Triggered assembly transmembrane nucleic acid nanostructures

Abstract

A nucleic acid nanostructure is provided comprising a plurality of component modules, each component module comprising a nucleic acid sequence and at least one membrane anchor. The plurality of component modules can be controlled to assemble to form the nanostructure in response to an external stimulus, and can also penetrate the semi-fluid membrane upon or after controlled assembly. Methods of assembling the nanostructures and uses in sensors, drug delivery, and imaging substance release are also provided.

Description

Technical Field

[0001] The present invention relates to novel nucleic acid nanostructures and their uses. In particular, it relates to nucleic acids that can act as dynamic functional membrane-associated nanopores. Background Art

[0002] DNA nanostructures have shown the potential to advance nanotechnology and life sciences. Compared with other materials, DNA nanostructures have a highly controllable structure based on predictable folding using base pairing rules (Rothemund P.W. Nature 440, 297-302 (2006), Seeman, NC; Sleiman, HF Nat. Rev. Mater. (2017), 3, 17068; Hong, F. et al., Chem. Rev. 2017, 117, 12584-12640; Praetorius, F. et al., Nature (2017), 552, 84-87; Sacca, B.; Niemeyer, CM Angew. Chem. Int. Ed. 2012, 51, 58-66). By utilizing these characteristics, functional DNA nanostructures are increasingly designed to be beneficial to fields other than DNA nanotechnology. Examples include DNA scaffolds, which precisely position proteins and other biomolecular assemblies for research applications in biophysics and molecular biology. In addition, predictable changes in DNA nanostructures have been used as smart biosensing devices for measuring intracellular pH (Bhatia, D. et al., Nat. Commun. (2011), 2, 339) or cellular DNA nanocages for delivering bioactive substances (Walsh, AS et al., ACS Nano (2011), 5, 5427-5432).

[0003] Replicating complex biological functions by simple and tunable synthetic means has attracted great interest in science and technology. Countless biological nanopores and other membrane proteins are powerful inspirations for this endeavor. By forming water-filled channels, protein nanopores shuttle bioactive substances across cell membranes and provide scientific insights into transport and molecular interactions within confined spaces. Such nanopores have found applications in portable and scalable DNA sequencing devices by allowing a single nucleic acid strand to pass through a reading head. Nanopores are also used for sensing of non-DNA analytes. Reflecting these advantages, synthetic nanopores have been created with self-assembling peptides, organic molecules, or solid-state inorganic materials to expand the sensing range and understand how transport is affected by pore chemistry, shape, and size that are not accessible in biology (e.g., see Howorka, S. Nat. Nanotechnol. 2017, 12, 619-630; Xue, L. et al., Nat. Rev. Mater. 2020, 5, 931-951; and Wei, RS et al., Nat. Nanotechnol. 2012, 7, 257-263).

[0004] However, the traditional configuration of transmembrane nanopores is usually constitutively open, which limits their functional complexity. To address this limitation, barrel-shaped pores are equipped with a lid that can be removed or opened upon external stimulation. However, such pores are rarely completely closed, allowing a small amount of current to flow from the cis side of the membrane to the trans side or even laterally across the membrane as long as the nanostructure penetrates and spans the membrane, which can affect sensing applications. In addition, even in the nominally closed state, the presence of a constitutively open pore lumen allows material to leak out or leads to further leakage during insertion into bilayers and associated semifluid membranes. Understandably, leakage reduces the potential of the pore for use in analyte sensing, which can lead to an increase in background noise and also reduce its application in drug delivery or targeted cell lysis.

[0005] There is a need to further develop further improved and optimized methods and compositions for assembling and inserting nanopores into bilayer and semifluid membranes. The present invention addresses deficiencies in the art. These and other uses, features and advantages of the present invention should be clear to those skilled in the art from the teachings provided herein. Summary of the invention

[0006] In various embodiments, the present invention provides DNA nanotechnology to construct functional advanced membrane pores, which are assembled from multiple membrane surface-associated subunits after defined trigger activation. The controlled formation of nanopores integrates the molecular recognition between trigger and inactive subunits, the repositioning of activated subunits within the membrane, and their assembly into a functional transmembrane channel.

[0007] A first aspect of the present invention provides a nucleic acid nanostructure comprising:

[0008] a plurality of component modules, each component module comprising a nucleic acid sequence and at least one membrane anchor,

[0009] wherein the plurality of component modules are capable of controlled assembly in response to external stimuli to form the nanostructure; and

[0010] The nanostructures are configured to penetrate the semifluid membrane upon or after controlled assembly.

[0011] Optionally, the plurality of component modules can be associated with the semifluid membrane before or after controlled assembly. Suitably, the nanostructures are configured to penetrate the semifluid membrane upon, during or after controlled assembly.

[0012] A second aspect of the present invention provides a nucleic acid nanostructure comprising:

[0013] a plurality of building blocks, each building block comprising a deoxyribonucleic acid (DNA) sequence, wherein the DNA sequence comprises at least a portion of a double helix defining a secondary structure and at least a portion of a single-stranded sequence defining an assembly interface, and at least one hydrophobic anchor; and

[0014] a plurality of single-stranded nucleic acid lock sequences capable of hybridizing to the single-stranded sequences at the assembly interface;

[0015] wherein the component modules are capable of associating and interacting with a surface of a semifluid membrane via the anchors, and

[0016] The modules are capable of controlled assembly in response to an externally applied stimulus, which initiates the dissociation of the plurality of lock sequences to reveal an assembly interface, thereby associating the plurality of component modules to form a nanostructure.

[0017] In a specific embodiment, the component modules are configured to associate and interact with the surface of the semifluid membrane via anchors.

[0018] A third aspect of the present invention provides a semifluid membrane having bonded thereto a nucleic acid nanostructure as described in any of the embodiments described herein.

[0019] A fourth aspect of the present invention provides a semifluid membrane having a plurality of nanostructured component modules bonded thereto, each component module comprising a deoxyribonucleic acid (DNA) sequence, wherein the DNA sequence comprises at least a portion of a double helix defining a secondary structure and at least a portion of a single-stranded sequence defining an assembly interface, and at least one hydrophobic anchor; and a plurality of single-stranded nucleic acid lock sequences capable of hybridizing with the single-stranded sequence of the assembly interface.

[0020] In a particular embodiment, the semifluid membrane comprises a synthetic amphiphilic membrane or a lipid bilayer membrane.

[0021] A fifth aspect of the present invention provides a sensor device comprising the nucleic acid nanostructure as described in any of the embodiments described herein.

[0022] A sixth aspect of the present invention provides a sensor device, wherein the sensor device comprises a semifluid membrane as defined in any of the embodiments described herein.

[0023] A seventh aspect of the present invention provides a semifluid membrane vesicle, wherein the vesicle comprises a nucleic acid nanostructure as defined herein.

[0024] An eighth aspect provides a pharmaceutical composition comprising a semifluid membrane vesicle as defined herein and a pharmaceutically acceptable carrier.

[0025] A ninth aspect of the present invention provides a method for assembling a nucleic acid nanostructure, the method comprising:

[0026] Provides multiple component modules, each of which includes

[0027] - a nucleic acid sequence, wherein the nucleic acid sequence comprises at least a portion of a double helix defining a secondary structure and at least a portion of a single-stranded sequence defining an assembly interface, wherein the assembly interface of each of the plurality of building blocks comprises a sequence that is complementary to and capable of hybridizing with the assembly interface of another building block; and

[0028] - at least one hydrophobic anchor;

[0029] and, combining multiple component modules under conditions that allow the nanostructures to spontaneously assemble.

[0030] A tenth aspect of the present invention provides a method for triggering the assembly of a nucleic acid nanostructure, the method comprising:

[0031] providing a plurality of component modules, each component module comprising a nucleic acid sequence, wherein the nucleic acid sequence comprises at least a portion of a double helix defining a secondary structure and at least a portion of a single-stranded sequence defining an assembly interface, wherein the assembly interface of each component module in the plurality of component modules comprises a sequence complementary to and capable of hybridizing with the assembly interface of another component module, and at least one hydrophobic anchor,

[0032] Providing a plurality of single-stranded nucleic acid lock sequences that hybridize with the single-stranded sequences of the assembly interface, thereby forming a plurality of locked component modules that are inhibited from initiating spontaneous assembly into nucleic acid nanostructures;

[0033] exposing the plurality of locking component modules to an external stimulus, wherein the external stimulus triggers the plurality of single-stranded nucleic acid lock sequences to dissociate from the plurality of locking component modules, thereby exposing the plurality of assembly interfaces; and

[0034] Providing conditions that allow the spontaneous assembly of nanostructures.

[0035] Within the scope of the present application, it is expressly intended that the various aspects, embodiments, examples and alternatives listed in the preceding paragraphs, claims and / or the following description and drawings, in particular their individual features, may be adopted independently or in any combination. That is, all embodiments and / or features of any embodiment may be combined in any manner and / or combination, unless these features are incompatible. BRIEF DESCRIPTION OF THE DRAWINGS

[0036] One or more embodiments of the present invention will now be described by way of example with reference to the accompanying drawings, in which:

[0037] Figure 1 Assembly of a nanostructure in the form of a DNA nanopore A·B triggered by components A and B at the membrane interface. (a) Components carrying locks LA and LB. Inactive components ALA and BLB are anchored to the membrane via cholesterol anchors (grey crescents). Addition of key chains KA and KB undoes the locks, allowing the activated A and B to reorient relative to the bilayer, forming a transmembrane ion channel. (b) Top and side views of the A·B pore, where the four DNA helices are arranged in a square lattice. The membrane cholesterol anchors are embedded and bent into the lipid bilayer.

[0038] FIG2 is a diagram of an embodiment of the present invention, showing a 2D chain diagram of the direct and triggered assembly of A·B. (a) 2D chain diagram illustrating components A and B, and assembly into the 4HB nanopore A·B. (b) Locking component AL A and BL B Schematic diagram of a 2D chain diagram showing the addition of a key (K A and K B ) to remove the lock (L A and L B ) and then assembled onto A·B. The resulting lock-and-key duplex is shown on the right. The squares represent the 5' end of the DNA, the triangles represent the 3' end of the DNA, and the asterisks represent the cholesterol anchor modification attached to the 3' end of the designated strand via a tri(ethylene) glycol (TEG) linker. The lines that make up the pores are shown in alternating light and dark grey. The crossovers at the top and bottom of each strand consist of four thymine bases.

[0039] Figure 3 shows a schematic diagram of the trigger mechanism and components in solution and on the membrane. (a) Assembling the locking pre-pore assembly (AL A and BL B ) are respectively composed of hole components (A and B) plus chains (LA and L B ) that prevent holes from forming. Add keychains (K A and K B ), each chain is unlocked by a toehold mechanism, leaving unlocked components (A and B). (b) A and B can freely self-assemble in solution (top) or on the membrane surface (bottom) to form an active ion channel.

[0040] Figure 4 shows the results of nanostructure A·B formed by direct and triggered assembly. (a) In the key chain K A and K B In the presence or absence of A and BL B (b) PAGE-based analysis of the formation of A·B by direct (dark lines) and triggered (light lines) mechanisms. ΔC Kinetic analysis of the triggered assembly. The inset shows a representative PAGE gel of the triggered assembly.

[0041] Figure 5 shows the PAGE characterization results of A·B pore formation. Confirmation of nanopore (A·B) ΔC Assembly from components in PBS (left) and KCl (300 mM KCl, 15 mM Tris-HCl, pH 7.4; right). Assembly from unlocked components assessed by native PAGE (A·B) ΔC The locking mechanism controls and triggers the efficiency of pore assembly. A was added in a 1:1 stoichiometric ratio ΔC and B ΔC Will lead to (A·B) ΔC Assembly (lanes 1-3). A was added in a 1:1 stoichiometric ratio ΔC L A +B ΔC L B No interaction occurs but it remains in its component parts (sequencing lanes 4-6). However, the key K is added in a 1:1 stoichiometric ratio A and K B will eliminate the lock and lead to the formation of (A·B) ΔC (Lane 7). Addition of the key in a 1:1 stoichiometric ratio was also sufficient to completely remove the corresponding lock (Sequencing Lane 7). Samples were incubated at 30°C for 30 min and then loaded onto the gel. 10% (left) and 12% (right) native PAGE were run at 4°C in 1× TBE buffer at 90 V for 115 min.

[0042] Figure 6 shows the results of gel electrophoretic mobility shift assays for the key-specificity and concentration-dependency of triggering pore assembly. (a) Key K A Assemble with locking assembly A ΔC L A A ΔC L:K A The gel showed that A ΔC L A Quantitative conversion to A in a 1:1 stoichiometric ratio ΔC The extra keys will not be associated with A ΔC (b) The key K B A Δ C L:K B Add to the assembled locking component A at a molar ratio of 1:0.5 to 1:10 ΔC L A The gel showed that A ΔC L A It is not unlocked, but forms a new structure A with a 1:1 stoichiometric ratio ΔC L A K B Once a stoichiometric structure is formed, the excess key will not interact further. (c) Key, K A and K B Assemble with locking assembly A ΔC L A and B ΔC L B The gel showed that when the stoichiometric ratio of all four elements was 1:1:1:1, the pores (A·B) ΔC and two double-chain K A L A and K B L B As the number of keys increased, other products emerged, such as the incomplete A ΔC B ΔC D, where D = L A , L B , K A or K B At a stoichiometric ratio of 1:1:10:10, the key inhibits the formation of the hole, resulting in a large amount of assembled locked A ΔC L A and B ΔC L B , free keys and a small amount of product A ΔC B ΔC D and assembled (A·B) ΔC .

[0043] FIG7 shows UV melting curve plots. (a) UV melting curves of nanostructure A·B when pre-annealed (light blue) or directly assembled from A and B (light red). The melting curves were evaluated in solution or when inserted into small unilamellar vesicles (SUVs) composed of DPhPC lipids with a diameter of 170 nm. 0.2 μM A·B was evaluated with 200 μM DPhPC lipids in 150 μL PBS buffer. The samples were melted using a temperature gradient of 1°C / min from 20°C to 80°C. Each trace represents the average of three independent replicates. (b) The melting temperature (T) used to determine each structure. m )'s first derivative of the melting curve.

[0044] Figure 8 shows the results of gel electrophoresis mobility shift analysis of A·B nanostructure formation. (a) Representative 3% agarose gel showing the binding of AΔC and BΔC in solution. As the concentration of AΔC increases, (A·B)ΔC nanopore complexes are formed, which are larger than the individual components and migrate more slowly in the gel. (b) Fitting of the binding curve of the (A·B)ΔC complex to the Langmuir curve shows K d is 1.54±0.23×10-7M. Data represent the mean and standard deviation of three independent experiments. (c) Representative 2% agarose gel showing the binding of A-SUV and BΔC on the membrane surface. As the concentration of A increases, a (A·B)1C membrane-bound complex is formed, which cannot migrate into the gel matrix but remains in the pores. (d) Binding curve of the (A·B)1C complex. The fitted Langmuir curve shows K d The molecular weight of the gel was 1.64 ± 0.14 × 10-7 M. The data represent the mean and standard deviation of three independent experiments. All gels were run in 1 × TAE buffer at 60 V for 60 min. The main marker of the 100 bp DNA molecular weight marker is shown on the left side of each gel. The bands are identified at the top and right.

[0045] Fig. 9 The nanostructure assembly Figure 1 shows the results of a FRET analysis. Components A and B carry the FRET pairs Cy3 and Cy5, respectively. Cy5 B (FRET acceptor) added to Cy3 A (FRET donor), resulting in the assembly of Cy3 A. Cy5 B, thus forming close proximity and FRET, which is manifested as a decrease in the emission intensity of Cy3 and an increase in the emission intensity of Cy5. (a) A in solution ΔC With B ΔC The ratio of A ΔC:B ΔC (b) (A·B) based on (a) shown as a percentage of binding ΔC The binding curve of the pore assembly. (c) The membrane system A-SUV and B Δc The ratio of A:B ΔC (d) (A·B) based on (c) 1C Binding curves for pore assembly, shown as percent binding. (e) Titration of pore assembly of A versus B on membrane surface. Ratio A:B (f) Binding curves for A·B pore assembly based on (e), shown as percent binding. Data shown in (b, d, and f) represent the mean and standard deviation of three independent experiments.

[0046] Figure 10 shows the two-color fluorescence cross-correlation spectroscopy analysis of the pore assembly. Assembly A ΔC and B ΔC Carrying Alexa488 and Alexa647 fluorophores respectively. (a)(A·B) ΔC Example of binding curve for pore formation, as Alexa488 A ΔC Function of concentration (grey dots). Alexa647 B ΔC The concentration of was kept constant at 16 nM. Each point represents a single sample measurement. The solid line represents the Langmuir isotherm fitted to the data, yielding the K d =6.39±1.07x10 -8 M. K generated by the average of two replicates d 7.17±0.77×10 -8 M, maximum observed binding A = 101%. The grey dots on the left represent the same experiment, but Alexa647 B ΔC Keep constant at 1nM. (b) (A·B) ΔC Example of binding curve for pore formation, as Alexa647 B ΔC Function of concentration (grey dots). Alexa488 A ΔC The concentration of was kept constant at 15 nM. The solid line represents the Langmuir isotherm fitted to the data, yielding K d =7.99±1.68x10 -8 M. The average of three replicates resulted in K d 6.22±1.25×10 -8 M, maximum observed binding B = 84%.

[0047] Figure 11 shows PAGE kinetic electrophoretic mobility shift analysis of direct and triggered pore assembly. (a) PAGE titration was used to assess A ΔC +B ΔCA·B formed by direct assembly. (b) Quantitative analysis of the kinetic data in (a) was performed by plotting the normalized amount of assembled A·B versus time. The amount of assembled A·B extracted from the brightness of the gel band was 1-(I A -I 背景 ), where I = intensity. (c) PAGE titration from AL A +BL B +K A +K B (d) Quantitative analysis of the kinetic data in (a) by plotting the normalized amount of assembled A·B versus time. The amount of assembled A·B extracted from the brightness of the gel band is 1-(I AL -I 背景 ), where I = intensity. 12% native PAGE was run in 1×TBE buffer at 4°C, 115V for 90 minutes. Time points were 1, 5, 10, 15, 20, 25 and 30 minutes. Each track represents data from three independent replicates. The main marker of the 100 bp DNA molecular weight marker is shown on the left.

[0048] Figure 12 shows the nanostructure assembly dynamics monitored by FRET. (a) When an assembly is attached to the membrane (A-SUV+B ΔC Figure 2. Fluorescence analysis of the kinetics of pore assembly in solution over time when immobilized on the membrane surface (A-SUB+B, blue) and on the membrane surface (A-SUB+B, purple). ΔC and buffer added control trace (grey). (b) Same as (a), but the initial rapid drop in A-SUV+B (purple) has been removed. Each trace is the average of three independent replicates.

