Cell-Free Expression of De Novo Designed
Peptides That Form β‑Barrel Nanopores
Shoko Fujita, Izuru Kawamura, and Ryuji Kawano*

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sı Supporting Information*

ABSTRACT: Nanopore sensing has attracted much attention
as a rapid, simple, and label-free single-molecule detection
technology. To apply nanopore sensing to extensive targets
including polypeptides, nanopores are required to have a sizeand structure suitable for the target. We recently designed a denovo β-barrel peptide nanopore (SVG28) that constructs astable and monodispersely sized nanopore. To develop the sizes
and functionality of peptide nanopores, systematic explorationis required. Here we attempt to use a cell-free synthesis systemthat can readily express peptides using transcription and translation. Hydrophilic variants of SVG28 were designed andexpressed by the PURE system. The peptides form a monodispersely sized nanopore, with a diameter 1.1 or 1.5 nm smaller
than that of SVG28. Such cell-free synthesizable peptide nanopores have the potential to enable the systematic custom designof nanopores and comprehensive sequence screening of nanopore-forming peptides.
KEYWORDS: nanopore, cell-free synthesis, lipid bilayer, peptide sensing, molecular dynamics simulations

P

design of nanopore-forming materials is a promising method toenable custom-made nanopores for a given target molecule,leading to recent reports on the de novo design of nanoporeforming proteins/peptides. For example, a de novo α-helicalbarreled peptide ion channel and nanopore have beenreported.30−32 Baker and co-workers have reported a β-barrel
protein with eight strands, engineered using computationalsequence design with several strategies and optimization of theloop sequence.33
We previously reported a de novo designed β-hairpin peptide(SVG28) that assembles to form a β-barrel structure in the
BLM with a diameter of predominantly 1.7 nm, and with theability to discriminate poly-L-lysine of different chain lengths.34
To functionalize and improve the resolution of the detection
signals for future polypeptide detection, we would like toinvestigate systematic design based on SVG28 sequences andto evaluate their capabilities. The SVG28 has 28 residues and issynthesized by solid-phase synthesis. The high hydrophobicity
and aggregation propensity of SVG28 make it difficult to purify
using typical methods; therefore, the isoacyl dipeptides
method35 was needed to synthesize SVG28. The complex

ore-forming proteins reconstituted in a bilayer lipid
membrane (BLM) offer characteristic information onmolecules passing electrophoretically through the pore,
via the level of the blocking current. There has been
considerable attention given to this nanopore sensing
technology because of its potential as a powerful tool forlabel-free single-molecule detection.1−5 The α-hemolysin(αHL) nanopore, which is derived from Staphylococcus aureus
and is one of the most commonly used nanopore proteins todate, has been applied in the single-molecule recognition of
nucleotides,6−10 peptides,11−14 proteins,15,16 etc.17,18Mycobacterium smegmatis porin A (MspA) was reported as nearly ideal
for the detection of single-stranded nucleic acid strands19 andhas been investigated in the realization of a nanopore
sequencer. Based on these studies, the company Oxford
Nanopore Technology has recently commercialized a nanopore DNA sequencer using a CsgG nanopore instead of the
MspA nanopores.20 Having achieved DNA sequencing using
nanopore technology, the next target is the sequencing ofproteins that have amino acids with fewer structural differences
than DNA.21−23 Several attempts toward peptide and protein
sequencing using aerolysin,24 FraC,25 and MspA26,27 nanopores have recently been reported.
The selectivity of nanopore sensing is mainly dependent on
the pore configuration and the interaction between the poresurface and the target molecules. Although several poreforming proteins in nature have been investigated forpolypeptide detection,28,29 challenges remain. The bottom-up© XXXX The Authors. Published byAmerican Chemical Society

Received: August 10, 2022
Accepted: January 3, 2023

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ArticleFigure 1. Hydrophilic variants of SVG28. (A−F) Amino acid sequences of SVG28 (A) and the position of hydrophilic mutations in SVG28D4 (B), SVG28-N4 (C), SVG28-D2 (D), and SVG28-N2 (E) and the control with valine substitutions in the transmembrane region (F). The
β-sheet region predicted by MINNOU43 is highlighted by black lines. (G−L) Side and top view of final structure of 6-mer SVG28 (G),SVG28-D4 (H), SVG28-N4 (I), SVG28-D2 (J), SVG28-N2 (K), and the control (L) after 200 ns simulation. SVG28-D4, N4, D2, N2 and thecontrol have a starting methionine for cell-free synthesis.

