pubs.acs.org/ac

ArticleNanopore Filter: A Method for Counting and Extracting Single DNA
Molecules Using a Biological Nanopore
Asuka Tada, Nanami Takeuchi, Kan Shoji,* and Ryuji Kawano*
Cite This: https://doi.org/10.1021/acs.analchem.3c00573

ACCESS

Metrics & More

Read Online

Article Recommendations

sı Supporting Information*

ABSTRACT: This paper describes a method for the real-time
counting and extraction of DNA molecules at the single-molecule
level by nanopore technology. As a powerful tool for electrochemical single-molecule detection, nanopore technology eliminates the need for labeling or partitioning sample solutions at the
femtoliter level. Here, we attempt to develop a DNA filteringsystem utilizing an α-hemolysin (αHL) nanopore. This systemcomprises two droplets, one filling with and one emptying DNA
molecules, separated by a planar lipid bilayer containing αHLnanopores. The translocation of DNA through the nanopores isobserved by measuring the channel current, and the number oftranslocated molecules can also be verified by quantitative
polymerase chain reaction (qPCR). However, we found that the issue of contamination seems to be an almost insolvable problem
in single-molecule counting. To tackle this problem, we tried to optimize the experimental environment, reduce the volume ofsolution containing the target molecule, and use the PCR clamp method. Although further efforts are still needed to achieve a singlemolecule filter with electrical counting, our proposed method shows a linear relationship between the electrical counting and qPCR
estimation of the number of DNA molecules.

■

INTRODUCTION

through the pore subsequently produces a characteristic ion
current blockage, which can be utilized to identify the analytethrough electrical means. The inner diameter of biological
nanopores typically ranges from sub-nanometer to several
nanometers.4−8 α-Hemolysin (αHL), a toxin produced by
Staphylococcus aureus, has been widely utilized to detect singlestranded nucleic acid molecules since its initial reported use by
Kasianowicz et al. in 1996, due to its ability to reproducibly
form rigid nanopores with diameters of around 1.4 nm.9
Researchers in the field of nanopore technology have been
particularly focused on the application of nanopores for DNA
sequencing,10−14 with a DNA sequencer becoming commercially available in 2015 and being updated with the use of the
Escherichia coli transport channel CsgG15 in 2016.
In previous research, we have applied αHL nanopores in thecontext of DNA computing technology,16−21 utilizing them inthe decoding of oligonucleotides. DNA computing involves theperformance of logical operations or arithmetic calculations
using DNA molecules.22−26 To accomplish this, we prepared

The use of nanopore technology as a powerful tool forelectrochemical single-molecule detection at the femtoliter
level without the need for labeling or partitioning sample
solutions has been well established.1−3 The basic configuration
for nanopore measurements involves connecting two separated
electrolytes through a nanopore, with an ion current generated
by applying a voltage (Figure 1). The migration of analytes

Received: February 7, 2023
Accepted: May 23, 2023

Figure 1. (a) Photograph of the microdevice used for lipid bilayer
preparation. Two chambers are separated by a separator with aparylene film. (b) Illustration of the droplet contact method andcollection of single molecules of DNA. Immediately after a single
molecule of DNA passes through the αHL pore in the lipid bilayer,
the separator is slid across to intercept the bilayer.
© XXXX American Chemical SocietyA

https://doi.org/10.1021/acs.analchem.3c00573
Anal. Chem. XXXX, XXX, XXX−XXX

Analytical Chemistrypubs.acs.org/ac

water-in-oil droplets27−30 for each of the input DNA,
computational DNA, and output DNA and used αHLnanopores to translocate the DNA molecules between thedroplets.18 Specifically, the input DNA in the input droplet
moves into the computational droplet, and the output DNA in
the computational droplet moves into the output droplet,molecule by molecule, through the pore. In the case of singlemolecule detection using nanopore technology, the target
molecules are counted in real-time as they pass through thepore and are quantified according to the number of passing
molecules per unit time.16,31 Our previous research using
nanopore experiments has led us to consider the nanopore
system as a valuable tool for counting and collecting single
molecules in real time. In this study, we utilized a nanopore
system with the αHL nanopore as a molecular filtering device.
The αHL nanopore was reconstituted in a lipid bilayer that
separated the measurement solution into two compartments,and single-stranded DNA (ssDNA) was passed through thepore, moving from the sample droplet (cis-side compartment)
to the collecting droplet (trans-side compartment). The
translocation of ssDNA could be monitored through pulsed
blockage signals, with the number of signals corresponding to
the number of translocated ssDNA molecules. This device has
the potential to allow for real-time counting of a specificnumber of molecules and the simultaneous extraction of a
specific number of molecules by collecting the droplet on thetrans side. To evaluate the reliability of the nanopore filter
system, the number of ssDNA molecules in the collected
solution was verified through a quantitative polymerase chain
reaction (qPCR) assay.

