Systems and methods for capturing and stretching polynucleotide chains

Abstract

Systems and methods for capturing polynucleotide strands at the surface of a channel. The method may include providing a polymer to the channel surface and physically interacting the polymer with the channel surface. The method can also include providing a sample comprising the polynucleotide strand to the polymer that interacts with the channel surface, and applying an electric field to the polynucleotide strand to promote a physical interaction between the polynucleotide strand and the polymer to capture the polynucleotide strand at the channel surface.

Description

[0001] Cross-reference This application claims the benefit of U.S. Provisional Application Serial No. 63 / 702,095, filed October 1, 2024, the entire contents of which are incorporated herein by reference. Technical Field

[0002] This invention relates to systems and methods for controlling, capturing, and stretching polynucleotide chains. Background Technology

[0003] The control, capture, stretching, and imaging of single-molecule DNA flow and migration could enable fundamental research into DNA polymer physics and the interaction of DNA with fluids and electric fields. Current techniques require complex fabrication, lack throughput, and are not cost-effective. Furthermore, current methods typically analyze moving molecules within nanoscale channels, which degrades detection quality. Therefore, there is a need for cost-effective systems and methods that allow for high-throughput sample processing to facilitate the control, capture, and visualization of single-molecule polynucleotide chains. Summary of the Invention

[0004] In at least one aspect, a method is provided for capturing a polynucleotide chain on a channel surface. The method may include: providing a polymer to the channel surface; causing the polymer to physically interact with the channel surface; providing a sample containing a polynucleotide chain to the polymer interacting with the channel surface; and applying an electric field to the polynucleotide chain to promote physical interaction between the polynucleotide chain and the polymer, thereby capturing the polynucleotide chain on the channel surface. The physical interaction between the polynucleotide chain and the polymer may occur at a vertex of the polynucleotide chain. After the electric field is applied, a first end and a second end of the polynucleotide chain may each extend outward from the vertex. The polymer may have the ability to modulate electroosmotic flow in the channel. The polymer may be a neutral and water-soluble polymer. A neutral and water-soluble polymer may be a polyvinylpyrrolidone (PVP) polymer. A PPVP polymer may have a molecular weight greater than 100 kDa. A neutral and water-soluble polymer may be hydroxyethyl cellulose. A neutral and water-soluble polymer may be polyethylene glycol. A neutral and water-soluble polymer may be polyvinyl alcohol. The channel may have at least one dimension perpendicular to the direction of the electric field, less than 5 micrometers. The strength of the electric field can be from 10 to 1,000 V / cm. The method may further include varying the strength of the electric field applied to the polynucleotide chain to promote the release of the polynucleotide chain from its physical interaction with the polymer. Changing the electric field may include altering its strength. The method may further include providing a second sample containing the polynucleotide chain to the polymer interacting with the channel surface, and changing the electric field to promote physical interaction between the polynucleotide chain and the polymer, thereby trapping the polynucleotide chain at the channel surface. The method may further include applying a pulse of pressure-driven flow to the polynucleotide chain to promote its release from its physical interaction with the polymer.

[0005] In at least another aspect, a system for capturing polynucleotide chains is provided. The system may include a device comprising at least one reservoir, sized to accommodate an electrode, an electrolyte, and / or a sample containing the polynucleotide chain; and a fluidic chip having at least one channel in fluid communication with the at least one reservoir, allowing the sample to be transported through the at least one channel. The system may also include a voltage source capable of applying an electric field to the channel. The system may further include a controller programmed to direct the transport of a polymer to the device to facilitate polymer delivery to the channel surface, enabling the polymer to physically interact with the channel surface; to direct the transport of a sample from the at least one reservoir to the channel; and to interact with the voltage source to generate a voltage difference between the electrode and at least one other electrode to form an electric field and facilitate physical interaction between the polynucleotide chain and the polymer to capture the polynucleotide chain on the channel surface. The controller may be further programmed to reduce the electric field strength applied to the polynucleotide chain to facilitate the release of the polynucleotide chain from the physical interaction with the polymer. The controller may also be further programmed to direct the transport of the released polynucleotide chain through the channel to remove the polynucleotide chain from the channel region where it was captured. The controller can also be further programmed to guide the transfer of a second sample containing polynucleotide chains from at least one reservoir in fluid communication with the channel, and to increase the electric field strength to capture the polynucleotide chains from the second sample. The controller can also be further programmed to provide pulses of pressure-driven flow to the device to increase the release rate of the polynucleotide chains. Attached Figure Description

[0006] Figure 1A and Figure 1B A method for capturing and stretching polynucleotide chains is shown.

[0007] Figure 2A and Figure 2B A method for capturing and stretching polynucleotide chains is shown.

[0008] Figure 3A and Figure 3B A system for capturing and stretching polynucleotide chains is shown.

[0009] Figure 4 A channel with more than one fluid layer is shown.

[0010] Figure 5 An example of a channel with both low electric (E) field regions and high electric field regions is shown.

[0011] Figure 6 Examples are shown of channels with a polymer-adsorbable surface and a defined region of patterned material that is not polymer-adsorbable.

[0012] Figure 7A and Figure 7B A method is shown for optical imaging of vertex-captured polynucleotide chains for the purpose of determining their sequence.

[0013] Figure 8 A system is shown for optical imaging of vertex-captured polynucleotide chains for the purpose of determining their sequence.

[0014] Figures 9A to 9C A DNA capture method according to one embodiment is shown.

[0015] Figures 10A to 10B A summary of the image processing workflow for quantifying DNA capture amounts is shown.

[0016] Figure 11A and Figure 11B The relaxation dynamics analysis of the vertex-fixed DNA after the electric field was removed is shown. Figure 11C and Figure 11D The statistical moments of the characteristic length of a fixed DNA molecule during relaxation are shown.

[0017] Figure 12 This demonstrates a representative cycle of DNA vertex fixation under a high electric field.

[0018] Figures 13A to 13G This demonstrates that the vertex of a single DNA molecule is fixed to the wall, and its position changes randomly in different experimental runs.

[0019] Figures 14A to 14F A microfluidic interface device customized according to one implementation is shown.

[0020] Figure 15A and Figure 15B This illustrates the case where a straight channel with a depth of 3 μm is fixed at the DNA apex on the bottom and top surfaces under an axial electric field strength of 150 V / cm.

[0021] Figure 16A A schematic diagram of a commercially available glass chip purchased from Microfluidic ChipShop is shown. Figure 16B and Figure 16C This image shows a series of original fluorescence micrographs of the electromigration of DNA in a commercial glass channel at a depth of 37 μm. Figure 16D The global temporal median image of the image sequence is shown.

[0022] Figure 17 A single 20 kbp DNA molecule is shown as apex-fixed in a microfluidic channel 0.9 μm deep filled with a linear polymer buffer solution and subjected to an axial electric field.

[0023] Figures 18A to 18D The area-average α-shaped boundary strength is shown. Comparison with manual molecule counting results.

[0024] Figure 19 The experimental quantitative results show the change in the number of vertex-fixed single-molecule DNA molecules as a function of electric field strength.

[0025] Figures 20A to 20D The experimental quantitative results show the changes in the number of vertex-fixed single-molecule DNA over time and electric field strength.

[0026] Figure 21 The relaxation dynamics of vertex-fixed single-molecule DNA are shown.

[0027] Figures 22A to 22J A summary of automated image processing for quantifying 48.5 kbp DNA relaxation is presented.

[0028] Figures 23A to 23J A summary of automated image processing for quantifying 20 kbp DNA relaxation is presented.

[0029] Figures 24A to 24C The results of colocalization analysis of image intensity for vertex-fixed DNA across three different experiments are shown.

[0030] Figures 25A to 25C Colocalization analysis of image intensity for vertex-fixed DNA across three consecutive images from an experimental run is shown.

[0031] Figure 26 Example frames from the video are shown, illustrating the difference between the original video and a moving median video, which was processed to display only non-moving DNA molecules. Detailed Implementation

[0032] Detailed embodiments of this disclosure are disclosed herein as needed; however, it should be understood that these disclosed embodiments are merely exemplary implementations of this disclosure, and various different forms and alternatives may be adopted. The drawings are not necessarily drawn to scale; some features may be enlarged or reduced to show details of specific components. Therefore, the specific structural and functional details disclosed herein should not be construed as limiting, but rather as representative guidance for those skilled in the art to implement this disclosure in various ways.

[0033] Except as expressly stated in the Examples section or otherwise, all numerical values ​​representing amounts of materials or reaction conditions and / or usage conditions in this specification should be understood as modified by the word "about". The initial definition of acronyms or other abbreviations applies to all subsequent uses of the same abbreviations herein and to normal grammatical variations of the originally defined abbreviations; and, unless expressly stated to the contrary, measurements of a property should be performed using the same technique previously or subsequently cited for the same property.

[0034] Unless otherwise stated, all technical and scientific terms used herein have the same meaning as commonly understood by one of ordinary skill in the art to which this disclosure pertains.

[0035] It should also be understood that this disclosure is not limited to the specific embodiments and methods described below, as specific components and / or conditions can certainly vary. Furthermore, the terminology used herein is for describing particular embodiments only and is not intended to be limiting in any way.

[0036] It must also be noted that, unless the context clearly indicates otherwise, the singular forms "a," "an," and "the" as used in the specification and appended claims contain plural references. For example, referring to a component in the singular is intended to include multiple components.

[0037] The terms "or" and "and" are used interchangeably and can be understood as meaning "and / or".

[0038] The term "comprising" is synonymous with "including," "having," "containing," or "characterized by." These terms are inclusive and open-ended and do not exclude additional elements or methodological steps not listed.

[0039] The phrase "consisting of" excludes any element, step, or component not specified in the claim. When this phrase appears in a clause of the body of a claim, rather than immediately following the preamble, it only limits the elements listed in that clause; other elements are not excluded from the entire claim.

[0040] The phrase "consistent mainly of..." limits the scope of the claim to the specified materials or steps, as well as those materials or steps that do not substantially affect the essential and novel features of the claimed subject matter.

