Patent Publication Number: US-9835539-B2

Title: Biopolymer separation using nanostructured arrays

Description:
DOMESTIC PRIORITY 
     This application claims priority to U.S. Non-Provisional application Ser. No. 14/697,269, filed Apr. 27, 2015, which claims benefit to U.S. Provisional Application Ser. No. 62/084,653, filed Nov. 26, 2014, which is incorporated herein by reference in their entirety. 
    
    
     BACKGROUND 
     The present invention relates to a continuous flow size-based separation of entities, and more specifically, to separating entities using a nanopillar array structure. 
     The separation and sorting of biological entities, such as cells, proteins, deoxyribonucleic acid (DNA), ribonucleic acid (RNA), etc., is important to a vast number of biomedical applications including diagnostics, therapeutics, cell biology, and proteomics. 
     Protein and DNA/RNA separation for analytical purposes is traditionally done by gel electrophoresis, where a protein mix is subjected to a strong electric field (typically 30 volts per centimeter (V/cm)). Proteins or DNA/RNA move through the gel at a rate depending on their size and surface charge. The gels are prepared from agarose or acrylamide polymers that are known to be toxic. The outcome of the electrophoresis experiment is revealed optically from staining the proteins with dye, or staining the DNA/RNA with ethydium bromide which is extremely carcinogenic. Gels require sufficient quantities of material for the outcome of the electrophoresis to be detectable, but bad cross-linking in the gel matrix often leads to inconclusive results and the complete loss of the samples. If the gel matrix size is not adapted to the sample molecule size or if the electrophoresis is left to run for too long, the sample is also lost. 
     For separation of macromolecules, such as DNA, RNA, proteins, and their fragments, gel electrophoresis is widely employed. Gel electrophoresis currently has a market with world-wide sales greater than $1 billion dollars per year. Gel electrophoresis applied to medical diagnostic represents a multibillion dollar market. 
     In comparison with traditional techniques, silicon (Si) nanofabrication technology offers much more precise and accurate control in nano-structural dimensions and positioning of the same, and thus can lead to reliable sorting of particles based on their sizes. To date, Si-based Lab-on-a-Chip approaches using Si pillars arrays have shown promise. However, only sorting in the micron (10 6  or micrometer (μm)) range has been demonstrated using these techniques, which does not access the nanometer dimensions required for sorting DNA, proteins, etc. 
     SUMMARY 
     According to one embodiment, a method of sorting biopolymers is provided. The biopolymers are introduced into a nanopillar array. The biopolymers include a first population and a second population, and the nanopillar array includes nanopillars arranged to have a gap separating one from another. The biopolymers are sorted through the nanopillar array by transporting the first population of the biopolymers less than a predetermined bumping size according to a fluid flow direction and by transporting the second population of the biopolymers at least the predetermined bumping size according to a bumped direction different from the fluid flow direction. The nanopillar array is configured to employ the gap with a gap size less than 300 nanometers in order to sort the biopolymers. 
     According to one embodiment, a method for configuring a cascaded array structure is provided. A first stage is configured to output entities to a second stage according to a fluid flow direction and configured to bump the entities to the second stage according to a bumped direction. The second stage is configured to output the entities to a third stage according to the fluid flow direction and configured to bump the entities to the third stage according to the bumped direction. The third stage is configured to output the entities according to the fluid flow direction and configured to bump the entities according to the bumped direction. Separate outlets are provided for which to collect the entities from the third stage according to sorting through the first stage to the third stage. 
     According to one embodiment, an apparatus for sorting entities is provided. The apparatus comprises a first stage including a first nanopillar array corresponding to a first critical bumping dimension, and a second stage including a second nanopillar array and a third nanopillar array. The second nanopillar array has a second critical bumping dimension and the third nanopillar array has a third critical bumping dimension. The first nanopillar array is coupled to the second nanopillar array and configured to output the entities less than the first critical bumping dimension to the second nanopillar array. The first nanopillar array is coupled to the third nanopillar array and is configured to output the entities meeting the first critical bumping dimension to the third nanopillar array. 
     According to one embodiment, a method of separating and identifying unknown biopolymers is provided. Unknown biopolymers and biopolymer-binding proteins are introduced into a nanopillar array. When the unknown biopolymers bind with the biopolymer-binding proteins, biopolymer-protein complexes are formed. When the unknown biopolymers do not bind with the biopolymer-binding proteins, the unknown biopolymers and biopolymer-binding proteins remain unbound. The unknown biopolymers and the biopolymer-binding proteins are sorted through the nanopillar array by transporting any of the unknown biopolymers and the biopolymer-binding proteins which are unbound in a fluid flow direction, as the unknown biopolymers and the biopolymer-binding proteins which are unbound are less than a predetermined bumping size. The unknown biopolymers and the biopolymer-binding proteins are sorted through the nanopillar array by transporting the biopolymer-protein complexes in a bumped direction, as the biopolymer-protein complexes are at least the predetermined bumping size. The nanopillar array is configured to employ the gap with a gap size less than 300 nanometers for sorting. 
     According to one embodiment, a method of separating and identifying unknown proteins is provided. Unknown proteins and protein-binding biopolymers are introduced into a nanopillar array. When the unknown proteins bind with the protein-binding polymers, biopolymer-protein complexes are formed, and when the unknown proteins do not bind with the protein-binding biopolymers, the unknown proteins and biopolymers remain unbound. The unknown proteins and the protein-binding biopolymers are sorted through the nanopillar array by transporting any of the unknown proteins and the protein-binding biopolymers which are unbound in a fluid flow direction, as the unknown proteins and the protein-binding biopolymers which are unbound are less than a predetermined bumping size. The unknown proteins and the protein-binding biopolymers are sorted through the nanopillar array by transporting the biopolymer-protein complexes in a bumped direction, as the biopolymer-protein complexes are at least the predetermined bumping size. The nanopillar array is configured to employ the gap with a gap size less than 300 nanometers for sorting. 
     Additional features and advantages are realized through the techniques of the present invention. Other embodiments and aspects of the invention are described in detail herein and are considered a part of the claimed invention. For a better understanding of the invention with the advantages and the features, refer to the description and to the drawings. 
    
    
     
       BRIEF DESCRIPTION OF THE SEVERAL VIEWS OF THE DRAWINGS 
       The subject matter which is regarded as the invention is particularly pointed out and distinctly claimed in the claims at the conclusion of the specification. The forgoing and other features, and advantages of the invention are apparent from the following detailed description taken in conjunction with the accompanying drawings in which: 
         FIG. 1  is a schematic of a deterministic lateral displacement (DLD) array showing definitions of the array parameters. 
         FIG. 2A  illustrates a schematic of particle trajectories at the interface between a neutral region and a micro fluidic metamaterial element. 
         FIG. 2B  illustrates the simplest metamaterial element is an asymmetric array of posts tilted at an angle +α relative to the channel walls and bulk fluid flow. 
         FIG. 2C  illustrates a cross-sectional SEM image showing the microfabricated post array. 
         FIG. 2D  illustrates equivalent microfluidic birefringence based on particle size showing the time-trace of a 2.7-μm red fluorescent transiting the interface and being deflected from the normal. 
         FIG. 3A through 3G  illustrate schematics of a process flow for nanopillar array fabrication according to an embodiment, in which: 
         FIG. 3A  illustrates a hard mask layer disposed on a substrate; 
         FIG. 3B  illustrates disposing a resist layer on the hard mask layer; 
         FIG. 3C  illustrates patterning the resist layer; 
         FIG. 3D  illustrates patterning the hard mask layer; 
         FIG. 3E  illustrates etching the substrate into the pillar array; 
         FIG. 3F  illustrates the pillar array with the hard mask pattern removed; and 
         FIG. 3G  illustrates disposing an oxide layer on the pillar array. 
         FIGS. 4A and 4B  are scanning electron microscope images of the same wafer to illustrate the result of reactive ion etching before hard masks are removed according to an embodiment. 
         FIGS. 4C and 4D  are scanning electron microscope images of a parallel processed wafer to illustrate the result of reactive ion etching after hard masks are removed according to an embodiment. 
         FIGS. 5A and 5B  are scanning electron microscope images of another wafer to illustrate a fabricated nanopillar array without a thermal oxide according to an embodiment. 
         FIGS. 5C, 5D, and 5E  are scanning electron microscope images of a parallel processed wafer to illustrate the impact of growing a thermal oxide on nanopillar arrays according to an embodiment. 
         FIGS. 6A and 6B  are scanning electron microscope images of another wafer to illustrate starting with a smaller gap size according to an embodiment. 
         FIGS. 6C and 6D  are scanning electron microscope images of a parallel processed wafer to illustrate the oxidation process when the initial gap size is small according to an embodiment. 
         FIG. 7A  illustrates a general chemical schematic of chemical modification to a pillar array to form sorting array surfaces according to an embodiment. 
         FIG. 7B  illustrates a chemical schematic for chemical modification by applying metal to a pillar array to form sorting array surfaces according to an embodiment. 