Articlereports,4,17,32 we prepared planar bilayer membranes (pBLMs)
by the droplet contact method using a microdevice in which an
infinity symbol-shaped micro chamber is separated into two
micro chambers by a 5 μm thick parylene film with a single
hole of 100 μm diameter (Figure 1). For electrical measurements of nanopores, Ag/AgCl electrodes were prepared at thebottom of each chamber. We basically used a brand new device
for each experiment.
pBLM formation using this device was performed with thefollowing procedure. First, 10 mg/mL DPhPC solution (in ndecane) (0.9 μL) was poured into both chambers. Next, thebuffer solution (4.7 μL) with 100 nM αHL was added into oneof the chambers which was connected to the ground terminal.
Then, the buffer solution (4.7 μL) with 160 nM target DNA
was added into the other chamber. A few minutes after adding
the buffer solution, the pBLM spontaneously formed bycontacting two lipid monolayers.
Contact Bubble Bilayer Method Using Glass Pipettes.
The contact bubble bilayer (CBB) method33,34 was performed
under an inverted microscope (IX71; Olympus, Tokyo, Japan),and images were recorded using a digital camera (MS-200; Bio
Craft, Tokyo, Japan). Glass pipettes for bubble formation andperfusion inside the bubble were fabricated by pulling aborosilicate glass capillary (BF100-50-10; OD/ID; 1.0/0.5
mm, Sutter Instrument, Novato, CA) with a micropipettepuller (PC-100; Narishige, Tokyo, Japan). Then, the tips of thepipettes were cut to obtain a tip diameter of around 30 μm andlightly polished using a micro-forge (MF-900; Narishige).
First, 10 mg/mL DPhPC solution (in n-decane) (100 μL) was
put on a slide glass. Next, two glass pipettes, one filled with thebuffer solution with 100 nM αHL and the other filled with thebuffer solution containing 100 nM target DNA were immersed
into the DPhPC solution. Then, two water bubbles wereformed from the pipettes by applying pressure inside thepipettes. Finally, the pBLM was formed by manipulating thepipettes so the two water bubbles came into contact. The
pipette position was controlled by micromanipulators (UM3C; Narishige) under the microscope. The pressure in thepipette was regulated by a micro injector (IM-9B; Narishige).
Ag/AgCl electrodes were placed in both glass pipettes. The
ionic current was measured by applying a transmembranepotential of +180 mV.
Nanopore Filter Experiments. We prepared 30 pM PNA,
10 pM target DNA, 100 nM αHL, [0.1 M NH4Cl solution with
1 mM MOPS at pH 7.0], and [1 M NH4Cl solution with 10
mM MOPS at pH 7.0] for the nanopore filter experiments.
After forming pBLMs, αHL nanopores were spontaneously
reconstituted into the pBLM, and this nanopore-integrated
pBLM acted as a nanopore filter by translocating DNA. The
reconstitution of nanopores and the translocation of DNA
were confirmed by measuring the ionic current. When αHLnanopores were reconstituted into the pBLMs, step-likeincreases in current were obtained. Then, blockage signals
were observed when DNA translocated through the nanopores.
We counted signals with a blocking ratio of over 95% as thetarget DNA signals in experiments without PNA. With PNA,
we identified signals with a blocking ratio more than around
70% as target signals. After confirming the translocation ofDNA, the solution in which DNA flowed was collected. We
added 0.5 μL of lipid/oil to the cis-side chamber, and gently
collected a droplet in the trans-side chamber using a gelloading tip with a 0.3 mm tip diameter (Thermo Fisher
Scientific, Waltham, MA, USA). To check that the membrane■

EXPERIMENTAL SECTION
Reagents and Chemicals. In this study, we used thefollowing reagents: 1,2-diphytanoyl-sn-glycero-3-phosphocholine (DPhPC; Avanti. Polar Lipids, AL, USA); 1,2-dioleoyl-snglycero-3-phospho-rac-(1-glycerol) (DOPG; Avanti. Polar
Lipids); n-decane (Wako Pure Chemical Industries, Osaka,
Japan); 3-morpholinopropane-1-sulfonic acid (MOPS; Nacalai
Tesque, Kyoto, Japan); potassium chloride (KCl; Nacalai
Tesque), sodium chloride (NaCl; Nacalai Tesque), ammonium chloride (NH4Cl; Wako Pure Chemical Industries),
lithium chloride (LiCl; Nacalai Tesque), calcium chloride(CaCl2; Nacalai Tesque), magnesium chloride (MgCl2;
Nacalai Tesque), and streptavidin (Wako Pure Chemical
Industries). Buffered electrolyte solutions were prepared using
ultrapure water (MilliQ; Merck Millopore, Burlington, MA,
USA). Wild-type αHL (Sigma-Aldrich, St. Louis, MO, USA,and List Biological Laboratories, Campbell, CA, USA) was
obtained as a monomeric polypeptide, isolated from S. aureus.
For use, samples were diluted to the designated concentration
using a buffered electrolyte solution and stored at 4 °C. Highperformance liquid chromatography-grade DNA oligonucleotides and peptide nucleic acid (PNA) were synthesized by
Eurofins Genomics (Tokyo, Japan) and PANAGEN (Daejeon,South Korea), respectively, and stored at −20 °C. TBE buffer(10×) was obtained from Takara Bio (Shiga, Japan) and was
diluted 10-fold for gel electrophoresis. TB Green Premix Ex
Taq II (Takara Bio) and KOD SYBR qPCR Mix (TOYOBO,
Osaka, Japan) were used for the real-time polymerase chain
reaction (real-time PCR).
Planar Bilayer Membranes for Nanopore Filter
Experiments Prepared with the Droplet Contact
Method Using a Microdevice. As shown in our previous
Bhttps://doi.org/10.1021/acs.analchem.3c00573
Anal. Chem. XXXX, XXX, XXX−XXX

Analytical Chemistrypubs.acs.org/ac

ArticleFigure 2. (a) Schematic illustration of the nanopore filter experiments. After observing 30 blockage signals, the solution in the chamber connectedto the recording terminal of a patch-clamp amplifier was collected. Then, the DNA was quantified by qPCR assay. (b) Amplification curves
obtained from the filtered target DNA and 106 molecules of the target DNA by real-time PCR assay. Similar Ct values were obtained from thesesamples. (c) Amplification curves obtained from 102 molecules of the target DNA in different salt solutions. DNA amplification was observed only
when using NH4Cl and MilliQ with the PCR solution. (d) Calibration curves of the Ct value versus the molecular number with 94 mM NH4Cl andMilliQ with the PCR solution. The Ct value can fit linearly with the natural logarithmic value of the DNA number. The literature value wasreferenced from N. Jothikumar et al., J. Virol. Methods, 2006.39