[0049] Fig.13 Confocal microscopy images demonstrating cholesterol-mediated membrane binding of (A·B)1C to GUVs and assembly of nanostructures on the membrane surface. Component A was modified with Cy3 fluorophore and B with Cy5 fluorophore. Images were acquired using a 96× oil objective, scale bar 5 μm. (Top left) Bright field image of two GUVs composed of POPC lipids. (Top right) The green channel shows a ring around the GUVs, indicating that Cy3A is attached to the membrane. (Bottom left) The red channel shows that Cy5BΔC, which cannot interact with the membrane by itself, has bound to the membrane-tethered component A. (Bottom right) Overlay of the red and green channels demonstrates the formation and membrane attachment of (A·B)1C.

[0050] Figure 14 Nanostructures assembled in the form of DNA nanopores, A and B consisting of fluorescently labeled components. Using fluorescence microscopy and FRET analysis of membrane interfaces Cy3 A and Cy5B. (a)(A·B) 1C Confocal microscopy images of pores assembled on SUVs composed of DPhPC lipids. Bright field images (top) and overlay (bottom) of Cy5 (red) and Cy3 (green) channels. (b) Emission spectra of pores assembled on SUVs composed of DPhPC lipids using 545 nm excitation. When added at a 1:1 ratio (grey) Cy5 B, 563nm Cy3 The A emission peak (pink) drops to the same level as the pre-annealed A·B pore (dark grey). (c) The concentration-dependent change in Cy3 emission indicates pore assembly in solution (A Δc With B ΔC ) and the pore assembly at the membrane interface (A-SUV and B ΔC and A-SUV and B). (n=3).

[0051] Figure 15 shows a graph showing the results of a FRET control experiment between assembled locking components on the membrane surface. (a) Assembled locking component AL A and BL B Fluorescence spectra of AL were used to confirm pore assembly. This control was also used to assess the effect of Cy3-Cy5 proximity on the membrane surface. (b) Normalized histogram comparing the assembled locked components AL from a 1:1 stoichiometric ratio on the membrane surface. A and BL B The observed FRET signal is comparable to that observed from direct assembly of A + B with a 1:1 stoichiometry. Each trace is the average of three independent replicates, with standard deviations shown as error bars in the histograms.

[0052] Fig.16 Shown are smFRET and single particle tracking confirmation of A+B assembly on the membrane surface. Single molecule FRET (smFRET) and single particle tracking images confirm nanostructured assembly of monomeric assemblies A and B (top) compared to premixed control A·B (bottom). Bright green spots indicate the formation of A·B in the smFRET images (e.g., gray boxes). A carries a Cy3 fluorophore at the 5' end of chain A1 and B carries a Cy5 fluorophore at the 5' end of chain B1. Pore assembly of A and B was monitored on a supported lipid bilayer membrane at 62.5 pM. Single particle tracking experiments of A+B and premixed A·B were monitored for 500 and 1000 seconds, respectively.

[0053] Fig.17 Schematic diagram showing the model system used to investigate the effects of steric hindrance on DNA hybridization. Schematic diagram showing the model system used to investigate the effects of steric hindrance on DNA hybridization on membrane surfaces. The simplest of the four conditions, hybridization of a receptor (R) and a ligand (S), is shown on the left. The most sterically hindered of the four conditions, vesicle-anchored receptors (RSUV ) and the ligand attached to the nanopore (S NP ) as shown on the right. The cholesterol lipid anchor is shown in orange and inserted into the lipid bilayer in grey. The SUV used was composed of DOPC and DOPE in a 7:3 molar ratio and had an extrusion diameter of approximately 100 nm. For clarity, only two of the four conditions are shown.

[0054] Figure 18S NP The structure and assembly of S NP 2D chain diagram of DNA. All chains shown are numbered. The square represents the 5' end of the DNA and the circle represents the 3' end of the DNA. (b) 2% agarose gel confirms the presence of R and S NP Assembly.

[0055] FIG19 shows the results of a gel electrophoretic mobility shift assay to assess the binding of a receptor (R) to a ssDNA ligand (S) or a nanopore-bound ligand (S NP ). (a) 3% agarose gel shift assay. Addition of increasing concentrations of R to S induces the formation of an R·S dsDNA complex, which appears as a slower migrating band. S carries a Cy5 fluorophore and can be imaged at low concentrations. (b) Binding curves of the R·S complex are shown as percent binding. Data were fitted with a Langmuir binding curve and represent the mean and standard deviation of three independent experiments. (c) 2% agarose gel shift assay measuring R·S NP Complex formation. To improve separation, cholesterol-lipid anchor-modified R was used. Although S NP Much larger than R, but the cholesterol modification interacts with the gel matrix, resulting in R·S NP The mobility is significantly reduced. (d) R·S NP The binding curves of the composites are shown as percent binding. Data were fitted with a Langmuir binding curve and represent the mean and standard deviation of three independent experiments. All gels were run at 60 V in 1× TAE buffer for 60 min. The primary marker of a 100 bp DNA molecular weight marker is shown on the left.

[0056] Figure 20 shows the R and S labeled with Cy3 and Cy5, respectively, using FRET NP Fluorescence analysis results of the interaction. (a) R (black) and S NP (Blue) hybridization produces composite R·S NP (red) Integrated fluorescence measurement. (b) Hybridization overtime leads to Cy3 R and Cy5 S NP Kinetic fluorescence measurement of intensity changes. Cy3 R: Cy5 S NP The stoichiometric ratio is measured.

[0057] FIG21 shows the results of a gel electrophoretic mobility shift assay to evaluate the activity of ssDNA ligand (S) or nanopore-bound ligand (S NP ) binds to a receptor (R) attached to the surface of the SUV. (a) 3% agarose gel shift assay. Increasing the concentration of R results in R SUV · Formation of S membrane-bound dsDNA complexes that are unable to migrate into the gel matrix and remain in the wells. S carries a Cy5 fluorophore to improve resolution at low concentrations. The gel was imaged using a fluorescent gel scanner. (b) R SUV The binding curves of the S complex are shown as percent binding. The data were fitted with a Langmuir binding curve and represent the mean and standard deviation of three independent experiments. (c) 2% agarose gel shift assay: increasing R concentration results in R SUV ·S NP Formation of membrane-bound complexes that are unable to migrate into the gel matrix and remain in the pores. (d) R SUV ·S NP The binding curves of the composites are shown as percent binding. Data were fitted with a Langmuir binding curve and represent the mean and standard deviation of three independent experiments. All gels were run at 60 V in 1× TAE buffer for 60 min. The primary marker of a 100 bp DNA molecular weight marker is shown on the left.

[0058] Fig. 22 It shows the R and S (or S) of the solution and membrane surface NP ) is a graph showing the FRET kinetic analysis of hybridization. NP Add points to normalize the fluorescence signal to F / F0. For R+S (bottom line) and R+S NP (second from the bottom line), R was added at 1.25 nM, S and S were added at 125 nM in 100-fold excess NP . R SUV with S (top line at 150 seconds) and R SUV With S NP (second from the top line at 150 s), R was added at a concentration of 5 nM, S and S NP A 20-fold excess was added at a concentration of 100 nM. SUV (100 nm) was added at a concentration of 22.2 pM, resulting in a 225:1 R:SUV loading or a 1167:1 lipid:R ratio.

[0059] Fig.23 The GUV surface S is shown NP Confocal microscopy images of hybridization with R. on the surface of giant unilamellar vesicles (GUVs) composed of POPC lipids Cy3 R GUV and Cy5 SNP Combined confocal laser scanning microscopy images. Three channels are shown: the bright field in the first row confirms the presence of GUVs, the second row shows the green channel that excites the Cy3 fluorophore on R, and the third row shows the green channel that excites the S NP (a) The fluorescent green halo around the GUV confirms that Cy3 R is inserted and bound into the bilayer. (b) The green and red fluorescence halos around the GUV confirm both (A) and Cy5 S NP and Cy3 R GUV (c) The absence of any fluorescent halo confirms that there is no Cy3 In the case of R, Cy5 S NP Will not interact with GUVs.

[0060] Fig.24 is a graph showing the results of digestion analysis confirming nanopore insertion. In solution, (A·B) 1C Both A·B were completely and rapidly digested by BAL-31 DNA digestion enzyme. 1C When A·B and A·B were first incubated with LUVs composed of DPhPC, digestion was slowed in both cases, probably due to steric hindrance of the vesicles. A·B inserts into the lipid bilayer, making the strand gaps of DNA inaccessible, thus significantly inhibiting digestion. 1C In this case, the presence of a membrane reduces access to the notch, but only on one side of the cholesterol lipid anchor.

[0061] Figure 25 shows the stability and dynamics of membrane-bound DNA and transmembrane DNA nanostructures. (a) A·B (dark grey) and control (A·B) 4C (light gray), (A·B) 1C (gray) and (A·B) ΔC (black). (b) In the presence of SUVs composed of POPC, A·B (dark gray), (A·B) 4C (medium gray), (A·B) 1C (light grey) and (A·B) ΔC (c) Representative snapshots of the DNA nanostructures from four simulation trajectories. Component A in solution (upper left) is attached to the membrane (upper right), and A·B in solution (lower left) spans the membrane A·B (lower right). The simulated POPC bilayer is shown in grey, with solvent atoms omitted for clarity. The DNA nanostructures are plotted by per-residue RMSF 10 Shading to indicate regions of structural flexibility (left to right in scale). The location of the hydrophobic membrane anchor is indicated by a black arrow.

[0062] Fig.26 The eigenvector indices showing the slowest modes of the simulation trajectories are shown. The eigenvector indices were generated by principal component analysis using gmx_cocovariance and gmx_aneaig, demonstrating that the top ten slowest modes describe more than 90% of the structural dynamics of the DNA heavy atoms.

[0063] Figure 27 shows the time-dependent root mean square deviation (RMSD) of DNA heavy atoms for four simulation trajectories. (a) RMSD of nanostructure A·B in 1 M KCl solution (top) or embedded in a POPC bilayer (bottom). (b) RMSD of component A in 1 M KCl solution (top) or attached to a POPC bilayer (bottom). The RMSD was calculated using an initial idealized structure consisting of a rigid B-form DNA helix. Data are plotted as a running average of 10 ns. Error bars represent standard deviations.

[0064] Fig.28 The root mean square fluctuations per residue of the DNA nanostructures from four simulation trajectories are shown. 10 ). The residue index on the Y axis has been sorted to match that of the associated strand plot (top: A; bottom: A·B). The squares represent the 5' end of each DNA strand and the triangles represent the 3' end of each DNA strand. The grey circles indicate cholesterol anchor modifications attached to the 3' end of the DNA strand via a tri(ethylene) glycol (TEG) linker.

[0065] Fig.29 It is RMSF 10 Box plots showing the values ​​of component A and nanostructure A·B in solution and in interaction with the membrane. Box plots showing the arithmetic mean RMSF of DNA heavy atoms for each of the four simulation trajectories 10 value

[0066] Figure 30 shows the effect of membrane anchoring of component A and nanopore A·B on the lipid bilayer analyzed by molecular dynamics simulation. (a, b) Representative snapshots of the equilibrium region of the trajectories of (a) membrane-attached component A and (b) transmembrane A·B. The inset shows the initial configuration. DNA is shown in purple and the TEG cholesterol linker is shown in gold. The POPC bilayer is shown in white (head) and gray (lipid tail). For clarity, the solvent is omitted. The average bilayer density of the A and A·B trajectories is plotted in the upper right corner of each snapshot, and the dashed line represents the core density of the membrane lacking the transmembrane nanopore (0.425 g / mol / A 3). (c, d) Top views corresponding to panels (a) and (b), respectively, with DNA omitted for clarity. Each hexagon is colored according to the RMSF per lipid molecule (black = low, white = high). The position of the cholesterol-lipid anchor is indicated at the beginning (orange) and end (yellow) of the trajectory. (e) Simulated channel lumen mapping using HOLE analysis for a series of clustered snapshots of the A·B channel in a POPC bilayer in 1 M KCl. The proportion of the trajectory represented by each cluster is indicated by the transparency of the points, and a trend line is drawn to estimate the average channel shape. The two dashed black lines indicate the approximate position of the bilayer head groups compared to the nanopore channel coordinates in the densest cluster.

[0067] Fig.31 is a schematic diagram of the top-down images showing representative structures from four simulation trajectories, according to the per-residue RMSF 10 Coloring. From left to right; A subunit in solution, A subunit attached to a POPC bilayer, A·B nanopore in solution, and A·B nanopore embedded in a POPC bilayer. For clarity, the solvent was omitted, making the bilayer transparent. The position of the hydrophobic cholesterol anchor is indicated by a black arrow.

[0068] Fig.32 A plot of the average angle of the lipid head groups relative to the bilayer normal is shown. Insertion of the A·B nanostructure results in the rapid formation of toroidal lipids. The head groups of the trajectory with the transmembrane A·B DNA nanopore are shown in red, and the head groups of the trajectory with the membrane-tethered component A are shown in blue. The lipids are split by the leaflet, lipids more than 4 nm from the DNA structure are discarded, and the angle is calculated with gmx_gangle and time averaged to indicate the direction of motion. The initial region of the NvT equilibrium trajectory must be used to capture this motion, as it occurs primarily within the first 20 ns of the simulation.

[0069] Fig.33 Shown is the output of the evolution of the simulated per-residue distance between the lipid bilayer and the DNA nanostructure A·B and assembly A. Calculated using gmx_minist on the initial NvT equilibrium trajectory to demonstrate the DNA nanopore-induced lipid remodeling.

[0070] Fig.34 Analysis of different clustering cutoffs for clustering cavity analysis of membrane-embedded DNA nanostructure pores A·B is shown. After discarding the initial 10% of the trajectories, the NpT regions of the trajectories were clustered using the gromos method (see Examples). The final clustering cutoff used was 0.25 nm, which was chosen to ensure that there were no more than 50 clusters, that clusters greater than 10 contained 90% of the trajectories, and that fewer than 3 clusters contained a single species.

[0071] Figure 35 shows the conductance characteristics of the nanostructured pore A·B. (a) Schematic diagram of the preassembled A·B inserted into a planar lipid bilayer. (b) Schematic diagram of the triggering assembly at the membrane interface induced by the addition of a keychain. (c) Representative ion current trace of a single A·B nanopore inserted into a planar lipid bilayer composed of DPhPC lipids in 1M KCl, 10mM HEPES, pH 7.4 at a membrane potential of -50mV. (d) Current-voltage relationships in the voltage range of -100mV to +100mV with a step size of 20mV show the ohmic characteristics of the A·B channel and show the mean ± SEM of 16 independent pore insertions. (e) Channel conductance histograms obtained from 16 independent single channel recordings at membrane potentials of +20mV to +50mV. (f) Representative ion current traces showing the ion currents in a planar lipid bilayer containing AL·B. A and BL B How the keychain is added to the membrane to form nanopores A and B at +90 mV. (g) Channel conductance histogram obtained from 8 independent single-channel recordings of triggered assembled nanopores at a membrane potential of 20 mV.

[0072] Figure 36 shows graphs demonstrating that the transport-active A·B channel is assembled from inactive components. (a) Fluorescence traces of sulforhodamine B (SRB) efflux kinetics resulting from direct assembly of A and B and from the inactive subunit AL. A and BL B A·B triggered assembly was performed. The fluorescence signal originates from SRB, which is contact quenched at high concentrations within DPhPC SUVs but regains emission when released into the environment at low concentrations. 100% of the signal was obtained by disrupting the SUVs by adding detergent. Each trace is the average of three independent replicates obtained at 400 nM. (b) (A·B) inserted into the membrane 4C Ca 2+ Kinetic fluorescence trace of influx of Ca-containing 2+ Sensitive dye Fura-2 in POPC SUVs. The traces were repeated 3, 2, 1, and 3 times for 75 nM, 38 nM, 25 nM, and 0 nM, respectively. The maximum influx was recorded after rupturing the vesicles by adding detergent.

[0073] Figure 37 shows the graph of A·B concentration-dependent SRB release in SUV. (a) When A·B is composed of the unlocked component A+B (b) and the locked component AL after adding the key A +BL B (c) Concentration dependence of the formation of prefolded A·B. (d) Bar graph summarizing the net fluorescence increase results of (a)-(c). 0nM A·B represents the control group: 400nM A, 400nM (AL A +BLB ) and buffer. Data represent the mean and standard deviation of three independent replicates.

[0074] Figure 38 shows the 2+ Characteristic diagram of Fura-2 encapsulated vesicles in the influx assay. 2+ concentration, Fura-2 is excited at 380 nm. 2+ As the concentration increases, the excitation maximum shifts to 340 nm. The emission wavelength remains constant at 510 nm. Therefore, the Ca concentration can be assessed by monitoring the excitation intensity ratio at 340 / 380 nm. 2+ (a) Fluorescence scanning of purified SUVs with encapsulated Fura-2 (100 μM) demonstrated the absence of Ca in the vesicles. 2+ , the 340 / 380 ratio was low (0.95). Addition of 250 μM CaCl2 resulted in a slight increase in the 340 / 380 ratio (1.13), consistent with some unpurified dyes and confirming that the vesicles are responsive to Fura-2 and Ca 2+ After adding detergent (Triton X-100), the vesicles are lysed to remove Fura-2 and Ca 2+ The barrier between them leads to maximum binding, and the 340 / 380 ratio increases significantly (5.62). (B) In the presence of Ca 2+ Dynamic light scattering analysis of vesicles formed by POPC with encapsulated Fura-2 (100 μM) in the presence and absence of Ca 2+ In the presence of Ca 2+ Under the conditions of Ca 2+ The induced membrane compression is consistent. DETAILED DESCRIPTION

[0075] Before describing the present invention, a number of definitions are provided to aid in understanding the present invention.

[0076] The practice of the present invention employs, unless otherwise indicated, conventional techniques of chemistry, molecular biology, microbiology, recombinant DNA techniques and chemical methods, which are within the capabilities of ordinary skill in the art. These techniques are also explained in the literature, for example, MR Green, J. Sambrook, 2012, Molecular Cloning: A Laboratory Manual, Fourth Edition, Books 1-3, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY; Ausubel, FM et al., (Current Protocols in Molecular Biology, John Wiley & Sons, Online ISSN: 1934-3647); B. Roe, J. Crabtree, and A. Kahn, 1996, DNA Isolation and Sequencing: Essential Techniques, John Wiley & Sons; JM Polak and James O'D. McGee, 1990, In Situ Hybridisation: Principles and Practice, Oxford University Press; MJ Gait (Editor), 1984, Oligonucleotide Synthesis: A Practical Approach, IRL Press; and D. MJ Lilley and JEDahlberg, 1992, Methods of Enzymology: DNA Structure Part A: Synthesis and Physical Analysis of DNA Methods in Enzymology, Academic Press; Synthetic Biology, Part A, Methods in Enzymology, Edited by Chris Voigt, Volume 497, pages 2-662 (2011); Synthetic Biology, Part B, ComputerAided Design and DNAAssembly, Methods in Enzymology, Edited by Christopher Voigt, Volume 498, Pages 2-500 (2011.Each of these general texts is incorporated herein by reference.