         FIGS. 8A through 8D  are cross-sectional views illustrating chemical modification of sorting arrays as a means of modifying the gap size between pillars according to an embodiment, in which: 
         FIG. 8A  illustrates the gap size between pillars before chemical modification; 
         FIG. 8B  illustrates the reduced gap size between pillars after chemical modification; 
         FIG. 8C  illustrates an enlarged view of a reactive site in  FIG. 8A ; and 
         FIG. 8D  illustrates an enlarged view of the monolayer in  FIG. 8B . 
         FIG. 9A  is a top view illustrating particle flow in a chemically modified sorting array with particles that have no affinity for the surface monolayer compared to particles that do have affinity for the surface monolayer according to an embodiment. 
         FIG. 9B  is an enlarged view of a cross-section of the nanopillar, monolayer, and particle with affinity according to an embodiment. 
         FIG. 10A  is a cross-sectional view illustrating pillars having gap variation according to an embodiment. 
         FIG. 10B  is a cross-sectional view illustrating the oxidation process that removes the gap variation according to an embodiment. 
         FIG. 11  is a top view illustrating a chip (fluidic device) having the pillar array according to an embodiment. 
         FIG. 12  is a method of providing a fluidic apparatus (e.g., chip) according to an embodiment. 
         FIG. 13  is a method of forming a nanopillar array according to an embodiment. 
         FIG. 14  is a top view of a schematic representing an arrangement of the pillars in the nanopillar array according to an embodiment. 
         FIG. 15  is a schematic of the chip now with two inlets and with particles of different sizes traversing through the nanopillar array according to an embodiment. 
         FIG. 16A  is a scanning electron microscope image of particle trajectories for 70 nanometer diameter beads according to an embodiment. 
         FIG. 16B  is a plot of trajectory angle as a function of velocity for the 70 nanometer beads according to an embodiment. 
         FIG. 16C  is a scanning electron microscope image of particle trajectories for 50 nanometer diameter beads according to an embodiment. 
         FIG. 16D  is a plot of trajectory angle as a function of velocity for the 50 nanometer beads according to an embodiment. 
         FIG. 17  is a chart of example data according to an embodiment. 
         FIG. 18  is a method of sorting entities according to an embodiment. 
         FIG. 19  is a method of sorting entities according to an embodiment. 
         FIG. 20  is a method of sorting entities according to an embodiment. 
         FIG. 21A  is a top view of a nanopillar array sorting long biopolymers from short biopolymers according to an embodiment. 
         FIG. 21B  is an enlarged view of  FIG. 21A  according to an embodiment. 
         FIG. 22A  is a top view of a nanopillar array according to an embodiment. 
         FIG. 22B  is a scanning electron microscope image of biopolymers deflected through the nanopillar array in  FIG. 22A  according to an embodiment. 
         FIG. 22C  is a scanning electron microscope image of biopolymers further deflected through the nanopillar array in  FIG. 22A  according to an embodiment. 
         FIG. 22D  is a scanning electron microscope image of one of the biopolymers exiting the nanopillar array in  FIG. 22A  according to an embodiment. 
         FIG. 22E  is a scanning electron microscope image of one of the biopolymers still being deflected through the nanopillar array in  FIG. 22A  according to an embodiment. 
         FIGS. 23A, 23B, 23C, and 23D  are consecutive fluorescence images illustrating DNA molecules being deflected through a nanopillar array according to an embodiment. 
         FIGS. 24A, 24B, 24C, 24D, 24E, 24F, 24G, and 24H  are consecutive fluorescence images illustrating that shorter DNA molecules are not deflected while traversing through the nanopillar array according to an embodiment. 
         FIG. 25  is a schematic of a cascaded pillar array design to separate biopolymers according to an embodiment. 
         FIG. 26  is a method of sorting biopolymers according to an embodiment. 
         FIG. 27  is a method for configuring a cascaded array structure according to an embodiment. 
     
    
    
     DETAILED DESCRIPTION 
     Sorting in the micron (10 6  μm) range has been demonstrated using Si-based Lab-on-a-Chip approaches. Additional information in this regard is further discussed in a paper entitled “Hydrodynamic Metamaterials: Microfabricated Arrays To Steer, Refract, And Focus Streams Of Biomaterials” by Keith J. Morton, et al., in PNAS 2008 105 (21) 7434-7438 (published ahead of print May 21, 2008), which is herein incorporated by reference. 
     The paper “Hydrodynamic Metamaterials: Microfabricated Arrays To Steer, Refract, And Focus Streams Of Biomaterials” discusses that their understanding of optics came from viewing light as particles that moved in straight lines and refracted into media in which the speed of light was material-dependent. The paper showed that objects moving through a structured, anisotropic hydrodynamic medium in laminar, high-Peclet-number flow move along trajectories that resemble light rays in optics. One example is the periodic, micro fabricated post array known as the deterministic lateral displacement (DLD) array, a high-resolution microfluidic particle sorter. This post array is asymmetric. Each successive downstream row is shifted relative to the previous row so that the array axis forms an angle α relative to the channel walls and direction of fluid flow as shown in  FIG. 1 . During operation, particles greater than some critical size are displaced laterally at each row by a post and follow a deterministic path through the array in the so-called “bumping” mode. The trajectory of bumping particles follows the array axis angle α. Particles smaller than the critical size follow the flow streamlines, weaving through the post array in a periodic “zigzag” mode. 
       FIG. 1  is a schematic of a deterministic lateral displacement (DLD) array showing definitions of the array parameters: The posts are periodically arranged with spacing λ, and each downstream row is offset laterally from the previous row by the amount δ breaking the symmetry of the array. This array axis forms an angle α=tan −1  (δ/λ)=tan −1 (ε) with respect to the channel walls and therefore the direction of fluid flow. Because of the array asymmetry, fluid flow in the gaps between the posts G is partitioned into 1/ε slots. Each of these slots repeats every 1/ε rows so the flow through the array is on average straight. Particles transiting the gap near a post can be displaced into an adjacent streamline (from slot  1  to slot  2 ) if the particles radius is larger than the slot width in the gap. Therefore, larger particles are deterministically displaced at each post and migrate at an angle α to the flow. Smaller particles simply follow the streamline paths and flow through the array in the direction of fluid flow. 
       FIG. 2A  demonstrates size-based birefringence of particles flowing through a hydrodynamic medium of channel-spanning microfabricated posts. Two differently sized particles are normally incident on an interface between a symmetric post array (left half of channel) and an asymmetric post array (right half). Pressure-driven fluid flow through the arrays is from left to right, its overall direction determined by the larger micro fluidic channel.  FIG. 2B  illustrates a schematic of particle trajectories at the interface between a neutral region and a microfluidic metamaterial element. Particles larger than a critical size follow the array asymmetry, whereas smaller particle follow the fluid flow.  FIG. 2B  illustrates the simplest metamaterial element is an asymmetric array of posts tilted at an angle +α relative to the channel walls and bulk fluid flow. Shown is a top-view scanning electron micrograph (SEM) of the interface between a neutral array (α=0°) and an array with array angle α=11.3° (the gap G=4 μm and post pitch λ=11 μm are the same for both sides).  FIG. 2C  illustrates a cross-sectional SEM image showing the microfabricated post array.  FIG. 2D  illustrates equivalent microfluidic birefringence based on particle size showing the time-trace of a 2.7-μm red fluorescent transiting the interface and being deflected from the normal. Smaller 1.1-μm green beads are not deflected at the interface. 
     Array elements can be tailored to direct specific particle sizes at an angle to the flow by building arrays with design parameters shown in  FIG. 1 , which include obstacle size D, spacing between the posts G, and post pitch λ. Asymmetry is determined by the magnitude of the row-to-row shift δ and is characterized by the slope ε=δ/λ. The final array angle is then α=tan −1 (ε). For a given array angle, the critical particle size for the bumping mode is determined by the ratio between the particle diameter and the post spacing or gap. This critical particle size has been previously delineated for a range of array angles between 1.0° and 16°. For a given gap size, the critical size of bumping is larger at steeper angles. By using these design criteria, streams of beads, cells, and DNA have all been moved deterministically for size-based separation applications. For the example given in  FIG. 1 , which has an array angle of 11.3°, gap G=4 μm, and post pitch λ=11 μm, the threshold particle size is ≈2.4 μm. Therefore, 2.7-μm red beads travel along the array axis angle in the bumping mode, and the 1.0-μm green beads travel along streamlines in the zigzag mode, as shown. The array elements and any ancillary microfluidic channels and reservoirs are fabricated in silicon wafers by using standard microfabrication techniques including photolithography and etching. Arrays can also be molded in PDMS by using a similarly crafted silicon master. For the silicon etch, an optimized deep reactive ion etch (DRIE) is used to maintain smooth, vertical side walls, ensuring uniform top-to-bottom spacing between posts as shown in  FIG. 2C . 