was not ruptured, a voltage of −180 mV was maintained
throughout the collection experiments.
Real-Time PCR for Identifying the Number of DNA
Molecules Collected in the Nanopore Filter Experiments. The number of DNA molecules in the collected
solution was confirmed by a real-time PCR assay. In this study,
we performed SYBR Green real-time PCR with two different
reaction volumes (25 and 50 μL), and the chemicalcomponents of each reaction solution were as below. The 25
μL reaction solution contained 12.5 μL of TB Green Premix
Ex Taq II, target DNA (nanopore filtered or specificconcentration), 0.4 μM primers, 94 mM KCl, and 0.94 mM
MOPS (pH 7.0). The 50 μL reaction solution contained 25 μL
of KOD SYBR qPCR Mix, DNA (nanopore filtered or specificconcentration), 0.2 μM primers, 94 mM NH4Cl, and 0.94 mM
MOPS (pH 7.0). Thermal Cycler Dice Real Time System Lite
(Takara Bio) was used for all real-time PCR tests. The DNA
amplification cycles for each reaction volume were performedby the following protocols: 40 cycles at 95 °C (5 s) and 60 °C(30 s) for the use of the 25 μL reaction solution (TB Green
Premix Ex Taq II) and cycles at 98 °C (10 s), 57 °C (20 s),
and 68 °C (30 s) for the use of the 50 μL reaction solution(KOD SYBR qPCR Mix). Real-time PCR data were obtained
after the reaction, and the crossing point (CP) was calculatedby the Thermal Cycler Dice system software. For generation ofstandard quantitation curves using 10 to 105 copies of thetarget DNA, the Ct values were plotted proportionally to thelogarithm of the input copy number. Negative controls were
included in each run.
Polyacrylamide Gel Electrophoresis for Analysis of
PCR Products. Amplification products of real-time PCR wereanalyzed by denaturing polyacrylamide gel electrophoresis(PAGE, containing 19/1 acrylamide/bis (w/w)) on 8% gels in
1× TBE buffer (89 mM Tris-borate, 2 mM ethylenediaminetetraacetic acid, pH 8.3) at a constant power of 7.5 W for 30 min
at room temperature. After electrophoresis, the gel was stainedwith diluted SYBR Green II solution (Takara Bio) for 30 min.
Then, the images were obtained using an LED transilluminator (Bio Craft).
PCR Clamp Using PNA Molecules. A duplex of PNA(5′TGATTCTCAGCCCTTCGC3′) and the target DNA
were prepared by mixing them at a specific concentration
ratio, followed by annealing in a 95 °C heat block for 5 min
and annealing to room temperature over 20 h. SYBR Green
real-time PCR amplifications were performed with the 50 μL
reaction solution including 25 μL KOD SYBR qPCR Mix, thePNA-DNA duplexes (nanopore filtered or specific concentration), 0.2 μM primers, 94 mM NH4Cl, and 0.94 mM MOPS
(pH 7.0). The DNA amplification protocol of 45 cycles at 94°C (0 s), 70 °C (5 s), 57 °C (5 s), and 65 °C (10 s) was
conducted with the real-time PCR equipment.
Measurement of PNA Concentration to Check PNA
Adsorption on the Surface of Microtubes and Pipet
Tips. The PNA concentrations were measured using a UV−vis
spectroscopy microvolume spectrophotometer (Nanodrop;
Thermo Fisher Scientific).
Channel Current Measurements and Data Analysis.
Channel currents were monitored using a patch-clamp
amplifier (PICO2; Tecella, Foothill Ranch, CA, USA) with a
7.9 kHz low-pass filter at a sampling frequency of 40 kHz.
These settings were carefully chosen to enable the detection of
DNA translocations in the experiments.35,36 Constant transmembrane potentials of +180 and +150 mV were applied tothe cis-side droplet in the case of DNA and PNA-DNA duplex
C

https://doi.org/10.1021/acs.analchem.3c00573
Anal. Chem. XXXX, XXX, XXX−XXX

Analytical Chemistrypubs.acs.org/ac

measurements, respectively. When αHL nanopores werereconstituted into the pBLMs, current increases were observed
due to ion passage through the nanopores. Current blockages
were also observed when DNA molecules translocated throughthe nanopores. Analysis of channel current signals was
performed using Clampfit software ver. 10.7 (Molecular
Devices, CA, USA).

Articlewas shorter than those for DNA #1 to #6, resulting in lesscomplementarity between the primers, which inhibited thegeneration of primer dimers. However, 100 molecules of DNA#7 were not amplified when using a 94 mM KCl buffer thatwas transferred from the nanopore experiments, which
required high electrolyte concentrations to ensure ion
conductance. We therefore optimized the buffer conditionsof the nanopore filter by changing the electrolytes. In general,
the optimal experimental conditions for PCR experiments are
known;40 generally, the pH in the PCR buffer is 8 to 9.5, and 1
to 4 mM Mg2+ is needed as a cofactor for polymerase. The
optimal concentrations of Na+ (enhances primer annealing)
and NH4+ (improves the specificity of primers) are 50 and 20
mM, respectively.
We then tested PCR experiments using six different
electrolytes (KCl, NaCl, NH4Cl, LiCl2, CaCl2, and MgCl2)
in the PCR solution. The salt concentration and number of
DNA #7 were 94 mM and 100 molecules, respectively. We
only obtained an amplification of the target DNA with use of
the NH4Cl solution (Figure 2d). NH4+ ions compete withhydrogen bonds between DNA strands and destabilize weakhydrogen bonds between mismatched primer/template basepairs.40 Hybridization of the specific primer and target DNA
was promoted by NH4+ ions, which enabled amplification from100 target molecules. Although the Ct value using the NH4Cl
solution was slightly higher than that using only the PCR
solution, we decided to use NH4Cl as the electrolyte because
electrolytes must be added to the solution to obtain ioniccurrents in the nanopore filter experiments. We prepared acalibration curve of the Ct value vs molecular number tocalibrate the DNA number when using a 94 mM NH4Cl
solution (Figure 2c). The Ct value versus the natural
logarithmic value of the DNA number was linearly fit with ahigh correlation factor (R2) (mean R2 value of 0.97), with agradient of −1.3. Moreover, we confirmed that the target DNA
was amplified by qPCR experiments from the melting curve
analysis and gel electrophoresis (Figure S1).
Problem (ii): Investigation of Contaminated Pathways and Implementation of Negative Control Experiments. Improvement of the Experimental Environment.
Since DNA contamination was strongly suspected, weinvestigated the contamination pathways and attempted toreduce it. First, we investigated DNA contamination from theambient atmosphere. To prevent contamination from the air,
we improved the experimental environment as follows:■

