Systems and methods for capturing, extending, and detecting polynucleotide chains

Abstract

This invention provides a system and method for capturing and detecting polynucleotide chains. [Solution] The method may include the step of labeling at least one subsequence within a polynucleotide chain to obtain a labeled polynucleotide chain having at least one label. The method may further include the step of supplying a polymer to the surface of a channel and causing the polymer to physically interact with the surface of the channel. The method may further include the step of supplying a sample containing a labeled polynucleotide chain to the channel, the step of applying an electric field to the labeled polynucleotide chain to promote a physical interaction between the labeled polynucleotide chain and the polymer on the surface of the channel and to capture the labeled polynucleotide chain on the surface of the channel, and the step of detecting at least one label within the labeled polynucleotide chain.

Description

Technical Field

[0004] , , ,

[0003] , , ,

[0001] Cross-reference This application claims the benefit of U.S. Provisional Application Serial No. 63 / 702,095, filed on October 1, 2024, the disclosure of which is incorporated herein by reference in its entirety.

[0002] The present disclosure relates to systems and methods for controlling, capturing, stretching, and detecting polynucleotide chains.

Background Art

[0003] By controlling the flow and movement, capturing, stretching, and imaging of single-molecule DNA, it may be possible to conduct basic research on the physical properties of DNA polymers and the interaction between DNA and flow or electric fields. Current technologies require complex configurations, lack throughput, and are cost-ineffective. Furthermore, current methods often involve analyzing molecules moving within nanometer-scale channels, resulting in a decrease in the quality of detection. Therefore, there is a need for systems and methods that are cost-effective and enable high-throughput processing of samples to facilitate the control, capture, and visualization of single-molecule polynucleotide chains.

Summary of the Invention

[0004] In at least one embodiment, a method is provided for capturing and detecting a polynucleotide chain within a channel. This method may include the steps of: labeling at least one subsequence within a polynucleotide chain to obtain a labeled polynucleotide chain having at least one label; supplying a polymer to the surface of a channel; physically interacting the polymer with the surface of the channel; supplying a sample containing the labeled polynucleotide chain to the channel; applying an electric field to the labeled polynucleotide chain to facilitate a physical interaction between the labeled polynucleotide chain and the polymer on the surface of the channel to capture the labeled polynucleotide chain on the surface of the channel; and detecting at least one label within the labeled polynucleotide chain. At least one label may be optically detectable. At least one label may be a fluorescent label. At least one label may be detectable by a fluorescence microscope. At least one label may be detectable by one of the following methods: light scattering microscopy, total internal reflection microscopy (TIRF), super-resolution structural illumination microscopy (SR-SIM), stimulated emission suppression microscopy (STED), stochastic optical reconstruction microscopy (STORM), or single-molecule localization microscopy (SMLM). The labeled polynucleotide chain may be trapped at its vertices. The first and second free ends of the labeled polynucleotide chain may each extend outward from the vertex in the direction opposite to the electric field. The labeled polynucleotide chain may be greater than 1 kilobase pair. The channel surface may be flat. The polymer can alter the electroosmotic flow within the channel. The polymer may be a neutral, water-soluble polymer. The polymer may be a polyvinylpyrrolidone polymer. The polyvinylpyrrolidone polymer may have a molecular weight greater than 100 kDa. The neutral, water-soluble polymer may be hydroxyethylcellulose. The neutral, water-soluble polymer may be polyethylene glycol. The neutral, water-soluble polymer may be polyvinyl alcohol. The channel may have at least one dimension perpendicular to the electric field that is less than 5 microns. The electric field may be between 10 and 1,000 V / cm.The method may further include the step of supplying a pulse of pressure-driven flow to the labeled polynucleotide chain to facilitate the release of the labeled polynucleotide chain from physical interaction with the polymer. The method may further include the step of reducing the intensity of the electric field applied to the labeled polynucleotide chain to facilitate the release of the labeled polynucleotide chain from physical interaction with the polymer so that the labeled polynucleotide chain moves through the channel. The method may further include the steps of supplying a second sample containing a second labeled polynucleotide chain to the surface of the channel, increasing the intensity of the electric field to facilitate physical interaction between the second labeled polynucleotide chain and the polymer on the surface of the channel to trap the second labeled polynucleotide chain on the surface of the channel, and detecting at least one label within the second labeled polynucleotide chain to visualize the second labeled polynucleotide chain. This method may further include the steps of: reducing the intensity of an electric field applied to a second labeled polynucleotide chain to facilitate the release of the second labeled polynucleotide chain from physical interaction with the polymer so that the labeled polynucleotide chain moves through the channel; providing a sample containing the labeled polynucleotide chain; increasing the intensity of the electric field to capture and detect the labeled polynucleotide chain; and repeating the steps of reducing the intensity of the electric field to release the labeled polynucleotide chain so that the labeled polynucleotide chain moves through the channel n times.

[0005] In another embodiment, a system is provided for capturing, extending, and detecting a labeled polynucleotide chain. The system may comprise a device comprising at least one reservoir sized to accommodate a sample containing an electrode and / or a labeled polynucleotide chain, a fluid tip having at least one channel, the at least one channel being in fluid communication with at least one reservoir so that the sample can flow through at least one channel, and a channel cover. The system may also comprise a voltage source capable of applying an electric field to the channel and at least one detector capable of detecting a signal from the device. The system may also comprise a controller programmed to guide a flow of polymer into the device to facilitate the delivery of the polymer to the channel surface so that the polymer can physically interact with the channel surface, to guide a flow of sample from at least one reservoir to the channel, to form an electric field to facilitate physical interaction between the labeled polynucleotide chain and the polymer, to generate a voltage difference between an electrode and at least one other electrode to capture the labeled polynucleotide chain on the channel surface, and to interact with at least one detector to collect, record, and store a signal received from at least one detector, and to process the signal received from at least one detector. The detector may be a camera. The controller may be further programmed to reduce the intensity of the electric field applied to the labeled polynucleotide chain to facilitate the release of the labeled polynucleotide chain from physical interaction with the polymer. The controller may be further programmed to allow the released polynucleotide chain to move through the channel. The controller may be further programmed to induce a flow of a second sample containing a second polynucleotide chain from at least one reservoir fluid-communicated with the channel, and to increase the electric field intensity to capture the second polynucleotide chain. The controller may be further programmed to supply pulses of pressure-driven flow to the device to increase the release rate of the polynucleotide chain. The channel cover may be transparent.

[0006] In yet another embodiment, a method is provided for capturing and detecting polynucleotide chains within a channel. The method may include the step of supplying a polymer to the channel surface. The method may further include the step of physically interacting the polymer with the channel surface and the step of supplying a sample containing polynucleotide chains to the channel. The method may also include the step of labeling at least one subsequence within a polynucleotide chain to obtain a labeled polynucleotide chain having at least one label. The method may further include the steps of applying an electric field to the labeled polynucleotide chain to facilitate a physical interaction between the labeled polynucleotide chain and the polymer on the channel surface to capture the labeled polynucleotide chain on the channel surface and the step of detecting at least one label within the labeled polynucleotide chain. [Brief explanation of the drawing]

[0007] [Figure 1] Figures 1A and 1B illustrate methods for capturing and extending polynucleotide chains. [Figure 2] Figures 2A and 2B illustrate methods for capturing and extending polynucleotide chains. [Figure 3] Figures 3A and 3B show a system for capturing and extending polynucleotide chains. [Figure 4] Figure 4 shows a channel having multiple fluid layers. [Figure 5] Figure 5 shows an example of a channel having a low electric field (E) region and a high electric field region. [Figure 6] Figure 6 shows an example of a channel having a surface to which a polymer can be adsorbed and a defined region of patterned material to which the polymer does not adsorb. [Figure 7] Figures 7A and 7B show a method for optically imaging polynucleotide chains trapped at their vertices for the purpose of determining the sequence of the polynucleotide chains. [Figure 8] Figure 8 shows a system for optically imaging polynucleotide chains captured at their vertices, with the aim of determining the sequence of the polynucleotide chains. [Figure 9] Figures 9A to 9C show a DNA capture method according to one embodiment. [Figure 10] Figures 10A-10B show an overview of the image processing flowchart used to quantify the amount of DNA captured. [Figure 11] Figures 11A and 11B show the analysis of the relaxation dynamics of vertex-pinned DNA when the electric field is removed. Figures 11C and 11D show the characteristic length statistical moments when the pinned DNA molecule relaxes. [Figure 12] Figure 12 shows a typical cycle of DNA vertex pinning under high electric fields. [Figure 13] Figures 13A to 13G show that a single DNA molecule is apex-pinned to the wall at a position that changes randomly in each example. [Figure 14] Figures 14A to 14F show a custom microfluidic interface device according to one embodiment. [Figure 15] Figures 15A and 15B show DNA apex pinning at the bottom and top surfaces of a 3 μm deep linear channel under an axial electric field strength of 150 V / cm. [Figure 16] Figure 16A is a schematic diagram of a commercially available glass chip purchased from Microfluidic ChipShop. Figures 16B and 16C show serial epifluorescence images of DNA electrophoresed through a commercially available glass channel with a depth of 37 μm. Figure 16D shows the global temporal median of the image sequence. [Figure 17] Figure 17 shows a vertex-pinned monomolecule of 20 kbp DNA in a 0.9 μm deep microfluidic channel filled with linear polymer buffer, to which an axial electric field has been applied. [Figure 18] Figures 18A-18D show a comparison between the area-averaged alpha-shape boundary intensity I and the results obtained by manually counting molecules. [Figure 19]Figure 19 shows the experimental quantification of the amount of vertex-pinned single-molecule DNA as a function of electric field strength. [Figure 20] Figures 20A to 20D show the experimental quantification of the amount of vertex-pinned single-molecule DNA as a function of time and electric field strength. [Figure 21] Figure 21 shows the relaxation dynamics of vertex-pinned single-molecule DNA. [Figure 22] Figures 22A-22J show an overview of the automated image processing used to quantify 48.5 kbp DNA relaxation. [Figure 23] Figures 23A to 23J show an overview of the automated image processing used to quantify 20kbpDNA relaxation. [Figure 24] Figures 24A-24C show the colocalization analysis of image intensity of vertex-pinned DNA in three different examples. [Figure 25] Figures 25A-25C show the colocalization analysis of image intensity of vertex-pinned DNA across three consecutive images from a single example. [Figure 26] Figure 26 shows an example of a frame from this video, illustrating the difference between the raw video and the moving median video, which has been processed to display only stationary DNA molecules. [Modes for carrying out the invention]

[0008] Where necessary, detailed embodiments of the Disclosure are disclosed herein, but it should be understood that the disclosed embodiments are merely illustrative examples of the Disclosure, which can be carried out in various alternative forms. The figures are not necessarily to scale, and some features may be exaggerated or minimized to illustrate the details of certain components. Accordingly, the details of certain structures and functions disclosed herein should not be construed as limiting, but merely as representative grounds to teach those skilled in the art to employ the Disclosure in various ways.

