymer. To maintain the volume of the droplet, these flow rates must be balanced. Therefore, the dependence of the flow rates against the microchannel dimensions (width, depth, and length) was independently investigated. Figure S3 in the supplementary material shows that the inflow and outflow can be balanced in the range of 0.03–5 μL/s. Based on this, we decided to use a flow rate of 0.24 μL/s for solution exchange, because high flow rates led to the rupture of the lipid bilayer.12 Besides, the flow driven by the superabsorbent polymer was constant for more than 40 min at this flow rate [Fig. S3(c) in the supplementary material]. The previous study demonstrated microfluidic flow using some kind of paper as absorbents and showed the microfluidic flow following the classic Washburn equation,18 suggesting that our concept is not limited to only the superabsorbent polymer. In addition, the previous study suggests the possibility of controlling the flow using absorbents.

We fabricated a device as shown in Fig. S4 of the supplementary material. To demonstrate the solution exchange capacity of the device, blue ink in the well was replaced by a buffer solution and red ink. First, the droplet contact method was applied with blue ink [Fig. 2(a)]. In this state, no significant microfluidic flow into the empty wells was observed for a few minutes. Subsequently, the buffer solution was added to the reservoir, and the superabsorbent polymer was added into the absorbent well. After this, the blue ink in the droplet connected to the microchannels was washed out gradually, leaving a transparent solution [Fig. 2(b)]. At the same time, the white superabsorbent polymer expanded and turned blue because of the outflow from the microchannel [Fig. 2(b)]. Furthermore, red ink was added to the reservoir, as shown in Fig. 2(c). The color of the droplet connected to the microchannels and that of the superabsorbent polymer became red [Fig. 2(d)], indicating that the reservoir solution had flowed into the well and the absorbent well. It is to be noted that the opposite well, isolated from the microchannels, retained the blue droplet. The entire process is shown in a 24 times fast-forward movie [Fig. 2 (Multimedia view)].

FIG. 2.

FIG. 2.

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Photographs of the solution exchange experiment. (a) Lipid bilayer formation between two blue-ink droplets with n-decane containing DPhPC. (b) A superabsorbent polymer and a buffer solution were added to the absorbent well and reservoir, respectively, which exchanged the blue ink with the buffer solution (transparent) in the bottom droplet. (c) Red ink was added to the reservoir. (d) The bottom droplet was exchanged with red ink. The figures at the bottom are cross-sectional illustrations corresponding to the photographs above. Multimedia view: https://doi.org/10.1063/1.5123316 Download video file (4.9MB, avi) DOI: 10.1063/1.5123316.1

To examine the duration of the solution exchange, a fluorescent molecule (calcein) was added to the droplet, and the time course of the variation in the fluorescence intensity was monitored [Fig. (3)]. To quantify the concentration of fluorescent in the detector well, we obtained the calibration curve (Fig. S5 in the supplementary material). In this experiment, the fluorescent solution was exchanged with the buffer solution in the reservoir. Figure 3 shows that the concentration of calcein decreased owing to solution exchange. To understand the behavior of the solution exchange, the continuous stirred-tank reactor model was applied, assuming the complete mixing of the solution in the droplet,19 as given by

C=C0exp(−v0Vt). (1)

FIG. 3.

FIG. 3.

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Time dependence of the decrease in fluorescent dye concentration. The inset shows the fluorescence images of the area around the bilayer at 0, 2, and 10 min.

In Eq. (1), C is the concentration of the fluorescent dye in the droplet at retention time t, C0 is the initial concentration of the fluorescent dye, v0 is the flow rate, and V is the volume of the droplet. In this experiment, C0, v0, and V were 322 nM, 0.24 μL/s, and 31 μL, respectively. The theoretically estimated values are plotted as a solid line in Fig. 3. The experimental data fit the equation well (R2 = 0.98), suggesting that the solution in the droplet was properly mixed during the solution exchange; therefore, the concentration around the bilayer can be assumed to be the same as that of the bulk solution. The duration of the solution exchange was about 8 min (less than 10 nM).

To demonstrate solution exchange with nanopore sensing, we utilized αHL nanopores and s7βCD as a blocker. We investigated the dependence of the blockade rate on the blocker concentration. First, to discriminate between the blockade current originating from the blockers (signal) and the accidental spikelike currents (noise), the dwell time was plotted against the blockade current level, as shown in Fig. S6 of the supplementary material. The dwell time and the blockade current attributed to the blockade by s7βCD were 78 ± 90 ms and 44 ± 3 pA (average ± standard deviation), respectively. Although the blockade current was almost consistent with previous reports,12,20 the dwell time was different (this could be attributed to the different experimental parameters such as the applied voltage21,22). We defined a blockade current (>30 pA) and dwell time (>20 ms) for the blockade by s7βCD, such that the spikelike current signals were eliminated. This definition excluded 10% signals with s7βCD. The spikelike currents were hardly observed (noise frequency: 5 min−1). The definition enabled us to automatically discriminate the signals using Clampfit software, as shown in Fig. S7 of the supplementary material. The αHL nanopores were spontaneously formed over time, and thus, it is important to know the relationship between the blockade rate (the blocking frequency of nanopores) and the number of nanopores. The analysis of multiple nanopores is well discussed in a previous study.23 The blockade rate linearly increased with the number of nanopores (Fig. S8 in the supplementary material), indicating that the blockade rate can be analyzed for multiple nanopores independently. The blockade rate per nanopore against the concentration of s7βCD was plotted to obtain the calibration curve (Fig. 4).