RESULTS AND DISCUSSION
Separation of DNA by Nanopore Filters andQuantification with qPCR. Single DNA molecules weretranslocated through the αHL nanopore in the droplet contact
device (Figure 1). To facilitate the subsequent PCR reaction, adroplet solution containing electrolytes and buffer (typically 1
M KCl, 10 mM MOPS, pH 7.0) was collected, with dilution by
1/2 in the PCR solution. We first attempted to create
calibration curves of the target DNA under these solution
conditions. The target DNA and PCR primers were selectedfrom a random sequence using Primer3Plus (https://primer3plus.com/). We confirmed the formation of secondary
structures and self-pairing of all primers using Primer3Plus.
The design parameters of primers and target DNAs were
determined as follows: the lengths of primers and target DNAs
were 17 to 25 and 70 to 80 nt, respectively. The GC-content
and Tm value were 40 to 60% and 60 to 65 °C, respectively. As
a result, six different target DNA sequences were designed(Table S1). Then, we performed qPCR assays for all
sequences. When using DNA #1, #2, #3, #4, #5, and #6 at
low concentrations of less than 105 molecules, nonspecificamplification such as primer dimers was observed, and we did
not obtain linear calibration curves. Even primers designed
using mathematical models sometimes form nonspecificproducts in negative controls and require PCR optimization.37,38
As an initial trial, the nanopore filter experiment was
performed using six different types of ssDNA, DNA #1−6,
under conventional conditions for nanopore sensing. Clear
blocking signals of ssDNA (160 nM) translocation events wereobserved for the buffer condition of 1 M KCl, 10 mM MOPS,
pH 7.0. The result for DNA #2 serves as a typical example forall DNA #1−6. First, we pipetted out the solution in the transside chamber after observing 30 blockage signals (identified as
the translocation of 30 target DNA molecules) (Figure 2a).
Then, we quantified the copy number of the DNA in thesolution by qPCR assays. As a result, we obtained amplification
curves of DNA #2 comparable to a positive control experimentusing 106 molecules of the DNA (Figure 2b). As the 30
molecules of DNA were detected to be more than 30,000 times
the number of molecules, we concluded that the preparation ofcalibration curves and DNA contamination were significant
issues in this experiment. In the following section, we aimed tosolve these problems regarding (i) the difficulty in obtaining
calibration curves in the low-concentration range and (ii) thelack of a negative control and contamination of the target
DNA.
Problem (i): Optimization of Experimental Conditions for Generating the Calibration Curve in the qPCR
Assay. To generate a calibration curve in the lowconcentration range, we utilized a new DNA #7 as the target
DNA instead of DNA #1 to #6. DNA #7 originated fromhepatitis E virus genomes, and its calibration curve waspreviously obtained in the range of 10−106 molecules (TableS2 and Figure 2c).39 The length of the primers for DNA #7

1. Separation of the experimental space: We divided theoperation area into a place to handle the template DNA(outside of a laminar flow cabinet) and a place to use
other solutions (inside the laminar flow cabinet) toprevent contamination of the template DNA, as reported
previously.41
2. Consideration of experimental instruments: Since DNA
solution and aerosol may enter the pipette,41 we utilized
micropipette tips with a hydrophobic porous filter tohandle the PCR solutions. Thus, the DNA solution andaerosol were blocked by the filter in the pipette tip,
preventing contamination. In addition, we used
segregated pipettes for handling the PCR solution andthe template DNA.
3. Decontamination of work-areas: Before using thelaminar flow cabinet, exposure to ultraviolet (UV) light
was carried out for 15 min to degrade DNA molecules inDhttps://doi.org/10.1021/acs.analchem.3c00573
Anal. Chem. XXXX, XXX, XXX−XXX

Analytical Chemistrypubs.acs.org/ac

ArticleFigure 3. (a) Amplification curves of the negative control before and after improvements. In the improved situation, there was no DNA
amplification after 45 cycles. (b) DNA numbers obtained from controlled experiments. The DNA numbers increased with increasing concentrationof DNA added to the droplet. (c) Amplification curves with and without an applied negative transmembrane potential (−180 mV) and negatively
charged lipid in the oil. The Ct value were 14.0, 17.9, and 15.6 for the normal, −180 mV, and the negatively charged lipid conditions, respectively.

between DNA and DOPG was expected. DOPG and DPhPC(1:1, mol/mol) in n-decane were mixed to form the pBLM as amembrane of solely DOPG was not stable. As a result of thecontrolled experiment performed with 500 nM target DNA,
contamination of 107 DNA molecules was still observed(Figure 3b). Moreover, the applied potential was inverted andapplied as −180 mV to the collecting side (droplet-B) to
prevent electrophoretic transport of DNA molecules; however,
we did not observe a critical improvement in the DNA
decontamination (Figure 3c). The DNA contaminations
therefore could not be prevented by changing the phospholipids and the applied potential.
Problem (ii): Nanopore Filter Experiments Using the
CBB Method. The results suggest that DNA contamination
may originate from the lipid membrane and lipid/oil solution.
The thin layer of a lipid membrane may constitute the primarypathway for contamination. To investigate this further, we
employed the CBB method to form pBLMs (Figure 4a).33,34
This method involves the formation of pBLMs by bringing two
water-in-oil emulsions, prepared by bubbling water into an oil/lipid mixture with glass micropipettes, into contact. The size ofthese emulsions is significantly smaller (approximately 50 μm