     Unlike the state-of-the-art, embodiments are designed to create manufacturable silicon pillar arrays with uniform gaps between the pillars (also referred to as posts) with dimensions in the sub-100 nanometer (nm) regime. These pillar arrays can be used, for example, in a bumper array configuration as described above for the sorting and separation of biological entities at these dimensions, such as DNA, RNA, exosomes, individual proteins, and protein complexes. Particularly, the pillar arrays are designed with an oxide coating, such as a SiO 2  coating which can be used to “heal” variation in the gap size along the entire axis of the pillars. Uniform gap sizes are utilized to obtain efficient sorting, e.g., to sort a 20 nm particle from a 10 nm particle. This is particularly challenging for gaps in the sub-100 nm regime where there is inherent variation in gap size greater than the dimensions of the particles to be sorted, which is limited by the reactive-ion etch (RIE) process at this scale. Demonstrated sorting pillar gaps found in the state-of-the-art have dimensions in the micron range, and therefore, the state-of-the-art cannot sort close to this fine of a scale disclosed in embodiments. Even for a pillar array with a very small angle pitch (also referred to as array angle and critical angle), e.g. 5.7 degrees, where sorting efficiency is highest, only a particle greater that 12% of the gap will sort. Therefore, consistent gaps in the nanometer regime are required to sort, for example, a protein aggregate. Sorting of individual proteins (e.g., size range of 1-10 nm) is traditionally performed using ion exchange chromatography or gel electrophoresis, which are load-and-sort techniques rather than a continuous flow Si-based solution. However, the state-of-the-art technique has no existing solution for sorting entities in 10-100 nm scale, but the embodiments provide a solution in both of these ranges (e.g., the 1-10 nm range and the 10-100 nm range). Embodiments also include chemical modification of the pillars via attachment and/or grafting of molecules to further decrease a given gap to a tailored size. 
     For ease of understanding, sub-headings may be utilized at times. It should be noted that the sub-headings are for explanation purposes only and not limitation. 
     Pillar Array Fabrication 
       FIGS. 3A through 3G  illustrate schematics of a process flow for nanopillar array fabrication according to an embodiment. In  FIG. 3A , process flow  301  illustrates a substrate  302 . A hard mask  304  is disposed on top of the substrate  302 . The substrate  302  may be a wafer, such as, e.g., a silicon (Si) wafer. The oxide hard mask  304  may be silicon dioxide (SiO 2 ) that is used for etching. Although oxide is one example, nitride or another hard material may be utilized. The oxide hard mask  304  may be disposed by deposition and/or growth on bulk silicon (substrate  302 ). The thickness of the oxide hard mask  304  may range from tens to several hundred nanometers, depending on the etch depth needed to create the height of the pillars and the selectivity of the RIE chemistry for the substrate  302  versus the hard mask material  304 . Other materials may be utilized for the substrate  302  and the hard mask layer  304 . 
     In  FIG. 3B , process flow  303  illustrates disposing a resist  306  on top of the oxide hard mask  304 . The resist  306  may be a positive resist or a negative resist. The thickness of the resist  306  may range from 100 nm-1 μm, depending on the resist  306 , hard mark ( 304 ) etch selectivity, the thickness of the hard mask  304 , and nanopillar gap resolution needed. For narrow sub-100 nm gaps and shallow pillar depths, a resist thickness range of 100-500 nm is utilized to achieve higher resolution features with less variability in gap size. The resist  306  may also be a multi-layer resist stack comprised of two or more layers each with different etch selectivity to improve resolution. 
     In  FIG. 3C , process flow  305  illustrates patterning the resist  306  into a resist pattern  308 . The resist pattern  308  may be defined but is not limited to using electron-beam lithography, nanoimprint lithography, interference lithography, extreme ultraviolet lithography, and/or deep ultraviolet lithography or a combination of these techniques. The resist pattern  308  is formed into resist pillars in the pattern of the future nanopillar array. In one case, the resist pattern  308  may include multiple patterns for different nanopillar arrays. 
     Process flow  307  illustrates pattern transfer from the resist pattern  308  to the oxide hard mask  304  to result in the etched hard mask pattern  312  in  FIG. 3D . The pattern transfer to the hard mask  304  may be performed using reactive ion etching (RIE). Process flow  307  shows the resist pattern  308  on top of the corresponding etched hard mask pattern  312 . 
     In  FIG. 3E , process flow  309  illustrates patterning the nanopillars  314  to be defined in the substrate  302  underneath the etched hard mask pattern  312 . The nanopillars  314  may be etched using reactive ion etching. The resist pattern  308  may be removed from on top of the etched hard mask pattern  312  before patterning the nanopillars  314  in the substrate  302  or after patterning the nanopillars  314 . Removing the resist pattern  308  after etching the nanopillars  314  may be performed as it can serve to avoid hard mask pattern  312  erosion that can occur during the nanopillar  314  RIE process. Hard mask erosion, in turn, may lead to pillars with a tampered (undesired) sidewall angle. 
     Process flow  311  illustrates removal of the hard mask pattern  312  in  FIG. 3F . The hard mask pattern  312  may be removed in dilute hydrofluoric (DHF) acid, if the hard mask material is SiO 2 . Process flow  311  shows a nanopillar array  320  of nanopillars  314 . 
     To further reduce the size of gaps between each of the nanopillars  314  and to reduce gap variation, process flow  313  illustrates disposing oxide  316  to cover the surface of the nanopillar array  320  formed in the substrate  302  in  FIG. 3G . In one case, thermal oxidation may be utilized to grow silicon dioxide  316  to cover of the surface of the nanopillar array  320  in order to narrow the gaps. In another case, the oxide  316  may be deposited on the nanopillar array  320  (made of silicon), for example using atomic layer deposition. 
     In general, pillar arrays include a dense array of silicon pillars defined by RIE followed by an oxidation operation (e.g., process flow  313 ) that serves to narrow the gaps between the pillar posts and minimize gap variation. Nanopillar array fabrication may also include an optional chemical modification operation where further gap scaling (i.e., reduction in size) may be required. These pillar and/or gap arrays can be implemented into angled pillar designs to concentrate a sample or separate a heterogeneous mixture of biological entities at the single molecule level, similar to work demonstrated by the paper “Hydrodynamic Metamaterials: Microfabricated Arrays To Steer, Refract, And Focus Streams Of Biomaterials” for cell or large particle sorting. The process flow for nanopillar array fabrication in  FIGS. 3A and 3B  can be utilized to create arrays of nanopillars  314  shifted in any desired gap G spacing between the nanopillars  314 , desired pillar pitch λ, desired row-to-row shift δ, and desired array angle α (also referred to as the critical angle α) (as shown in  FIG. 1 ). 
     Multiple nanopillar arrays  320  (e.g., 1-N, where N is the last number of nanopillar arrays  320 ) may be fabricated as discussed in  FIGS. 3A and 3B  on the same substrate  302 . The first nanopillar arrays  320  may have a first set of parameters (desired gap G spacing between the nanopillars  314 , desired pillar pitch λ, desired row-to-row shift δ, and desired array angle α). The second nanopillar arrays  320  may have a second set of parameters (desired gap G spacing between the nanopillars  314 , desired pillar pitch λ, desired row-to-row shift δ, and desired array angle α), where one or more of the first set of parameters can be different from the second set of parameters. The third nanopillar arrays  320  may have a third set of parameters (desired gap G spacing between the nanopillars  314 , desired pillar pitch λ, desired row-to-row shift δ, and desired array angle α), where one or more of the first set of parameters can be different from and/or the same as some of the second set of parameters, and one or more of the third set of parameters can be different from and/or the same as some of the first and second set of parameters. This same analogy can apply through the last (N) nanopillar arrays  320  which may have a last (N) set of parameters (desired gap G spacing between the nanopillars  314 , desired pillar pitch λ, desired row-to-row shift δ, and desired array angle α), where one or more of the last set of parameters can be different from and/or the same as any one of first, second, third, and N−1 set of parameters. 
     To define the pillars and gaps, a negative-tone nanoscale lithography technique may be better to ensure a patterned gap size less than (&lt;) 100 nm to begin with, e.g., the pillars and gaps are defined in the resist pattern  308  shown in process flow  305 . Electron-beam lithography is one option where pillars array patterns are smaller. However, the more manufacturable approach of nanoimprint lithography can also be applied as well as extreme ultraviolet (EUV) and deep ultraviolet (DUV) lithography under well controlled dose conditions. To achieve a high aspect ratio pillars, the written pattern (i.e., resist pattern  308 ) must be transferred to the hard mask  304  (hard mask pattern  312 ) before etching the (Si) substrate  302 . High aspect ratio pillars permit larger fluidic throughput and can reduce clogging issues associated with micro/nanofluidic features. High aspect ratio pillars are therefore a useful feature to have so long as the gap size can be maintained between adjacent pillars. By defining the pillars in the resist pattern  308  and transferring them to the etched hard mask pattern  312  first, the benefit of etch selectivity increases the aspect ratio while maintaining a more consistent gap size when the pillar array ( 320 ) etch is performed. 
     Some experimental data is discussed below as example implementations. The experimental data is for explanation and not limitation. In this case, electron-beam lithography was utilized to define the pillar dimensions (e.g., resist pattern  308 ) in hydrogen silsesquioxane (HSQ) as part of a double layer resist stack (e.g., resist  306 ), which is then transferred to a 150 nm undensified low-temperature oxide (LTO) hard mask (e.g., etched hard mask pattern  312 ). Densified LTO, thermal oxide and/or SiO 2 /SiN/SiO 2  hard masks may also be considered. The experiment then used a RIE-based Si etch process to define the pillars (e.g., pillars  314 ) in the substrate. Further details of the RIE process are now described. 
