Plasmon nanoantennas and their use in single-molecule sequencing

Abstract

This disclosure relates, in general, to a method for sequencing nucleic acid molecules, such as DNA, and more particularly, a single molecule of DNA, using a polymerase enzyme positioned within an electric field-enhancing area. This disclosure also relates to nucleic acid sequencing, a method for producing the nanoantennas, an array containing the nanoantennas, and a method for using the nanoantennas and array in a single-molecule sequencing technique. In one embodiment, the fluorescence signature of a labeled nucleotide is not enhanced outside the electric field-enhancing area.

Description

[Technical Field]

[0001] Related application data This application claims priority to Australian Provisional Patent Application No. 2023901389, filed on 8 May 2023, which is incorporated herein by reference in its entirety.

[0002] Technical field This disclosure generally relates to methods for sequencing nucleic acid molecules, plasmon nanoantennas, methods for producing said nanoantennas, arrays containing said nanoantennas, and methods for using said nanoantennas and arrays in single-molecule sequencing techniques. [Background technology]

[0003] background DNA sequencing is the process of reading the sequence in which the four nucleotide bases—adenine (A), thymine (T), cytosine (C), and guanine (G)—are arranged in the long biopolymer that codes for the genes of all living organisms. The ability to sequence DNA has revolutionized biological science and is rapidly expanding its impact on various industries, including pharmaceuticals, agriculture, and biotechnology. The DNA sequencing market is projected to reach $60 billion by 2030. Nevertheless, the current wave of industry-leading high-throughput sequencing methods faces significant challenges, including the need for large amounts of DNA, the limited maximum length of DNA that can be read sequentially, and the computational resources required to assemble sequences from raw data. Therefore, a highly desirable goal is to accurately read the sequence of a single DNA molecule in real time. [Overview of the project] [Problems that the invention aims to solve]

[0004] overview In recent years, single-molecule sequencing technology has advanced significantly, with two new technologies currently leading the market: Oxford Nanopore's "current-blocking" sequencing and PacBio's "zero-mode waveguide" sequencing. Each company is already valued at approximately US$2 billion. However, both technologies use methods that have fundamental limitations in their ability to precisely manipulate a single DNA molecule and to limit the observation volume to a single nucleotide. These limitations result in a high error rate, and therefore, accurate sequencing still requires multiple sequencing reactions to achieve statistically accurate results. Thus, true single-molecule DNA sequencing is needed. [Means for solving the problem]

[0005] The use of metal nanoparticles and plasmons can function as auxiliary antennas to enhance molecular fluorescence, for example, in the field of optical sensors. By coupling electromagnetically irradiated molecules with nanoparticle plasmons, it is possible to influence the intensity and dynamics of the interaction between molecules and light, thereby increasing the emission rate of fluorescent molecules. Therefore, detection based on nanoparticle plasmon resonance is a label-free technique developed to detect the presence of analytes in samples and / or to investigate biomolecular interactions in real time.

[0006] Plasmon resonance-based detection methods and compositions for use in improved DNA molecular sequencing are presented herein. The plasmon resonance-based detection methods and compositions presented herein also encompass solutions to challenges in the development of plasmon resonance-based detection methods for sequencing, where such challenges have not been previously considered or known. While previous attempts have been made to apply plasmon resonance-based detection to complex biomolecules such as proteins at the single-molecule level (e.g., US20130252825A1), challenges remain due to the difficulty in immobilizing large biomolecules, such as enzymes, within hotspots formed by nanoparticles in a manner that preserves their function (e.g., polymerase enzymes). Therefore, there is a need for new methods and tools for analyzing single molecules using nanoparticle plasmon resonance-based techniques. This disclosure is based in part on the inventors' recognition that, despite the development of plasmon nanoantennas and their usefulness in various applications, previous efforts to use plasmon nanoantennas in biological applications have been limited by the types of molecules that can be tethered to the DNA scaffold within the “plasmon hotspot” that defines the electric field enhancement area. The limitations of existing plasmon nanoantenna technologies stem in part from the fact that the relatively small size of the plasmon hotspot required for meaningful signal enhancement limits the size of molecules that can be bound within the hotspot, as well as the size of analytes that can access the hotspot during use.Very recent attempts have resulted in plasmon nanoantennas with so-called "cleared space" within plasmon hotspots. However, there is currently no evidence that enzymes bind to the plasmon hotspots of plasmon nanoantennas, much less that enzymes bound within the hotspot can maintain their activity under the steric constraints caused by the limited space within the DNA origami structure. However, we have now developed a plasmon nanoantenna containing a nanoscale DNA scaffold to which two plasmon nanoparticles are immobilized, forming a plasmon hotspot between them. We have demonstrated for the first time that an active protein (i.e., DNA polymerase) can be immobilized within the unoccupied area (or "cleared space") of the plasmon hotspot. We have also experimentally demonstrated that DNA polymerase retains its biological function when immobilized within the plasmon hotspot of the nanoantenna design, and that the plasmon hotspot, while containing DNA polymerase, maintains a sufficient electric field intensity to enhance the fluorescence signal. In this regard, the inventors have shown that the intensity of light emitted from fluorophores attached to or positioned with protein molecules within a hotspot can be enhanced compared to the intensity of light emitted from corresponding fluorophores outside the hotspot. However, most importantly, the inventors have experimentally found that when DNA polymerase is tethered within the plasmon hotspot of these nanoantennas in the presence of fluorescently labeled deoxynucleotide triphosphates (dNTPs) and a DNA template, the order in which specific fluorescently labeled dNTPs are incorporated into the growing DNA strand by the DNA polymerase enzyme during DNA synthesis can be detected within the plasmon hotspot by detecting the enhanced, unique fluorescence signals emitted from each dNTP when they are incorporated. Therefore, the inventors have shown that the plasmon nanoantennas of this disclosure can be used for single-molecule DNA sequencing applications.

[0007] Therefore, in one embodiment, the present disclosure is a method for sequencing nucleic acid analytes, (I) A step of contacting nucleic acid polymerase with the nucleic acid analyte and the labeled nucleotide for a period of time and under conditions such that the labeled nucleotide is sequentially incorporated by the polymerase into a polynucleotide having a sequence complementary to the polynucleotide sequence of the nucleic acid analyte, wherein the polymerase is located within an electric field enhancement area and each labeled nucleotide is (i) adenine nucleotide (A), guanosine nucleotide (G), thymine nucleotide (T), or cytosine nucleotide (C), (ii) Fluorophores, and (iii) Polyphosphate linkers that bind nucleotides to fluorophores Includes, A, G, T, and C are each independently linked to the fluorophore via a polyphosphate. Each of the labeled nucleotides A, G, T, and C has a distinct fluorescence signature when the fluorophore to which the respective nucleotide is linked is excited. The distinct fluorescent signature of the labeled nucleotide is enhanced when the labeled nucleotide is incorporated into a sequence complementary to the polynucleotide sequence of the nucleic acid analyte by nucleic acid polymerase. Steps and (II) A step of detecting the sequence of enhanced, distinct fluorescence signatures when labeled nucleotides are sequentially incorporated by nucleic acid polymerase into polynucleotides having a sequence complementary to the polynucleotide sequence of the nucleic acid analyte, thereby determining the sequence of the nucleic acid analyte by determining the sequence of nucleotides incorporated into polynucleotides having a sequence complementary to the polynucleotide sequence of the nucleic acid analyte, and This provides a method that includes [something].

[0008] g

[0009] In one embodiment, the method includes contacting a nucleic acid polymerase with a nucleic acid analyte in the presence of each of labeled nucleotides A, G, T, and C.

[0010] In some embodiments, the nucleotide of the labeled nucleotide can further be selected from synthetic nucleotides. In one embodiment, the synthetic nucleotide is selected from 5-methylcytidine, N-methylcytidine, and N6-methyladenosine.

[0011] The synthetic nucleotide is preferably linked to a fluorophore via a polyphosphate. According to an example where the method uses a synthetic nucleotide linked to a fluorophore via a polyphosphate, each of nucleotides A, G, T, and C and the synthetic nucleotide has a distinct fluorescence emission signature when the fluorophore to which each nucleotide is linked is excited.

[0012] In one embodiment, the distinct enhanced fluorescence signature of the labeled nucleotide is enhanced by a factor of 2 or more (e.g., about 3 or more, or about 4 or more, or about 5 or more, or about 6 or more, or about 7 or more, or about 8 or more, or about 9 or more) compared to the distinct fluorescence signature of the corresponding labeled nucleotide outside the electric field enhancement area. In some embodiments, the enhanced, distinct fluorescence signature of the labeled nucleotide is enhanced by an order of magnitude or more (e.g., by about 10 times or more, or about 20 times or more, or about 30 times or more, or about 40 times or more, or about 50 times or more, or about 60 times or more, or about 70 times or more, or about 80 times or more, or about 90 times or more, or about 100 times or more) compared to the distinct fluorescence signature of the corresponding labeled nucleotide outside the electric field enhancement area. In further embodiments, the enhanced separate fluorescence signature of the labeled nucleotide is enhanced by at least about 100 times or more (e.g., at least about 150 times, or at least about 200 times, or at least about 250 times, or at least about 300 times, or at least about 350 times, or at least about 400 times, or at least about 450 times, or at least about 500 times, or at least about 550 times, or at least about 600 times, or at least about 650 times, or at least about 700 times, or at least about 750 times, or at least about 800 times, or at least about 850 times, or at least about 900 times, or at least about 950 times, or at least about 1000 times) compared to the separate fluorescence signature of the corresponding labeled nucleotide outside the electric field enhancement area.

[0013] The distinct fluorescence signatures described herein may be distinguishable from one another based on differences in peak emission wavelength, fluorescence emission intensity, fluorescence emission duration, fluorescence lifetime, duration between sequential fluorescence emissions, or any combination thereof.

[0014] In one embodiment, each fluorophore has a peak emission wavelength independently selected from emission wavelengths in the visible spectrum, ultraviolet (UV) spectrum, infrared (IR) spectrum, and near-infrared spectrum. In another embodiment, each fluorophore has a peak emission wavelength independently selected from emission wavelengths between approximately 350 nm and 850 nm. In some embodiments, one or more or each of the fluorophores have low quantum efficiency (or low quantum yield).

[0015] In one embodiment, the distinct fluorescence signatures of two or more species of labeled nucleotides are distinguishable based on differences in their peak emission wavelengths. In one embodiment, each different species of nucleotide is linked to a fluorophore having a distinct peak emission wavelength. According to any example in which the labeled nucleotides are distinguishable based on differences in their peak emission wavelengths, the peak emission wavelengths of the labeled nucleotides are separated from each other by 10 nm or more. For example, the peak emission wavelengths may be separated from each other by about 25 nm or more. For example, the peak emission wavelengths may be separated from each other by about 50 nm or more. For example, the peak emission wavelengths may be separated from each other by about 75 nm or more. For example, the peak emission wavelengths may be separated from each other by about 100 nm or more.

[0016] In one embodiment, distinct fluorescent signatures of two or more species of labeled nucleotides can be distinguished based on differences in fluorescence lifetime.

[0017] Alternatively, or in addition to that, nucleotides of two or more species are linked to the same fluorophore, and nucleotides of different species are distinguished based on differences in fluorescence intensity and / or fluorescence duration. In one embodiment, nucleotides of different species are distinguished based on differences in fluorescence intensity. In one embodiment, nucleotides of different species are distinguished based on differences in fluorescence duration.

[0018] As described herein, each labeled nucleotide used in the methods of this disclosure comprises a nucleotide linked to a fluorophore via a polyphosphate. In one embodiment, the polyphosphate is a triphosphate, tetraphosphate, pentaphosphate, or hexaphosphate. For example, the polyphosphate is a triphosphate. For example, the polyphosphate is a tetraphosphate. For example, the polyphosphate is a pentaphosphate. For example, the polyphosphate is a hexaphosphate.

[0019] In some embodiments, the fluorophore is bound to the phosphate furthest from the nucleotide.

[0020] In some embodiments, the method includes the step of contacting a nucleic acid polymerase with a nucleic acid analyte and a labeled nucleotide in the presence of one or more quenching agents.

[0021] In one embodiment, the method includes the step of providing a source of electromagnetic waves to an electric field enhancement area, wherein the electromagnetic waves have a wavelength sufficient to excite a fluorophore linked to a labeled nucleotide.

[0022] In each of the above examples, the step of detecting the sequence of the enhanced distinct fluorescence signatures in (II) is performed using a fluorescence microscope or fluorophotometer. For example, the step of detecting the sequence of the enhanced distinct fluorescence signatures in (II) is performed using a total internal reflection fluorescence (TIRF) microscope. For example, the step of detecting the sequence of the enhanced distinct fluorescence signatures in (II) is performed using a confocal microscope. For example, the step of detecting the sequence of the enhanced distinct fluorescence signatures in (II) is performed using an epifluorescence microscope.

[0023] In each of the examples described above, the nucleic acid analyte may be DNA. In one example, the DNA is genomic DNA. In another example, the DNA is complementary DNA (cDNA) obtained from RNA. Following the example where the nucleic acid analyte is cDNA, the method may include the step of determining the RNA sequence based on the corresponding cDNA sequence.

[0024] In some embodiments, the method further includes the step of contacting DNA with an oligonucleotide primer capable of specifically hybridizing with a region of DNA to initiate synthesis by a nucleic acid polymerase, such as DNA polymerase. In some embodiments, the region of DNA that the oligonucleotide primer hybridizes is an oligonucleotide adapter, and the oligonucleotide primer is substantially complementary to the oligonucleotide adapter. In certain examples, the DNA to be sequenced contains an oligonucleotide adapter at its 3' end. For example, the DNA containing an oligonucleotide adapter at its 3' end may be single-stranded DNA. For example, the DNA containing an oligonucleotide adapter at its 3' end may be double-stranded DNA. In other examples, the DNA to be sequenced is double-stranded DNA flanked by two hairpin loops, the hairpin loops being formed of single-stranded DNA, and at least one of the hairpin loops containing an oligonucleotide adapter. The method may further include the step of ligating the oligonucleotide adapter to the DNA to be sequenced prior to the contacting step of (I).

[0025] In another example, the DNA to be sequenced is single-stranded DNA containing a hairpin oligonucleotide adapter at its 3' end, where the hairpin oligonucleotide adapter contains a sequence that is internally complementary and forms a hairpin containing a double-stranded region, which can initiate synthesis by a nucleic acid polymerase, such as DNA polymerase. The method may further include the step of ligating the hairpin oligonucleotide adapter to the DNA to be sequenced prior to the contacting step of (I).

[0026] In each of the examples described above, the nucleic acid analyte may be a single molecule. Therefore, the method may include the step of contacting a nucleic acid polymerase with a single nucleic acid molecule, such as a single molecule of DNA.

[0027] In each of the aforementioned examples, the electric field enhancement area can be generated by a plasmon hotspot, which is generated by a plasmon nanoantenna. The plasmon nanoantenna may contain at least two plasmon nanoparticles.

[0028] In one particular example, the plasmon nanoantenna used to generate an electric field enhancement area in the method is, Two plasmon nanoparticles, Nanoscale nucleic acid scaffolds and Nucleic acid polymerase and Including, here, (a) Two plasmon nanoparticles are attached to a nanoscale nucleic acid scaffold and are arranged such that a plasmon hotspot exists between the two plasmon nanoparticles relative to each other. (b) Plasmon hotspots include regions not occupied by nanoscale nucleic acid scaffolds, (c) Nucleic acid polymerase is bound to a nanoscale nucleic acid scaffold and is located in a region not occupied by the nanoscale nucleic acid scaffold. (d) If necessary, the nucleic acid analytes are bound to a nanoscale nucleic acid scaffold.

[0029] This disclosure is, Two plasmon nanoparticles, Nanoscale nucleic acid scaffolds and Nucleic acid polymerase and A plasmon nanoantenna including, (a) Two plasmon nanoparticles are bound to a nanoscale nucleic acid scaffold and are arranged such that a plasmon hotspot exists between the two plasmon nanoparticles relative to each other. (b) Plasmon hotspots include regions not occupied by nanoscale nucleic acid scaffolds, (c) Nucleic acid polymerase is bound to a nanoscale nucleic acid scaffold and is located within a region not occupied by the nanoscale nucleic acid scaffold. (d) If necessary, the nucleic acid analyte is bound to a nanoscale nucleic acid scaffold. We also offer plasmon nano-antennas.

[0030] In the methods and / or other examples of plasmon nanoantennas of the present disclosure, the plasmon nanoantenna may contain more than two plasmon nanoparticles, each bound to a nanoscale nucleic acid scaffold and arranged relative to one another such that plasmon hotspots exist between the plasmon nanoparticles. For example, the plasmon nanoantenna may contain three plasmon nanoparticles. For example, the plasmon nanoantenna may contain four plasmon nanoparticles. For example, the plasmon nanoantenna may contain five plasmon nanoparticles. For example, the plasmon nanoantenna may contain six plasmon nanoparticles. For example, the plasmon nanoantenna may contain seven plasmon nanoparticles. For example, the plasmon nanoantenna may contain eight plasmon nanoparticles. For example, the plasmon nanoantenna may contain nine plasmon nanoparticles. For example, the plasmon nanoantenna may contain ten or more plasmon nanoparticles.

[0031] According to the method and / or example of a plasmon nanoantenna comprising two plasmon nanoparticles, the region within the plasmon hotspot not occupied by the nanoscale nucleic acid scaffold is defined by a three-dimensional space having a length (L) between the two plasmon nanoparticles, a height (H) perpendicular to (L) at the midpoint (L / 2), and a width (W) perpendicular to L and H at L / 2, where, at L / 2, W is selected from about 20 nm to about 100 nm, H is selected from about 10 nm to about 100 nm, and L is measured at the shortest distance between the two plasmon nanoparticles. In some examples, L is selected from about 20 nm to about 100 nm. In other examples, L is selected from about 20 nm to about 50 nm.

[0032] In other examples, the volume of the region of a plasmon hotspot not occupied by the nanoscale nucleic acid scaffold is at least about 1 zL, for example, from about 1 zL to about 10 zL, or from about 5 zL to about 10 zL. In certain specific examples, the volume of the region of a plasmon hotspot not occupied by the nanoscale nucleic acid scaffold is greater than or equal to about 10 zL. For example, the volume of the region of a plasmon hotspot not occupied by the nanoscale nucleic acid scaffold is from about 10 zL to about 50 zL. For example, the volume of the region of a plasmon hotspot not occupied by the nanoscale nucleic acid scaffold is from about 20 zL to about 40 zL.

[0033] As in the examples above, the region of the plasmon hotspot not occupied by the nanoscale nucleic acid scaffold is large enough to accommodate the nucleic acid polymerase and to retain its biological activity. In some examples, the region not occupied by the nanoscale nucleic acid scaffold is less than or equal to 50% of the plasmon hotspot. However, in other examples, the region not occupied by the nanoscale nucleic acid scaffold is greater than 50% of the plasmon hotspot. In some examples, the region not occupied by the nanoscale nucleic acid scaffold is 60% or more of the plasmon hotspot. In certain specific examples, the plasmon hotspot is substantially not occupied by the nanoscale nucleic acid scaffold. For example, the region not occupied by the nanoscale nucleic acid scaffold may be 70% or more of the plasmon hotspot. In some examples, the region not occupied by the nanoscale nucleic acid scaffold is 75% or more of the plasmon hotspot. In some examples, the region not occupied by the nanoscale nucleic acid scaffold is 80% or more of the plasmon hotspot. In some examples, the region not occupied by the nanoscale nucleic acid scaffold accounts for 85% or more of the plasmon hotspot. In some examples, the region not occupied by the nanoscale nucleic acid scaffold accounts for 90% or more of the plasmon hotspot.

[0034] In each of the examples described above, the region not occupied by the nanoscale nucleic acid scaffold may have a different shape or form. In one embodiment, the region not occupied by the nanoscale nucleic acid scaffold may be amorphous. Alternatively, the region not occupied by the nanoscale nucleic acid scaffold may have a substantially defined shape. For example, the region not occupied by the nanoscale nucleic acid scaffold may be spherical, elliptical, cubic, or rectangular.

[0035] According to the methods and / or some examples of plasmon nanoantennas of this disclosure, the average interparticle distance between plasmon nanoparticles is about 10 nm to about 100 nm, where the interparticle distance between two particles is determined as the shortest distance between the surfaces of each particle. For example, the average interparticle distance between plasmon nanoparticles may be about 10 nm to 80 nm. For example, the average interparticle distance between plasmon nanoparticles may be about 10 nm to 50 nm. For example, the average interparticle distance between plasmon nanoparticles may be about 20 nm to 80 nm. For example, the average interparticle distance between plasmon nanoparticles may be about 20 nm to 50 nm. For example, the average interparticle distance between plasmon nanoparticles may be about 30 nm to 50 nm. For example, the average interparticle distance between plasmon nanoparticles may be about 20 nm to 40 nm. For example, the average interparticle distance between plasmon nanoparticles may be about 30 nm to 40 nm. In one particular example, the average interparticle distance between plasmon nanoparticles is about 30 nm.

[0036] In some embodiments of the methods and / or plasmon nanoantennas of the present disclosure, the plasmon nanoantenna comprises a nucleic acid analyte bound to a nanoscale nucleic acid scaffold. For example, the nucleic acid analyte may be non-covalently bonded to the nanoscale nucleic acid scaffold (e.g., by hydrogen bonds).

[0037] The plasmon nanoparticles of the plasmon nanoantennas described herein may be formed from any material having plasmon resonance properties. Suitable plasmon nanoparticles include, but are not limited to, metal nanoparticles, metal alloy nanoparticles, polymer nanoparticles and their derivatives and composites. In one embodiment, each of the plasmon nanoparticles is independently selected from metal nanoparticles. For example, each of two or more plasmon nanoparticles is a metal nanoparticle independently selected from the group consisting of nanoparticles formed from gold nanoparticles, silver nanoparticles, aluminum nanoparticles, copper nanoparticles, bismuth nanoparticles, nickel nanoparticles, palladium nanoparticles and platinum nanoparticles, or alloys of any one of these. In some embodiments, the metal nanoparticles are solid and formed entirely from a metallic substance. In other embodiments, the metal nanoparticles are formed entirely from a metallic substance but have a hollow core. In other embodiments, the metal nanoparticles include a core (e.g., glass, polymer or composite core) coated with a metallic substance so that a metallic surface is formed. In one embodiment, the plasmon nanoparticles are gold nanoparticles or gold alloy nanoparticles. In one embodiment, the plasmon nanoparticles are silver nanoparticles or silver alloy nanoparticles. In other examples, plasmon nanoparticles are formed from non-metallic materials or metal oxides that are doped to increase the total carrier load of the nanoparticles, thereby increasing their plasmon resonance.

[0038] The plasmon nanoparticles of the plasmon nanoantennas described herein may vary in shape and geometry. Therefore, the plasmon nanoparticles can be independently selected from nanospheres, nanorods, nanoprisms, nanocubes, nanoshells, nanotubes, or nanostars. The plasmon nanoparticles of the plasmon nanoantennas may be homogeneous or heterogeneous. In one embodiment, the plasmon nanoparticles are nanospheres.

[0039] The plasmon nanoparticles of the plasmon nanoantennas described herein may vary in size. In one embodiment, the maximum diameter of each plasmon nanoparticle is approximately 5 nm to approximately 500 nm. In another embodiment, the maximum diameter of each plasmon nanoparticle is approximately 10 nm to approximately 250 nm. In another embodiment, the maximum diameter of each plasmon nanoparticle is approximately 20 nm to approximately 200 nm. In another embodiment, the maximum diameter of each plasmon nanoparticle is approximately 50 nm to approximately 150 nm. In another embodiment, the maximum diameter of each plasmon nanoparticle is approximately 80 nm to approximately 120 nm. In one particular embodiment, the maximum diameter of each plasmon nanoparticle is approximately 80 nm. In yet another particular embodiment, the maximum diameter of each plasmon nanoparticle is approximately 100 nm. In yet another particular embodiment, the maximum diameter of each plasmon nanoparticle is approximately 120 nm.

[0040] In some cases, plasmon nanoparticles have the same maximum diameter. In other cases, the maximum diameter differs among plasmon nanoparticles in the same plasmon nanoantenna. According to examples where the maximum diameter differs among plasmon nanoparticles in the same plasmon nanoantenna, the average maximum diameter of plasmon nanoparticles can range from approximately 5 nm to approximately 500 nm. For example, the average maximum diameter of plasmon nanoparticles in a plasmon nanoantenna can range from approximately 10 nm to approximately 250 nm. For example, the average maximum diameter of plasmon nanoparticles in a plasmon nanoantenna can range from approximately 20 nm to approximately 200 nm. For example, the average maximum diameter of plasmon nanoparticles in a plasmon nanoantenna can range from approximately 50 nm to approximately 150 nm. For example, the average maximum diameter of plasmon nanoparticles in a plasmon nanoantenna can range from approximately 80 nm to approximately 120 nm. In one particular example, the average maximum diameter of plasmon nanoparticles in a plasmon nanoantenna can be approximately 80 nm. In one specific example, the average maximum diameter of plasmon nanoparticles in a plasmon nanoantenna could be approximately 100 nm. In another specific example, the average maximum diameter of plasmon nanoparticles in a plasmon nanoantenna could be approximately 120 nm.

[0041] In one particular example, the plasmon nanoantenna described herein comprises gold or gold-coated nanoparticles (e.g., nanospheres) having an average maximum diameter of about 100 nm.

[0042] In each of the aforementioned examples describing the use of the plasmon nanoantennas and / or methods of the present disclosure, each plasmon nanoparticle is coated by a coating. Preferably, the coating substantially covers the surface of each plasmon nanoparticle. In one embodiment, the coating covers about 75% or more of the surface of each plasmon nanoparticle. In one embodiment, the coating covers about 80% or more of the surface of each plasmon nanoparticle. In one embodiment, the coating covers about 85% or more of the surface of each plasmon nanoparticle. In one embodiment, the coating covers about 90% or more of the surface of each plasmon nanoparticle. In one embodiment, the coating covers about 95% or more of the surface of each plasmon nanoparticle.

[0043] In some embodiments, coating reduces the aggregation of plasmon nanoparticles compared to uncoated plasmon nanoparticles. In one embodiment, coating reduces the aggregation of plasmon nanoparticles by 35% or more compared to uncoated plasmon nanoparticles. In another embodiment, coating reduces the aggregation of plasmon nanoparticles by 50% or more compared to uncoated plasmon nanoparticles. In yet another embodiment, coating reduces the aggregation of plasmon nanoparticles by 65% ​​or more compared to uncoated plasmon nanoparticles.