[0009] Unless otherwise indicated in the Examples or explicitly stated, all numerical values indicating amounts of substances, reactions and / or conditions of use in this specification are to be understood as being modified by the term "about". The initial definition of an acronym or other abbreviation applies to all subsequent uses of the same abbreviation in this specification and also to normal grammatical variations of the initially defined abbreviation. Also, unless explicitly stated to the contrary, measurements of properties are determined by the same techniques as those previously or later referenced for the same property.

[0010] Unless otherwise indicated, all technical and scientific terms used in this specification have the same meaning as commonly understood by one of ordinary skill in the art to which this disclosure pertains.

[0011] It should also be understood that, because specific components and / or conditions may of course vary, the present disclosure is not limited to the specific embodiments and methods described below. Further, the terms used in this specification are used only to describe particular embodiments and are not intended to be limiting in any sense.

[0012] It should also be noted that, as used in this specification and the appended claims, the singular forms "a", "an", and "the" include plural referents unless the context clearly dictates otherwise. For example, reference to a single component is intended to include a plurality of components.

[0013] The words "or" and "and" can be used interchangeably and are to be understood to mean "and / or".

[0014] The term "comprising" is synonymous with "including", "having", "containing", or "characterized by". These terms are inclusive and open-ended and do not exclude additional, unrecited elements or method steps.

[0015] The phrase "consists of..." excludes elements, steps, or components not explicitly stated in the claim. If this phrase appears in a clause of the claim text rather than immediately following the preamble, it limits the claim to only the elements specified in that clause, and does not exclude other elements from the claim as a whole.

[0016] The phrase "essentially consisting of" limits the scope of the claim to the specified materials or steps, in addition to those that do not affect the fundamental and novel characteristics of the subject matter of the claim.

[0017] The terms “polynucleotide,” “nucleotide,” “nucleotide sequence,” “nucleic acid,” “polynucleic acid,” and “oligonucleotide” may be used interchangeably in this disclosure. These refer to polymeric forms of nucleotides of any length, deoxyribonucleotides or ribonucleotides, or analogs thereof. Polynucleotides have any three-dimensional structure and may perform any known or unknown function. Non-limiting examples of polynucleotides include single-stranded, double-stranded, or multi-stranded DNA or RNA, genomic DNA, cDNA, DNA amplicon products, DNA restriction products, peptide nucleic acids (PNAs) and Loc nucleic acids (LNAs), synthetic non-natural polynucleotide chains, DNA-RNA hybrids, or polymers consisting of purine and pyrimidine bases, or other natural, chemically or biochemically modified, non-natural, or derivatized nucleotide bases. The terms “polynucleotide” and “nucleic acid,” when applied to the embodiments described, should be understood to include single-stranded (e.g., sense or antisense) and double-stranded polynucleotides. Polynucleotides may include one or more modified nucleotides, such as methylated nucleotides or nucleotide analogs. If present, modifications to the nucleotide structure may be conferred before or after polymer assembly. The nucleotide sequence may be interrupted by non-nucleotide components. Polynucleotides may be further modified after polymerization, such as by binding with labeling components.

[0018] The term "complementarity" refers to the ability of a nucleic acid to form hydrogen bonds with another nucleic acid sequence, either through the conventional Watson-Crick model or other unconventional methods. The percentage of complementarity indicates the proportion of residues in a nucleic acid molecule that can form hydrogen bonds (e.g., Watson-Crick base pairs) with a second nucleic acid sequence (for example, 4, 5, and 6 out of 6 would represent 66.67%, 83.33%, and 100% complementarity, respectively). "Perfectly complementary" means that every consecutive residue in one nucleic acid sequence forms hydrogen bonds with the same number of consecutive residues in the second nucleic acid sequence. As used herein, “substantially complementary” refers to a degree of complementarity over 4, 5, 6, 7, and 8 nucleotide regions of at least 40%, 50%, 60%, 62.5%, 70%, 75%, 80%, 85%, 90%, 95%, 97%, 98%, 99%, or 100%, or any percentage in between, or to two nucleic acids that hybridize under stringent conditions.

[0019] Unless otherwise expressly stated, the term “polymer” includes “oligomer,” “copolymer,” “terpolymer,” etc., the molecular weight of a polymer refers to the weight-average molecular weight unless otherwise specified, and in relation to the present invention, a description of a group or class of materials that is suitable or preferred for a given purpose means that a mixture of any two or more components of that group or class is equally suitable or preferred, the initial definition of an acronym or other abbreviation applies to all subsequent use of the same abbreviation herein and also applies mutatis mutandis to ordinary grammatical variations of the abbreviation initially defined. Furthermore, unless otherwise expressly stated, the measurement of properties is determined by the same method as previously or later referenced for the same property.

[0020] Furthermore, it should be understood that an integer range explicitly includes all intervening integers. For example, the integer range 1 to 10 explicitly includes 1, 2, 3, 4, 5, 6, 7, 8, 9, and 10. Similarly, the range 1 to 100 includes 1, 2, 3, 4...97, 98, 99, and 100. Likewise, if an arbitrary range is required, the difference between the upper and lower limits divided by 10 can be used as a substitute for the upper or lower limit. For example, if the range is 1.1 to 2.1, then 1.2, 1.3, 1.4, 1.5, 1.6, 1.7, 1.8, 1.9, and 2.0 can be selected as the lower or upper limit. In the specific examples described herein, concentrations, temperatures, and reaction conditions (e.g., pressure, pH, etc.) can be carried out with a tolerance of plus or minus 50% of the indicated values, rounded to three significant figures. In further improvements, the concentration, temperature, and reaction conditions (e.g., pressure, pH, etc.) can be adjusted to within plus or minus 30% of the indicated values, rounded to three significant figures. In further improvements, the concentration, temperature, and reaction conditions (e.g., pH, etc.) can be adjusted to within plus or minus 10% of the indicated values, rounded to three significant figures.

[0021] In the examples described herein, concentrations, temperatures, and reaction conditions (e.g., pressure, pH, flow rate, etc.) may be expressed with two significant figures, rounded or truncated to within plus or minus 50% of the indicated values. Further improvements may be made, and concentrations, temperatures, and reaction conditions (e.g., pressure, pH, flow rate, etc.) may be expressed with two significant figures, rounded or truncated to within plus or minus 30% of the indicated values. Further improvements may be made, and concentrations, temperatures, and reaction conditions (e.g., pressure, pH, flow rate, etc.) may be expressed with two significant figures, rounded or truncated to within plus or minus 10% of the indicated values.

[0022] The capture and extension of single-molecule DNA may enable fundamental research into the physical properties of DNA polymers and the interaction between DNA and fluids or electric fields. Methods for capturing, extending, and simultaneously visualizing single-molecule DNA include using optical or magnetic tweezers to manipulate microbeads attached to the DNA, applying stretching flow strain to the DNA within microfluidic devices, introducing chemical reactions to tether DNA to a flat surface (such as a DNA curtain) and exposing it to fluid flow or electric fields, and using a moving air / water interface to comb DNA on silanized glass. The sealing and capture of single-molecule DNA, combined with high-resolution imaging, is also applied to genetic analysis such as karyotype analysis and DNA mapping. For example, electrophoresis within nanochannels and imaging of long (over 100kb) genomic DNA using sequence-specific labeling are used for optical mapping in medical diagnostics.

[0023] However, these techniques also have drawbacks. For example, throughput is insufficient. Optical tweezers and magnetic tweezers can only study a single molecule, or at best a few molecules. These techniques also require expensive and complex imaging and control systems. Molecular combing and DNA curtain assays can image many molecules, but they cannot process multiple samples cyclically. Therefore, these assays cannot resample the same population or analyze new samples. Thus, multiplexing of data analysis is necessary to capture information about fast-moving molecules. Furthermore, because molecules are not stationary (moving molecules), it is difficult to obtain high-quality images or study dynamic interactions in these techniques. Other problems with using electrophoresis in nanochannels (including so-called nanoslit channels) for DNA isolation include the difficulty of channel fabrication and sealing, and the difficulty of controlling the flow within the system, such as liquid filling, pressure-driven flow generation, and dynamic current generation. Also, depending on the nanochannel fabrication method, optical access to the channel may be poor. Furthermore, the walls of such nanochannels may have roughness elements on the order of the channel's cross-sectional dimensions. Another challenge is that the non-uniform surface properties of nanochannels lead to non-uniform zeta potentials, resulting in non-uniform electroosmotic flow velocities and the creation of velocity gradients within the fluid. Current methods for DNA capture (with or without DNA visualization), including optical and magnetic tweezers as described above, and / or DNA adsorption to liquid / air interfaces using so-called DNA combing or DNA curtain techniques, can be cumbersome manual processes and are unsuitable for high-throughput applications where numerous DNA molecules need to be individually captured and spread. Therefore, there is a need for cost-effective systems and methods that facilitate the control and detection of single-molecule polynucleotide chains, enabling high-throughput processing of samples.

[0024] This specification discloses systems and methods for capturing, extending, releasing, and / or recapturing numerous single polynucleotide chains. The disclosed systems and methods can facilitate the control and detection (including visualization) of single polynucleotide chains.

[0025] This specification discloses methods for capturing, extending, releasing, and / or recapturing multiple single-molecule polynucleotide chains. A polynucleotide chain is a chain of linked nucleotides, such as DNA or RNA. Polynucleotide chains are also called chains or molecules. One or more methods can facilitate the control and visualization of single molecules. According to embodiments of various methods, polynucleotide chains in a sample can be fixed to the wall of a microchannel at their vertices along their length. The polynucleotide chains can be captured, held in place, and extended using an applied axial electric field. This enables high-quality imaging and quantification of the three-dimensional structure of the polynucleotide chains. In one or more embodiments, this disclosure presents a set of conditions for the capture of polynucleotide chains, quantifying the capture dynamics, and quantifying the relaxation dynamics, separation, and release from the pinned state.