[0041] The terms "polynucleotide," "nucleotide," "nucleotide sequence," "nucleic acid," "polynucleotide," and "oligonucleotide" are used interchangeably in this disclosure. They refer to polymeric forms of nucleotides of any length, which may be deoxyribonucleotides or ribonucleotides, or analogs thereof. Polynucleotides can have any three-dimensional structure and can perform any known or unknown function. The following are non-limiting examples of polynucleotides: single-stranded, double-stranded, or multi-stranded DNA or RNA, genomic DNA, cDNA, DNA amplicon products, DNA restriction products, synthetic non-natural polynucleotide chains (including peptide nucleic acids (PNAs) and locked nucleic acids (LNAs)), DNA-RNA hybrids, or polymers containing purine and pyrimidine bases or other natural, chemically or biochemically modified, non-natural, or derived nucleotide bases. The terms "polynucleotide" and "nucleic acid" should be understood to include single-stranded (e.g., sense or antisense) and double-stranded polynucleotides as applicable to the described embodiments. Polynucleotides may contain one or more modified nucleotides, such as methylated nucleotides and nucleotide analogs. If modifications are present, the modification of the nucleotide structure may be performed before or after polymer assembly. Nucleotide sequences can be broken down by non-nucleotide components. Polynucleotides can be further modified after polymerization, for example, by binding to labeled components.

[0042] The term "complementarity" or "complementarity" refers to the ability of a nucleic acid to form hydrogen bonds with another nucleic acid sequence via conventional Watson-Crick or other non-conventional methods. The complementarity percentage indicates the percentage of residues in a nucleic acid molecule that can form hydrogen bonds with a second nucleic acid sequence (e.g., Watson-Crick base pairing) (e.g., complementarity rates of 66.67%, 83.33%, and 100% for 4, 5, and 6 residues out of 6 residues, respectively). "Complete complementarity" means that all adjacent residues in the nucleic acid sequence will form hydrogen bonds with the same number of adjacent residues in the second nucleic acid sequence. As used herein, "basic complementarity" refers to a complementarity of at least 40%, 50%, 60%, 62.5%, 70%, 75%, 80%, 85%, 90%, 95%, 97%, 98%, 99%, or 100% in regions of 4, 5, 6, 7, and 8 nucleotides, or somewhere in between, or refers to hybridization of two nucleic acids under stringent conditions.

[0043] Unless otherwise expressly stated to the contrary: the term "polymer" includes "oligomer," "copolymer," "terpolymer," etc.; the molecular weight provided for any polymer refers to the weight-average molecular weight unless otherwise stated; describing a group or class of materials as suitable or preferred for a particular purpose related to the present invention means that a mixture of any two or more members of that group or class is equally suitable or preferred; when describing components in chemical terms, it refers to the components added to any combination specified in the specification, and does not necessarily exclude chemical interactions between the components of the mixture after mixing; the initial definition of acronyms or other abbreviations applies to all subsequent uses of the same abbreviation herein and to normal grammatical variations of the originally defined abbreviations; and unless otherwise expressly stated to the contrary, measurements of a property shall be performed using the same technique as previously or subsequently cited for the same property.

[0044] It should also be understood that the integer range explicitly includes all integers within it. For example, the integer range 1-10 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, 100. Similarly, when any range is needed, the difference between the upper and lower limits divided by 10 can be used as an optional upper or lower limit. For example, if the range is 1.1 to 2.1, the following numbers can be selected as the lower or upper limits: 1.2, 1.3, 1.4, 1.5, 1.6, 1.7, 1.8, 1.9, and 2.0. In the specific embodiments described herein, concentrations, temperatures, and reaction conditions (e.g., pressure, pH, etc.) may be rounded to three significant figures within ±50% of the values ​​shown. In an improvement, concentrations, temperatures, and reaction conditions (e.g., pressure, pH, etc.) may be rounded to three significant figures within ±30% of the values ​​shown in the embodiments provided. In another improvement, concentration, temperature, and reaction conditions (e.g., pH) can be rounded to three significant figures within ±10% of the values ​​shown in the examples.

[0045] In the embodiments described herein, concentration, temperature, and reaction conditions (e.g., pressure, pH, flow rate, etc.) can be adjusted within ±50% of the values ​​shown, rounded or truncated to two significant figures of the values ​​provided in the embodiments. In one improvement, concentration, temperature, and reaction conditions (e.g., pressure, pH, flow rate, etc.) can be adjusted within ±30% of the values ​​shown, rounded or truncated to two significant figures of the values ​​provided in the embodiments. In yet another improvement, concentration, temperature, and reaction conditions (e.g., pressure, pH, flow rate, etc.) can be adjusted within ±10% of the values ​​shown, rounded or truncated to two significant figures of the values ​​provided in the embodiments.

[0046] Single-molecule DNA capture and stretching enable fundamental research into DNA polymer physics and the interaction of DNA with fluids and electric fields. Methods for capturing and stretching single-molecule DNA and simultaneously visualizing it include manipulating microbeads bound to DNA using optical or magnetic tweezers; subjecting DNA to stretching flow strain in microfluidic devices; binding DNA to a flat surface with binding chemicals and then subjecting it to fluid flow or an electric field, much like a so-called DNA curtain; and performing so-called DNA combing on silanized glass using a moving air / water interface. The combination of single-molecule DNA confinement and capture with high-quality imaging has also been applied to genetic analysis, including karyotype analysis and DNA mapping. For example, electromigration within nanochannels and imaging of long (>100kb) genomic DNA with sequence-specific markers are used for optical mapping in medical diagnostics.

[0047] However, such techniques also have certain drawbacks. For example, they lack throughput. Optical tweezers and magnetic tweezers can only study single molecules or at most a small number of molecules. These techniques also require additionally expensive and complex imaging and control systems. Molecular combing and DNA curtain analysis can image many molecules, but cannot cycle through multiple samples. Therefore, these analyses cannot resample the same population or analyze new samples. Thus, multiplexed data analysis is required to capture information from fast-moving molecules. Furthermore, because the molecules in these techniques are non-stationary (moving molecules), it is difficult to obtain high-quality images or study dynamic interactions. Other difficulties in using electromigration within nanochannels (including so-called nanoslit channels) for DNA separation include the difficulty in fabricating and sealing the channels and the difficulty in controlling the fluids in these systems, including liquid filling, generating pressure-driven flow, and generating electrofluids. Additionally, the fabrication methods of nanochannels can result in poor optical pathways within the channels. Furthermore, the walls of such nanochannels may have roughness comparable to the cross-sectional dimensions of the channel. Another challenge is that the surface properties of nanochannels may also be non-uniform, which can lead to non-uniform potentials and thus non-uniform electroosmotic flow rates, which generate velocity gradients in the fluid. Current methods for capturing DNA (with or without DNA visualization) include optical tweezers and magnetic tweezers as described above, and / or using so-called DNA combing or DNA curtain techniques to adsorb DNA onto a surface via a liquid / gas interface. These methods may require cumbersome manual steps and are not suitable for high-throughput applications that require the individual capture and stretching of multiple DNA molecules. Therefore, there is a need for systems and methods that facilitate the control and detection of single-molecule polynucleotide chains, which are both cost-effective and enable high-throughput sample processing.

[0048] This article discloses systems and methods for capturing, stretching, releasing, and / or recapturing multiple single-molecule polynucleotide chains. The disclosed systems and methods facilitate the control and detection (including visualization) of single-molecule polynucleotide chains.

[0049] This document discloses methods for capturing, stretching, releasing, and / or recapturing multiple single-molecule polynucleotide chains. A polynucleotide chain can refer to a chain of linked nucleotides and may include, for example, DNA or RNA. A polynucleotide chain may be referred to as a chain or a molecule. One or more methods facilitate single-molecule control and visualization. According to various method embodiments, polynucleotide chains in a sample can be immobilized along their length at the apex to the walls of a microchannel. Polynucleotide chains can be captured, immobilized, and stretched using an applied axial electric field. This enables high-quality imaging and quantification of polynucleotide chain conformations. In one or more embodiments, this disclosure demonstrates a set of conditions for capturing polynucleotide chains; quantifying capture kinetics; and quantifying relaxation kinetics, untangling, and release from the immobilized state.

[0050] According to at least one embodiment, a method for capturing and stretching polynucleotide chains is disclosed. This method facilitates the capture of polynucleotide chains at the surface of a channel. The channel may be a microfluidic channel. As used herein, the term channel also includes a fluid chamber in fluid communication with at least one port, said at least one port possibly for filling the chamber with liquid. A channel may be part of a microfluidic device comprising one or more channels. The one or more channels may each have a single inlet and a single outlet. Multiple independent channels may exist. Samples in each of the independent channels may be processed in parallel. The surface of the channel may be a dielectric material including oxides. For example, the channel surface may be silicon oxide, aluminum oxide, titanium oxide, or various other oxide surfaces. For example, the channel surface may also be a nitride, such as silicon nitride. The surface may also be glass, including borosilicate glass or quartz. The channel surface may also be a dielectric material having regions of non-dielectric material (e.g., metal or semiconductor). The dielectric and non-dielectric materials may be arranged in a specific pattern on the surface. Polynucleotide chains may be captured in the dielectric regions but not in the non-dielectric regions. The channel may also include a channel cap. The channel cap may be borosilicate glass. The channel may include a bottom and one or more walls extending outward from the bottom. The walls and / or cap of the channel may be planar. The surface of the channel may include surfaces etched into the substrate or the channel cap. The surface of the channel may alternatively include a defined portion of either the substrate or any wall of the channel or the channel cap. In some embodiments, the channel may have a depth of 500 nm to 3 μm. Channel depths within this range may, for example, facilitate the easy acquisition of high-quality images of polynucleotide chains examined by epifluorescence microscopy. In other embodiments, the channel may have a depth greater than 3 μm. For example, with channel depths greater than 3 μm, high-quality images of polynucleotide chains may be obtained by total internal reflection fluorescence (TIRF) microscopy. In various embodiments, the channel may also have a nominal transverse width greater than 1 μm. The channel may optionally include one or more pillars supporting the cap and / or a castellation pattern on one or more sidewalls of the channel. The channel may have at least one dimension of less than 5 micrometers perpendicular to the applied electric field.

[0051] The transfer of reagents and / or liquids in the system may be accomplished using processes including electromigration, electroosmotic flow, pressure-driven flow, dielectrophoresis, magnetophoresis, electrowetting, induced charge electroosmotic flow, alternating current (AC) electrodynamic flow, gravity-driven flow, and surface tension-driven flow.