the cabinet. UV lights are classified into UV-A
(wavelength: 315−400 nm), UV-B (wavelength: 280−
315 nm), and UV-C (wavelength: under 280 nm)
according to their wavelength. Since DNA absorbs
wavelengths of light between 240 and 280 nm,42 we
exposed the work area to UV light that includes UV-C.
In addition, a decontamination reagent for DNA (PCR
Clean; Minerva Biolabs Berlin, Germany) and sodium
hypochlorite solution43 were sprayed on work areas
before experiments. We evaluated the effect of these
improvements by comparing the PCR results of thenegative control, where the target DNA was not added,
before and after these improvements. As a result,although a Ct value of 33.96 was obtained before these
improvements, which indicates 21 DNA molecules were
included in the solution, no amplification of DNA wasobserved after improving the experimental environment(Figure 3a). Thus, DNA contamination from the air wascompletely prevented by these improvements of thework area. The number of contaminated DNA
molecules from the air (21 molecules) was muchsmaller than the result of the nanopore filter experiments(106 molecules). This result suggests that there isanother pathway of contamination other than the air.
Prevention of ssDNA Permeation through the Lipid
Bilayer Using a Negatively Charged Lipid Membrane. We
next investigated DNA contamination during the nanopore
filter experiments. To confirm DNA contamination during theexperiment, we performed controlled experiments without
nanopores as follows: we prepared only pBLMs with two
droplets, one containing the target DNA with a concentrationof 500 nM, 1 nM, 1 pM, 1 fM, or 0 M (droplet-A) and theother without the target DNA (droplet-B). We then sampledfrom droplet-B 10 min after the pBLM was formed. The pBLM
formation was confirmed by measuring the membrane
capacitance, which we monitored to ensure that pBLMs did
not rupture during the experiments. The number of moleculesin the collected droplet-B was evaluated by the qPCR assay.
The number of DNA molecules in droplet-B increased with anincrease in the DNA concentration added to droplet-A (Figure3b). The DNA molecules were considered to be contaminantspassed from another droplet through the oil/lipid mixture, in
addition to those from the air (21 molecules mentioned
above). We assume that there may be multiple pathways ofcontamination, with n-decane being one of the possibilities. To
prevent DNA contamination through the oil/lipid solution, thetypes of phospholipids were investigated. DOPG, a negatively
charged phospholipid, was used because electrostatic repulsion

Figure 4. (a) Schematic illustration of the CBB system. A lipid bilayer
is formed by the contact of two water droplets bubbled from glass
pipettes on the microscope. (b) Microscopic image during pBLM
formation by the CBB method. The diameters of the emulsion andthe lipid bilayer were around 50 and 5 μm, respectively. (c) Control
data of the nanopore filter experiments using the CBB method. One
droplet contained 100 nM (1011 molecules) or 0 M target DNA. 10
min after forming pBLMs, another droplet was collected, and thenumber of the target DNA was quantified by qPCR assay.
E

https://doi.org/10.1021/acs.analchem.3c00573
Anal. Chem. XXXX, XXX, XXX−XXX

Analytical Chemistrypubs.acs.org/ac

ArticleFigure 5. (a) Schematic illustration of the nanopore filter using PNA-DNA duplexes. Because the contaminated DNA is hydrolyzed with PNA,
PCR amplification is inhibited. (b) Typical recorded channel current obtained by translocating PNA-DNA duplexes through the αHL nanopore.
(c) Number of DNA molecules obtained from controlled experiments with and without 3 nM PNA. (d) Amplification curves and the quantitated
DNA numbers for the nanopore-filtered DNA solution and the control experiment with 10 pM DNA and 30 pM PNA. The R2 value is 0.9.

in diameter) than the droplets used in the DCM (approximately 2000 μm in diameter, Figure 4b), leading to a
reduction in the number of DNA molecules present in thedroplets by a factor of several ten thousand. If 100 nM target
DNA is used, the numbers of molecules in a droplet of the
DCM method and in an emulsion of the CBB method are
calculated to be roughly 3 × 1011 and 4 × 106 molecules,
respectively. The contact area between the emulsions and theoil/lipid mixture was around 5 μm in diameter (with a contact
area of about 2 × 10−5 mm2). In contrast, the DCM
experiments employed a parylene hole with a 100 μm diameterto separate the two droplets and reduce the lipid membrane
area (with a contact area of about 4 × 10−3 mm2). As a control
experiment, the pBLM was formed by bringing two water-inoil emulsions into contact, one with and one without the 100nM target DNA. After 10 min, the water-in-oil emulsion wascollected by aspiration and the DNA number was evaluated by
qPCR assay. Unfortunately, even this method yielded 109
DNA molecules (Figure 4c). The reason for this contamination may be attributed to the dissolution of DNA moleculesin the oil/lipid mixture during the preparation of theemulsions, as well as the collection of contaminated oil/lipid
mixture during the collection of the emulsions.
Problem (ii): Nanopore Filter Applied with the PCR
Clamp Method. After conducting several experiments, as
previously discussed, it became apparent that it was difficult toeliminate DNA contamination at the single-molecule level via
the oil/lipid mixture. We subsequently attempted to utilize a
PCR clamp method44,45 for exclusion of DNA contamination(as depicted in Figure 5a).
The mechanism is as follows: the PNA probe (5′TGATTCTCAGCCCTTCGC-3′) (as depicted in FigureS2), which possesses positive charges and a complementary
sequence to the center of the target DNA, is hybridized withthe target DNA. As the PNA-DNA duplexes translocate
through the nanopore, they unzip46,47 and the target DNA

moves to another droplet. In contrast, the PNA-DNA duplex
structure is maintained when PNA-DNA duplexes arecontaminated in the oil/lipid mixture. While the translocatedtarget DNA is amplified, the contaminated PNA-DNA
duplexes are not amplified, thus inhibiting PCR and allowing
for exclusion and disregard of DNA contamination. Prior to
conducting experiments, we defined the clamping efficiency(ηc) as the inhibitory efficiency of PCR by the PNA-DNA
duplex.
ij ni na yz
jzz × 100zz
k ni {