[0026] According to at least one embodiment, a method for capturing and extending polynucleotide chains is disclosed. This method facilitates the capture of polynucleotide chains on the surface of a channel. The channel may be a microfluidic channel. As used herein, the term channel also includes a fluid chamber that is in fluid communication with at least one port used to fill the chamber with a liquid. The channel may be part of a microfluidic device having one or more channels. One or more channels may each have one input and one output. There may be multiple independent channels. Samples of each independent channel may be processed in parallel. The surface of the channel may be a dielectric, including an oxide. For example, the channel surface may be silicon oxide, aluminum oxide, titanium oxide, or various other oxide surfaces. The surface of the channel may be a nitride, such as silicon nitride. Alternatively, the surface may be glass, including borosilicate glass, or quartz. The surface of the channel may be dielectric, and there may be non-dielectric (such as metal or semiconductor) regions. Dielectric and non-dielectric materials may be arranged on the surface in a specific pattern. Polynucleotide chains may be captured in the dielectric regions but not in the non-dielectric regions. The channel may include a channel cover. The channel cover may be borosilicate glass. The channel may include a base and one or more walls extending outward from the base. The walls and / or cover of the channel may be flat. The surface of the channel may include a surface etched onto the substrate or channel cover. Alternatively, the surface of the channel may include a defined portion of the substrate, any wall of the channel, or the cover of the channel. In certain embodiments, the channel depth may be from 500 nm to 3 μm. Channel depths in this range can facilitate the easy acquisition of high-quality images of polynucleotide chains, for example, by epifluorescence microscopy. In other embodiments, the channel may have a depth greater than 3 μm. High-resolution images of polynucleotide chains can be acquired, for example, using TIRF microscopy (total internal reflection fluorescence) with a channel depth greater than 3 μm. In various embodiments, the channel may also have a nominal span width greater than 1 μm.The channel may optionally include one or more supports for supporting the cover, and / or castellation patterns on one or more side walls of the channel. The channel may have at least one dimension perpendicular to the applied electric field that is less than 5 microns.

[0027] The transport of reagents and / or liquids within a system can be achieved using processes including electrophoresis, electroosmotic flow, pressure-driven flow, dielectrophoresis, magnetophoresis, electrohumidification, induced charge electroosmotic flow, alternating current (AC) dynamic current, gravity-driven flow, and surface tension-driven flow.

[0028] This method may include the step of supplying a polymer to the channel surface. The polymer can suppress electroosmotic flow within the channel. The polymer may be a neutral, water-soluble linear polymer. The polymer may be, for example, polyvinylpyrrolidone (PVP). PVP is a neutral, water-soluble linear polymer used as an additive in buffer systems to suppress electroosmotic flow (EOF) for electrodynamic experiments. The polymer may also be polyethylene glycol (PEG), also known as polyethylene oxide (PEO) in the field of polymer chemistry. PEG is used in gas chromatography as well as for coating and treating silica surfaces to suppress EOF. PEG with a sufficiently large molecular weight can also achieve DNA apex pinning. The polymer may also be hydroxyethylcellulose (HEC). HEC can suppress EOF as a linear polymer. The polymer may also be polyvinyl alcohol (PVA). PVA can be used for the separation of DNA restriction fragments and effectively suppresses EOF at pH less than 8.

[0029] This method may further include a step of physically interacting the polymer with the channel surface. This step can be achieved by coating the surface with the polymer. Coating can be achieved by delivering the polymer to the surface and adsorbing the polymer onto the surface. The surface may include the floor or walls of the channel, a portion of the floor or walls of the channel, the channel cover or a portion of the channel cover, or a surface that is not in physical contact with the channel cover. For example, the polymer can adsorb to the silicon oxide walls of the channel or the borosilicate glass walls of the channel cover. In at least some embodiments, a metal to which the polymer does not adsorb may be deposited in a pattern on the oxide substrate. In this way, only regions of the oxide substrate (e.g., not patterned with metal) where there is a sufficient electric field and linear polymers interacting with the surface can capture polynucleotide chains.

[0030] The method may further include the step of supplying a sample containing polynucleotide chains to a polymer on the surface of a channel. The sample may contain multiple polynucleotide chains. The polynucleotide chains may be isolated or synthesized. The polynucleotide chains may originate from the same source or from different sources. For example, different sources may contain polynucleotide chains from different organisms (e.g., humans) or from the same organism. A single sample may contain polynucleotide chains from both a control and an experiment. The polynucleotide chains in the sample may include a barcode that identifies the source or type of the polynucleotide chain. Sources may include, for example, human, plant, and animal cells and tissues, organoids, spheroids, tumor cells and other microsystems, bacterial and protozoan cells, and synthetic reactions that produce polynucleotide chains. The sample containing polynucleotide chains may further include one or more buffering components. In at least one embodiment, the sample supplied to the channel may include polynucleotide chains, buffering components, and a polymer.

[0031] The method further includes the step of applying an electric field to the polynucleotide chain to promote physical interaction between the polynucleotide chain and the polymer, thereby trapping the polynucleotide chain on the surface of the channel (also called pinning in one or more embodiments). The electric field can be 30–400 V / cm. The electric field can be applied to the electrolyte by two or more electrodes supporting the Faraday reaction. Such a Faraday reaction can promote ionic currents in the channel. The applied magnetic field associated with these ionic currents may be substantially parallel to the wall in which the polynucleotide chain is trapped. For example, the inner wall of the channel may be a dielectric material, and the applied electric field may be parallel to the wetted wall of the channel, driving the movement of the polynucleotide chain parallel to the channel wall (including electrophoresis). Alternatively, the electric field promoting the interaction between the polynucleotide chain and the wall may be applied using electrodes capacitively coupled to the electrolyte, thereby avoiding or mitigating the Faraday reaction in the system. The electric field is applied to the channel and affects the channel, as well as the contents of the channel, including the fluid sample. For example, applying an electric field to a channel will apply the electric field to the polynucleotide chains in the sample flowing through the channel. Physical interactions between the polynucleotide chains and the polymer can occur at the vertices of the polynucleotide chains. Physical interactions can occur at a single vertex along the length of the polynucleotide chain. The ends of the polynucleotide (also called arms) can extend outward from the vertex in the opposite direction to the applied electric field.

[0032] Pinning and extension of polynucleotide chains occurs when a polymer with a sufficiently large molecular weight (e.g., 100 kDa or more) is supplied to the channel surface and a sufficiently large electric field (e.g., 30 V / cm) is applied axially. The electric field threshold decreases with increasing molecular weight. For example, the electric field threshold is low for high molecular weight PVP polymers. The phenomena of capture and extension occur only on the surface and occur on both thermally oxidized silicon and glass surfaces.

[0033] While not supporting any specific theory, under sufficiently high electric fields, polynucleotide chains can become entangled with polymers adsorbed on their surface, and strong electric forces can form a hydrated layer between the nucleic acid and the polymer, resulting in a high-friction state. When the electric field is removed, the space between the nucleic acid and the polymer is rehydrated, and the resulting decrease in friction allows Brownian motion to separate the polynucleotide chains from the polymer. Since no pinpoint spatial correlation is observed in each experiment, this phenomenon is suggested to be independent of specific channel shapes.

[0034] The method may also include a step of reducing or removing the electric field to below an empirically determined threshold to encourage the release of the polynucleotide chain from physical interaction with the polymer. Once the electric field above the empirically determined threshold is removed, the polynucleotide chain relaxes toward and around the pinning site. The pinning site may be a single-point entanglement between the polynucleotide chain and the polymer. After the polynucleotide chain relaxes toward or around the pinning site, it detaches from the polymer and begins to move through the channel. The movement of the detached polynucleotide chain through the channel may occur by diffusion or electrophoresis. The relaxation, separation, and movement of the polynucleotide chain allow it to be removed from the channel, and a second sample containing the polynucleotide chain can be supplied to the channel for capture and extension. Thus, the method may include a periodic configuration comprising the steps of introducing a polynucleotide chain into a region of interest, capturing the polynucleotide chain with a high electric field, imaging the polynucleotide chain one or more times (including detecting various wavelengths), lowering the electric field to relax the polynucleotide chain and make it separable, applying an electric field to electrophoresthetize the polynucleotide chain with little or no capture, and repeating the process. In this way, the next sample containing a polynucleotide chain can be input into the channel. For example, after the first sample is captured, relaxed, separated, and cleared, a second sample can be supplied to the channel for capture. After that sample is captured, relaxed, separated, and cleared, the next sample can be supplied to the channel for capture. The process of capture, relaxation, separation, and clearing can be repeated for n samples. This periodic configuration enables high-throughput processing of a large number of independent samples, and also allows for resampling the same sample multiple times to improve output. The second sample, subsequent sample, or nth sample may contain polynucleotide chains that were present in the first or previous sample. Furthermore, introducing pulses of pressure-driven flow can lead to faster separation.In some embodiments, the step of deactivating the electric field may be replaced with a step of supplying pulses of pressure-driven flow to the channel. The periodic configuration of the steps in the method enables high-throughput capture and analysis of a large number of polynucleotide chains.

[0035] Figure 1A shows a method 100 for capturing and extending a polynucleotide chain according to an embodiment. This method includes step 102 of supplying a polymer to the surface of a channel. This method includes an additional step 104 of causing the polymer to physically interact with the surface of the channel. Step 106 includes supplying a sample containing a polynucleotide chain to the polymer that interacts with the surface of the channel. This method further includes step 108 of applying an electric field to the polynucleotide chain to facilitate a physical interaction between the polynucleotide chain and the polymer, thereby capturing the polynucleotide chain on the surface of the channel.

[0036] Figure 1B shows a method 101 for capturing and extending polynucleotide chains at high throughput according to an embodiment. This method facilitates the capture, extension, and release of polynucleotides and allows for sequential processing of multiple samples. The method includes step 102 of supplying a polymer to the surface of a channel. The method includes an additional step 104 of physically interacting the polymer with the surface of the channel. Step 106 includes supplying a sample containing polynucleotide chains to the polymer interacting with the surface of the channel. The method further includes step 108 of applying an electric field to the polynucleotide chains to facilitate physical interaction between the polynucleotide chains and the polymer, thereby capturing the polynucleotide chains on the surface of the channel. Step 109 includes changing the electric field to release the captured polynucleotide chains from physical interaction with the polymer. Lowering the electric field causes the polynucleotide chains to relax, resulting in their unwinding, free diffusion and electrophoresis occurring to clear the polynucleotide chains, and allowing the next sample containing polynucleotide chains to be introduced into the channel. The steps of the method can then be repeated n times to achieve high throughput processing of polynucleotide chains. In an alternative embodiment, step 109 may include supplying a pulse of pressure-driven flow to the channel. The pulse can also release the polynucleotide chains by promoting relaxation, separation, free diffusion, and electrophoresis of the polynucleotide chains.

[0037] Figure 2A shows an additional method 110 for capturing and extending a polynucleotide chain according to one embodiment. This method includes step 112 of supplying a sample containing a polymer and a polynucleotide chain to the surface of a channel. Step 114 includes physically interacting the polymer with the channel surface. An additional step 116 includes applying an electric field to the polynucleotide chain to facilitate physical interaction between the polynucleotide chain and the polymer, thereby capturing the polynucleotide chain on the surface of the channel.

[0038] Figure 2B shows an additional method 111 for capturing and extending polynucleotide chains at high throughput according to one embodiment. This method facilitates the capture, extension, and release of polynucleotides, allowing for the continuous processing of multiple samples. The method includes step 112 of supplying a sample containing a polymer and polynucleotide chains to the surface of a channel. Step 114 includes physically interacting the polymer with the channel surface. An additional step 116 includes applying an electric field to the polynucleotide chains to facilitate physical interaction between the polynucleotide chains and the polymer, thereby capturing the polynucleotide chains on the surface of the channel. Step 117 includes changing the electric field to release the captured polynucleotide chains from physical interaction with the polymer. Lowering the electric field causes the polynucleotide chains to relax, resulting in their unwinding, free diffusion and electrophoresis occurring to clear the polynucleotide chains, and allowing the next sample containing polynucleotide chains to be introduced into the channel. The steps of the method can then be repeated n times to achieve high-throughput processing of polynucleotide chains. In an alternative embodiment, step 117 may include supplying a pulse of pressure-driven flow to the channel. Pulses can also release polynucleotide chains by promoting relaxation, separation, free diffusion, and electrophoresis of the polynucleotide chains.