[0052] This method may involve providing a polymer to the surface of the channel. The polymer is capable of suppressing electroosmotic flow in 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 that may be used as an additive in a buffer system to suppress electroosmotic flow (EOF) for performing electrodynamic experiments. The polymer may alternatively be polyethylene glycol (PEG), which, in the context of polymer chemistry, may also be called polyethylene oxide (PEO). PEG may be used in gas chromatography and may also be used to coat or treat silica surfaces to suppress EOF. PEG with a sufficiently large molecular weight may also allow for DNA vertex fixation. The polymer may alternatively be hydroxyethyl cellulose (HEC). HEC, as a linear polymer, may suppress EOF. The polymer may alternatively be polyvinyl alcohol (PVA). PVA may be used for the separation of DNA restriction fragments and effectively suppresses EOF below about pH 8.

[0053] The method may further include physically interacting the polymer with the surface of the channel. This step may be achieved by coating the surface with the polymer. Coating may be achieved by delivering the polymer to the surface and allowing the polymer to adsorb onto the surface. The surface may include the bottom or wall of the channel, a portion of the bottom or wall of the channel, a channel cap or a portion of the channel cap, or a surface not in physical contact with the channel cap. For example, the polymer may be able to adsorb onto the silica wall of the channel or the borosilicate glass wall of the channel cap. In at least some embodiments, metals to which the polymer cannot adsorb may be patterned and deposited onto the oxide substrate. In this way, only regions of the oxide substrate with a sufficient electric field and the presence of a linear polymer interacting with the surface (e.g., not patterned with metal) are likely to trap polynucleotide chains.

[0054] The method may further include providing a sample comprising polynucleotide chains to a polymer at the surface of the channel. The sample may comprise multiple polynucleotide chains. The polynucleotide chains may be isolated or synthesized. The polynucleotide chains may originate from the same or different sources. For example, different sources may include polynucleotide chains from different organisms (e.g., humans) or the same organism. A single sample may comprise polynucleotide chains from both control and experimental sources. The polynucleotide chains in the sample may include barcodes identifying the source or type of the polynucleotide chains. Sources may include, for example, human, plant, or animal cells or tissues, microphysical systems including organoids, spheroids, tumor organoids and similar systems, bacterial or protozoan cells, or synthetic reactions that produce polynucleotide chains. The sample comprising polynucleotide chains may additionally include one or more buffer components. In at least one embodiment, the sample provided to the channel may comprise polynucleotide chains, buffer components, and a polymer.

[0055] The method further includes applying an electric field to the polynucleotide chain to promote physical interactions between the polynucleotide chain and the polymer, thereby capturing (also referred to as immobilizing) the polynucleotide chain at the surface of the channel. The electric field may be 30 to 400 V / cm. The electric field may be applied to the electrolyte via two or more electrodes supporting a Faraday reaction. These Faraday reactions can support ionic currents in the channel. The applied electric field associated with these ionic currents may be substantially parallel to the wall on which the polynucleotide chain to be captured is located. For example, the inner wall of the channel may be a dielectric material such that the applied electric field is parallel to the wetting wall of the channel and drives electromigration (including electrophoresis) of the polynucleotide chain parallel to the wall of the channel. Alternatively, the electric field promoting the interaction between the polynucleotide and the wall may be applied using an electrode coupled to the electrolyte capacitance, thereby avoiding or mitigating Faraday reactions in the system. The electric field may be applied to the channel, thereby affecting the channel and its contents, including any fluid sample. For example, applying an electric field to the channel may result in the electric field being applied to the polynucleotide chain in a sample flowing through the channel. Physical interactions between polynucleotide chains and polymers can occur at the vertices of the polynucleotide chain. These interactions can occur at a single vertex along the length of the polynucleotide chain. The two ends of the polynucleotide (also known as arms) can extend outward from the vertex in the opposite direction of the applied electric field.

[0056] When a polymer with a sufficiently large molecular weight (e.g., greater than 100 kDa) is provided to the surface of the channel, and a sufficiently large axial electric field (e.g., >30 V / cm) is applied, trapping of the polynucleotide chain via vertex fixation and stretching occurs. For larger molecular weight polymers, the threshold with respect to the electric field may be lower. For example, the threshold with respect to the electric field is lower for larger molecular weight PVP polymers. The trapping and stretching phenomena occur only at the surface and on both thermally oxidized silica and glass surfaces.

[0057] Without favoring any particular theory, under sufficiently high electric fields, polynucleotide chains may become entangled with surface-adsorbed polymers, and the strong electric force may dissipate the hydrated layer between the nucleic acid and the polymer, leading to a high-friction state. After the electric field is removed, the space between the nucleic acid and the polymer may become rehydrated, and the resulting lower friction may allow Brownian motion to untangle the polynucleotide chains from the polymer. The spatial correlation across which no vertex fixation occurs suggests that this phenomenon is independent of specific channel geometry.

[0058] The method may also include reducing the electric field below an empirically determined threshold or removing the electric field to facilitate the release of the polynucleotide chain from its physical interaction with the polymer. After removing the electric field above the empirically determined threshold, the polynucleotide chain may relax towards and around a fixed vertex. Fixed positioning may be a single-point entanglement between the polynucleotide chain and the polymer. After the polynucleotide chain relaxes towards and around the fixed vertex, it may unwrap from the polymer and resume movement through the channel. The movement of the unwrapped polynucleotide chain through the channel may occur by diffusion or electromigration. The relaxation, unwrapping, and movement of the polynucleotide chain may clear the polynucleotide chain from the channel, such that a second sample including the polynucleotide chain may be provided to the channel for capture and stretching. In this way, the method may include a cyclical arrangement comprising the following steps: introducing the polynucleotide chain into the region of interest, capturing the polynucleotide chain under a high electric field, imaging the polynucleotide chain once or multiple times (including detecting various wavelengths, reducing the electric field to relax the polynucleotide chain (which may lead to unwrapping), applying an electric field that induces electromigration of the polynucleotide chain with minimal or no capture), and then repeating the process. In this way, the next sample, including a polynucleotide chain, may be fed into the channel. For example, after the first sample has been captured, relaxed, untangled, and cleared, a second sample may be provided to the channel for capture. The capture, relaxation, untangling, and clearing process may be repeated for n samples. This cyclical arrangement may allow for high-throughput processing of many independent samples and may also allow for multiple resampling of the same sample to enhance output. The second, next, or nth sample may include a polynucleotide chain present in the first or previous sample. Further, a pulse of pressure-driven flow may be introduced, resulting in much faster untangling. In some embodiments, the step of deactivating the electric field may be replaced by a step of providing a pulse of pressure-driven flow to the channel. This cyclical arrangement of steps in the method may enable high-throughput capture and analysis of multiple polynucleotide chains.

[0059] Figure 1A A method 100 for capturing and stretching polynucleotide chains according to one embodiment is illustrated. The method includes step 102 of providing a polymer to the surface of a channel. The method further includes step 104 of physically interacting the polymer with the surface of the channel. Step 106 includes providing a sample comprising the polynucleotide chain to the polymer interacting with the surface of the channel. The method further includes step 108 of applying an electric field to the polynucleotide chain to facilitate physical interaction between the polynucleotide chain and the polymer to capture the polynucleotide chain at the surface of the channel.

[0060] Figure 1B A method 101 for high-throughput capture and stretching of polynucleotide chains is illustrated according to one embodiment. This method may facilitate the capture, stretching, and release of polynucleotides, allowing for sequential processing of multiple samples. The method includes a step 102 of providing 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 providing a sample comprising a polynucleotide chain to the polymer interacting with the surface of the channel. The method further includes a step 108 of applying an electric field to the polynucleotide chain to facilitate physical interaction between the polynucleotide chain and the polymer to capture the polynucleotide chain at the surface of the channel. Step 109 includes adjusting the electric field to release the captured polynucleotide chain from the physical interaction with the polymer. Reducing the electric field may cause relaxation of the polynucleotide chain, which in turn leads to untangling and free diffusion and electromigration to clear the polynucleotide chain, allowing a next sample comprising the polynucleotide chain to be introduced into the channel. The steps of this method may then be repeated n times to achieve high-throughput processing of polynucleotide chains. In an alternative embodiment, step 109 may include providing a pulse of pressure-driven flow to the channel. Pulses may also release polynucleotide chains by promoting relaxation, untangling, free diffusion, and electromigration of polynucleotide chains.

[0061] Figure 2A Another method 110 for capturing and stretching polynucleotide chains according to one embodiment is shown. The method includes step 112 of providing a sample comprising a polymer and a polynucleotide chain to the surface of a channel. Step 114 includes physically interacting the polymer with the surface of the channel. A further step 116 includes applying an electric field to the polynucleotide chain to facilitate physical interaction between the polynucleotide chain and the polymer to capture the polynucleotide chain at the surface of the channel.

[0062] Figure 2B An additional method 111 for high-throughput capture and stretching of polynucleotide chains is illustrated according to one embodiment. This method may facilitate the capture, stretching, and release of polynucleotides, allowing for sequential processing of multiple samples. The method includes step 112 of providing a sample comprising a polymer and a polynucleotide chain to the surface of a channel. Step 114 includes physically interacting the polymer with the surface of the channel. An additional step 116 includes applying an electric field to the polynucleotide chain to facilitate physical interaction between the polynucleotide chain and the polymer to capture the polynucleotide chain at the surface of the channel. Step 117 includes adjusting the electric field to release the captured polynucleotide chain from the physical interaction with the polymer. Reducing the electric field may cause relaxation of the polynucleotide chain, which in turn leads to untangling and free diffusion and electromigration to clear the polynucleotide chain, allowing a next sample comprising the polynucleotide chain to be introduced into the channel. The steps of this method may then be repeated n times to achieve high-throughput processing of the polynucleotide chain. In an alternative embodiment, step 117 may include providing a pulse of pressure-driven flow to the channel. Pulses may also release polynucleotide chains by promoting relaxation, untangling, free diffusion, and electromigration of polynucleotide chains.