     RIE Process Details: Dry etching was carried out in an Applied Materials DPSII ICP etch chamber for pattern transfer to fabricate 400 nm high Si pillars from the e-beam resist pattern. First, the developed negative tone e-beam resist (HSQ) is used to etch through an organic planarization layer (OPL) mask using a N 2 /O 2 /Ar/C 2 H 4  chemistry at 400 watts (W) source power, 100 W bias power, and 4 millitorr (mTorr) pressure at 65° C. Then, the pattern is transferred further into a SiO 2  hard mask using CF 4 /CHF 3  chemistry at 500 W source power, 100 W bias power, and 30 mTorr pressure at 65° C. The carbon hard mask is then stripped using O 2 /N 2  chemistry in an Applied Materials Axiom downstream asher at 250° C. Using the SiO 2  hard mask, Si pillars are etched to 400 nm depth using the DPS II by first a CF 4 /C 2 H 4  breakthrough step and then Cl 2 /HBr/CF 4 /He/O 2 /C 2 H 4  main etch at 650 W source power, 85 W bias power and 4 mTorr pressure at 65° C. It is noted that three masks were utilized to eventually etch the pillars, and the three masks were the developed HSQ e-beam resist (mask), the OPL mask, and the SiO 2  hard mask. 
     Gap Analysis 
       FIGS. 4A, 4B, 4C, and 4D  are scanning electron microscope images of the result of this RIE process for two separate instances.  FIGS. 4A and 4B  illustrate the pillars (e.g., pillars  314 ) before the hard mask (e.g., hard mask pattern  312 ) is removed (such as in process flow  309 ), and the tops of the pillars (pillars  314  with hard mask pattern  312  on top) have a rounded shape. The 150 nm LTO (undensified) hard mask was utilized together with a RIE etch to produce the pillars  314  in  FIGS. 4A and 4B .  FIGS. 4C and 4D  illustrate pillars (e.g., pillars  314 ) after hard mask (e.g., hard mask pattern  312 ) removal by dilute hydrofluoric acid carried out on different wafers, and the tops of the pillars  314  are flat in  FIGS. 4C and 4D . In both cases, the Si pillars bow inward at the center due to the high density of the pillars in the array. That is, the gaps between pillars widen at the center of the pillars  314  because the diameter of the pillars are reduced at the center. The pillars have an inward-bowed shape or an hour glass shape. It is noted that pillars at the boundaries of the array are very vertical (not shown). This highlights the problem of gap non-uniformity at the nanometer scale where approximately (˜) 100 nm gap sizes have approximately 50 nm of gap variation from the top of a pillar to bottom (i.e., depth or height) of the same pillar as seen in  FIGS. 4C and 4D . The close proximity of pillars in the array as defined by the gap caused the pillars to bow inward at the center, producing gap variation that inhibits further scaling. This effect has been observed on gap sizes with dimensions of 250 nm and below for the etch process described above (i.e., prior to disposing the oxide layer  316 ). 
     According to an embodiment,  FIGS. 5A and 5B  are scanning electron microscope images of the fabricated nanopillar array of wafer  5  without a 50 nm thick thermal oxide.  FIGS. 5C, 5D, and 5E  are scanning electron microscope images of wafer  7  showing the impact of growing a 50 nm thick thermal oxide (e.g., oxide layer  316 ) on nanopillar arrays embedded in Si according to an embodiment. On the side of the pillars, there is a right wall  505  (shown in  FIGS. 5A and 5C ), a bottom  510  of the substrate, and a left wall  515  (shown in  FIG. 5B ). 
     The processing of pillars in  FIGS. 5A and 5B  for wafer  5  are identical to the processing of pillars on wafer  7  in  FIGS. 5C, 5D, and 5E  except for the final oxidation step (only performed on wafer  7  in  FIGS. 5C-5E ). In the case of  FIG. 5B  (wafer  5 ), there is 26 nm of variation for the gap size of approximately 186 nm while  FIG. 5D  (wafer  7 ) shows only a 13 nm variation in gap size after oxidation with the gap size narrowing to approximately 138 nm in this case. This healing effect of oxidation occurs as a result of oxide non-uniformity on these non-planar structures (i.e., pillars) as shown in  FIG. 5E .  FIG. 5E  shows that relative to two pillars (from a side-by-side perspective in the x-axis), the gap size between those two pillars can only vary by 13 nm from top to bottom (i.e., along the vertical axis of the y-axis) because the oxide has filled in the inward-bowed shape. Using the etch process applied to  FIGS. 5A and 5B  (wafer  5 ), uneven oxidation on pillar features is found to “heal” gap variation as shown in  FIGS. 5C, 5D, 5E  (wafer  7 ) as the oxidation proceeds more rapidly at the center of the pillars (instead of at the top and bottom), and this is shown further in  FIGS. 10A and 10B . 
       FIGS. 6A and 6B  (wafer  5 ) illustrate starting with a smaller gap size such as 80-89 nm (varies by 9 nm) in which no oxide is disposed to fill in the hour glass shape.  FIGS. 6C and 6D  (wafer  7 ) illustrate the 50 nm oxidation step applied when the original gap size is 80-89 nm (varies by 9 nm). The impact of oxidation is very apparent in FIGS.  6 C and  6 D where the same 50 nm oxidation step (discussed above in  FIGS. 5C, 5D, 5E ) reduces the gap size from 80-89 nm to just 21-25 nm (gap variation 4 nm) with a 12:1 (depth:gap) ratio. As seen in  FIGS. 6C and 6D , oxidation on smaller starting gap sizes (e.g., such as 80-89 nm (or smaller) before the oxidation step to narrow the gap and remove the inward-bow) yields approximately 25 nm gaps with only a few nanometers variation (4 nm) over approximately a 300 nm etch depth, where the depth to gap ratio of 300:25 results in the 12:1 ratio. This small amount of gap variation (e.g., 4 nm) and process opens up the opportunity to make custom, tunable gap sizes, particularly when these nanopillars are combined with chemical modification processes. The term high aspect ratio can pertain to structures with a depth to gap ratio of greater than 4:1, which can be difficult to achieve at this scale in a manufacturable process. 
     By disposing the oxide on the pillar array as discussed herein, embodiments are configured to provide a pillar array with a gap size that is uniform along the vertical axis (i.e., the depth) of two pillars that are side-by-side (e.g., the gap size between the two side-by-side pillars varies less than 5 nm (such as by 4 nm, 3 nm, 2 nm)). For example,  FIGS. 10A and 10B  are cross-sectional views illustrating the healing process that removes (reduces) the gap variation and creates a uniform gap size in the pillar array  320  according to an embodiment. For illustration purposes only, two pillars  314  are shown side-by-side but the illustration applies to all of the pillars  314  in the pillar array  320 . The height of the pillars  314  is shown on the y-axis, and the width/diameter is shown on the x-axis. The z-axis represent to length of the array  320 , and additional pillars  314  (not shown) in the array are positioned in front of and behind the two pillars  314 .  FIG. 10A  shows two example pillars  314  made out of their substrate material (substrate  302 ). The pillars  314  are bowed inward to have an hour glass shape. In  FIG. 10A , two gap sizes G 1  and G 2  are shown but there may be additional gap sizes between gap sizes G 1  and G 2 . The gap size G 1  is at (near) the top and bottom of the pillars  314 . The gap size G 2  is at (near) the center of the pillars  314 . The close proximity of pillars  314  as defined by the gap size G 1  in the array  320  may cause the hour glass shape because of the dimensional constraints of the gap size G 1  imposed on the impinging flux of reactive ions during the RIE process. 
       FIG. 10B  shows the two example pillars  314  after disposing the oxide layer  316 . Because of the non-planar architecture, nanosize of the pillars, and the tight nano-spacing between the pillars  314  in the pillar array  320 , the oxide layer  316  does not distribute evenly on the pillars  314 . Instead, more oxide  316  is formed more rapidly in the center (cavities) of the pillars  314  than at the top and bottom of the pillars  314  in the y-axis. In other words, the bowed-in centers are filled in at a faster rate than the tops and bottoms of the pillars  314 . This uneven distribution of the oxide  316  formed on the pillars  314  serves to straighten the individual pillars  314  changing them from the hour glass shape to a cylinder-like shape, which in turn makes the gap size G 5  uniform between the two pillars  314  (and any other two pillars  314  side-by-side in the x-axis). Accordingly, all of the gaps G (representing the general gap size of the array) are uniform throughout the pillar array  320 . 
     Chemical Modification 
     Interaction between the particles to be sorted and the surfaces of the array can be tailored by using chemical modification. In general, this can involve the attachment and/or grafting of molecules to the surfaces of the pillar array, through physical adsorption and/or formation of chemical bonds. Also, the chemical modification of the pillar array can include application of a layer(s) of material such as a metal, polymer, and/or ceramic coating, as well as changes to the oxidation state of the array surface. Surfaces (for chemical modification) can include the areas of the sorting pillars, the walls, the ceiling, and/or the floors of the fluidic pillar array. Additionally, chemical modification can be on any surfaces present in the inlets, outlets, drive mechanisms, and/or other fluidic channels attached to the nanofluidic device (e.g., one or more pillar arrays). 