and time-consuming procedures required for SVG28 synthesis
is a major challenge in the systematic sequence optimization,
so herein we attempted to use the cell-free protein synthesis
system, which enables the synthesis of proteins and peptides invitro using transcription−translation.
Since around 1960, arbitrary polypeptides have beensynthesized by adding mRNA or nucleic acid fractions to celllysate.36−38 However, peptides do not form folded structures
and tend to be degraded by proteases contained in the lysate,39
and therefore are not suitable in this cell-free synthesis.
Shimizu and co-workers have reported a reconstituted cell-freesynthesis system (PURE system) using the individually
purified proteins that contribute to transcription and translation, resulting in a higher yield than cell lysate.40 This systemdoes not contain protease and can also synthesize peptides.
In this study, we introduced hydrophilic substitutions to
SVG28 and expressed the designed peptides using the PURE
system. The pore-forming ability, pore size, and noise ofhydrophilic variants were analyzed using electrophysiology toevaluate the effect of the hydrophilic mutations. The
applicability of the hydrophilic nanopores to protein
sequencing was also verified by single molecule detection ofoligopeptides.

four different types of more hydrophilic variants of SVG28
which were mutated from Ser to Asp or Asn. Asp (D,hydrophobicity: −3.50) and Asn (N, −3.50) are morehydrophilic residues than that of Ser (S, −0.80)41 and haverelatively compact side chains compared to other hydrophilicamino acids. The four-residue mutated variants named SVG28D4 (Figure 1B) and SVG28-N4 (Figure 1C) and the tworesidue mutated variants named SVG28-D2 (Figure 1D) and
SVG28-N2 (Figure 1E) were designed. The hydrophilic
residues were introduced at the edge of the β-sheet region toprevent its structure collapse and to avoid the increase of porenoise which would result from adding amino acid residues in
the loop and terminal regions.42 To design SVG28-D2 and
SVG28-N2, two hydrophilic residues at the interior side of thepore were removed from the SVG28-D4 and SVG28-N4
structures, respectively.
We performed all-atom molecular dynamics (MD) simulations to confirm whether the barreled structure is stable in
the lipid membrane. The hexametric structure of each peptide
was embedded in a system with the same lipid compositionand salt concentration as in the experimental study. The pore
structure of each variant was retained for 200 ns (Figure 1G−K) compared to the control structure with all-valine
substitution in the transmembrane region (Figure 1F,L). The
detailed analysis such as the root-mean-square distance(RMSD), root-mean-square fluctuation (RMSF), and othercontrol structures are shown in Figure S4 and S5.
Cell-Free Synthesis and Confirmation of SVG28-D4,
SVG28-N4, SVG28-D2, and SVG28-N2. A reconstituted
cell-free synthesis system (PURE system) was used in thisstudy because it does not contain proteases that degrade thepeptide chain.39,40 We designed E. coli codon-optimized
synthetic genes of the mutants (Supporting Information) and

RESULTS AND DISCUSSION
Design of Hydrophilic Variants of SVG28. We initiallyinvestigated the cell-free synthesis of the most hydrophobic
peptide, SVG28 (Figure 1A), as we previously reported.34 The
synthesis of SVG28 was not confirmed using a MALDI-TOF/
MS measurement (Figure S3). The sequence of SVG28 issuggested to be too hydrophobic to obtain enough yield withthe cell-free system, or to purify without aggregation. To
synthesize peptides using the cell-free system, we redesigned
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ArticleFigure 2. Confirmation of peptide expression. (A−D) MALDI-TOF/MS mass spectra of SVG28-D4 (A), SVG28-N4 (B), SVG28-D2 (C), andSVG28-N2 (D) expressed using cell-free synthesis. Obs. mass and Calc. mass stand for observed and calculated [M + H]+ values of thepeptides. (E−G) CD spectra of SVG28-D4 (E), SVG28-N4 (F), and SVG28-D2 (G).

Figure 3. Analysis of signals obtained from SVG28, SVG28-D4, SVG28-N4, and SVG28-D2. (A−D) The typical current−time trace of
SVG28 synthesized by solid-phase synthesis (A) and SVG28-D4 (B), SVG28-N4 (C), SVG28-D2 (D) synthesized by cell-free synthesis.
(E,F) Step (E) and square-top (F) signals were defined as corresponding to stable pore formation. (G,H) Multi (G) and erratic (H) signals
were defined as corresponding to unstable pore formation. (I) The ratio of step, square-top, multi, and erratic signals from the channel
current analysis. SVG28 used here was synthesized by solid-phase synthesis.

tried to express them using the PURE system. MALDI-TOF/
MS measurements confirmed the expression of SVG28-D4,
N4, and D2 (Figure 2A−C), but not SVG28-N2 (Figure 2D).
Although the hydrophobicity index of Kyte and Doolittle,41
based on experimental data, indicates the same hydrophobicity
for Asp and Asn, only SVG28-D2 of the two-residue mutatedvariants was successfully synthesized. However, the hydrophobicity index of Engelman predicts that transmembrane αhelical structures have different hydrophobicity for Asn andAsp.44 This suggests that there was in fact a hydrophobicity
difference between SVG28-D2 and SVG28-N2 in this
synthesis.