[0039] Figures 3A and 3B illustrate a system for capturing and extending polynucleotide chains. Figure 3A shows a system 118 comprising a device 120, a voltage source 122, and a controller 124 programmed to interact with the device 120 and the voltage source 122. The device 120 may include at least one reservoir sized to accommodate a sample containing polynucleotide chains. The reservoir may include electrodes. Alternatively, the device 120 may include a first reservoir sized to accommodate a sample containing polynucleotide chains and a second reservoir sized to accommodate electrodes. These reservoirs may be electrically and / or ionically connected to each other or to other electrodes in the system. The electrodes (either alone in the reservoir or together with the sample containing polynucleotide chains) may be, for example, metals, metallic alloys, or semiconductor materials. Other possible materials include graphite, graphene, or other forms of carbon, including activated porous carbon. Electrodes can be fabricated using microfabrication techniques such as vapor deposition, lift-off methods, or doping. The electrodes may be electroplated or coated. These electrodes can drive a Faraday reaction (including water splitting at the electrodes) to introduce an electric field into the system's channels. The electrodes may also have high capacitance, including porous carbon electrodes that operate at low voltages to avoid the Faraday reaction. In at least some embodiments, the electrodes may be platinum electrodes. As previously mentioned, one or more electrodes may be carbon-based. For example, the electrodes may be monolayer or multilayer graphene. The electrodes may be graphite. The electrodes may also be inkjet-printed carbon-deposited lead. Alternatively, the electrodes may be gold, silver, titanium, palladium, copper, stainless steel, titanium nitride, or silver / silver chloride. The electrodes may also be fabricated from doped silicon. The voltage waveform applied from the electrodes and the resulting electric field may be direct current (DC), alternating current (AC), pulse-modulated, or a very complex waveform. For example, the waveform may contain both DC and AC components and may be periodic, aperiodic, antiperiodic, or non-antiperiodic.

[0040] The device may include a chip. The chip may be a silicon substrate chip. The chip may have two isolated (independent) single-input, single-output channels. The channels may be manufactured so that they can be visualized simultaneously within the same field of view near the geometric center of the silicon chip. The microfluidic channels may be dry-etched into a silicon wafer to a depth of, for example, 1 μm. The microfluidic channels may be in fluid contact with a reservoir. There may be multiple independent channels. Samples from each independent channel may be processed in parallel. The device may include a channel cover. The channel cover may be made of glass. The channel cover may be, for example, borosilicate glass. One or more walls of the channel may be optically transparent and / or facilitate efficient transmission to electromagnetic radiation such as infrared and ultraviolet. One or more walls of the channel may facilitate significant transmission, reflection, refraction, and / or diffraction of electromagnetic radiation. Electromagnetic radiation as defined herein includes optical (visible) wavelengths, infrared, and ultraviolet. The system may allow and focus a magnetic field that can impart magnetism to the device containing magnetic particles.

[0041] The system may also include a voltage source and a controller. The voltage source can apply an electric field to the polynucleotide chain in the channel. The voltage source can apply a voltage using electrodes in a reservoir. The voltage source can also measure current. The controller may be programmed to interact with the voltage source. For example, the controller may be programmed to instruct the voltage source to apply a voltage or to measure current. The controller may be programmed to change the electric field applied to the channel. Furthermore, the controller may be programmed to guide a flow of polymer to the device to facilitate the delivery of the polymer to the channel surface so that the polymer can physically interact with the channel surface. The controller may similarly guide a flow of sample to the device with or without polymer to facilitate the delivery of the sample to the channel surface. The controller may also be programmed to instruct electrophoresis of the polynucleotide chain through the channel. Figure 3B shows a system 119 for capturing and extending a polynucleotide chain according to one embodiment. The voltage source 122 is part of the controller 124.

[0042] The processes, methods, or algorithms disclosed herein are available for or implemented in processing units, controllers, or computers, which may include existing programmable electronic control units or dedicated electronic control units. Similarly, the processes, methods, or algorithms can be stored as executable data and instructions by a controller or computer in many forms, including, but not limited to, information permanently stored in non-writable storage media such as ROM devices, and information modifiablely stored in writable storage media such as floppy disks, magnetic tapes, CDs, RAM devices, or other magnetic or optical media. The processes, methods, or algorithms can also be implemented as executable software objects. Alternatively, the processes, methods, or algorithms can be embodied, in whole or in part, using appropriate hardware components such as ASICs (Application-Specific Integrated Circuits), FPGAs (Field-Programmable Gate Arrays), state machines, controllers, or other hardware components or devices, or combinations of hardware, software, and firmware components.

[0043] Methods for spatially patterning the locations within a channel where polynucleotide chains can be trapped are also provided herein. These methods allow polynucleotides to be trapped within a specific region of interest. In one embodiment, the method includes the step of supplying a polymer to a specific location in a fluid system. For example, laminar flow functionality may be utilized to supply the polymer to only a portion of the subsurfaces, preventing the polymer from physically interacting with other surfaces. This method can utilize hydrodynamic focusing to pattern the locations of the polymer-treated walls. Figure 4 shows a channel 126 having multiple fluid layers 128a, 128b, and 128c. The first fluid layer 128a contains polynucleotide chains but no polymer. The second fluid layer 128b contains both polynucleotide chains and polymer. The third fluid layer 128c contains polynucleotide chains but no polymer. Because the second fluid layer 128b contains polymer, the polynucleotides are trapped in the second fluid layer 128b. In the first layer 128a and the third layer 128c, the polynucleotide chains move in a 3D ball shape.

[0044] In alternative embodiments, a method for spatially patterning the locations within a channel where a polynucleotide chain can be trapped may include shaping the channel walls and / or adding features that affect the electric field in a specific defined region. For example, obstacles may be added to have a local effect on the channel's electric field. This method takes advantage of the fact that polynucleotide chain trapping occurs above a certain threshold electric field. Figure 5 shows an example of a channel 130 having low electric fields 132a, 132b and a high electric field 134. In the high electric field region, only the vertices of the polynucleotide chain are fixed and extended, while in the low electric field region, the polynucleotide chain moves in a 3D ball shape.

[0045] In yet another embodiment, a method for spatially patterning locations within a channel where polynucleotide chains can be captured may include patterning a metal-coated region on a fluid tip where the polymer does not adsorb. Figure 6 shows an example of a channel 136 having a polymer-adsorbable surface 138 and a defined region of patterned material 140 where the polymer does not adsorb.

[0046] Systems and methods for capturing, extending, and detecting polynucleotide chains The systems and methods of one or more embodiments disclosed herein describe systems and methods for apex pinning, relaxation, separation, and free electrophoresis cycles for capturing and extending single polynucleotide chains. Apex pinning and extension may cycle through many polynucleotide chains, interspersed with relaxation, separation, and low-field clearing of the polynucleotide chains. These systems and methods may be further modified to detect polynucleotide chains. For example, these systems and methods can be modified to achieve high-throughput capture, extension, and imaging of polynucleotide chains. They may also be modified for the purpose of determining the sequence of polynucleotide chains. The systems and methods can facilitate the capture and extension of polynucleotide chains in a specific region of interest within a channel. The specific region of interest may be, for example, a field of view within a channel.

[0047] Figures 7A and 7B illustrate a method 142 for optically imaging a vertex-captured polynucleotide chain for the purpose of visualizing a subsequence of the polynucleotide chain. Step 144 includes providing a sample containing at least one polynucleotide chain and labeling the polynucleotide chain to create a labeled polynucleotide chain. The polynucleotide chain may be significantly longer than 1 kb. For example, the polynucleotide chain may be longer than 300 kb. A labeled subsequence can be formed by labeling at least one sequence of 4 to 10 base pairs within the polynucleotide chain. Thus, a labeled polynucleotide chain is a polynucleotide chain having at least one labeled subsequence. As an example, the polynucleotide chain may be DNA. The region containing the subsequence may be labeled with a fluorescent label. Multiple subsequences may be labeled on a DNA chain. Subsequences are also called target sequences, subsequences, codes, or barcode elements. Multiple types of labeling can be used, including fluorescent labels with different emission wavelengths. Step 146 includes capturing and visualizing the labeled polynucleotide chain. Step 146 includes substeps 147, 148, 150, and 152. Substep 147 includes supplying the channel with a polymer that can alter the electroosmotic flow within the channel. Substep 148 includes supplying the channel with a sample containing a labeled polynucleotide chain. In some embodiments, the polymer may be contained in the sample containing the labeled polynucleotide chain. The polymer may be neutral and water-soluble. Next, an electric field exceeding an empirically determined threshold is applied to the polynucleotide chain in the channel (substep 150). This threshold may depend on the polymer used and its molecular weight. The labeled polynucleotide chain physically interacts with the polymer at the channel surface and is captured. Once captured, the free end (also called the arm) of the labeled polynucleotide chain extends in the opposite direction to the electric field. During capture, the labeled polynucleotide chain can be imaged by a detector (step 152).As an example, fluorescently labeled DNA strands can be imaged using a fluorescence microscope, a scientific complementary metal-oxide-semiconductor (scientific-CMOS) camera, a charge-coupled device (CCD), or an electron-multiplier charge-coupled device (EMCCD) camera. Other methods (other than fluorescence microscopy) that can be used to image captured polynucleotide strands include light scattering microscopy, total internal reflection microscopy (TIRF), super-resolution structural illumination microscopy (SR-SIM), stimulated emission suppression microscopy (STEP), stochastic photoreconstruction microscopy (STORM), or single-molecule localization microscopy (SMLM).

[0048] In at least one embodiment, labeled apex-pinning DNA can be captured and unfolded in a shallow microfluidic channel under an applied electric field to achieve high-quality single-molecule imaging. Step 154 ​​includes programming a controller to collect, record, and store the signal obtained from the detector. The signal is then processed (step 156). Within the computer, the image of the DNA can be interpreted and the sequence of labels can be analyzed. For example, the distance between labels forms a detectable and analyzable pattern. Of the captured DNA, regions that do not overlap with other DNA regions are easier to interpret than regions where parts of the DNA overlap with other DNA regions. Overlapping regions of "hairpin" shaped DNA can be analyzed in silico by computer, and the DNA pattern can be computationally "unfolded" to determine the pattern along the unfolded DNA. In other words, the pattern of labels associated with unfolded DNA that has not been observed can be inferred from the imaged labels and can be reconstructed computationally.