[0044] In each of the aforementioned examples describing the coating of plasmon nanoparticles, the coating may contain multiple oligonucleotides. In some examples, each nucleotide within each oligonucleotide of the multiple oligonucleotides is selected from pyrimidine nucleotides. In one example, each oligonucleotide of the multiple oligonucleotides contains about 20 to about 50 nucleotides. In another example, each oligonucleotide of the multiple oligonucleotides contains about 20 to about 30 nucleotides.

[0045] In each of the aforementioned examples describing the use of the plasmon nanoantennas and / or methods of the present disclosure, the nanoscale nucleic acid scaffold may be substantially composed of double-stranded DNA. For example, the double-stranded DNA may be interconnected parallel strands of a double helix. In some embodiments, the nanoscale nucleic acid scaffold consists of a DNA origami structure, e.g., a 3D DNA origami structure. In one embodiment, the DNA origami structure includes a scaffold strand and complementary staple strands derived from M13mp18. In certain examples, at least a portion of the DNA origami structure is assembled to form a U-shaped or curved structure.

[0046] In one embodiment, the nanoscale nucleic acid scaffold includes a first surface to which one of two plasmon nanoparticles is anchored, and a second surface to which a second plasmon nanoparticle of the two plasmon nanoparticles is anchored. The first and second surfaces of the scaffold may be on the same side or adjacent sides of the nanoscale nucleic acid scaffold. Alternatively, the first and second surfaces of the scaffold may be on opposite sides of the nanoscale nucleic acid scaffold.

[0047] According to the method and / or example of a plasmon nanoantenna of the present disclosure, the plasmon nanoantenna comprises two plasmon nanoparticles, wherein one plasmon nanoparticle is anchored to a first surface of a scaffold via one or more nucleic acid linkers, and the other plasmon nanoparticle is anchored to a second surface of the scaffold via one or more nucleic acid linkers, where each nucleic acid linker is formed of an oligonucleotide that hybridizes with a polynucleotide that forms part of or extends from a nanoscale nucleic acid scaffold, which is coated on one surface of the plasmon nanoparticle.

[0048] The sequences of each oligonucleotide and polynucleotide capable of hybridizing are complementary or substantially complementary to each other. According to this example, the nanoscale nucleic acid scaffold comprises one or more polynucleotides extending from a first face of the scaffold, each of which comprises a sequence complementary to the sequence of one or more oligonucleotides coating one of the two plasmon nanoparticles. Furthermore, the nanoscale nucleic acid scaffold comprises one or more polynucleotides extending from a second face of the scaffold, each of which comprises a sequence complementary to the sequence of one or more oligonucleotides coating one of the two plasmon nanoparticles. The polynucleotides forming part of the nanoscale nucleic acid scaffold or extending from the nanoscale nucleic acid scaffold may each be single-stranded DNA. Preferably, the polynucleotides extending from the first and second faces of the scaffold each contain regions of sufficient length and complementarity to hybridize with the oligonucleotides coating the surface of the plasmon nanoparticles. In one embodiment, the polynucleotides extending from the first and second surfaces of the scaffold each contain about 20 to about 50 nucleotides. In another embodiment, the polynucleotides extending from the first and second surfaces of the scaffold each contain about 20 to about 30 nucleotides. In each of these embodiments, the polynucleotides extending from the first and second surfaces of the scaffold each contain a region of about 10 to about 25 nucleotides in length that can hybridize with a region of the corresponding length in the oligonucleotide coating the surface of the plasmon nanoparticle.

[0049] In each of the aforementioned examples describing the use of the plasmon nanoantennas and / or methods of the present disclosure, a nucleic acid polymerase is bound to or immobilized on a nanoscale nucleic acid scaffold via a nucleic acid linker. In one embodiment, the nucleic acid linker is double-stranded and comprises (i) a polynucleotide covalently or non-covalently bound to the nucleic acid polymerase via amino acids within the nucleic acid polymerase, and (ii) a polynucleotide forming part of or extending from the nanoscale nucleic acid scaffold, wherein the polynucleotides of (i) and (ii) comprise single-stranded DNA sequences that are complementary or substantially complementary to each other and capable of hybridizing. In one embodiment, the polynucleotide bound to the nucleic acid polymerase comprises a peptide tag, the peptide tag being bound to the nucleic acid polymerase via an isopeptide bond formed between an amino acid within the nucleic acid polymerase and an amino acid within the peptide tag. In one embodiment, the isopeptide bond can be formed spontaneously. For example, the isopeptide bond may be formed between Lys and Asp, or between Lys and Asn. In some embodiments, the amino acid within the nucleic acid polymerase to which the nucleic acid linker binds is positioned in the N-terminal domain of the nucleic acid polymerase.

[0050] Each polynucleotide of the nucleic acid linker that binds or immobilizes nucleic acid polymerase to a nanoscale nucleic acid scaffold contains regions of sufficient length and complementarity to hybridize with each other. In one example, each polynucleotide of the nucleic acid linker contains about 20 to about 50 nucleotides. In another example, each polynucleotide of the nucleic acid linker contains about 20 to about 30 nucleotides. In each of these examples, each polynucleotide of the nucleic acid linker contains a region of about 10 to about 25 nucleotides in length that is complementary to each other and capable of hybridizing.

[0051] In each of the aforementioned examples describing the use of the plasmon nanoantennas and / or methods thereof in the present disclosure, the nucleic acid polymerase may be a DNA polymerase.

[0052] In any example described herein, where a nucleic acid polymerase (e.g., DNA polymerase) is conjugated or immobilized to a nanoscale nucleic acid scaffold of a plasmon nanoantenna via a nucleic acid linker, the nucleic acid polymerase can be modified to express an amino acid sequence capable of binding to a peptide tag conjugated with the nucleic acid linker.

[0053] This disclosure also provides arrays comprising multiple plasmon nanoantennas as described herein.

[0054] In one embodiment, the array comprises a solid substrate, and a plurality of plasmon nanoantennas are immobilized on the solid substrate. For example, each plasmon nanoantenna in the array can be immobilized on the solid substrate via its nucleic acid scaffold. In some embodiments, the solid substrate is glass or silica. For example, the solid substrate may be glass or silica in the form of a slide or chip.

[0055] In some embodiments, less than 30% of the plasmon nanoantennas in the array are clustered in aggregates of two or more plasmon nanoantennas. For example, less than 25% of the plasmon nanoantennas in the array are clustered in aggregates of two or more plasmon nanoantennas. For example, less than 20% of the plasmon nanoantennas in the array are clustered in aggregates of two or more plasmon nanoantennas. For example, less than 15% of the plasmon nanoantennas in the array are clustered in aggregates of two or more plasmon nanoantennas. For example, less than 10% of the plasmon nanoantennas in the array are clustered in aggregates of two or more plasmon nanoantennas.

[0056] In certain examples, the arrays of this disclosure can be used in the nucleic acid analyte sequencing methods described herein. Thus, in any of the aforementioned examples describing nucleic acid analyte sequencing methods, an array comprising multiple plasmon nanoantennas described herein can be used, for example, to simultaneously sequence multiple nucleic acid analytes.

[0057] Those skilled in the art will understand that numerous variations and / or modifications can be made to the above embodiments without departing from the broad general scope of this disclosure. Accordingly, these embodiments should be considered in all respects to be illustrative and not limiting.

[0058] The following drawings form part of this specification and are included to further demonstrate certain aspects of the disclosure. This disclosure can be better understood by referring to one or more of these drawings in combination with a detailed description of the particular embodiments presented herein. [Brief explanation of the drawing]

[0059] [Figure 1] Figure 1 illustrates the design of the plasmon nanoantenna DNA sequencer described herein.

[0060] [Figure 2] Figure 2 illustrates the design of the NanoSpanno DNA origami scaffold described herein.

[0061] [Figure 3]Figure 3 illustrates the synthesis of the DNA origami NanoSpanno scaffold described herein. (A) Agarose gel stained with RedSafe® DNA staining solution. The gel ladder contains DNA molecular weight standards, lane 2 M13 contains a single-stranded M13 phage genome (7249 nucleotides), and lane 3 NS contains the synthesized NanoSpanno sample. (B) A typical TEM image at 38,000x magnification containing approximately 20 NanoSpanno particles. (C) A 3D model showing the 2D class average and proposed orientation for the image.

[0062] [Figure 4] Figure 4 shows visualizations of thiolated DNA coating on gold nanoparticles. (A) shows agarose gel electrophoresis under white transmitted light (top) and agarose gel electrophoresis with DNA-specific staining (bottom), where colloidal gold is observable as a red band. (B) TEM micrographs of bare gold nanoparticles (left) and gold nanoparticles coated with thiolated DNA (right).

[0063] [Figure 5] Figure 5 illustrates the stability of DNA-coated gold nanoparticles. DNA-coated gold nanoparticles were suspended in 1 × TAE + 6 mM MgCl2. The agarose gel was stained with a DNA-specific stain. The photograph shows the corresponding color of the sample.

[0064] [Figure 6] Figure 6 shows the expression vector map of recombinant DNA polymerases. The petDuet-1 vector is shown on the left, with relevant features labeled. MCS-1, the site from which all three recombinant polymerases were cloned, is partially enlarged on the right. Restriction sites and components of each recombinant protein are shown in order from the N-terminus to the C-terminus.

[0065] [Figure 7]Figure 7 presents exemplary data from IMAC and SEC purification of Krenow-SpyCatcher, and from SEC purification of Alexa647-labeled Krenow-SpyCatcher. (A) shows the UV280 chromatogram from IMAC purification. The green line is the gradient from the sample pump, and the orange line is the conductivity, presented as a measure of the imidazole concentration of the sample. (B) SDS PAGE of the eluted fraction from IMAC purification. (C) shows the UV280 chromatogram from SEC purification. (D) SDS PAGE of the eluted fraction from SEC purification. (E) shows the chromatogram from SEC purification of Alexa647-labeled Krenow-SpyCatcher. The blue line is the absorbance at 280 nm, and the purple line is the absorbance at 650 nm. (F) SDS PAGE of the eluted fraction from SEC purification of Alexa647-labeled Krenow-SpyCatcher. The gel on the left was used to scan for Alexa647, while the gel on the right was used for bright-field scanning.

[0066] [Figure 8] Figure 8 illustrates the two-step process used for the synthesis of the DNA-SpyTag peptide. The left side shows the reaction to generate maleimide-DNA using SMCC. The right side shows the reaction between the maleimide DNA and the thiol-cysteine ​​side chain of the peptide to obtain the DNA-peptide conjugate.

[0067] [Figure 9] Figure 9 shows data related to the HPLC purification of maleimide DNA. (A) HPLC chromatograph of 5'-maleimide-15nucleotide DNA at 260 nm; (B) HPLC chromatograph of 3'-maleimide-15nucleotide DNA; (C) Absorbance spectra and absorbance of 5'Cy5-3'-maleimide-15nucleotide DNA at 260 nm (blue) and 650 nm (orange).

[0068] [Figure 10]Figure 10 shows HPLC chromatographic data for peptide-DNA conjugates. (A) HPLC chromatograph of 5'-SpyTag-15 nucleotide DNA at 260 nm. (B) HPLC chromatograph of 3'-SpyTag-15 nucleotide DNA at 260 nm. (C) Absorbance spectra and absorbance of 5'Cy5-3'-SpyTag-15 nucleotide DNA at 260 nm (blue) and 650 nm (orange).

[0069] [Figure 11] Figure 11 shows an SDS-PAGE gel showing the conjugation of SpyTag-DNA and polymerase-SpyCatcher. (A) The image on the left shows Alexa 647 imaging, while the image on the right shows bright-field imaging of the gel after protein-specific staining. Top: 5'-SpyTag-DNA conjugate conjugated with Alexa 647-labeled Krenow-SpyCatcher. Bottom: 3'-SpyTag-DNA conjugate conjugated with Alexa 647-labeled Krenow-SpyCatcher. In each image, the left lane shows Alexa 647-labeled Krenow, and the right lane shows Alexa 647-labeled Krenow after conjugation with SpyTag-DNA. (B) The gel shows a 3'-SpyTag-DNA-5'Cy5 conjugate conjugated with unlabeled Krenow-SpyCatcher. The image on the left shows the merged imaging, the image in the middle shows the Cy5 imaging, and the image on the right shows the bright-field image of the gel after protein-specific staining. In each image, the left lane shows the Krenow-SpyTag-DNA conjugate, and the right lane shows Krenow.

[0070] [Figure 12]Figure 12 illustrates DNA-labeled polymerase function. (A) 15% Tris-glycine undenatured gel for DNA polymerase (Krenow, Tag, Phi29) activity assay. (B) 15% Tris-glycine undenatured gel for Krenow-SpyCatcher-SpyTag-DNA conjugation (left: after staining with Alexa488, right: after staining with SYBR® Gold Nucleic Acid Gel Stain).

[0071] [Figure 13] Figure 13 shows the binding dynamics of Krenow (top row) and Phi29 (bottom row) to the target DNA strand being primed. The association curves (A, D) and dissociation curves (B, E) were fitted to the biphasic exponential curve (red) described for the range of protein concentrations shown. The plateau of the association curve was plotted against protein concentration (C, F) and fitted to a single-site binding model (red).

[0072] [Figure 14] Figure 14 shows the assembly of the plasmon nanoantenna of this disclosure. (A) i: Photograph showing the color of the solution; ii: Photograph of agarose gel electrophoresis. The red band corresponds to colloidal gold; iii: Agarose gel electrophoresis image after DNA-specific staining of the gel. (B) Examples of photographs of well-formed dimers using various sequences with EM and cryo-EM. (C) Examples of photographs of monomers and aggregates.

[0073] [Figure 15]Figure 15 shows the yields with different configurations and different sequences: (A) 100 nm AuNP-25AC + NS-25TG (2 binding sites); (B) 100 nm AuNP-25AC + NS-25TG (1 binding site); (C) 100 nm AuNP-25AC + NS-25TG (0 binding sites); (D) 100 nm AuNP-25TG + NS-25AC (2 binding sites); (E) 100 nm AuNP-25TG + NS-25AC (1 binding site); (F) 100 nm AuNP-25TG + NS-25AC (0 binding sites).

[0074] [Figure 16] Figure 16 illustrates DNA polymerase bound to a cavity in a DNA origami NanoSpanno scaffold via DNA hybridization.

[0075] [Figure 17] Figure 17 shows the specific co-localization of DNA polymerase with the DNA origami scaffold. 2% agarose gel and negative control 2% agarose gel are shown to confirm the conjugation of the DNA origami scaffold with Klenow.

[0076] [Figure 18-1] Figure 18 shows the quantification of co-localization yields of DNA polymerase and DNA origami scaffolds. Data for Alexa488-labeled DNA origami scaffolds and Alexa647-labeled Krenow are shown. (A) shows an example of fluorescence image using intensity traces from the enclosed spots shown in (B). (C) shows the distribution of the first Alexa-647 relative to the first Alexa-488 intensity. (D) shows the experimentally determined co-localization yields of DNA polymerase bound to different locations in the cavities of the DNA origami scaffolds, shown in the inset photograph. NStop corresponds to molar excess, DNA origami annealed with Alexa647 DNA complementary to staple extension at the cavity edges. [Figure 18-2] Same as above.

[0077] [Figure 19-1] Figure 19 shows the quantification of fluorescence enhancement in the presence of AuNP. (A) Exemplary photobleached trace shown. (B) Distribution of photobleached step heights for measuring the intensity of a single Alexa647 fluorophore in the absence (blue) and presence (red) of AuNP binding sites in a DNA origami scaffold. (C) Quantification of photobleached rate from the data shown in (B). (D) Scattering distribution of 2×AuNP binding site measurements from 488 and 647 channels. A threshold selected using Gaussian fitting of the 488 channel was used. (E) Distribution of particle step heights from 2×AuNP experiments separated based on 488 scattering, similar to (D). (F) Photobleached rate from the data

number

[0078] [Figure 20] Figure 20 shows the results of repeated measurements of fluorophore intensity within the hotspot region by sequential binding and unbinding of a short Alexa647-labeled DNA strand (imager strand) in the absence (top) or presence (bottom) of 2AuNPs.

[0079] [Figure 21] Figure 21 shows the results of repeated intensity measurements of plasmon nanoantennas in the absence (orange) and presence (blue) of a 1 nM imager chain.

[0080] [Figure 22] Figure 22 shows the amplification of a single fluorescent DNA polymerase bound within the hotspot.

[0081] [Figure 23]Figure 23 shows the polarization-dependent change in scattering intensity of a vertical plasmon nanoantenna orientation. (A)i. Simulation of TIRF background intensity as the polarization angle of the incident light is increased. ii. Experimentally measured background intensity as the polarization angle is increased. (B)i. Predicted scattering cross-section of dimeric AuNPs aligned along (black) or perpendicular (blue) the axis of TIRF field propagation. ii. The particle intensity measured as the polarization angle is increased allows for estimation of particle number and orientation. Dimeric AuNPs or monomeric AuNPs (green) aligned along (black) or perpendicular (blue) the axis of TIRF field propagation.

[0082] [Figure 24] Figure 24 shows the traces of individual plasmon nanoantennas in the absence of the imager strand after incubation with DNA-bound AuNPs and complementary biotin-DNA. A significant reduction in variation is observed compared to Figures 21 and 22.

[0083] [Figure 25] Figure 25 illustrates simulated enhancement of Alexa647 fluorophores within a hotspot, positioned with various orientations of plasmon nanoantennas relative to the polarization of incident light.

[0084] [Figure 26-1] Figure 26 shows the fluorescent labeling of free dNTP molecules, including an HPLC chromatogram from the purification of fluorescently labeled dNTPs. [Figure 26-2] Same as above. [Figure 26-3] Same as above. [Figure 26-4] Same as above. [Figure 26-5] Same as above.

[0085] [Figure 27] Figure 27 shows the configuration of the plasmon nanoantenna DNA sequencer of this disclosure for the single-molecule DNA sequencing reaction described.

[0086] [Figure 28] Figure 28 shows an example of traces obtained from a two-color DNA sequencing experiment. The traces are from two independent plasmon nanoantenna DNA sequencer molecules. The sequencing reaction was performed using Alexa647-labeled dGTP and Alexa561-labeled dATP, with dCTP and dTTP nucleotides unlabeled. Bases identified by intensity values ​​significantly above the background by more than three standard deviations are shown in the traces. [Modes for carrying out the invention]

[0087] Detailed explanation general Throughout this specification, unless otherwise specified or the context requires a different interpretation, any reference to a single step, feature, composition of a substance, group of steps, or group of features or compositions of a substance shall be interpreted as encompassing (i.e., one or more) any of those steps, features, compositions of a substance, group of steps, or group of features or compositions of a substance.

[0088] Those skilled in the art will understand that this disclosure permits variations and modifications other than those specifically described herein. It should be understood that this disclosure encompasses all such variations and modifications. This disclosure also encompasses all steps, features, compositions and compounds that are individually or collectively referred to or shown herein, as well as any combination or any two or more of such steps or features.

[0089] This disclosure is not limited to the scope of the specific examples described herein, which are intended solely for illustrative purposes. Functionally equivalent products, compositions, and methods obviously fall within the scope of this disclosure.

[0090] Any example in this disclosure shall be construed to apply to any other example in this disclosure, with any necessary modifications, unless otherwise specified.

[0091] Unless otherwise defined, all scientific and technical terms used herein shall be construed to have the same meaning as those commonly understood by those skilled in the art (e.g., cell culture, molecular genetics, immunology, immunohistochemistry, protein chemistry, and biochemistry).

[0092] Unless otherwise specified, the recombinant DNA, recombinant proteins, cell cultures, and immunological techniques used in this disclosure are standard procedures well known to those skilled in the art. Such techniques are described in, for example, J. Perbal, A Practical Guide to Molecular Cloning, John Wiley and Sons (1984), J. Sambrook et al. Molecular Cloning: A Laboratory Manual, Cold Spring Harbor Laboratory Press (1989), TA Brown (editor), Essential Molecular Biology: A Practical Approach, Volumes 1 and 2, IRL Press (1991), DM Glover and BD Hames (editors), DNA Cloning: A Practical Approach, Volumes 1-4, IRL Press (1995 and 1996), and FM Ausubel et al. (editors), Current Protocols in Molecular Biology, Greene Pub. Associates and Wiley-Interscience (1988, including all revisions to date), Ed Harlow and David Lane (editors) Antibodies: A Laboratory Manual, Cold Spring Harbor Laboratory, (1988), and JE Coligan et al. (editors) Current Protocols in This is described and explained throughout the literature of sources such as Immunology, John Wiley & Sons (including all revisions to date).

[0093] Those skilled in the art will understand that this disclosure permits variations and modifications other than those specifically described herein. It should be understood that this disclosure includes all such variations and modifications. This disclosure also encompasses all steps, features, compositions and compounds that are individually or collectively referred to or shown herein, as well as any combination of any two or more of the aforementioned steps or features.

[0094] This disclosure is not limited to the scope of the specific embodiments described herein, which are intended solely for illustrative purposes. Functionally equivalent products, compositions, and methods obviously fall within the scope of the disclosure described herein.

[0095] Each feature of any particular aspect or embodiment of this disclosure may be applied to any other aspect or embodiment of this disclosure with necessary modifications.

[0096] Throughout this specification, unless otherwise specified or the context requires a different interpretation, references to a single step, composition of a substance, group of steps, or group of compositions of a substance shall be construed as encompassing one or more of those steps, compositions of a substance, group of steps, or group of compositions of a substance (i.e., one or more).

[0097] Throughout this specification, unless contextually required to interpret it differently, the word “comprise,” or its inflections such as “comprises” or “comprising,” shall be understood to mean that the explicitly stated step or element or integer or group of steps or elements or integers is included, but no other step or element or integer or group of elements or integers is excluded.

[0098] The terms "and / or," for example, "X and / or Y," shall be understood to mean either "X and Y" or "X or Y," and shall be construed as providing explicit support for both meanings or either meaning.

[0099] Plasmon nanoantenna This disclosure is, (i) Two plasmon nanoparticles, (ii) Nanoscale nucleic acid scaffolds, (iii) nucleic acid polymerase and A plasmon nanoantenna including, (a) Two plasmon nanoparticles are bound to a nanoscale nucleic acid scaffold and are arranged such that a plasmon hotspot exists between them relative to each other. (b) Plasmon hotspots include regions not occupied by nanoscale nucleic acid scaffolds, (c) Nucleic acid polymerase is bound to a nanoscale nucleic acid scaffold and is located within a region (plasmon hotspot) not occupied by the nanoscale nucleic acid scaffold. If necessary, including nucleic acid analytes bound to a nanoscale nucleic acid scaffold, We provide plasmon nanoantennas.

[0100] A plasmon nanoantenna according to one exemplary embodiment of the present disclosure is illustrated in Figure 1.

[0101] As used herein, the term “plasmon nanoparticles” refers to particles having nanoscale dimensions whose electron density can be coupled to electromagnetic radiation of wavelengths much larger than those of the particle due to the properties of the dielectric-metal interface between the medium and the particle. Plasmon nanoparticles can exhibit a variety of scattering, absorbance, and coupling properties based on their geometry and relative positions, and it will be understood that a variety of geometries are intended in this disclosure. The nanoparticles of this disclosure may be formed from any material having plasmon resonance properties. Suitable nanoparticles include, but are not limited to, metal nanoparticles, metal alloy nanoparticles, polymer nanoparticles and their derivatives and composites. In one particular example, each of the plasmon nanoparticles is independently selected from metal nanoparticles.

[0102] As used herein, the term “metal nanoparticles” shall be understood to mean particles formed of a metallic substance having nanoscale dimensions. It will be understood that nanoparticles can be formed into any suitable or desired shape and size. Examples thereof are given herein. Metal nanoparticles may be formed entirely or partially of any metallic material having plasmon resonance properties. The metal nanoparticles of the plasmon nanoantennas described herein may be formed entirely or partially of any metallic material known in the art having plasmon resonance properties. Metal materials known to have plasmon resonance properties include noble metals and non-noble metals. Suitable examples, but not limited to, include gold, silver, aluminum, copper, bismuth, nickel, palladium, and platinum nanoparticles or alloys of any one of these. For example, one or both or each of the metal nanoparticles may be gold nanoparticles, gold alloy nanoparticles, nanoparticles with a gold coating, or nanoparticles with a gold alloy coating. For example, one or both or each of the nanoparticles may be silver nanoparticles, silver alloy nanoparticles, nanoparticles with a silver coating, or nanoparticles with a silver alloy coating. For example, one or both or each of the nanoparticles may be aluminum nanoparticles, aluminum alloy nanoparticles, nanoparticles with an aluminum coating, or nanoparticles with an aluminum alloy coating. For example, one or both or each of the nanoparticles may be copper nanoparticles, copper alloy nanoparticles, nanoparticles with a copper coating, or nanoparticles with a copper alloy coating. For example, one or both or each of the nanoparticles may be bismuth nanoparticles, bismuth alloy nanoparticles, nanoparticles with a bismuth coating, or nanoparticles with a bismuth alloy coating. For example, one or both or each of the nanoparticles may be nickel nanoparticles, nickel alloy nanoparticles, nanoparticles with a nickel coating, or nanoparticles with a nickel alloy coating.For example, one or both or each of the nanoparticles may be palladium nanoparticles, palladium alloy nanoparticles, nanoparticles with a palladium coating, or nanoparticles with a palladium alloy coating. For example, one or both or each of the nanoparticles may be platinum nanoparticles, platinum alloy nanoparticles, nanoparticles with a platinum coating, or nanoparticles with a platinum alloy coating.

[0103] The core may be formed of glass, polymer, or composite material or other suitable material, as plasmon nanoparticles include a core coated with a metallic substance having plasmon resonance properties as described herein.

[0104] Nanoparticles formed from nonmetallic materials and metal oxides are also intended for use as plasmonic nanoparticles. For example, one or more of the plasmonic nanoparticles, or each of the plasmonic nanoparticles, may be formed from a nonmetallic material or metal oxide that is doped to have or enhance plasmonic resonance properties. Exemplary nonmetallic materials include, but are not limited to, metal oxides, chalcogenides, phosphides, nitrides, and silicon. Doped plasmonic nanoparticles and methods for producing them, but are not limited to, photodoping, chemical doping, hierarchical doping, and combinations thereof, are described in the literature and are known to those skilled in the art.