The secondary structure was confirmed using circulardichroism (CD) spectroscopy. Figure 2E−G suggests formation of β-sheet-like structures in SVG28-D4, N4, and D2
with 0.2 mM n-dodecyl-β-D-maltoside (DDM). The expressedamount of each peptide was estimated by UV−Vis spectroscopy (Table S1, Figure S6), and these yields of peptides were
sufficient for a channel current measurement using our lipid
bilayer system, which only requires a small amount of sample.
The amount of expressed peptides was almost equal to that ofgreen fluorescent protein (GFP) and dihydrofolate reductase(DHFR) expressed using the PURE system (Table S2).45,46
Electrophysiological Measurements and Evaluationof the Pore-Formation. The pore-formation of SVG28-D4,Chttps://doi.org/10.1021/acsnano.2c07970
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ArticleFigure 4. Analysis of pore current of SVG28, SVG28-D4, SVG28-N4, and SVG28-D2. (A−D) Histograms of the current conductance of openlevels for SVG28 synthesized by solid-phase synthesis (A), SVG28-D4 (B), SVG28-N4 (C), and SVG28-D2 (D) at +50, 100, 150, and 200
mV. (E−H) The expanded current histogram in the range of 0−2 nS of SVG28 synthesized by solid-phase synthesis (E), SVG28-D4 (F),SVG28-N4 (G), and SVG28-D2 (H). (I−L) The histogram of the normalized RMS noise of SVG28 synthesized by solid-phase synthesis (I),SVG28-D4 (J), SVG28-N4 (K), and SVG28-D2 (L) at all voltages. (M−O) Visualization of the structure of SVG28-D4 (M), SVG28-N4 (N),and SVG28-D2 (O) by HOLE.49 The 11th and 17th residues (Asp of SVG28-D4, Asn of SVG28-N4, Ser of SVG28-D2) are visualized in red,showing that the substituted residues generate a constriction in the pentamer pore.

N4, and D2 were investigated by channel current measurements in the lipid bilayer system. The pore-opening states inwhich the current momentarily increased were observed for all
peptides (Figure 3A−D), indicating that their pore-forming
ability was retained despite the introduction of hydrophilicmutations. To determine the probability of opening stable
pores, we defined and classified the current signal into fourtypes:47 step and square-top signals corresponding to stablepore-formation (Figure 3E,F), and multiple and erratic signals
corresponding to unstable pore-formation (Figure 3G,H). The
definition of the signal classifications is described in the
Supporting Information (Figure S2). The ratio of stable poreformation for the hydrophilic mutant decreased compared tothat of SVG28 synthesized by solid-phase synthesis (Figure 3I
and S7). When looking at stable pore-formation, thehydrophilic mutation did not decrease in step signals,
associated with single stable pore formation, with 49% of
SVG28, 58% of SVG28-D4, 41% of SVG28-N4, and 37% of
SVG28-D2 under every voltage application. However, thesquare-top signals decreased, with 39% of SVG28, 3% of

SVG28-D4, 10% of SVG28-N4, and 5% of SVG28-D2 (Figure3I). The square-top signal would reflect the pore-formation
with intermittent monomer insertion and dissociation; therefore, the reduction of the square-top ratio suggests inhibitionof the larger pore-formation for these hydrophilic peptides.
The pore diameter of each nanopore was calculated by the
conductance of the initial open-pore current from the baseline
level, such as shown in Figure 3A. The histograms of the pore
conductance of each peptide are shown in Figure 4A,B and the
expanded histograms are shown in Figure 4E−H. The peakconductance of the SVG28-D4 and SVG28-N4 occurred ataround 0.4 nS (Figure 4F,G), and that of SVG28-D2 at around
0.4 nS and 0.8 nS (Figure 4H), which were less than that of
SVG28 at 1.1 nS34 (Figure 4A). We estimated pore size using
the relationship between the current conductance and the pore
diameter of natural β-barrel proteins as we previously
proposed34 (Figure S8). The SVG28-D4 and N4 were
estimated approximately to have a 5-mer structure with adiameter of ca. 1.1 nm, and the SVG28-D2 was estimated toform 5- and 6-mer structures with a diameter of ca. 1.1 and 1.5
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ArticleFigure 5. Single-molecule detection. (A) Illustration of a typical blocking current. (B,C) The current and time trace of SVG28-D2 without
R7G (B) and with 500 nM R7G (C). The upper dashed lines indicate the threshold for the R7G translocation events and lower lines indicate
the baseline (≅0 A). (D) Histogram of the blocking current after bootstrapping of peptide without R7G (control, red, 0.42 nS) and with
R7G under 100 mV (green, 0.46 nS). (E) The event frequency of the R7G translocation as a function of the applied voltage. The error bars
show the mean ± SE. (F) Pore diameter at each z-axis of the 5-, 6-, and 7-mer pore calculated using HOLE. The dashed lines indicate
approximate pore areas. (G) Cross section of the pore of 5-, 6-, and 7-mer SVG28. Average pore diameter was calculated using HOLE49 andthe peak conductance. The R7G structure was predicted by Alphafold2.50