     Although the chemical modification can be applied as discussed above, the better application is the chemical modification of the sorting pillars themselves, as this allows design of the interactions between the particles with the sorting array surfaces. 
     In one example, a small organic molecule or polymer, termed a ligand, can be chemically grafted to the surface of the pillars, such as through condensation of chlorosilane and/or alkoxysilanes on the pillars&#39; native silicon oxide as illustrated in  FIG. 7A . Also, the ligand can be chemically grafted to the surface of the pillars, such as through thiols, amines, and/or phosphines on pillars coated with a thin layer (e.g., 10 nm) of gold or silver as illustrated in  FIG. 7B . The resulting layer of ligand molecules is a single molecule thick, i.e., a monolayer. The terminal groups of the monolayer, which are in direct contact with the fluid and particles, determine the physochemical interactions felt by the particles as they pass through the array. Changing the terminal group of the ligand therefore allows tailoring of the surface interactions within the array. 
       FIGS. 7A and 7B  illustrate the general chemical schematic of methods of chemical modification of sorting array surfaces according to an embodiment. Referring to  FIG. 7A , for a generic substrate (array pillar) reactive sites (X) on the surface can be used to form chemical bonds and/or physical absorption of small molecule ligands. The attachment of ligands to the surface forms a new layer, which is a single molecule thick (i.e., the monolayer). A general ligand consists of (i) a bonding group (Z) which interacts with the substrate reactive site (X), (ii) a backbone which consists of a number of spacer molecules (n) that determine in-large the thickness of the final monolayer, and (iii) a terminal group (A) which interacts with the interface between the monolayer and the fluid/particles in the array. The terminal group (A) interacts with the particles to be sorted. Although  FIG. 7A  shows the bonding group Z and reactive site X, this is just one example and the chemical modification is not meant to be limited to the one type of reaction mechanism in this example. There are two other general mechanistic possibilities: (1) direct bond formation, i.e. the Z group bonds to the reactive site X in a Z—X bond, and/or (2) bond formation with elimination, i.e. the reactive group Z—W bonds to the reactive site X—V in a Z—X bond, with the byproducts W, V eliminated. For example, the reaction of chlorosilanes R—Si—Cl with a silanol on the silica surface, H—O—Si, forms the R—Si—O—Si bond with the elimination of HCl. 
     Referring to  FIG. 7B , monolayers can be formed on metal layers (M) pre-deposited onto the array of pillars. For example, one or more metal layers (M) can be deposited on the pillars (e.g., after the oxidization process that creates the uniform gap size), such that the pillars now have a metal surface (M) over the substrate (and/or over the oxide layer that fills in the inward bow). In  FIG. 7B , the bonding group is identified with ‘Q’ as opposed to ‘Z’ in  FIG. 7A . Ligands (e.g., with the bonding group (Q)) can form coordination complexes directly with the metal surface (M) of the pillar array, forming a tightly packed monolayer. 
     Chemical modification can be used to tune the pillar array to sort smaller particles by decreasing the gap size as illustrated in  FIGS. 8A and 8B . The surfaces of the sorting pillars  314  can be modified with molecules of various length, including aliphatic or aromatic oligomers/polymers, which effectively increase the thickness of the pillars and thereby reduce the gap space between them. By selecting longer ligands, the gap size can be made smaller and therefore the effective cut-off particle size lowered (i.e., smaller particles can be sorted). The backbone of the ligand can be selected to provide a range of mechanical properties between either a rigid, tight packed molecular layer and/or a flexible, disordered layer. Ligands can include small organic molecules, proteins, peptides, nucleic acids, oligosaccharides, and/or synthetic polymers. In one example, pillar surfaces are modified with oligomers of polyethylene glycol (PEG) through siloxane linkages. At approximately 0.36 nm per ethylene oxide residue, for a 12 residue PEG oligomer, this produces an approximately 9 nm decrease in the gap size; for a 20 residue PEG oligomer this is approximately a 14 nm decrease in the gap size. 
       FIGS. 8A through 8D  illustrate schematics of chemical modification of sorting arrays as a means of modifying the gap size between pillars according to an embodiment. Referring to  FIG. 8A , for pillars  314  with their native oxide, a grown oxide layer, and/or a deposited layer of alternative material, e.g. metal, ceramic, polymer, there are reactive sites (X) on the surface of the pillars. The pillars  314  have an initial gap width denoted by g. There is an array floor  805  (which is the floor of the substrate  302  on which the pillars stand).  FIG. 8C  shows an enlarged view  820  that depicts the empty reactive site (X) in  FIG. 8A . In view  820 , the reactive site (X) is not attached to a ligand, but the ligand is to be applied to the pillar array  320  as shown in  FIG. 8B . 
     Referring to  FIG. 8B , chemical attachment of the ligand  810  to the pillars&#39; surfaces forms a monolayer  815  which has a thickness determined by the properties of the ligand packing. The added thickness of the monolayer  815  reduces the gap width (from initial gap width g) to a new effective gap width (g e ). Adjustment of the ligand structure, in particular the backbone, as well as the packing and defect density of the monolayer  815 , can tailor the thickness of the monolayer  815  and thus the tailor the effective gap (g e ). The effective gap (g e ) is the new physical gap size experienced by the particles as they flow through the array  320 , and is formed from the combination of the physical barrier of the pillars plus the added steric barrier of the monolayer. The effective gap is, in general, an approximate value, dependent on the structural, mechanical, and dynamic properties of the monolayer under the operation conditions of the particle sorting.  FIG. 8D  shows an enlarged view  820  in which the reactive sites (X) have been attached to the ligands  810 , thereby extending the diameter of the pillars  314 . 
     Further improvement and refining of the sorting array can be introduced through the terminal group(s) (A) of the ligands, which can be selected to have specific interactions with the fluid and/or particles to be sorted as shown in the schematic of  FIG. 9 . When the particles flow through the pillar array  320 , interactions with the terminal groups of the monolayer  915  leads to increased adhesion and temporary retention on the pillar walls of pillars  314 . These interactions slow down the particle&#39;s flow, as well as causes the particles, on average, to be positioned more at the walls of the pillars, therefore reducing the amount of the flow field it samples. As the pitch of the array is asymmetric with respect to the average fluid flow, particles (such as particles  910 ) that retain and transition between pillars  314  are effectively moved along the critical angle of the array and are sorted out. In one example, thiol terminal groups at the end of PEG-type ligands can be used to formed disulfide linkages between transiting particles such as proteins or other molecules labeled with thiols. In combination with a suitable catalyst agent in the fluid, when proteins (such as particles  910 ) flow through the array  320  they can form disulfide bonds with the pillars  314 , temporarily arresting their flow. In another example, small segments of a chemically stable nucleic acid such as peptide nucleic acid (PNA) can be attached to the pillar walls, to selectively delay and sort out DNA or RNA analytes through reversible base pairing. In another example, patches of hydrophobic ligands embedded within hydrophilic monolayers can be introduced onto pillars, one such pair being aliphatic hydrocarbon ligands and PEG. The hydrophobic patches can be used to interact with hydrophobic domains on proteins, to selectively sort them from solution. 
       FIG. 9A  provides illustration of particle flow in chemically modified sorting array with particles  905  that have no affinity for the surface monolayer  915  and particles  910  which interact with the monolayer  915 . Particles  905  with no affinity follow the flow lines through the array  320  (i.e., exhibit a zigzag mode) and are not subject to any strong interactions with the pillars  314 . The trajectory of these particles  905  is unaffected on average, and they flow without sorting in the array  320 . For example, the particles  905  flow into an outlet  940 . However, particles  910  with a physochemical affinity, caused by molecules on their surface, experience interactions with the molecules of monolayer  915  on the surface of the pillars  314 . The interactions can temporarily bind these particles  910  to the surface of the pillars  314 , and cause particles  910  to, on average, remain closer to the pillar walls of the pillars  314 . Through several sequential binding and dissociation events, the particles  910  are transferred along the direction of the pillars  314  (i.e., exhibit a bump mode in the direction of the critical angle α) and are sorted by the array  320  due to chemical affinity. The particles  910  are sorted into an outlet  945 .  FIG. 9B  is an enlarged view of a cross-section of the nanopillar  314 , monolayer  915 , and particle  910  with affinity according to an embodiment. 
     To chemically modify the pillar array  320 , the ligand can be introduced through chemical vapor deposition (CVD) and/or wet chemistry. To apply the metal, CVD, sputtering, and/or wet chemistry may be utilized. Two detailed examples of chemically modifying the pillars  314  by adding a monolayer discussed for explanation purposes and not limitation, and the two examples using wet chemistry are provided below. 
     For illustration purposes, an example method of modification of a microfluidic device using poly(ethyleneoxide) (PEG) ligand modifiers is provided below: All glassware to be exposed to chlorosilanes, is first washed in an isopropanol bath saturated with potassium hydroxide for at least 24 hours, then rinsed thoroughly with deionized water and dried in an oven at 140° C. for 12 hours. 