[0063] Figure 3A and 3B A system for capturing and stretching polynucleotide chains is shown. Figure 3A System 118 is shown, comprising device 120, voltage source 122, and controller 124 programmed to interact with device 120 and voltage source 122. Device 120 may include at least one reservoir sized to receive a sample comprising a polynucleotide chain. The reservoir may also include electrodes. Alternatively, device 120 may include a first reservoir sized to receive a sample comprising a polynucleotide chain and a second reservoir sized to receive an electrode. These reservoirs may have electrical and / or ion communication between themselves or among other electrodes in the system. For example, the electrodes (either alone in the reservoir or together with the sample comprising a polynucleotide chain) may be metals, metal alloys, or semiconductor materials. Other possible materials include graphite or graphene or other forms of carbon, including active porous carbon. Electrodes may be fabricated using microfabrication techniques including vapor deposition, lift-off techniques, or doping. Electrodes may also be electroplated or coated. These electrodes may drive Faraday reactions (including water splitting at the electrode) to introduce an electric field within the channels of the system. The electrodes may also exhibit significant capacitive properties, including porous carbon electrodes operating at low voltages to avoid Faraday reactions. 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 wires. Alternatively, the electrodes may be gold, silver, titanium, palladium, copper, stainless steel, titanium nitride, or silver / silver chloride. The electrodes may also be made of doped silicon. The voltage waveform applied by the electrodes and the resulting electric field can be direct current (DC), alternating current (AC), pulse-modulated, and can be very complex waveforms. For example, the waveform may include both DC and AC components and can be periodic, aperiodic, antiperiodic, or non-antiperiodic.

[0064] The device may also include a chip. The chip may be a silicon substrate chip. The chip may have two separate (independent) single-input, single-output channels. The channels may be fabricated such that they can be simultaneously visualized in 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 one or more reservoirs. Multiple independent channels may exist. Samples in each independent channel may be processed in parallel. The device may also include channel caps. The channel caps may be glass. The channel caps may be, for example, borosilicate glass. One or more walls of the channels may be optically transparent and / or may facilitate efficient transmission of electromagnetic radiation such as infrared or ultraviolet. One or more walls of the channels may also facilitate significant transmission, reflection, refraction, and / or diffraction of electromagnetic radiation. As defined herein, electromagnetic radiation includes optical (visible) wavelengths, infrared, and ultraviolet light. The system may allow and focus magnetic fields, which may be within the device, including applying magnetic forces to magnetic particles.

[0065] The system may also include a voltage source and a controller. The voltage source may apply an electric field to the polynucleotide chains in the channel, and may apply a voltage using electrodes in a reservoir. The voltage source may also measure current. The controller may be programmed to interact with the voltage source. For example, the controller may be programmed to direct the voltage source to apply a voltage or measure a current. The controller may be programmed to modulate the electric field applied to the channel. Additionally, the controller may be programmed to direct a polymer flow to the device to facilitate polymer delivery to the surface of the channel, allowing the polymer to physically interact with the surface of the channel. The controller may similarly direct a sample flow, with or without polymer, to the device to facilitate sample delivery to the surface of the channel. The controller may also be programmed to guide the electromigration of polynucleotide chains through the channel. Figure 3B A system 119 for capturing and stretching polynucleotide chains according to one embodiment is shown. Voltage source 122 is part of controller 124.

[0066] The processes, methods, or algorithms disclosed herein may be deliverable to / implemented by a processing device, controller, or computer, which may include any existing programmable electronic control unit or dedicated electronic control unit. Similarly, processes, methods, or algorithms may be stored in various forms as data and instructions executable by a controller or computer, including but not limited to information permanently stored on non-writable storage media such as ROM devices, and information modifiable stored on writable storage media such as floppy disks, magnetic tapes, CDs, RAM devices, and other magnetic and optical media. Processes, methods, or algorithms may also be implemented in an executable software object. Alternatively, processes, methods, or algorithms may be embodied, wholly or partially, using suitable hardware components such as application-specific integrated circuits (ASICs), field-programmable gate arrays (FPGAs), state machines, controllers, or other hardware components / devices, or a combination of hardware, software, and firmware components.

[0067] This document also provides methods for spatially patterning the localization of potentially trapped polynucleotide chains within channels. These methods allow polynucleotides to be trapped within specific regions of interest. In one embodiment, the method includes providing a polymer to a designated location within a fluid system. For example, laminar flow features may be used to provide the polymer only to certain subsurfaces and prevent the polymer from physically interacting with other surfaces. This approach may utilize hydrodynamic focusing to pattern the localization of wall surfaces processed by the polymer. Figure 4 Channel 126 is shown having more than one fluid layer 128a, 128b, 128c. The first fluid layer 128a comprises polynucleotide chains but not polymers. The second fluid layer 128b comprises both polynucleotide chains and polymers. The third fluid layer 128c comprises polynucleotide chains but not polymers. Because the second fluid layer 128b comprises polymers, the polynucleotides are trapped in the second fluid layer 128b. In the first layer 128a and the third layer 128c, the polynucleotide chains migrate in a 3D spherical shape.

[0068] In alternative implementations, methods for spatially patterning the localization of potentially trapped polynucleotide chains within the channel may include shaping the channel walls and / or adding features to influence the electric field in a specific defined region. For example, barriers may be added to locally influence the electric field within the channel. This approach leverages the fact that polynucleotide chain trapping occurs above a certain threshold electric field. Figure 5 An example of channel 130 is shown, which has low electric (E) field regions 132a and 132b and a high electric field region 134. Polynucleotide chains are fixed and stretched at vertices only in the high E field regions, while polynucleotide chains in the low E field regions migrate in a 3D spherical shape.

[0069] In other embodiments, the method for spatially patterning the location of potentially trapped polynucleotide chains within the channel may include patterning a polymer-free metal coating region on the fluid chip. Figure 6 An example of channel 136 is shown, which has a surface 138 to which a polymer may be adsorbed, and a defined region of patterned material 140 to which the polymer is not adsorbed.

[0070] Systems and methods for capturing, stretching, and detecting polynucleotide chains. The systems and methods disclosed herein in one or more embodiments describe systems and methods for vertices, relaxation, unwinding, and free electromigration cycles to capture and stretch single-molecule polynucleotide chains. Vertex fixation and stretching may be alternated with polynucleotide chain relaxation, unwinding, and low-field clearance to cycle through multiple polynucleotide chains. These systems and methods may be further modified to detect polynucleotide chains. For example, these systems and methods may be modified to achieve high-throughput capture, stretching, and imaging of polynucleotide chains. The systems and methods may also be modified for the purpose of determining the sequence of polynucleotide chains. The systems and methods may facilitate the capture and stretching of polynucleotide chains at specific regions of interest within a channel. Specific regions of interest may, for example, be fields of view within the channel.

[0071] Figure 7A and 7B A method 142 for optical imaging of a polynucleotide chain for vertex capture is illustrated, the optical imaging being used to visualize a subsequence of the polynucleotide chain. Step 144 includes providing a sample comprising at least one polynucleotide chain and labeling the polynucleotide chain to produce a labeled polynucleotide chain. The polynucleotide chain may be significantly larger than 1 kb. For example, the length of the polynucleotide chain may exceed 300 kb. At least one sequence of 4 to 10 base pairs within the polynucleotide chain may be labeled to form a labeled subsequence. In this way, the labeled polynucleotide chain is a polynucleotide chain having at least one labeled subsequence. In one instance, the polynucleotide chain may be DNA. Regions containing the subsequence may be labeled with a fluorescent label. More than one subsequence may be labeled on the DNA chain. The subsequence may also be referred to as a target sequence, subsequence, coding, or barcode element. Multiple types of labels may be used, including fluorescent labels with various emission wavelengths. Step 146 includes capturing and visualizing the labeled polynucleotide chain. Step 146 includes substeps 147, 148, 150, and 152. Sub-step 147 involves providing the channel with a polymer capable of modulating electroosmotic flow within the channel. Sub-step 148 involves providing the channel with a sample comprising a labeled polynucleotide chain. In some embodiments, the polymer may be included in the sample comprising the labeled polynucleotide chain. The polymer may be a neutral and water-soluble polymer. An electric field higher than an empirically determined threshold is then applied to the polynucleotide chain in the channel (sub-step 150). The threshold may depend on the polymer used and the molecular weight of the polymer. The labeled polynucleotide chain then physically interacts with and is trapped by the polymer at the surface of the channel. After being trapped, the free ends (also referred to as arms) of the labeled polynucleotide chain extend in the opposite direction of the electric field. Simultaneously with the trapping, the labeled polynucleotide chain may be imaged by a detector (step 152). In one example, the fluorescently labeled DNA chain may be imaged using an epifluorescence microscope and a scientific-grade complementary metal-oxide-semiconductor (scientific-grade-CMOS) camera, a charge-coupled device (CCD) camera, or an electron-multiplying charge-coupled device (EMCCD) camera. Other methods that may be used to image the captured polynucleotide chains (in addition to epifluorescence microscopy) include light scattering microscopy, total internal reflection microscopy (TIRF), super-resolution structure illumination microscopy (SR-SIM), stimulated emission depletion microscopy (STEP), stochastic optical reconstruction microscopy (STORM), or single-molecule localization microscopy (SMLM).

[0072] In at least one embodiment, the apex-fixed DNA of the label may be captured and elongated in a shallow microfluidic channel under an applied electric field for high-quality single-molecule imaging. Step 154 ​​includes programming a controller to collect, record, and store signals derived from the detector. The signals may then be processed (step 156). On a computer chip (in silico), the image of the DNA may be interpreted to analyze the sequence of the label. For example, the label spacing forms patterns that can be detected and analyzed. Regions of captured DNA that do not overlap with other DNA regions are easier to interpret than regions where one segment of DNA overlaps with another segment of DNA. Overlapping regions in "hairpin"-shaped DNA can be analyzed in a computer (on a computer chip), and the pattern of the DNA can be "unfolded" computationally to determine the pattern along the DNA in its unfolded form. That is, the label pattern associated with unobserved unfolded DNA can be deduced from the imaged labels and may be reconstructed computationally.