[0049] As mentioned above, applications may include imaging of fluorescently labeled markers on captured DNA, such as DNA mapping and detection of specific sequences. Such applications require detection algorithms and methods designed to detect these codes. Given the vertex capture and folded nature of DNA described here, there are at least two types of shapes for DNA captured in this way. Many (perhaps most) DNAs are entangled at some point along their length, resulting in a region where two DNA arms substantially overlap. In this overlapping region, a subsequence in the 3' to 5' direction may significantly overlap with a subsequence in the 5' to 3' direction. The two observable subsequences in this first overlapping region converge near the vertex. The two separate sequences in this overlapping region can be read by reading the overlapping region and then computationally unfolding the subsequent sequences. This involves (computer programmatically) analyzing the 3' to 5' overlap near the 5' to 3' sequence. Once the overlapping sequences are determined, the two overlapping regions can be interpreted in the computational domain.

[0050] In addition to this overlapping region, most DNA has a substantial second region located away from the vertex, where the longer DNA arm does not overlap with the shorter arm. This region contains a subsequence of either 3' to 5' or 5' to 3'. This sequence can be read and interpreted correctly.

[0051] Methods for interpreting overlapping DNA arm regions include first identifying non-overlapping regions and then using this information to interpret overlapping regions. Another method for interpreting overlapping DNA arm regions involves maintaining a database (on a computer) of codes associated with overlapping or partially overlapping regions and comparing the overlapping regions to these codes.

[0052] A system for optically imaging labeled vertex-captured polynucleotide chains for the purpose of determining the sequence of the polynucleotide chain is also provided herein. Figure 8 shows a system 158 according to one embodiment. The system 158 may include a device 160, at least one detector 162, a voltage source 164, and a controller 166. The device 160 may include one or more reservoirs. Any of the one or more reservoirs may be large enough to accommodate a sample containing a polynucleotide chain. Any of the one or more reservoirs may be large enough to accommodate at least one electrode. The electrode (either alone in the reservoir or in the reservoir with the sample containing the polynucleotide chain) may be, for example, a metal, a metal alloy, or a semiconductor material. In at least some embodiments, the electrode may be a platinum electrode. The electrode may be carbon. For example, the electrode may be monolayer or multilayer graphene. The electrode may be graphite. The electrode may also be inkjet-printed carbon-deposited lead. Alternatively, the electrode may be gold, silver, titanium, palladium, copper, stainless steel, titanium nitride, or silver / silver chloride. The electrodes can also be fabricated from doped silicon. The device may include a chip. The chip may be a microfluidic chip. The chip may be a silicon substrate chip. The chip may have two isolated (independent) single-input, single-output channels. The channels may be fabricated so that they can be visualized simultaneously within the same field of view near the geometric center of the silicon chip. The microfluidic channels may be dry-etched into the silicon wafer to a depth of, for example, 1 μm. The microfluidic channels may be in fluid contact with a reservoir. The device may include a channel cover. The channel cover may be made of glass. The channel cover may be, for example, borosilicate glass.

[0053] The system may also include at least one detector, which may include an optical detector configured to visualize the polynucleotide chains trapped within the channel. For example, the optical detector could be an epifluorescence microscope system. The epifluorescence microscope system may include a water immersion lens, an illumination source, and at least one camera.

[0054] The system may also include a voltage source for applying an electric field to the polynucleotide chain within the channel. The voltage source can apply a voltage using electrodes in the reservoir. The voltage source can also measure current.

[0055] The system may also include a controller. The controller may include a voltage source component. The controller may be programmed to interact with the device and / or with pump or flow component devices, either external to or internal to the device. The controller may also be programmed to interact with at least one detector and, if included, a separate voltage source. For example, the controller may guide a flow of polymer into the device and facilitate the delivery of the polymer to the channel surface so that the polymer can physically interact with the channel surface. Similarly, the controller may guide a flow of a sample containing labeled polynucleotide chains, with or without polymer, into the device and facilitate the delivery of the sample to the channel surface. The controller may further be programmed to interact with at least one detector. For example, the controller may be programmed to receive, record, and / or store signals obtained by at least one detector. The controller may also be programmed to process signals obtained from at least one detector or other sensors integrated into the system. The other sensors may be inside or outside the fluid device. [Examples]

[0056] The following embodiments illustrate various embodiments of the present disclosure. Those skilled in the art will recognize many modifications within the spirit and scope of the present disclosure.

[0057] Example 1: Capture of long single-molecule DNA strands Figures 9A-9C illustrate a DNA capture method according to one embodiment. In this example, a 48 kbp unimolar DNA molecule is shown apex-pinned in a microfluidic channel filled with a linear polymer buffer, with an applied axial electric field. Figure 9A is a perspective view of the apex-pinned DNA with two arms extending opposite the applied electric field. Figure 9B shows experimental epifluorescence images of the apex-pinned unimolar DNA. The images show extension at 350 V / cm, 20 V / cm, and without an applied electric field, where the DNA is relaxed in a Brownian coil shape around the apex. Figure 9C shows three consecutive images of unimolar DNA pinning with an applied 300 V / cm. The fourth and fifth images are two additional experimental examples of the capture process. The solution is 4 pM of YOYO-1 labeled λ-DNA in 1×TBE buffer containing 2% w / w 1300 kDaPVP and 2% v / v β-mercaptoethanol. The channel depth is 0.9 μm and the nominal span width is 30 μm (see Figure 14 for details). One side of the channel has a castellation pattern that is not necessary for capture (see Figures 15 and 16 for details). Figure 9A is a schematic diagram of each DNA pinned at its vertex to the wall, with two relaxed arms extending in the opposite direction to the applied electric field, corresponding to the electrostatic force of the polymer and counteracting electroosmotic flow (EOF). Figure 9B is a series of false-color epifluorescence images of a single 48.5 kbp DNA molecule at electric fields of 350, 20, and 0 V / cm. First, the DNA is captured at the high electric field. As the electric field is reduced, the DNA capture persists, and the overlap of the DNA arms decreases due to Brownian motion. The electric field is released at t=3.5 sec, and the DNA molecule coils into a three-dimensional (3D) cloud centered on the aforementioned vertex pinpoint (t=7 sec). The leftmost part of Figure 9C is a bright-field image of the channel shape. This figure also shows three consecutive images capturing the accumulation of vertex-pinned single-molecule DNA. By repeating this process, a large number of DNA molecules within the same field of view can be visualized at high throughput. The last two images in Figure 9C show two subsequent experimental examples of the capture process. Similar data for 20kbp DNA is also available in SM (see Figure 17).

[0058] Experiments were conducted with various buffer chemical compositions, including electric fields ranging from 19 to 455 V / cm and the presence or absence of linear polymer additives commonly used to suppress electroosmotic flow. DNA capture by apex pinning was observed only in sufficiently high electric fields and in the presence of polyvinylpyrrolidone (PVP) with a molecular weight of 360 kDa or greater. In one or more embodiments, the PVP polymer adsorbs at least partially onto the glass and / or oxide channel walls, thereby increasing the localized bulk viscosity of the solution within the electrical bilayer. Individual DNA molecules become entangled with one or more adsorbed PVP molecules, and this entanglement almost always occurs at a single point along the DNA, resulting in a configuration as shown in Figure 9. Pinning occurs on the silicon oxide wall of the channel and on the surface of the glass wall that seals the channel (see Figure 15). Figures 15A and 15B show DNA apex pinning at the bottom and top surfaces of a 3 μm deep linear channel under an axial electric field strength of 150 V / cm. Figure 15A shows apex-pinned DNA at the bottom surface, which is thermal oxide grown by thermal oxidation. Figure 15B shows apex-pinned DNA on the upper surface, which is anodized borosilicate glass. Figures 15A and 15B show two sequential images obtained from one implementation. The depth of focus is estimated to be 0.7 μm.

[0059] Figures 16A–16D show single-molecule DNA molecules apex-pinned by applying an axial electric field in a commercially available glass microfluidic channel 37 μm deep and filled with a linear polymer solution. Figure 16A is a schematic diagram of a commercially available glass chip purchased from Microfluidic ChipShop. Each chip contains four parallel glass channels, each 58.5 mm long, sealed with a 210 μm lid. All materials are glass. Figures 16B and 16C show sequential epifluorescence raw images of DNA electrophoresed through a commercially available glass channel 37 μm deep. The depth of field is estimated to be approximately 1 μm, and most of the DNA is out of focus. Apex-pinned DNA is observed in the background near the wall. Figure 16D shows the overall temporal median of the image sequence. The image magnifies the apex-pinned DNA at the wall.

[0060] Figure 17 shows vertex-pinned monomolecules of 20 kbp DNA in a 0.9 μm deep microfluidic channel filled with linear polymer buffer, to which an axial electric field has been applied. These are wide-field continuous raw images of monomolecular DNA pinning at an applied 254 V / cm. The solution is 4 pM of YOYO-1 labeled 20 kbp DNA in 1 × TBE buffer containing 2% w / w 1300 kDaPVP and 2% v / v β-mercaptoethanol. Raw images at t=0, 1.4, 11.5, and 28.4 seconds are shown. Faint lines in the background are due to streaking of 20 kbp DNA by high-speed electrophoresis.

[0061] Example 2: Experiment on the effect of electric fields on DNA trapping (experimental quantification of vertex-pinned single-molecule DNA quantity as a function of electric field intensity) The effect of the magnitude of the applied electric field E on the initiation and rate of DNA capture was investigated. A custom image processing algorithm was developed to quantify the number of pinned DNAs as a function of E. Figure 10A shows an overview of this analysis and additional details. Figure 10A shows an overview of the image processing flowchart used to quantify the amount of DNA capture. Subsequently, the median values ​​of a small number of images were adaptively thresholded. Using the binarized images, an alpha-shaped boundary mask was created, and the raw image data was integrated within the region enhanced by the binarized mask. Figure 10B shows the area-averaged time median intensity obtained from a 30-second experiment at each applied E.<I_med> The data is shown below. All data are for 4 pM YOYO-1 labeled λ-DNA (48.5 kbp) in 1× TBE buffer containing 2% v / v β-mercaptoethanol. The dark gray curves with circles and the light gray curves with squares represent experiments using solutions prepared with PVP molecular weights of 1300 kDa and 360 kDa (both at a concentration of 2% w / w), respectively. The dashed vertical lines highlight the approximate threshold field observed when DNA capture begins. Briefly, moving median images were calculated from small image sets to highlight the signal from stationary DNA (3 image sequences for the largest E, and 21 image sequences for the smallest E). Adaptive local thresholding was then performed to obtain binarized images. The binarized images were used as input to an alpha-shape algorithm, where they were expanded and contracted to obtain an alpha-shape boundary mask. This mask was applied to the aforementioned moving median image sequences and integrated to obtain scalar measurements (vs. time) of stationary DNA for each E. In Figure 18, this scalar value correlates well with examples of manual counting of captured DNA (obtained by manually analyzing each frame). Figures 18A–18D show a comparison between area-averaged alpha-shape boundary intensity and the results of manual molecule counting. The curves show the relationship between alpha-shape boundary intensity and time at electric field strengths of 18, 99, 164, and 235 V / cm, respectively. The scatter points show the manual count of each entangled DNA at the corresponding time step for electric field strengths of 18, 99, 164, and 235 V / cm, respectively.All data are for 4 pM YOYO-1 labeled λ-DNA (48.5 kbp) in 1× TBE buffer with 2% v / v β-mercaptoethanol and 1300 kDa PVP polymer. The alpha-shape boundary intensity against time shows a similar trend to the results of manual counting. For each E, the median of the scalar time series data from automated analysis over 30 seconds was calculated. Figure 10B shows the median of these scalar measurements as a function of E for 4 pM YOYO-1 labeled λ-DNA in 1× TBE buffer with 2% β-mercaptoethanol. Data for 2% w / w PVP with molecular weights of 1300 (circular curve) or 360 kDa (square curve) are shown. The respective E field data (360 kDa and 1300 kDa) indicate the approximate threshold E value required to initiate capture (see also Figures 19 and 20). The observed thresholds were 70 V / cm for 1300 kDa PVP and 180 V / cm for 360 kDa PVP. Experiments were also conducted using buffer solutions containing 2% w / w of PVP with molecular weights of 10 kDa and 58 kDa. No DNA capture was observed in solutions prepared using commercially available low-weight PVP. Referring again to the aforementioned PVP entanglement hypothesis, these observations suggest that sufficiently long PVP is required to adsorb to the channel surface and form a PVP scaffold that enables DNA entanglement with the PVP.