[0077] As used herein, the term "comprising" means that any one of the elements described must be included, and other elements may be optionally included. "Substantially consisting of..." means that any of the cited elements must be included, and elements that significantly affect the basic and novel characteristics of the listed elements are excluded, and other elements may be optionally included. "Consisting of..." means that all elements except the listed elements are excluded. Embodiments defined by each of these terms are within the scope of the present invention.

[0078] The term "modular" as used herein refers to the use of one or more units or components to design or construct all or part of a larger complex nanostructure. In the context of the present invention, it refers to the use of a single component module, subunit or building block to build a nanostructure, appropriately a nanostructure configured to penetrate or pass through a semifluid or lipid bilayer membrane. These modules can be the same or different. In order to form a nanostructure, each module is constructed to include an assembly interface region that helps to assemble into a larger nanostructure complex by complementary base pairing at the assembly interface.

[0079] The modular design of a nanostructure may include a framework or frame of modules, and additional, typically smaller sub-modules connecting or supporting the framework, which act as struts or support members.

[0080] Modular components or submodules are composed of nucleic acids, typically DNA, RNA and synthetic nucleic acids or their analogs (e.g., LNA or PNA). Each individual unit can be assembled using appropriately selected scaffold strands and staple strands by DNA / RNA origami techniques described elsewhere herein to create higher-order structures, such as secondary structures with defined geometric parameters. Such secondary structures may include the formation of A-, B- or Z-shaped double helices (double helices), triple helices, quadruple helices, hairpin loops and trilobal structures, as well as combinations of these structures.

[0081] The term "nucleic acid" as used herein is a single-stranded or double-stranded covalently linked nucleotide sequence, wherein the 3' and 5' ends on each nucleotide are connected by a phosphodiester bond. Polynucleotides can be composed of deoxyribonucleotide bases or ribonucleotide bases. Nucleic acids can include DNA and RNA, and are usually synthetically manufactured, but can also be separated from natural sources. Nucleic acids can also include modified DNA or RNA, such as methylated or chemically modified DNA and RNA, such as 5', 3'-processed with 7-methylguanosine or its analogs, such as cutting and polyadenylation, splicing or labeling with fluorophores or other compounds. Nucleic acids can also include synthetic nucleic acids (XNA) or nucleic acid analogs, such as hexitol nucleic acids (HNA), cyclohexene nucleic acids (CeNA), threose nucleic acids (TNA), glycerol nucleic acids (GNA), locked nucleic acids (LNA) and peptide nucleic acids (PNA). Therefore, when the terms "DNA" and "RNA" are used in this article, it should be understood that these terms are not limited to only naturally occurring nucleotides. The size of a nucleic acid, also referred to herein as a "polynucleotide," is generally expressed as the number of base pairs (bp) for double-stranded polynucleotides, or as the number of nucleotides (nt) in the case of single-stranded polynucleotides. One thousand bp or nt equals a kilobase (kb). Polynucleotides less than about 100 nucleotides in length are generally referred to as "oligonucleotides."

[0082] As used herein, the terms "3'" ("3prime") and "5'" ("5prime") have their usual meanings in the art, i.e., distinguishing the ends of a polynucleotide. A polynucleotide has a 5' and a 3' end, and polynucleotide sequences are usually written in a 5' to 3' direction. The term "complement of a polynucleotide molecule" refers to a polynucleotide molecule having a complementary base sequence and an orientation opposite to that of a reference sequence.

[0083] The term "double-stranded" as used herein refers to a double-stranded nucleic acid hybrid molecule, such as DNA (dsDNA), which means that the nucleotides of two complementary DNA sequences are bound together and then curled to form a double helix (assuming an A-, B-, or Z-shape), or also refers to a single-stranded RNA (ssRNA), which anneals to a complementary DNA sequence to produce an RNA-DNA hybrid (RDH) double strand. The RDH nanostructure can include a single RNA scaffold sequence with multiple shorter hybrid DNA sequences (e.g., DNA oligonucleotides) as staple strands, forming a series of RDH double strands along the length of the RNA scaffold, thereby defining a higher-order structure.

[0084] According to the present invention, the homology with nucleotide sequence described herein is not limited to 100%, 99%, 98%, 97%, 95% or even 90% sequence identity. Although sequence identity is obviously lower, many nucleotide sequences can prove each other's biochemical equivalence. In the present invention, homologous nucleic acid sequences are considered to sequences (Sambrook J. et al., Molecular Cloning:a Laboratory Manual, Cold Spring Harbor Press, Cold Spring Harbor, NY) that hybridize with each other under low stringency conditions. However, in some cases, it may be necessary to distinguish between two sequences that can hybridize with each other but contain some mispairings-"imprecise matching", "incomplete matching" or "imprecise complementation"-and two sequences that can hybridize with each other without mispairing-one "precise matching", "complete matching", or "precise complementation". In addition, possible mismatching degrees are also considered.

[0085] As used herein, the term "nanostructure" refers to a geometrically predefined or "predesigned" two- or three-dimensional molecular structure, typically composed of a biopolymer, suitably a naturally or non-naturally occurring nucleic acid, which structure has at least one dimension or an aspect of its geometric configuration that is within the nanometer scale (i.e., 10 -9 m). The nanoscale structure suitably has a size or geometry of less than about 100 nm, typically less than about 50 nm, and most suitably about 20 nm. The nanoscale structure suitably has a size or geometry greater than about 0.1 nm, typically greater than about 1 nm, and optionally greater than about 2 nm. The assembly of nucleic acid nanostructures can occur spontaneously in solution, for example by heating and cooling a mixture of DNA chains of a preselected sequence, or may require the presence of additional cofactors, including but not limited to nucleic acid scaffolds, nucleic acid aptamers, nucleic acid staple chains, coenzymes, and molecular chaperones. When the desired nanostructure is produced by one or more pre-designed spontaneous self-folding nucleic acid molecules (such as DNA or RNA), this is generally referred to as nucleic acid "origami". Rational design and folding of DNA to create two-dimensional or three-dimensional nanoscale structures and shapes is known in the art (e.g., Rothemund (2006) Nature 440, 297-302). The term "geometric predefined" is used to indicate that the geometry of the nanostructure is predefined so that when assembled, the nanostructure conforms to the shape and configuration desired by the designer. For example, the choice of scaffold and staple sequences enables iterative confirmation of the rational design of the nanostructures after hybridization is complete.

[0086] In the classical scaffold-and-staple method, one or more long biosource scaffold chain components are folded into a defined nucleic acid nanostructure, optionally with a staple component composed of many shorter synthetic staple chain oligonucleotides. The classical DNA nanostructure is formed by parallel arranged DNA double-stranded bundles, which are arranged in bundles or barrels, can close the central pore channel and pierce the semi-fluid membrane. Suitably, certain scaffold structures can be based on all or part of the M13 or phiX174 sequences, using which multiple smaller staple chains and linker sequences are configured to achieve the desired three-dimensional nanostructure geometry. In embodiments of the present invention, alternative scaffolds can be used and can include artificial or non-natural sequences designed specifically for nanostructure modular assembly tasks. Typically, such sequences will be non-repetitive and have optimized base selection to promote nucleic acid hybridization between component modules under conditions that are conducive to nanostructure assembly.

[0087] The nucleic acid sequences that form the nanostructures are usually synthetically manufactured, although they can also be obtained by conventional recombinant nucleic acid techniques. A DNA construct containing the desired sequence can be contained in a vector grown in a microbial host organism (such as E. coli). This will allow large quantities of DNA or RNA to be prepared in a bioreactor and then harvested using conventional techniques. The vector can be isolated, purified to remove foreign matter, and the desired DNA sequence can be excised and separated by restriction endonucleases, for example by using chromatography or electrophoresis.

[0088] As used herein, the term "hydrophobic" refers to molecules, including organic molecules and polymers, having non-polar characteristics. For example, saturated or unsaturated hydrocarbons. The molecule may have amphiphilic properties.

[0089] As used herein, the term "hydrophobic modification" refers to modifying (connecting, bonding or otherwise connecting) a polynucleotide chain with one or more hydrophobic parts. "Hydrophobic part" as defined herein is a hydrophobic organic molecule, which can be synonymous with the term "lipophilic", indicating that the molecule has affinity for lipids, particularly the lipid core of a membrane bilayer. The hydrophobic part can be any part comprising a non-polar or low-polarity aliphatic, aliphatic aromatic or aromatic chain. Suitably, the hydrophobic part used in the present invention includes molecules such as long-chain carbocyclic molecules, polymers, block copolymers and lipids. The term "lipid" as defined herein relates to fatty acids and derivatives thereof (including triglycerides, diglycerides, monoglycerides and phospholipids), as well as sterol-containing metabolites such as cholesterol. The hydrophobic part included in the embodiments of the present invention can form non-covalent attractive interactions with amphipathic semifluid membranes or phospholipid bilayers (such as the lipid basement membrane of cells), and act as a membrane anchor for nanostructures. According to certain embodiments of the present invention, suitable hydrophobic moieties, such as lipid molecules, having membrane anchoring properties may include sterols (including cholesterol, cholesterol derivatives, phytosterols, ergosterols and bile acids), alkylated phenols (including methylated phenols and tocopherols), flavonoids (including flavonoid-containing compounds such as 6-hydroxyflavone), saturated and unsaturated fatty acids (including derivatives such as lauric acid, oleic acid, linoleic acid and palmitic acid) and synthetic lipid molecules (including dodecyl β-D-glucoside). The anchor of the polymer membrane may be the same as or different from the lipid bilayer. The specific hydrophobic moiety anchor may be selected based on the binding properties of the selected membrane.

[0090] In embodiments of the present invention, the disclosed nanostructures may include one or more hydrophobic or lipophilic anchors for attaching or linking or anchoring hydrophilic nucleic acid nanostructures to a generally hydrophobic membrane, such as a semifluid or lipid bilayer of a vesicle. The lipid anchor is attached to the nanostructure or contained in a module that forms a portion of the entire nanostructure. A suitable connection is by an oligonucleotide carrying a lipid anchor (suitably cholesterol) at the 5' or 3' end. In the chain synthesis reaction, a modified phosphoramidite can be used to functionalize the polynucleotide or oligonucleotide, which is easily compatible with adding reactive groups (such as cholesterol and lipids) or linking groups (including thiols and biotin). Enzymatic modification using terminal transferase can also be used to bind oligonucleotides to the 3' of single-stranded nucleic acids (such as ssDNA), and the oligonucleotides are combined with modifications such as anchors. These lipid-modified anchor chains can be hybridized with the corresponding portion of the nucleic acid sequence that forms the nanostructure scaffold portion through an "adaptor" oligonucleotide. Alternatively, lipid anchors are assembled with nanostructures using lipid-modified oligonucleotides as scaffolds or staple chains. Combinations of anchoring methods using two or more membrane anchors may also be employed, wherein the anchors are incorporated into one or all of the scaffold strands, staple strands, and linker oligonucleotides. Cholesterol has been found to be a lipid that is particularly suitable for use as an anchor in the present invention (see Figure 1). Other lipids may be contemplated for use as anchors, although for a given membrane chemistry, particular preferences for particular lipids and a given number of membrane anchors may be expected.

[0091] In an alternative embodiment of the invention, the hydrophobic modification is contained within one or more synthetic nucleic acids (XNA) that are integrated into the nanostructure itself.

[0092] Nanostructures according to embodiments of the present invention are capable of binding to cells, membranes, or microsomes or exosomes in a subject. In one embodiment of the present invention, the nanostructures are bound to the membrane by inserting at least one associated hydrophobic anchoring moiety into the membrane bilayer. According to this embodiment of the present invention, most of the nanostructures are located on the outer surface of the membrane, but do not penetrate or pierce the membrane in the manner of transmembrane nanopores.

[0093] Suitably, the nanostructures of the invention comprise one or more polynucleotide chains providing a functional scaffold component, wherein the polynucleotide chains contained within the scaffold component include a polynucleotide backbone; and multiple polynucleotide chains providing multiple functional staple components. The scaffold chain interacts and hybridizes with itself or multiple staple nucleotide chains, for example, by appropriate Watson-Crick base pairing hybridization, to form a three-dimensional structure of the nanostructure, referred to as a secondary structure.

[0094] Nanostructures according to embodiments of the present invention may include nucleic acid nanostructures, such as nanotubes. Thus, nucleic acids may form a bundle, or a series of modules consisting of bundles, which work together to define the desired geometry of the nanostructure. The geometry may include a combination of secondary structural patterns and more open unstructured areas. The geometry may change from pre-assembly to post-pre-assembly.

[0095] All configurations of the nanostructures of the present invention can be configured by the "scaffold and staple chain" method. In this important route to nucleic acid nanostructures, DNA or RNA is used as a building material to create nanoscale three-dimensional shapes. The arrangement of complex nanostructures composed of multiple unhybridized linear molecules is often referred to as "nucleic acid origami". The nucleic acid origami process generally involves folding one or more elongated "scaffold" chains into a specific shape by self-hybridization and / or using multiple rationally designed "staple" oligonucleotide chains. The scaffold chain can have any sufficiently non-repetitive sequence. The sequence of the staple chain is designed to include a sequence that hybridizes to a specific defined portion or region of the scaffold chain, and in doing so, the two components cooperate to force the scaffold chain to assume a specific structural configuration. The staple chain is usually made of DNA, but may also contain RNA or other synthetic nucleic acids as described above. Methods that can be used to make DNA origami structures can be found in, for example, Rothemund, PW, Nature 440: 297-302 (2006); Douglas et al, Nature 459: 414-418 (2009); Dietz et al, Science 325: 725-730 (2009); and U.S. Patent Application Publication Nos. 2007 / 0117109, 2008 / 0287668, 2010 / 0069621 and 2010 / 0216978, the entire contents of which are incorporated herein by reference. For example, the design of staple strand sequences can be accomplished using CaDNAno software (available at http: / / www.cadnano.org) or DAEDALUS online platform (website address is http: / / daedalus-dna-origami.org).

[0096] In an embodiment of the present invention, the staple chain and / or the support assembly also include a plurality of hydrophobic / lipophilic membrane anchoring molecules attached thereto. The hydrophobic anchor (or part of a sequence) contributes to the combination of the nanostructure assembly before or after the semifluid membrane assembly. In or after the process of being assembled into a complete nanostructure, the presence of the membrane anchoring molecule promotes the penetration of the nanostructure into the membrane, and contributes to the nanostructure in the stable membrane. Typically, the membrane anchoring molecule can be connected to the nanostructure assembly at a substantially equatorial position (e.g., at a position along the midline of the nanostructure when embedded in the membrane) so that the anchoring molecule extends radially outward from the nanostructure into the surrounding semifluid membrane. According to the size and stability requirements of the module, and the final fully assembled transmembrane nanostructure, each individual component module can include at least one, two, three or more membrane anchors.

[0097] According to one embodiment of the present invention, a kind of nucleic acid nanostructure is provided, it comprises a plurality of component modules.Suitably, each module comprises nucleic acid sequence (such as DNA) and at least one attached membrane anchoring molecule.The nucleic acid sequence comprises at least a portion / region of double helix, and it is used to define the secondary structural element with defined length and structural rigidity level.The secondary structure can comprise one or more nucleic acid double-stranded bundles.In a specific embodiment of the present invention, the double-stranded bundle can be oriented 5' to 3' substantially perpendicular to the planar axis of semifluid film.In another embodiment of the present invention, the double-stranded bundle can be oriented 5' to 3' substantially coaxially with the planar axis of semifluid film.In some nanostructures, according to the requirements of the resulting nanostructure, there may be a module combination comprising two orientation combinations.

[0098] The nucleic acid sequence also includes at least a portion of a single-stranded sequence that defines an assembly interface. The assembly interface may be exposed, which facilitates the spontaneous assembly of the nanostructure in solution or when complementary component modules are co-located on a membrane surface. After assembly, the nanostructure changes conformation, enabling it to penetrate and cross a semifluid membrane.

[0099] In another embodiment, a plurality of single-stranded nucleic acid lock sequences capable of hybridizing with single-stranded sequences of the assembly interface are provided. The lock sequence is used to inhibit the spontaneous assembly of the nanostructure in solution or when the component module is applied to the membrane surface. Upon addition of an external stimulus (e.g., by a triggering mechanism), the lock sequence may be displaced or induced to separate from the assembly interface sequence, thereby allowing the spontaneous assembly of the nanostructure and subsequent membrane penetration. Therefore, the controlled initiation of a three-dimensional nanostructure from a plurality of dissociated component modules in response to a predefined stimulus is referred to as "triggered" assembly. The external stimulus may include a trigger molecule selected from one or more of the following: a nucleic acid sequence; an aptamer; a small molecule; an antibody or antibody fragment; an antibody mimetic; a peptide; a polypeptide; a polysaccharide; and an oligosaccharide. In a specific embodiment, the external stimulus includes a nucleic acid sequence that hybridizes with all or a portion of the lock sequence, or hybridizes with a portion of the lock sequence sufficient to induce separation from the assembly interface sequence of the component module.

[0100] In an embodiment of the present invention, the lock sequence may not completely hybridize with the assembly interface sequence, for example, there are one or more base pair mismatches. In an alternative or additional embodiment, the lock sequence may include one or more connected and sterically hindered parts. In these two embodiments, energetics may be conducive to the dissociation of the lock sequence in the presence of external stimuli (such as a fully matched trigger nucleic acid sequence). In another embodiment of the present invention, the lock sequence may include or be connected to an aptamer that binds to a trigger molecule as described above. After the trigger molecule binds, the aptamer may undergo a conformational change, resulting in the separation of the lock sequence from the assembly interface sequence.

[0101] Therefore, in a plurality of embodiments, nanostructure of the present invention is formed or built by a plurality of discrete component modules, and these component modules cooperate spontaneously or in response to external stimulation, and form complete transmembrane complex.In a plurality of embodiments, nanostructure can be formed by the module arrangement that forms basic framework or framework.In a plurality of embodiments, the module of framework is supported by the extra, usually smaller submodule that is connected and supports framework structure.In a plurality of embodiments of the present invention, module and submodule can comprise a plurality of substantially similar supports and short nucleic acid structures that assemble in the same manner, and they combine to form repeated structural motifs.