[0073] As described above, applications may include imaging of fluorescently labeled markers on the 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 nature described herein and the folding properties of DNA, there are at least two classes of DNA shapes associated with DNA captured in this manner. For many (and most likely most) DNAs wrapped around a point along their length, there will be regions where two DNA arms substantially overlap. In this overlapping region, some subsequences oriented from 3' to 5' can largely overlap with some subsequences oriented from 5' to 3'. The two observable subsequences in this first overlapping region converge near the vertex. By reading the overlapping region and then unfolding the adjacent sequences using computational methods, the two separate sequences in this overlapping region can be read. This will involve (in a computer program) analyzing the 3' to 5' overlap, its neighboring 5' to 3' type sequences. Once the overlapping sequences are determined, the two overlapping regions can be interpreted in a computational domain.

[0074] In addition to the overlapping region, most DNA sequences also have a significant second region far from the vertex, in which the longer DNA arms do not overlap with the shorter arms. This region contains subsequences that can be of type 3' to 5' or 5' to 3'. This sequence can then be read and interpreted normally.

[0075] Methods for interpreting overlapping DNA arm regions may include first identifying non-overlapping regions and then using this information to help interpret overlapping regions. Alternative methods for interpreting overlapping DNA arm regions may include maintaining a database (in a computer) of codes associated with overlapping or partially overlapping regions and comparing the overlapping regions with these codes.

[0076] This article also provides a system for optical imaging of labeled vertices to capture polynucleotide chains for the purpose of determining the sequence of the polynucleotide chains. Figure 8 A system 158 according to one embodiment is shown. System 158 may include a device 160, at least one detector 162, a voltage source 164, and a controller 166. Device 160 may include one or more reservoirs. Any of the one or more reservoirs may be sized to receive a sample comprising a polynucleotide chain. Any of the one or more reservoirs may be sized to receive at least one electrode. For example, the one or more electrodes (either alone in the reservoir or together with the sample comprising a polynucleotide chain) may be metal, metal alloy, or 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 a single-layer or multi-layer graphene. The electrode may be graphite. The electrode may also be an inkjet-printed carbon-deposited wire. Alternatively, the electrode may be gold, silver, titanium, palladium, copper, stainless steel, titanium nitride, or silver / silver chloride. The electrode may also be made of doped silicon. The device may also include a chip. The chip may be a microfluidic chip. The chip may be a silicon-based chip. The chip may have two separate (independent) single-input, single-output channels. The channels may be fabricated such that they can be simultaneously visualized 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 one or more reservoirs. The device may also include channel caps. The channel caps may be glass. The channel caps may be, for example, borosilicate glass.

[0077] The system may also include at least one detector. This may include an optical detector configured to visualize polynucleotide chains captured in the channel. For example, the optical detector might be an epifluorescence microscopy system. An epifluorescence microscopy system might include a water immersion objective, an illumination source, and at least one camera.

[0078] The system may also include a voltage source for applying an electric field to the polynucleotide chains in the channel. The voltage source may apply a voltage using electrodes in the reservoir. The voltage source may also measure current.

[0079] 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 pumps or flow component components, either external to or within the device. The control system may also be programmed to interact with at least one detector and a separate voltage source (if included). For example, the controller may direct a polymer flow to the device to facilitate polymer delivery to the surface of a channel, enabling the polymer to physically interact with the surface of the channel. Similarly, the controller may direct a sample flow, including labeled polynucleotide chains (with or without polymer), to the device to facilitate sample delivery to the surface of the channel. The controller may also be additionally programmed to interact with at least one detector. For example, the controller may be programmed to receive, record, and / or store signals acquired by at least one detector. The controller may also be programmed to process signals from at least one detector or from other sensors integrated into the system. These other sensors may be within or external to the fluid device. Example

[0080] The following examples illustrate various embodiments of this disclosure. Those skilled in the art will recognize many variations within the spirit and scope of this disclosure.

[0081] Example 1: Capturing long single DNA strands Figures 9A-9C A method for DNA capture according to one embodiment is illustrated. In this example, a vertex-fixed, single-molecule 48 kbp DNA is displayed in a microfluidic channel filled with a linear polymer buffer solution and subjected to an applied axial electric field. Figure 9A An isometric schematic diagram of DNA fixed at a vertex is shown, the DNA having two arms extending in the opposite direction to the applied electric field. Figure 9B Experimental epifluorescence images of a single DNA molecule with its vertex fixed are shown. The images show stretching at 350 V / cm, 20 V / cm, and no applied electric field, where the DNA relaxes around the vertex into a Brownian coil. Figure 9C Three consecutive images of single-molecule DNA fixation after application of 300 V / cm are shown. The fourth and fifth images are two additional experimental runs of the capture process. The solution was 4 pM YOYO-1 labeled λ-DNA in 1X TBE buffer containing 2% w / w 1300 kDa PVP and 2% v / v β-mercaptoethanol. The channels had a depth of 0.9 μm and a nominal transverse width of 30 μm (see Figure 14 for more details). The channels included a zigzag pattern on one side, which was not essential for capture (see Figures 15 and 16 for further discussion). Figure 9A A schematic diagram of each DNA fixed at the apex of the wall is shown, the DNA having two relaxed arms extending in the opposite direction of the applied electric field, consistent with the electrostatic force on the polymer and resisting any electroosmotic flow (EOF). Figure 9B Serial pseudo-color epifluorescence images of a single 48.5 kbp DNA molecule are shown under electric fields of 350, 20, and 0 V / cm. Initially, the DNA is captured at the high field. As the field decreases, DNA capture persists, and the DNA arms overlap to a lesser extent due to Brownian motion. The electric field is turned off at t = 3.5 s, and the DNA molecule coils into a three-dimensional (3D) cloud-like structure centered on the aforementioned vertex fixation point (at t = 7 s). Figure 9C The leftmost small image shows a bright-field image of the channel geometry. The image also shows three consecutive images of aggregates of single-molecule DNA with fixed capture vertices. The process can be repeated, allowing for high-throughput visualization of many DNA molecules within the same field of view. Figure 9C The last two images show two subsequent experimental runs of the capture process. Similar data for 20 kbp DNA are available in SM (reference). Figure 17 Provided in ).

[0082] Experiments were conducted under electric fields ranging from 19 to 455 V / cm and under various buffering chemistry conditions, including those containing and without linear polymer additives typically used to suppress electroosmotic flow. DNA capture via vertex fixation was observed only under sufficiently high electric fields and only in the presence of polyvinylpyrrolidone (PVP) with a molecular weight (MW) of 360 kDa or greater. In one or more embodiments, the PVP polymer is at least partially adsorbed onto the glass and / or oxide channel walls, thereby increasing the local solution bulk viscosity within the electrical bilayer. Individual DNA molecules may become entangled with one or more adsorbed PVP molecules, and this entanglement most often occurs at a single point along the DNA, resulting in the configuration described in Figure 9. Fixation occurs on the silica walls of the channel and on the surface of the glass walls used to seal the channel (see Figure 15). Figure 15A and 15B DNA apex fixation at the bottom and top surfaces of a straight channel 3 μm deep is shown under an axial electric field strength of 150 V / cm. Figure 15A DNA is shown fixed at its apex on a bottom surface, which is a thermally grown thermal oxide. Figure 15B DNA is shown fixed at the apex on the top surface, which is anodically bonded borosilicate glass. Figure 15A and 15B Two consecutive images from a single experimental run are shown. The depth of focus is estimated to be 0.7 μm.

[0083] Figures 16A to 16D A single DNA molecule is shown fixed at its apex in a 37 μm deep commercial glass microfluidic channel filled with a linear polymer solution and subjected to an axial electric field. Figure 16A A schematic diagram of a commercially available glass chip purchased from Microfluidic ChipShop is shown. Four parallel glass channels, each 58.5 mm long, were fabricated on each chip and sealed with a 210 μm cap. All materials are glass. Figure 16B and 16C This image shows a raw, continuous epifluorescence image of DNA electromigration 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. Vertex-fixed DNA can be observed against the background near the wall surface. Figure 16D The global temporal median of the image sequence is shown. The image is magnified to show DNA fixed at the vertices of the wall.

[0084] Figure 17 A single 20 kbp DNA molecule immobilized at its apex in a 0.9 μm deep microfluidic channel filled with a linear polymer buffer and subjected to an applied axial electric field is shown. A wide-field continuous raw image of the single DNA molecule immobilized after an application of 254 V / cm is presented. The solution was 4 pM YOYO-1 labeled 20 kbp DNA in 1X TBE buffer containing 2% w / w 1300 kDa PVP and 2% v / v β-mercaptoethanol. (See related information.) t = 0, 1.4, 11.5, and 28.4 s original images. The faint lines in the background are due to the streaking effect of 20 kbp DNA under rapid electromigration.