     A 100 mL round bottom flask is removed from the 140° C. oven and quickly sealed with a septum. A nitrogen gas purge is set up through the septum using needles, and the flask allowed to purge for 10 minutes. 30 mL of anhydrous toluene is transferred into the flask via cannula. Via syringe, 600 μL of n-octyldecyltrichlorosilane is injected to form a 49 mM solution. The flask is momentarily vortexed to mix the reagents homogenously. This forms the passivation solution. A 500 mL reactor and 3-neck head are removed from the 140° C. oven and then quickly sealed together, with each inlet closed with a septum. A nitrogen gas purge is set up through the septum using needles, and the flask allowed to purge for 10 minutes. Via cannula, 20 mL of the passivation solution in the 100 mL flask is transferred to the reactor. The reactor is gently shaken to swish the passivation solution around the walls of the reactor thoroughly. The same is done for the 100 mL flask using the remaining passivation solution. This gentle shaking is repeated every 10-15 minutes, for 1 hr. Between shaking, the glassware is allow to sit at ambient temperature. This procedure is to passivate the walls of the glassware against further silizanizaiton. The passivation solution is then poured out of the reactor, and the reactor washed sequentially, 3× each, with toluene, acetone, isopropanol and deionized water. The same is done for the 100 mL flask. The glassware is then returned to the 140° C. oven and allowed to dry 12-14 hours. 
     The 100 mL round bottom flask is removed from the 140° C. oven and quickly sealed with a septum. A nitrogen gas purge is set up through the septum using needles, and the flask allowed to purge for 30 min. 100 mL of anhydrous toluene is transferred into the flask via cannula. Via syringe, 100 μL of 2-(methoxypoly(ethyleneoxy) 6-9 propyl)dimethylchlorosilane is injected to form an approximately 2 mM solution. The flask is momentarily vortexed to mix the reagents homogenously. This modification solution is used within the day of its preparation. 
     Silica/silicon based microfluidic devices (chips) are cleaned for 30 min in an oxygen plasma to remove organic surface contamination. The chips are transferred then to a 0.1M aqueous nitric acid solution for 10 min to hydrolyze any surface siloxane bonds to silanols. The chips are then washed sequentially, using a squeeze bottle stream, in deionized water, acetone, ethanol, and then isopropanol. The chip is then set face-up on a fresh texwipe and immediately dried off using a stream of nitrogen gas, pushing solvent from the middle to outside of the chip. The chips are then set on a custom glass holder (which sets the chips horizontal/face-up inside the reactor, as described below). 
     A 500 mL reactor and 3-neck head are removed from the 140° C. oven. A stir bead along with the glass holder and chips are set into the reactor, and then quickly sealed together, with each inlet closed with a septum. A nitrogen gas purge is set up through the septum using needles, and the reactor allowed to purge for 30 minutes. 
     Via a cannula, the modification solution (with the ligand) is transferred into the reaction flask until the solution level is above the chips. Nitrogen positive pressure is then maintained using a bubbler. The reaction is allowed to run for 2 hours, at ambient temperature, with stirring. The reactor is then opened and the chips cleaned (one-by-one) by rinsing sequentially, using a squeeze bottle stream, toluene, acetone, isopropanol, then deionized water. The chip is then set face-up on a fresh texwipe and immediately dried off using a stream of nitrogen gas, pushing solvent from the middle to outside of the chip. The chips are then set in a glass holding jar with a septum. A nitrogen gas purge is set up through the septum using needles, and the chips allowed to dry overnight (approximately 12-14 hours). 
     Use of the sub-headings is now discontinued.  FIG. 11  illustrates a chip  1100  (fluidic device) having the pillar array  320  according to an embodiment. The chip  1100  has an inlet  1105  to receive fluid containing the different sized particles (i.e., biological entities) to be sorted. The inlet  1105  may be an opening or hole in the walls around the nanopillar array  320  or may span the width of the nanopillar array  320  through which fluid (e.g. water, electrolyte solutions, organic solvents, etc.) and particles (e.g., biological entities) can flow. Particles having a size greater than the critical dimension are bumped (i.e., bumped mode) through the pillar array  320  in the direction of the critical angle, and these particles larger than the critical dimension are collected at outlet  940 . The critical dimension is the size (e.g., diameter) of a round shaped particle and/or persistence length of chain structure, such as DNA, that is too large to zigzag through the nanoarray  320 . Particles having a size smaller than the critical dimension zigzag (i.e., zigzag mode) through the pillar array  320  in the direction of fluid flow, and these smaller particles are collected at the outlet  945 . The particles having the size smaller than the critical dimension follow the direction of the fluid flow, and are sorted through the outlet  945 . In one case, the pillars  314  may have the chemical modification as discussed herein, which can further reduce the gap size and/or sort particles having affinity to the chemical modification. The outlets  940  and  945  may be openings through which the sorted particles can flow and be collected in bins after sorting. 
       FIG. 12  is a method  1200  of providing a fluidic apparatus  1100  (e.g., chip  1100 ) is provided according to an embodiment. Reference can be made to  FIGS. 1-11  discussed above. At block  1205 , the inlet  1105  is configured to receive a fluid. At block  1210 , the outlet (e.g., outlets  940 ,  945 ) is configured to exit the fluid. The nanopillar array  320  is coupled to the inlet and the outlet, and the nanopillar array  320  is configured to allow the fluid to flow from the inlet to the outlet at block  1215 . 
     At block  1220 , the nanopillar array  320  comprises nanopillars  314  arranged to separate biological entities (particles) by size. At block  1225 , the nanopillars  314  are arranged to have a gap G separating one nanopillar  314  from another nanopillar  314 , and the gap is constructed to be in a nanoscale range (e.g., sub-100 nm). 
     The one nanopillar is to the side of the other nanopillar, such that the gap G is in between. The gap between the one nanopillar and the other nanopillar is uniform along a vertical axis of the one nanopillar and the other nanopillar (such as, e.g., gap G 5  as shown in  FIG. 10B ). 
     The nanopillar array comprises an oxide layer  316  applied on the nanopillars, and the oxide layer  316  causes the gap to be uniform along a vertical axis of the one nanopillar and the another nanopillar (e.g., the gap G 5  is uniform up and down the space between the two nanopillars  314  in  FIG. 10B ). 
     The oxide layer  316  causes a size of the gap (e.g., gap G 5 ) to be as small as about 20 nanometers while the gap remains uniform along the vertical axis (e.g., y-axis in  FIG. 10B ). The oxide layer  316  causes unevenness in a diameter (e.g., the diameter of pillar  314  is not uniform in  FIG. 10A ) of the nanopillars to be uniform in  FIG. 10B , resulting in the gap being uniform along the vertical axis of the one nanopillar and the other nanopillar. An increase in a thickness of the oxide layer  316  causes a decrease in a size of the gap. 
     In one case, the size of the gap ranges from 20-300 nm. In another case, the size of the gap may be formed to be less than 100 nm, may be less than 80 nm, may be less than 60 nm, may be less than 40, may be less than 30, may be less than 25, etc., according to the desired size of the particles to be separated. For example, 100 nm particles can be sorted/separated with 240 nm size gaps according to an embodiment. 
     A monolayer (e.g., the monolayer in  FIGS. 7A, 7B , monolayer  815  in  FIG. 8B , and/or monolayer  915  in  FIG. 9A ) is applied to the nanopillars  314  to reduce a size of the gap. The gap having a reduced size is configured to separate smaller entities relative to when the monolayer is not applied to the nanopillars. 
       FIG. 13  is a method  1300  of forming a nanopillar array  320  according to an embodiment. Reference can be made to  FIGS. 1-12 . 
     At block  1305 , the hard mask layer  304  is disposed on the substrate  302 . At block  1310 , the resist layer  306  is patterned into a pattern (resist pattern  308 ) of the nanopillar array  320  in which the resist layer  306  was disposed on the hard mask layer  304 . 
     At block  1315 , the resist layer (resist pattern  308 ) is utilized to pattern the hard mask layer  304  into the pattern (hard mask pattern  312 ) of the nanopillar array  320 , such that both the resist layer and the hard mask layer have the pattern of the nanopillar array  320 . 
     At block  1320 , the substrate  302  is patterned into the pattern of the nanopillar array  320  such that the nanopillar array  320  is now formed, wherein the resist layer (resist pattern  308 ) and the hard mask layer (hard mask pattern  312 ) are removed and wherein nanopillars  314  in the nanopillar array have a first gap size (e.g., gap size G 1  and/or G 2  in  FIG. 10A ) in a side-to-side relationship relative to each other. At block  1325 , the first gap size is reduced to a second gap size (e.g., gap size G 5 ) by disposing the oxide layer  316  on the nanopillar array  320 . 
     The resist layer is patterned into the pattern (resist pattern  308 ) of the nanopillar array  320  by at least one of electron-beam lithography and/or nanoimprint lithography or another form of lithography. 
     Utilizing the resist layer to pattern the hard mask layer into the pattern of the nanopillar array comprises performing reactive ion etching to etch the hard mask into the pattern (hard mask pattern  312 ) of the nanopillar array  320 . 
     Patterning the substrate  302  into the pattern of the nanopillar array such that the nanopillar array is formed comprises performing reactive ion etching to etch the substrate into the nanopillar array  320 . 