[0062] Figure 19 shows the experimental quantification of the amount of vertex-pinned single-molecule DNA as a function of electric field strength. The median intensity of each result after applying an electric field for 30 seconds is plotted on the vertical axis, and the axial electric field strength of each result is plotted on the vertical axis. Unlike Figure 10B, the intensity here is normalized by the applied axial electric field strength. All data are from 4 pM YOYO-1 labeled λ-DNA (48.5 kbp) in 1× TBE buffer with 2% v / v β-mercaptoethanol. The curves marked with circles and squares represent experiments using samples with PVP of molecular weights of 1300 kDa and 360 kDa added at a concentration of 2% w / w, respectively.

[0063] Figures 20A–20D experimentally quantify the amount of vertex-pinned single-molecule DNA as a function of time and electric field strength. The relationship between alpha-shape boundary mask intensity and time is shown for experiments performed in multiple electric fields. All data were obtained from 4 pM YOYO-1 labeled λ-DNA (48.5 kbp) in 1×TBE buffer with 2% v / v β-mercaptoethanol. Figures 20A and 20B show the buffer with 2% w / w 1300 kDaPVP polymer added, while Figures 20C and 20D show the buffer with 2% w / w 360 kDaPVP polymer added. The intensities in Figures 20B and 20D are normalized by the axial electric field strength applied to each experiment.

[0064] This data suggests that a high electric field is necessary to generate high frictional force between DNA and PVP polymers. Considering the threshold characteristics of the required electric field, a sufficiently high electric force may be needed on the DNA to eliminate the hydration layer between the DNA and PVP molecules. The absence of the hydration layer can increase solid friction between the charged DNA polymer and the uncharged linear PVP polymer. This may lead to the persistent high-friction entanglement observed at fixed vertices. This hypothesis is consistent with the fact that persistent pinning occurs at vertices near the ends of the DNA molecule, resulting in significantly different forces due to the large difference in the lengths of the two relaxed arms. Studies of DNA interacting with agarose crosslinked polymers have suggested the removal of the hydration layer between single DNA molecules and polymer molecules. DNA electrophoresis studies via 3D agarose gel networks have shown that DNA is trapped under high electric fields. Another study observed DNA trapping within agarose gels under electric fields above a certain threshold, leading to the "knot" trapping hypothesis. However, in these previous studies, the tangled DNA formed complex three-dimensional shapes within the gel, not the "clean" vertices with two linear arms pinned to the wall (in a linear polymer solution) as in this study.

[0065] Example 3: Testing the relaxation dynamics of apex-pinned DNA The relaxation dynamics of vertex-pinned DNA after electric field removal were analyzed. Figure 11A shows time-series images. The DNA was observed to be pinned at random positions along the length of the DNA molecule, resulting in various relaxation timescales. An automated image processing method based on adaptive thresholding, elliptic fitting of DNA images, and extraction of the major and minor axes of the best-fitting ellipse was employed. Figure 11B is a magnified image of the best-fitting multiple ellipses. Specifically, Figure 11A shows an example sequence of raw epifluorescence microscope images of pinned λ-DNA (48.5 kbp) immediately after removal of the applied electric field (t=0 s). Figure 11B shows an example of a raw image with the best-fitting ellipses superimposed, used to quantify the length and width of the DNA. Figures 11C and 11D plot the statistics of the major axis length L of the ellipse against the relaxation of 48.5 kbp and 20 kbp DNA molecules against time from electric field removal. The black curve shows the instantaneous mean of L. The dark gray and light gray diagonal lines represent one and two standard deviations of the distribution, respectively.

[0066] Figures 11C and 11D show characteristic length statistical moments when pinned DNA molecules relax. Figures 11C and 11D show data for 1300 kDa λ-(48.5 kbp) or 20 kbp DNA labeled with YOYO-1 dye in 1× TBE buffer containing 2% v / v β-mercaptoethanol and 2% w / w PVP, respectively. Since most DNA was pinned to the vertices quite far from the ends of the molecule, the initial mean value of L is lower than the expected full contour length. The vertex-pinned 48.5 kbp DNA molecule took approximately 8 seconds to fully relax, and the vertex-pinned 20 kbp DNA took approximately 3 seconds to fully relax. Figure 21 shows the epifluorescence image of relaxation of vertex-pinned 20 kbp DNA. Figure 21 shows the relaxation dynamics of vertex-pinned single-molecule DNA. This is an example sequence of raw epifluorescence microscope images immediately after electric field removal (t=0 seconds) of pinned 20kbp DNA.

[0067] Figures 22A–22J show an overview of the automated image processing used to quantify 48.5 kbp DNA relaxation. Figure 22A is an example image of vertex-pinned DNA before relaxation. Figure 22B is the binarized image of Figure 22A after adaptive thresholding and alpha-shape calculation. Figure 22C is the image obtained by integrating Figure 22A with the mask of Figure 22B. Figure 22D is a magnified view of Figure 22A. Figure 22E is an example image showing the automated detection of each island. The elliptic curve shows the result of elliptic approximation. The black and light gray lines within the ellipse indicate the principal and secondary axes of the approximated elliptic shape, respectively. Figures 22F–22J are images showing the same procedure as Figures 22A–22E, except that images were selected during the DNA relaxation process.

[0068] Figures 23A–23J show an overview of the automated image processing used to quantify 20kbp DNA relaxation. Figure 23A is an example image of vertex-pinned DNA before relaxation. Figure 23B shows the binarized image of Figure 23A after adaptive thresholding and alpha-shape calculation. Figure 23C is the image obtained by integrating Figure 23A with the mask of Figure 23B. Figure 23D is a magnified view of Figure 23A. Figure 23E is an example image showing the automated detection of each island. The elliptic curve shows the result of elliptic approximation. The black line and light gray line show the principal and secondary axes of the approximated elliptic shape, respectively. Figures 23F–23J are images showing the same procedure as Figures 23A–23E, except that images were selected during the DNA relaxation process.

[0069] Figures 24A–24C show the colocalization analysis of image intensity of vertex-pinned DNA in three different examples. Colocalization data is shown for three independent experimental examples of DNA capture. Experimental examples 1, 2, and 3 are the DNA vertex-pinned DNA data from Figures 13A, 13B, and 13C in the text, respectively. Figures 24A–24C show colocalization scatter plots (correlation plots) comparing pairs of each experimental example. As shown by the intensity bars on the right, the heatmap is a count based on the raw intensity values ​​of colocalized pixels obtained from two images of two experimental examples. The Pearson correlation coefficients were calculated between each pair, and their values ​​are 0.30, 0.25, and 0.22, respectively.

[0070] Figures 25A-C show the colocalization analysis of image intensity of vertex-pinned DNA across three consecutive images obtained from one experimental example. Figure 25A shows three consecutive epifluorescence images from one experimental example of a DNA vertex-pinning experiment. The first and second subpanels are separated by 5 seconds. The second and third subpanels are separated by 2 seconds. Figures 25B and 25C are confocal scatter plots (correlation plots) comparing each pair of consecutive images. The Pearson correlation coefficients of the two time-series images are 0.81 and 0.86, respectively.

[0071] As described above, upon removal of the (high) electric field, the DNA consistently relaxes around the pinned point, towards the apex. This further supports the hypothesis of a single-point entanglement between DNA and PVP. The persistence of such a pinning point (for 30 seconds) suggests that the observed DNA capture is not due to electroadsorption forces such as dielectrophoretic forces arising from wall defects (such as roughness elements). Such electroadsorbed DNA molecules are expected to be immediately "released" upon removal of the electric field. In this example, the DNA is not captured at the convex corner of the wall structure within the channel.

[0072] Example 4: Cycle test of DNA capture, relaxation, and separation Figure 12 shows a typical cycle of DNA vertex pinning under a high electric field, relaxation due to electric field termination, separation after approximately 36 seconds, and subsequent free diffusion and electrophoresis. The DNA molecule was initially in a three-dimensional Brown coil at t=0 seconds. At t=2.4 seconds, an electric field of 190 V / cm was applied, initiating capture. The electric field was terminated at t=33.8 seconds, and the DNA molecule rapidly relaxed towards the vertex / pinpoint (see Figure 11). After approximately 50 seconds, the relaxed DNA was released from its fixation to the wall and began to diffuse and electrophores freely again. At t=88.6 seconds, a low axial electric field of 19 V / cm was applied to clear the field of view (FOV) of the previously pinned DNA. This process can be repeated multiple times with a period of approximately 60 seconds. Figure 12 shows a sequence of images of a single DNA molecule (from top to bottom). The DNA molecule initially has the expected random coil shape. To initiate vertex pinning, an electric field of E=190V / cm was applied at t=2.4s. E was stopped at t=33.8s, and the DNA molecules were either rapidly released or slowly relaxed towards the vertex. After approximately 50 seconds, more than 95% of the DNA was released from the wall. At t=88.6s, a relatively low axial electric field of 11V / cm was applied, and the DNA molecules that had been vertex-pinned underwent electrophoresis. Note the displacement of the correlation pattern across the last three images.

[0073] The data in Figure 12 further supports the entanglement between DNA and PVP. Removing the electric field rehydrates the space between DNA and PVP, and the resulting low-friction configuration and Brownian motion facilitate separation after several tens of seconds. Pulses of pressure-driven flow can induce even faster separation, down to the order of seconds.