jj
c = j

(1)

where ni and na are the number of initial PNA-DNA duplexes
and amplified DNA, respectively. We prepared a solution inwhich PNA and DNA were mixed at a molar ratio of 1:1 anddiluted the solution to contain between 10 and 105 PNA-DNA
duplexes. We then performed qPCR assays and evaluated theamplified copy number (as listed in Table S3). Although the ηc
was greater than 90% at high concentrations, the ηc at low
concentrations was lower than at higher numbers of PNADNA duplexes. We subsequently determined that this decreasein efficiency may be attributed to PNA adsorption on thesurface of microtubes and pipette tips during pipetting, as
determined through UV−vis spectroscopy measurements of
PNA concentration before and after dilution (Table S4). To
address this issue, we altered the PNA/DNA concentration
ratio to 3:1, 10:1, and 100:1 (mol/mol) and reevaluated theclamping efficiency. As a result, the clamping efficiency
improved at every concentration ratio and reached saturation
at a molar ratio greater than 3:1 (as summarized in Table S5).
We next confirmed that PNA-DNA duplexes could
successfully translocate with unzipping through αHL nanopores, as evidenced by the observation of blockage signals
indicating such translocation (as depicted in Figure 5b). It is
important to note that PNA itself does not possess charged
phosphate groups, and the electrophoretic force acting on PNA
F

https://doi.org/10.1021/acs.analchem.3c00573
Anal. Chem. XXXX, XXX, XXX−XXX

Analytical Chemistrypubs.acs.org/ac

Article■is significantly weaker than on DNA.48 Consequently, few PNA
translocation signals would be expected in nanopore experiments. In order to investigate contamination through the oil/lipid mixture in the device, we performed a control experimentusing only pBLM without nanopores. PNA (3 nM) and target
DNA (1 nM) were added to droplet-A. After 10 min, thesolution of droplet-B was collected, and the copy number of
DNA analyzed via qPCR assay. The results indicated that DNA
contamination was reduced by 99.98% in comparison to when
PNA was not added (as depicted in Figure 5c). Despite thisreduction, 105 contaminant molecules were still present evenin these conditions. To further reduce the number ofcontaminated DNA molecules for single-molecule separation,
we lowered the added DNA concentration from 1 nM to 10
pM, as this reduced contamination when testing the CBB
method. Under these conditions, the amount of contaminatedDNA was dramatically reduced; only four DNA molecules
were obtained from the droplet, within an acceptable range fortolerance (as depicted in Figure 5d).
Building upon these results, we next sought to conduct thenanopore filter experiment. It is important to note that utilizing
solutions with low DNA concentrations in nanopore experiments can result in a reduction of translocation frequency.31
To address this, we employed asymmetric salt conditions in
the droplets to enhance the frequency of translocation. In thenanopore filter experiments, we counted the different numberof blockage signals that corresponded to the translocation of29, 46, and 89 DNA molecules and subsequently analyzed thedroplet-B via qPCR. The results indicated that the DNA
numbers obtained were 67, 184, and 222 from each experiment(as shown in Figure 5d) with a Tm value of 83.3 ± 0.10 °C,
indicating the elimination of the amplification from otherartifacts, and a correlation between the nanopore-filtered andquantitated DNA numbers was observed (as depicted in Figure5d). It was found that the quantification of DNA numbers via
qPCR yielded higher results than those obtained throughnanopore filtration. This discrepancy may be attributed to themigration of unclamped DNA in droplet-A (calculated to bearound 105 molecules, Table S5) to droplet-B by a number offactors, including lipid/oil, air, or the collection step. While theresults obtained through qPCR and filtration were not entirely
congruent, a correlation coefficient of 0.9 with a linear
relationship was observed, and the difference in values wasless than 50% in the region from 0 to 100 molecules (see Text
S1).
Conducting the nanopore filter experiments necessitated theuse of a brand new, uncontaminated device for eachexperiment to prevent contamination. Additionally, in order
to count and collect the molecules that passed through thenanopore, we needed to (i) form a stable lipid bilayer within
the device without rupturing it, (ii) reconstitute a single
nanopore within the lipid bilayer, and (iii) collect the solution
from the chamber without disrupting the integrity of the lipid
bilayer. In this study, we performed a total of 68 nanopore
filter experiments. Of these, 60 experiments (88%) resulted inmembrane rupture or failure to form a stable bilayer, and 4
experiments (6%) resulted in the formation of a lipid bilayer
but with the presence of more than one pore or no pore at all.
Only in 4 experiments (6%) were we able to successfullycollect the solution from the nanopore filter. The current
challenges associated with conducting nanopore filter experiments thus highlight the need for technological advancements
to make nanopores a viable filter system.

CONCLUSIONS
Biological nanopores have garnered significant attention as
highly sensitive DNA and RNA sensors and have already been
implemented as DNA sequencers.49−51 In this study, we
focused on the single-molecule sensitivity of biological
nanopore sensors and developed a biological nanopore filteringsystem capable of counting the number of filtered DNA
molecules from the blockage signals. For the development of ananopore filter system with single-molecule accuracy, it isessential to completely prevent DNA contamination. We here
investigated two different pathways of DNA contamination:from the air and through the oil/lipid mixture. To prevent
DNA contamination from the air, we implemented “areaseparation,” “improvement of experimental instrumentation,”and “decontamination of work areas” and were able tosuccessfully reduce DNA contamination from the air.
However, despite our efforts to reduce DNA contamination
through the oil/lipid mixture by using negatively charged
phospholipids, applying reversed transmembrane potentials,
and forming pBLMs via the CBB method, DNA contamination
was not significantly reduced. We then attempted to eliminate
the contamination by utilizing PNA-DNA duplexes that were
not amplified in a PCR assay. In this system, PNA-DNA
duplexes are unzipped when passing through the nanopores,
resulting in the filtered solution containing only singlestranded target DNA. In contrast, PNA-DNA duplexes
contaminated through the oil/lipid mixture preserve their
duplex structure. The PCR amplification of PNA-DNA
duplexes is inhibited by the PCR clamp mechanism, thereby
allowing only the filtered target DNAs to be amplified. Using
this system, we observed a linear relationship between thenumber of counted molecules and the qPCR assay in the rangeof 0−100 molecules in the solution.