[0102] Although any arrangement of modules can be envisioned, it is appropriate that after assembly, the modules can be arranged to form a series of transmembrane nanostructures with polygonal cross-sections. The component modules are arranged in parallel, thereby defining the geometric configuration of the entire nanostructure. The module can have an adjustable side length (in this case, the side length is defined as the longest dimension of the module), and when a suitable final overall shape is selected, nanostructures of different sizes and / or shapes can be prepared. For example, the nanostructure defined by the assembly of the component modules can include a series of three-dimensional geometric shapes, appropriately selected from regular or irregular polyhedrons, whose cross-sections define annular or solid shapes, such as circles, ellipses, triangles, quadrilaterals (e.g., squares, rectangles or trapezoids), pentagons, hexagons, heptagons, octagons, etc. As disclosed herein, in certain embodiments, the nanostructure is similar to a nanorod or a helical bundle. However, it should be understood that the geometry of the nanostructure can be selected to accommodate a range of factors. These factors may depend on the inherent properties of the membrane thickness, such as the length required to span the membrane; or they may be defined by stability or functional factors, such as the desired diameter of the enclosed lumen in the case of transmembrane nanopores.

[0103] Generally, the length of side or the maximum dimension of the module included in the nanostructure or the nanostructure is between 5nm and 50nm. Suitably, the length of side of the module can be at least 2nm, 3nm, 4nm, 5nm, 6nm, 7nm, 8nm, 9nm or 10nm. The size of the component submodule can be determined by the number of modules required for complete assembly, or the relative spacing between the modules is determined by the shape of the nanostructure and the size and quantity of the modules adopted. Suitably, the length of side of the component module can be at least 0.5nm, 1nm, 1.5nm, 2nm, 2.5nm, 3nm, 3.5nm, 4nm or 5nm. Suitably, the length of side of the component module can be at most 20nm, 10nm, 9nm, 8nm, 7.5nm, 7nm, 6nm or 5nm.

[0104] According to one embodiment of the application, the nanostructure defines at least one inner cavity extending along the central axis of the nanostructure, thereby defining at least the first and second openings. Therefore, the assembled nanostructure can be a transmembrane nanopore. The first and second openings can be referred to as holes (such as the first and second holes), and allow fluid to be communicated through the inner cavity or central channel of the hole. Suitably, the first hole is located at the cis side of the nanopore, and the second hole is located at the trans side. When the nanopore is embedded in the membrane, the fluid communication through the hole allows measurable charged ions to pass through the hole from cis to trans or vice versa, i.e., measurable current can pass through the inner cavity of the nanopore through the membrane. It should be understood that before the nanostructure is assembled and penetrates the membrane, measurable current will not pass through the membrane. In this way, sensing or drug delivery functions are controlled by the ability of the nanostructure to be assembled from its component modules. In the embodiment of the present invention controlled by external stimulation assembly, sensing or drug delivery capabilities are therefore affected by the presence of stimulation.

[0105] The three-dimensional configuration of the assembled nanopore of the present invention defines at least one channel, suitably a single channel across the membrane, the channel having an inner cavity, the minimum internal width of the inner cavity being at least about 0.2nm, suitably 0.5nm, optionally 0.75nm. When observed perpendicular to the plane of the membrane where the hole is located, the fully assembled nanopore of the present invention can generally have a single channel located at least substantially in the center of the pore structure. The channel defines a cavity passing through the nanopore, the cavity being perpendicular to the plane axis defined by the membrane. The minimum opening or pore size (e.g., minimum contraction) of the channel in the cross section is suitable for promoting a close fit interaction with folded proteins or other analytes in the solution, or allowing small molecules to pass. Typically, the minimum opening of the cavity is at least 1nm, 2nm, 3nm, 5nm, 7.5nm, 8nm, 8.5nm, 9nm, 9.5nm, 10nm, 11nm, 12nm, 13nm, 14nm or 15nm or larger. Suitably, the width of the cavity is between about 1nm and about 20nm. Suitably, the maximum opening (ie smallest constriction) of the channel is at most 200 nm, 150 nm, 100 nm, 75 nm, 50 nm, 40 nm, 30 nm, 20 nm, 18 nm, 15 nm, 12 nm or 10 nm.

[0106] Application of Nanostructures in Nanopore Sensing Applications

[0107] In one embodiment of the invention, a signal readout is generated by measuring an ionic current that flows from a first side of the membrane to a second side (e.g., cis to trans or trans to cis) through a gradient of soluble ions present in solution. The flow of this current is measurable over a given period of time. This makes the nanostructures of the invention useful in nanopore sensing devices, such as those used in "chip-based" nanopore sequencing and analytical sensor applications. For example, the Oxford Nanopore For sale systems and other devices; GS for sale and GS system; Sales GenomeAnalyzer and System; Iorn sold by Life Technologies System and Iorn Proton Beckman Sales System; and Pacific PacBio for sale and System. It should also be understood that there may be alternative readings for identifying when the analyte molecule is optimally located in the pore cavity. For example, alternative detection methods based on optical methods such as field effect transistors (FETs), quantum tunneling, and fluorescence and plasma sensing can be used. For example, a combination of a solid-state FET nanopore with adjacent nanobelts, nanotubes, or nanowires allows sensing of analyte molecules that interact with the pore cavity, thereby destroying the local ionic current through the hole. In alternative embodiments, lateral electrical measurements of voltage, current, or impedance across the membrane can be performed to generate detectable signal readings in the presence of an analyte. It should be understood that no matter which readout technology is used, a triggered assembly mechanism can prevent current leakage in the absence of stimulation, and can therefore be used to reduce background noise and / or false results.

[0108] Non-naturally occurring amphiphiles and amphiphiles forming semi-fluid amphiphilic membrane layers are known in the art, for example, including block copolymers (Gonzalez-Perez et al., Langmuir, 2009, 25, 10447-10450). Block copolymers can be diblocks (composed of two monomer subunits), but can also be composed of more than two monomer subunits to form a more complex arrangement that exhibits amphiphilicity. Copolymers can be triblock, tetrablock or pentablock copolymers. The membrane can select one of the membranes disclosed in PCT / GB2013 / 052767, the entire contents of which are incorporated herein by reference. Amphiphilic molecules can be chemically modified or functionalized to facilitate the insertion of nanostructures. The membrane can include lipids and amphiphilic polymers, as disclosed in PCT / US2016 / 040665.

[0109] Polymer-based semifluid membranes can be formed by any suitable material. Typically, synthetic membranes are composed of amphiphilic synthetic block copolymers. Examples of hydrophilic block copolymers are polyethylene glycol (PEG / PEO) or poly (2-methyloxazoline), and examples of hydrophobic blocks are polydimethylsiloxane (PDMS), polycaprolactone (PCL), polylactic acid (PLA) or polymethyl methacrylate (PMMA). In a number of embodiments, the polymer membrane used can be formed by amphiphilic block copolymer poly-2-(methacryloyloxy) ethylphosphorylcholine-b-diisopropylamino)ethyl methacrylate (PMPC-b-PDPA). The membrane is generally planar, although in certain embodiments, it can be curved or shaped. Amphiphilic membrane layers can also be supported. Suitably, the membrane is a lipid bilayer or monolayer. Methods for forming a lipid bilayer are known in the art, such as disclosed in International Application No. PCT / GB2008 / 000563. Lipid bilayers are generally formed by the method of Montal and Mueller (Proc. Natl. Acad. Sci. USA., 1972; 69: 3561-3566).

[0110] In a particular embodiment, one or more component modules of the nanostructure may also include one or more binding moieties, which may be composed of affinity binding components or molecules capable of binding affinity binding molecules (such as antigens). When fully assembled, the binding moiety is appropriately located on the component module near the cavity or in the cavity of the nanopore. This helps to bind to the analyte near the first hole. Alternatively, the affinity binding moiety can respond to external stimuli and start the nanostructure assembly process. In another embodiment, the binding moiety is connected to the fully assembled nanopore at a position that allows the binding moiety to extend completely or partially out of the cavity. After the analyte is bound, the current flow through the nanostructure cavity is hindered, resulting in all or most of the current through the cavity being blocked. The level of the signal reading (such as current blocking) can be measured to determine that the analyte has been detected. One or more binding moieties can be attached to the nanostructure and inserted into the membrane by covalent or non-covalent connection (such as by avidin-biotin or His-tag type interaction) before or after assembly.

[0111] The affinity binding moiety may comprise a polynucleotide or polypeptide capable of binding an analyte, which may be used as an external stimulus, present in the solution surrounding the assembly module (before assembly) or the membrane in which the fully assembled nanopore is embedded. When the binding molecule comprises a polypeptide or polynucleotide attached to the assembly module or the nanopore, whether in the cavity, in the pore or close to the cis or trans side of the nanopore, it may be selected from one of the following:

[0112] I. Enzymes - including polymerases, helicases, gyrases and telomerases, as well as nucleic acid binding subdomains or their derivatives

[0113] II. Synthetic or naturally derived affinity binding proteins and peptides, including affimers, antigen binding microproteins, engineered multi-repeat proteins, ankyrin binding domains, lactoferrin, cathepsins, ficolins, collectins, T cell receptor domains and defensins;

[0114] III. Antibodies - including polyclonal, monoclonal, humanized and camelid antibodies, or antigen-binding fragments and derivatives thereof, including Fab, scFv, Bis-scFv, VH, VL, V-NAR, VhH or any other antigen-binding single domain antibody fragment;

[0115] IV. Synthetic or naturally derived affinity binding nucleic acids and nucleic acid analogs, including oligonucleotide probes, aptamers, and ribozymes;

[0116] V. Natural or synthetic small molecules, including drugs, fluorophores, metabolites, and chemokines;

[0117] VI. Antigens or antigenic fragments; and

[0118] VII. Signaling molecules and / or polypeptide receptors thereof, including receptor binding domains and receptor complexes.

[0119] Alternatively, the affinity binding molecule may comprise a small molecule, a lipid moiety, a polysaccharide moiety, a polymer, or any other natural or synthetic moiety capable of undergoing a specific affinity binding interaction with an analyte in solution.

[0120] The assembled nanopore structures of the present invention are suitable for use in sensor applications that allow for the detection of a variety of potential analytes that may be present in the solution being tested. Exemplary analytes may include:

[0121] o Peptides / polypeptides / proteins - folded, partially / fully unfolded;

[0122] oEnzyme;

[0123] oProtein / nucleic acid constructs;

[0124] o Molecules defined by size;

[0125] o within a specific size range (e.g., selected from 1-10 kD, 1-50 kD, 1-100 kD, 10-50 kD,

[0126] 10-100 kD, 20-50 kD and 20-100 kD ranges) of macromolecules;

[0127] o Multiprotein complexes;

[0128] o Antigen or its antibody;

[0129] o Glycoproteins;

[0130] o Carbohydrates;

[0131] o Biopolymers;

[0132] o Toxins;

[0133] o Metabolites and their by-products;

[0134] o Cytokines;

[0135] o Nucleic acids.

[0136] The nanopore structures of the present invention can be incorporated into a plurality of improved devices and sensors. Such devices and sensors are useful in applications where various materials and analytes need to be sensed and characterized. As non-limiting examples, particularly useful applications include genome sequencing, protein sequencing, other biomolecule sequencing, and detection of ions, molecules, chemicals, small molecules, biomolecules, metal atoms, pollutants, polymers, nanoparticles, etc. Such detection and characterization can in turn be used to diagnose disease, develop drugs, identify contamination or adulteration or food or water supplies, and quality control and standardization.

[0137] According to an exemplary embodiment of the present invention, the sensor device generally includes a substrate, the substrate includes a membrane, one or more assembled nanopores are embedded therein, or one or more component modules are anchored on the membrane before being exposed to the assembly trigger. The substrate is placed to facilitate contact with a fluid (optional electrolyte) containing an analyte or assembly trigger molecule. At least one, optionally multiple, device is positioned relative to the substrate, wherein a given device generates a signal (e.g., mechanical, electrical and / or optical) in response to detecting the binding of one or more analytes to the nanopore and / or passing through the nanopore. Multiple devices can be greater than 2, up to 100, or up to 20, or up to 10, or between 2 and 8 devices. Each device may be selected from one or more of the following: field effect sensors; plasma sensors; laser-based sensors; interferometric sensors; waveguide sensors; cantilever sensors; acoustic sensors; quartz crystal microbalance (QCM) sensors; ultrasonic sensors; mechanical sensors; thermal sensors; optical dye-based sensors; fluorescence sensors; calorimetric sensors; photometric sensors; graphene sensors; quantum dot sensors; quantum well sensors; photoelectric sensors; 2D material sensors; nanotube or nanowire sensors; enzyme sensors; electrochemical sensors, including FET or BioFET sensors; potentiometric sensors; conductivity sensors; capacitive sensors; and electron spin sensors. These devices may work together in an array, allowing multiplexed testing of multiple analytes. The sensor device may also include dedicated hardware and systems (e.g., circuits, processors, memory, GUI, etc.) that perform specified functions or actions, or a combination of dedicated hardware and computer instructions so that a properly functioning sensor device can provide meaningful readings to a user.

[0138] Therefore, according to embodiments of the present invention, appropriately configured nanopore devices can achieve various different types of sensor measurements. Generally, electrical measurements include: current measurement, impedance measurement, tunneling measurement (Ivanov AP et al., Nano Lett. 2011 Jan 12; 11 (1): 279-85)) and FET measurement (International Application WO 2005 / 124888). Optical measurements can be combined with electrical measurements or based on electrical measurements (Soni GV et al., Rev Sci Instrum. 2010 Jan; 81 (1): 014301), for example, by converting ionic current into a fluorescent signal from an indicator dye (such as Fluo-8), which is a Ca2+-ionized ion that passes through the nanopore. 2+Flux caused by (Huang et al., Nat Nanotechnol. 2015 Nov; 10 (11): 986-991). As mentioned above, the measurement can be a transmembrane current or voltage measurement, such as measuring the ionic current flowing through the nanopore. Alternatively, the signal can be obtained by measuring the change of the transverse membrane current, voltage and / or impedance value over time.

[0139] Application of Nanostructures in Drug Delivery

[0140] The nanostructures described in the embodiments of the present invention can also be used as membrane-embedded composite sub-gates that open under specific stimuli. Stimuli can allow the incorporation of defined molecular recognition systems into vesicle / liposome-based drug or imaging material delivery platforms. Therefore, the composite sub-gates consist of fully assembled nanopores that regulate the flux of biologically active or other detectable substances on the vesicle / liposome membrane. Bioactive substances, referred to as "pharmaceuticals", may include small molecules, such as drug compounds; or biological products, such as coding or non-coding mRNA, siRNA, aptamers, monoclonal or polyclonal antibodies or fragments thereof and their mimetics. Lanphere et al. (2021)(2021) Ang. Chemie, 60(4): 1903-1908 describe the controlled delivery of therapeutic drugs to the cellular environment under biologically relevant exogenous triggers. Imaging substances may include substances that can be used in nuclear medicine and / or clinical diagnosis, such as radionuclides of technetium, thallium, gallium, iodine and / or xenon. For example, biological imaging substances can be used for more basic research, including in vivo imaging of biological molecules labeled with fluorescent probes. Different imaging materials are suitable for visualizing target molecules, cells, tissues and organs in different ways, depending on the diagnostic or clinical need. The imaging material can be included in a companion diagnostic composition that provides information that is essential for the safe and effective use of the corresponding therapeutic product. For example, the imaging material can include a radioisotope or fluorescently labeled antibody, antibody fragment of its mimetic.

[0141] When administered to a subject, the therapeutic assembly is appropriately administered as a part of the in vivo delivery composition, and a pharmaceutically acceptable carrier may also be included to prepare a pharmaceutical composition. An acceptable pharmaceutical carrier may be a liquid, such as water and oil, including liquids from petroleum, animal, plant or synthetic sources, such as peanut oil, soybean oil, mineral oil, sesame oil, etc. The pharmaceutical carrier may be saline, gum arabic, gelatin, starch paste, talcum, keratin, colloidal silicon dioxide, urea, etc. In addition, auxiliary, stabilizer, thickener, lubricant and colorant may be used. When applied to a subject, a pharmaceutically acceptable carrier is preferably sterile. When the compounds of the present invention are injected intravenously, water is a suitable carrier. Saline solutions and aqueous solutions of glucose and glycerol may also be used as liquid carriers, particularly for injection solutions. Suitable pharmaceutical carriers also include excipients, such as starch, glucose, lactose, sucrose, gelatin, malt, rice, flour, chalk, silica gel, sodium stearate, glyceryl monostearate, talcum, sodium chloride, skim milk powder, glycerol, propylene, ethylene glycol, water, ethanol, etc. If necessary, the pharmaceutical composition may also contain a small amount of a wetting or emulsifying agent or a buffer.

[0142] The drugs and pharmaceutical compositions of the present invention can be in the form of liquids, solutions, suspensions, gels, modified release formulations (such as sustained release or sustained release), emulsions, capsules (e.g., capsules containing liquids or gels), liposomes, microparticles, nanoparticles, or any other suitable formulations known in the art. Other examples of suitable pharmaceutical carriers are described in Remington's Pharmaceutical Sciences, Alfonso R. Gennaro ed., Mack Publishing Co. Easton, Pa., 19th ed., 1995, for example, see pages 1447-1676.

[0143] For any compound or composition described herein, the therapeutically effective amount can be initially determined by in vitro cell culture assays.The target concentration will be that concentration of the active component that is capable of achieving the methods described herein, as measured using methods described herein or known in the art.

[0144] As is known in the art, therapeutically effective amounts for human subjects can also be determined from animal models. For example, a dose can be formulated for humans to achieve a concentration that is effective in animals. As described above, human doses can be adjusted by monitoring the effectiveness of the compound and adjusting the dose upward or downward. Adjusting the dose to achieve maximum efficacy in humans based on the above methods and other methods is well within the capabilities of those of ordinary skill in the art.

[0145] It is contemplated that embodiments of the present invention may include compositions formulated for medical use. Therefore, the compositions of the present invention may be suspended in a biocompatible solution to form a composition that can target a position on a cell, in a tissue, or in a patient or animal (i.e., the composition can be used in vitro, in vitro, or in vivo). Suitably, the biocompatible solution may be a phosphate buffered saline solution or any other pharmaceutically acceptable carrier solution. One or more additional pharmaceutically acceptable carriers (such as diluents, adjuvants, excipients, or excipients) may be combined with the compositions of the present invention in a pharmaceutical composition. EWMartin describes suitable pharmaceutical carriers in Remington's Pharmaceutical Sciences. The formulation of the pharmaceutical preparations and compositions of the present invention meets regulatory standards and may be administered orally, intravenously, topically, intratumorally, or subcutaneously, or by other standard routes. The mode of administration may be systemic, topical, intranasal, or intrathecal. In particular, the specific compositions of the present invention may be administered intravenously, intralesionally, intratumorally, subcutaneously, intramuscularly, intranasally, intrathecally, intraarterially, and / or by inhalation.

[0146] The present invention is further illustrated by the following non-limiting examples.

[0147] Example

[0148] This example demonstrates the use of an exogenous trigger / stimulus to initiate in situ assembly of two independent component modules to form a transmembrane nanopore.