[0085] Example 2: Testing the effect of electric field on DNA capture (based on the effect of field strength on the amount of single DNA molecules fixed at the vertex). (Experimental Quantitative) The effect of the applied electric field magnitude E on the initiation and rate of DNA capture was investigated. A custom image processing algorithm was developed and used to quantify the number of DNA cells fixed according to E. Figure 10A This analysis is summarized and further details are provided. Figure 10A A flowchart outlining the image processing used for quantifying DNA capture is shown. The median of a small number of images is then subjected to adaptive thresholding. The binarized image is used to create an α-shaped boundary mask, and the original image data is integrated within the region highlighted by the binarized mask. Figure 10B The area-mean-time median intensity of the experiment for a total duration of 30 seconds under each applied E is shown.<I_med> All data represent 4 pMYOYO-1 labeled λ-DNA (48.5 kbp) in 1X TBE buffer containing 2% v / v β-mercaptoethanol. Dark gray curves with circle markers and light gray curves with square markers represent experiments using solutions prepared by PVP MW (both 2% w / w concentration) at MW 1300 or 360 kDa. Dashed vertical lines highlight the approximate threshold field observed at the beginning of DNA capture. In short, moving median images were computed from a small set of images to enhance the signal from stationary DNA (3-image sequences with respect to the maximum E and 21-image sequences with respect to the minimum E). Adaptive local thresholding was then applied to obtain binarized images. These binarized images were used as input for the α-shape algorithm, followed by dilation and erosion to obtain an α-shape boundary mask. This mask was applied to the aforementioned moving median image sequences, and the product was integrated to obtain a scalar measurement (as of time) of the stationary DNA with respect to each E. In Figure 18, this scalar is well correlated with a sample manual count of the captured DNA (obtained by manually analyzing each frame). Figures 18A to 18D The area-averaged α-shape boundary strength is shown. Comparison with results from manual molecular counting. Curves show the α-shape boundary strength over time for electric field strengths of 18, 99, 164, and 235 V / cm. Scatter plots show the manual counts of each wrapped DNA at the corresponding time steps for electric field strengths of 18, 99, 164, and 235 V / cm. All data are for 4 pM YOYO-1 labeled λ-DNA (48.5 kbp) in 1XTBE buffer containing 2% v / v β-mercaptoethanol supplemented with 1300 kDa PVP polymer. The α-shape boundary strength over time shows a similar trend to the manual counting results. For each E, we calculated the median of the scalar time-series data from the automated analysis over a duration of 30 seconds. Figure 10B These median scalar measurements of E are shown for 4 pM YOYO-1-labeled λ-DNA in 1X TBE buffer containing 2% β-mercaptoethanol. Data for 2% w / w PVP are shown for MW of 1300 (curve with circle markers) or 360 kDa (curve with square markers). The E-field data (for 360 and 1300 kDa) each represent the approximate threshold E value required for initiating capture (see also...). Figure 19 (See Figure 20). The observed thresholds were 70 V / cm for 1300 kDa PVP and 180 V / cm for 360 kDa PVP. We also performed experiments using a buffer containing 2% w / w PVP with 10 MW and 58 kDa PVP. No DNA capture was observed with solutions prepared using these lower, commonly used, commercially available PVP weights. Referring again to the aforementioned PVP entanglement hypothesis, these observations suggest that sufficiently long PVP is required for adsorption to the channel surface and the presentation of a PVP scaffold that allows DNA entanglement on the PVP.

[0086] Figure 19 Experimental quantification of the amount of single-molecule DNA at a fixed vertex is shown based on the electric field strength. The median intensity of the electric field for each experimental run with the field applied for 30 seconds is plotted on the ordinate relative to the axial electric field strength for each experimental run on the x-axis. Figure 10B Unlike other methods, the intensity here is normalized by the applied axial field strength. All data are for 4 pM YOYO-1 labeled λ-DNA (48.5 kbp) in 1X TBE buffer containing 2% v / v β-mercaptoethanol. Curves with circle markers and curves with square markers represent experiments using PVP with MW of 1300 and 360 kDa, respectively, supplemented with 2% w / w concentration.

[0087] Figures 20A to 20D Experimental quantification of the amount of vertex-fixed, single-molecule DNA based on time and field strength is shown. The α-shaped boundary mask strength as a function of time is shown for experimental runs performed under multiple separate electric fields. All data were obtained from 4 pM YOYO-1 labeled λ-DNA (48.5 kbp) in 1X TBE buffer containing 2% v / v β-mercaptoethanol. Figure 20A and 20B With the addition of 2% w / w 1300 kDa PVP polymer in the buffer, while Figure 20C and 20D The addition of 2% w / w 360 kDa PVP polymer to the buffer solution. Figure 20B and 20D The intensity is normalized by applying the axial electric field intensity for each experimental run.

[0088] The data suggests that high-magnitude electric fields are required for the high frictional forces between DNA and PVP polymers. Given the threshold nature of the required field, sufficiently strong electric forces may be needed on the DNA to dislodge the hydration layer between the DNA and PVP molecules. The removal of the hydration layer may increase solid-state friction between the charged DNA polymer and the non-charged linear PVP polymer. This could lead to persistent high-frictional tangles observed at fixed vertices. This hypothesis is consistent with the fact that persistent fixation can occur at vertices near the ends of DNA molecules, resulting in two loose arms with significantly different lengths and thus experiencing vastly different forces. The removal of the hydration layer between individual DNA molecules and polymer molecules has been addressed in studies of DNA interactions with agarose crosslinked polymers. Studies of DNA electrophoresis through 3D agarose gel networks have shown that DNA can be trapped under strong electric fields. Another study observed DNA trapping within agarose gels for electric fields exceeding a specific threshold and proposed the "knot" trapping hypothesis. However, in these previous studies, the tangled DNA formed complex 3D shapes inside the gel, rather than being fixed at "clean" vertices with two straight arms (in the linear polymer solution) at the walls, as in the current work.

[0089] Example 3: Testing the relaxation dynamics of vertex-fixed DNA The relaxation dynamics of vertex-fixed DNA after the electric field was removed were analyzed. Figure 11A The time series of the images is shown. We observed that DNA is fixed at random locations along the length of the DNA molecule, resulting in a series of relaxed time scales. We employed an automated image processing method based on adaptive thresholding, fitting the DNA image with an ellipse, and extracting the major and minor axes of the best-fit ellipse. Figure 11B A close-up image showing the best-fitting ellipse is displayed. More specifically, Figure 11A It shows the closing of the applied electric field (in) t Example sequence of raw epifluorescence microscopy images of fixed λ-DNA (48.5 kbp) after 0 s. Figure 11B An example original image of a best-fitting ellipse with superposition is shown for quantifying DNA length and width. Figure 11C and 11D Showing relaxations for 48.5 and 20 kbp DNA molecules, the length of the major axis of the ellipse. L A graph showing the statistics relative to the time since the field closed. The black curve shows... L instantaneous mean 。 The dark gray shaded curve and the light gray shaded curve show the distribution with one standard deviation and two standard deviations, respectively.

[0090] Figure 11C and 11D The statistical moments of the characteristic lengths of fixed DNA molecules when they are relaxed are shown. Figure 11C and 11D Data are shown for λ- (48.5 kbp) or 20 kbp DNA labeled with YOYO-1 dye in 1X TBE buffer with 2% v / v β-mercaptoethanol and 2% w / w PVP at MW 1300 kDa. The initial average length of L was lower than the expected full profile length because most of the DNA was anchored at the vertices far from the molecular ends. The vertex-fixed 48.5 kbp DNA molecule took approximately 8 seconds to fully relax, while the vertex-fixed 20 kbp DNA took approximately 3 seconds. Figure 21 The image includes epifluorescence images of 20 kbp DNA relaxed with vertices fixed. Figure 21 This illustrates the relaxation dynamics of a vertex-fixed, single-molecule DNA. The relaxation dynamics occur precisely at the close of the applied electric field (in...). t Example sequence of raw epifluorescence microscopy images of fixed 20 kbp DNA after 0 s.

[0091] Figures 22A to 22J An overview of automated image processing for quantifying 48.5 kbp DNA relaxation is shown. Figure 22A An example image of DNA with pre-relaxed vertex fixation is shown. Figure 22B This shows the results after adaptive thresholding and α-shape calculation. Figure 22A The binarized image. Figure 22C The image in the image will be obtained by Figure 22A and Figure 22B It is obtained by integrating the mask in the middle. Figure 22D It shows Figure 22A A magnified image. Figure 22E Example images are shown to illustrate the automated detection of each island structure. Elliptical curves represent the results of elliptical shape fitting. The black and light gray lines within the ellipses represent the principal and secondary axes of each fitted ellipse shape, respectively. Figures 22F to 22J The display shows the relationship with the data from... Figures 22A to 22E Images from the same procedure, except that the images are selected during the DNA relaxation process.

[0092] Figures 23A to 23J An overview of automated image processing for quantifying 20 kbp DNA relaxation is shown. Figure 23A An example image of DNA with pre-relaxed vertex fixation is shown. Figure 23B This shows the results after adaptive thresholding and α-shape calculation. Figure 23A The binarized image. Figure 23C The image in the image will be obtained by Figure 23A Images and Figure 23B It is obtained by integrating the mask in the middle. Figure 23D It shows Figure 23A A magnified image. Figure 23E Example images are shown to illustrate the automated detection of each island structure. Elliptic curves show the results of elliptical shape fitting. The black and light gray lines represent the principal and secondary axes, respectively, for each fitted ellipse shape. Figures 23F to 23J The display shows the relationship with the data from... Figures 23A to 23E Images from the same procedure, except that the images are selected during the DNA relaxation process.

[0093] Figures 24A to 24C Colocalization analysis of image intensity for vertex-fixed DNA across three different experimental runs is shown. Colocalization data for three pairs of DNA captures from independent experimental runs are presented. Experimental runs 1, 2, and 3 are respectively from the text. Figure 13A , 13B DNA data fixed at the 13C vertex. Figures 24A to 24C A colocalization scatter plot (correlation plot) showing the comparison of each pair of experimental runs is shown. As indicated by the intensity bar on the right, the heatmap is based on the count of the raw intensity values ​​of the colocalized pixels from the two images from the two experimental runs. The Pearson correlation coefficients between each pair are calculated, and the values ​​are 0.30, 0.25, and 0.22, respectively.

[0094] Figure 25A C shows a colocalization analysis of the image intensity of DNA across three consecutive images from a single experimental run, with the vertices fixed. Figure 25A Three raw, consecutive epifluorescence images from a single run of the DNA vertex fixation experiment are shown. The first and second sub-images are spaced 5 seconds apart. The second and third sub-images are spaced 2 seconds apart. Figure 25B and 25C A scatter plot (correlation diagram) comparing the colocalization of each consecutive image pair is shown. The Pearson correlation coefficients calculated for the two time-series image pairs are 0.81 and 0.86, respectively.

[0095] As described above, after the (high) electric field is removed, the DNA uniformly aligns and relaxes around the fixed vertices. This further supports the hypothesis of single-point entanglement between DNA and PVP. The persistence of such fixed points (around 30 seconds) suggests that the observed DNA trapping phenomenon is not due to electroadsorption forces such as dielectrophoresis, but may be manifested by wall defects (e.g., roughness elements). Such electroadsorbed DNA molecules are expected to be rapidly "released" after the field is turned off. In this instance, the DNA was not trapped at the convex corners of the serrated features in the channel.