nm, compared to the SVG28 7-mer structure with a diameterof 1.7 nm. The decreased size of the pore and the ratio ofsquare-top signals in the hydrophilic variant suggest that thedestabilization of the peptide prevented it from making large
and dynamic pore structures, also supporting our proposal of adecrease in the number of monomers.
We next considered the pore noise of the open-pore state.
The open state of the step currents seemed to show increased
current noise compared to that for the SVG28 pore (Figure
S9A). To assess the precise current noise of pores, we
attempted to quantify the noise using root-mean-square(RMS) analysis. The inherent RMS noise of a pore current
depends on the magnitude of the conductance due to thedifferences in pore resistance.29 In the precise analysis for thedifferent conductance of the signals, we made an approximate
curve that was obtained from the plot of RMS noise versus
conductance (n = 23), and the slope was used for theestimation (Figure S9C). The histogram of the RMS noise isshown in Figure 4I−L, and the law data is presented in Figure
S9B for reference. When applying this to the nanopore
measurements, we defined the pores with RMS noise of lessthan 4 to have sufficiently low noise and appropriate forrecognition of the translocation of single molecules. The orderof the ratio of the RMS with less than 4 was as follows:[SVG28-N4:31%] < [SVG28-D4:33%] < [SVG28-D2:34%] <[SVG28:38%], and overall RMS trends of SVG28-D2 was also
improved. This result suggests that the introduction ofhydrophilic residues destabilizes the pores in accordance with
the number of substitutions. In addition, SVG28-D4 and N4
have the mutation at the inside of the membrane-anchoringtyrosine48 (11th and 17th residues), which is positioned at theinterface between the hydrophilic lipid head and thehydrophobic lipid tail. The protrusion of bulky hydrophilic
residues (Asp and Asn) inside the barrel may have led toincreased noise based on the MD simulations (Figure 4M−O).
Single-Molecule Detection Using the SVG28-D2
Nanopore. The SVG28-D2 formed relatively large poresand had low noise, and thus we attempted use it to detect anoligopeptide. In a previous study, 13 of the 20 natural aminoacids positioned at the N-terminus of a heptaarginine (R7-X)
were identified using the aerolysin nanopore.24 To test thedetection capabilities of SVG28-D2, we used RRRRRRRG(R7G) as the target. The SVG28-D2 mainly forms a nanopore
with conductance at around 0.8 nS (Figure 4D) which weestimate to have a diameter of ca. 1.5 nm. Although theblocking signal was not observed in the nanopore with thisconductance, clear blocking signals (Figure 5A) appeared in
the case of much larger nanopores with conductance at around2 nS (Figure 5B,C). Because SVG28-D2 sometimes shows
intrinsic blocking-like noise signals, the precise identification of
the signals from the translocation of peptides was a challenge.
We compared the blocking amplitudes between the experiments with and without R7G, and investigated the event
frequency of the blocking signals against voltage applications.
As a result, the histogram of the blocking conductanceprovides the discrimination of the blocking signal and the porederived noise (Figure 5D and S10), and the frequency ofblocking events increased with an increase in applied voltage,suggesting that the blocking signals reflect the translocation ofR7G (Figure 5E). Although the large distribution in signal
duration makes it difficult to strictly discuss, the duration timeEhttps://doi.org/10.1021/acsnano.2c07970
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ArticleFigure 6. Single-molecule detection of another peptide, R7W. (A) The current and time trace of SVG28-D2 with 500 nM R7W. The upperdashed lines indicate the threshold for the R7W translocation events and lower lines indicate the baseline (≅0 A). (B) Scatter plots ofblocking current and duration time after bootstrapping of SVG28-D2 without R7G (control, red, 1.65 ms, 0.42 nS), with R7G under 50 mV(green, 1.30 ms, 0.45 nS), and R7W under 50 mV (orange, 1.86 ms, 0.59 nS) The center of the plot is represented by crosses. (C) The event
frequency of the noise of SVG28-D2 and the R7G and R7W translocation as a function of the applied voltage. The error bars show the mean
± SE.

of translocation signal under 100 mV was slightly longer thanunder 50 mV. See Figure S10 for details.
In order to detect R7G, a nanopore with diameter muchlarger than ca. 1.3 nm is required, based on the HOLE analysis
of MD simulations in this experiment (Figure 5F,G). The
results of the blocking signal observed only at higher
conductance also indicates that the decreased conductance ofhydrophilic variants was due to a decreasing number ofmonomers. The aerolysin nanopore previously used for thedetection of the same oligopeptide has at its narrowest a 1.0
nm constriction calculated by HOLE analysis, suggesting that alarger vestibule region for capturing a target molecule isneeded for frequent nanopore-sensing with narrow constrictions. Moreover, the upper three negative charges permonomer in SVG28-D2 have been suggested to prevent theinsertion of the cationic peptide into the pore, compared to thedetection using SVG28 (data not shown), and the reduction of
one positive charge at the pore exit have also contributed to
the faster translocation of the oligopeptide.
As in the previous study of aerolysin,24 the change in theblocking amplitude depending on the side chain volume was
examined using RRRRRRRW(R7W), which has a larger side
chain than R7G. Clear blocking signals were observed onlywith ca. 2 nS pore, similar to the single-molecule measurementof R7G (Figure 6A). The blocking conductance of R7W wasclearly discriminated from pore-derived noise (Figure 6B), andthe frequency of blocking events increased compared to porenoise (Figure 6C), suggesting the blocking signals reflect thetranslocation of R7W. The blocking amplitude was increased
with the increase of the volume of the side chain (Figure 6B),
indicating that the SVG28-D2 nanopore has the potential for
peptide discrimination at one amino acid resolution, although
this is not a measurement of its main conductance.