[0149] The controlled assembly of non-spanning subunits into barrel-shaped nanopores is functionally complex and provides a clear opening signal that avoids leakiness of covered nanopores. The novel triggered assembly pores also add scientific breadth by integrating several fundamental processes that underpin their formation: (i) molecular recognition (between trigger and pore subunits) - activating them for interaction, (ii) conformational changes (of pore subunits on the membrane) - preparing them for interaction, and (iii) molecular assembly (of activated subunits) - forming a single nanostructure that defines the pore channel across the membrane. By integrating these three processes, synthetic triggerable nucleic acid nanostructured pores are thus provided that mimic the actions of many specialized natural membrane proteins that have evolved to perform these tasks.

[0150] DNA nanotechnology offers a versatile route to biomimetic design. DNA nanotechnology offers high structural precision, tunability, and dynamic nanomechanical control, as well as chemical modifications for extended functional interactions with biomolecules, including bilayers. Based on these advantages, rational design of DNA has generated barrel-shaped membrane pores with tunable lumen diameters. Molecular dynamics (MD) simulations have been used to investigate the structural dynamics of DNA nanopores and their molecular interactions with bilayer membranes, complementing computational studies of biological pores. DNA design has also led to pores that unlock the channel lumen in response to stimuli, such as oligonucleotides, proteins, or temperature, as well as controllable capping of nanotubes. However, to date, no DNA membrane nanopores with in situ controlled assembly have been constructed. As mentioned above, these DNA structures could be used for sensing, cell biology studies, or drug delivery.

[0151] Here, the inventors used DNA nanotechnology to construct a functionally advanced membrane pore that is assembled from two unique subunits upon trigger activation (Figure 1). The controlled formation integrates the molecular recognition process between the trigger and inactive subunits, the repositioning of the activated subunits within the membrane, and their assembly into a functional channel (Figure 1). The affinity and kinetics of pore formation were determined to reveal any effects of the lipid bilayer on this pathway. In addition, molecular dynamics simulations explored the structural dynamics associated with the repositioning of membrane-bound DNA components during the assembly process. Finally, fluorophore and ion fluxes were detected to probe the transport efficiency of the assembled DNA channel.

[0152] result

[0153] DNA nanopore design. A DNA nanopore capable of assembly from constituent components at a membrane interface (Figure 1) and in solution was designed. Table 6 lists the nucleic acid sequences and Table 7 describes the various components. The nanopore labeled A·B consists of a bundle of four DNA helices assembled from components A and B. Each component consists of two DNA strands forming a central duplex, a ssDNA loop, and two ssDNA arms (Figures 1, 2, and 3). The pore is assembled from the components by hybridization between the ssDNA arms of one component and the loop of another component, thereby forming two additional duplexes. The nominal dimensions of the resulting pore are 7.5 nm in height and 5.1 nm in external width. The width of the internal channel lumen can be up to 0.8 nm.

[0154] To control the formation of holes, two chains L can be used A and L B Make these components lose their assembly activity. A and BL BIn the case of the ssDNA arms, the lock strands sequester them in the second duplex (Figure 1, Figure 3), making them inaccessible to other components. However, the lock strands have 10-nucleotide overhangs, which allow their selective removal by toehold-mediated strand displacement to add key strands (Figure 1, Figure 3) 54 . Add keychain K A and K B The chains are isolated and an assembly interface is displayed, restoring the ability of components A and B to cooperate in forming a pore (Figure 1). To facilitate membrane binding, each component carries a lipophilic cholesterol anchor. After pore assembly, the cholesterol anchors are symmetrically positioned on opposite sides of the pore (Figures 1, 3).

[0155] A·B assembled directly or by trigger activation. Direct pore assembly was first evaluated in solution. Using gel electrophoresis as a readout, separated components A and B appeared as fast-migrating single bands ( Figure 4a , sequencing lanes 1-2), which means there is a homogenous population. In contrast, a mixture of A and B ( Figure 4a , sequencing lane 3) resulted in a clear band shift, indicating the formation of a larger assembled 4-double-stranded hole A·B ( Figure 4a , sequencing lane 3). The lack of other significant bands indicates that the pore assembly is quantitative.

[0156] Pore ​​formation also occurs via triggered assembly in solution. A and BL B ( Figure 4a , lanes 4-5) have higher mobilities than the lockless assemblies A and B, which is consistent with the expectation of a larger mass structure. More consistently, when mixed, the locked assemblies AL A and BL B Migration without additional bands indicative of interaction ( Figure 4a , sequencing lane 6). However, adding a stoichiometric amount of the key K A and K B Causes AL A and BL B The band is completely removed and a higher-mobility A·B band is simultaneously formed that matches the direct pore formation ( Figure 4a , sequencing channel 7). The assembly byproduct, i.e., double-stranded L A K A and L B K B , migrated to the bottom of the gel ( Figure 4a , sequencing lane 7). This further proves that the cholesterol-free component A ΔC , B ΔC and corresponding locking assembly A ΔC L A and B ΔC L BThe data showed that pore formation in solution was not affected by the presence or absence of the cholesterol tag. UV thermal melting analysis also confirmed the equivalence of the pores formed by direct assembly with those of the annealing control. ΔC +B ΔC and (A·B) ΔC The melting curves are very close to ( Figure 7a ), resulting in the same melting temperature (T m ) were 63.1±0.1℃ and 63.1±0.2℃ (n=3, Figure 7b ).

[0157] After successfully confirming that the pore assembly was triggered, the effect of varying the concentration of the key on the assembly was investigated. A Add to corresponding assembly locking component A ΔC L A Even a 10-fold excess of the key will result in the expected component unlocking without any other interaction ( Figure 6a ). When the non-restricted key K B Added to A ΔC L A When the ternary complex A ΔC L A K B The formation of Figure 6b ). In this complex, considering component A ΔC Partial sequence complementarity, K B This is an implicit consequence of the design triggering the assembly of double-stranded bundles. A and B ΔC L B In further experiments, the two keys were added to the two assembled lock components at increasing concentrations ( Figure 6c ). As mentioned before, a 1:1 ratio of key: assembly lock components resulted in quantitative pore formation. However, when an excess of both keys was added, pore assembly was inhibited. The byproduct was the ternary complex A ΔC L A K B and B ΔC L B K A , where the bound mismatched key impedes assembly into the pore. This experiment also led to the formation of a new complex that is slightly above the folded pore (A·B) ΔC Belt ( Figure 6c The new complex is an incompletely folded open barrel structure in which the bound key prevents assembly into the 4-double-stranded pore ( Figure 6c ). This effect could be addressed by adjusting the key design or properties of the triggering stimulus in subsequent hole arrangements.

[0158] Affinity and kinetics of pore assembly in solution. After confirmation of pore assembly, the equilibrium dissociation constant K was determined. d and the kinetic rate constant k on Electrophoretic mobility shift assay (EMSA) was used to derive K d In A ΔC :B ΔC The expected binding curves were obtained by visually tracking pore formation over a range of ratios. Figure 8a , B) and a 1:1 binding stoichiometry. Plot the gel band intensity ( Figure 8a , b) derive the Langmuir-derived K d It is 154±23nM (n=3).

[0159] Since the EMSA-derived K d The limited sensitivity of ethidium bromide staining may have affected the Resonance Energy Transfer (FRET) Detection Method. For this analysis, component A ΔC and B ΔC The pores are labeled with the FRET donor dye Cy3 and the acceptor dye Cy5, respectively. When the dyes are brought into close proximity after pore formation, mixing of the components results in the expected FRET signal. In particular, the Cy3 emission at 563 nm decreases and the Cy5 emission at 670 nm increases ( Fig.14b ). Using the same ratio as for EMSA, at a constant Cy3 A ΔC Upper titration Cy5 B ΔC , but the concentration was 2.5 times lower, and the fluorescence signal was quantitatively analyzed to obtain K d 53.1±5.4nM (n=3, Fig. 9 a, b).

[0160] The FRET-derived K d Using a concentration range 10-fold lower than that of FRET, the K measured by FCCS d 62.2±12.5nM (n=3, Fig.10a ), K obtained in FRET d The results are within the error range (Table 1). A further 5-fold reduction in the concentration of the labeled component did not result in binding saturation ( Fig.10a In further analysis, FCCS did not show any significant difference in the binding strength of the two components; ΔC The concentration of A remains constant, while ΔC When K changes d 71.7±7.7nM (n=2) ( Fig.10b ), in other K dThe error range is within 100%. d The affinity obtained is more than one order of magnitude higher than that obtained for other DNA duplexes of comparable length, which may reflect the molecular difference between the duplex formation restricted in the DNA nanopore and the binding of two conformationally unlocked single-stranded DNA chains.

[0161] Table 1. Use Resonance energy transfer (FRET) from components A and B in solution (A ΔC +B ΔC ), membrane interface (A-SUV+B ΔC Summary of the equilibrium and kinetic data for nanopores A·B assembled on the surface of a membrane (A-SUW+B).

[0162]

[0163] a Mean values ​​and standard deviations were obtained from at least three independent replicates. b k extracted from the kinetic traces after removing the initial fluorescent droplet on

[0164] In determining K d Then, the kinetic rate constant k of pore assembly was measured. on Using EMSA, we examined whether the kinetics of triggered assembly differed from direct assembly. Direct assembly (4.5 ± 0.4 × 10 3 M -1 s -1 , n = 3, Figure 4b , Fig.11a , b) and triggered assembly (4.7±0.5×10 3 M -1 s -1 , n = 3) to obtain k on . Figure 4b ,insert; Fig.11c ,d) is within the error range. This indicates that the triggering mechanism is not rate-limiting.

[0165] We confirmed the EMSA-derived kinetic data using a FRET-based sequential kinetic analysis. The time dependence of the FRET signal after component mixing yielded 11.9 ± 2.8 × 10 3 M -1 s -1 K on (n=3, Table 1, Figure 12), close to the value obtained by EMSA. 3 M -1 s -1The rate constants are two to three orders of magnitude slower than those of typical DNA duplex hybridization. The slower kinetics are likely due to slow nucleation caused by the presence of additional sequences in the non-target arms and loops, and slow zippering caused by significant secondary structure due to the interaction of the arms and loops.

[0166] Pore ​​assembly at membrane interfaces. After characterizing pore formation in solution, we investigated pore assembly at membrane interfaces. We first incubated giant unilamellar vesicles (GUVs) with cholesterol-labeled Cy3 A, then non-cholesterol modified Cy5 B ΔC The overlapping Cy3 and Cy5 fluorescence halos around the GUVs indicate that the two components assemble into pores at the membrane interface (A·B) 1C ( Fig.14a and Fig.13 ).

[0167] To obtain quantitative information on pore assembly on the membrane, we performed FRET assays using small unilamellar vesicles (SUVs). As a baseline, we first Cy3 A was added to SUV, and the Cy3 emission spectrum was obtained ( Figure 3b , pink, Fig. 9 ). Place the second component in a 1:1 ratio. Cy5 B added to the membrane anchor Cy3 A, resulting in a FRET-induced decrease in Cy3 and increase in Cy5 emission, respectively, implying that pore assembly ( Fig.14b , purple). The emission spectrum is overlaid with a control trace of an A·B pore pre-annealed before addition to the SUV ( Fig.14b , dark purple, Fig. 9 ). The spectral equivalence indicates quantitative assembly on the membrane surface. Part of the fluorescence change may be due to Cy3 A and Cy5 B is caused by the proximity of the fluorophore when it is bound to the membrane but not assembled. Adding the assembly in a 1:1 ratio in SUV is inactive Cy3 AL A and Cy5 BL B After that, the FRET change was weaker, resulting in a FRET change equivalent to 32.0 ± 3.9% of the components (n = 3, Fig.15a In contrast, A+B resulted in a 96.3±6.6% signal change compared to the pre-annealing control, confirming pore assembly ( Fig.14c , Fig.15b ).

[0168] To further probe the insertion of the A·B pore into the SUV membrane, melting curves were analyzed, as bilayer insertion is known to result in increased DNA pore stability and higher T m32,65 T of A·B assembled on the membrane and pre-annealed before SUV incubation m The value was 3 °C higher than that of the non-SUV sample (Figure 6), which implies membrane insertion.

[0169] Affinity and kinetics of pore assembly in membranes. Cy3 A anchored SUV adds components Cy5 B and measured the change in FRET to obtain the equilibrium dissociation constant K for pore formation at the bilayer interface d The normalized FRET signal is plotted as Cy5 Function of B concentration ( Fig. 9 e, f), and K d is 55.8±6.2 nM (n=3, Table 1). This is due to the pore formation and cholesterol-free Cy5 B ΔC Combined with SUV Cy3 A assembled K d The value error range is within (Table 1, Fig. 9 ad). Consistently, the pore assembly yields obtained from FRET efficiency (E) calculations indicated that pore assembly proceeded with high yields under all three conditions (Table 2). Similar equivalence in affinity values ​​for pore formation (K) was obtained using EMSA analysis. d =1.64±0.14×10-7M, n=3, Figure 8c , d).

[0170] Table 2 summarizes the FRET efficiency (E) data of pore assembly derived from FRET pore assembly binding titration and pore assembly kinetics. Pre-annealing controls for all three conditions were derived from the FRET binding data and used as a benchmark. Means and standard deviations were calculated from at least three independent replicates.

[0171]

[0172]

[0173] a Derived from the dynamic trajectory of the initial droplet,

[0174] b Derived from the dynamic trajectory after removing the initial droplet

[0175] In contrast, the membrane Cy3 A and Cy5 Kinetic FRET analysis of B assembly (Figure 12) showed that the binding rate constant k on 1.9±0.3×10 3 M -1 s -1(n = 3), which is one order of magnitude slower than in solution (Table 1). It is likely that pore formation on the membrane surface is sterically hindered by the membrane anchor. Cy5 B and carrying membrane anchor Cy3 When the vesicles of A are mixed, k on Values ​​are corrected for the initial sharp drop in Cy3 emission ( Fig.12a ). It most likely happens because Cy5 The rapid binding of B to the membrane leads to Cy3 In support of this interpretation, the magnitude of the rapid initial drop obtained by normalizing the intensity (F / F0) to 0.91 ± 0.03 ( Fig.12a ), close to the previously discussed assembly blocking AL A +BL B The rapid process of membrane binding occurs before the slower pore assembly.

[0176] Experimentally determined K d and k on For calculating the dissociation rate constant k using equation (1) off , assuming a second-order system:

[0177]

[0178] k of pore assembly on membrane off 1.94±0.53×10 -4 s -1 , than 10 -1 -10 -3 s -1 Typical values ​​for simple DNA hybridization between pores are two to three orders of magnitude slower. Given the multiple dissociations required to separate the pore into its two components, a lower k off Other contributions come from the movement of isolated assemblies against lateral membrane pressure and from the reorientation of isolated pore assemblies from a membrane-spanning orientation to a fixed orientation. Quantitative kinetic analysis is complemented by visually tracking pore assembly on supported lipid bilayers using single-molecule FRET (smFRET) and single-particle tracking ( Fig.16 The FRET efficiency obtained from the analysis was 0.39 ± 0.3, which was very close to the value of assembly on the vesicle membrane (Table 2).

[0179] The influence of spatial factors on hybridization was explored. The kinetic analysis of A·B pore formation on the membrane showed that k on and k offThe reason for this is that duplex formation and dissociation require DNA assemblies to change from a membrane-adherent state to a membrane-spanning state and back again, respectively. This theory is confirmed by model systems in which DNA hybridization occurs outside the membrane and is therefore expected to be less affected by steric factors. The model is based on the DNA duplex hybridization of a 20nt DNA strand S with a complementary strand R ( Fig.17 , Figure 18, Table 8 and Table 9). To explore the effect of steric volume, S optionally carried a 15.5×5.5 nm six-double-stranded DNA barrel, and R optionally was cholesterol anchored to the SUV membrane. Hybridization was assessed under all conditions using EMSA and FRET (Figure 19, Figure 20, Figure 21, Fig. 22 As expected, EMSA analysis of DNA hybridization (Figures 19 and 21) showed that K d was largely unaffected by the presence or absence of nanobarrels (Table 3, Figure 19, Figure 21). Membrane anchoring resulted in a weakened affinity, but in the presence of nanobarrels, affinity was unaffected. Similarly, k on The values ​​were also not affected by volume or membrane anchoring (Table 3, Fig. 22 , Fig.23 ). These values ​​are also consistent with literature studies of DNA duplex formation. This suggests that the slower kinetics of A·B pore assembly is largely a result of the slow nucleation and zippering steps discussed previously. Duplex insertion in the bilayer may also have an impact on the kinetics.

[0180] Table 3: Summary of thermodynamic and kinetic data obtained under four conditions for the model system. R vs. S and R vs. S NP The assembly of R in solution and when R was combined with SUV composed of DOPC:DOPE in a 7:3 molar ratio, R SUV Assembly with S at the membrane interface; and R SUV With S NP The mean and standard deviation were obtained from three independent replicates.

[0181]

[0182] As further insight into the DNA duplex model, the extent of FRET-derived membrane interface assembly decreased by approximately 40% compared to solution (Table 4), consistent with previous reports. In contrast, the yield of A·B pore assembly was not only not significantly affected by the membrane, but was also higher than that of the model system under all conditions (Table 2). This high yield is attributed to the very favorable pore formation mentioned previously (Tables 1 and 2).

[0183] Table 4. Summary of FRET efficiencies based on hybridization kinetics between the four components of the model system. Means and standard deviations were calculated from at least three independent replicates.

[0184]

[0185] To investigate the orientation of A·B at the membrane interface, we first used a nuclease digestion assay to probe the orientation of A·B relative to the bilayer membrane ( Fig.24 ). In this assay, DNA pores inserted into the membrane are partially protected from nuclease digestion compared to DNA pores that are more spatially accessible in solution. In fact, having zero, one, and two cholesterols (A·B) ΔC 、(A·B) 1C and the solvation pores of A·B are quickly and completely digested ( Fig.24 In contrast, incubation of the wells with large unilamellar vesicles (LUVs) prior to nuclease incubation resulted in the expected cholesterol-dependent reduction in digestion rate and extent ( Fig.24 ). Non-cholesterol (A·B) ΔC is not affected because it is not bound to the space protective membrane. In contrast, (A·B) 1C Obtained LUV protection ( Fig.24 ), because cholesterol alone is expected to orient the pore parallel to the membrane (Fig. Figure 3b ), making it more inaccessible to nucleases. Finally, dicholesterol-labeled A·B experienced the greatest nuclease protection in the pores expected to span the lipid bilayer ( Fig.24 )(Figure 1, Figure 3b ). These measurements do not indicate what the percentage of (A·B) pores is.