[0096] Example 4: Testing DNA capture, relaxation, and unwinding cycles Figure 12 The diagram shows a representative cycle of DNA vertex fixation under a high electric field; relaxation after the electric field is turned off; unwinding after approximately 36 seconds; and then a state of free diffusion and electromigration. At t = 0 s, the DNA molecule initially forms a three-dimensional Brownian coil. At t = 2.4 s, an electric field of 190 V / cm is activated to initiate trapping. At t = 33.8 s, the electric field is turned off and the DNA molecule rapidly relaxes toward its vertex / fixation point (see Figure 11). After more than approximately 50 seconds, the relaxed DNA becomes unfixed from the wall and again freely diffuses and electromigrates. At t = 88.6 s, a low axial electric field of 19 V / cm is applied to clear the field of view (FOV) of the previously fixed DNA. This process can be repeated in multiple subsequent cycles with an estimated period of approximately 60 seconds. Figure 12 This shows a sequential (top-down) image of a single DNA molecule. The DNA molecule initially appears in the expected random coiled shape. t = 2.4 s, apply E = 190 V / cm fixed at the initial vertex. t = Closed at 33.8 s E DNA molecules are rapidly released or slowly relax towards the apex. After approximately 50 seconds, more than 95% of the DNA becomes unfixed from the wall. t At 88.6 s, a relatively low axial electric field of 11 V / cm was applied to cause the previously vertex-fixed DNA molecules to electromigrate away. Note the displacement of the relevant patterns across the last three images.

[0097] Figure 12 The data further support the entanglement between DNA and PVP. After the electric field is removed, the space between DNA and PVP may become rehydrated, and the associated low-friction configuration and Brownian motion facilitate disentanglement after tens of seconds. Pulses of pressure-driven flow may induce disentanglement much faster, down to the order of seconds.

[0098] The study also investigated whether DNA fixation sites were spatially correlated across experimental runs. Figures 13A to 13G The image shows that the vertex of a single DNA molecule is fixed to the wall, and its location varies randomly between different experimental runs. Figures 13A-13C Three representative experimental runs are shown. The first is displayed in inverted grayscale, while the other two are original grayscale images. Between experimental runs, the DNA was allowed to fully relax and untangle. Figure 13D The image shows a superposition of the images from Experiment Run 1 and Experiment Run 2; and Figure 13E The image shows an overlay of the images from Experiment 1 and Experiment 3. Figure 13D and 13E The magnified image is displayed in Figure 13F and 13G middle. Figures 13A to 13G All data shown were obtained from λ-DNA labeled with 4 pM YOYO-1 containing 2% w / w 1300 kDa PVP. Figures 13A to 13C Images of the DNA capture in three separate experimental runs are shown using inverted grayscale (Experimental Run 1), grayscale (Experimental Run 2), and grayscale (Experimental Run 3) schemes. DNA was captured for 15 seconds at 300 V / cm. The DNA was allowed to relax and untangle between experiments (see reference). Figure 4 Furthermore, a low electric field is used to confirm untangling and remove previously captured DNA. Figure 13C and 13D The superimposed images of experimental runs 1, 2, and 3 are shown, illustrating the weak spatial correlation of vertex capture points across two and three experimental runs, respectively. Figure 13F and Figure 13G They are Figure 13D and Figure 13E A magnified image.

[0099] We observed a weak correlation in DNA capture localization across all experimental runs studied. This result further supports the hypothesis that the observed capture is not due to specific features of the channel geometry, such as roughness elements or nanoscale surface pits. The data support the assumption that DNA molecules become entangled with adsorbed PVP molecules. This entanglement is ultimately reversible, and new DNA molecules become anchored at new, unrelated localizations. The approximately linear increase in capture rate shown in Figure 10 suggests that the DNA capture sites have not saturated for the duration of the experiments.

[0100] Materials and methods used in Examples 1 to 4: Customized microfluidic interface devices Figures 14A to 14D A custom microfluidic interface device according to one embodiment is shown. Figure 14A The components of the PDMS reservoir, 170 μm thick borosilicate glass, and silicon substrate chip are shown. Figure 14B The assembled custom microfluidic device is shown. Borosilicate glass is anoly bonded to a silicon substrate chip, and then plasma-bonded to the borosilicate glass in a PDMS reservoir. Figure 14C The assembled custom microfluidic device is shown attached to a glass slide with tape and then visualized via a 60X water immersion objective with a numerical aperture of 1.2. The illumination source is a blue LED. Platinum electrodes are inserted into a reservoir filled with liquid. Figure 14D The layout of the entire silicon substrate chip is shown, along with magnified images of two regions. The central region exhibits a serrated feature near one side. A large microchannel region leading to the reservoir features a pillar array with a diameter of 10 μm. The pillar array is designed to support the borosilicate glass for anodic bonding. More specifically, Figure 14A The diagram shows three main components of a microfluidic device according to one or more embodiments: four polydimethylsiloxane (PDMS) reservoirs, a borosilicate glass plate (channel cap), and a silicon substrate chip. Figure 14B An isometric view of the assembly after bonding and assembly is shown. Figure 14C Images of the experiment show the chip, platinum electrodes (connected to alligator clips) in the reservoir, and the water immersion objective. Figure 14D The channel layout in a silicon-based chip is shown. The chip has two separate (independent) single-input, single-output channels. The channels are fabricated such that they can be simultaneously visualized in the same field of view near the geometric center of the silicon chip. Fabrication was performed on a 4-inch n-type silicon wafer (4 Ohm-cm resistivity). Following standard photolithography, the microfluidic channels were dry etched to a depth of 1 μm within the silicon wafer using HBr and BCl3 (the actual depth obtained by profilometry was 0.9 μm). Residual photoresist was removed using oxygen plasma, followed by a 20-minute piranha clean at 120°C. A 300 nm thick thermal oxide layer was then grown on the wafer by Rogue Valley Microdevices. The wafer was diced using a wafer saw into nine devices, each with an in-plane dimension of 22 x 24 mm. These devices then underwent another round of piranha cleaning. The cover glass was made of 3.3mm borosilicate glass (No. 1 thickness, purchased from Kemtech America Inc.). Hole drilling was performed using a 1.1mm diameter, three-wave diamond drill bit (purchased from Arrowhead Lapidary & Supple) and a hand drill press. The drilled cover glass was then anodized. Following anodizing, a PDMS reservoir with a 5.0mm outer diameter (OD) and a 1.5mm inner diameter (ID) was cut using a biopsy punch and bonded to the glass using air plasma. The drilled holes in the glass were aligned with the cut holes on the PDMS to establish fluid connectivity.

[0101] Sample preparation λ-DNA samples were purchased from ThermoScientific (Waltham, MA) 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. 10X TBE buffer was purchased from Invitrogen and consisted 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. β-mercaptoethanol (MW = 78.13, >98.0% purity) was purchased from TCI America. The 10X TBE buffer was initially diluted to 1X TBE buffer with biological grade water (Fisher Bioreagents, Pittsburgh, PA). Next, β-mercaptoethanol was added to the 1X TBE buffer as an oxygen scavenger to achieve a concentration of 2% v / v. PVP was then added to the buffer at a concentration of 2% w / w according to its respective MW. The final buffer consisted of 1X TBE, 2% v / v β-mercaptoethanol, and 2% w / w PVP with MWs of 10, 58, 360, or 1300 kDa. The conductivity of the loading buffer was measured to be 1487 μS / cm using a conductivity probe (Advanced Electrochemistry Meter from ThermoScientific Inc.). All previous steps were performed at room temperature (20 ± 1 °C). Next, the raw DNA sample was diluted in the loading buffer to a final concentration of 4 pM, and YOYO-1 dye was added to achieve a 5:1 base pair / dye stoichiometric ratio. The samples were then incubated at 40 °C for 1 hour before use. The samples were then individually incubated at 40 °C for 1 hour before use to promote dye intercalation.

[0102] Measuring device Figure 14C Example images of an experiment using the assembled microfluidic device are shown. Visualization was performed using a standard OLYMPUS BX60 upright epifluorescence microscope. We used a 60X water immersion objective (OLYMPUS UPlanApo) with a numerical aperture of 1.2. The illumination source was a high-power blue LED (SOLIS-470C) from ThorLabs, controlled by a ThorLabs DC200 controller, and the epifluorescence filter set had the following excitation / dichroic / emission components: EX460-490, DM505, EM510IF (OLMPUS U-MWIB2). Two cameras were used for image acquisition. The first camera was a scientific-grade complementary metal-oxide-semiconductor (sCMOS) camera (ORCA-Flash 4.0LT) manufactured by Hamamatsu, controlled using HCImageLive software. The sCMOS camera was used to acquire all data in the main manuscript and most of the data in the supplementary material (SM). The second camera was an Andor Electron Multiplier Charge-Coupled Device (EMCCD) camera (iXon Ultra 897), which was controlled using Andor SOLIS software. The EMCCD camera was used to acquire image data for Figure S3 (images within all glass channels of the wet etching process described below).

[0103] A voltage was applied using a platinum electrode inserted into the reservoir at the end of the insertion channel. The electrode was a platinum wire (0.368 mm in diameter, hard, 99.95% metal content), purchased from ThermoScientific. The voltage was obtained and the current was measured using a Keithley 2400 SourceMeter. The SourceMeter was triggered and controlled via an RS232 interface through a personal computer (PC), using a custom script we wrote in MATLAB (MATLAB 2023b, Mathworks Inc., Natick, MA, USA).