than that of SVG28, and the amount of pore noise increased
according to the number of hydrophilic mutations. Thechanges in conductance and pore noise due to the introductionof the mutations provides insight into the assembly state of βbarrels formed by the association of β-hairpins and provides
knowledge that can be applied in nanopore design. Singlemolecule detection of oligopeptides and discrimination ofthem at one amino acid resolution were achieved with a ca. 2
nS nanopore, but not with the main conductance of SVG28D2. However, through this sequencing experiment we
confirmed that our intended outcomes were successful in thedesign and fabrication of cell-free synthesizable nanopores.
Although previous synthesis of nanopore proteins using a cellfree system and with the addition of liposomes has beenreported,51,52 our cell-free synthesizable peptide nanopores do
not require lipid components and the pore properties can beeasily modulated through sequence modification, thus enabling
the easy custom design of nanopore sequences andcomprehensive sequence screening of de novo nanoporeforming β-hairpin peptides.

METHODS
Reagents and Chemicals. The following reagents were used:
Purefrex2.0 (GeneFrontier, Japan); GL-Tip SDB (GL Sciences,
Japan); Trifluoroacetic acid (TFA, FUJIFILM Wako Pure Chemical
Industries, Ltd. (Wako), Japan); acetonitrile (ACN, FUJIFILM Wako
Pure Chemical Industries, Ltd. (Wako), Japan), α-cyano-4-hydroxycinnamic acid (CHCA, Tokyo Chemical Industry, Japan); polyethylene Glycol 4,000 (PEG4000, Tokyo Chemical Industry, Japan);n-dodecyl-β-D-maltoside (DDM, FUJIFILM Wako Pure Chemical
Industries, Ltd. (Wako), Japan); 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC, Avanti Polar Lipids, USA); cholesterol (SigmaAldrich, USA); n-decane (FUJIFILM Wako Pure Chemical Industries,
Ltd. (Wako), Japan); 3-morpholinopropane-1-sulfonic acid (MOPS,
Nacalai Tesque, Japan); potassium chloride (KCl, Nacalai Tesque,
Japan); potassium hydroxide (KOH, FUJIFILM Wako Pure Chemical
Industries, Ltd. (Wako), Japan); dimethyl sulfoxide (DMSO,FUJIFILM Wako Pure Chemical Industries, Ltd. (Wako), Japan).
SVG28 was synthesized and purified by Fmoc synthesis. DNA
templates for cell-free synthesis were synthetic genes provided byIntegrated DNA Technologies, USA. DOPC was diluted to 10 mg/
mL in n-decane. Buffered electrolyte solutions were prepared fromultrapure water, which was obtained from a Milli-Q system (Millipore,
Billerica, USA). SVG28 was dissolved at a concentration of 100 μM in
ultrapure water:DMSO = 1:1 (v/v) and stored at −30 °C. The solidphase synthesis of SVG28 and confirmation of synthesis is described