[0105] Therefore, in one embodiment, one or more of the plasmon nanoparticles are formed from a photodoped nonmetallic material or metal oxide. A method for photodoping nanoparticles to impart or enhance plasmon resonance properties, which can be used in this disclosure, is described in Petrini et al., (2023) J. Phys. Chem. C. Nanomater Interfaces, 127(3):1576-1587. In another example, one or more of the plasmon nanoparticles are formed from a hierarchically doped nonmetallic material or metal oxide. A method for hierarchically doping nanoparticles to impart or enhance plasmon resonance properties, which can be used in this disclosure, is described in Kim et al., (2023) Nano Letters, 23(16):7633-7641 and Russo et al. (2018) Materials & Designs, 156:311-319. In another example, one or more plasmon nanoparticles are formed from a nonmetallic material or metal oxide that has been chemically doped, for example, by introducing metal atoms into the matrix lattice or structure. Examples of materials that can be chemically doped to increase carrier density and plasmon resonance properties include metal oxides, chalcogenides, phosphides, nitrides, and silicon. Methods for chemically doping nanoparticles to impart or enhance plasmon resonance properties that can be used in this disclosure are described in Chowdhury et al., (2017) Nanoscale, 9:15591-15597 and Liu et al., (2019) Nature Communications, 10: 1394.

[0106] In some embodiments, the plasmon nanoparticles are formed from or contain the same material. For example, each plasmon nanoparticle may be a metal plasmon nanoparticle as described herein. For example, each plasmon nanoparticle may be a gold nanoparticle or a nanoparticle coated with gold to have a gold surface. For example, each plasmon nanoparticle may be a silver nanoparticle or a nanoparticle coated with silver to have a silver surface.

[0107] In other examples, plasmon nanoparticles may be formed from or contain different materials from one another. For example, one nanoparticle may be a gold nanoparticle or a gold-coated nanoparticle, and one or more other nanoparticles may be formed from or coated from another material (e.g., another precious metal (e.g., silver) or another metal having the plasmon resonance properties described herein).

[0108] The plasmon nanoparticles described herein can be formed into any suitable or desired shape and size, for example, as nanospheres, nanorods, nanoprisms, nanocubes, nanoshells, nanotubes, or nanostars. It should be understood that these are merely representative examples of nanoparticle shapes and are not limiting the disclosure. In one embodiment, one or more of the plasmon nanoparticles or each of the plasmon nanoparticles are nanospheres. In one embodiment, one or more of the plasmon nanoparticles or each of the plasmon nanoparticles are nanorods. In one embodiment, one or more of the plasmon nanoparticles or each of the plasmon nanoparticles are nanoprisms. In one embodiment, one or more of the plasmon nanoparticles or each of the plasmon nanoparticles are nanocubes. In one embodiment, one or more of the plasmon nanoparticles or each of the plasmon nanoparticles are nanoshells. In one embodiment, one or more of the plasmon nanoparticles, or each of the plasmon nanoparticles, are nanotubes. In one embodiment, one or more of the plasmon nanoparticles, or each of the plasmon nanoparticles, are nanostars. The plasmon nanoparticles of the plasmon nanoantenna may be the same shape or may be heterogeneous.

[0109] In addition to shape, it will be understood by those skilled in the art that the size of the nanoparticles in the plasmon nanoantenna can vary. In some embodiments, the plasmon nanoparticles may have the same or substantially the same maximum diameter. For example, the maximum diameter of each plasmon nanoparticle may be in the range of about 5 nm to about 500 nm (e.g., in the range of about 10 nm to about 250 nm, or in the range of about 20 nm to about 200 nm, or in the range of about 50 nm to about 150 nm, or in the range of about 80 nm to about 120 nm). In some embodiments, the maximum diameter of each plasmon nanoparticle is about 80 nm. In some embodiments, the maximum diameter of each plasmon nanoparticle is about 100 nm. In some embodiments, the maximum diameter of each plasmon nanoparticle is about 120 nm.

[0110] On the other hand, in other examples, the maximum diameter can differ among plasmon nanoparticles of the same plasmon nanoantenna. Following the example where the maximum diameter differs among plasmon nanoparticles of the same plasmon nanoantenna, the average maximum diameter of plasmon nanoparticles may be in the range of approximately 5 nm to approximately 500 nm. For example, the average maximum diameter of the plasmon nanoparticles in a plasmon nanoantenna may be in the range of approximately 10 nm to approximately 250 nm (e.g., in the range of approximately 20 nm to approximately 200 nm, or approximately 30 nm to approximately 200 nm, or approximately 40 nm to approximately 200 nm, or approximately 50 nm to approximately 200 nm, or approximately 60 nm to approximately 200 nm, or approximately 70 nm to approximately 200 nm, or approximately 80 nm to approximately 200 nm, or approximately 90 nm to approximately 200 nm, or approximately 100 nm to approximately 200 nm, or approximately 50 nm to approximately 150 nm, or approximately 75 nm to approximately 150 nm, or approximately 100 nm to approximately 120 nm, or approximately 100 nm to approximately 150 nm). In one embodiment, the average diameter of the nanoparticles is approximately 100 nm. In one particular example, the plasmon nanoparticles are gold or gold-coated nanoparticles (e.g., nanospheres) with an average diameter of approximately 100 nm. In another example, plasmon nanoparticles are silver or silver-coated nanoparticles (e.g., nanospheres) with an average diameter of about 100 nm.

[0111] As described herein, the plasmon nanoantennas of this disclosure comprise two plasmon nanoparticles, each attached to a nanoscale nucleic acid scaffold, and are arranged relative to each other such that a plasmon hotspot exists between the plasmon nanoparticles. However, in some embodiments, the plasmon nanoantennas may comprise more than two plasmon nanoparticles (e.g., three, four, five, six, seven, eight, nine, ten, or more nanoparticles), each attached to a nanoscale nucleic acid scaffold, and are arranged relative to each other such that a plasmon hotspot exists collectively between the plasmon nanoparticles. In the example where the plasmon nanoantennas may comprise more than two plasmon nanoparticles, two or more of the plasmon nanoparticles may aggregate.

[0112] The terms “plasmon hotspot,” “plasmon-enhanced hotspot,” or similar terms are used herein to define an area or region between two or more plasmon nanoparticles in which the electric field is enhanced compared to the electric field outside the area or region defined between two or more plasmon nanoparticles. In one embodiment, a “plasmon hotspot” is an electric field-enhanced area resulting from plasmon resonance induced by an incident oscillating electric field (e.g., light), i.e., polarization of plasmon nanoparticles.

[0113] Plasmonic nanoparticles are attached to a nanoscale nucleic acid scaffold and are arranged relative to each other such that plasmon hot spots exist that contain regions not occupied by the nanoscale nucleic acid scaffold between the plasmonic nanoparticles. As used herein, the term "region not occupied by the nanoscale nucleic acid scaffold", or similar phrase, is understood to refer to a region or space within the plasmon hot spot that is accessible when a nucleic acid polymerase, for example, is bound to the nanoscale nucleic acid scaffold. This unoccupied region may also be referred to as the "empty space" or "empty hot spot" within the plasmon hot spot. The "region not occupied by the nanoscale nucleic acid scaffold" is preferably large enough so that the enzymatic activity of the polymerase is retained when the nucleic acid polymerase is placed therein. In other words, the enzymatic activity of the polymerase is not lost due to steric effects (e.g., blockage) by the nanoscale nucleic acid scaffold. Methods for measuring and / or determining the dimensions and / or volume of the plasmon hot spot between at least two plasmon particles and the accessible empty space of the analyte are known in the art and are described, for example, in Close et al., (2022) Advanced Materials Interfaces, 9 (24) (especially in relation to Supplementary Figure 12 (S12)). For example, in the case of the plasmonic nanoantenna described herein where the empty space has a cubic form with dimensions of L = W = H = 30 nm, the empty volume of the hot spot can be calculated as 2.7×10 4 nm 3 = 2.7×10 -20 L, which can be calculated as approximately 27 zL, where 1 nm 3 is equal to 1×10 -24 L and 1 zL is equal to 1×10 -21 L. In the case of the plasmonic nanoantenna described herein where the empty space has a spherical form with a radius (r) of 15 nm, the empty volume can be calculated as 4 / 3πr 3 = approximately 14 zL, where 1 nm 3 is equal to 1×10 -24 L and 1 zL is equal to 1×10-21 It is equal to L. It will also be understood by those skilled in the art that the dimensions and / or volume of the region not occupied by the nanoscale nucleic acid scaffold can be determined based on the volume of the nanoscale nucleic acid scaffold that overlaps with the volume and / or dimensions of the plasmon hotspot. Those skilled in the art can also take into account the following factors when designing plasmon nanoparticles with a hotspot of sufficient size and intensity: the dimensions and design of the nanoscale nucleic acid scaffold, the wavelength of the incident light polarization, the propagation of the incident light, the material of the nanoparticles, the shape of the nanoparticles, the volume of the nanoparticles, the interparticle distance, the number of nanoparticles, the relative position of the nanoparticles, the relative permittivity of the surrounding medium, and / or the wavelength of emission detection.

[0114] In some embodiments, the region within the plasmon hotspot not occupied by the nanoscale nucleic acid scaffold is defined by a three-dimensional space having a length (L) between two plasmon nanoparticles, a height (H) perpendicular to (L) at the midpoint (L / 2), and a width (W) perpendicular to L and H at L / 2, where at L / 2, W is selected from approximately 20 nm to approximately 100 nm (e.g., approximately 30 nm to approximately 100 nm, or approximately The height (H) is selected from 40 nm to approximately 100 nm or approximately 50 nm to approximately 100 nm, or approximately 20 nm to approximately 50 nm or approximately 30 nm to approximately 50 nm, H is selected from approximately 10 nm to approximately 100 nm (for example, approximately 30 nm to approximately 100 nm, or approximately 40 nm to approximately 100 nm or approximately 50 nm to approximately 100 nm, or approximately 20 nm to approximately 50 nm or approximately 30 nm to approximately 50 nm), and L is measured as the shortest distance between two plasmon nanoparticles. It will be understood by those skilled in the art that the shape of the nanoparticles (examples thereof are described herein) and the geometry of the nucleic acid scaffold can affect the height and width of the plasmon hotspot, as well as the height (H) and width (W) of the unoccupied region within the plasmon hotspot. Those skilled in the art will understand that the interparticle distance of plasmon nanoparticles attached to a plasmon nanoantenna can be varied (or "adjusted"), thereby altering the size and magnitude of plasmon hotspots and regions not occupied by the nanoscale nucleic acid scaffold. As used herein, the term “interparticle distance” refers to the distance between any two plasmon nanoparticles on the nanoscale DNA scaffold of the plasmon nanoantenna. In some embodiments, the interparticle distance L between plasmon nanoparticles is about 10 nm to about 100 nm (e.g., selected from about 10 nm to about 80 nm, or about 20 nm to about 80 nm, or about 30 nm to about 80 nm, or about 40 nm to about 80 nm, or about 50 nm to about 80 nm). In another example, the interparticle distance L between plasmon nanoparticles is approximately 20 nm to 100 nm (for example, selected from approximately 20 nm to 50 nm, or approximately 30 nm to 50 nm, or approximately 20 nm to 40 nm, or approximately 30 nm to 40 nm).In one embodiment, the interparticle distance L of plasmon nanoparticles is about 20 nm to about 50 nm. In one particular example, the interparticle distance L of plasmon nanoparticles is about 30 nm. In one particular example, the volume of the region of the plasmon hotspot not occupied by the nanoscale nucleic acid scaffold is at least about 1 zL, e.g., from about 1 zL to about 10 zL, or from about 5 zL to about 10 zL. In one particular example, the volume of the region of the plasmon hotspot not occupied by the nanoscale nucleic acid scaffold is greater than or equal to about 10 zL. In some embodiments, the volume of the region of the plasmon hotspot not occupied by the nanoscale nucleic acid scaffold is at least about 10 zL (e.g., at least about 15 zL, or at least about 20 zL, or at least about 25 zL, or at least about 30 zL, or at least about 35 zL, or at least about 40 zL, or at least about 50 zL). For example, the volume of the region of a plasmon hotspot not occupied by the nanoscale nucleic acid scaffold can be about 10 zL to about 50 zL (for example, it can be selected from about 10 zL to about 50 zL, or about 20 zL to about 50 zL, or about 30 zL to about 50 zL, or about 40 zL to about 50 zL, or about 10 zL to about 40 zL, or about 20 zL to about 40 zL, or about 30 zL to about 40 zL, or about 10 zL to about 30 zL, or about 20 zL to about 30 zL, or about 40 zL to about 50 zL). In one embodiment, the region of a plasmon hotspot not occupied by the nanoscale nucleic acid scaffold is about 20 zL to about 40 zL in volume.

[0115] Alternatively, or in addition to that, the region of the plasmon hotspot not occupied by the nanoscale nucleic acid scaffold can be expressed as a relative percentage of the total volume of the plasmon hotspot. In some embodiments, the region not occupied by the nanoscale nucleic acid scaffold is 70% or more of the plasmon hotspot (e.g., 75%, 80%, 85%, 90%, 95%, or more).

[0116] Those skilled in the art will understand that plasmon hotspots, and the regions of plasmon hotspots not occupied by the nanoscale nucleic acid scaffold, can take on different shapes or forms depending, for example, the selection of nanoparticles and the geometry of the nucleic acid scaffold. In one embodiment, the regions not occupied by the nanoscale nucleic acid scaffold may be amorphous. Alternatively, the regions not occupied by the nanoscale nucleic acid scaffold may have a substantially defined shape. For example, the regions not occupied by the nanoscale nucleic acid scaffold may be spherical, elliptical, cubic, or cuboid.

[0117] Plasmon nanoparticles may be immobilized at predetermined positions on the surface of a nanoscale nucleic acid scaffold by hybridization between (i) DNA (i.e., oligonucleotide) coated plasmon nanoparticles and (ii) DNA (i.e., polynucleotides) positioned at predetermined positions and extending from the surface of the nanoscale nucleic acid scaffold that are complementary or substantially complementary to the DNA-coated plasmon nanoparticles (i.e., the nanoparticles may be immobilized by covalent bonding via Watson-Crick base pairing). In this way, the interparticle distance of plasmon nanoparticles on the surface of the nanoscale nucleic acid can be controlled by appropriate design and arrangement of the polynucleotides extending from the surface of the nanoscale nucleic acid scaffold.

[0118] In some embodiments, plasmon nanoparticles may be coated with or conjugated / linked with multiple oligonucleotides capable of hybridizing with polynucleotides extending from the surface of a nanoscale nucleic acid scaffold. For example, each plasmon nanoparticle may be coated with multiple DNA oligonucleotides selected from 2 to about 100 oligonucleotides (e.g., 2 to about 75 oligonucleotides, or 2 to about 50 oligonucleotides, or 2 to about 25 oligonucleotides, or 2 to about 10 oligonucleotides). In other examples, each plasmon nanoparticle may be coated with 5 or more DNA oligonucleotides (e.g., 6, 7, 8, 9, 10, 11, 12, 13, 14, or 15, or more). The DNA oligonucleotide coating preferably substantially covers the surface of each plasmon nanoparticle. For example, the DNA oligonucleotide coating may cover about 75% or more of the surface of each plasmon nanoparticle (e.g., selected from about 80%, 85%, 90%, 95%, or more).

[0119] The DNA oligonucleotide coating the plasmon nanoparticles facilitates the immobilization of the coated plasmon nanoparticles onto the surface of the nanoscale nucleic acid scaffold, and also reduces the aggregation of plasmon nanoparticles during the preparation of plasmon nanoantennas compared to uncoated plasmon nanoparticles. Exemplary methods for measuring nanoparticle aggregation are known in the art, but are not limited to, including ultraviolet-visible spectroscopy, differential centrifugal sedimentation, particle counters and / or dynamic light scattering, particle tracking analysis, and single-particle inductively coupled plasma mass spectrometry. These exemplary methods are described in Minelli et al., (2019) Langmuir, 35 (14):4927-4935. In one example, coating with DNA oligonucleotides reduces the aggregation of plasmon nanoparticles by 35% or more (e.g., 40% or 45% or more) compared to uncoated plasmon nanoparticles. In one embodiment, coating with DNA oligonucleotides reduces the aggregation of plasmon nanoparticles by 50% or more (e.g., 55%, 60%, or more) compared to uncoated plasmon nanoparticles. In another embodiment, coating with DNA oligonucleotides reduces the aggregation of plasmon nanoparticles by 65% ​​or more (e.g., 70%, 75%, or more) compared to uncoated plasmon nanoparticles.

[0120] Coating or conjugation of plasmon nanoparticles with one or more DNA molecules (e.g., oligonucleotides) can be carried out by any method known in the art. However, in one embodiment, one or more oligonucleotides are linked to the plasmon nanoparticles by a linker. In one particular example, the oligonucleotides can be thiolated according to the examples disclosed herein. However, it will be understood that other functional anchoring groups, such as -COOH, -NH2, and -OH, can also be used to covalently attach DNA oligonucleotides to the surface of nanoparticles.

[0121] Plasmon nanoparticles coated with DNA oligonucleotides are designed so that their nucleotide sequences are complementary or substantially complementary to the sequences of congeneral DNA polynucleotides extending from the surface of a nanoscale nucleic acid scaffold, enabling hybridization with them. In certain examples, more than 50% (e.g., more than 50%, 60%, 70%, or 80%) of the nucleotides in plasmon nanoparticles coated with oligonucleotides or multiple oligonucleotides are selected from pyrimidine nucleotides (i.e., thymine and cytosine). In some examples, each nucleotide in plasmon nanoparticles coated with oligonucleotides or multiple oligonucleotides is selected from pyrimidine nucleotides (i.e., thymine and cytosine).

[0122] DNA oligonucleotides attached to plasmon nanoparticles, and complementary or substantially complementary DNA polynucleotides extending from or within a nanoscale nucleic acid scaffold, may be modified or unmodified. Suitable modifications are known in the art. These modifications include, for example, azo modifications known in the art that can be used to achieve photoresponsive reversible binding and unbinding of each DNA molecule. Further modifications include modifications that modulate the strength of interactions between each DNA molecule, such as skeletal modifications to alter surface charge (e.g., peptide nucleic acids (PNAs)) or structural flexibility to induce hybridization efficiency (e.g., locked nucleic acids (LNAs)). Yet another modification is the use of photocleavable structures known in the art.

[0123] As described herein, plasmon nanoantennas include nanoscale nucleic acid scaffolds for immobilizing or binding plasmon nanoparticles and nucleic acid polymerase molecules. The nanoscale nucleic acid scaffold may be a nanoscale DNA scaffold. The nanoscale DNA scaffold may include or be formed of one or more DNA origami structures. “DNA origami structure” and similar terms are understood to refer to scaffolds or structures formed by DNA molecules that self-assemble and are fabricated by so-called DNA origami techniques. Scaffolded DNA origami is a technique proposed by Paul Rothemund in 2006, building upon earlier methods of structural DNA nanotechnology and resulting in the utilization of finite DNA nanomaterials. In this technique, single-stranded circular DNA molecules, typically 7.25 kb in size, are folded into helical arrays by a periodic cross arrangement using staple strands. Extending this concept to assemble 3D DNA structures has led to the establishment of a robust and reliable method for assembling DNA nanostructures (or "nanoscale DNA scaffolds") with dimensions in the range of 20–100 nm. However, larger structures are also possible. DNA origami structures may be bundles of tubes or pipes, e.g., 3-helix bundles, 6-helix bundles, or 12-helix bundles, and may have a substantially flat, rectangular shape. Thus, by combining DNA origami structures of various sizes and dimensions (e.g., stacking or layering), nanoscale DNA scaffolds of any particular shape can be constructed. The origami structures are addressable to several thousand nanometers, allowing any object of interest (OOI) to be positioned with a "single-pixel" resolution of approximately 6 nanometers. 2It can be used as a “molecular pegboard” having a surface area. Furthermore, the shape, dimensions, and contour of the surface of the origami structure can be adjusted so that the nanoscale DNA scaffold has a geometrically suitable surface for attaching the target molecule (e.g., plasmon nanoparticles and polymerases described herein). Methods for generating DNA origami structures, including those having DNA sequence strands protruding from the structure, are known in the art. In one embodiment, the nanoscale DNA scaffold comprises one or more 3D DNA origami structures. In a particular example, at least a portion of the DNA origami structures are assembled to form a U-shaped or curved structure. Exemplary U-shaped structures are illustrated in Figures 1-3.

[0124] Following the example of the nanoscale DNA scaffold being assembled in a U-shape, the nanoscale DNA scaffold includes a first surface to which one of two plasmon nanoparticles is anchored via a nucleic acid linker, and a second surface to which a second plasmon nanoparticle of the two plasmon nanoparticles is anchored via a nucleic acid linker. The first and second surfaces of the scaffold may be on the same or adjacent surfaces of the nanoscale DNA scaffold, or they may be on opposite sides.

[0125] In one embodiment, the nanoscale DNA scaffold comprises (i) one or more DNA polynucleotides extending from a first surface of the scaffold, each containing a sequence complementary to the sequence of one or more DNA oligonucleotides coating one of two plasmon nanoparticles, and (ii) one or more DNA polynucleotides extending from a second surface of the scaffold, each containing a sequence complementary to the sequence of one or more DNA oligonucleotides coating one of two plasmon nanoparticles. Each of the DNA polynucleotides forming part of the nanoscale DNA scaffold or extending from the nanoscale DNA scaffold may be single-stranded DNA or may contain single-stranded DNA. Each of the DNA polynucleotides extending from the first and second surfaces of the scaffold contains a region of single-stranded DNA that is long enough to allow hybridization and is complementary to the plasmon nanoparticles coating the DNA oligonucleotides. When congeneral DNA polynucleotides and DNA oligonucleotides hybridize to form a nucleic acid linker, plasmon nanoparticles containing the DNA oligonucleotide are anchored to the surface of a nanoscale DNA scaffold. Each nucleic acid linker anchoring the plasmon nanoparticles to the nanoscale DNA scaffold is formed by one of the DNA oligonucleotides coated on the surface of a plasmon nanoparticle hybridized with a DNA polynucleotide that forms part of or extends from the nanoscale DNA scaffold. In one embodiment, the DNA polynucleotides extending from the first and second surfaces of the scaffold contain a scaffold sequence derived from M13mp18, and the plasmon nanoparticles coating the DNA oligonucleotide contain a staple sequence complementary to its scaffold sequence.

[0126] The lengths of the DNA polynucleotides extending from the first and second surfaces of the scaffold can be varied as desired. However, in certain examples, the DNA polynucleotides extending from the first and second surfaces of the scaffold each contain approximately 20 to 50 nucleotides (e.g., approximately 20 to 40 nucleotides, or approximately 20 to 30 nucleotides, or approximately 30 to 50 nucleotides, or approximately 30 to 50 nucleotides). In one embodiment, the DNA polynucleotides extending from the first and second surfaces of the scaffold contain approximately 20 to 30 nucleotides.

[0127] In a particular example, each plasmon nanoparticle is anchored to a nanoscale DNA scaffold via multiple nucleic acid linkers (e.g., via two, three, four, five, six, seven, eight, nine, ten, or more nucleic acid linkers). It will be understood that the number of nucleic acid linkers connecting each plasmon nanoparticle is positively correlated with the strength of binding of the plasmon nanoparticle to the nanoscale DNA scaffold. Thus, the number of DNA polynucleotides extending from the nanoscale DNA scaffold and hybridizing with the DNA oligonucleotides coating the plasmon nanoparticles can be freely selected, depending on the specific application of the purpose, to achieve the desired binding strength and / or desired hybridization efficiency. Furthermore, the length of the Watson-Crick pairing region in the nucleic acid linker (i.e., between the DNA polynucleotide(s) extending from the nanoscale DNA scaffold and the DNA oligonucleotide(s) attached to the plasmon nanoparticles) can be varied to achieve desired binding specificity and / or binding strength and / or desired hybridization efficiency, depending on the specific application of interest. In certain examples, the Watson-Crick pairing region is approximately 8–30 nucleotides long, or approximately 10–25 nucleotides long, or approximately 12–20 nucleotides long. The terms “hybridize,” “to hybridize,” or similar terms, as used herein, refer to the process by which two complementary or substantially complementary nucleic acid sequences anneal to each other according to the Watson-Crick base pairing rules.

[0128] Those skilled in the art will understand that the DNA oligonucleotides attached to the plasmon nanoparticles, and the complementary or substantially complementary DNA polynucleotides extending from or within the nanoscale DNA scaffold (together hybridizing to form a nucleic acid linker that anchors the plasmon nanoparticles to the nanoscale DNA scaffold), may be of any length. For example, each DNA molecule may have a length independently selected from between about 20 and about 50 nucleotides, or between about 20 and about 40 nucleotides, or between about 20 and about 30 nucleotides. The Watson-Crick pairing region between the DNA polynucleotide sequence(s) extending from the nanoscale nucleic acid scaffold (e.g., DNA origami structure) and the DNA oligonucleotide sequence attached to the plasmon nanoparticles is preferably, for example, about 8 to 30 nucleotides, or about 10 to 25 nucleotides, or about 12 to 20 nucleotides.

[0129] When used herein, “hybridize,” “to hybridize,” or similar terms refer to the process by which two complementary or substantially complementary nucleic acid sequences anneal to one another according to the Watson-Crick base pairing rules.

[0130] As used herein, the term “substantially complementary” refers to two sequences that are fully or partially complementary to each other and therefore hybridize under appropriately stringent hybridization conditions. In other words, the term “substantially complementary” is used to indicate a sufficient degree of complementarity or tight pairing between two nucleic acid sequences, for example, between a DNA sequence attached to a plasmon nanoparticle and a complementary or substantially complementary DNA sequence in a nanoscale DNA scaffold. It is understood that a nucleic acid sequence does not need to be 100% complementary to its target or complement sequence (although this may be one preferred example). For example, a nucleic acid sequence may be at least 80%, or at least 85%, or at least 90%, or at least 95%, or at least 96%, or at least 97%, or at least 98%, or at least 99% complementary to its target or complement sequence.