     Reducing the first gap size (e.g., gap size G 1  and G 2 ) to the second gap size (gap size G 5 ) by disposing the oxide layer  316  on the nanopillar array  320  comprises reducing the second gap size (e.g., to less than 300 nanometers, to less than 100 nanometers, etc.). 
     Reducing the first gap size to the second gap size by disposing the oxide layer on the nanopillar array causes each of the nanopillars to have a uniform shape and causes the second gap size to be uniform throughout the nanopillar array for the side-to-side relationship of the nanopillars (as shown in  FIGS. 10A and 10B ). Before reducing the first gap size to the second gap size by disposing the oxide layer, the nanopillars have an inward-bowed shape at a middle of the nanopillars at a nanoscale level. Reducing the first gap size to the second gap size by disposing the oxide layer both fills in the inward-bowed shape at the middle and straightens the nanopillars into a cylinder-like shape. 
     As discussed herein, embodiments provide silicon chips with nanopillars and nanogaps that can separate molecules and particles by size from the micron regime down to the nanometer regime. The size of two or more entities (particles) that can be separated depends on the size of the gaps (i.e., nanogaps) between the nanopillars. The state-of-the-art has no technologies for sorting entities by size in the 10-100 nm scale. However, embodiments described herein provide a mechanism for sorting entities within, above, and below this range (10-100 nm). For example, embodiments can sort a 30 nm particle from a 40 nm particle. Furthermore, embodiments provide continuous flow bio-separation, which means that particle sorting is continuous as fluid and the entities (to be separated) are introduced into one or more inlets of the nanopillar array  320 , and the continuous flow bio-separation nanopillar array  320  continuously sorts the entities without requiring any type of reset. 
     For example, the technology of embodiments can be used to stream a solution mix through the chip  1100 , obtaining a continuous separation of particles within a specified size range. A heterogeneous particle solution is introduced at the inlet of the chip  1100  and a solution flow carries the particles through a pillar network (i.e., pillar array  320 ). Particles of larger sizes bounce off the nanopillars  314  according to a preset angle (i.e., critical angle α) defined by the offset δ and the pitch λ of the nanopillars  314 . In this way, the trajectory of the larger particles is directed (bump mode) toward a specific microchannel exit (e.g., outlet  940 ) where the separated sample can be extracted, while smaller particles will zigzag through the nanopillars  314  parallel to the direction of fluid flow where the smaller particles exit the chip  1100  through a different microchannel (e.g., outlet  945 ). 
     The improvements in embodiments allow for this type of continuous flow separation to operate at the nanometer scale, permitting efficient separation of bio-markers, bio-molecules, sub-cellular components, exosomes, viruses, immuno-assays, drug screening, and protein aggregates on a Si chip (such as, e.g., chip  1100 ). Embodiments are a significant scale down from the micron scale in state-of-the-art. The improvement over the state-of-the-art was achieved through the nanofabrication of nanopillars capable of sorting particles at the nanoscale. Embodiments also demonstrate that, at this new scale, a different flow regime applies and improves the separation method. At this scale, dead flow areas between nanopillars are proportionally significant with respect to the nanopillar size. The presence of these dead flow areas contributes to a narrower fluidic gap between nanopillars than the physical gap (G) defined by the nanopillar wall to wall distance. This results in the ability to sort a particle size smaller to what the original theory predicts. 
       FIG. 14  is a top view of a schematic representing an arrangement of the pillars  314  in the nanopillar array  320  according to an embodiment. In this example, the pillar array  320  may be considered as multiple pillar arrays. For example, the pillar array  320  includes a symmetric part/arrangement  1405  of pillars  314  and an asymmetric part/arrangement  1410  of pillars  314 . The symmetric part  1405  has a critical angle that is (virtually) 0°, while the asymmetric part  1410  has a critical angle α (defined with respect to the z-axis in  FIG. 14 ). 
     In  FIG. 14 , the flow stream (i.e., fluid flow direction) is horizontal on the average, and the pillar rows are tilted to an angle (i.e., forming the critical angle α) in the asymmetric part  1410  of the nanopillar array  320 . At sufficiently slow flow rates of the fluid, the distance (gap G) between the pillars  314  together with the critical angle define the size (smaller than a critical dimension) of the particles that are able to follow the flow direction by zig-zagging through the pillars  314 , and the size (equal to and/or greater than the critical dimension) of the particles that will be displaced (bumped) by the angle of the pillar rows. In one case, a slow flow rate may correspond to a flow slower than 500 μm/s. 
     The pillars  314  have a diameter, a pillar pitch λ, a gap (G), and a row-to-row shift (δ). The row-to-row shift (δ) is in the asymmetric part  1410  because there is no row-to-row shift in the symmetric part  1405 . In the example of  FIG. 14 , two example particles of different sizes are traversing through the pillar array  320 . The larger particle  1450  is displaced (indicated by a dash line) across the array  320  according to the pillar angle (i.e., critical angle α), while the smaller particle  1455  follows the deterministic flow (solid line) through the array  320  zig-zagging through the pillars  314 . 
       FIG. 15  is a schematic of the chip  1100  now with two inlets and with two particles of different sizes traversing through the pillar array  320  according to an embodiment. The larger particle  1450  is displaced along the dashed line across the array  320  according to the pillar angle (critical angle), while the smaller particle  1455  follows the deterministic flow through the array by zig-zagging through the pillars  314 . The large nanoparticle  1450  and the smaller nanoparticle  1455  exit the array through separate microfluidic channels. For example, the large nanoparticle  1450  (e.g., equal to and/or above the critical dimension) exits the array through outlet  940 , while the small nanoparticle  1455  (e.g., below the critical dimension) exits the through outlet  945 . In this example, the fluid, which may be a buffer solution, can be introduced through the buffer in inlet  1105 . The buffer solution (also referred to as a pH buffer or hydrogen ion buffer) is an aqueous solution consisting of a mixture of a weak acid and its conjugate base, or vice versa. The sample, which includes the nanoparticles  1450  and  1455  to be sorted, is introduced through the same inlet  1510 . Although only two particles are shown, the same sorting process applies to numerous particles having the different sizes introduced into the sample inlet  1510 . 
       FIGS. 16A, 16B, 16C, and 16B  illustrate experimental results of passing two populations of nano-beads through the nanopillar array  320  according to an embodiment.  FIGS. 16A and 16B  correspond to a population of 70 nm diameter beads, while  FIGS. 16C and 16D  correspond to the population of 50 nm diameter beads. In  FIGS. 16A and 16C  the respective beads&#39; trajectories are recorded with a video camera on a fluorescence microscope. In this example, the gap size (G) between the nanopillars  314  is 210 nm. 
       FIG. 16A  is the image of particle trajectories for 70 nm beads being displaced in a 5.7° critical angle array ( 320 ).  FIG. 16A  shows that the bead trajectory of 70 nm diameter beads is angled with respect to the flow direction. The average trajectory angle observed for the 70 nm beads is a 5.7° angle. The marked trajectories of three 70 nm particles are shown. The angle between the flow direction (i.e., fluid flow direction) and particle trajectory describes the degree to which the particle is bumping in the array  320 . For these 70 nm particles, a plot of trajectory angle as a function of velocity shows a positive value close to the critical angle in  FIG. 16B . In other words, the average trajectory angle of the 70 nm beads plotted in  FIG. 16B  is approximately 5.7° which is expected for the 5.7° critical angle of the array  320  in the experiment. 
       FIG. 16C  is the image of particle trajectories for 50 nm diameter beads sorted in the same array  320 . Under the same conditions (including the same nanopillar array  320  with the 5.7° critical angle), the 50 nm beads are introduced in the array  320 , and their trajectories are recorded in  FIG. 16C . These 50 nm particles are not displaced (i.e., not in bump mode) in the array  320 , which is observed in the trajectory angle and velocity plot in  FIG. 16D . The plot in  FIG. 16D  shows a near-symmetric distribution of trajectory angles around 0° (i.e., in accord with the flow axis in the array  320 ). Also,  FIG. 16C  shows that the average trajectory angle followed by the 50 nm beads is close to 0°. 
     As seen in  FIGS. 16A, 16B, 16C, and 16D , the nanopillar array  320  is configured to sort the 50 nm beads from the 70 nm beads, by outputting the 50 nm beads in a first direction along the flow axis, while outputting the 70 nm beads in a second direction along the critical angle of the pillar array  320 . 
       FIG. 17  is a chart illustrating example data of the approximate gap sizes to separate a particle of one size from a particle of another size utilizing the nanopillar array  320  according to an embodiment. It is noted that the example data in chart in  FIG. 17  is meant for illustration purposes and not limitation. The gap sizes (G) between the nanopillars are listed horizontally, and particle diameters are listed vertically. The result of each experiment is described as either No displacement, Partial displacement, or displacement 100%. Displacement 100% means that the trajectory angle of the particles over the array is the same as the set pillar critical angle, or within 15% of this angle. Partial displacement accounts for particles trajectories ranging from 15 to 85% of the nanopillar critical angle. No displacement represents any experiment were the trajectory angle of the particles is less than 15% of the nanopillar array critical angle. 