[0186] To confirm the cholesterol-dependent orientation of our DNA nanopores, dichroism spectroscopy was used. The helical structure of the nanopore was determined using circular dichroism (CD) spectroscopy. In the absence of a membrane, the CD spectra of the A·B constructs containing 4, 1, or no cholesterol showed the characteristic features expected for B-form DNA, with a negative peak at 245 nm and a positive peak at 280 nm ( Fig.25a ). The lack of competing peaks in the CD spectra indicated that all structures formed typical B-type helical structures.

[0187] Linear dichroism (LD) was then used to probe the orientation of the A·B pore relative to the SUV membrane. The positive peak at 260 nm in the LD spectrum indicated that the A·B ( Fig.25b , dark purple) oriented perpendicular to the bilayer 73-74 , indicating a transmembrane orientation. Further in agreement, the pore variant (A·B)4C containing four cholesterols produced similar spectra ( Fig.25b , upper line). Control wells without cholesterol (A·B) ΔC The spectrum has flat spectral lines ( Fig.25b, bottom line), lacks membrane interaction as expected. 1C The broad negative LD peak ( Fig.25b , gray) indicates that its dynamic orientation is parallel to the bilayer. The noisy nature of the spectrum is believed to be partly due to the effects of light scattering by SUVs with a diameter of approximately 180 nm.

[0188] Molecular dynamics (MD) simulations provided insight into the dynamics of the pore assembly and pore structure. The membrane-dependent interactions of component A and the assembled A·B were further investigated using atomistic molecular dynamics. Insights from experimental data were used to inform the initial configuration of the simulation trajectories. The per-residue root mean square fluctuations (RMSF 10 ) studied structural dynamics ( Fig.26 Figure 27 Fig.28 Baseline analysis of component A in solution yielded a highly unstable structure with an average area fluctuation of 0.72 ± 0.38 nm ( Fig.25c , top left; Fig.28 However, the addition of lipid bilayer stabilized the mean area fluctuation of component A at 0.49 ± 0.17 nm ( Fig.25c , upper right corner; Fig.28 This stabilization may be caused by the proximity of the DNA nanostructure to the membrane mediated by the cholesterol tag, resulting in electrostatic interactions between the DNA duplex and the lipid head groups ( Fig.29 ).

[0189] Compared to component A, pore A·B in solution is significantly more stable but still dynamic, with an average area RMSF of 0.42 ± 0.14 nm ( Fig.25c Figure 27 Fig.28 The simulation also showed that the A·B structure is connected by terminal cross-links, and the gap in each double strand is a hinge point ( Fig.25c , Figure 2). Consistent with previous reports, the TEG cholesterol group is highly mobile and hovers within attached or adjacent duplexes. In contrast, membrane interactions stabilize the A·B pore, resulting in a reduced regional average RMSF of 0.35 ± 0.15 nm and a compact pore geometry ( Fig.25c , Fig.28 As a possible reason for the stabilization, the lipid bilayer reduces the structural flexibility and chain scission in the central cavity of the pore.

[0190] The stability of the duplex (RMSF 0.2-0.3 nm) located between two cholesterol-modified duplexes (RMSF 0.3-04 nm) was also improved ( Fig.25c , Figure 27). This stabilization may be due to the proximity of the linker cholesterol group to the unmodified duplex, which reduces the dynamics of the surrounding phospholipid molecules.

[0191] The membrane-induced change from a spherical structure to a dense pore structure was confirmed by comparison with the average intrafluorophore distance of Cy3-Cy5 on A·B (Table 5), which is a useful indicator of pore size. The Cy3-Cy5 distance is derived from the FRET efficiency as described 78 The experimental data support the membrane-induced compression of the inserted pore, because the 7.10 ± 0.50 nm non-transmembrane pore (A·B) of the A·B pore in the SUV ΔC The intrafluorophore distance of the fluorophore decreased slightly to 6.63 ± 0.15 nm. Consistently, the control wells (A·B) in the membrane-bound but non-compressed state 1C A solution-phase distance of 7.05 ± 0.14 nm was maintained.

[0192] Table 5. Calculated Cy3-Cy5 distances as an approximation of the average nanopore diameter A·B for each condition. When Cy3-Cy5 has R0 = 5.6 or 6 nm, the distances were calculated using the FRET efficiencies derived from the prefolding controls in Table S3.

[0193]

[0194] Membrane-pore interactions alter lipid bilayer structure and dynamics. MD simulations were also used to assess whether and to what extent the membrane-interacting component A and pore A·B alter lipid bilayer structure and dynamics. After simulations, the attachment of component A to the membrane resulted in the smallest change in bilayer structure ( Fig.30a ). After membrane bonding, component A becomes flat on the membrane surface ( Fig.30c , Fig.29 ).

[0195] In contrast, the transmembrane orientation of A·B is achieved by forming a toroidal lipid array around the pore ( Fig.30b , inset shows the initial state) leads to significant lipid remodeling, consistent with previous modeling. The ring-shaped arrangement protects the hydrophobic membrane core from the hydrophilically charged DNA helices ( Fig.31 , Fig.32 As a result of lipid rearrangement, the bilayer widens, as indicated by the increase in area per lipid head group (Table 6). This widening is a result of the geometric constraints imposed by lipid packing and maximization of interactions between the DNA phosphate backbone and the lipid head groups. The formation of the ring-shaped arrangement is accompanied by the alignment of the cholesterol anchors parallel to the fatty acid tails of the phospholipids ( Fig.30b ), and the upward motion of A·B relative to the bilayer plane. The cholesterol moiety stabilizes the surrounding bilayer, as shown by the decrease in RMSF near the lipid anchor ( Fig.30d ).

[0196] Table 6. Area per lipid (APL) values ​​for the two membrane simulations. APL was calculated using MEMBPlugin in VMD and averaged across the NpT production simulations, discarding the initial 10% of trajectories. These values ​​compare favorably to literature values ​​for POPC membranes, but show increased APL for trajectories of transmembrane DNA nanostructures.

[0197]

[0198]

[0199] MD simulations were used to map the channel lumen. MD simulations were used to elucidate the shape of the channel lumen. Cluster analysis of the transmembrane A·B pore trajectories was performed to generate a series of coordinates, which were then analyzed using HOLE software - J. Mol. Graph. 1996, 14, 354-360 ( Fig.30e , Fig.33 , Fig.34 ). The simulations showed that the pore has a dynamically changing cavity in the middle section, with a diameter ranging from 0.60 ± 0.19 to 1.02 ± 0.18 nm, with an overall average of 0.83 ± 0.14 nm. The pore is narrower at the top and bottom where the component duplexes are physically cross-linked. However, there are also differences in the diameter of these two regions. A third narrowing of the channel is observed at the location of the cholesterol tag, which may be due to dynamic compression induced by the bilayer.

[0200] The triggered assembly of A·B on the membrane surface leads to functional nanopores. After characterizing the pore formation and its interaction with the membrane, the pore activity was characterized. In particular, it was determined whether the A·B formed on the membrane surface played a role in the bilayer spanning the nanopore ( Fig.35a , b). Single channel current recording (SCCR) addressed this question. Pre-annealed A·B was first examined to establish a reference for the conductivity of a single nanopore. Addition of A·B to a planar lipid bilayer resulted in a current shift to -54pA at -50mV ( Fig.35c The current was recorded as a function of voltage for 16 independent insertions to confirm the presence of the DNA pore. The ohmic current-voltage dependence was between ±100 mV ( Fig.35d ) matches the expected signal for perpendicular symmetry A·B, as does other similar DNA nanopores in the field. The average conductivity obtained from the analysis was 0.70±0.27nS( Fig.35e This is consistent with the theoretical conductance of 0.67 nS obtained based on MD simulation cavity analysis with a nominal cavity diameter of 0.8 nm ( Fig.30d ).

[0201] After characterizing the pre-annealed A·B, we investigated whether triggering A·B assembly on the membrane surface would result in similar pore properties compared to directly assembled pores.A and BL B Incubation with a planar lipid bilayer did not result in a change in current, indicating a lack of membrane puncture ( Figure 35f ). However, the key K A and K B The addition of leads to the assembly of pores on the membrane surface, as evidenced by the characteristic increase in current to 71 pA at +90 mV ( Figure 35f ), consistent with the insertion of membrane A·B. A small additional current step of 4.0 pA ( Figure 35f , asterisks) indicate that when the pore is inserted into the lipid bilayer, the duplex undergoes a small-scale local rearrangement. Further analysis of eight independently inserted triggered assembled pores yielded an average pore conductivity of 0.69 ± 0.12 nS ( Figure 35g ), which is within the error range of the preassembled pore. Electrical recordings confirmed that triggering the assembly of A·B after adding the key produced a nanopore that was functionally identical to the directly assembled pore.

[0202] A·B transports molecular substances through the lipid bilayer. According to MD simulations and SCCR experiments, the central pore width of the assembled nanostructure is about 0.8nm, which should support the flow of small molecules. To investigate this, direct assembly of components A and B or locking component AL was studied. A and BL B Trigger assembly and adding unlocking key K A and K B Figure 3 Molecular transport through the formed A·B pore. In the transport assay, the fluorophore sulforhodamine B (SRB), with a molecular weight of 558.7 g / mol, was encapsulated within the SUVs, where it was contact quenched but increased in brightness as it flowed across the membrane pore into the environment. As expected, there was no dye flux ( ) when the SRB-filled SUVs were incubated with either the individual components or the assembled locked components without the key. Fig.36a , Figure 37).

[0203] Addition of 400 nM of direct and triggered assembly A·B to vesicles resulted in very similar dye efflux (3.57 ± 0.14% and 3.33 ± 0.26%, respectively) ( Fig.36a ), the dye efflux was lower when using a 200 nM pore (Figure 37). When using a 400 nM pre-annealed pore A·B, the flux was 5.25 ± 1.05%. The data indicate that the in situ assembled A·B formed a functional nanopore. The lower transmission at about 3-5% is consistent with the expected slow diffusion of SRB in a relatively narrow lumen of 0.8 nm diameter (about 0.7 nm32). Designing a pore with a wider diameter lumen channel would allow for faster diffusion.

[0204] A·B forms a synthetic Ca2+ The dye flux analysis above indicates that the narrow cavity of A·B makes it suitable for transporting substances smaller than fluorescent dyes. Due to its small size (0.23 nm) and positive charge, it is expected that Ca 2+ The transport efficiency in the pore is even higher than that of SRB, thus complementing the data on ion transport from SCCR. 2+ The sensitive ratiometric dye Fura-2 was encapsulated inside the SUV at a concentration of 100 μM, and 250 μM CaCl2 was added to the ambient fluid. In the absence of pores, the SUV membrane is responsive to Ca 2+ is impermeable, as confirmed by fluorescence analysis ( Fig.38a In addition, dynamic light scattering determined that CaCl2 did not disrupt the integrity of the vesicles or significantly change the diameter of the vesicles ( Figure 38b ). Add 25nM pre-assembled pores (A·B) 4C After that, a large amount of Ca 2+ Inflow, reaching 79.2% signal after 10 minutes ( Fig.36b ). After only 70 seconds, 75nM (A·B) 4C The flux of Ca2+ was 90.3±3.1% (n=3) higher than that of pores. 2+ Internal flow (0.94±0.13%, n=3).

[0205] in conclusion

[0206] The present embodiment pioneered the development of a synthetic DNA nanopore that is formed by triggering the assembly of inactive pre-pore components. Previous DNA pores were preformed in solution and integrated into the membrane as complete pores. The formation of the present pore is either carried out by direct assembly of pore subunits or by reactivating the pore assembly by activating the assembly lock component with a DNA key (as an external stimulus). Both routes produce the same assembly yield and pore function. Pore formation controlled by nucleic acid subunits and external triggers does not occur in nature. But this concept is related to biological protein pores, which can be assembled from membrane-connected subunits. Oligomeric pores are usually formed through an intermediate non-transmembrane pre-pore state, which matures into a membrane-piercing pore through spontaneous conformational changes (such as hemolysin pores) or changes triggered by proteases (usually found in membrane attack complexes). However, it is understandable that the dynamics of protein pore assembly in vivo do not make the efficient ability of nucleic acid nanostructures to assemble into nanopores under current conditions less surprising and unexpected.

[0207] The biomimetic DNA pores described in this embodiment provide insight into the processes that underlie controlled pore formation. By analyzing the affinity and kinetics of nanopore assembly, the effects of the membrane on molecular interactions and assembly were determined. DNA hybridization is slowed down by an order of magnitude because the assembly of the pore subunits requires a change from membrane connection to membrane spanning direction. Conversely, the rate at which the pore decomposes into subunits is slowed down because this requires a transition from the connection direction to the spanning direction. This also reduces the structural flexibility of the DNA structure due to the stabilizing effect of the pores around the lipid bilayer. Previously known nanopores do not involve similar changes in DNA binding or dissociation. Therefore, this embodiment allows the creation of a variety of dynamic functional nanostructures at the membrane interface and helps to further understand the molecular processes on the membrane. By aiming to establish synergies between chemistry and life sciences, this embodiment makes a wide range of customizable nanodevices possible for use in biomedicine, synthetic biology, sensor technology, and chemical biology.

[0208] 1. Experimental information

[0209] 1.1. Materials

[0210] Unmodified, fluorophore-labeled, and cholesterol-modified DNA oligonucleotides were purchased from Integrated DNA Technologies at a 100 nmol scale and subjected to HPLC purification. 1,2-Dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE), 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphohydroquinone (POPC), and 1,2-diphenyloyl-sn-glycerol (DPhPC) were purchased from Avanti Polar Lipids (USA). All other reagents and solvents were purchased from Merck (UK) unless otherwise stated.

[0211] 1.2. DNA assembly

[0212] An equimolar mixture of DNA oligonucleotides (1 μL of each, stock concentration of 100 μM) (see Table S2 for the composition of DNA pores and assemblies) was dissolved at 1 μM in buffer A (1× PBS; 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7.4), buffer B (300 mM KCl, 15 mM Tris-HCl, pH 7.4). Folding was achieved on a BioRad T100 thermal cycler (UK) using a program that involved heating to 95°C for 0.5 min, followed by cooling to 75°C over 5 min, holding for 1 min, and then cooling to 4°C at a rate of 0.5°C per 1 min. Samples were stored at 4°C for up to 1 week.

[0213] 1.3.PAGE

[0214] The assembled DNA nanostructures and component DNA oligonucleotides were analyzed using commercial 10% polyacrylamide gels (BioRad, UK) in 1×TBE buffer (100mM Tris, 90mM boric acid, 1mM EDTA, pH 8.3). For gel loading, the DNA nanopore solution (2μL, 1μM) was mixed with folding buffer (13μL, 2mM MgCl2, 0.6x TAE, pH 7.4) and 6x gel loading dye (5μL, New England Biolabs, UK). The gel was run at 115V for 90 minutes at 4°C. The gel bands were observed by ethidium bromide staining and UV irradiation. A 100bp marker (New England Biolabs, UK) was used as a reference standard.

[0215] 1.4. Agarose gel electrophoresis

[0216] The assembled DNA nanostructures and component DNA oligonucleotides were analyzed with 2-3% agarose (Invitrogen, UK) gels in 1×TAE buffer (40mM Tris, 20mM acetic acid, 1mM EDTA, pH 8.3). For gel loading, the DNA nanostructure solution (2 μL, 1 μM) was mixed with folding buffer (13 μL) and 6x gel loading dye (5 μL, New England Biolabs, UK). The gel was run at 60V for 60 minutes at 4°C. The gel bands were observed by ethidium bromide staining and UV irradiation. A 100 bp marker (New England Biolabs, UK) was used as a reference standard.

[0217] 1.5. Preparation of small unilamellar vesicles (SUV)

[0218] DPhPC (100 μL, 10 mM) or POPC (100 μL, 10 mmol) in chloroform was added to a 5 mL round-bottom flask. The solvent was removed using a rotary evaporator (Buchi, Newmarket, UK) to obtain a thin film, which was further dried under high vacuum for 1 hour. The lipids were resuspended in 1 mL of buffer A or buffer B. The solution was sonicated at 30 ° C for 20 minutes, then equilibrated for 1 hour, and then extruded 25 times through a 0.1 μm polycarbonate membrane (AvantiPolar Lipids, US) using an extruder kit (Avanti Polar Lipids, US). The SUVs were then stored at 4 ° C and used within 48 hours.

[0219] 1.6. Analysis of melting temperature (T) using UV-Vis spectroscopy m )

[0220] UV melting curves were obtained using 10 mm quartz cuvettes (HellmaAnalytics, Southen-on-Sea, UK) in a Varian Cary 300Bio UV-Vis spectrophotometer (Agilent, Cheadle, UK) equipped with a Peltier element (Agilent, Cheadle, UK). Samples were analyzed at a concentration of 200 nM, and SUVs composed of DPhPC were analyzed at a lipid concentration of 200 μM. Samples were analyzed by monitoring the change in absorbance at 260 nm when the temperature was increased from 20°C to 80°C at a rate of 1°C / min. The melting curves were then background corrected and the first derivative was calculated to determine T m .

[0221] 1.7. Electrophoretic mobility shift analysis

[0222] For binding titrations, add component A ΔC or A (5 μL, 1 μM) with component B ΔC or B (1 μM stock solution) in buffer A in a final volume of 20 μL at concentrations ranging from 0 to 0.5 μM. ΔC In the case of A or B, component A (5 μL, 1 μM) was first added to SUV (5 μL, 100 nm, 16.7 nm). After incubation at 30 °C for 30 min, 6x gel loading dye (5 μL, New England Biolabs, Hitchin, UK) was added, the samples were mixed and loaded onto a thermally equilibrated 2-3% agarose gel. The gel was run for 60 min at 4 °C, 60 V, 1× TAE buffer, pH 8.3. Staining and molecular labeling were as described in Section 1.4. Band intensities were analyzed using ImageJ and normalized to (1-(I A -I 背景 )). The normalized intensity is then fitted to a Langmuir curve to determine K d .

[0223] Kinetic assembly titration, component A ΔC (5 μL, 1 μM) and B ΔC (5 μL, 1 μM) was mixed in buffer A to a final volume of 20 μL. Samples were incubated at 30°C for 0, 1, 5, 10, 15, 20, 25 and 30 minutes while shaking at 500 rpm. Samples were prepared in reverse chronological order and after all samples were prepared, they were crushed in ice to prevent pore formation. For assembly of locking assembly A ΔC L A With B ΔC L B , a key K is added at each time point A and KB (1 μL, premixed, 5 μM). The samples were mixed with 6x gel loading dye (5 μL) and then loaded onto a thermally equilibrated 10% PAGE. The gel was run at 115 V at 4°C in 1× TBE buffer for 90 minutes. Staining and molecular labeling were as described in Section 1.3.