■

ASSOCIATED CONTENT

sı Supporting Information*

The Supporting Information is available free of charge athttps://pubs.acs.org/doi/10.1021/acs.analchem.3c00573.
Results of melting curve analysis and gel electrophoresis
of DNA #7, design of the PNA probe, sequence ofDNAs, sequence of DNA #7 primers, clamp efficiency of
PNA, PNA concentration before and after dilution, andcomparison of nanopore filters and qPCR (PDF)■

AUTHOR INFORMATION

Corresponding Authors

Kan Shoji − Department of Biotechnology and Life Science,
Tokyo University of Agriculture and Technology, Koganei,
Tokyo 184-8588, Japan; Department of MechanicalEngineering, Nagaoka University of Technology, Nagaoka,
Niigata 940-2188, Japan; orcid.org/0000-0002-71989683; Email: kshoji@mech.nagaokaut.ac.jp
Ryuji Kawano − Department of Biotechnology and Life
Science, Tokyo University of Agriculture and Technology,
Koganei, Tokyo 184-8588, Japan; orcid.org/0000-00016523-0649; Email: rjkawano@cc.tuat.ac.jp
Authors

Asuka Tada − Department of Biotechnology and Life Science,
Tokyo University of Agriculture and Technology, Koganei,
Tokyo 184-8588, JapanG

https://doi.org/10.1021/acs.analchem.3c00573
Anal. Chem. XXXX, XXX, XXX−XXX

Analytical Chemistrypubs.acs.org/ac

Nanami Takeuchi − Department of Biotechnology and Life
Science, Tokyo University of Agriculture and Technology,
Koganei, Tokyo 184-8588, Japan(24) Chen, Y. Q.; Song, Y. Y.; Wu, F.; Liu, W. T.; Fu, B. S.; Feng, B.
K.; Zhou, X. Chem. Commun. 2015, 51, 6980−6983.
(25) Benenson, Y.; Gil, B.; Ben-Dor, U.; Adar, R.; Shapiro, E. Nature
2004, 429, 423−429.
(26) Park, K. S.; Seo, M. W.; Jung, C.; Lee, J. Y.; Park, H. G. Small
2012, 8, 2203−2212.
(27) Hwang, W. L.; Holden, M. A.; White, S.; Bayley, H. J. Am.
Chem. Soc. 2007, 129, 11854−11864.
(28) Tsuji, Y.; Kawano, R.; Osaki, T.; Kamiya, K.; Miki, N.;
Takeuchi, S. Anal. Chem. 2013, 85, 10913−10919.
(29) Heron, A. J.; Thompson, J. R.; Cronin, B.; Bayley, H.; Wallace,
M. I. J. Am. Chem. Soc. 2009, 131, 1652−1653.
(30) Funakoshi, K.; Suzuki, H.; Takeuchi, S. Anal. Chem. 2006, 78,
8169−8174.
(31) Zhang, H. L.; Hiratani, M.; Nagaoka, K.; Kawano, R. Nanoscale
2017, 9, 16124−16127.
(32) Ohara, M.; Takinoue, M.; Kawano, R. ACS Synth. Biol. 2017, 6,
1427−1432.
(33) Iwamoto, M.; Oiki, S. Sci. Rep. 2015, 5, 9110.
(34) Iwamoto, M.; Oiki, S. Sci. Rep. 2017, 7, 6857.
(35) Meller, A.; Nivon, L.; Branton, D. Phys. Rev. Lett. 2001, 86,3435−3438.
(36) Mathe, J.; Visram, H.; Viasnoff, V.; Rabin, Y.; Meller, A.
Biophys. J. 2004, 87, 3205−3212.
(37) Garafutdinov, R. R.; Galimova, A. A.; Sakhabutdinova, A. R.
Nucleosides, Nucleotides Nucleic Acids 2020, 39, 1251−1269.
(38) Garafutdinov, R. R.; Galimova, A. A.; Sakhabutdinova, A. R.
Anal. Biochem. 2017, 518, 126−133.
(39) Jothikumar, N.; Cromeans, T. L.; Robertson, B. H.; Meng, X. J.;
Hill, V. R. J. Virol. Methods 2006, 131, 65−71.
(40) Elizabeth Van Pelt-Verkuil, A. v. B.; Heys, J. P. Principles andTechnical Aspects of PCR Amplification; Springer Science + BusinessMedia B.V., 2018.
(41) Dieffenbach, C. W.; Dveksler, G. S. Genome Res. 1993, 3, S2−
S7.
(42) Grskovic, B.; Zrnec, D.; Popovic, M.; Petek, M. J.; Primorac,
D.; Mrsic, G. Croat. Med. J. 2013, 54, 263−271.
(43) Hawkins, C. L.; Davies, M. J. Chem. Res. Toxicol. 2002, 15, 83−92.
(44) Araki, T.; Shimizu, K.; Nakamura, K.; Nakamura, T.; Mitani, Y.;
Obayashi, K.; Fujita, Y.; Kakegawa, S.; Miyamae, Y.; Kaira, K.; et al. J.
Mol. Diagn. 2010, 12, 118−124.
(45) Chiou, C. C.; Luo, J. D.; Chen, T. L. Nat. Protoc. 2006, 1,
2604−2612.
(46) Perera, R. T.; Fleming, A. M.; Peterson, A. M.; Heemstra, J. M.;
Burrows, C. J.; White, H. S. Biophys. J. 2016, 110, 306−314.
(47) Ciuca, A.; Asandei, A.; Schiopu, I.; Apetrei, A.; Mereuta, L.;
Seo, C. H.; Park, Y.; Luchian, T. Anal. Chem. 2018, 90, 7682−7690.
(48) Dragomir, I. S.; Bucataru, I. C.; Schiopu, I.; Luchian, T. Anal.
Chem. 2020, 92, 7800−7807.
(49) Ju, Y.; Pu, M. J.; Sun, K.; Song, G. Q.; Geng, J. Chem.�Asian J.
2022, 17, No. e202200774.
(50) Sun, K.; Liu, Y.; Zhou, X.; Yin, C. L.; Zhang, P. J.; Yang, Q. Q.;
Mao, L. F.; Shentu, X. P.; Yu, X. P. Front. Microbiol. 2022, 13, 939666.
(51) Werner, D.; Acharya, K.; Blackburn, A.; Zan, R. X.; Plaimart, J.;
Allen, B.; Mgana, S. M.; Sabai, S. M.; Halla, F. F.; Massawa, S. M.;
et al. Water 2022, 14, 2491.