[0131] As described herein, this disclosure is based in part on the ability to immobilize nucleic acid polymerase within an unoccupied area (or “empty space”) of a plasmon hotspot formed by a plasmon nanoantenna as described herein, while simultaneously maintaining the biological activity of the nucleic acid polymerase. For example, the immobilization of nucleic acid polymerase on the surface of a nanoscale nucleic acid scaffold is achieved by a nucleic acid linker. In one embodiment, the nucleic acid linker that tethers nucleic acid polymerase to the surface of a nanoscale nucleic acid scaffold (e.g., a nanoscale DNA scaffold as described herein) is a double-stranded DNA linker comprising (i) DNA polynucleotides covalently or non-covalently bonded to the nucleic acid polymerase via amino acids within the nucleic acid polymerase, and (ii) DNA polynucleotides forming part of the surface of the nanoscale nucleic acid scaffold or extending from the surface of the nanoscale nucleic acid scaffold, wherein the polynucleotides of (i) and (ii) each comprise single-stranded DNA sequences that are complementary or substantially complementary to each other and capable of hybridizing with each other. The polynucleotide bound to nucleic acid polymerase may contain a peptide tag, which is further bound to the nucleic acid polymerase via an isopeptide bond formed between an amino acid in the nucleic acid polymerase and an amino acid in the peptide tag. In this case, the inventors used the SpyCatcher-SpyTag bioconjugation system described in Hatlem et al (2019) Int. J. Mol. Sci, 20(9):2129 to attach the DNA polynucleotide of a double-stranded DNA linker to nucleic acid polymerase. Following an example of using the SpyCatcher-SpyTag system, the polymerase is modified to express the SpyCatcher protein on its surface. Then, a homologous SpyTag is conjugated to the polynucleotide by any means known in the art.Methods for preparing oligonucleotide-peptide conjugates are known in the art, for example, described in Klabenkova et al (2021) Molecules, 26(17):5420, and include, but are not limited to, conjugation via thioether or disulfide bonds, native ligation, oxime linkage, thiazolidinedione linkage or hydrazone linkage, amine bond formation, click chemistry, Diels-Alder reaction, and thiol-maleimide conjugation. However, in one particular example, a DNA polynucleotide is modified to contain maleimide, and SpyTag is conjugated with the DNA polynucleotide by thiol-maleimide conjugation (for example, as described in the examples herein). Once the DNA polynucleotide containing SpyTag is prepared, the DNA polynucleotide can be contacted with a polymerase modified to express the SpyCatcher protein. Upon recognition, SpyCatcher and SpyTag form a covalent isopeptide bond between the lysine side chain of SpyCatcher and the aspartic acid of SpyTag, resulting in the binding of the polymerase to the DNA polynucleotide of the double-stranded DNA linker. While the SpyCatcher-SpyTag system is used in this disclosure, it will be understood by those skilled in the art that alternative enzyme-mediated conjugation systems (including modified forms of the Catcher-Tag system) may be used, and that alternative amino acid combinations known to form isopeptide bonds and covalently bind the DNA polynucleotide of the double-stranded DNA linker may be relied upon. Those skilled in the art will also be aware of alternative chemistry for protein ligation and bioconjugation that can be used, which are intended in this disclosure.

[0132] In some embodiments, the amino acid within the polymerase to which the polynucleotide linker binds is positioned in the N-terminal domain of the polymerase.

[0133] Each DNA polynucleotide contained within the double-stranded DNA linker that binds or immobilizes the nucleic acid polymerase to the nanoscale nucleic acid scaffold contains a region of sufficient length and complementarity to hybridize with one another. Each DNA polynucleotide contained within the double-stranded DNA linker may be about 20 to about 50 nucleotides long (e.g., about 20 to about 30 nucleotides, or about 20 to about 40 nucleotides, or about 30 to about 40 nucleotides). In one embodiment, the DNA polynucleotides within the double-stranded DNA linker are between about 20 and about 30 nucleotides long. The Watson-Crick pairing region between each DNA polynucleotide contained within the double-stranded DNA linker that binds or immobilizes the polymerase to the nanoscale nucleic acid scaffold is preferably about 8 to 30 nucleotides long, or about 10 to 25 nucleotides long, or about 12 to 20 nucleotides long. In this regard, those skilled in the art can adjust the length of the Watson-Crick pairing region within the double-stranded DNA linker so that a desired binding specificity and / or binding strength and / or desired hybridization efficiency are achieved.

[0134] DNA polynucleotides contained within a double-stranded DNA linker that binds or immobilizes nucleic acid polymerase to a nanoscale nucleic acid scaffold may include one or more modifications. Suitable modifications are known in the art. Examples of such modifications include, for example, azo modifications known in the art that can be used to achieve photoresponsive, reversible binding and unbinding of each DNA sequence. Further modifications include modifications that modulate the strength of interactions between each DNA sequence, such as skeletal modifications to alter surface charge (e.g., peptide nucleic acids (PNAs)) or structural flexibility to induce hybridization efficiency (e.g., locked nucleic acids (LNAs)). Yet another modification is the use of photocleavable structures known in the art.

[0135] In each of the aforementioned examples describing the plasmon nanoantennas of this disclosure, the nucleic acid polymerase may be a DNA polymerase.

[0136] The terms “DNA polymerase” or “DNA polymerase molecule,” as used herein, refer to the enzyme that synthesizes a new strand of DNA in the 5'-3' direction from a primer hybridized to a DNA template strand. DNA polymerases are well known and commercially available in the art. DNA polymerase “reads” the template in the 3'→5' direction and adds individual nucleotides (bases) to the new or nascent strand in the 5'→3' direction. The polymerase requires the 3'OH group from the primer to initiate the elongation of the new DNA strand. As the new DNA strand is synthesized by DNA polymerase, individual nucleotides (dNTPs, or dATP, dCTP, dTTP, dGTP, or A, C, T, G) are repeatedly added to the elongating DNA molecule. The specific bases (A, C, T, or G) depend on the sequence of the template DNA, and therefore the new bases hybridize to the nucleotides of the template strand through Watson-Crick interactions. DNA polymerase switches between an "open" and a "closed" conformation. DNA polymerase is in the open position with the primer-template DNA complex. When a new nucleotide enters the active site, the polymerase switches to the closed position.

[0137] Various DNA polymerases can be used, many of which are commercially available. At least five families of DNA-dependent DNA polymerases are known, but the majority belong to families A, B, and C. Most family A polymerases are single-chain proteins that may contain multiple enzymatic functions, including polymerase activity, 3'→5' exonuclease activity, and 5'→3' exonuclease activity. Family B polymerases generally have a single catalytic domain with polymerase activity and 3'→5' exonuclease activity, as well as accessory factors. Family C polymerases are generally multi-subunit proteins with polymerization activity and 3'→5' exonuclease activity. In E. coli, three types of DNA polymerases have been found: DNA polymerases I, II, and III (similar to families A, B, and C, respectively). In eukaryotic cells, three distinct family B polymerases, DNA polymerases α, δ, and ε, are associated with nuclear replication, while polymerase γ, a family A polymerase, is used for mitochondrial DNA replication. Other types of DNA polymerases include phage polymerases. Any of these polymerases, combinations of all or some of these polymerases, and chimeras or hybrids between two or more such polymerases, or their equivalents, can be used to form some or all of the DNA polymerase molecules of the plasmon nanoantennas described herein.

[0138] Examples of DNA polymerases that can be used include, but are not limited to, phi-29, Taq, T7, E. coli Klenow (derived from DNA pol I), E. coli DNA pol III, and Baccilus stearothermophilus (Bst) DNA pol. DNA polymerases may also be genetically modified, such as hybrids (e.g., Phusion DNA polymerases in which a domain with strong dsDNA binding affinity is fused to the DNA polymerase to enhance processing capacity). Many useful DNA polymerases are commercially available (e.g., T7 DNA pol, Sequenase version 2.0™). Highly processable polymerases include phi29 and T7 DNA polymerases, and Moloney's mouse leukemia virus (M-MLV) reverse transcriptase. Those skilled in the art will understand that DNA polymerases are structurally similar and that recombinant and hybrid polymerases can be manipulated using homologous domains derived from different polymerases.

[0139] In some embodiments, to avoid the potential for variations in the integration efficiency of fluorescently labeled dNTPs, DNA polymerases and labels can be used that maximize integration efficiency and minimize integration variability. Some DNA polymerases can efficiently integrate fluorescently labeled dNTPs along a DNA template to produce highly uniform fragments regardless of sequence conditions, and these polymerases are used in the most automated Sanger / dideoxy-based methods using capillary-array DNA sequencers. Several natural and engineered DNA polymerases can incorporate fluorescently labeled dNTPs (Reeve & Fuller Nature 376, 796-97 (1995); Rosenblum et al., Nucleic Acids Res 25, 4500-04 (1997); Ramanathan et al. Anal Biochem 337, 1-11 (2005); Aksyonov et al., Anal Biochem 348, 127-138 (2006); Tabor & Richardson, J Biol Chem 265, 8322-28 (1990); Zhu et al., Nucleic Acids Res 22, 3418-22 (1994); Zhu & Waggoner Cytometry 28, 206-211 (1997); Randolph & Waggoner Nucleic Acids Res 25, 2923-29 (1997);Mitra et al., Anal Biochem 320, 55-65 (2003);Anderson et al. Biotechniques 38, 257-264 (2005)).

[0140] As described herein, DNA polymerase can be modified to express an amino acid sequence (e.g., CatcherTag or other binding motif) capable of binding to a conjugate homologous peptide tag (e.g., SpyTag or other tag recognized by a binding motif) within a nucleic acid linker (e.g., a double-stranded DNA linker) described herein for anchoring the polymerase to the surface of a nanoscale nucleic acid scaffold.

[0141] In some embodiments, the plasmon nanoantennas described herein can be immobilized on a substrate via their nucleic acid scaffold. In certain examples, the substrate is a solid substrate. For example, the plasmon nanoantennas described herein can be immobilized on glass or silica. For example, the plasmon nanoantennas described herein can be immobilized on a glass or silica slide or a glass or silica chip. However, other solid substrates are known to those skilled in the art and are intended in this disclosure. In some embodiments, the substrate can be coated with an antifouling material.

[0142] Array of plasmon nanoantennas This disclosure also provides arrays comprising the plurality of plasmon nanoantennas described herein. For example, the plurality of plasmon nanoantennas of this disclosure can be immobilized on a substrate. For instance, each plasmon nanoantenna in the array can be immobilized on a solid substrate via its nucleic acid scaffold. Each plasmon nanoantenna can be immobilized on a solid substrate via a nanoscale DNA scaffold using means known in the art. For example, a nanoscale DNA scaffold can be immobilized on a solid substrate via a DNA linker or via a DNA origami structure. However, it will be understood by those skilled in the art that there are many methods for immobilizing biological analytes, such as the plasmon nanoantennas of this disclosure, on a substrate, whether by covalent or non-covalent bonding, via a linker portion, or by tethering to the immobilized portion. These methods are well known in the fields of solid-phase synthesis and microarrays (Beier et al., Nucleic Acids Res. 27:1970-1-977 (1999)). Non-limiting exemplary binding moieties for attaching either nucleic acids or proteins to a solid support include, among others, streptavidin or avidin / biotin linkages, carbamic acid linkages, ester linkages, amides, thiol esters, (N)-functionalized thiourea, functionalized maleimide, amino, disulfide, amide, and hydrazone linkages. Furthermore, silyl moieties can be attached to nucleic acids using methods known in the art and directly attached to substrates such as glass. Additional methods for immobilization are presented in U.S. Patent Applications 11 / 645,125 and 11 / 645,135, both filed on December 21, 2006; and U.S. Patent Publication 20080199932, all of which are incorporated herein in their entirety for all purposes.In one specific, non-limiting example, avidin / biotin linkages are used to immobilize each plasmon nanoantenna on the surface of a substrate (e.g., glass or silica piece), thereby conjugating biotin molecules to each plasmon nanoantenna (e.g., via a linker as needed), and immobilizing the nanoantennas on the substrate via avidin (e.g., streptavidin) immobilized on the surface.

[0143] The array may include a plurality of plasmon nanoantennas placed on a substrate at a desired density and spacing in a desired pattern (e.g., at constant intervals).

[0144] The substrate may be made of any suitable material to which plasmon nanoantennas can be attached covalently or non-covalently, such as glass, nylon, carbohydrates such as dextran, plastics such as polystyrene or polypropylene, materials including polyacrylamide, latex, silicon, paper, metals such as gold, chromium, titanium or tin, titanium oxide, tin oxide, or cellulose. According to a particular example, the substrate is glass, for example, a glass slide or chip. Any suitable glass can be used, but a preferred substrate has at least a silicon dioxide surface layer. In another particular example, the substrate is a flexible plastic. For example, the flexible plastic may include nitrocellulose acetate. The flexible plastic substrate may also include a flexible plastic strip. The use of a flexible plastic substrate allows for greater portability and increased adaptability to interfacing with other devices in the array. For example, it can be wrapped around optical fibers and optical cables to increase sensitivity.

[0145] The substrate can further be provided in any desired form, including chips, beads, wells, flow cells, nanowells, capillary tubes, slides, wafers, filters, fibers, porous media, porous nanotubes, or columns.

[0146] In some embodiments, the assay can be configured to assay (e.g., sequence) two or more samples simultaneously. In this regard, the solid substrate on which the plasmon nanoantennas are immobilized may be partitioned, with each region containing a separate array of plasmon nanoantennas. For example, the substrate may be a microwell plate or a microchannel plate. The microwell plate or microchannel plate may also be configured to be readable by a standard plate reader, such as a fluorometer.

[0147] In some embodiments, the surface of the substrate can be modified to enable or enhance the covalent or noncovalent adhesion of plasmon nanoantennas as described herein. The substrate and process for attaching plasmon nanoantennas to the substrate are preferably stable to repeated bonding, washing, imaging, and elution steps (if desired). In some embodiments, the surface of the substrate can be modified to have a positive or negative charge. In some embodiments, the surface of the substrate can be functionalized by modifying with specific functional groups such as maleic or succinic moieties, or derivatized by modifying with chemically reactive groups such as amino, thiol, or acrylate groups, for example, by silane treatment. Suitable silane reagents include aminopropyltrimethoxysilane, aminopropyltriethoxysilane, and 4-aminobutyltriethoxysilane. The surface can be functionalized with N-hydroxysuccinimide (NHS) functional groups. The glass surface can also be derivatized using other reactive groups, such as acrylates or epoxys, for example, epoxysilane, acrylatesilane, or acrylamidesilane.

[0148] In some embodiments, a solid substrate can be modified to reduce the nonspecific adhesion of plasmon nanoantennas to the surface of the solid substrate. In some embodiments, a solid support can be modified to reduce the nonspecific binding of biological and / or chemical entities to the surface of the substrate. For example, the surface of the substrate can be antifouling to prevent nonspecific adsorption to the surface of the substrate. Suitable antifouling reagents are known in the art and are intended in this disclosure. However, in one particular example, the surface of the substrate is treated with APTES. In some embodiments, the substrate can be passivated, and / or the surface of the substrate can be passivated. For example, the passivation layer may include diamond-like carbon, hexa-methyldisilane, Teflon®, fluorocarbons, polymers such as polyethylene glycol (PEG), and / or parylene. In some embodiments, the substrate can be passivated by depositing polyethylene glycol (PEG) molecules across the substrate. In some embodiments, the passivation component does not have to be covalently bonded to the surface of the substrate. For example, the substrate can be passivated by coating it with bovine serum albumin (BSA) or casein (e.g., derived from powdered milk).

[0149] In some embodiments, the solid substrate can be modified across the entire surface to which plasmon nanoantennas are attached. In other embodiments, the surface of the solid substrate may include areas that are modified to allow plasmon nanoantenna attachment and areas that are not modified. Alternatively, or in addition to these, the surface of the solid substrate may include areas that are modified to reduce plasmon nanoantenna attachment and areas that are not modified. Alternatively, or in addition to these, the surface of the solid substrate may include areas that are modified to increase plasmon nanoantenna attachment and areas that are modified to decrease plasmon nanoantenna attachment. In some cases, the plasmon nanoantenna attachment sites can be created in an array, for example, in a regular array.

[0150] Regular arrays of attachment sites can be created, for example, by photographic plate, dip-pen nanolithography, nanoimprint lithography, nanosphere lithography, cluster lithography, nanopillar array, nanowire lithography, scanning probe lithography, thermochemical lithography, thermal scanning probe lithography, localized oxidation nanolithography, molecular self-assembly, stencil lithography, or electron beam lithography. The attachment sites of the regular array can be positioned at any density and / or spatial arrangement.For example, in a regular array of attachment sites, each attachment site is less than 20 nm, or about 20 nm, or about 50 nm, or about 75 nm, or about 100 nm, or about 125 nm, or about 150 nm, or about 175 nm, or about 200 nm, or about 225 nm, or about 250 nm, or about 275 nm, or about 300 nm, or about 325 nm, or about 350 nm, or about 375 nm, or about 400 nm, or about 425 nm, or about 450 nm, or about 475 nm, or approximately 500 nm, or approximately 525 nm, or approximately 550 nm, or approximately 575 nm, or approximately 600 nm, or approximately 625 nm, or approximately 650 nm, or approximately 675 nm, or approximately 700 nm, or approximately 725 nm, or approximately 750 nm, or approximately 775 nm, or approximately 800 nm, or approximately 825 nm, or approximately 850 nm, or approximately 875 nm, or approximately 900 nm, or approximately 925 nm, or approximately 950 nm, or approximately 975 nm, or approximately 1000 nm, or approximately 1025 nm, or approximately 1050nm, or approximately 1075nm, or approximately 1100nm, or approximately 1125nm, or approximately 1150nm, or approximately 1175nm, or approximately 1200nm, or approximately 1225nm, or approximately 1250nm, or approximately 1275nm, or approximately 1300nm, or approximately 1325nm, or approximately 1350nm, or approximately 1375nm, or approximately 1400nm, or approximately 1425nm, or approximately 1450nm, or approximately 1475nm, or approximately 1500nm, or approximately 1525nm, or approximately 1550nm, and It can be positioned to be farther away from approximately 1575nm, or approximately 1600nm, or approximately 1625nm, or approximately 1650nm, or approximately 1675nm, or approximately 1700nm, or approximately 1725nm, or approximately 1750nm, or approximately 1775nm, or approximately 1800nm, or approximately 1825nm, or approximately 1850nm, or approximately 1875nm, or approximately 1900nm, or approximately 1925nm, or approximately 1950nm, or approximately 1975nm, or approximately 2000nm, or 2000nm.

[0151] In some embodiments, the spacing between attachment sites on the surface of the solid substrate can be selected according to the size of the plasmon nanoantennas to be attached.

[0152] In some embodiments, less than 30% of the plasmon nanoantennas in the array are clustered in aggregates of two or more plasmon nanoantennas. For example, less than 25% of the plasmon nanoantennas in the array are clustered in aggregates of two or more plasmon nanoantennas. For example, less than 20% of the plasmon nanoantennas in the array are clustered in aggregates of two or more plasmon nanoantennas. For example, less than 15% of the plasmon nanoantennas in the array are clustered in aggregates of two or more plasmon nanoantennas. For example, less than 10% of the plasmon nanoantennas in the array are clustered in aggregates of two or more plasmon nanoantennas.

[0153] In some embodiments, the substrate may be optically opaque. In some embodiments, the substrate may be optically transparent at one or more wavelengths. In some examples, the substrate may be partially optically transparent, or optically transparent in some areas. For example, a solid substrate may be optically opaque in unfunctionalized areas and optically transparent in functionalized areas.

[0154] Methods for determining DNA sequences The inventors have shown for the first time that when sequencing a single DNA molecule, a DNA polymerase molecule can be positioned within an electric field-enhanced area, and therefore, during DNA synthesis, it is possible to detect the incorporation of specific fluorescently labeled dNTPs into the growing strand of DNA by the DNA polymerase enzyme.

[0155] Therefore, in one embodiment, the present disclosure is a method for sequencing nucleic acid analytes, (I) A step of contacting nucleic acid polymerase with the nucleic acid analyte and the labeled nucleotide for a period of time and under conditions such that the labeled nucleotide is sequentially incorporated by the polymerase into a polynucleotide having a sequence complementary to the polynucleotide sequence of the nucleic acid analyte, wherein the polymerase is located within an electric field enhancement area and each labeled nucleotide is (i) adenine nucleotide (A), guanosine nucleotide (G), thymine nucleotide (T), or cytosine nucleotide (C), (ii) Fluorophores, and (iii) Polyphosphate linkers that bind nucleotides to fluorophores Includes, A, G, T, and C are each independently linked to the fluorophore via a polyphosphate. Each of the labeled nucleotides A, G, T, and C has a distinct fluorescence signature when the fluorophore to which the respective nucleotide is linked is excited. The distinct fluorescent signature of the labeled nucleotide is enhanced when the labeled nucleotide is incorporated into a sequence complementary to the polynucleotide sequence of the nucleic acid analyte by nucleic acid polymerase. Steps and (II) A step of detecting the sequence of enhanced, distinct fluorescence signatures when labeled nucleotides are sequentially incorporated by nucleic acid polymerase into polynucleotides having a sequence complementary to the polynucleotide sequence of the nucleic acid analyte, thereby determining the sequence of the nucleic acid analyte by determining the sequence of nucleotides incorporated into polynucleotides having a sequence complementary to the polynucleotide sequence of the nucleic acid analyte, and This provides a method that includes [something].

[0156] For fluorophores attached to nucleotides to emit a fluorescent signal when colocalized with polymerase, an electromagnetic energy source must be supplied to the sequencing reaction. Therefore, the method includes the step of supplying an electromagnetic energy source to the labeled nucleotide when it colocalizes with DNA polymerase. In one embodiment, the electromagnetic energy is supplied by one or more lasers. The one or more lasers are configured to supply electromagnetic energy (e.g., light) to the labeled nucleotide at a wavelength corresponding to the excitation wavelength of the fluorophores attached to the nucleotide. As used herein, the phrase “electric field enhancement area” refers to an area or region where the electric field is enhanced compared to the electric field outside a defined area. In some cases, and in the case of DNA sequencing, the enhancement of the electric field is due to plasmon resonance induced by an incident oscillating electric field (e.g., light).

[0157] In some embodiments, the electric field enhancement area is generated by a plasmon hotspot, which is generated by a plasmon nanoantenna described herein. Thus, in some embodiments, the method includes the step of sequencing a nucleic acid analyte using a plasmon nanoantenna described herein, in accordance with the method described herein. In this example, step (I) of the method includes contacting the nucleic acid analyte and the labeled nucleotide with a plasmon nanoantenna described herein for a sufficient amount of time and under conditions to allow the labeled nucleotide to be sequentially incorporated by a nucleic acid polymerase, wherein the polymerase is bound to a nanoscale nucleic acid (e.g., DNA) scaffold and is located within a region of the plasmon hotspot not occupied by the nanoscale nucleic acid scaffold.

[0158] In one embodiment, the method includes the step of contacting a nucleic acid polymerase with a nucleic acid analyte in the presence of labeled nucleotides A, G, T, and C, respectively. In this regard, the method preferably includes the step of enabling the detection of the incorporation of each type of naturally occurring nucleotide and discriminating the incorporation of each nucleotide from the incorporation of other types of nucleotides, thereby yielding the nucleotide sequence of the nucleic acid analyte. In other examples, the nucleotides of the labeled nucleotides can be further selected from synthetic nucleotides. In one embodiment, the synthetic nucleotides are selected from 5-methylcytidine, N-methylcytidine, and N6-methyladenosine. Thus, the method may enable the discrimination of the incorporation of the synthetic nucleotides described herein from the incorporation of other types of nucleotides when determining the nucleotide sequence of the nucleic acid analyte.

[0159] The method further includes the step of contacting a nucleic acid analyte, which is a DNA molecule, with an oligonucleotide primer that can specifically hybridize with a region of DNA and initiate synthesis by a nucleic acid polymerase, such as DNA polymerase.

[0160] When used in the context of DNA sequencing methods, the terms “primer” or “oligonucleotide primer” or similar terms are understood to mean an oligonucleotide or nucleic acid fragment capable of specifically hybridizing to a template DNA sequence of a DNA molecule to initiate synthesis using DNA polymerase. Primers may be of any length depending on the specific technique used. However, primers for priming DNA synthesis in DNA polymerase reactions (e.g., PCR) are generally between 10 and 40 nucleotides in length. Those skilled in the art can easily design primers having sequences with appropriate complementarity for specific hybridization to a target template DNA sequence using methods and software known in the art. In some examples, oligonucleotide primers are designed to be substantially complementary to a universal adapter template sequence attached to the 3' end of the template DNA sequence (e.g., by ligation). In other examples, oligonucleotide primers are designed to be substantially complementary to hairpin adapter sequences attached to the 3' and 5' ends of the template DNA sequence (e.g., by ligation). The use of universal primers means that only one primer sequence is needed to initiate DNA synthesis in any number of DNA molecules. Following the example of preparing a library of DNA molecules or fragments for sequencing, each of those DNA molecules or fragments may have a universal adapter template sequence attached to its 3' end (e.g., by ligation) to provide a template for a universal primer that specifically hybridizes.

[0161] When used in the context of hybridization, the terms "selectively" or "specifically" refer to the binding, double-stranding, or hybridization of an oligonucleotide primer to a specific (usually predetermined) site of high affinity in the template DNA sequence, for example, under stringent conditions compared to other (non-specific) sites within the template DNA sequence. Those skilled in the art will understand that specific hybridization between nucleotides usually relies on Watson-Crick pair binding between complementary nucleotide sequences.

[0162] The nucleic acid polymerase is preferably a DNA polymerase. As used herein, the term “DNA polymerase” refers to the enzyme that synthesizes a new strand of DNA from a DNA template strand and a primer hybridized in the 5' to 3' direction. The DNA polymerase “reads” the template in the 3'→5' direction and adds individual nucleotides (bases) to the new or nascent strand in the 5'→3' direction. The polymerase requires the 3'OH group from the primer to initiate the elongation of the new DNA strand. As the new DNA strand is synthesized by the DNA polymerase, individual nucleotides (dNTPs, or dATP, dCTP, dTTP, dGTP, or A, C, T, G) are repeatedly added to the elongating DNA molecule. The specific bases (A, C, T, or G) depend on the sequence of the template DNA, and therefore the new bases hybridize to the nucleotides of the template strand through Watson-Crick interactions. DNA polymerase switches between an "open" and a "closed" conformation. The DNA polymerase is in an open position with the primer-template DNA complex. When a new nucleotide enters the active site, the polymerase switches to a closed position. DNA polymerases are well known in the art, and various types are commercially available. Exemplary DNA polymerase enzymes that can be used in the methods of this disclosure are described herein in the context of plasmon nanoantennas and, unless otherwise specified, are construed to be applicable to any example of sequencing methods described, with necessary modifications.