[0224] FRET analysis of pore assembly

[0225] For binding titrations, the assembly of A·B was investigated using a fluorescence spectrophotometer (Cary Eclipse, Agilent, Cheadle, UK). ΔC ,A or AL A (12 μL, 1 μM), B ΔC , B or BL B (0 μL, 1.2 μL, 2.4 μL, 6 μL, 12 μL, 24 μL; 1 μM), SUV (0 μL, 6 μL; 1 mM lipid, 7.22 nM SUV) and buffer B to a final volume of 120 μL. The tubes were then incubated at 30 °C for 30 min while shaking at 750 rpm. The combined solution was then added to a 10 mm quartz cuvette (Hellma Analytics, Southend on Sea, UK), placed in a fluorescence spectrophotometer and scanned (ex545 nm, em555-725 nm). Prefolded A·B was used as a control for maximum assembly. In the case of SUV, A or AL were placed in A Mix with SUV and add B ΔC , B or BL B The donor (Cy3) emission intensity is A ΔC or A and pre-folded control wells (A·B), (A·B) 1C Normalization was performed between A and B. A 1:2 A:B was used as an internal control. Since the donor (Cy3) was able to donate to multiple acceptor (Cy5) molecules, it was set to the same binding level as the pre-assembled control and used as the K d Anchor point for determination.

[0226] For kinetic assembly, the assembly of A·B was studied by monitoring the Cy3 emission (ex550nm, em570nm) using a fluorescence spectrophotometer (Cary Eclipse, Agilent, Cheadle, UK). ΔCor A (2.5 μL, 1 μM) was added to SUV (0 μL, 1.25 μL; 1 mm lipid, 7.22 nM SUV) and buffer B (97.5, 96.25 μL) in a 10 mm quartz cuvette (HellmaAnalytics, Southend on Sea, UK), and the signal was allowed to stabilize for 5 min. Then, B was quickly added ΔC Or B (50 μL, 1 μM) and mixed. Pore formation was monitored for 1 hour. In the case of SUVs, A and SUVs were mixed and left to bind for 10 minutes before the start of the run.

[0227] FRET efficiency (E) was calculated using equation (2):

[0228]

[0229] I DA is the donor intensity when both donor and acceptor are present; I D is the donor intensity in the absence of the acceptor.

[0230] The interfluorophore (Cy3-Cy5) distance was calculated using equation (3):

[0231]

[0232] E represents the FRET efficiency, r is the donor-acceptor separation distance, and R0 is Distance, where E = 50%.

[0233] 1.9. Two-color FCCS analysis for measuring K of pore assembly d

[0234] Two-color fluorescence cross-correlation measurements were performed on a commercial laser scanning microscope (ConfoCor 3, Carl Zeiss, Jena, Germany) equipped with a 40× water immersion objective. ΔC and B ΔC The samples were labeled with spectrally non-overlapping fluorophores Alexa488 and Alexa647, respectively. A 635 nm secondary dichroic mirror was used for cross-correlation measurements, a 505-540 nm bandpass filter was used in the green channel, and a 650 nm longpass filter was used in the red channel. The laser power was adjusted so that the brightness ratio of Alexa647 to Alexa488 was approximately 3:1.

[0235] At a 1:1 binding stoichiometry, Alexa488 A ΔC (X g )or Alexa647 B ΔC (X r) can be calculated based on the number of dual-labeled particles (N rg ) is calculated relative to the total number of each particle (N g express Alexa488 A ΔC , N r express Alexa647 B ΔC ), using equations 4 and 5:

[0236]

[0237]

[0238] detected in the green or red channel, respectively Alexa488 A ΔC or Alexa647 B ΔC The number of particles is calculated by the single-component model of two-dimensional translational diffusion shown in Equation 6 for the corresponding autocorrelation (for N g and N r ) and cross-correlation (for N rg ) function G(τ) is fitted to obtain:

[0239]

[0240] where represents the diffusion time through the confocal volume. Considering the green and red detection volumes V by using the following formula g and V r The size difference of the bound particles is X g and X r To make a correction:

[0241]

[0242]

[0243] Where V eff represents the effective cross-correlation volume, which is defined by the following equation:

[0244]

[0245] where S is a structural parameter that is set equal for both channels. g =S r = 6, ω is the corresponding lateral radius ω of the confocal volume of the green or red confocal volume g or r The confocal volume V in the red and green channels is calculated as follows:

[0246] V=π 3 / 2 *ω 3 *S (10)

[0247] We measured the markers Rhodamine 6G (Merck, Germany) and Alexa647 maleimide (Jena Bioscience, Germany) with known diffusion coefficients (D) by calibration experiments using the following equation:

[0248] ω 2 =4Dτ D (11)

[0249] On each day of data collection, the maximum achievable cross-correlation was determined using DNA duplexes carrying Alexa488 and Alexa647 fluorophores (IDT, USA) to represent 100% binding.

[0250] For binding measurements, Alexa488 A ΔC and Alexa647 B ΔC Mix in Buffer B to a final volume of 60 μL. In other measurements, Alexa647 B ΔC The concentration of Alexa488 A ΔC The concentration of (7.5 μL, 100 nM) was kept constant. The samples were then incubated with shaking at 30°C for 30 minutes. After incubation, the mixtures of each ratio were measured separately. Alexa647 B ΔC was kept constant (9.6 μL, 100 nM), while Alexa488 A ΔC Measurement results of concentration changes.

[0251] To determine K d , the volume correction fraction of bound particles X g / v and X r / v Fit to the Langmuir isotherm:

[0252]

[0253] where C represents the concentration of the changing component.

[0254] By design, dc-FCCS experiments showed saturation values ​​below 100%, with a slight overestimation of the cross-correlation observed in the red channel ( Fig.10a). The former originates from an incomplete overlap of the red and green laser confocal volumes. The latter originates from the bleeding of green-labeled particles into the red channel at a higher green to red particle ratio, which is evident at ratios above 10:1.4. To minimize the effect, we performed a control measurement using a cross-correlation standard before each FCCS experiment. This is a fluorescent dual-labeled DNA duplex. The control measurement showed the same deviation from 100% saturation within a precision of 5%, with the same saturation ratio between the red and green channels.

[0255] 1.10. Preparation of giant unilamellar vesicles (GUVs)

[0256] A POPC lipid solution (5 μL, 10 mM in chloroform) was added to an indium tin oxide (ITO) coated glass slide. Within 5 minutes, the solvent evaporated to form a dry lipid film. The glass slide was then inserted into a vesicle preparation device (Nanion Technologies, Munich, Germany). An O-ring was added around the membrane. Sucrose (300 μL, 1 M aqueous solution) was added to the lipid membrane restricted by the O-ring. Finally, another ITO glass slide was applied from the top to form a sealed chamber. An AC electric field was applied between the two glass slides for 120 minutes using a voltage program of 3 V, 5 Hz. The solution was collected and stored at 4 °C.

[0257] 1.11. Confocal laser scanning microscopy

[0258] The GUV suspension (10 μL, 130 M lipid concentration) was added to a FluoroDish (World Precision Instruments, Hitchin, UK) containing buffer (500 μL, 1× TAE, 500 mM NaCl, pH 8.1). The solution was gently mixed. After adding component A (10 μL, 1 M) to the dish, the solution was thoroughly mixed and allowed to stand for 10 minutes to ensure membrane binding. Component B was then added. ΔC (10 μL, 1 M), and then mix the solution. The mixture was left for 5 minutes to allow the GUVs to sink to the bottom of the dish. The FluorDish (World Precision Instruments, Hitchin, UK) was placed under a microscope and the GUVs were visualized using a 96x optical zoom. The sample was then observed by bright field. Cy3 A is 570nm, Cy5 B is 670nm, and images are collected.

[0259] 1.12. Preparation of planar lipid bilayers on glass slides and smFRET and single particle tracking

[0260] Planar lipid bilayers were formed on glow-discharged slides provided by Oxford NanoImaging (Oxford, UK). SUVs consisting of DPhPC in buffer A (15 μL, 1 mM) were placed on a support and allowed to stand for 15 minutes. Some of the solution (~5 μL) was then replaced with H2O and allowed to stand for 1-2 minutes. Repeat 3 times. After the third wash with H2O, the solution was washed with buffer A. The slides were used within 1 hour and topped up with buffer A as needed. smFRET and single-particle tracking were performed by Jon Shewrew of Oxford NanoImaging using a NanoImager S (Oxford NanoImaging, Oxford, UK). The constructs (1 μL, 1 nM in buffer A) were added to the planar lipid bilayer consisting of DPhPC on the slide.

[0261] 1.13. Enzyme digestion

[0262] A chloroform solution of lipid DPhPC (100 μL, 10 mM) was added to a 5 mL round-bottom flask. The solvent was removed using a rotary evaporator (Buchi, Newmarket, UK) to obtain a thin film. The lipid was resuspended in buffer B (1 mL), sonicated at 30 °C for 20 min, and then equilibrated for 3 h. ΔC 、(A·B) 1C or A·B (2 μL, 1 μM), nuclease-free water (58 μL) and 2× Bal-31 buffer (60 μL; 40 mM Tris-HCl, 1.2 M NaCl, 24 mM MgCl2, 24 mM CaCl2, 2 mM EDTA) were added to 10 quartz test tubes (HellmaAnalytics, Southend-on-Sea, UK). For vesicle assays, (A·B) were first added. ΔC 、(A·B) 1C Or A·B (2 μL, 1 μM) was incubated with LUV (18 μL, 1 mM DPhPC) for 1 hour and then added to a 10 mm quartz cuvette with nuclease-free water (40 μL) and 2x Bal-31 buffer (60 μL, New England Bioscience, Hitchin, UK). Fluorescence was monitored at 570 nm using a Varian Cary Eclipse fluorescence spectrophotometer (Agilent, Cheadle, UK) and excited at 555 nm. After 5 minutes, Bal-31 (0.75 μL, New England Bioscience, Hitchin, UK) was added and fluorescence emission was monitored for 15 minutes.

[0263] Circular dichroism

[0264] Solution phase circular dichroism (CD) spectroscopy was performed on a Jasco-810 spectropolarimeter (Kromatec Ltd, Great Dunmow, UK). A microvolume quartz Couette flow cell (Kromatec Ltd, Great Dunmow, UK) with an annular gap of approximately 0.5 mm and quartz capillaries was used. CD spectra of DNA nanopores (1.4 μM) were obtained between 320-190 nm.

[0265] 1.15. Linear dichroism

[0266] Solution phase flow linear dichroism (LD) spectroscopy was performed on a Jasco-810 spectropolarimeter (Kromatec Ltd, Great Dunmow, UK) using a photoelastic modulator 1 / 2 wave plate. A microvolume quartz Couette flow cell with an annular gap of approximately 0.5 mm and a quartz capillary was used (both from Kromatec Ltd, Great Dunmow, UK). Molecular alignment was achieved by applying a constant flow of the sample solution between two coaxial cylinders, a fixed quartz rod and a rotating cylindrical capillary. LD spectra were obtained with laminar flow obtained by maintaining the rotation speed at 3000 rpm and processed by subtracting the non-rotating baseline spectrum. DNA nanopores were detected at a lipid concentration of 1.4 μM and SUVs composed of POPC were detected at a lipid concentration of 500 μM.

[0267] 1.16. Simulation preparation

[0268] The DNA nanopore A·B and component A were recreated in caDNAno and then converted to all-atom models in python. The polythymine linker regions at the ends of the pore were then constructed using the MolSoft ICM software suite. The TEG cholesterol lipid anchor was parameterized using cgenff7 and connected using pyMol. A topology file compatible with CHARMM36 was then generated using psfgen. The initial structures of A·B and A were minimized in vacuum for 10,000 steps (2 fs) and then simulated for 100,000 steps (2 ns) using an elastically constrained network exported from the ENRG web server.

[0269] Simulated DNA nanopores A·B and assembly A in 1 M KCl and TIP3 water prepared in VMD.

[0270] Nanopore A·B was simulated in a 13×11×15 nm box with a total of 437k atoms, and component A was simulated in a 16×14×19 nm box with a total of 6.5k atoms. A 1ns NpT equilibration was performed to balance the box size and pressure before a 50ns NvT equilibration to further relax the DNA nanostructure. Production simulations were then run in the NpT ensemble.

[0271] For the simulations of the membrane-tethered assembly A and the membrane insertion A·B, VMD was used to generate membranes and orient the DNA nanostructures while maintaining a favorable cholesterol content. The orientation of each structure was determined by experimental data on linear dichroism. The transmembrane nanopore A·B was simulated in a 12 × 12 × 12 nm 1 M KCl box bisected by a bilayer composed of POPC lipids, for a total of 141 k atoms. Under the same conditions, the membrane-tethered assembly A was simulated in a 15 × 15 × 16 nm box for a total of 303 k atoms. Fixed atom constraints were placed on all atoms except the lipid tails when the temperature was increased to 301 K, followed by thermal equilibrium within a 0.5 ns dynamic range in the NvT ensemble. The fixed atom constraints have been replaced by harmonic constraints with an elastic constant of The simulation box size and pressure were equilibrated in the NpT ensemble for 3.5 ns, and the harmonic restraints were reduced every 0.5 ns. Unconstrained dynamics in the NvT ensemble brought the system to full equilibrium, and production simulations were performed in the NpT ensemble.

[0272] The production simulations were performed at 301 K and 1.013 bar pressure, maintained using a Langevin thermostat and the Nosé-Hoover Langevin piston method. The simulations were performed in NAMD, a smooth switching algorithm with a switching distance of The cut-off distance is The pairing list distance is for van der Waals interactions. A 2fs time step was used, and the SETTLE and SHAKE algorithms were used to constrain hydrogen bond lengths. Particle-mesh Ewald electrostatics were performed with The simulations were performed on a cubic grid with spaced and periodic boundary conditions. The equilibrium simulations were performed on a single GPU 1080Ti workstation and the production runs were parallelized on 850 CPU cores of the UCL GraceHPC facility.

[0273] 1.17. Simulation analysis

[0274] Production simulations were analyzed using GROMAC and VMD tools, and graphs were drawn using ggplot and RStudio after discarding the initial 10 ns.

[0275] 1.17.1.RMSF10

[0276] gmx_conver and gmx_aenig are used to investigate the top ten quasi-harmonic modes of the root mean square fluctuations (RMSF) of the heavy atoms of the DNA backbone, averaged per residue, to interrogate the structural dynamics of DNA nanostructures while accounting for thermal noise and random motions.

[0277] Clustering

[0278] gmx_cluster was used to prepare snapshots of the transmembrane A·B DNA nanostructure trajectory. Clustering was performed using the gromos method with a cutoff of 0.35 nm.

[0279] 1.17.3. Cavity analysis

[0280] The cluster coordinates were analyzed using HOLE with a channel end radius of 0.8 nm and a sampling distance of 0.25 nm. To account for the asymmetry of the DNA nanostructure, the coordinates were then rotated and analyzed again.

[0281] Lipid analysis

[0282] In initial equilibrium simulations, gmx_rms was used to measure the angles of phosphate and nitrogen atoms in lipid head groups that were split by the lipid leaflet compared to bilayer normal values. Production simulations were analyzed using gmx_rms, VMD plugins density_profile_too, and MEMBPLUGIM to determine lipid RMSF, mean lipid density, and area per lipid, respectively.

[0283] 1.18. Nanopore current recording

[0284] Single-channel current measurements were achieved using an integrated chip-based parallel bilayer recording device (Orbit Mini, Nanion Technologies, Munich, Germany) and a multi-electrode chamber array (MECA) chip (IONERA, Freiburg, Germany). The bilayer was formed from DPhPC (10 mg / mL in octane). The electrolyte solution was 1 M KCl and 10 mM HEPES, pH 7.4. To achieve pore insertion, a 2:1 mixture of nanopore A·B and 0.5% OPOE (n-octyl oligoethylene oxide in 1 M KCl, 10 mM HEPES, pH 7.4) was added to the cis side of the bilayer. Successful insertion was observed by detecting the current step. For triggered assembly, the membrane was attached to AL A and BL Band 0.5% (v / v) OPOE preincubation. After 10 min, the keychain was added and successful insertion was observed by monitoring the current step. The current traces were not Bessel filtered and were acquired at 10 kHz using Element Data Recorder software (Elements.rl, Cesena, Italy). Single channel analysis was performed using Clampfit software (Molecular Devices, Sunnyvale, CA, USA).

[0285] Preparation of SUVs loaded with fluorophores and dye release assays

[0286] A chloroform solution of DPhPC lipid (100 μL, 10 mM) was added to a 5 mL round-bottom flask. The solvent was removed using a rotary evaporator (Buchi, Newmarket, UK) to obtain a thin film, which was further dried under high vacuum (Buchi) for 1 hour. The lipid was resuspended in buffer A containing the fluorophore sulforhodamine B (SRB, 50 mM), sonicated at 30 ° C for 20 minutes, and then balanced at 4 ° C for 3 hours. The SUV was then extruded 25 times through a 100 nm polycarbonate membrane (Avanti Polar Lipids, US) using an extruder kit (Avanti Polar Lipids, US). Unencapsulated SRB was removed using a NAP-25 column (Cytivia, UK), and the SUV was exchanged into buffer D (0.2 M KCl, 10 mM Tris pH 7.4). The purified SUV was used within 48 hours and gently resuspended immediately before use.

[0287] For release assays, A·B was folded at 1 μM in buffer C using a 15-hour folding protocol, while components A and B were folded at 2 μM. SUV suspensions with encapsulated SRB (10 μL) and buffer D (110 μL, 80 μL) were added to 10 mm quartz cuvettes (HellmaAnalytics, Southend on Sea, UK). Fluorescence was monitored at 586 nm using a Varian Cary Eclipse fluorescence spectrophotometer (Agilent, Cheadle, UK) with excitation at 565 nm. After 5 minutes, A, A+B, A·B, AL A +BL B or AL A +BL B +K A +K B(30 μL or 60 μL; 1 μM in buffer C) was added to a final volume of 150 μL. After 55 minutes of fluorescence monitoring, the samples were mixed with 1% (v / v) Triton X-100 (10 μL) solution to lyse all vesicles to determine the maximum release of SRB. The release extent (%) was calculated using the maximum fluorescence emission and the fluorescence before the addition of A·B (or component).