[0074] We also investigated whether the DNA pinning sites were spatially correlated between experiments. Figures 13A–13G show single DNA molecules apex-pinned to the wall at randomly varying positions for each experiment. Figures 13A–13C show three representative experimental examples. The first is shown in inverted grayscale, while the other two are raw grayscale images. Between experiments, the DNA was fully relaxed and separated. Figure 13D is a superposition of images from experimental examples 1 and 2, and Figure 13E is a superposition of images from experimental examples 1 and 3. Enlarged images of Figures 13D and 13E are shown in Figures 13F and 13G. All data shown in Figures 13A–13G are from YOYO-1 labeled λ-DNA from 4 pM, 2% w / w 1300 kDaPVP. Figures 13A–13C show images of three separate experimental examples of DNA capture using inverted grayscale (experiment 1), grayscale (experiment 2), and grayscale (experiment 3) schemes, respectively. The DNA was captured at 300 V / cm for 15 seconds. The DNA was relaxed and separated between experiments (see Figure 4), and separation was confirmed using a low electric field to clear the previously captured DNA. Figures 13C and 13D are superimposed images of Experiments 1, 2, and 3, showing low spatial correlation of vertex capture points across two and three experimental examples, respectively. Figures 13F and 13G are enlarged images of Figures 13D and 13E, respectively.

[0075] The DNA capture sites were observed to be uncorrelated in all the experiments examined. This result further supports the hypothesis that the observed capture is not due to specific channel shape features such as roughness elements or nanoscale surface pits. This data supports the idea that DNA molecules become entangled with adsorbed PVP molecules. This entanglement is ultimately reversible, with new DNA molecules being fixed to new, uncorrelated locations. The nearly linear increase in capture rates shown in Figure 10 suggests that the DNA capture sites did not reach saturation during the experimental period.

[0076] Materials and methods for Examples 1-4: Custom microfluidic interface devices Figures 14A–14D show a custom microfluidic interface device according to one embodiment. Figure 14A shows pieces of a PDMS reservoir, 170 μm thick borosilicate glass, and a silicon substrate chip. Figure 14B shows the assembled custom microfluidic device. The borosilicate glass was anodized to the silicon substrate chip, and the PDMS reservoir was plasma-bonded to the borosilicate glass. Figure 14C shows the assembled custom microfluidic device taped to a glass slide and visualized with a 1.2 numerical aperture 60x immersion objective lens. The illumination source is a blue LED. A platinum electrode was inserted into the reservoir filled with liquid. Figure 14D shows the overall layout of the silicon substrate chip, with magnified images of two regions. The central region has a wall structure near one side. The large microchannel region connected to the reservoir has a 10 μm diameter column array. The column array is designed to support the anodized borosilicate glass. More specifically, Figure 14A shows three main components of a microfluidic device in one or more embodiments: four polydimethylsiloxane (PDMS) reservoirs, a borosilicate glass slip (channel cover), and a silicon substrate chip. Figure 14B is a perspective view of the device after bonding and assembly. Figure 14C is an experimental image showing the chip, platinum electrodes in the reservoir (connected to alligator clips), and a water immersion objective lens. Figure 14D shows the channel layout within the silicon substrate chip. This chip has two isolated (independent) single-input, single-output channels. The channels were fabricated so that they could be visualized simultaneously within the same field of view near the geometric center of the silicon chip. Fabrication was performed on a 4-inch n-type silicon wafer (resistivity 4 Ω-cm). After standard photolithography, the microfluidic channels were dry-etched into the silicon wafer to a depth of 1 μm using HBr and BCl3 (Oxford Instruments PlasmaLab III-V etcher) (the actual depth measured with a profile meter was 0.9 μm). The residual photoresist was removed with oxygen plasma, followed by a piranha wash at 120°C for 20 minutes.Subsequently, a 300nm thick thermal oxide film was grown on the wafer using Rogue Valley Microdevices. The wafer was diced with a wafer saw to produce nine devices with in-plane dimensions of 22×24mm. These devices were subjected to piranha cleaning again. The glass coverslip was made of No. 1 thickness 3.3 borosilicate glass (purchased from Kemtech America Inc.). Holes were drilled in the glass using a 1.1mm diameter triple-ripple diamond drill bit (purchased from Arrowhead Lapidary & Supple) and a manual drill press. The drilled coverslip was anodized. After anodizing, a PDMS reservoir with an outer diameter of 5.0mm and an inner diameter of 1.5mm was cut with a biopsy punch and bonded to the glass with air plasma. Fluid connections were formed by aligning the holes drilled in the glass and the holes drilled in the PDMS.

[0077] Sample preparation λ-DNA samples were purchased from ThermoScientific (Waltham, Massachusetts) at a stock concentration of 0.3 μg / uL. 20 kbp DNA samples were purchased from ThermoScientific at a stock concentration of 0.5 μg / uL. YOYO-1 dye was purchased from Invitrogen at a stock concentration of 1 mM. 10×TBE buffer was purchased from Invitrogen and consists of 1.0 M Tris, 0.9 M boric acid, and 0.01 M EDTA. Polyvinylpyrrolidone (PVP) with molecular weights (MW) of 10, 58, 360, and 1300 kDa was purchased from ThermoScientific. α-Mercaptoethanol (MW=78.13, purity 98.0%) was purchased from TCI America. The 10×TBE buffer was first diluted to 1×TBE buffer with molecular biology grade water (Fisher Bioreagents, purchased from Pittsburgh, Pennsylvania). Next, β-mercaptoethanol was added to 1×TBE buffer to a concentration of 2% v / v as an oxygen scavenger. Then, PVP was added to the buffer at a concentration of 2% w / w in each of the aforementioned MWs. The final buffer consisted of 1×TBE, 2% v / v β-mercaptoethanol, and 2% w / w PVP, with MWs of 10, 58, 360, or 1300 kDa, respectively. The conductivity of the loading buffer was measured at 1487 μS / cm using a conductivity probe (Advanced Electrochemistry Meter, ThermoScientific Inc.). All steps up to this point were performed at room temperature (20±1℃). Next, the raw DNA samples were diluted in the loading buffer to a final concentration of 4 pM, and YOYO-1 dye was added to achieve a stochastic ratio of 5:1 between base pairs of dyes. The samples were then incubated at 40℃ for 1 hour before use. Subsequently, each sample was incubated at 40℃ for 1 hour before use to promote dye intercalation.

[0078] Measuring device Figure 14C shows an example image from one experiment using an assembled microfluidic device. Visualization was performed using a standard OLYMPUS BX60 upright epifluorescence microscope. A 1.2 numerical aperture 60x immersion objective lens was used (OLYMPUS UPlanApo). The illumination source was a ThorLabs high-power blue LED (SOLIS-470C) controlled by a ThorLabs DC200 controller, and the excitation / dichroism / emission components of the epifluorescence filter cube were EX460-490, DM505, and EM510IF (OLMPUS U-MWIB2). Two cameras were used for image acquisition. The first camera was a Hamamatsu Photonics scientific complementary metal-oxide-semiconductor (sCMOS) camera (ORCA-Flash 4.0LT), controlled using HCImageLive software. The sCMOS camera was used to acquire all the data in this book and most of the data in the supplementary materials (SM). The second camera was an Andor electron-multiplier charge-coupled device (EMCCD) camera (iXon Ultra 897), controlled using Andor's SOLIS software. The EMCCD camera was used to acquire image data in Figure S3 (image of the wet-etched entire glass channel, described later).

[0079] The voltage was applied using platinum electrodes inserted into the reservoir at the channel end. The electrodes were platinum wires (0.368 mm diameter, hard, 99.95 metal base) purchased from ThermoScientific. Voltage application and current measurement were performed using a Keithley 2400 SourceMeter. This SourceMeter was controlled and triggered from a personal computer (PC) via an RS232 interface using a custom script written in MATLAB (MATLAB 2023b, Mathworks Inc., Natick, Massachusetts, USA).

[0080] Image processing for quantifying vertex-pinned DNA quantity All data in the following image processing section were acquired using a Hamamatsu Photonics Orca-Flash4.0LT sCMOS camera at 20 fps with an exposure time of 0.49 seconds. The camera's sensor pixel size is 6.5 × 6.5 μm. Using a 60x objective lens without a reduction lens, the pixel size on the image plane was approximately 0.11 × 0.11 μm per pixel. The raw TIFF sequences from each experiment were imported into MATLAB for image analysis. Each 30-second sequence consists of 600 images. First, a sequence consisting of a time-shifted median calculation is obtained. This shifted median replaces the pixel value of each image with the median of the same pixel across a small number of preceding and succeeding images. This operation has the effect of highlighting and retaining nearly stationary image data (pinned and stretched DNA) and rejecting moving objects (electrophoretic DNA). A maximum applied voltage of 363 V / cm correlated with the highest speed. Therefore, each median calculation considered only three image groups (i.e., each image pixel value was replaced with the median of the previous frame, the current frame, and the next frame). Median calculations for the lowest applied electric field (slowest moving DNA) gradually used more images within each group. Up to 21 frames were considered at the lowest applied electric field of 19 V / cm. The upper subplot in Figure 10A shows an example of frames obtained from the moving median image sequence. For example, footage taken for comparison between raw footage and moving median footage shows how image data from stationary objects (pinning DNA) is enhanced. Figure 26 is an example of a frame from this footage, showing the difference between raw footage and moving median footage, processed to show only non-moving DNA molecules. Local mean-based adaptive thresholding was performed on the moving median image sequence using the Bradley method. For this purpose, the MATLAB command adaptthresh was used with a sensitivity parameter of 0.45, followed by imbinarize to obtain binarized images of each frame. The pixel values ​​of the binarized image were set to zero if they were below a threshold and to 1 if they were above a threshold. Next, the data for the pixel coordinates for which the threshold was set was identified and saved using MATLAB commands.Subsequently, an alpha shape operation was performed on the binarized image to efficiently remove non-DNA image noise from the analysis. The alpha shape is a generalization of the convex hull concept and is a subset of Delaunay triangles. The alpha shape method is used in computational graphics pattern recognition to identify the (proximity-based) relationships between points on a graph. To obtain the alpha shape, the binarized pixel coordinates and an alpha radius of 2.5 were used as input to the MATLAB command alphaShape. The obtained alpha shape was projected onto a binary mask. The binary mask was first expanded and then condensed using stream lines of the same 10-pixel length as structuring elements (again, because the DNA image was pinned almost horizontally). After condensation, the algorithm obtained an alpha shape boundary mask. The alpha shape method successfully reduced background noise caused by adaptive thresholding. The alpha shape operation, expansion, and condensation also smoothed the noisy edges of the identified object. The central sub-image in Figure 10A shows an example of the obtained mask. At each time frame, the binarized mask was used on the moving median image to obtain a sequence of thresholded moving median images. The lower sub-image in Figure 10A shows an example of the calculation result between the alpha-shape join mask and one of the median images. Next, the (original, unaltered) intensities of the DNA detected within the obtained threshold image were spatially integrated to obtain a scalar signal that changes over time at a frame rate of 20 fps (scalar I in Figure 11).