Complete contact information is available at:https://pubs.acs.org/10.1021/acs.analchem.3c00573
Author Contributions

A.T. and N.T. contributed equally to this work. R.K. conceived
the original idea. T.A. performed all experiments, and T.A. and
N.T. conducted the analyses. A.T., N.T., K.S., and R.K. wrote
the entire manuscript. All authors have given approval to thefinal version of the manuscript.
Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS
This research was partially supported by JSPS KAKENHI grant
numbers 15H00803 (R.K.), 19H00901 (R.K.), 19K15418(K.S.), 21H05229 (R.K.), and 22KJ1231 (N.T.) and the JST
FOREST Program Grant Number JPMJFR2028 (K.S.).

■

ArticleREFERENCES(1) Tan, C. S.; Fleming, A. M.; Ren, H.; Burrows, C. J.; White, H. S.
J. Am. Chem. Soc. 2018, 140, 14224−14234.
(2) Shoji, K.; Kawano, R.; White, R. J. ACS Nano 2019, 13, 2606−
2614.
(3) Denuga, S.; Whelan, D. E.; O’Neill, S. P.; Johnson, R. P.
Electrochem. Sci. Adv. 2022, 2, No. e2200001.
(4) Watanabe, H.; Gubbiotti, A.; Chinappi, M.; Takai, N.; Tanaka,
K.; Tsumoto, K.; Kawano, R. Anal. Chem. 2017, 89, 11269−11277.
(5) Miyagi, M.; Takiguchi, S.; Hakamada, K.; Yohda, M.; Kawano, R.
Proteomics 2022, 22, No. e2100070.
(6) Mayer, S. F.; Cao, C.; Dal Peraro, M. iScience 2022, 25, 104145.
(7) Zhang, M.; Chen, C.; Zhang, Y. J.; Geng, J. Proteins: Struct.,
Funct., Bioinf. 2022, 90, 1786−1799.
(8) Cao, C.; Long, Y. T. Acc. Chem. Res. 2018, 51, 331−341.
(9) Kasianowicz, J. J.; Brandin, E.; Branton, D.; Deamer, D. W. Proc.
Natl. Acad. Sci. U.S.A. 1996, 93, 13770−13773.
(10) Stoddart, D.; Franceschini, L.; Heron, A. J.; Bayley, H.; Maglia,
G. Nanotechnology 2015, 26, 084002.
(11) Stoddart, D.; Heron, A. J.; Klingelhoefer, J.; Mikhailova, E.;
Maglia, G.; Bayley, H. Nano Lett. 2010, 10, 3633−3637.
(12) Stoddart, D.; Heron, A. J.; Mikhailova, E.; Maglia, G.; Bayley,
H. Proc. Natl. Acad. Sci. U.S.A. 2009, 106, 7702−7707.
(13) Cherf, G. M.; Lieberman, K. R.; Rashid, H.; Lam, C. E.;
Karplus, K.; Akeson, M. Nat. Biotechnol. 2012, 30, 344−348.
(14) Manrao, E. A.; Derrington, I. M.; Laszlo, A. H.; Langford, K.
W.; Hopper, M. K.; Gillgren, N.; Pavlenok, M.; Niederweis, M.;
Gundlach, J. H. Nat. Biotechnol. 2012, 30, 349−353.
(15) Goyal, P.; Krasteva, P. V.; Van Gerven, N.; Gubellini, F.; Vanden Broeck, I.; Troupiotis-Tsailaki, A.; Jonckheere, W.; PehauArnaudet, G.; Pinkner, J. S.; Chapman, M. R.; et al. Nature 2014, 516,
250−253.
(16) Hiratani, M.; Ohara, M.; Kawano, R. Anal. Chem. 2017, 89,2312−2317.
(17) Yasuga, H.; Kawano, R.; Takinoue, M.; Tsuji, Y.; Osaki, T.;
Kamiya, K.; Miki, N.; Takeuchi, S. PLoS One 2016, 11, No. e0149667.
(18) Yasuga, H.; Inoue, K.; Kawano, R.; Takinoue, M.; Osaki, T.;
Kamiya, K.; Miki, N.; Takeuchi, S. PLoS One 2017, 12, No. e0180876.
(19) Kawano, R. Biotechnol. J. 2018, 13, 1800091.
(20) Hiratani, M.; Kawano, R. Anal. Chem. 2018, 90, 8531−8537.
(21) Takeuchi, N.; Hiratani, M.; Kawano, R. JACS Au 2022, 2,
1829−1838.
(22) Adleman, L. M. Science 1994, 266, 1021−1024.
(23) Benenson, Y.; Paz-Elizur, T.; Adar, R.; Keinan, E.; Livneh, Z.;
Shapiro, E. Nature 2001, 414, 430−434.
Hhttps://doi.org/10.1021/acs.analchem.3c00573
Anal. Chem. XXXX, XXX, XXX−XXX