[0288] 1.20. Preparation of Fura-2-filled SUVs and Ca 2+ Influx assay

[0289] A chloroform solution of lipid POPC (100 μL, 10 mM) was added to a 5 mL round-bottom flask. The solvent was removed using a rotary evaporator (Buchi, Newmarket, UK) to obtain a film, which was further dried for 1 hour under high vacuum (Buchi). The lipid was resuspended in buffer E (500 mM NaCl, 100 mM HEPES, pH 7.4) containing the fluorophore Fura-2 (100 μM). The solution was sonicated at 30 ° C for 20 minutes and then balanced for 3 hours. SUV was extruded 25 times through a 100 nm polycarbonate membrane (Avanti Polar Lipids, USA) using an extruder kit (Avanti Polar Lipids, USA). Unencapsulated dyes were removed using an Illustra MicroSpin S-400 spin column (Cytivia, UK). Dynamic light scattering of SUV was then performed with Malvern Zetasizer Nano S (Malvern Pananalytical, Malvern, UK) to confirm the diameter of the vesicle. Purified SUVs should be used within 48 hours and gently resuspended immediately before use. 2+ For influx assay, SUV suspensions containing encapsulated Fura-2 (30 μL) and buffer E (138.3 μL, 145.8 μL, 148.33 μL) were added to 10 mm quartz cuvettes (HellmaAnalytics, Southend on Sea, UK). Fluorescence was monitored at 510 nm using a Varian Cary Eclipse fluorescence spectrophotometer (Agilent, UK) with excitation at 340 and 380 nm. After 2.5 min, CaCl2 (16.7 μL, 3 mM in H2O) was added and allowed to stabilize for another 2.5 min. At 5 min, (A·B) was added. 4C(15 μL, 7.5 μL, 5 μL, 1 μM in buffer B) to a final volume of 200 μL. After monitoring fluorescence for 30 min, the samples were mixed with 1% Triton X-100 (10 μL) solution (v / v) to lyse all vesicles to determine the maximum Ca 2+ internal flow. Ca 2+ Influx was monitored as the ratio of emission changes at each excitation wavelength, i.e. 340 / 380 nm. The maximum 340 / 380 nm ratio after addition of Triton X-100 was used to normalize all traces.

[0290] Table 6. Names, modifications and sequences of DNA oligonucleotides used for folding nanostructures A, B, A·B and variants

[0291]

[0292]

[0293] Cy3X = Cy3 fluorophore; Cy5X = Cy5 fluorophore; Alexa488X = Alexa488 fluorophore; Alexa647X = Alexa647 fluorophore

[0294] Table 7. Names and strand compositions of structures for exemplary DNA nanopores with biomimetic triggered assembly

[0295]

[0296]

[0297] Table 8. For folding R, S and S NP The names, modifications and sequences of the DNA oligonucleotides.

[0298]

[0299]

[0300]

[0301] Table 9. Names and chain compositions of structures used in model systems

[0302] composition NP 1，2，3，4，5，6，7，8 <![CDATA[S NP ]]> 1, 2, 3, 4, 5-ext, 6, 7, 8 <![CDATA[ Cy5 S NP ]]> <![CDATA[1,2,3,4, Cy5 5-ext,6,7,8]]> <![CDATA[R ΔC ·S]]> <![CDATA[R ΔC ,S]]> R.S. R, S <![CDATA[R ΔC ·S NP ]]> <![CDATA[R ΔC ,1,2,3,4,5-ext,6,7,8]]> <![CDATA[R·S NP ]]> R,1,2,3,4,5-ext,6,7,8 <![CDATA[ Cy3 R ΔC · Cy5 S]]> <![CDATA[ Cy3 R ΔC , Cy5 S]]> <![CDATA[ Cy3 R· Cy5 S]]> <![CDATA[ Cy3 R, Cy5 S]]> <![CDATA[ Cy3 R ΔC · Cy5 S NP ]]> <![CDATA[ Cy3 R ΔC ,1,2,3,4, Cy5 5-ext,6,7,8]]> <![CDATA[ Cy3 R· Cy5 S NP ]]> <![CDATA[ Cy3 R,1,2,3,4, Cy5 5-ext,6,7,8]]>

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[0426] Although specific embodiments of the present invention are disclosed in detail herein, this is only by way of example and is for illustrative purposes only. The above examples are not intended to limit the scope of the appended claims. For example, for those skilled in the art who are familiar with the embodiments currently described, the selection of nucleic acid starting materials is considered to be a routine matter. The inventors anticipate that various substitutions, changes and modifications may be made to the present invention without departing from the spirit and scope of the present invention as defined in the claims.

Claims

1. A nucleic acid nanostructure, comprising: a plurality of component modules, each component module comprising a nucleic acid sequence and at least one membrane anchor, and wherein the plurality of component modules are capable of controlled assembly in response to external stimuli to form the nanostructure; and The nanostructures are configured to penetrate the semifluid membrane upon or after controlled assembly.

2. The nucleic acid nanostructure of claim 1, wherein the plurality of component modules are capable of being combined with a semifluid membrane prior to controlled assembly.

3. The nucleic acid nanostructure according to claim 2, wherein the plurality of component modules can be combined with a semifluid membrane after controlled assembly.

4. A nucleic acid nanostructure according to any preceding claim, wherein the external stimulus comprises a molecule binding to at least one of the plurality of component modules.

5. The nucleic acid nanostructure of claim 4, wherein the external stimulus comprises one or more molecules selected from the group consisting of: nucleic acid sequences; aptamers; small molecules; antibodies or antibody fragments; antibody mimetics; peptides; polypeptides; polysaccharides; oligosaccharides; macromolecules in the following size ranges: 1-10 kD, 1-50 kD, 1-100 kD, 10-50 kD, 10-100 KD, 20-50 kD, and 20-100 kD; carbohydrates; biopolymers; toxins; metabolites; and / or cytokines.

6. The nucleic acid nanostructure according to claim 4 or 5, wherein the external stimulus comprises a molecule selected from the group consisting of: I. Enzymes - including polymerases, helicases, gyrases and telomerases, as well as nucleic acid binding subdomains or their derivatives; II. Synthetic or naturally derived affinity binding proteins and peptides, including affibodies, antigen binding microproteins, engineered multi-repeat proteins, ankyrin binding domains, lactoferrin, cathepsins, ficolins, collectins, T cell receptor domains, and defensins; III. Antibodies - including polyclonal, monoclonal, humanized and camelid antibodies, or antigen-binding fragments and derivatives thereof, including Fab, scFv, Bis-scFv, VH, VL, V-NAR, VhH or any other antigen-binding single domain antibody fragment; IV. Synthetic or naturally derived affinity binding nucleic acids and nucleic acid analogs, including oligonucleotide probes, aptamers, and ribozymes; V. Natural or synthetic small molecules, including drugs, fluorophores, metabolites, and chemokines; VI. antigens or antigen fragments; and VII. signaling molecules and / or polypeptide receptors thereof, including receptor binding domains and receptor complexes.

7. The nucleic acid nanostructure according to any one of claims 2 to 6, wherein the semifluid membrane comprises a synthetic amphiphilic membrane or a lipid bilayer membrane.

8. A nucleic acid nanostructure according to claim 7, wherein the nanostructure is a nanopore and optionally the nanopore comprises a central cavity having a minimum internal width of at least 0.2 nm, suitably at least 0.5 nm, optionally at least 0.75 nm.

9. The nucleic acid nanostructure of claim 7, wherein the nanostructure is a nanopore and optionally the nanopore comprises a central cavity having a maximum internal width of at most about 20 nm, suitably at most about 10 nm, optionally at most about 5 nm.

10. The nucleic acid nanostructure according to any one of the preceding claims, wherein the nucleic acid nanostructure consists of DNA.

11. The nucleic acid nanostructure of claim 10, wherein each of the plurality of component modules comprises a DNA sequence, at least a portion of which comprises a double helix defining a secondary structure, and at least a portion of which comprises a single-stranded sequence defining an assembly interface.

12. The nucleic acid nanostructure of claim 11, wherein the assembly interface of each of the plurality of component modules comprises a sequence that is complementary to and capable of hybridizing with the assembly interface of another component module.

13. A nucleic acid nanostructure according to any one of the preceding claims, wherein the membrane anchor comprises a hydrophobic molecule, suitably wherein the hydrophobic molecule comprises a cholesterol molecule or an analog thereof; a sterol; an alkylated phenol; a flavonoid; a saturated or unsaturated fatty acid; a synthetic lipid molecule; or dodecyl-β-D-glucoside.

14. A nucleic acid nanostructure according to any preceding claim, wherein the nanostructure consists of at least two component modules.

15. A nucleic acid nanostructure according to any preceding claim, wherein the nanostructure is composed of at least three component modules.

16. A nucleic acid nanostructure according to any preceding claim, wherein the assembly of the nanostructure and / or its components is performed by DNA origami technology.

17. A nucleic acid nanostructure comprising: a plurality of building blocks, each building block comprising a deoxyribonucleic acid (DNA) sequence, wherein the DNA sequence comprises at least a portion of a double helix defining a secondary structure and at least a portion of a single-stranded sequence defining an assembly interface, and at least one hydrophobic anchor; as well as a plurality of single-stranded nucleic acid lock sequences capable of hybridizing to the single-stranded sequences at the assembly interface; wherein the component modules are capable of associating and interacting with a surface of a semifluid membrane via the anchors, and The modules are capable of controlled assembly in response to an externally applied stimulus, which initiates the dissociation of the plurality of lock sequences to reveal an assembly interface, thereby associating the plurality of component modules to form a nanostructure.

18. The nucleic acid nanostructure of claim 17, wherein the nanostructure is configured to penetrate the semifluid membrane during or after the controlled assembly.

19. A nucleic acid nanostructure according to claims 17 and 18, wherein the externally applied stimulus comprises a molecule selected from one or more of the following: a nucleic acid sequence; an aptamer; a small molecule; an antibody or antibody fragment; an antibody mimetic; a peptide; a polypeptide; a polysaccharide; an oligosaccharide; a macromolecule in the following size range: 1-10 kD, 1-50 kD, 1-100 kD, 10-50 kD, 10-100 KD, 20-50 kD and 20-100 kD; a carbohydrate; a biopolymer; a toxin; a metabolite; and / or a cytokine.

20. The nucleic acid nanostructure of claims 17 to 19, wherein the externally applied stimulus comprises a molecule selected from the group consisting of: I. Enzymes - including polymerases, helicases, gyrases and telomerases, as well as nucleic acid binding subdomains or their derivatives; II. Synthetic or naturally derived affinity binding proteins and peptides, including affibodies, antigen binding microproteins, engineered multi-repeat proteins, ankyrin binding domains, lactoferrin, cathepsins, ficolins, collectins, T cell receptor domains, and defensins; III. Antibodies - including polyclonal, monoclonal, humanized and camelid antibodies, or antigen-binding fragments and derivatives thereof, including Fab, scFv, Bis-scFv, VH, VL, V-NAR, VhH or any other antigen-binding single domain antibody fragment; IV. Synthetic or naturally derived affinity binding nucleic acids and nucleic acid analogs, including oligonucleotide probes, aptamers, and ribozymes; V. Natural or synthetic small molecules, including drugs, fluorophores, metabolites, and chemokines; VI. antigens or antigen fragments; and VII. signaling molecules and / or polypeptide receptors thereof, including receptor binding domains and receptor complexes.

21. The nucleic acid nanostructure according to any one of claims 17 to 20, wherein the semifluid membrane comprises a synthetic amphiphilic membrane or a lipid bilayer membrane.

22. The nucleic acid nanostructure according to any one of claims 17 to 21, wherein the nanostructure formed after the controlled assembly is a transmembrane nanopore.

23. A nucleic acid nanostructure according to claim 22, wherein the nanopore comprises a central cavity having a minimum internal width of at least 0.2 nm, suitably at least 0.5 nm, optionally at least 0.75 nm.

24. The nucleic acid nanostructure of claim 22, wherein the nanopore comprises a central cavity having a maximum internal width of at most about 20 nm, suitably at most about 10 nm, optionally at most about 5 nm.

25. The nucleic acid nanostructure of any one of claims 17 to 24, wherein the assembly interface of each of the plurality of building blocks comprises a sequence that is complementary to and capable of hybridizing with the assembly interface of another of the building blocks.

26. The nucleic acid nanostructure according to any one of claims 17 to 25, wherein the hydrophobicity comprises: Sterols; alkylated phenols; flavonoids; saturated or unsaturated fatty acids; or synthetic lipid molecules.

27. The nucleic acid nanostructure of claim 26, wherein: - sterol selected from cholesterol; cholesterol derivatives; phytosterols; ergosterol; bile acids; - alkylated phenols are selected from methylated phenols; dolicholine and tocopherol; - flavonoids are selected from: flavanone-containing compounds; 6-hydroxyflavone; - saturated and unsaturated fatty acids selected from lauric acid derivatives; oleic acid; linoleic acid; palmitic acid; and / or -The synthetic lipid molecule is dodecyl-β-D-glucoside.

28. A semifluid film having bonded thereto the nanostructure of any one of claims 1 to 27.

29. A semifluid membrane having a plurality of nanostructured component modules bonded thereto, each component module comprising a deoxyribonucleic acid (DNA) sequence, wherein the DNA sequence comprises at least a portion of a double helix defining a secondary structure and at least a portion of a single-stranded sequence defining an assembly interface, and at least one hydrophobic anchor; and a plurality of single-stranded nucleic acid lock sequences capable of hybridizing with the single-stranded sequence of the assembly interface.

30. The semifluid membrane of claim 29, wherein the semifluid membrane comprises a synthetic amphiphilic membrane or a lipid bilayer membrane.

31. A semifluid membrane according to claim 30, wherein the membrane is formed from a polymer.

32. A semifluid membrane according to claim 31, wherein the polymer forming the semifluid membrane consists of an amphiphilic synthetic block copolymer suitably selected from a hydrophilic copolymer block and a hydrophobic copolymer block.

33. A sensor device, wherein the sensor comprises the nucleic acid nanostructure of any one of claims 1 to 27.

34. The sensor device of claim 33, wherein the nucleic acid nanostructure is a transmembrane nanopore.

35. The sensor device of claim 34, wherein the transmembrane nanopore defines an ion channel.

36. The sensor device of any one of claims 33 to 35, wherein the nucleic acid nanostructure is contained within a flow cell.

37. The sensor device according to any one of claims 33 to 36, further comprising an electrical measuring device and / or a fluorescence measuring device.

38. A sensor device, wherein the sensor device comprises the semifluid membrane of any one of claims 29 to 32.

39. The sensor device of claim 38, wherein the semifluid membrane is contained within a flow cell.

40. The sensor device according to any one of claims 38 or 39, further comprising an electrical measuring device and / or a fluorescence measuring device.

41. The sensor device of claim 40, wherein the electrical measurement device measures a parameter selected from the group consisting of: current measurement; impedance measurement; tunneling measurement; or field effect transistor (FET) measurement.

42. A semifluid membrane vesicle, wherein the vesicle comprises the nucleic acid nanostructure of any one of claims 1 to 27.

43. The semifluid membrane vesicle of claim 42, wherein the vesicle further comprises a pharmaceutical agent.

44. The semifluid membrane vesicle of claim 42 or 43, wherein the membrane vesicle comprises a synthetic amphiphilic membrane or a lipid bilayer membrane.

45. A pharmaceutical composition comprising the semifluid membrane vesicle of any one of claims 42 to 44 and a pharmaceutically acceptable carrier.

46. ​​A method of assembling a nucleic acid nanostructure, the method comprising: Provides multiple component modules, each of which includes - a nucleic acid sequence, wherein the nucleic acid sequence comprises at least a portion of a double helix defining a secondary structure and at least a portion of a single-stranded sequence defining an assembly interface, wherein the assembly interface of each of the plurality of building blocks comprises a sequence that is complementary to and capable of hybridizing with the assembly interface of another building block; and - at least one hydrophobic anchor; and combining multiple component modules under conditions that allow the spontaneous assembly of nanostructures.

47. A method for triggering assembly of a nucleic acid nanostructure, the method comprising: providing a plurality of component modules, each component module comprising a nucleic acid sequence, wherein the nucleic acid sequence comprises at least a portion of a double helix defining a secondary structure and at least a portion of a single-stranded sequence defining an assembly interface, wherein the assembly interface of each of the plurality of component modules comprises a sequence that is complementary to and capable of hybridizing with the assembly interface of another component module, and at least one hydrophobic anchor, Providing a plurality of single-stranded nucleic acid lock sequences that hybridize with the single-stranded sequences of the assembly interface, thereby forming a plurality of locked component modules that are inhibited from initiating spontaneous assembly into nucleic acid nanostructures; exposing the plurality of locking component modules to an external stimulus, wherein the external stimulus triggers the plurality of single-stranded nucleic acid lock sequences to dissociate from the plurality of locking component modules, thereby exposing the plurality of assembly interfaces; and Providing conditions that allow the spontaneous assembly of nanostructures.

48. The method of claim 47, wherein: The plurality of sub-assembly modules are introduced into the semi-fluid membrane prior to exposure to the external stimulus under conditions that allow the plurality of locking assembly modules to be bonded to the semi-fluid membrane via the at least one membrane anchor.

49. The method of claim 47 or 48, wherein the external stimulus comprises a molecule selected from one or more of the following: a nucleic acid sequence; an aptamer; a small molecule; an antibody or antibody fragment; an antibody mimetic; a peptide; a polypeptide; a polysaccharide; and an oligosaccharide.

50. The method of claim 47 or 48, wherein the externally applied stimulus comprises a molecule selected from the group consisting of: I. Enzymes - including polymerases, helicases, gyrases and telomerases, as well as nucleic acid binding subdomains or their derivatives; II. Synthetic or naturally derived affinity binding proteins and peptides, including affibodies, antigen binding microproteins, engineered multi-repeat proteins, ankyrin binding domains, lactoferrin, cathepsins, ficolins, collectins, T cell receptor domains, and defensins; III. Antibodies - including polyclonal, monoclonal, humanized and camelid antibodies, or antigen-binding fragments and derivatives thereof, including Fab, scFv, Bis-scFv, VH, VL, V-NAR, VhH or any other antigen-binding single domain antibody fragment; IV. Synthetic or naturally derived affinity binding nucleic acids and nucleic acid analogs, including oligonucleotide probes, aptamers, and ribozymes; V. Natural or synthetic small molecules, including drugs, fluorophores, metabolites, and chemokines; VI. antigens or antigen fragments; and VII. signaling molecules and / or polypeptide receptors thereof, including receptor binding domains and receptor complexes.

51. A method according to any one of claims 47 to 50, wherein when assembled, the nanostructures define membrane-penetrating nanostructures.

52. The method of claim 51, wherein the membrane-penetrating nanostructures define transmembrane nanopores.

53. A method according to claim 52, wherein the transmembrane nanopore comprises a central cavity having a minimum internal width of at least 0.2 nm, suitably at least 0.5 nm, optionally at least 0.75 nm.

54. A method according to claim 52, wherein the nanopore comprises a central cavity having a maximum internal width of at most about 20 nm, suitably at most about 10 nm, optionally at most about 5 nm.

55. The method of any one of claims 47 to 54, wherein the membrane-penetrating nanostructures define an ion channel.

56. A method according to any one of claims 47 to 55, wherein the membrane-penetrating nanostructures define channels that allow fluid communication through the semifluid membrane.

57. The method of claim 56, wherein the membrane-penetrating nanostructures serve as a release mechanism for a drug substance.

58. The method of claim 56, wherein the membrane-penetrating nanostructures serve as a release mechanism for imaging substances.

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