[0163] As used herein, the term “labeled nucleotide” refers to any nucleotide (including naturally occurring, non-naturally occurring, or “synthetic nucleotides”) to which a fluorophore is directly or indirectly attached. “Fluorophore,” “fluorescently labeled,” or similar terms refer to a signaling moiety of one or more molecules that transmits information through its fluorescent absorption and / or luminescence properties. Such fluorescent properties include fluorescence intensity, fluorescence lifetime, emission spectral characteristics, and energy transfer.

[0164] The fluorophore is bound to the nucleotide by a polyphosphate linker. In one embodiment, the polyphosphate is a triphosphate, tetraphosphate, pentaphosphate, or hexaphosphate. For example, the polyphosphate is a triphosphate. For example, the polyphosphate is a tetraphosphate. For example, the polyphosphate is a pentaphosphate. For example, the polyphosphate is a hexaphosphate.

[0165] In some embodiments, the fluorophore is bound to the phosphate furthest from the nucleotide.

[0166] Examples of fluorophores that can be conjugated or attached to nucleotides used in sequencing methods include, but are not limited to, fluorescein / Oregon Green, fluorescein isothiocyanate (FITC), 6-carboxyfluorescein, tetramethylrhodamine, Texas Red, Dansyl, Alexa Fluor 488, BODIPY FL, Lucifer Yellow, and Alexa Fluor 405 / Cascade Blue fluorophores.

[0167] Examples of commercially available fluorescent nucleotide analogs that can be easily incorporated into DNA molecules during DNA polymerase synthesis include Cy3-dCTP, Cy3-dUTP, Cy5-dCTP, Cy5-dUTP (Amersham Biosciences, Piscataway, NJ), fluorescein-12-dUTP, tetramethylrhodamine-6-dUTP, TEXAS RED™-5-dUTP, CASCADE BLUE™-7-dUTP, BODIPY™FL-14-dUTP, BODIPY™MR-14-dUTP, BODIPY™TR-14-dUTP, RHODAMINE GREEN™-5-dUTP, OREGON GREEN™488-5-dUTP, TEXAS RED™-12-dUTP, and BODIPY™ 630 / 650-14-dUTP, BODIPY™ 650 / 665-14-dUTP, ALEXA FLUOR™ 488-5-dUTP, ALEXA FLUOR™ 532-5-dUTP, ALEXA FLUOR™ 568-5-dUTP, ALEXA FLUOR™ 594-5-dUTP, ALEXA FLUOR™ 546-14-dUTP, Fluorescein-12-UTP, Tetramethylrhodamine-6-UTP, TEXAS RED™ 5-UTP, mCherry, CASCADE BLUE™-7-UTP, BODIPY™ FL-14-UTP, BODIPY™ MR-14-UTP, BODIPY™ TR-14-UTP, RHODAMINE Examples include GREEN(trademark)-5-UTP, ALEXA FLUOR(trademark)488-5-UTP, and ALEXA FLUOR(trademark)546-14-UTP (Molecular Probes, Inc. Eugene, Oreg.). Protocols for the custom synthesis of nucleotides with other fluorophores are available. Henegariu et al., "Custom Fluorescent-Nucleotide Synthesis as an Alternative Method for Nucleic Acid Labeling," Nature Biotechnol. 18:345-348 (2000).

[0168] Other fluorophores available for post-synthesis attachment to dNTPs include, in particular, ALEXA FLUOR® 350, ALEXA FLUOR® 532, ALEXA FLUOR® 546, ALEXA FLUOR® 568, ALEXA FLUOR® 594, ALEXA FLUOR® 647, BODIPY 493 / 503, BODIPY FL, BODIPY R6G, BODIPY 530 / 550, BODIPY TMR, BODIPY 558 / 568, BODIPY 558 / 568, BODIPY 564 / 570, BODIPY 576 / 589, BODIPY 581 / 591, BODIPY 630 / 650, BODIPY 650 / 665, Cascade Blue, Cascade Yellow, Dansyl, Lisamin Rhodamine B, Marina Blue, Oregon Green 488, Oregon Green 514, Pacific Blue, Rhodamine 6G, Rhodamine Green, Rhodamine Red, Tetramethylrhodamine, DYLIGHT® DYES (e.g., DYLIGHT® 405, DYLIGHT® 488, DYLIGHT® 549, DYLIGHT® 594, DYLIGHT® 633, DYLIGHT® 649, DYLIGHT® 680, DYLIGHT® 750, DYLIGHT® 800, etc.) (available from Thermo Fisher Scientific, Rockford, Ill.), Texas Red (available from Molecular Probes, Inc., Eugene, Oreg.), as well as Cy2, Cy3.5, Cy5.5, and Cy7 (Amersham Biosciences, Piscataway, NJ). (Available from the USA and other sources.)

[0169] The fluorophore(s) ligated to the nucleotide may have low quantum efficiency and / or yield. In this regard, low-emitters with low quantum yield can benefit from strong fluorescence enhancement within plasmon resonance hotspots compared to fluorophores with strong emitters. The use of fluorophores with low quantum yield can help reduce background noise from fluorescently labeled nucleotides outside the electric field enhancement area that are not incorporated into the new DNA strand.

[0170] The term "fluorescent signature," as used herein, refers to the cumulative fluorescence properties of a labeled nucleotide when excited during the sequencing process of a nucleic acid analyte described herein, including, but not limited to, peak emission wavelength, fluorescence intensity, and fluorescence duration. Thus, a "distinct fluorescent signature" means that the labeled nucleotide is distinguishable from another fluorescent signature when excited. For example, a "distinct fluorescent signature" as described herein may be a fluorescent signature for a labeled nucleotide that is distinguishable from another labeled nucleotide used in the sequencing method described herein based on differences in one or more of the following, or any combination thereof: differences in peak emission wavelength, differences in fluorescence intensity, differences in fluorescence duration, differences in duration between sequential fluorescence emissions.

[0171] In some embodiments, distinct fluorescence signatures are based on peak emission wavelengths, and based on the difference in peak emission wavelengths, two or more (e.g., two, three, or four) species of labeled nucleotides are distinguishable. In one embodiment, each different species of nucleotide is linked to a fluorophore having a distinct peak emission wavelength. According to any example in which labeled nucleotides are distinguishable based on the difference in peak emission wavelengths, the peak emission wavelengths of the labeled nucleotides are separated from each other by 10 nm or more. For example, the peak emission wavelengths may be separated from each other by about 25 nm or more. For example, the peak emission wavelengths may be separated from each other by about 50 nm or more. For example, the peak emission wavelengths may be separated from each other by about 75 nm or more. For example, the peak emission wavelengths may be separated from each other by about 100 nm or more. In a particular example, each different species of nucleotide is linked to a fluorophore having a distinct peak emission wavelength, and the peak emission wavelengths of each fluorophore are separated from each other by about 100 nm or more.

[0172] Each labeled nucleotide may comprise a fluorophore having a peak emission wavelength independently selected from emission wavelengths in the visible spectrum, the ultraviolet (UV) spectrum, the infrared (IR) spectrum, and the near-infrared spectrum. In one embodiment, one or more fluorophores, or each fluorophore, have a peak emission wavelength independently selected from emission wavelengths between approximately 350 nm and 850 nm.

[0173] Alternatively, or in addition to the above, nucleotides of different species are distinguished from one another based on differences in fluorescence emission intensity. For example, differences in fluorescence intensity can be obtained by differences in the amount of fluorophores (e.g., the same fluorophore or different fluorophores) linked to nucleotides of different species. Following this example, two or more species of labeled nucleotides used in the method can be labeled with the same fluorophore and they can be distinguished based on differences in fluorescence emission intensity. In one embodiment, two or more species of labeled nucleotides used in the method are distinguishable based on differences in fluorescence emission intensity.

[0174] In another example, nucleotides of different species are distinguished from one another based on differences in fluorescence duration. In this regard, it is known that different nucleotides have different residence times when incorporated into the growing DNA strand by DNA polymerase during synthesis. These differences in residence times affect the duration of fluorescence emission from the fluorophore linked to the nucleotide. Therefore, in some embodiments, the distinct fluorescence signatures used to identify nucleotide species during the methods of this disclosure may include fluorescence durations specific to that nucleotide species, or may be unique to a particular nucleotide species in combination with a specific fluorophore. Following this example, two or more species of labeled nucleotides used in the method can be distinguished based on differences in fluorescence duration.

[0175] As described herein, the distinct fluorescence signature of each labeled nucleotide incorporated into the growing DNA sequence by nucleic acid polymerase is enhanced compared to the fluorescence signature of the corresponding labeled nucleotide outside the electric field enhancement area. In one embodiment, the distinct fluorescence signature of the labeled nucleotide incorporated by polymerase is enhanced by a factor of 2 or more (e.g., about 3 or more, or about 4 or more, or about 5 or more, or about 6 or more, or about 7 or more, or about 8 or more, or about 9 or more) compared to the fluorescence signature of the corresponding labeled nucleotide outside the electric field enhancement area. In some embodiments, the fluorescence signature of a separate labeled nucleotide incorporated by polymerase is enhanced by an order of magnitude or more (e.g., by about 10 times or more, or about 20 times or more, or about 30 times or more, or about 40 times or more, or about 50 times or more, or about 60 times or more, or about 70 times or more, or about 80 times or more, or about 90 times or more, or about 100 times or more) compared to the fluorescence signature of the corresponding labeled nucleotide outside the electric field enhancement area.In further embodiments, the distinct fluorescence signature of the labeled nucleotide incorporated by the polymerase is enhanced by at least about 100 times or more (e.g., at least about 150 times, or at least about 200 times, or at least about 250 times, or at least about 300 times, or at least about 350 times, or at least about 400 times, or at least about 450 times, or at least about 500 times, or at least about 550 times, or at least about 600 times, or at least about 650 times, or at least about 700 times, or at least about 750 times, or at least about 800 times, or at least about 850 times, or at least about 900 times, or at least about 950 times, or at least about 1000 times) compared to the fluorescence signature of the corresponding labeled nucleotide outside the electric field enhancement area.

[0176] In some embodiments, the method further includes the step of contacting a nucleic acid polymerase with a nucleic acid analyte and a labeled nucleotide in the presence of one or more quenchers. As used herein, “quencher” or “quencher” refers to any fluorescence-modifying moiety that can attenuate or reduce the light emitted from a fluorophore. This attenuation or reduction of emission is referred to as “quenching.” Therefore, in some embodiments, one or more quenchers may be supplied to help reduce background noise from a fluorophore conjugated to the labeled nucleotide outside the electric field enhancement area. In some embodiments, the quenchers are supplied in a free, unconjugated form. Quenchers that may be useful in the methods of this disclosure include, but are not limited to, Black Hole Quencher Dyes (Biosearch Technologies, e.g., BHQ-0, BHQ-1, BHQ-2, BHQ-3, BHQ-10); QSY Dye fluorescent quenchers (Molecular Probes / Invitrogen), e.g., QSY7, QSY9, QSY21, QSY35; and other quenchers, e.g., Dabeyl and Dabsyl; Cy5Q and Cy7Q and Dark Cyanine dyes (GE Healthcare), which can be used in conjunction with donor fluorescence such as Cy3B, Cy3, or Cy5; DY-Quencher (Dyomics), e.g., DYQ-660 and DYQ-661; and ATTO fluorescent quenchers (ATTO-TEC GmbH), e.g., ATTO 540Q, 580Q, 612Q.

[0177] The term “nucleic acid analyte” refers to any nucleic acid molecule whose sequence is determined using the method of this disclosure. “Nucleic acid” in its broadest form refers to a polymer having multiple nucleotide monomers. A nucleic acid analyte may be single-stranded or double-stranded and may be DNA (e.g., complementary DNA, genomic DNA, or mitochondrial DNA), RNA, or a hybrid polymer (e.g., DNA / RNA). The term “nucleic acid” does not refer to a polymer of any particular length. Rather, a nucleic acid is composed of nucleotides and may be of any length, e.g., longer than about 2 nucleotides, longer than about 10 nucleotides, longer than about 100 nucleotides, longer than about 500 nucleotides, longer than 1000 nucleotides, longer than 10,000 nucleotides, longer than 100,000 nucleotides, longer than about 1,000,000 nucleotides, or longer than that.

[0178] Nucleic acid analytes to be sequenced can be obtained from any source of interest and may include DNA, mRNA, and their mimics, analogs, and derivatives. They can be isolated from cells, cell cultures, tissue samples, bodily fluids, viral samples, genomic nucleic acid samples, cDNA preparations, environmental samples, forensic samples, or synthetic sources. Nucleic acids can be cloned, amplified, transcribed, ligated, fragmented, or otherwise manipulated according to standard methods to obtain nucleic acids to be sequenced, and these manipulations do not render the nucleic acids unsuitable for subsequent sequencing as described herein. It is understood that such nucleic acids may include modified, non-standard, and / or non-natural nucleotides or nucleotide analogs, many of which are described in U.S. Patent Application No. 12 / 945,767, filed November 12, 2010, which is incorporated herein by reference in whole for any purpose. Thus, the methods of this disclosure can be used in a variety of sequencing applications, including, but are not limited to, genome sequencing, transcriptome sequencing, RNA sequencing, single-molecule sequencing, single-cell sequencing, methylation sequencing, and combinations thereof.

[0179] In each of the examples described above, the sequence of the nucleic acid analyte is determined by determining the order of nucleotides in the DNA molecule. In one example, the DNA is genomic DNA. In another example, the DNA is complementary DNA (cDNA) obtained from RNA. Following the example where the nucleic acid analyte is cDNA, the method may include the step of determining the sequence of the RNA based on the sequence of the corresponding cDNA.

[0180] In each of the examples described above, the nucleic acid analyte may be subjected to one or more processing steps before localization with the plasmon nanoantenna. These processing steps may include, but are not limited to, purification steps, reverse transcription (e.g., for RNA), fragmentation to achieve a template DNA of a desired size, enrichment steps (e.g., enrichment of the sequence or architecture of interest), depletion steps (e.g., to remove unwanted sequences), amplification steps (e.g., to increase the amount of template DNA), ligation steps (e.g., to attach the adapter of interest and other sequences), and combinations thereof. The selection of processing steps and the techniques used may vary depending on the application. In some examples, combinations of these processing steps can be used to prepare one or more libraries (e.g., DNA template libraries) for sequencing using the methods of this disclosure. Protocols and methods for preparing nucleic acid samples for sequencing reactions are abundant in the literature, and those skilled in the art can easily determine the necessary steps and methods.

[0181] In some embodiments, the nucleic acid analyte is a double-stranded DNA molecule that can be denatured to produce a single-stranded DNA molecule (i.e., “target DNA sequence,” “template DNA,” or “template DNA sequence”) that the DNA polymerase enzyme can use to synthesize a new strand. The terms “target DNA” or “template DNA” refer to a sequence of DNA that can be sequenced according to this method. This term encompasses both complementary strands of the target DNA molecule. As will be understood by those skilled in the art, from the well-known Watson-Crick base pairing, the sequence of one strand of the target DNA reveals the sequence of the other strand. For example, one or more oligonucleotide primers or adapters can be attached to the target DNA to facilitate synthesis. However, in other embodiments, the double-stranded DNA molecule is not denatured, but rather retains its double-stranded form.

[0182] Sequence-determinable DNA molecules (including cDNA, single-stranded DNA, and double-stranded DNA) can be fragmented or sheared into various sizes (0.5kb–10kb) using hydrodynamic mechanical shear with a relatively narrow size distribution (Thorstenson et al., Genome Res 8, 848-855 (1998); Roe, Methods Mol Biol 255, 171-187 (2004)). Methods for fragmenting and / or shearing DNA are known in the art, including, for example, sonication, spraying, and enzymatic methods. Specialized devices for size-limited DNA shearing are commercially available (e.g., Bioruptor® from Diagenode). These methods are further described in Sambrook (Molecular Cloning: A Laboratory Manual. New York: Cold Spring Harbor Laboratory Press; 1989) and Ausubel (Current Protocols in Molecular Biology. New York: John Wily; 2001), which are incorporated herein by reference in their entirety for all purposes.

[0183] The sequenced DNA molecule may be of any length. For example, the sequenced DNA molecule may be 10 nucleotides or more in length, e.g., about 20 nucleotides, or about 50 nucleotides, or about 100 nucleotides, or about 200 nucleotides, or about 300 nucleotides, or about 400 nucleotides, or about 500 nucleotides, or about 1000 nucleotides, or longer (e.g., 2000, or 5000, or 10,000 or more nucleotides). In some embodiments, the sequenced DNA molecule is about 10 to 10000 nucleotides long. In some embodiments, the sequenced DNA molecule is about 100 to 5000 nucleotides long. In some embodiments, the sequenced DNA molecule is about 200 to 2000 nucleotides long. In some embodiments, the sequenced DNA molecule is about 400 to 1000 nucleotides long. As described herein, in some embodiments, large DNA molecules can be fragmented or sheared to various smaller sizes to facilitate sequencing.

[0184] The sequenced DNA molecule includes one or more oligonucleotide primers and one or more adapter sequences that hybridize with DNA polymerase and provide a binding site for initiating synthesis by DNA polymerase. Therefore, the method may include a further step of ligating adapters to one or more sequenced DNA molecules (e.g., a library thereof) using the method described herein. For example, the method may include ligating a universal adapter sequence to the 3' end of a sequenced DNA molecule. In some embodiments, the method may include ligating hairpin adapters to both ends of a double-stranded DNA molecule to obtain a sequenced template that includes both the complementary and non-complementary strands of the DNA molecule as single-stranded circular constructs, which can be repeatedly sequenced to obtain overlapping sequenced information from both strands. With regard to single-molecule sequencing reactions, such as Pacific Biosciences' SMRT® Sequencing, where sequence data is generated from a single template molecule, statistical analysis of overlapping information is used to generate a consensus sequence to a target region from a single sequenced template. Further details regarding overlapping and annular sequencing templates are presented, for example, in U.S. Patents 7,476,503 and 8,153,375, both of which are incorporated herein by reference in their entirety for all purposes.

[0185] In other examples, the method may include the step of ligating a hairpin adapter to only one end of a double-stranded DNA molecule. By ligating a hairpin adapter to only one end of a double-stranded DNA molecule, a sequencing template containing both the complementary and non-complementary strands of the DNA molecule is provided as a single-stranded linear construct, which can be sequenced to obtain sequence reads from each strand. Such templates are particularly useful in sequencing techniques where single-stranded linear templates are preferred, for example, in sequencing techniques using nanopore-based sensors, which have been described in detail in the art and are currently being developed by several companies, including Oxford Nanopore and Genia. However, the use of templates with stem-loop adapters at both ends is also intended with respect to sequencing in nanopore-based methods, for example, when using a single-stranded circle resulting from separating the complementary strand as a template for rolling circle replication, for example, when a nascent strand or released phosphate group is directed to or passes through the nanopore.

[0186] In each of the examples described above, step (II) of the method for sequencing the nucleic acid analyte includes detecting the sequence of enhanced distinct fluorescence signatures as the labeled nucleotides are sequentially incorporated by nucleic acid (e.g., DNA) polymerase into a polynucleotide (i.e., complementary strand) having a sequence complementary to the polynucleotide sequence of the nucleic acid analyte. The enhanced distinct fluorescence signatures can be detected by any means known in the art for detecting fluorescence, including, but not limited to, confocal microscopy, confocal laser scanning microscopy, internal total internal reflection (TIR), total internal reflection illumination fluorescence (TIRF), epifluorescence microscopy, near-field scanning microscopy, far-field confocal microscopy, wide-field epifluorescence, light scattering, dark-field microscopy, photoconversion, wide-field fluorescence, single-photon and / or multiphoton excitation, spectral wavelength discrimination, evanescent light illumination, scanning two-photon, scanning wide-field two-photon, Nipkow spinning disk, and / or multifocal multiphoton. In some examples, a combination of these methods is used for detecting fluorescence. In one embodiment, a confocal laser scanning microscope is used to detect the enhanced, distinct fluorescence signature of a nucleotide as it is incorporated into the complementary strand by DNA polymerase. In another embodiment, internal total internal reflection (TIR) ​​is used to detect the enhanced, distinct fluorescence signature of a nucleotide as it is incorporated into the complementary strand by DNA polymerase. In yet another embodiment, total internal reflection illumination fluorescence (TIRF) is used to detect the enhanced, distinct fluorescence signature of a nucleotide as it is incorporated into the complementary strand by DNA polymerase.

[0187] Distinct fluorescence signatures emitted from different species of labeled nucleotides can be separated using fluorescence resonance energy transfer measurements; photoconversion; fluorescence lifetime measurements; polarization; fluorescence lifetime determination; correlation / anticorrelation analysis; Raman; intensity; ratiometric; time-resolved methods; anisotropy; near-field or far-field microscopy; photobleaching-after fluorescence recovery (1-RAP); spectral wavelength discrimination; fluorescence lifetime measurement and separation; fluorophore identification; background suppression, parallel multicolor imaging, or any suitable discrimination method based on any combination thereof. See, for example, JR Lakowitz 2006, "Principles of Fluorescence Spectroscopy", Third Edition. When different nucleotides are labeled with different energy transfer or reporter moieties, separation of the emitted signals can be used to distinguish between different nucleotides that bind to and / or are incorporated by polymerase. In one particular example, each labeled nucleotide emits a fluorescence signal at a different wavelength, and thus the distinct fluorescence signatures of nucleotide species are identified by their emission wavelength, e.g., peak emission wavelength.

[0188] To detect distinct fluorescence signatures emitted from each of different species of labeled nucleotides, the method may further utilize a multifluorescence imaging system capable of detecting multiple distinct fluorescence signatures emitted from different species of labeled nucleotides during the sequencing reaction. Such a system may include a combination of special filters for each excitation line and / or each emission band. In one embodiment, the detection system includes tunable excitation and / or tunable emission fluorescence imaging.

[0189] When the method of the present disclosure is implemented with an array, for example, an array of plasmon nanoantennas as described herein, the detection system may include an optical train that guides signals emitted from the organized array to different locations on an array-based detector to detect multiple optical signals from multiple locations. The optical train generally includes an optical grating and / or wedge prisms to simultaneously guide and separate signals having different spectral characteristics from different addressable locations on the array to different locations on an array-based detector, for example, a CCD.

[0190] After detecting the sequence of separate fluorescence signatures in step (II), the sequence of the nucleic acid analyte can be determined by correlating the sequence of separate fluorescence signatures with the sequence of nucleotides sequentially incorporated into the synthesized DNA.

[0191] kit This disclosure also provides kits and reaction mixtures for performing single nucleic acid molecular sequencing as described herein. The kits, while their components and their configurations may vary, generally include a plasmon nanoantenna as described herein or an array of such plasmon nanoantennas attached to a substrate, and a reaction mixture. The reaction mixture may include one or more fluorescently labeled nucleotides (e.g., each of dATP, dCTP, dGTP, and dTTP as described herein), one or more buffers (e.g., Tris), various salts (e.g., KCl, NaCl, (NH4)2SO4, MnCl2, Zn salts, MgCl2), and often stabilizers, surfactants, DMSO, and DTT. The reaction mixtures of this disclosure may also include additives to increase the specificity and efficiency of the polymerase reaction. The reaction mixtures may include oligonucleotide primers for primer DNA synthesis by DNA polymerase. In the case of universal primers that hybridize with a universal adapter sequence, the primers may be provided in the reaction mixture (though not required). In the case of custom-made primer sequences, it may be preferable that the primers be provided separately, for example, by the user. It will be understood that the kits of this disclosure may encompass any combination of the above components.

[0192] The kits of this disclosure may also include instructions for carrying out the method for determining the sequence of a single DNA molecule as described herein.

[0193] Nothing relating to any documents, acts, materials, devices, articles, or similar matters contained herein should be construed as meaning that any or all of these matters constituted part of the fundamental principles of the prior art or were common knowledge in the art relating to this disclosure prior to the respective priority dates of the attached claims. [Examples]

[0194] (Example 1) General materials and methods Design and synthesis of DNA origami The inventors used caDNAno to design a DNA staple sequence for folding an M13mp18 single-stranded DNA template to create a DNA origami nanospanno scaffold.

[0195] FEM simulation for estimating fluorescence enhancement All simulations are performed using the Boundary Element Method (MNPBEM) in MATLAB® 2020b. 22 The simulation of metal nanoparticles was performed using the toolbox. The optical properties of silver and gold were analyzed using Johnson and Christy's tools. 23 The background medium dielectric constant was set to 1.77 to match the experimental aqueous buffer solution. To simulate measurements with a total internal reflection fluorescence (TIRF) microscope, a glass substrate layer with a dielectric constant of 2.25 was placed 5 nm below the nanoparticles on the z axis, and an incident plane wave was propagated at an angle of 65° to the perpendicular to the glass layer. The emission characteristics of the dye were modeled using a vibrating dipole with a frequency corresponding to the dye's maximum emission wavelength.

[0196] DNA origami synthesis The synthesis of DNA origami requires folding a single long template DNA strand, composed of a single-stranded phage vector M13mp18, by hybridizing it with approximately 250 short "staple" DNA strands that crossbridge the DNA helix at precisely defined locations, thereby forming the desired shape. 21、24 The synthesis requires an annealing process to prevent the formation of undesirable kinetically trapped structures and to maximize the yield of well-formed structures consistent with the design. The synthesis conditions were optimized to the following: 1× Origami buffer (10 mM Tris, pH 8 and 1 mM EDTA) + 20 mM magnesium chloride (MgCl2), 15-fold excess of staples (50-fold excess of extensional staples), annealing for 22 hours at a favorable temperature range of 48–43°C. Details of the temperature gradient of the annealing reaction are shown in Table 1 below. Table 1. Temperature gradient for DNA origami synthesis [Table 1]

[0197] Agarose gel electrophoresis 10 μL of a 5 nM sample (DNA origami or M13mp18 single-strand template) or molecular weight standard was mixed with 2 μL of loading dye, and this was loaded onto an agarose gel prepared by dissolving 1.125–3 g of agarose in 150 mL of buffer, with or without 7.5 μL of RedSafe dye. Electrophoresis was performed over 3 hours at 70 V in pre-cooled 1×TAE buffer with 6 mM MgCl2. The gel was then scanned using Alexa Fluorophore 647 / 488 if the sample was labeled with Alexa Fluorophore, or with RedSafe dye if RedSafe was added to the gel. After staining with SYBRgold, the gel was scanned again using SYBRgold.

[0198] Purification of DNA origami DNA origami was purified either by extraction of the purified sample from an agarose electrophoresis gel, or by polyethylene glycol (PEG) precipitation.