[0081] Image processing for relaxation analysis of vertex-pinned DNA This section describes the image processing analysis performed to obtain the DNA relaxation data shown in Figures 11B, 11C, and 11D of this book. First, the TIFF sequence was temporally Gaussian filtered with a 3-frame wide filter using the MATLAB built-in function imgaussfilt3. Next, an alpha-shape boundary mask for the DNA relaxation image was obtained in the same way as described above, except that it did not include expansion and contraction to preserve the shape of the DNA. Once the boundary mask was obtained, the MATLAB command bwconncomp followed by regionprops was used to characterize the distinct connection regions within the alpha-shape boundary mask. To ensure that image noise and DNA fragments do not remain in the subsequent analysis, connection regions with total intensities less than 100,000 and 20,000 were removed for the 48.5kbp and 20kbp DNA samples, respectively. These thresholds were determined empirically and are designed to remove objects with significantly lower intensities than intact DNA fragments. Figures 21A–21C and 21F–21H show examples of the aforementioned image processing procedure for vertex-pinning DNA before and during relaxation, respectively. Next, ellipses were fitted to each of the remaining regions using the MATLAB region props command again. These ellipses have the same normalized second-order central moment as the fitted regions. Examples of ellipses fitted to various DNA fragments are shown in Figure 11B. For each frame after the electric field was turned off in each DNA relaxation experiment, a collection of fitted ellipses was obtained that ideally represented the shapes of various DNA fragments at a given time in that experiment. Figures 21D–21E and 21I–21J show examples of the aforementioned island identification and principal axis estimation. Figures 21D and 21E show examples of vertex-pinning DNA before relaxation, and Figures 21I and 21J show examples of vertex-pinning DNA during relaxation. Elliptic approximations are shown as a superposition of gray ellipses, and the major axes of the ellipses are shown as two mutually orthogonal lines. From this collection of fitted ellipses, the following two values ​​were recorded. This refers to the average length of the major axis of the fitted ellipse used as the characteristic length of relaxed DNA, and the standard deviation of the major axis lengths of fitted ellipses whose major axis length is longer or shorter than the average length. Figures 11C and 11D show the analysis results for 1 and 2 standard deviations.The black lines indicate feature lengths, while the dark and light gray shaded areas represent data that are one or two standard deviations from the mean of the long axis length. The only difference between the relaxation analyses of 48.5kbp and 20kbp DNA is the threshold chosen to remove selected regions that appear to be DNA fragments or image noise.

[0082] Multiple MW PVP and DNA contour line length The outline lengths of unstained λ-DNA (48.5 kbp) and 20 kbp DNA have been reported to be approximately 16 μm and 6.6 μm, respectively. When λ-DNA and 20 kbp DNA are stained with YOYO-1 dye, their outline lengths become approximately 21 μm and 8.7 μm, respectively. The outline lengths of PVP with MWs of 10, 58, 360, and 1300 kDa are approximately 25, 145, 900, and 3250 nm, respectively.

[0083] spatial average intensity The symbol <···> is used to indicate the spatial average of all pixels within a two-dimensional region. For example, it is used to show the spatial integration or two-dimensional correlation of the intensity values ​​of raw images. For a quantity q in each time frame, the spatial average is given by equation (1): TIFF2026064217000002.tif17170 Here, Nx and Ny represent the number of pixels in the horizontal and vertical directions, respectively.

[0084] Estimation of the magnitude of the axial electric field The conductivity of the aforementioned buffer solution was measured using a commercially available conductivity meter. The current during the experiment was measured using a Keithley SourceMeter. The channel depth was measured using a profilometer and found to be approximately 0.9 μm. The channel width was identified using a photolithography mask, and measured from the bright-field image, as shown in the leftmost subplot of Figure 9C, it was found to be approximately 20 μm. The axial electric field intensity can be related to the current by Ohm's law, as shown in equation (2): j=σE (2) Here, j is the current density and σ is the conductivity of the buffer. Dividing the measured current I by the estimated cross-sectional area A gives the current density j. Then, the electric field strength was estimated using E = j / σ.

[0085] The U.S. patent applications (RBPA0517PUSP and RBPA0517PUSP2) are related to this application and are incorporated in their entirety by reference.

[0086] While exemplary embodiments have been described above, these embodiments are not intended to describe all possible forms of the Disclosure. Rather, the terms used herein are descriptive, not limiting, and it should be understood that various modifications may be made without departing from the spirit and scope of the Disclosure. Furthermore, features of various embodiments can be combined to constitute further embodiments of the Disclosure.

Claims

1. A method for detecting polynucleotide chains by capturing them within a channel, The steps include labeling at least one subsequence within a polynucleotide chain to obtain a labeled polynucleotide chain having at least one label, The steps include supplying the polymer to the surface of the channel, The steps include physically interacting the polymer with the surface of the channel, The steps include supplying a sample containing the labeled polynucleotide chain to the channel, The steps include applying an electric field to the labeled polynucleotide chain to promote physical interaction between the labeled polynucleotide chain and the polymer on the surface of the channel, thereby trapping the labeled polynucleotide chain on the surface of the channel, A method characterized by comprising the step of detecting at least one label within the labeled polynucleotide chain.

2. The method according to claim 1, wherein the at least one marker is optically detected.

3. The method according to claim 1, wherein the at least one of the labels is a fluorescent label.

4. The method according to claim 1, wherein the at least one label is detectable by a fluorescence microscope.

5. The method according to claim 1, wherein the at least one label is detectable by one of the following methods: light scattering microscopy, total internal reflection microscopy (TIRF), super-resolution structural illumination microscopy (SR-SIM), stimulated emission suppression microscopy (STED), stochastic optical reconstruction microscopy (STORM), or single-molecule localization microscopy (SMLM).

6. The method according to claim 1, wherein the labeled polynucleotide chain is captured at the vertex of the labeled polynucleotide chain.

7. The method according to claim 6, wherein the first free end and the second free end of the labeled polynucleotide chain each extend outward from the vertex in the direction opposite to the electric field.

8. The method according to claim 1, wherein the labeled polynucleotide chain is greater than one kilobase pair.

9. The method according to claim 1, wherein the surface of the channel is flat.

10. The method according to claim 1, wherein the polymer can alter the electroosmotic flow within the channel.

11. The method according to claim 1, wherein the polymer is a neutral and water-soluble polymer.

12. The method according to claim 11, wherein the polymer is a polyvinylpyrrolidone polymer.

13. The method according to claim 12, wherein the polyvinylpyrrolidone polymer has a molecular weight of more than 100 kDa.

14. The method according to claim 11, wherein the neutral and water-soluble polymer is hydroxyethylcellulose.

15. The method according to claim 11, wherein the neutral and water-soluble polymer is polyethylene glycol.

16. The method according to claim 11, wherein the neutral and water-soluble polymer is polyvinyl alcohol.

17. The method according to claim 1, wherein the channel has at least one dimension perpendicular to the electric field that is less than 5 microns.

18. The method according to claim 1, wherein the electric field is 10 to 1,000 V / cm.

19. The method according to claim 1, further comprising the step of supplying pulses of a pressure-driven flow to the labeled polynucleotide chain to promote the release of the labeled polynucleotide chain from physical interaction with the polymer.

20. The method according to claim 1, further comprising the step of reducing the intensity of the electric field applied to the labeled polynucleotide chain in order to facilitate the release of the labeled polynucleotide chain from physical interaction with the polymer so that the labeled polynucleotide chain moves through the channel.

21. The method according to claim 20, further comprising the steps of: supplying a second sample containing a second labeled polynucleotide chain to the surface of the channel, increasing the intensity of the electric field to promote physical interaction between the second labeled polynucleotide chain and the polymer on the surface of the channel to capture the second labeled polynucleotide chain on the surface of the channel; and detecting at least one label within the second labeled polynucleotide chain to visualize the second labeled polynucleotide chain.

22. The method according to claim 21, further comprising the steps of: reducing the intensity of an electric field applied to the second labeled polynucleotide chain so that the labeled polynucleotide chain moves through the channel, thereby promoting the release of the second labeled polynucleotide chain from physical interaction with the polymer; providing a sample containing the labeled polynucleotide chain; increasing the intensity of the electric field to capture and detect the labeled polynucleotide chain; and repeating the steps of reducing the intensity of the electric field to release the labeled polynucleotide chain so that the labeled polynucleotide chain moves through the channel, n times.

23. In a system for capturing, extending, and detecting labeled polynucleotide chains, It is a device, A reservoir of a size to accommodate a sample including electrodes and / or labeled polynucleotide chains, A fluid tip having at least one channel, wherein the at least one channel is in fluid communication with the at least one reservoir so that a sample can flow through the at least one channel, and A device equipped with a channel cover, A voltage source capable of applying an electric field to the channel, At least one detector capable of detecting a signal from the aforementioned device, It is a controller, Guide the flow of polymer into the device and facilitate the delivery of the polymer to the surface of the channel so that the polymer can physically interact with the surface of the channel. The flow of the sample from the at least one reservoir to the channel is guided, A voltage difference is generated between the electrode and at least one other electrode to form an electric field, thereby promoting the physical interaction between the labeled polynucleotide chain and the polymer, and trapping the labeled polynucleotide chain on the surface of the channel. Interacting with the at least one detector, the system collects, records, and stores the signal received from the at least one detector. A system comprising a controller programmed to process signals received from at least one of the detectors.

24. The system according to claim 23, wherein the detector is a camera.

25. The system according to claim 23, wherein the controller is further programmed to reduce the intensity of the electric field applied to the labeled polynucleotide chain to facilitate the release of the labeled polynucleotide chain from physical interaction with the polymer.

26. The system according to claim 25, wherein the controller is further programmed to move the released polynucleotide chain through the channel.

27. The system according to claim 26, wherein the controller is further programmed to induce a flow of a second sample containing a second polynucleotide chain from at least one reservoir in fluid communication with the channel, and to increase the intensity of the electric field to capture the second polynucleotide chain.

28. The system according to claim 23, wherein the controller is further programmed to supply pulses of pressure-driven flow to the device in order to increase the release rate of the polynucleotide chain.

29. The system according to claim 23, wherein the channel cover is transparent.

30. A method for detecting polynucleotide chains by capturing them within a channel, The steps include supplying the polymer to the surface of the channel, The steps include physically interacting the polymer with the surface of the channel, The steps include supplying a sample containing a polynucleotide chain to the channel, The steps include labeling at least one subsequence within the polynucleotide chain to obtain a labeled polynucleotide chain having at least one label, The steps include applying an electric field to the labeled polynucleotide chain to promote physical interaction between the labeled polynucleotide chain and the polymer on the surface of the channel, thereby trapping the labeled polynucleotide chain on the surface of the channel, A method characterized by comprising the step of detecting at least one label within the labeled polynucleotide chain.

Images