[0104] Image processing for quantifying the number of DNA molecules with fixed vertices. All data for the image processing section below were acquired using a Hamamatsu Orca-Flash 4.0LT sCMOS camera at 20 frames per second (fps) with an exposure time of 0.49 seconds. This camera has a sensor pixel size of 6.5 x 6.5 μm. Since we used a 60X objective lens and no downsizing lenses, the pixel size of the imaging plane is approximately 0.11 x 0.11 μm / pixel. The raw TIFF sequences from each experimental run were imported into MATLAB for image analysis. Each 30-second sequence contained 600 images. First, a processed sequence was obtained through a time-shifted median operation. This median operation replaces each pixel value in each image with the median of the same pixel in a small number of previous and subsequent images. This operation has the effect of enhancing and preserving near-static image data (fixed, stretched DNA) and excluding moving objects (electromigrated DNA). The fastest speed was achieved when a maximum electric field of 363 V / cm was applied. Therefore, each median operation considers only three frames (i.e., each pixel value is replaced by the median of the previous, current, and next frames). For the median operation with the lowest applied electric field (where DNA moves the slowest), an increasing number of frames are used in each group. For the lowest electric field of 19 V / cm, a maximum of 21 frames are considered. Figure 10A The subplot above shows example frames obtained from a moving median image sequence. For example, captured video was used to compare the original video with the moving median video, demonstrating how image data from a stationary object (stationary DNA) can be augmented. Figure 26 Example frames from the video are shown, illustrating the difference between the original video and a moving median video, which has been processed to display only non-moving DNA molecules. The Bradley method is used for adaptive thresholding of the moving median image sequence based on local means. For this, the MATLAB command `adaptthresh` with a sensitivity parameter of 0.45 is used, followed by `imbinarize` to obtain a binarized image for each frame. Pixel values ​​in the binarized image are set to zero below the threshold and one above the threshold. The threshold pixel coordinates are then identified and stored via a MATLAB command. An α-shape operation is then performed on the binarized image to effectively exclude non-DNA image noise from the analysis. The α-shape is a generalization of the convex hull concept and a subset of Delaunay triangulation. The α-shape method is used for computational pattern recognition in computer graphics to identify the correlation of points on a graph (via proximity). To obtain the α-shape, the binarized pixel coordinates and an α radius of 2.5 are used as input to the MATLAB command `alphaShape`. The resulting α-shape is projected onto a binary mask. The binary mask is first dilated, then eroded using streamlines of the same length (10 pixels) as structuring elements (again, to support even roughly equivalent fixed DNA images). After erosion, the algorithm obtains an α-shaped boundary mask. The α-shape method successfully reduces background noise caused by adaptive thresholding. The α-shape operation, along with dilation and erosion, also smooths noisy edges of the identified objects. Figure 10A The middle sub-image shows the resulting mask instance. At each time frame, a binarized mask is used for the moving median image to obtain a series of thresholded moving median images. Figure 10A The subplot below shows an example operation after overlaying an α-shaped boundary mask with one of the median images. The (original, unchanged) intensity of the DNA detected in the resulting thresholded image is then spatially integrated to obtain a scalar signal (scalar I in Figure 11) that varies over time at a frame rate of 20 fps.

[0105] Image processing for analyzing vertex-fixed DNA relaxation This section describes how to obtain the text. Figure 11B , 11C Image processing analysis was performed on the DNA relaxation data shown in 11D. First, the TIFF sequence was temporally Gaussian filtered using the MATLAB built-in function `imgaussfilt3` with a 3-frame-wide filter. Then, an α-shaped boundary mask for the DNA relaxation video was obtained using the same method described above, but without dilation and subsequent erosion operations to preserve the DNA shape. After obtaining the boundary mask, the MATLAB command `bwconncomp` and subsequent `regionprops` were used to identify the properties of different connected regions within the α-shaped boundary mask. To prevent image noise and DNA fragment remnants in subsequent analysis, we removed connected regions with total intensities less than 100,000 and 20,000 from the 48.5 kbp and 20 kbp DNA samples, respectively. These thresholds were heuristically determined and aimed to remove objects with intensities significantly lower than those of essentially intact DNA fragments. Figures 22A to 22C and Figures 22F to 22H Examples of the above image processing steps for vertex-fixed DNA before and during relaxation are shown. Then, we again use the MATLAB command `regionprops` to fit an ellipse to each remaining region. These ellipses have the same normalized second-order central moments as the regions they fit. Examples of ellipses fitting various DNA fragments are shown in... Figure 11B As can be seen, for each DNA relaxation experimental run in each frame after the electric field is turned off, a set of fitted ellipses is obtained, which ideally represent the shapes of various DNA fragments obtained at a given time for a given experimental run. Figures 22D to 22E Examples of the aforementioned island identification and principal axis estimation are shown in Figures 22I to 22J. Figure 22D and 22E An example of DNA fixation at the relaxed anterior vertex is shown, while Figure 22I and 22J An example of vertex-fixed DNA during relaxation is shown. Elliptical approximations are shown as superimposed gray ellipses, with the major axis of each ellipse represented by two mutually orthogonal line segments. From this set of fitted ellipses, we recorded two values: the mean length of the major axis of the fitted ellipse, which we used as a characteristic length of the DNA during relaxation; and the standard deviation of the major axis length regarding whether the major axis of the fitted ellipse is longer or shorter than the mean major axis length. Figure 11C and 11D The results of the analysis are shown in the figure, expressed as one and two standard deviations, respectively. The black line represents the feature length, and the dark and light gray shaded areas represent data with a long axis mean length of one or two standard deviations. The only difference between the 48.5 and 20 kbp DNA relaxation analyses is the selected threshold, which is used to remove selected regions that are considered DNA fragments or image noise.

[0106] PVP and the outline length of DNA of various molecular weights Notably, the outline lengths of unstained λ-DNA (48.5 kbp) and 20 kbp DNA were approximately 16 μm and 6.6 μm, respectively. When λ-DNA and 20 kbp DNA were stained with YOYO-1 dye, their complete outline lengths extended to approximately 21 μm and 8.7 μm, respectively. The outline lengths of PVPs with MW of 10, 58, 360, and 1300 kDa were approximately 25, 145, 900, and 3250 nm, respectively.

[0107] Spatial average intensity The symbol <⋯> is used to denote the spatial average of all pixels within a two-dimensional region. For example, to represent the spatial integral of the intensity values ​​of the original image or two-dimensional correlation. For the quantity q in each time frame, the spatial average is expressed as shown in Equation (1): Where Nx and Ny represent the number of pixels in the horizontal and vertical directions, respectively.

[0108] Estimation of axial electric field strength As mentioned above, the conductivity of the buffer solution was measured using a commercial conductivity meter. During the experiment, we measured the current using a Keithley SourceMeter. We measured the channel depth using a profilometer, and the measured depth was approximately 0.9 μm. The photolithographic mask determined the channel width, and this width could be measured from the brightfield image, such as... Figure 9C The leftmost subplot shows a value of approximately 20 μm. The axial electric field strength can be related to the current using Ohm's law, as shown in equation (2): (2) Where j is the current density and σ is the conductivity of the buffer solution. The current density j is obtained by dividing the measured current I by the estimated cross-sectional area A. Then, the electric field strength is estimated using E = j / σ.

[0109] The following applications are related to this application: U.S. Patent Application Serial Nos. _______________ (RBPA0517PUSP1 and RBPA0517PUSP2), both filed in _________, the entire contents of which are incorporated herein by reference.

[0110] While exemplary embodiments have been described above, they are not intended to describe all possible forms of this disclosure by way of these embodiments. Rather, the language used herein is descriptive and not restrictive, and it should be understood that various changes may be made without departing from the spirit and scope of this disclosure. Furthermore, features of various embodiments may be combined to form further embodiments of this disclosure.

Claims

1. A method for capturing polynucleotide chains on the surface of a channel, comprising: Provide a polymer to the channel surface; This allows the polymer to physically interact with the channel surface; Provide a sample containing polynucleotide chains to the polymer that interacts with the channel surface; as well as An electric field is applied to the polynucleotide chain to promote the physical interaction between the polynucleotide chain and the polymer, thereby capturing the polynucleotide chain on the channel surface.

2. The method of claim 1, wherein the physical interaction between the polynucleotide chain and the polymer occurs at a vertex of the polynucleotide chain.

3. The method of claim 2, wherein after an electric field is applied, the first and second ends of the polynucleotide chain extend outward from the vertex.

4. The method of claim 1, wherein the polymer is capable of regulating the electroosmotic flow in the channel.

5. The method of claim 1, wherein the polymer is a neutral and water-soluble polymer.

6. The method of claim 5, wherein the neutral and water-soluble polymer is a polyvinylpyrrolidone polymer.

7. The method of claim 6, wherein the polyvinylpyrrolidone polymer has a molecular weight greater than 100 kDa.

8. The method of claim 5, wherein the neutral and water-soluble polymer is hydroxyethyl cellulose.

9. The method of claim 5, wherein the neutral and water-soluble polymer is polyethylene glycol.

10. The method of claim 5, wherein the neutral and water-soluble polymer is polyvinyl alcohol.

11. The method of claim 1, wherein the channel has at least one dimension perpendicular to the electric field direction of less than 5 micrometers.

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

13. The method of claim 1, further comprising altering the electric field applied to the polynucleotide chain to facilitate the release of the polynucleotide chain from physical interaction with the polymer.

14. The method of claim 13, wherein changing the electric field includes changing the intensity of the electric field.

15. The method of claim 14, further comprising providing a second sample containing a polynucleotide chain to a polymer interacting with the channel surface, and altering the electric field to facilitate a physical interaction between the polynucleotide chain and the polymer to capture the polynucleotide chain on the channel surface.

16. The method of claim 1, further comprising providing a pulse of pressure-driven flow to the polynucleotide chain to facilitate the release of the polynucleotide chain from physical interaction with the polymer.

17. A system for capturing polynucleotide chains, comprising: An apparatus comprising: At least one reservoir, sized to accommodate electrodes, electrolytes, and / or samples containing the polynucleotide chains; and A fluidic chip having at least one channel, the at least one channel being in fluid communication with the at least one reservoir, such that a sample can be transported through the at least one channel; A voltage source capable of applying an electric field to the channel; and A controller, which is programmed to: The transport of the polymer to the device is guided to facilitate polymer delivery to the channel surface, enabling the polymer to physically interact with the channel surface; Instructing the transfer of the sample from the at least one storage tank to the channel; and Interacting with the voltage source to generate a voltage difference between the electrode and at least one other electrode to form an electric field and promote physical interaction between the polynucleotide chain and the polymer to capture the polynucleotide chain on the channel surface.

18. The system of claim 17, wherein the controller is further programmed to reduce the electric field strength applied to the polynucleotide chain to facilitate the release of the polynucleotide chain from physical interactions with the polymer.

19. The system of claim 18, wherein the controller is further programmed to direct the transport of the released polynucleotide chain through the channel to remove the polynucleotide chain from the channel region where the polynucleotide chain is trapped.

20. The system of claim 19, wherein the controller is further programmed to guide the transfer of a second sample containing polynucleotide chains from at least one reservoir in fluid communication with the channel, and to increase the electric field strength to capture the polynucleotide chains from the second sample.

21. The system of claim 17, wherein the controller is further programmed to provide pulses of pressure-driven flow to the device to increase the release rate of the polynucleotide chain.

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