[0199] Purification by gel extraction A 0.75% agarose gel was prepared by dissolving 1.125 g of agarose in 150 mL of 1×TAE buffer containing 11 mM MgCl2, and RedSafe stain was added for visualization. The sample, mixed with the loading dye, was loaded onto the gel, and electrophoresis was performed on ice at 70 V for 2.5 hours. After electrophoresis, the DNA origami bands were cut off with a clean blade and transferred to a 500 μL Eppendorf tube, which was then centrifuged at maximum speed for 5 minutes to pulverize the gel. The gel slurry was then transferred to a Freeze "N" Squeeze column and centrifuged at 4°C at maximum rcf for 10 minutes. Finally, the purified DNA origami was collected in the collection tube provided with the column.

[0200] Purification by polyethylene glycol precipitation (PEG) Unpurified DNA origami samples were pipetted into DNA-lowbind tubes and diluted to 400 μL with pre-PEG buffer (1× origami buffer and 20 mM MgCl2, filtered through a 0.45 μm syringe filter). 400 μL of 2× PEG precipitation buffer (15% (g / mL) PEG8000, 10 mM Tris, pH 8, 1 mM EDTA, 500 mM NaCl) was added, and the samples were centrifuged at 20°C and 21100 RCF for 25 minutes. Immediately after centrifugation, the supernatant was removed, and the samples were resuspended in 1× origami buffer with 6 mM MgCl2 (also filtered through a 0.45 μm syringe filter). After resuspending in 50 μL of 1× origami buffer with 6 mM MgCl2, the concentration was quantified using Nanodrop. The concentration in nM units is obtained by dividing the concentration in ng / μl units by 4.8.

[0201] Determining the dimensions of DNA origami To accurately determine the dimensions of the NanoSpanno, single-particle averaging (SPA) was performed using RELION. All images used for SPA were acquired using a Tecnai microscope at a consistent magnification of 19,000–38,000x.

[0202] Grid preparation A carbon / formvar grid was glow-discharged, and a droplet of 2% uranyl acetate (or uranyl formate) stain was placed on a piece of Parafilm. A 5 μL droplet of 1 nM sample (undiluted from gel purification) was applied to the dark, glossy side of the grid, and immediately absorbed from the grid by touching the edge of the grid with a piece of filter paper. The grid (dark, glossy side) was then brought into contact with the stain droplet, and the droplet was immediately absorbed with a piece of filter paper. The grid was then air-dried for several minutes.

[0203] Imaging conditions All images used in SPA were collected using a Tecnai microscope at a consistent magnification of 19,000 to 38,000x.

[0204] 2D particle averaging To accurately determine the dimensions of NanoSpanno, single-particle averaging (SPA) was performed using RELION.

[0205] Preparation of grids for cryoTEM To prepare the sample for observation using a cryo-electron microscope, an aqueous solution was applied to a glow-discharge treated (hydrophilic) perforated carbon film supported by an EM grid. Excess solution was carefully removed from one or both sides using filter paper. The blotted grid was then rapidly immersed in a cryogen pre-cooled to the temperature of liquid nitrogen. This rapid freezing process embeds biomolecules into a thin, amorphous ice film, making them observable with a cryo-electron microscope.

[0206] Imaging using cryo-TEM The frozen grid is placed in a cryotransfer holder equipped with a liquid nitrogen dewar in a cryoworkstation to avoid ice contamination. The ice-embedded specimen is then loaded into the cryo-electron microscope in a frozen state without contamination using the cryotransfer holder. Images of the holes are acquired using low-dose mode. The focus is adjusted, and then the target exposure is obtained at the desired preset magnification. DNA coating of spherical gold nanoparticles Materials used for DNA coating of gold nanoparticles • 100nm gold nanoparticles (AuNP) purchased from Nanopartz • Thiolized DNA purchased from IDT (Coralville USA) • Phosphate buffer [Prepared: 980 μL of MQ water mixed with 10 μL of Tween® 20 (10%) and 10 μL of potassium phosphate (a 4:5 mixture of 1 M KH2PO4 and 1 M K2HPO4)] • 10%Tween (Registered Trademark) 20 • 20 mM TCEP: 5.732 mg / mL (in MQW, adjusted to pH 3.0)

[0207] Preparation of thiolated DNA 25 μM thiol-DNA in 10 mM TCEP was prepared by adding 0.5 μL of monothiol-DNA (25AC, 500 μM in Milli-Q water) + 5 μL of TCEP (20 mM, pH=3) + 4.5 μL of Milli-Q water. This solution was incubated at room temperature for 1 hour. Then, 25 μL of Milli-Q water was added to obtain a total volume of 35 μL.

[0208] Coating of gold nanoparticles with thiolated DNA One mL of the nanoparticle solution (10 pM, with 10 μL of 5× phosphate buffer added) was centrifuged at 4500 RCF for 5 minutes. The supernatant was pipetteed and mixed with 35 μL of TCEP-treated DNA, then diluted and mixed in 10 μL of 5× phosphate buffer (total volume 50 μL). The nanoparticle-oligonucleotide mixture was then frozen at -20°C for 2 hours and then allowed to stand at room temperature. The final mixture consisted of gold nanoparticles with an oligonucleotide concentration of 0.2 nM and 5 μM of DNA in a 25,000-fold molar excess.

[0209] Excess DNA was removed from DNA-coated gold nanoparticles by repeated precipitation and supernatant removal. The mixture was centrifuged at 4500 RCF for 5 minutes, the supernatant was pipetted, and the particle pellet was diluted in 10 μL of 5× phosphate buffer, and then in 1 mL of MQW. The precipitation process was repeated 5 times to completely purify the nanoparticles from free oligonucleotides. Protein expression and purification material • 1000 x Ampicillin: 100 mg / ml (1 g / 10 mL) ·LB agar LB Bros • 1000 x IPTG: 1M (0.238 g / mL) • Krenow buffer: 10 mM Tris HCl, pH 8, 50 mM NaCl, 10 mM MgCl2, and 1 mM DTT • Phi29 buffer: 50 mM Tris HCl, pH 7.5, 50 mM NaCl, 10 mM MgCl2, and 4 mM DTT • Taq: 10mM Tris HCl, pH 8, 50mM NaCl, 1.5mM MgCl2 1M Imidazole 120mL • 500 mL of solubility buffer: 20 mM Tris, pH 7.5, 300 mM NaCl (420 mL for washing buffer) • Washing buffer 500ml: 20mM Tris, pH 7.5, 300mM NaCl, 20mM Imidazole • Elution buffer 200 mL: 20 mM Tris, pH 7.5, 300 mM NaCl, 500 mM Imidazole SEC buffer 1L: 20mM Tris, pH 7.5, 150mM NaCl, 1mM DTT • SEC buffer 100 mL (after fluorophore labeling): 20 mM Tris, pH 7.5, 150 mM NaCl • Dialysis buffer: 20mM Tris, pH 7.5, 150mM NaCl 0.1M TCEP, pH 7 • Alexa647 fluorophores in 50 nanomolar aliquots (resuspended in 10 μL of DMSO at a stock concentration of 5 mM)

[0210] Plasmid description Three petDuet-1 plasmids were ordered, each containing one of the recombinant polymerases: Krenow-SpyC, Phi29-SpyC, and Taq-SpyC (Genscript). Each recombinant polymerase construct was cloned into the petDuet-1 vector for expression in E. coli cells.

[0211] Transformation of T7 E. coli using expression plasmids LB agar plates containing ampicillin were prepared by heating 300 ml of Luria broth (LB) agar to a liquid state and then cooling it to room temperature. Before solidification, 300 μL of 100 mg / ml ampicillin was added to the LB agar to obtain a final concentration of 100 μg / ml ampicillin. The LB agar was then poured into a sterile petri dish and allowed to solidify.

[0212] 1 μL of plasmid was used to transform 20 μL of T7 E. coli-expressing cells. The cells were first incubated on ice for 30 minutes, and then 1 μL of the expression plasmid was added. The mixture was then incubated in a 42°C water bath for 10 seconds, and then returned to ice for 5 minutes. 20 μL of the transformed cells were spread onto a Luria broth (LB) agar plate containing 100 μg / ml ampicillin and incubated overnight at 37°C. Multiple single colonies containing the expression plasmid were observed.

[0213] Protein expression A single E. coli colony was placed in 100 ml of starter culture containing 100 μg / ml ampicillin and incubated overnight at 37°C. Approximately 25 ml of the starter culture was added to 2 L of LB broth containing 100 μg / ml ampicillin. The culture was grown on a shaking platform at 37°C and 180 rpm until the OD600 reached 0.5–0.6. The temperature was then reduced to 18°C, and IPTG was added to a final concentration of 1 mM to induce protein expression. The cells were then left overnight on an orbital shaker rotating at 180 rpm to produce protein.

[0214] Next, the cells were centrifuged at 4°C and 6000 RCF for 20 minutes (centrifuge VX22N, rotor R9A2, VWR). The cell pellet can be stored at -80°C for later use. The cell pellet was resuspended in 80 mL of lysis buffer (+1 Complete EDTA tablet + DNAse1), and the cells were lysed by sonication three times for 3 minutes each using a 50% duty cycle, power 7, 3. Proteins expressed in the cytoplasm were extracted by removing the solid material by centrifugation. The sample was centrifuged at 4°C and 3,600 RCF for 30 minutes. The pellet was discarded, and the cell lysate supernatant was collected for purification (centrifuge VX22N, rotor R9A2, VWR).

[0215] Protein purification by immobilized metal affinity purification (IMAC) Immobilized metal affinity chromatography (IMAC) was performed using a 2 × 5 mL HisTrap Fast Flow Crude column. The HisTrap column was first equilibrated with wash buffer (30 mL, 3 × column volume), then all contaminants were removed with 30 mL of elution buffer, and the column was re-equalized with 30 mL of wash buffer before the cell lysates were loaded.

[0216] Approximately 75 ml of cell lysate was filtered through a 0.22 μm membrane and then loaded onto a HisTrap column at a flow rate of 1 ml / min. The HisTrap column was then washed with wash buffer (20 mL), and the bound proteins were eluted. Gradient elution was performed over 10 column volumes with imidazole concentrations from 20 mM to 500 mM at a flow rate of 2 ml / min, and 2 ml fractions were collected.

[0217] Protein purification by size exclusion chromatography (SEC) The eluted fractions containing recombinant polymerase enzyme, purified by IMAC, were combined and concentrated to 5 mL using a 30KD Amicon Ultra Centrifugal Filter Unit, and further purified using SEC. SEC was performed using a HiLoad 16 / 600 Superdex 200pg column (GE Healthcare). The column was equilibrated with MQW (approximately 150 mL, 1.2 × column volume, 0.9 mL / min, <0.5 MPa). The SEC column was equilibrated with SEC buffer (approximately 150 mL, 1.2 × column volume, 1 mL / min, <0.5 MPa). Protein was eluted from the SEC column using SEC buffer (1.2 × column volume, 150 mL, <5 mL injection). The eluted fraction was passed through an SDS-PAGE gel as described above to confirm protein purity. The fraction containing pure polymerase was pooled, and the final concentration was determined by Denovix. Recombinant polymerase was stored at -80°C until needed.

[0218] Covalent attachment of fluorescent dyes to proteins The purified protein was dialyzed to remove DTT. 1 mL of approximately 80 μM protein was added to a dialysis button using a 6 kD cutoff snakeskin. The button was placed in 1 L of dialysis buffer and left overnight at 4°C with constant agitation, and the buffer was changed once. After dialysis, the concentration was determined by IR absorption using a Denovix instrument.

[0219] 400 μL of 50 μM dialyzed protein was added with 10 μL of TCEP (20 mM, 10 μL), and left on ice for about 30 minutes. Then the protein was labeled with a 5-fold molar excess of Alexa-647 maleimide dye. 400 μL of the protein with TCEP was added to 20 μL of a DMSO solution containing 100 nanomolar Alexa-647 dye (5-fold excess). The mixture was incubated overnight in the dark on a rotating wheel. Then the excess dye was purified, and the buffer was replaced with 20 mM Tris, pH 7.5, 150 mM NaCl using SEC.

[0220] SDS-PAGE A 4-12% SDS-PAGE gel was purchased from Invitrogen. Samples were run at 180 V for 30 minutes using 1×MES buffer. The protein molecular weight (MW) standard used in all SDS PAGE gels was SeeBlue Plus2 Pre-Stained Protein Standard. The gel was post-stained with Blue and imaged under UV light to identify which fractions contained pure protein.

[0221] High Performance Liquid Chromatography (HPLC) HPLC purification of maleimide DNA, DNA-peptide conjugates, and fluorophore-labeled dNTPs was performed using a Shimadzu LC-20AT HPLC system equipped with an Atlantis RP-HPLC C18 column (150 mm × 4.6 mm, 5 μm). Fractions were automatically collected, and fractions containing pure product were subjected to evaporation. The resulting residue was dissolved in water and quantified using a Denovix instrument.

[0222] HPLC method for maleimide DNA and DNA-peptide conjugates The sample was diluted with 0.1 M TEAA + 5% ACN, and 100 μL was injected into the HPLC. The HPLC buffer consisted of HPLC buffer A containing 0.1 M TEAA + 5% ACN, and HPLC buffer B consisting of 0.1 M TEAA + 70% ACN. The HPLC procedure was performed as follows: The sample was injected at t=0 minutes, and the buffer was flowed at a rate of 1 mL / min. The schedule was to pump in 0% of buffer B for 1 minute, then increase the concentration from 1 minute to 40 minutes, up to 100% of buffer B, and maintain it at 100% until 43 minutes. Then, decrease the concentration to 0% of buffer B at 43.01 minutes and maintain it until 48 minutes.

[0223] HPLC method for fluorophore-labeled dNTPs The sample was diluted with 0.1 M TEAA, and a volume of 100 μL was injected. The buffers used in the HPLC method were HPLC buffer A, consisting of 0.1 M TEAA, and HPLC buffer B, consisting of 100% ACN. The procedure involved injecting the sample at t=0 minutes and performing an HPLC method in which the buffers were flowed at a rate of 1 mL per minute according to a specific schedule. The schedule involved pumping in 2% of buffer B for 1 minute, then increasing the concentration to 7% of buffer B from 1 minute to 15 minutes, then increasing the concentration to 30% of buffer B from 15 minutes to 45 minutes, and finally increasing the concentration to 60% of buffer B from 45 minutes to 75 minutes, and maintaining it at 60% until 80 minutes. After 80.01 minutes, the concentration was returned to 2% of buffer B and maintained for 5.00 minutes until stopped by the controller at 85.01 minutes.

[0224] Synthesis of DNA-peptide conjugates maleimide-DNA synthesis and purification To synthesize maleimide-DNA, 2 mg of SMCC was resuspended in 300 μL of DMF, and 10 μL of DIPEA was added. Then, 5 μL of 2 mM aminated DNA (10 nmol) was added to the solution, and it was incubated on an orbital shaker at room temperature at 600 rpm for 45 minutes.

[0225] Next, SMCC-DNA was extracted by ethanol precipitation. The solution was divided into three approximately 105 μL aliquots, and 100 μL of Milli-Q water, 20 μL of 3M sodium acetate, pH 5.2, and 800 μL of ice-cold 100% EtOH were added to each tube and incubated at -80°C for 45 minutes. The DNA precipitate was pelletized by centrifugation at 4°C and 25000 RCF for 45 minutes, and the supernatant was removed. The pellets were dried (in an oven at 37°C for approximately 15 minutes) and stored at -20°C.

[0226] The final purification step was performed using high-performance liquid chromatography (HPLC). The pellet was resuspended in 100 μL of MilliQ water, and any undissolved precipitate was removed by centrifugation and filtration. The supernatant was loaded into the HPLC using Method 2.10.1, and eluted using an acetonitrile gradient. Generally, 5'-maleimide-15 nucleotide DNA and 3'-maleimide-15 nucleotide DNA elute in approximately 15 minutes, while 5'Cy5-3'maleimide-15 nucleotide DNA elutes in approximately 22 minutes. The fraction containing the pure product was evaporated, the residue was dissolved in water, and quantified using UV-Vis spectroscopy. The product was then divided into 100 pmol / tube portions and resuspended in 10 μL of MQW to create a 10 μM working stock. The sample was then dried using a SpeedyVac and stored at -20°C.

[0227] Conjugation with maleimide DNA peptide conjugates Maleimide-DNA was resuspended in 80 μl of MilliQ water, 10 μl of 200 mM Tris, pH 7.5, and 1.5 M NaCl. Then, 10 μl of 1 mM cysteine-containing peptide was added, and the mixture was incubated at room temperature for 2 hours, followed by purification by HPLC. 100 μl of the sample was loaded into the HPLC as described in the section titled "HPLC method for maleimide DNA and DNA-peptide conjugates" in Example 1.

[0228] Generally, 5'-SpyTag-15 nucleotide DNA and 3'-SpyTag-15 nucleotide DNA elute in approximately 17 minutes, while 5'Cy5-3'maleimide-15 nucleotide DNA elutes in approximately 20 minutes. The fraction containing the pure product was evaporated, the residue was dissolved in water, and quantified using UV-Vis spectroscopy. The product was then divided into 100 pmol / tube portions and resuspended in 10 μL of MQW to create a 10 μM working stock. The sample was then dried using Speedyvac and stored at -20°C.

[0229] Synthesis of DNA-protein conjugates Next, purified SpyTag-DNA was bound to purified polymerase-SpyCatcher to obtain a polymerase enzyme covalently linked to a DNA strand having a desired sequence, enabling specific immobilization of DNA origami nanoantennas into the cavities.

[0230] The buffering conditions are as follows: • Krenow 10x reaction buffer: 100mM Tris, pH 8, 0.5M NaCl, 100mM MgCl2, 10mM DTT • Binding and elution buffers: • 1M imidazole solution (68 mg / mL) • Binding and washing buffer: 20 mM Tris, pH 7.5, 300 mM NaCl, 10 mM Imidazole • Elution buffer: 20 mM Tris, pH 7.5, 300 mM NaCl, 500 mM Imidazole

[0231] SpyTag-DNA and SpyCatcher-protein were incubated at room temperature for 2 hours, followed by a reaction with 70 μL of SpyTag-DNA in a 2x molar excess relative to the SpyCatcher-protein. The reaction was carried out in separate reaction buffers for the SpyCatcher-protein. Excess SpyTag-DNA was removed by purifying the protein using IMAC. The mixture was diluted to one-fifth by adding 280 μL of binding buffer (final reaction volume = 350 μL). Tris nickel-NTA (NiNTA) beads were prepared as follows: the supernatant was removed from 100 μL of the bead slurry and washed three times with 300 μL of MilliQ water (centrifugation speed and time: 500 × g for 30 seconds). The beads were then equilibrated by exchanging 300 μL of binding and washing buffer twice. The solution containing SpyTag-DNA and polymerase-SpyCatcher was added to the NiNTA beads and incubated on a rotating platform at 4°C for 2 hours. The supernatant was removed by centrifugation (500 × g for 30 seconds), and the beads were washed three times with 300 μl of washing buffer (centrifugation speed and time: 500 × g for 30 seconds) to remove excess SpyTag-DNA. After incubation with 70 μl of elution buffer on a rotator at 4°C for 2 hours, polymerase-DNA was recovered from the supernatant NiNTA beads. The sample buffer was then replaced with the desired buffer for assembly on the DNA origami scaffold using a desalted spin SEC column, and the purity of the sample was evaluated using SDS-PAGE (where proteins were visualized using protein-specific staining and fluorescence signals from covalently attached fluorophores).

[0232] DNA polymerase activity assay DNA polymerase function was evaluated by measuring the extension of a fluorescently labeled primer bound to a target DNA strand. The reaction conditions are summarized below: [Table 5]

[0233] The reaction was stopped by adding 0.5 μl of 0.5 M EDTA, and then heat denaturation was performed at 95 °C for 15 minutes. The elongation of the primer by DNA polymerase was visualized by native PAGE.

[0234] Native PAGE It was run at 150 V for 10 minutes and then at 250 V for 45 minutes (until the dye reached 1.5 cm from the bottom) on a 15% Tris-Glycine Native-Polyacrylamide Gel (using a 5% stacking gel).

[0235] Assembly of gold nanoparticles on a DNA origami scaffold DNA-coated gold nanoparticles were resuspended in 1× TAE containing 6 mM MgCl₂ and mixed with purified DNA origami having a single-stranded DNA extension complementary to the thiolated DNA conjugated to the gold nanoparticles at 500 pM.

[0236] Assembly of DNA polymerase enzyme on a DNA origami scaffold 100 nM of polymerase-DNA conjugate and 20 nM of DNA origami scaffold were incubated at room temperature for 2 hours in a 10 μL reaction volume. The reaction buffer was 10 mM Tris HCl, pH 8, 50 mM NaCl, and 10 mM MgCl₂. Excess polymerase-DNA was removed using SEC as follows: · An S300 column containing 500 μL of slurry was prepared for each complex · The resin was rotated at 1000 g for 2 minutes once · The resin was washed 4 times with 500 μL of MQW at 1000 g for 1 minute · The resin was washed 3 times with 500 μL of imaging buffer at 1000 g for 1.5 minutes · The resin was rotated at 1000 g for 1.5 minutes · Approximately 10 μL was loaded onto a 1× S300 column · It was rotated at 1000 g for 4 minutes The concentration in nM units is the concentration in ng / μl units divided by 4.8.

[0237] Surface plasmon resonance (SPR) The experiments were performed using a Biacore S200 instrument (GE Healthcare Life Sciences). All experiments were conducted at room temperature using Krenow reaction buffer at a flow rate of 10 μL / min. The CM3 sensor tip was coupled to streptavidin to near saturation using an amine coupling kit (GE Healthcare Life Sciences) (typically between 4000 RU and 7000 RU). After streptavidin coupling, biotinylated DNA strands were injected into the reference flow cell and the experimental flow cell, and the excess biotin-binding sites in the Krenow reaction buffer were blocked with biotin. The DNA template strand was then loaded into the experimental flow cell. Experiments were performed using two flow cells on one tip, with the remaining flow cell serving as a reference cell without a DNA template. The surface was then conditioned by injecting Krenow reaction buffer twice, and then DNA polymerase at the indicated concentrations was injected. The data shown is after subtracting references, and all data has been fitted using MATLAB® 2020b.

[0238] Single-molecule total internal reflection fluorescence (TIRF) microscope General settings Microfluidic devices and coverslips functionalized with BSA-biotin and streptavidin were prepared as previously described. 25 The sample was added to each channel at a concentration of 10 pM in imaging buffer, and then washed with imaging buffer. Images were taken by McGuinness et al. 26 The custom-made TIRF microscope described above has an output density of approximately 1-3 W / cm². -2 The data was collected using (measured with the objective lens, with the laser beam perpendicular to the surface of the cover glass).

[0239] Single-particle photobleaching Single-particle photobleaching was performed in DNA origami imaging buffer using purified, pre-assembled DNA origami scaffolds and DNA polymerase, immobilized on coverslips to a density of approximately 1000 particles per field of view (FOV). Images were acquired using excitation at 488 nm (50 mW) and 647 nm (20 mW) with an exposure time of 200 ms and 200 frames per FOV. Three independent repeats of 20 FOVs were obtained. Images were analyzed using JIM-Immobilized-Microscopy-Suite (https: / / github.com / lilbutsa / JIM-Immobilized-Microscopy-Suite) to determine the height of single photobleaching steps and their corresponding steps.

[0240] Single-particle photobleaching was performed to measure fluorescence enhancement in the presence of metal nanoparticles. For single-particle photobleaching, a purified, pre-assembled DNA origami scaffold containing an Alexa-647 fluorophore within the hotspot region was used (Figure 1). This scaffold was then incubated for 2 hours in DNA origami imaging buffer at a concentration of 100 pM, along with a 100 nm AuNP coated with 500 pM DNA, and subsequently immobilized on a coverslip. Image acquisition was performed as described above.

[0241] Transient binding of fluorescent DNA strands to plasmon nanoantennas To measure the fluorescence enhancement of multiple individual fluorophores over time, a docking strand containing a 7-base pair ssDNA binding site (TCCTCCT)27 was incorporated into the hotspot region of a DNA origami scaffold. The DNA origami scaffold was purified, assembled with AuNPs, and immobilized on a coverslip as described above. Subsequently, a 1 nM imaging strand (Alexa647-AGGAGGA) was added, and 5000 frames were imaged at an exposure time of 200 ms and a laser power of 20 mW.

[0242] Measurement of gold scattering intensity with respect to polarization angle Variations in incident polarization were achieved by using a polarizing filter in the excitation light path and then passing the light through a zero-order half-wave plate. Here, the zero-order half-wave plate was rotated in 10° increments to achieve variations in polarization angle in 20° increments. Measurements were obtained using an exposure time of 1 second and a 639 laser power of 20 mW.

[0243] DNA sequencing measurement For sequencing measurements, the assembled nanoantennas were loaded onto coverslips as described above. Subsequently, 1 μM of biotinylated DNA complementary to the AuNP-coated DNA was added for 1 minute to immobilize the AuNPs on the coverslip, followed by washing with imaging buffer. Next, photobleaching measurements were performed as described to precisely localize the assembled particles. Then, sequencing measurements were initiated using 639 and 568 lasers at a laser power of 20 mW for 20,000 frames with an exposure time of 100 ms. Once the measurement started, 100 nM dNTPs were flowed through the channel at a rate of 50 μl / min for 2 minutes, after which the flow rate was reduced to 10 μl / min. Synthesis of fluorescently labeled dNTPs material • Amination of dNTPs: γ-(6-aminohexyl)-dGTP, γ-(6-aminohexyl)-dATP, γ-(6-aminohexyl)-dCTP, γ-(6-aminohexyl)-dTTP, all 50 μl (10 mM), stored at -20°C. • Alexa Fluor® 647 NHS ester: Freshly prepared in DMSO, 25 nanomoles / tube, dried by Speedyvac. • Alexa Fluor® 568 NHS ester: Freshly prepared in DMSO, 25 nanomoles / tube, dried by Speedyvac. • 10x reaction buffer: 1M NaHCO3, pH=8.30 (freshly prepared) • HPLC buffer A: 0.1M TEAA • HPLC buffer B: acetonitrile

[0244] Amine and ester crosslinking reactions Alexa Fluor® 568 / 647 NHS ester was dissolved in 5 μL of DMSO to a stock concentration of 5 mM. 10 μL of 0.4 mM Alexa Fluor® 568 / 647 NHS ester and 1 mM γ-(6-aminohexyl)-dNTPs were incubated in reaction buffer at room temperature for 2 hours, and then purified by HPLC as described above.