CONCLUSION
We used a cell-free system to express hydrophilic mutants of
the SVG28 peptide and establish a method for exploring thesystematic sequence design of peptide nanopores. The
hydrophilic variants were designed with two or four
substitutions from Ser to Asp or Asn. A surprising result wasthat the expressed variants retained the pore-forming ability
despite the substitution of four (about 14%) and two (about7%) of the total of 28 residues with more hydrophilic aminoacids. The pore conductance of the variants converged lessFhttps://doi.org/10.1021/acsnano.2c07970
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in our previous paper.34 Products of cell-free synthesis were dissolved
at a concentration of 10 μM in ultrapure water and stored at −30 °C.
Molecular Dynamics Simulations. All-atom molecular dynamics (MD) simulations of the β-barrel structure of the SVG28, SVG28D4, SVG28-N4, SVG28-D2, and SVG28-N2 were performed toconfirm the pore-forming ability of each peptide in the bilayer lipid
membrane. The 3D initial structures of each peptide were modeled
via a homology modeling technique using SWISS-MODEL.53 HASR
protein (PDB ID: 3CSL; chain A; 484−511) was selected as atemplate structure for the peptides, in keeping with our previous
study. The obtained monomeric structure was carefully aligned into
4-, 5-, 6-, and 7-mer structures, which were used as the masterstructure. Alignment of each monomer to the master structure was
performed by USCF Chimera.54 Each polymeric structure was used as
the starting structure and the remaining components were preparedby the CHARMM-GUI membrane builder.55 Each polymeric
structure was embedded in DOPC:Cholesterol = 2:1 (99 lipids perleaflet) with 1 M KCl at 298 K. Systems were initially equilibratedusing the protocol provided by CHARMM-GUI followed by anadditional 500 ps of equilibration to allow for the system to fully relax,
with a total simulation time of 200 ns. All simulations were executedwith GROMACS-2021.256 using the CHARMM36m force field57 andrun on a computer with Ryzen7 CPU and RTX3070 GPU. The
trajectory file was analyzed using the standard analysis tools installedin GROMACS. HOLE software49 was used to evaluate the pore
diameter in the trajectory.
Cell-Free Synthesis of β-Hairpin Peptides. Purefrex2.0(GeneFrontier, Japan), a reconstituted cell-free synthesis system,40
was used for cell-free synthesis. Template DNAs for the translation
were synthesized to contain the T7 promoter sequence, ribosome
binding site (SD sequence), target protein sequence and terminator(Supporting Information). The template DNAs were dissolved at aconcentration of 10 ng/μL in DNase-free water (Millipore, USA) andstored at 4 °C. Expression was performed according to the protocol
provided at 37 °C for 6 h on a heat block, and 1 μL of DNA template
diluted to 0.3 ng/μL was added to the reaction mixture. After thetranslation, the peptides were desalted with GL-tip SDB (GLSciences, Japan). The washing solution was 0.1% (v/v) TFA andthe eluting solution was 80% (v/v) ACN, 20% (v/v) 0.1% TFA. At
first, the tip column was conditioned with 20 μL of eluting solution(3000g, 2 min) and then equilibrated with 20 μL of washing solution(3000g, 2 min). After that, the reaction mixture was added to thecolumn and centrifuged (3000g, 5 min) and the column was washedwith 20 μL of washing solution (3000g, 2 min). Finally, 100 μL of
eluting solution was added to the column, centrifuged (3000g, 3 min),and collected in a collection tube. Then the acetonitrile in thedesalted solution was removed under a vacuum, and the remaining
sample was lyophilized using a Freezone 6-PLUS (Asahi Life Science
Corporation, Japan).
Matrix-assisted laser desorption or ionization with time-of-flight(MALDI-TOF) mass spectroscopy analyses were carried out using
the MALDI-TOF autoflex speed (Bruker, USA) in linear/positivemode. PEG4000 was used for external mass calibration. The
lyophilized peptides for 20 μL of the reaction mixture were dissolvedin 3 μL of TA30 (ACN:0.1% TFA = 3:7, v/v). 100 μg α-cyano-4hydroxycinnamic acid (CHCA) dissolved in 100 μL TA30 and the
peptide sample were mixed in a sample−matrix ratio of 1:2 (v/v), and1 μL of the solution was dropped on the plate. To confirm the βhairpin structure of the expressed peptide, CD (circular dichroism)
measurements were performed using a J-1100 CD Spectrometer(JASCO, Japan). The lyophilized peptides for 20 μL of the reaction
mixture were dissolved in 2 μL of 0.2 mM DDM and 1 mM MOPS. A
quartz microsampling disc with a 0.2 mm path length was used for themeasurements. The spectra were acquired every 0.5 nm from 190 to
300 nm with 20 mdeg/0.05 dOD of CD scale, 1 nm of bandwidth,and four times integration. Molar residue ellipticity (MRE [deg cm2
dmol−1 res−1]) was calculated from the observed ellipticity (mdeg) by
the following equation:

MRE = Y ×

Article

1
1
1
1
×
×
×
1000
L
C
n

MRE: Molar Residue Ellipticity [deg cm2 dmol−1 res−1], Y: observed
ellipticity [mdeg], L: path length [cm] C: peptide concentration[dmol cm−3] n: residue length of peptide. For each data set, baselines
from the same buffer and sampling disc were subtracted, and theobserved ellipticity of the purified and lyophilized cell-free solution(by replacing the DNA solution with purified water) was also
subtracted. The signal was smoothed by the Savitzky−Golay methodwith a convolution width of 15.
To confirm the amount of expressed peptide, absorbancemeasurements were carried out using a Nanodrop 2000c (Thermo
Scientific, USA) in UV−Vis mode. The absorption at 205 nm derived
from the peptide bond is used for the estimation of peptide
concentration. The absorption coefficient of each peptide wascalculated from the amino acid sequence of the peptide.58 Eachpeptide solution was aliquoted to 10 μM in 20 μL and stored at −30°C.
Preparation of Microdevice for BLM Formation. The
microdevice was fabricated by machining a 6.0 nm thick, 10 × 10
mm poly(methyl methacrylate) (PMMA) plate (Mitsubisi Rayon,
Japan) using a computer-aided design and computer-aided
manufacturing three-dimensional modeling machine (MM-100,
Modia Systems, Japan) as shown in Figure S1A. Two wells (2.0 nm
diameter and 4.5 nm depth) and a chase between the wells were
manufactured on the PMMA plate (Figure S1B). Each well had athrough-hole in the bottom and Ag/AgCl electrodes set into theseholes (Figure S1A). A polymeric film made of parylene C(polychloro-p-xylylene, Parylene Japan, Japan) with a thickness of 5
μm was patterned with single pore (100 μm diameter) using theconventional photolithography method, and then put between
PMMA sheets (0.2 mm thick) (Figure S1C). The sheets enclosing
the parylene C film were inserted into the chase to separate the twochambers.
Bilayer Lipid Membrane Preparation Using the Microfabricated Device. Bilayer lipid membranes (BLMs) were preparedusing the drop contact method in the microdevice. In the drop
contact method as we have been developing,59−65 the two lipid
monolayers containing the aqueous droplets come into contact in asmall hole punched in the parylene film to form lipid bilayers (Figure
S1D). Lipid bilayers were formed as follows: first, 0.6 μL of
DOPC:Cholesterol = 4:1 (w/w) dissolved in n-decane was added into
both chambers. Then 4.7 μL of 1 M KCl and 10 mM MOPS wasadded to a chamber on the voltage applied side and 4.7 μL of 1 M
KCl, 10 mM MOPS, and 1 μM SVG or 5 μM SVG28-D4, SVG28-N4,
or SVG28-D2 was added to a chamber on the ground side. When the
BLM was broken, a thin hydrophobic stick was used to trace back thelipid and reconstitute the BLM.
Channel Current Measurements. The channel current wasmonitored using a JET patch clamp amplifier (Tecella, USA). Signals
were detected using a 4 kHz lowpass filter at sampling frequency of 20
kHz. A constant voltage of +50, +100, +150, and +200 mV wasapplied to the recording chamber, and the chamber on the groundside was grounded. Data were analyzed using Clampfit 11.1
(Molecular Devices, USA), Excel (Microsoft, USA), OriginPro 2022
(Light Store, Japan), and Python 3.9 (Python Software Foundation,
USA). The channel current measurement was performed at 22 ± 2°C.
Current Signal Analysis of the Peptide Nanopores. To
evaluate the pore-forming ability of the peptides, current signals wereclassified into stable pore-formation signals and unstable poreformation signals, using the signal classification method previously
proposed in our laboratory47 and used for SVG28.34 The current
signals were classified into four types: step, square-top, multi, anderratic. The step signal, representing stable single pore formation,shows a sharp signal rise in less than 10 ms, and the 95% confidence
interval of the pore current value less than 10 times that of thebaseline. A signal lasting longer than 1 s was classified as a step signal(Figure S2A). The square-top signal, representing stable poreG

https://doi.org/10.1021/acsnano.2c07970
ACS Nano XXXX, XXX, XXX−XXXACS Nanowww.acsnano.org