FIG. 4.

FIG. 4.

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Calibration curve for s7βCD. The blockade rate per αHL was plotted against the concentration of s7βCD. Error bars are the standard deviation for three repeated experiments.

This calibration plot was linear, with R2 = 0.95 and slope = 0.003 s−1 μM−1.

To demonstrate the analyte exchange in the droplet using the superabsorbent polymer, s7βCD was repeatedly added to the droplet, and the exchange of s7βCD with the buffer solution in the reservoir was verified. The nanopore blockade by s7βCD and its washout were monitored. Figure S9 in the supplementary material shows that the lipid bilayer was intact for more than 1 h under the solution exchange condition. Representative data for solution exchange is shown in Fig. 5. Figure 5(a) shows the current before the addition of the blocker, where no current blockade was observed. The current blockade started after the addition of the blocker, as shown in Fig. 5(b) (5 min after the addition of s7βCD; s7βCD concentration: 74 μM), and the blockade rate decreased over time due to the exchange of s7βCD with a buffer solution. No current blockade was observed further after 25 min of s7βCD addition, as shown in Fig. 5(c) (s7βCD concentration: 3 μM). Figure 5(d) shows the variation in s7βCD concentration obtained from the calibration curve in Fig. 4, indicating the dynamics of solution exchange. After the injection of s7βCD, its concentration gradually increased. Increase in the concentration of s7βCD to the maximum is a time-consuming process because of the time required for the diffusion of s7βCD into the nanopore. The concentration reached a peak and then decreased to zero. This corroborated that the solution exchange in the droplet was successfully driven by the superabsorbent polymer, thereby demonstrating that the time-dependent monitoring of the s7βCD concentration could be achieved using an αHL nanopore sensor without inducing any membrane rupture.

FIG. 5.

FIG. 5.

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Solution exchange tests using the superabsorbent polymer. The solution exchange was conducted while maintaining 4–5 αHL nanopores in the lipid bilayer membrane. Current–time traces obtained in the electrical recordings are shown (a) before the addition of s7βCD, (b) after the addition of s7βCD (5 min), and (c) after completion of solution exchange (25 min). (d) Monitoring of blocker concentration.

This system is more compact than the previous system that was driven by two syringe pumps12 and can be feasibly mounted on a wireless amplifier24 (Fig. S10 in the supplementary material). In addition, continuous flow for more than 1 h can be achieved by this system, without any electricity. This is a significant progress compared with a pump-free system driven by Laplace pressure working for a few minutes.25 This is because our system is driven by the suction force of a superabsorbent polymer possessing high water absorption capacity. A biological nanopore can be applicable for early pathological diagnosis, food care,7,26 etc., and our system combining the droplet contact method can facilitate the point of care testing and on-site testing because of its portability.

IV. CONCLUSIONS

Compact solution exchange provides additional advantages of repetitive and time-dependent monitoring using portable nanopore sensors. We proposed a solution exchange system that is driven by the suction force of a superabsorbent polymer. The pump-free system allows us to exchange a solution inside a droplet without causing a bilayer rupture. Using our system, the concentration of a blocker, s7βCD, was monitored using αHL nanopores under solution exchange conditions. Based on our results, we believe that our pump-free system will be applicable to the development of portable nanopore sensors.

SUPPLEMENTARY MATERIAL

See the supplementary material for the (i) experimental procedure to measure the flow rate, (ii) design of the device, (iii) result of the flow rate, (iv) photograph of the device, (v) calibration curve, (vi) relationship of dwell time vs blockade current, (vii) detection of blockade signals, (viii) relationship of signal frequency vs the number of nanopores, (ix) solution exchange experiment, and (x) the photograph of the compact sensor.

ACKNOWLEDGMENTS

This study is based on the results obtained from a project commissioned by the New Energy and Industrial Technology Development Organization (NEDO), Japan.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

See the supplementary material for the (i) experimental procedure to measure the flow rate, (ii) design of the device, (iii) result of the flow rate, (iv) photograph of the device, (v) calibration curve, (vi) relationship of dwell time vs blockade current, (vii) detection of blockade signals, (viii) relationship of signal frequency vs the number of nanopores, (ix) solution exchange experiment, and (x) the photograph of the compact sensor.

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