[0245] (Example 2) Design and Embodiment of Plasmon Nano Antenna DNA Sequencer The plasmon nanoantenna consists of three components: firstly, metal nanoparticles (in this case, spherical gold nanoparticles with an average diameter of 100 nm); secondly, a DNA polymerase enzyme; and thirdly, a nanoscale scaffold constructed from DNA is used to control the spatial localization of the metal nanoparticles and DNA polymerase, thereby directly creating a field of electromagnetic field enhancement in a single DNA polymerase enzyme immobilized between the gold nanoparticles within a cavity (Figure 1A). To enable sequencing readouts, the dNTPs are labeled with fluorophores of different colors, and thus the sequential incorporation of these dNTPs by DNA polymerase can be detected. The scaffold also allows for the specific immobilization of the plasmon nanoantenna onto the surface of a glass coverslip for single-molecule fluorescence imaging (Figure 1B).

[0246] The following sections describe the synthesis of each component and the assembly of all components into a complete nano-antenna DNA sequencer.

[0247] Design and synthesis of DNA origami scaffolds Novel DNA sequencer designs require placing DNA polymerase directly within a plasmon-enhancing hotspot created by two 100 nm AuNPs spaced 30 nm apart. Therefore, a novel three-dimensional DNA origami structure was designed and folded from a scaffold strand and complementary staple strand derived from M13mp18.

[0248] Design of DNA Origami Scaffolding The DNA origami scaffold consists of 86 interconnected parallel DNA double helices designed using a honeycomb lattice to form a U-shaped structure with a central cavity, where each helix between AuNPs is set to make 7 turns or to be 72 bp long (24.5 nm). The dimensions of the cavity are approximately 25 × 12 × 24.5 nm (Figure 2A), which should provide space for polymerase binding. For immobilization on a coverslip for imaging using fluorescence microscopy, the U-shaped cavity has a long, rigid tail consisting of a bundle of four DNA helices, approximately 80 nm in length. At the base of the tail are one or more biotin-modified DNA staple strands, enabling specific immobilization via biotin-streptavidin interactions onto a BSA-biotin-coated coverslip (Figure 2A&C). The overall shape of the DNA origami scaffold resembles a nanoscale wrench, with the U-shaped cavity resembling the wrench head and the tail resembling the wrench handle. This DNA origami scaffold is called NanoSpanno(NS).

[0249] Single-stranded DNA strands can be designed to protrude from the DNA helix to create specific binding sites (stalks) for proteins or nanoparticles. These stalks are decorated with complementary (anti-stalk) DNA strands. In the head of the wrench, 10 DNA staples extend from the edge of either end of the cavity, providing anchoring sites for the binding of 100 nm AuNPs functionalized with DNA (Figure 2B&D). If gold nanoparticles are bound to both sides of the cavity, the estimated gap length is approximately 35 nm. Within the cavity between these gaps, another staple extension with an orthogonal DNA sequence is placed as a stalk that specifically binds to a polymerase enzyme covalently bound to the complementary DNA strand, as detailed in Example 1 (6E). This allows the polymerase to directly attach to the plasmon-enhancing hotspot of the plasmon nanoantenna (Figure 5B).

[0250] The signal amplification achievable in this NanoSpanno is calculated using a finite element model by directly incorporating fluorophore-labeled DNA staple strands (see below) into the plasmon hotspot of the NanoSpanno. These models demonstrate that fluorescent dyes within cavities between gold nanoparticles can be observed at more than 3000 times the brightness of fluorescent dyes not positioned within the hotspot (Figure 3).

[0251] Synthesis of DNA origami scaffolds DNA origami samples were purified and analyzed by agarose electrophoresis (AGE). Consistent with rigid, stable, and uniformly structured DNA origami, the NS moved at a constant rate in agarose electrophoresis, resulting in a single, distinct band. Furthermore, the migration speed was faster than that of the M13mp18 scaffold alone, thus indicating a smaller structure with a smaller radius of rotation (Figure 3A). These findings regarding AGE suggest a well-formed DNA origami structure.

[0252] To verify that the DNA origami structure was consistent with the design, the inventors directly visualized the purified structure using a transmission electron microscope. The NS structure was observed in various orientations. Examples of micrographs showing multiple single particles can be seen in Figure 3B, and the 2D class average of multiple particles in various orientations is shown in Figure 3C. The DNA origami particles appeared to be well formed with shape and dimensions consistent with the design, and therefore their structural integrity was verified.

[0253] Preparation of DNA-coated gold nanoparticles The binding of metal nanoparticles to specific sites on a DNA origami scaffold was facilitated by DNA hybridization. Single-stranded DNA on both sides of the cavity was subjected to thiol-gold chemical reaction. 28The single-stranded DNA used to coat the gold nanoparticles was designed to be complementary to the thiolated DNA. The optimized method for coating the metal nanoparticles is described herein. Co-localization of thiolated DNA and gold nanoparticles was demonstrated by agarose gel electrophoresis (Figure 4A), and a halo appeared on the surface of the DNA-coated gold nanoparticles, which was visualized by TEM. This halo was not observed in uncoated gold nanoparticles (Figure 4B). These data together are consistent with effective coating of gold nanoparticles by thiolated DNA.

[0254] Since 100nm gold nanoparticles tend to aggregate easily, a highly efficient method for coating the nanoparticles was extremely important. To ensure that these single-stranded DNA molecules achieved the following specifications, extensive optimization was required to determine their precise length and sequence: 1. Avoid causing aggregation of DNA origami structures. 2. It is possible to prevent aggregation of gold nanoparticles. This was particularly difficult for nanoparticles with a diameter of 100 nm. 3. To efficiently enable the specific coupling of nanoparticles to specific binding sites on DNA origami scaffolds.

[0255] Table 2 below details various DNA strands bound to gold nanoparticles for testing nanoparticle stability, DNA-gold binding efficiency, and assembly onto NS DNA origami scaffolds. Table 2. Details of various DNA strands bound to gold nanoparticles [Table 2]

[0256] Nanoparticle stability After functionalizing AuNPs with thiolated DNA, the pellet of the sample was carefully resuspended in 1×TAE buffer containing 6 mM MgCl2 to ensure uniform distribution of the sample. Stability was defined by the tendency to aggregate under 6 mM MgCl2, which was high enough to allow binding of gold nanoparticles to the DNA origami structure by DNA hybridization. Aggregation was detected by agarose gel electrophoresis, which yielded clear DNA staining bands (Figure 5) and solution color at the locations corresponding to the gold nanoparticles. Well-dispersed colloidal gold formed a clear red solution, while aggregated nanoparticles changed the solution to purple or clear, with aggregates visible in the tube. Figure 5 illustrates sequences 5-8 from Table 2. All of these sequences were 25 nucleotides long and, upon binding with gold nanoparticles, produced a stable colloidal solution, while sequences 1-3 produced a purple or colorless solution showing aggregation.

[0257] Design and synthesis of DNA polymerase enzymes Three polymerases were selected: DNA polymerase (Taq) from Thermus aquaticus, a large (Krenow) fragment of DNA polymerase I from Escherichia coli, and DNA polymerase (Phi29) from bacteriophage Phi29. Each polymerase possesses distinct properties that may be useful for the development of single-molecule sequencers. Recombinant versions of these polymerases were fused at their N-terminus via a SpyCatcher domain and a glycine-serine-glycine (GSG) linker, thereby enabling attachment to a DNA origami scaffold containing the corresponding SpyTag-DNA "stalk." An N-terminal polyhistidine tag (His tag) containing 6× histidine residues was also included to facilitate purification by IMAC (Figure 6).

[0258] Purification of protein constructs The proteins were purified using IMAC and SEC (see Example 1) and labeled with AlexFluor 647 to enable fluorescence visualization (see Example 1).

[0259] IMAC purification yielded a single elution peak (Figure 7A), and in that fraction, a band dominant at the molecular weight corresponding to the target protein was obtained in SDS-PAGE (Figure 7B). The peak fractions were combined and then further purified using SEC. The SEC chromatogram showed reasonable monodispersity (Figure 7C), and the corresponding SDS-PAGE showed a final purity of over 95% (Figure 7D). Alexa 647 labeling was visible in the SEC chromatogram, with a single elution peak visible in both A280 and A650 measurements (Figure 7C). In the corresponding SDS-PAGE analysis, a dominant band was shown in both the Alexa 647 channel and bright-field after staining (Figure 7D). The peak fractions were combined, and absorbance was measured at 280 nm and 650 nm. The results showed 100% Alexa 647 labeling efficiency.

[0260] Synthesis of DNA-SpyTag peptide conjugates To achieve specific binding to the cavities of the DNA origami scaffold via DNA hybridization, it was necessary to covalently attach the DNA strand to the DNA polymerase enzyme. Our method involved covalently linking a SpyTag peptide, which forms an isopeptide bond with a SpyCatcher protein fused to the DNA polymerase enzyme, to the DNA strand. DNA-SpyTag conjugation was achieved using a bifunctional crosslinking agent (SMCC) consisting of an N-hydroxysuccinimide (NHS) ester and a maleimide group. These parts were covalently bonded to a free amino group chemically added to the target DNA strand and to a sulfhydryl group on the cysteine ​​side chain of the peptide, respectively. In this way, the desired DNA-SpyTag construct was obtained (Figure 8).

[0261] maleimide DNA synthesis As detailed in Example 1 above, the synthesis of the DNA-peptide involved a two-step process. First, maleimide DNA was synthesized by reacting SMCC with aminated DNA (Figure 8A), and then purified by ethanol precipitation and HPLC. Maleimide could be selectively added to the 5' or 3' end of the DNA strand. Typical UV chromatograms obtained by HPLC purification are shown in Figures 9A and 9B for 5'-modified maleimide DNA and 3'-modified maleimide DNA, respectively, and in Figure 9C for 5'Cy5-labeled 3'-maleimide DNA. In both cases, the dominant peak corresponds to maleimide DNA.

[0262] Synthesis of peptide-DNA conjugates In the second step, maleimide DNA was conjugated to the free thiol at the N-terminal cysteine ​​residue of the target peptide and purified using HPLC (see Example 1). UV chromatograms are shown in Figures 10A and 10B for DNA-peptide conjugates with peptides attached to the 3' and 5' ends of the DNA, respectively, and in Figure 10C for 5'-Cy5 labeled DNA with the peptide attached to the 3' end. In all cases, a single dominant peak corresponding to the DNA-peptide conjugate was observed, indicating a high-purity target DNA-peptide conjugate.

[0263] Conjugation of SpyTag-DNA and polymerase-SpyCatcher The SpyTag-DNA conjugate was conjugated to a polymerase-SpyCatcher fusion protein and purified using IMAC as described in Example 1. Conjugation efficiency was visualized by SDS-PAGE, illustrating that the migration speed of DNA polymerase was significantly slower in the presence of SpyTag-DNA compared to DNA polymerase alone (Figure 11 - Krenow's example). The shift in migration speed occurred independently of whether the peptide was bound to the 5' or 3' end of the DNA (Figure 11). Co-localization of protein and DNA was demonstrated by a combination of protein-specific staining that specifically localizes either the fluorescently labeled DNA or the protein, and fluorescence imaging of the gel, thus confirming the successful assembly of the DNA-labeled polymerase enzyme.

[0264] Demonstration of DNA-labeled DNA polymerase function DNA polymerase function was evaluated by measuring the elongation of a fluorescently labeled primer bound to a target DNA strand. The reaction was stopped at specific time points, and primer elongation was visualized by undenatured PAGE. All polymerases appeared to successfully elongate the fluorescent primer, resulting in bands that moved more slowly at 0.5 to 1 hour compared to 0 hour (Figure 16A). Both the 10 nM Krenow-SpyCatcher-SpyTag-DNA conjugation and the 50 nM Krenow-SpyCatcher-SpyTag-DNA conjugation appeared to successfully elongate the fluorescent primer, resulting in bands that moved more slowly from 24 hours compared to 0 hour (Figure 16B). The inventors concluded that the DNA-labeled DNA polymerase enzyme retains its function.

[0265] Dynamic analysis of polymerase interactions with freely diffusing primed DNA To ensure the use of experimental conditions that enable highly efficient capture of target DNA molecules during sequencing reactions, it was crucial to determine the dynamic characteristics of the interaction between DNA polymerase and the target DNA strand being primed. Similar to single-molecule TIRF microscopy, binding dynamics are affected by proximity to the surface; therefore, binding dynamics were measured in bulk using surface plasmon resonance (SPR) (Example 1).

[0266] Regarding Krenow, the association curve (Figure 13A) is biphasic, and two concentration-dependent association rate constants were extracted. The relatively fast rate constant (k_(a,fast) = 3.7 ± 0.3 ×

[10] ) 6 mol (-1) s (-1) ) and slow velocity constant (k_(a, slow) = 7.4 ± 0.3 ×

[10] 4 mol (-1) s (-1) The dissociation curve (Figure 13B) is also biphasic, and the fast dissociation rate constant (k_(d,fast) = 1.8 ± 1.1 ×

[10] ) (-2) s (-1) ), the average dissociation time is 38 seconds, and the slow dissociation rate constant is (k_(d, slow) = 5.9 ± 1.1 ×

[10] (-4) s (-1) The average dissociation time was 20 minutes. The dissociation rate was determined from the proportion of polymerase bound in the steady state (K_D = 26 ± 3 nM, Figure 13C).

[0267] For Phi29, the association curve (Figure 13D) is similarly biphasic, and two concentration-dependent association rate constants were extracted. Another relatively fast rate constant (k_(a,fast) = 1.1 ± 0.3 ×

[10] 6 mol (-1) s (-1) ) and slower rate constants (k_(a, slow) = 2.1 ± 0.3 ×

[10] 5 mol (-1) s (-1) The dissociation curve (Figure 13E) is also biphasic, and the fast dissociation rate constant (k_(d,fast) = 1.4 ± 0.1 ×

[10] ) (-2) s (-1)), mean dissociation time 50 seconds, and slower dissociation rate constant (k_(d, slow) = 1.2 ± 0.2 ×

[10] (-3) s (-1) The average dissociation time was 10 minutes. The dissociation constant was determined from the proportion of polymerase bound in the steady state (K_D = 56 ± 9 nM, Figure 13F).

[0268] Plasmon nanoantenna assembly The primary challenge was determining the DNA strand sequence and length to enable stable passivation (pacification) to avoid aggregation of gold nanoparticles under 6 mM MgCl2, which is necessary to facilitate the binding of gold nanoparticles to the DNA origami scaffold via DNA hybridization, and to prevent aggregation of the DNA origami scaffold itself. Several combinations of DNA strand and nanoparticle size were tested. The effectiveness of these combinations and their assemblies in the target structure is summarized in Table 3 below. Table 3. Overview of gold nanoparticle sizes and the DNA sequences assembled on DNA origami scaffolds to coat those nanoparticles. [Table 3]

[0269] Assembled plasmon nanoantennas evaluated by electron microscopy. Various molar excesses of gold nanoparticles were tested across a DNA origami scaffold with different binding site configurations, including zero, one, or two binding sites to the gold nanoparticles. After incubation, the samples appeared red, suggesting that the solution was not occupied by undesirable aggregates (Figure 14A).

[0270] The samples were further analyzed using agarose gel electrophoresis. As expected, a single dominant band was obtained from DNA origami composed of 0, 1, or 2 binding sites. In the presence of excess gold nanoparticles, this band was not observable for DNA origami configurations with 1 or 2 gold binding sites. Importantly, when the DNA origami was composed without binding sites, the band corresponding to the DNA origami alone was still observable even in the presence of excess gold nanoparticles (Figure 14A). In summary, this suggests that gold nanoparticles specifically bind to DNA origami only when gold binding sites are present.

[0271] Gold nanoparticles alone produced ladder-like bands indicating oligomer formation. However, no additional bands were observed when gold nanoparticles were bound to DNA origami. Therefore, it is difficult to determine the number of gold nanoparticles bound to the DNA origami scaffold (Figure 14A).

[0272] To quantify the number of gold nanoparticles bound to the DNA origami scaffold and to confirm that these gold nanoparticles bound to their specific binding sites, the DNA origami-gold complex was visualized using a transmission electron microscope. An example of a well-assembled particle observed in an electron microscope image is shown in Figure 14B, where, as designed, two gold nanoparticles are bound to both sides of the DNA origami scaffold. In principle, this configuration of gold nanoparticles and DNA origami constitutes a functional plasmon nanoantenna that enhances the fluorescence intensity of fluorescent molecules within the cavities of the DNA origami scaffold. Examples of gold nanoparticle monomers and aggregates in electron microscope images are shown in Figure 14C. Some aggregates appear to have arisen from DNA origami-mediated daisy chaining, while others appear to have arisen from direct interactions between gold nanoparticles.

[0273] Assembly yield of plasmon nanoantenna particles evaluated by electron microscopy Using transmission electron microscopy images, we quantified the proportion of gold nanoparticles bound to DNA origami structures and their constituent structures (monomers, dimers, aggregates) using two different sulfhydryl DNA extensions (25-TG and 25-AC) with DNA origami scaffolds of different configurations (0, 1, or 2 binding sites).

[0274] When there were two binding sites, 100% of the DNA origami nanospanno structures bound to AuNP, and no free DNA origami structures were observed in electron microscopy. 22% of the molecules were dimers, approximately 10% were monomers, and approximately 65% ​​were aggregates or other poorly assembled structures (Figures 19A and D). For DNA origami structures with one binding site, only half bound to AuNP. No dimeric assemblies were observed, and 14–22% of AuNP contained bound single DNA origami structures (Figures 19B and E). For DNA origami structures without gold binding sites, only free gold and free NS were obtained in EM, and no evidence of assembled structures was found (Figures 19C and F). These data, when combined, demonstrate the successful assembly of a specific, desired plasmon nanoantenna architecture consisting of two 100 nm gold nanoparticles on either side of a cavity capable of immobilizing DNA polymerase.

[0275] Colocalization of DNA polymerase and DNA origami scaffolds This section describes the specific attachment of polymerase-DNA to the cavities of the DNA origami scaffold. This specific attachment occurs through hybridization between the DNA bound to the polymerase enzyme and the extension of complementary DNA staples positioned within the cavity (Figure 16).

[0276] Measurement of bulk co-localization using agarose gel electrophoresis The binding of DNA polymerase labeled with Alexa 488 fluorescent dye and DNA polymerase labeled with Alexa 647 dye was evaluated in bulk by observing their co-localization using agarose gel electrophoresis. Co-localization of DNA polymerase and DNA origami was observed only when the DNA polymerase was bound to SpyTag DNA. Furthermore, it was observed that the migration speed of both the DNA origami and DNA polymerase alone slowed down due to co-localization (Figure 17). These data suggest that the binding of DNA polymerase occurs through specific hybridization with DNA staple extension within the cavities of the DNA origami.

[0277] Quantification of colocalization yield using single-molecule fluorescence microscopy To quantify the co-localization yield of DNA polymerase and DNA origami scaffolds, single-molecule fluorescence microscopy was used to determine the proportion of single DNA origami molecules bound to a single DNA polymerase enzyme. The presence of a single DNA polymerase enzyme was determined by photobleaching. Here, only molecules exhibiting a single photobleaching step were used for yield analysis (Figure 18A and B). In contrast, we found that the Alexa 488 fluorophore bound to DNA origami was resistant to photobleaching. Therefore, the presence of single DNA origami particles was determined by the initial fluorescence intensity when excited with 488 nm light. By plotting the initial intensity distribution of all fluorescent particles, single DNA polymerase was easily distinguishable from oligomers or aggregates (Figure 18C). These measures allowed us to determine that the yield of single DNA origami scaffolds co-localized with single DNA polymerase molecules was approximately 90%, regardless of the location of the DNA polymerase binding site (Figure 18D). Note that, as a positive control, an excess amount of Alexa 647 DNA complementary to the DNA polymerase binding site was annealed. This provided a measure of the possible maximum occupancy of DNA polymerase, which was also 90% (Figure 18D).

[0278] (Example 3) Characterization of plasmon nanoantennas Enhancement of fluorescently bound fluorescent dyes at hotspots. To determine the function of plasmon nanoantennas, the intensity of fluorophores bound to DNA origami scaffolds, either individually or in conjunction with a single or pair of gold nanoparticles, was experimentally quantified.

[0279] Intensity distribution measured by single-molecule photobleaching The intensity of Alexa 647 fluorophores immobilized within plasmon nanoantennas was measured by a single-molecule photobleaching assay. Alexa 647-labeled DNA origami scaffolds with zero or two AuNP binding sites were incubated with 500 pM AuNP at 100 pM for 2 hours, and then diluted to 1 / 10 for TIRF microscopy (see the section titled "Single-molecule Total Internal Reflection Fluorescence (TIRF) Microscopy" in Example 1). Figure 19A shows an example of a single photobleaching step for DNA origami scaffolds without AuNPs (blue), whose intensity is substantially lower than that of a single fluorophore in DNA origami scaffolds with a pair of AuNPs (red). Figure 19B shows the intensity distribution for approximately 3000 individual assemblies that either lack AuNP binding sites (blue) or contain two AuNP binding sites (red). These data reveal a broad distribution of fluorescence enhancement, ranging from 1x to 200x, in the presence of two gold nanoparticles. This broad distribution is attributed to several factors, including the assembly yield of the 2×AuNP:1×DNA origami scaffold, the orientation of plasmon nanoantennas on the coverslip, possible anisotropy of fluorophore labeling, variations in the distance between nanoparticle pairs, and heterogeneity of nanoparticle size and shape. The fading rates (Figure 19C) are similar, indicating a higher total photon count from the fluorophore in the presence of AuNP.

[0280] TEM experiments showed a yield of 22% for precisely assembled nanoantennas. Scattering from gold nanoparticles was significant, and the scattering cross-section at 488 nm increased or decreased depending on the number of particles in the cluster, allowing for the identification of single DNA origami scaffolds with 0, 1, 2, or more gold nanoparticles (Figure 19D). Importantly, the proportion of dimerized gold nanoparticles was similar to that directly observed using electron microscopy (Figure 18D and Table 4). The distribution of fluorophore intensity in DNA origami scaffolds with 0, 1, or 2 gold nanoparticles, as measured by scattering, was separated. The average fluorophore intensity in DNA scaffolds with 2 gold nanoparticles was 10 times higher than that in DNA scaffolds without gold nanoparticles (Figure 19E). In particular, in the presence of 2 gold nanoparticles, the range of fluorescence enhancement varied substantially, with some particles showing no fluorescence enhancement and others showing enhancement of more than 100 times. This was not unexpected, as the plasmon enhancement effect relies on the orientation of the gold nanoparticle pair in harmony with the polarization of the incident light. Quantifying the rate of photobleaching, it was again found to be largely independent of the number of bound gold nanoparticles (Figure 19F).

[0281] Table 4. Quantification of populations of single Alexa647-labeled origami fading steps using corresponding scattering intensities of 0, 1, or 2 AuNPs. [Table 4]

[0282] Fluorescence intensity enhancement measured by DNA PAINT Photobleaching experiments are useful for determining the assembly composition, stoichiometry, and fluorescence intensity of static molecules, but DNA sequencing requires repeated measurements of the fluorescent dye because binding within the plasmon nanoantenna hotspot is transient. To establish the conditions necessary for DNA sequencing, we measured the repeated transient binding of fluorescent DNA strands to 7-base DNA staple extensions within the plasmon nanoantenna hotspot. Repeated measurements for the same molecule also revealed whether bright particles are always bright or whether there is significant variation in fluorescence intensity from a single plasmon nanoantenna.

[0283] Figure 20 shows fluorescence intensity spikes resulting from transient binding of the fluorescent DNA strand in the cavity of the DNA origami structure, both in the presence and absence of immobilized gold nanoparticle pairs. A 10- to 50-fold fluorescence enhancement was observed in the presence of gold pairs, thus confirming that plasmon nanoantennas function to enhance the fluorescence signal. Again, the wide range of enhancement intensity is likely due to variations in the orientation of the gold nanoparticles within the duration of the experiment. Fewer intensity spikes were observed in the absence of the imager strand (Figure 21). The origin of these spikes is thought to be due to variations in the orientation of the gold nanoparticles, which increases the scattering intensity and appears as a peak in the intensity trace, potentially leading to false-positive events in sequencing measurements.

[0284] The effect of polarization field rotation on the orientation of gold nanoparticles is experimentally characterized below.

[0285] Fluorescence enhancement of DNA polymerase in hotspots Next, to complete the synthesis of the plasmon nanoantenna DNA sequencer, a fluorescently labeled DNA polymerase molecule was bound to the hotspot of the plasmon nanoantenna. The intensity of a single fluorescent DNA polymerase enzyme bound to the cavity of the DNA origami scaffold was measured using single-molecule photobleaching, both without gold and with a pair of gold nanoparticles. Examples of bleaching traces without and with gold are shown on the left and right sides of Figure 22, respectively, demonstrating a two-order-of-magnitude increase in the fluorescence signal in the presence of the gold pair. As mentioned above, the complete distribution of plasmon enhancement with the gold nanoparticle pair is, as expected, extensive, due to the free rotation of the gold spheres. Nevertheless, these data provide direct evidence that the DNA polymerase was successfully immobilized within the hotspot of the plasmon nanoantenna. This is the first example of a protein molecule fluorescently enhanced by a plasmon nanoantenna.

[0286] Characterization of fluorescence enhancement at different polarization angles To determine the effect of incident light polarization on the scattering intensity of plasmon nanoantennas, simulations of scattering cross-section, light intensity, and fluorescence enhancement were performed for nanoantennas oriented perpendicularly to a glass surface at various incident polarization angles. Figure 23Ai shows how the TIR intensity above the coverslip varies between perfectly p-polarized light (parallel, 0°) where the polarization is perpendicular to the coverslip and perfectly s-polarized light (perpendicular, 90°) where the polarization is aligned with the coverslip. Using this, the polarization angle was experimentally calibrated based on the background intensity measured when the incident polarization was changed by a half-wave plate (see the section titled "Measurement of Gold Scattering Intensity and Polarization Angle" in Example 1) (Figure 23Aii). Figure 23Bi shows the simulated scattering cross-sections of nanoantennas oriented either in the direction of TIRF field propagation (blue) or perpendicular to propagation (black). In the blue orientation, there is no optimal alignment between the nanoantenna and the incident polarization; therefore, the scattering cross-section is low on average across all polarization angles, and the change with polarization angle follows the change in TIR intensity. In the black orientation, the change in scattering cross-section is out of phase (90°), and in this orientation, s-polarized light aligns with the nanoantenna, resulting in a larger average intensity. Figure 27Bii shows the experimentally measured intensities of scattered light for various particles when the polarization angle is changed. This allowed for measurements of nanoantennas aligned perpendicular to the propagation of the TIRF field (black), nanoantennas aligned parallel to it (blue), or a single AuNP nanoantenna (green).