formation involving insertion and dissociation of monomers, shows asharp signal rise in less than 10 ms and shoes a staircase current, with
the 95% confidence interval of the pore current value less than 10times that of the baseline for each step (Figure S2B). The multi signal,
representing unstable pore formation, shows a sharp signal rise in lessthan 10 ms, and the 95% confidence interval of the pore current value
is more than 10 times that of the baseline (Figure S2C). The erratic
signal, representing membrane disruption, shows a blunt signal rise in
more than 10 ms (Figure S2D). In addition, signals longer than 1 s
were included in the analysis because the hydrophilic variants showed
many short duration signals of less than 1 s. The signal distribution
was calculated by taking n = 20 data of the obtained signal from eachmeasurement, in the order of the longest signal duration, and n = 50
data of the sum of them, in the order of the longest duration. The
pore conductance was calculated by dividing the current value of thestep signal by the applied voltage. The current value is the average ofthe current values for 1 s after pore opening. To estimate the pore
diameter and the number of monomers, the relationship between the
conductance of the natural β-barrel proteins CymA, OmpA, OmpF,
OmpG, FhuA, and Vdac and the reported pore diameters34 was used
for the calculation (Figure S8), instead of the commonly used Hille’s
equation.66 To quantify the noise in the step signal in detail, the RMS
noise was calculated by subtracting the unbiased variance of thebaseline for approximately 1 s before the pore opening, from theunbiased variance of the current value for approximately 1 s after thepore opening, and taking the square root of the result. Since this value
tends to increase in response to pore conductance, an approximate
straight line was drawn using all conductance and RMS data (n =352) and normalized to the RMS when the pore was at 1 nS.
Solid-Phase Synthesis of R7X Peptide. R7G and R7W peptides
were synthesized by standard 9-fluorenylmethoxycarbonyl (Fmoc)based solid phase peptide method using N-α-(9-fluorenylmethoxycarbonyl)-glycine and tryptophan p-methoxybenzyl alcohol resins(Fmoc-Gly-alko and Fmoc-Trp-alko resins) in the following
procedure. To a swollen Fmoc-X-alko resin (0.25 mmol/g) in
NMP using 10 mL PTFE reactor vial were added N-α-(9fluorenylmethoxycarbonyl)-N-ω-(2,2,4,6,7-pentamethyldihydrobenzofuran-5-sulfonyl)-L-Arginine (Fmoc-L-Arg(Pbf)-OH, 4 equiv), 1,3diisopropylcarbodiimide (DIC, 4 equiv), and 1-hydroxybenzotriazole
monohydrate (HOBt, 0.5 M, 4 equiv) and the reaction mixtures werestirred for 5 min at 75 °C using microwave-assisted peptide
synthesizer (Biotage Initiator+ Alstra). Fmoc groups were removed
by 20% piperidine in NMP. The resins were filtered, washed 3 times
using dichloromethane (DCM), and dried under a high vacuum. The
cleavage from resin and deprotection were performed by stirring with
TFA, H2O, thioanisole, 1,2-ethanedithiol (35/2/2/1, v/v/v/v) for
120 min. The collected filtrate was evaporated and precipitated withcold ether. Finally, crude peptide was extracted using acetonitrile/
pure water (v/v = 7:3) from the filtrate. R7X peptide was purified byreverse phase HPLC H-Class (Waters) with WATERS ACQUITYHSS T3 (C18) 1.8 μm (2.1 × 150); detection: 220 nm, column
temperature: 298 K, elution: 0.1% TFA in water and 0.1% TFA in an
acetonitrile gradient system. The peaks of R7G and R7W wereobtained at retention times of 7.267 and 6.186 min (Figure S11A,B).
The purified peptides were obtained after lyophilization with 99.3%(R7G) and 98.5% (R7W) HPLC purity. The product m/z values wereconfirmed as 1168.65 [M + H]+ and 1297.98 [M + H]+ by LC/MS
with electrospray ionization mode (Waters H-Class QDa) as areference of ideal m/z = 1168.75 [M + H]+ and 1297.81 [M + H]+(Figure S11C,D).
Single-Molecule Detection of the Peptide Nanopores. 4.7
μL of 1 M KCl, 10 mM MOPS and 500 nM NH2-RRRRRRRGCOOH (R7G) or NH2-RRRRRRRW-COOH (R7W) was added tothe chamber on the voltage-applied-side and 4.7 μL of 1 M KCl, 10mM MOPS and 5 μM SVG28-D2 was added to the ground side. A
constant voltage of +50 or +100 mV was applied to the recording
chamber, and the chamber on the ground side was grounded. Because
the SVG28-D2 nanopore with 2 nS almost shows noise-like signal
longer than 3 nS (78%), the data were filtered to its duration <3 ms to
extract the translocation signal. Since the number of available noise

Articlesignals of SVG28-D2 with 2 nS is too small for valid comparison,
signals with 1 nS pore were used as control. The duration time of thetranslocation signal is the length of time of the current deflected fromthe mean open pore current (Figure 5A). To avoid including porenoise, the signal is thresholded to take a current value that has
decreased beyond a given value. The bootstrap method is a method
based on the resampling of the original random sample, drawn from a
population with an unknown distribution. We used the exact
bootstrap method, which availed of the entire space of resamples.
In the exact bootstrap method, accuracy verification is possible when
the sample number is greater than 30.67 In this study, our bootstrap
procedure randomly took 30 samples from the primary common
translocation data with 65,536 replacements and calculated the mean
for these samples. Three hundred samples were randomly extracted
from the bootstrapped data to produce the scatter plot of duration
time versus current blocking conductance (Figure S10A).

ASSOCIATED CONTENT
sı Supporting Information*

The Supporting Information is available free of charge athttps://pubs.acs.org/doi/10.1021/acsnano.2c07970.
DNA sequences of peptides in cell-free expression,
detailed information on signal classification, pore
analysis and RMS normalization, additional results of
MALDI-TOF/MS, MD simulation, UV spectrometry
and signal classification and single-molecule detection,yield of each expressed peptide, examples of signals at
each RMS value, and confirmation of solid-phase
synthesis of R7X peptide (PDF)

AUTHOR INFORMATION
Corresponding Author

Ryuji Kawano − Department of Biotechnology and Life
Science, Tokyo University of Agriculture and Technology,
Tokyo 184-8588, Japan; orcid.org/0000-0001-65230649; Email: rjkawano@cc.tuat.ac.jp
Authors

Shoko Fujita − Department of Biotechnology and Life Science,
Tokyo University of Agriculture and Technology, Tokyo 1848588, Japan
Izuru Kawamura − Graduate School of Engineering Science,
Yokohama National University, Yokohama 240-8501,
Japan; orcid.org/0000-0002-8163-9695
Complete contact information is available at:https://pubs.acs.org/10.1021/acsnano.2c07970
Author Contributions

R.K. conceived the original idea. S.F. performed all the otherexperiments and analysis. I.K. performed solid-phase synthesisof R7X peptide. S.F. and R.K. wrote the entire manuscript. I.K.
wrote the experimental details and results. All authors havegiven approval to the final version of the manuscript.
Funding

JSPS KAKENHI Grant Number 19H00901, 21H05229, andJST-CREST JPMJCR21B2.
Notes

The authors declare no competing financial interest.

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