[0287] When the TIRF field is s-polarized, the intensity measured based on the orientation of the nanoantennas changes significantly. To prevent this from happening during the duration of the sequencing experiment, AuNPs were immobilized on the coverslip via a biotin:streptavidin interaction. This was achieved by first attaching biotinylated DNA complementary to the DNA bound to the AuNPs to the functionalized coverslip via a biotinylated DNA origami scaffold, and then running the mixture. Measurement of the particles over approximately 30 minutes after this step showed a significant reduction in the number of intensity spikes (Figure 24).

[0288] The effect of nanoantenna orientation on Alexa-647 fluorophore enhancement was also simulated, showing enhancement changes ranging from less than 1x to 200x (Figure 25). The enhancement distribution for a randomly oriented population of nanoantennas was similar to that observed above.

[0289] (Example 4) Single-molecule DNA sequencing reaction Fluorophore-labeled dNTPs DNA sequencing was achieved by detecting the incorporation of a single fluorescent nucleotide by a DNA polymerase enzyme positioned within a plasmon hotspot, using each nucleotide (ATP, GTP, TTP, or CTP) labeled with a different color dye. Importantly, the fluorescent probe needed to be attached to the terminal phosphate of the nucleotide, which prevented termination of the polymerase reaction and meant that a phosphodiester bond would be formed and the fluorescent probe released upon nucleotide incorporation. Nucleotides with a 6-aminohexyl group attached to the distal phosphate were purchased from Jena Biosciences, allowing for succinimidyl fluorophore (Sigma) attachment using the NHS esterification reaction method before purification by HPLC.

[0290] Amination of dGTP typically elutes at approximately 12 minutes, while Alexa647-labeled dGTP typically elutes at approximately 32 minutes (Figure 26A). The fraction containing the pure product was evaporated and then co-evaporated with methanol:MQW = 1:1 (2×). The residue was dissolved in water, quantified by UV-Vis spectroscopy, then divided into 500 pmol / tube portions (resuspended in 10 μL of MQW to obtain a 50 μM working stock), dried, and stored at -20°C.

[0291] Alexa568 has two isomers, which generally elute at approximately 41 and 42 minutes, appearing blue at 260 nm and 578 nm (Figure 26B). These isomers have hydrolysis byproducts that elute at 30 and 35 minutes. Aminated dGTP generally elutes at approximately 12 minutes, appearing orange at 260 nm, while Alexa568-labeled dGTP generally elutes at approximately 31 and 36 minutes due to its two isomers, appearing orange at 260 nm and 578 nm (Figure 26B). Aminated dATP generally elutes at approximately 15 minutes, and Alexa568-labeled dTTP generally elutes at approximately 32 and 36 minutes. Amination-dTTP generally elutes at approximately 10 minutes, while Alexa568-labeled dTTP generally elutes at approximately 32 and 36 minutes. Finally, amination-dCTP typically elutes at approximately 5 minutes, while Alexa568-labeled dCTP generally elutes at approximately 32 and 36 minutes (Figure 26C).

[0292] Configuration of a plasmon nanoantenna for DNA sequencing As the first proof of concept for DNA sequencing experiments, a plasmon nanoantenna was configured to contain both a single DNA polymerase enzyme immobilized with a 71-base single-strand DNA linker and the target DNA strand to be primed (Figure 27). This configuration eliminates the need to capture freely diffusing DNA, thus minimizing experimental parameters. The length of the DNA reads is also sterically restricted.

[0293] Two-color DNA sequencing experiments using Alexa647-labeled dGTP and Alexa568-labeled dATP As a proof of concept, a plasmon nanoantenna DNA sequencer was assembled into the configuration illustrated in Figure 27 and immobilized on the surface of a coverslip for imaging using a TIRF microscope. To identify fully assembled plasmon nanoantenna sequencers, a single-molecule photobleaching assay was performed, and scattering intensity was measured to determine particles that had exactly one DNA origami scaffold, one DNA polymerase enzyme, and a pair of gold nanoparticles. The fluorescence intensity around these fully assembled particles was monitored over time with simultaneous excitation using 568 and 647 lasers to obtain two-color intensity traces. This resulted in a substantial increase in the fluorescence background, as expected. Furthermore, distinct fluorescence spikes well above the background were observed (Figure 28). Base calling was simply defined as any intensity spike that exceeded the background by more than three standard deviations. This base calling algorithm yielded multiple plasmon nanoantenna DNA sequencers that identified bases in strictly accurate sequences with read lengths of 8–16 bases. Two example traces are shown in Figure 28. Notably, in control experiments lacking a single crucial component, such as DNA polymerase or the target DNA template strand, the precise sequence could not be identified.

Claims

1. A method for determining the sequence of nucleic acid analytes, (I) A step of contacting nucleic acid polymerase with the nucleic acid analyte and the labeled nucleotide for a period of time and under conditions such that the labeled nucleotide is sequentially incorporated by the nucleic acid polymerase into a polynucleotide having a sequence complementary to the polynucleotide sequence of the nucleic acid analyte, wherein the nucleic acid polymerase is located within an electric field enhancement area, and each labeled nucleotide is (i) Adenine nucleotide (A), guanosine nucleotide (G), thymine nucleotide (T), or cytosine nucleotide (C), (ii) Fluorophores, and (iii) A polyphosphate linker that binds the nucleotide to the fluorophore. Includes, A, G, T, and C are each independently connected to a fluorophore via the polyphosphate linker. Each of the labeled nucleotides A, G, T, and C has a distinct fluorescence signature when the fluorophore to which each of the nucleotides is linked is excited. The separate fluorescent signature of the labeled nucleotide is enhanced when it is incorporated by the nucleic acid polymerase into a sequence complementary to the polynucleotide sequence of the nucleic acid analyte. Steps and (II) A step of detecting the sequence of enhanced separate fluorescence signatures when the labeled nucleotides are sequentially incorporated by the nucleic acid polymerase into polynucleotides having a sequence complementary to the polynucleotide sequence of the nucleic acid analyte, thereby determining the sequence of the nucleic acid analyte by determining the sequence of nucleotides incorporated into polynucleotides having a sequence complementary to the polynucleotide sequence of the nucleic acid analyte; Methods that include...

2. The method according to claim 1, wherein the distinct fluorescence signature of the labeled nucleotide is not enhanced outside the electric field enhancement area.

3. The method according to claim 1 or 2, wherein the enhanced separate fluorescence signature of the labeled nucleotide is enhanced by a factor of two or more compared to the separate fluorescence signature of the corresponding labeled nucleotide outside the electric field enhancement area.

4. The method according to claim 1, wherein the enhanced, distinct fluorescence signature of the labeled nucleotide is enhanced by an order of magnitude or more compared to the distinct fluorescence signature of the corresponding labeled nucleotide outside the electric field enhancement area.

5. The method according to any one of claims 1 to 4, wherein the electric field enhancement area is generated by a plasmon hotspot, and the plasmon hotspot is generated by a plasmon nanoantenna.

6. The method according to claim 5, wherein the plasmon nanoantenna comprises at least two plasmon nanoparticles.

7. Each plasmon nanoantenna Two plasmon nanoparticles, Nanoscale nucleic acid scaffolds and The nucleic acid polymerase and Includes, (a) The two plasmon nanoparticles are bound to the nanoscale nucleic acid scaffold and are arranged such that a plasmon hotspot exists between the two plasmon nanoparticles relative to each other. (b) The plasmon hotspot includes a region not occupied by the nanoscale nucleic acid scaffold. (c) The nucleic acid polymerase is bound to the nanoscale nucleic acid scaffold and is located in a region not occupied by the nanoscale nucleic acid scaffold. (d) If necessary, the nucleic acid analyte is bound to the nanoscale nucleic acid scaffold. The method according to claim 6.

8. The method according to claim 7, wherein the region not occupied by the nanoscale nucleic acid scaffold is defined by a three-dimensional space having a length (L) between the two plasmon nanoparticles, a height (H) perpendicular to (L) at the midpoint (L / 2), and a width (W) perpendicular to L and H at L / 2, wherein at L / 2, W is selected from 20 nm to about 100 nm, H is selected from 10 nm to about 100 nm, and L is measured at the shortest distance between the two plasmon nanoparticles.

9. The method according to claim 8, wherein L is selected from 20 nm to 100 nm.

10. The method according to claim 9, wherein L is selected from 20 nm to 50 nm.

11. The method according to any one of claims 7 to 10, wherein the volume of the region not occupied by the nanoscale nucleic acid scaffold is at least about 1 zL, preferably about 10 zL to about 50 zL.

12. The method according to any one of claims 7 to 10, wherein the region not occupied by the nanoscale nucleic acid scaffold is 80% or more of the plasmon hotspot.

13. The method according to claim 12, wherein the region not occupied by the nanoscale nucleic acid scaffold is 90% or more of the plasmon hotspot.

14. The method according to any one of claims 7 to 13, wherein the region not occupied by the nanoscale nucleic acid scaffold is amorphous.

15. The method according to any one of claims 7 to 13, wherein the region not occupied by the nanoscale nucleic acid scaffold is spherical or cubic.

16. The method according to any one of claims 7 to 15, wherein the plasmon nanoantenna comprises the nucleic acid analyte bound to the nanoscale nucleic acid scaffold.

17. The method according to claim 16, wherein the nucleic acid analyte is non-covalently bonded to the nanoscale nucleic acid scaffold.

18. The method according to claim 17, wherein the nucleic acid analyte is non-covalently bonded to the nanoscale nucleic acid scaffold by hydrogen bonds.

19. The method according to any one of claims 7 to 18, wherein each of the two plasmon nanoparticles is independently selected from metal nanoparticles.

20. The method according to claim 19, wherein each of the two plasmon nanoparticles is independently selected from the group consisting of gold, silver, aluminum, copper, palladium, and platinum nanoparticles or any one alloy thereof.

21. The method according to claim 20, wherein each of the two plasmon nanoparticles is a gold nanoparticle.

22. The method according to any one of claims 7 to 21, wherein the maximum diameter of each of the two plasmon nanoparticles is approximately 5 nm to approximately 500 nm.

23. The method according to claim 22, wherein the maximum diameter of each of the two plasmon nanoparticles is approximately 50 nm to approximately 150 nm.

24. The method according to claim 23, wherein the maximum diameter of each of the two plasmon nanoparticles is approximately 80 nm to 120 nm.

25. The method according to any one of claims 7 to 24, wherein each of the two plasmon nanoparticles is coated by a coating.

26. The method according to claim 25, wherein the coating substantially covers the surface of each of the two plasmon nanoparticles.

27. The method according to claim 25 or 26, wherein the coating reduces the aggregation of the two plasmon nanoparticles by 50% or more compared to the two plasmon nanoparticles without the coating.

28. The method according to any one of claims 25 to 27, wherein the coating comprises a plurality of oligonucleotides.

29. The method according to claim 28, wherein each of the plurality of oligonucleotides comprises about 20 nucleotides to about 50 nucleotides.

30. The method according to claim 29, wherein each of the plurality of oligonucleotides comprises about 20 to about 30 nucleotides.

31. The method according to any one of claims 28 to 30, wherein each nucleotide in each of the plurality of oligonucleotides is selected from pyrimidine nucleotides.

32. The method according to any one of claims 6 to 31, wherein the nanoscale nucleic acid scaffold is substantially composed of double-stranded DNA, preferably interconnected parallel strands of a double helix.

33. The method according to any one of claims 7 to 31, wherein the nanoscale nucleic acid scaffold is composed of a DNA origami structure.

34. The method according to claim 33, wherein at least a portion of the DNA origami structure is assembled to form a U-shaped or curved structure.

35. The method according to any one of claims 7 to 34, wherein the nanoscale nucleic acid scaffold includes a first surface to which the first plasmon nanoparticle of the two plasmon nanoparticles is anchored, and a second surface to which the second plasmon nanoparticle of the two plasmon nanoparticles is anchored.

36. The method according to claim 35, wherein the first surface of the scaffold and the second surface of the scaffold are on the same side or adjacent side of the nanoscale nucleic acid scaffold.

37. The method according to claim 35, wherein the first surface of the scaffold and the second surface of the scaffold are on opposite sides of the nanoscale nucleic acid scaffold.

38. The two plasmon nanoparticles are anchored to the first surface of the scaffold and the second surface of the scaffold, respectively, through one or more nucleic acid linkers. Each nucleic acid linker is formed by an oligonucleotide that hybridizes with a polynucleotide that forms part of the nanoscale nucleic acid scaffold or extends from the nanoscale nucleic acid scaffold, which is coated on the surface of the plasmon nanoparticles. The sequences of the oligonucleotide and polynucleotide that can hybridize are complementary or substantially complementary to each other. The method according to any one of claims 35 to 37.

39. The method according to any one of claims 35 to 38, wherein the nanoscale nucleic acid scaffold comprises one or more polynucleotides extending from the first surface of the scaffold, and each of the one or more polynucleotides comprises a sequence complementary to the sequence of one or more oligonucleotides coating one of the two plasmon nanoparticles.

40. The method according to any one of claims 35 to 39, wherein the nanoscale nucleic acid scaffold comprises one or more polynucleotides extending from the second surface of the scaffold, and each of the one or more polynucleotides comprises a sequence complementary to the sequence of one or more oligonucleotides coating one of the two plasmon nanoparticles.

41. The method according to claim 39 or 40, wherein the one or more polynucleotides extending from the surface of the scaffold are independently selected from single-stranded DNA.

42. The method according to any one of claims 7 to 41, wherein the nucleic acid polymerase is bound to or immobilized on the nanoscale nucleic acid scaffold via a nucleic acid linker.

43. The method according to claim 42, wherein the nucleic acid linker is double-stranded and comprises (i) a polynucleotide covalently or non-covalently bonded to the nucleic acid polymerase via amino acids in the nucleic acid polymerase, and (ii) a polynucleotide forming part of the nanoscale nucleic acid scaffold or extending from the nanoscale nucleic acid scaffold, wherein the polynucleotide in (i) and the polynucleotide in (ii) comprise a single-stranded DNA sequence that is complementary or substantially complementary to each other and capable of hybridizing.

44. The method according to claim 43, wherein the polynucleotide in (i) includes a peptide tag, and the peptide tag is bound to the nucleic acid polymerase via an isopeptide bond formed between an amino acid in the nucleic acid polymerase and an amino acid in the peptide tag.

45. The method according to claim 42 or 43, wherein the nucleic acid linker is bound to the nucleic acid polymerase via an amino acid in the N-terminal domain of the nucleic acid polymerase.

46. The method according to any one of claims 1 to 45, wherein the nucleic acid polymerase is DNA polymerase.

47. The method according to any one of claims 1 to 46, wherein the nucleotide of the labeled nucleotide is further selected from synthetic nucleotides.

48. The method according to claim 47, wherein the synthetic nucleotide is selected from 5-methylcytidine, N-methylcytidine, and N6-methyladenosine.

49. The method according to claim 47 or 48, wherein the synthetic nucleotide is linked by a fluorophore and a polyphosphate, and each of the nucleotides A, G, T, C and the synthetic nucleotide has a distinct fluorescence signature when the fluorophore to which each of the nucleotides is linked is excited.

50. The method according to any one of claims 1 to 49, wherein the distinct fluorescence signatures are distinguishable from one another based on differences in peak emission wavelength, differences in fluorescence emission intensity, differences in fluorescence emission duration, differences in duration between sequential fluorescence emission, or any combination thereof.

51. The method according to any one of claims 1 to 50, wherein each different species of nucleotide is linked to a fluorophore having a distinct peak emission wavelength.

52. The method according to claim 51, wherein the separate peak emission wavelengths are separated from each other by 10 nm or more.

53. The method according to any one of claims 1 to 50, wherein two or more species of nucleotides are linked to the same fluorophore, and the nucleotides of different species are distinguished based on differences in fluorescence intensity and / or fluorescence duration.

54. The method according to any one of claims 1 to 53, wherein the polyphosphate is a triphosphate, a tetraphosphate, a pentaphosphate, or a hexaphosphate.

55. The method according to claim 54, wherein the fluorophore is bound to the phosphate furthest from the nucleotide.

56. The method according to any one of claims 1 to 55, wherein the step of detecting the sequence of the enhanced separate fluorescence signatures of (II) is performed using a fluorescence microscope or a fluorophotometer.

57. The method according to any one of claims 1 to 56, wherein the step of detecting the sequence of the enhanced distinct fluorescence signatures of (II) is performed using a total internal reflection fluorescence (TIRF) microscope.

58. The method according to any one of claims 1 to 57, wherein the nucleic acid analyte is DNA.

59. The method according to any one of claims 1 to 57, wherein the nucleic acid analyte is complementary DNA (cDNA) derived from an RNA sample.

60. A method according to claim 5, or any one of claims 6 to 59 dependent on claim 5, comprising using an array of plasmon nanoantennas.

61. Two plasmon nanoparticles, Nanoscale nucleic acid scaffolds and Nucleic acid polymerase and A plasmon nanoantenna including, (a) The two plasmon nanoparticles are bound to the nanoscale nucleic acid scaffold and are arranged such that a plasmon hotspot exists between the two plasmon nanoparticles relative to each other. (b) The plasmon hotspot includes a region not occupied by the nanoscale nucleic acid scaffold, (c) The nucleic acid polymerase is bound to the nanoscale nucleic acid scaffold and is located in a region not occupied by the nanoscale nucleic acid scaffold. (d) If necessary, the nucleic acid analyte is bound to the nanoscale nucleic acid scaffold. Plasmon nano-antenna.

62. The plasmon nanoantenna according to claim 61, wherein the region not occupied by the nanoscale nucleic acid scaffold is defined by a three-dimensional space having a length (L) between the two plasmon nanoparticles, a height (H) perpendicular to (L) at the midpoint (L / 2), and a width (W) perpendicular to L and H at L / 2, wherein at L / 2, W is selected from 20 nm to about 100 nm, H is selected from 10 nm to about 100 nm, and L is measured at the shortest distance between the two plasmon nanoparticles.

63. The plasmon nanoantenna according to claim 62, wherein L is selected from 20 nm to 100 nm.

64. The plasmon nanoantenna according to claim 63, wherein L is selected from 20 nm to 50 nm.

65. The plasmon nanoantenna according to any one of claims 61 to 64, wherein the volume of the region not occupied by the nanoscale nucleic acid scaffold is at least about 1 zL, preferably about 10 zL to about 50 zL.

66. The plasmon nanoantenna according to any one of claims 61 to 65, wherein the region not occupied by the nanoscale nucleic acid scaffold is 80% or more of the plasmon hotspot.

67. The plasmon nanoantenna according to claim 66, wherein the region not occupied by the nanoscale nucleic acid scaffold is 90% or more of the plasmon hotspot.

68. The plasmon nanoantenna according to any one of claims 61 to 67, wherein the region not occupied by the nanoscale nucleic acid scaffold is amorphous.

69. The plasmon nanoantenna according to any one of claims 61 to 67, wherein the region not occupied by the nanoscale nucleic acid scaffold is spherical or cubic.

70. A plasmon nanoantenna according to any one of claims 61 to 69, comprising a polynucleotide that forms part of the nanoscale nucleic acid scaffold or extends from the nanoscale nucleic acid scaffold, which is capable of hybridizing with another polynucleotide contained in or ligated with the nucleic acid analyte of the target, and so that when each of the polynucleotides hybridizes, the nucleic acid analyte is non-covalently bonded to the nanoscale nucleic acid scaffold, preferably by hydrogen bonds.

71. The plasmon nanoantenna according to claim 70, comprising sequences of polynucleotides capable of hybridizing with each other that are complementary or substantially complementary to each other.

72. A plasmon nanoantenna according to any one of claims 61 to 71, comprising a nucleic acid analyte bound to the nanoscale nucleic acid scaffold.

73. The plasmon nanoantenna according to any one of claims 61 to 72, wherein each of the two plasmon nanoparticles is independently selected from metal nanoparticles.

74. The plasmon nanoantenna according to claim 73, wherein each of the two plasmon nanoparticles is independently selected from the group consisting of gold, silver, aluminum, copper, palladium, and platinum nanoparticles or any one alloy thereof.

75. The plasmon nanoantenna according to claim 74, wherein each of the two plasmon nanoparticles is a gold nanoparticle.

76. The plasmon nanoantenna according to any one of claims 61 to 75, wherein the maximum diameter of each of the two plasmon nanoparticles is approximately 5 nm to approximately 500 nm.

77. The plasmon nanoantenna according to claim 76, wherein the maximum diameter of each of the two plasmon nanoparticles is approximately 50 nm to approximately 150 nm.

78. The plasmon nanoantenna according to claim 77, wherein the maximum diameter of each of the two plasmon nanoparticles is approximately 80 nm to 120 nm.

79. The plasmon nanoantenna according to any one of claims 61 to 78, wherein each of the two plasmon nanoparticles is coated by a coating.

80. The plasmon nanoantenna according to claim 79, wherein the coating substantially covers the surface of each of the two plasmon nanoparticles.

81. The plasmon nanoantenna according to claim 80, wherein the coating reduces the aggregation of the two plasmon nanoparticles by 50% or more compared to the two plasmon nanoparticles in the absence of the coating.

82. The plasmon nanoantenna according to any one of claims 79 to 81, wherein the coating comprises a plurality of oligonucleotides.

83. The plasmon nanoantenna according to claim 82, wherein each of the plurality of oligonucleotides comprises about 20 nucleotides to about 50 nucleotides.

84. The plasmon nanoantenna according to claim 83, wherein each of the plurality of oligonucleotides comprises about 20 to about 30 nucleotides.

85. The plasmon nanoantenna according to any one of claims 82 to 84, wherein each nucleotide in each oligonucleotide of the plurality of oligonucleotides is selected from pyrimidine nucleotides.

86. The plasmon nanoantenna according to any one of claims 61 to 85, wherein the nanoscale nucleic acid scaffold is substantially composed of double-stranded DNA, preferably interconnected parallel strands of a double helix.

87. The plasmon nanoantenna according to any one of claims 61 to 86, wherein the nanoscale nucleic acid scaffold is composed of a DNA origami structure.

88. The plasmon nanoantenna according to claim 87, wherein at least a portion of the DNA origami structure is assembled to form a U-shaped or curved structure.

89. The plasmon nanoantenna according to any one of claims 61 to 88, wherein the nanoscale nucleic acid scaffold includes a first surface to which one of the two plasmon nanoparticles is anchored, and a second surface of the scaffold to which a second plasmon nanoparticle of the two plasmon nanoparticles is anchored.

90. The plasmon nanoantenna according to claim 89, wherein the first surface of the scaffold and the second surface of the scaffold are on the same side or adjacent side of the nanoscale nucleic acid scaffold.

91. The plasmon nanoantenna according to claim 89, wherein the first surface of the scaffold and the second surface of the scaffold are on opposite sides of the nanoscale nucleic acid scaffold.

92. The two plasmon nanoparticles are anchored to the first surface of the scaffold and the second surface of the scaffold, respectively, via one or more nucleic acid linkers. Each nucleic acid linker is formed by an oligonucleotide capable of hybridizing with a polynucleotide that forms part of the nanoscale nucleic acid scaffold or extends from the nanoscale nucleic acid scaffold, which is coated on the surface of the plasmon nanoparticles. The sequences of the oligonucleotide and polynucleotide that can hybridize are complementary or substantially complementary to each other. A plasmon nanoantenna according to any one of claims 89 to 91.

93. The plasmon nanoantenna according to any one of claims 61 to 92, wherein the nanoscale nucleic acid scaffold comprises one or more polynucleotides extending from the first surface of the scaffold, and each of the one or more polynucleotides comprises a sequence complementary to the sequence of one or more oligonucleotides coating one of the two plasmon nanoparticles.

94. The plasmon nanoantenna according to any one of claims 61 to 93, wherein the nanoscale nucleic acid scaffold comprises one or more polynucleotides extending from the second surface of the scaffold, and each polynucleotide comprises a sequence complementary to the sequence of one or more oligonucleotides coating one of the two plasmon nanoparticles.

95. The plasmon nanoantenna according to claim 93 or 94, wherein one or more polynucleotides extending from the surface of the scaffold are independently selected from single-stranded DNA.

96. The plasmon nanoantenna according to any one of claims 61 to 95, wherein the nucleic acid polymerase is bound to or immobilized on the nanoscale nucleic acid scaffold via a nucleic acid linker.

97. The plasmon nanoantenna according to claim 96, wherein the nucleic acid linker is double-stranded and comprises (i) a polynucleotide covalently or non-covalently bonded to the nucleic acid polymerase via amino acids in the nucleic acid polymerase, and (ii) a polynucleotide forming part of the nanoscale nucleic acid scaffold or extending from the nanoscale nucleic acid scaffold, wherein the polynucleotide in (i) and the polynucleotide in (ii) comprise a single-stranded DNA sequence that is complementary or substantially complementary to each other and capable of hybridizing.

98. The plasmon nanoantenna according to claim 97, wherein the polynucleotide in (i) includes a peptide tag, and the peptide tag is bound to the nucleic acid polymerase via an isopeptide bond formed between an amino acid in the nucleic acid polymerase and an amino acid in the peptide tag.

99. The plasmon nanoantenna according to claim 97 or 98, wherein the nucleic acid linker is bound to the nucleic acid polymerase via an amino acid in the N-terminal domain of the nucleic acid polymerase.

100. The plasmon nanoantenna according to any one of claims 61 to 99, wherein the nucleic acid polymerase is DNA polymerase.

101. An array comprising a plurality of plasmon nanoantennas according to any one of claims 61 to 100.

102. The array according to claim 101, wherein the array includes a solid substrate, and the plurality of plasmon nanoantennas are bonded to the solid substrate.

103. The array according to claim 102, wherein each plasmon nanoantenna is immobilized on the solid substrate via its nucleic acid scaffold.

104. The array according to claim 102 or 103, wherein the solid substrate is preferably glass or silica in the form of a slide or chip.

105. The array according to any one of claims 101 to 104, wherein less than 30% of the plasmon nanoantennas are clustered in aggregates of two or more plasmon nanoantennas.

106. The array according to claim 105, wherein less than 15% of the plasmon nanoantennas are clustered in aggregates of two or more plasmon nanoantennas.

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