     In one implementation, embodiments rely on manufacturable (silicon) pillar arrays  320  with uniform gaps between the pillars and with dimensions in the sub-100 nm regime. These arrays  320  are for the sorting and separation of biological entities at these dimensions, such as DNA, RNA, exosomes, individual proteins, and protein complexes. Uniform gap sizes are utilized to obtain efficient sorting, e.g., to sort a 20 nm particle from a 10 nm particle according to embodiment. This is particularly challenging for gaps in the sub-100 nm regime where there could be inherent variations greater than the dimensions of the particles to be sorted. This is usually caused by non-uniform nanopatterning at this scale, and feature variations in sizes and shapes due to the reactive-ion etch (RIE) process. Demonstrated sorting pillar gaps found in the state-of-the-art have dimensions in the micron range and therefore cannot sort even close to this fine of a scale. 
     Therefore, consistent gaps in the nanometer regime are required to sort, for example, a protein aggregate. Sorting of individual proteins (size range of 1-10 nm) is traditionally performed using ion exchange chromatography or gel electrophoresis, which are load-and-run techniques rather than continuous flow and thus much slower. However, embodiments provide a continuous flow separation process and mechanism, which is configured to sort individual proteins (or other particles) in the range of 1-10 nm, without requiring ion exchange chromatography or gel electrophoresis. 
       FIG. 18  is a method  1800  for sorting entities according to an embodiment. Reference can be made to  FIGS. 1-17 . 
     At block  1805 , the entities are introduced into the nanopillar array  320 , and the entities include a first population and a second population. The nanopillar array  320  includes nanopillars  314  arranged to have a gap separating one from another, and the nanopillars are ordered to have an array angle relative to a fluid flow direction. 
     At block  1810 , the entities are sorted through the nanopillar array  320  by transporting the first population of entities less than a predetermined critical size in a first direction (e.g., toward outlet  945 ) and by transporting the second population of entities at least the predetermined size in a second direction (e.g., toward outlet  940 ) different from the first direction. 
     At block  1815 , the nanopillar array  320  is configured to employ the gap with a gap size less than 300 nanometers or less than 100 nanometers in order to sort the entities having a sub-100 nanometer size. 
     When the entities have a nanometer size equal to or greater than 7 nanometers, the nanopillar array is configured accordingly to sort the entities having the nanometer size equal to or greater than 7 nanometers. When the entities have a nanometer size equal to or greater than 7 nanometers, the gap size is configured accordingly to sort the entities having the nanometer size equal to or greater than 7 nanometers. 
     A lower limit of the gap size may be about 20 nanometers. A thickness of an oxide layer  316  applied to the nanopillar array  320  causes the gap size of the gap to be about 20 nanometers while the gap remains uniform. In other words, the gap is uniform along the vertical axis (e.g., y-axis) between any two nanopillars  314  (i.e., no gap variation), and each of the gaps throughout the nanopillar array  320  has the same gap size. 
     The gap size of the gap is tuned to sort the first population of the entities less than the predetermined critical size in the first direction while sorting the second population of the entities at least the predetermined size in the second direction. Tuning the gap size is based on a thickness of the oxide layer  316  applied to the nanopillar array  320 . Further tuning the gap size can be based on a monolayer (e.g., without a metal applied in  FIG. 7A , and/or with a metal applied in  FIG. 7B ) applied to the nanopillars by chemical modification. The chemical modification forms a monolayer (e.g., such as monolayer  815 ,  915 ) on the nanopillars  314  such that the first population has an affinity to the monolayer and the second population has no affinity to the monolayer. Having the affinity to the monolayer directs the first population (e.g., such as entities  910 ) of the entities to be transported in the first direction (e.g., to outlet  945 ). Not having the affinity to the monolayer allows the second population (e.g., such as entities  905 ) to be bumped in the second direction to outlet  940 . In one case, both entities  905  and  910  may be about the same size, and the affinity of entities  910  causes the entities  910  to proceed toward the outlet  945 . The entities comprise at least one of bio-markers, bio-molecules, sub-cellular components, exosomes, viruses, immuno-assays, and/or protein aggregates. 
     Exosomes are becoming more and more important science but are too small, e.g., 30-100 nm, to be sorted by state-of-the-art arrays. Exosomes are now believed to be present in all body fluids, and represent a new way of thinking about cell signaling. These small extracellular vesicles are thought to play a role in a large number of biological functions. For example, exosomes are a messaging system and regulation system, which may contain and transfer DNA, RNA, protein, etc. In the nanopillar array  320 , the gap size can be narrowed by the oxide layer  316  to sort one size exosome from larger size exosome, and/or to sort the smaller exosome from a different (larger) particle. Additionally, exosomes have special affinity (i.e., attraction) to certain ligands. For example, a monolayer  815 ,  915  of the lipid membrane integrating ligands, such as [6-(pyren-2-yl)octyl]silane or 3-[(8-silyloctyl)oxy]cholesterol, can be applied to the pillars  314  to direct the exosomes in a first direction while directing the different particles in a second direction because the different particles do not have the special affinity. Therefore, even if the different particles have a same (or similar) size as the exosomes in one case, the exosomes can still be sorted because of their special affinity to the certain ligands. Although certain ligands having a special affinity to exosomes are discussed for explanation purposes, it is understood that the certain ligands having a special affinity to exosomes are not limited to these examples. 
       FIG. 19  is a method  1900  of sorting entities according to an embodiment. Reference can be made to  FIGS. 1-18 . 
     At block  1905 , entities to be sorted are introduced into the nanopillar array  320  (e.g., via inlet  1105  and/or inlet  1510 ), and the entities include a first population and a second population. The nanopillar array  320  includes nanopillars  314  arranged to have a gap G separating one from another, and the nanopillars are ordered to have an array angle (e.g., critical angle) relative to a fluid flow direction. 
     At block  1910 , the nanopillar array  320  is configured to receive the entities at the outlet (such as the outlet  940  and/or  945  where each outlet may be attached/coupled to a collection tray or collection bin) based on being sorted, such that the first population of the entities are output in a first direction and the second population of the entities are output in a second direction different from the first direction; 
     At block  1915 , a gap size of the gap G is tuned to sort the first population in the first direction and the second population in the second direction, and the gap size is tuned according to at least one of a thickness of an oxide layer  316  disposed on the nanopillar array  320  and/or a chemical modification (such as in  FIGS. 7-9 ) to the gap. 
     When the gap size is tuned by the oxide layer  316 , the oxide layer  316  reduces the gap size to a first dimension. When the gap size is tuned by the chemical modification, the chemical modification further reduces the gap size to a second dimension, and the second dimension is smaller than the first dimension. 
     The first dimension corresponds to the oxide layer  316  reducing the gap size to about 20 nanometers while the gap remains uniform. The second dimension corresponds to the chemical modification (e.g., attached ligand) further reducing the gap size below 20 nanometers (e.g., after the oxide layer  316  has been deposited). For the second dimension, the chemical modification may reduce the gap size to 18, 16, 14, 12, and/or 10 nanometers. In one case, the chemical modification may reduce the gap size to below 10 nanometers as the second dimension. In another case, the chemical modification (using longer ligands) may reduce the gap size to 8, 6, 4, and/or 2 nanometers as the second dimension. If desired, the chemical modification can nearly close the gap by reducing the gap size to less than 2 nanometers as the second dimension. 
     When the gap size is tuned by the chemical modification, the chemical modification reduces the gap size to a first dimension. It is contemplated that the chemical modification may be applied to the nanopillars  314  even in a case when the oxide layer  316  is not applied. 
     The chemical modification forms a monolayer on the nanopillars such that the first population has an affinity to the monolayer and the second population has no affinity to the monolayer. Having the affinity to the monolayer directs the first population of the entities to be output in the first direction. The entities comprise at least one of bio-markers, bio-molecules, sub-cellular components, exosomes, viruses, immuno-assays, and/or protein aggregates. 
     According to an embodiment,  FIG. 20  is a method  2000  of sorting entities. Reference can be made to  FIGS. 1-19 . 
     At block  2005 , entities are introduced into the nanopillar array  320 , and the entities including a first population and a second population. The nanopillar array  320  includes nanopillars  314  in an ordered arrangement. The nanopillars have a chemical modification. Various illustrations of chemical modifications have been discussed in  FIGS. 7-9 . 
     At block  2010 , the output (e.g., outlets  940  and  945 ) receives the entities after sorting, such that the first population of the entities are output in a first direction (e.g., outlet  945  in  FIG. 9A ) based on the first population having an affinity to the chemical modification and the second population of the entities are output in a second direction (e.g., outlet  940  in  FIG. 9A ) different from the first direction. Also, there may be an operator who receives the sorted entities now separated into/through one or more outlets (outlets  940  and  945 ). The operator may utilize or attach separate collection apparatuses to separately receive and hold the collected entities. 
     The second population does not have the affinity to the chemical modification, such as entities  905  in  FIG. 9A . By the second population not having the affinity to the chemical modification, the second population is output in the second direction (e.g., output outlet  940 ). 
     State-of-the-art technologies (e.g., such as silicon technologies) make use of a micro-structured and nano-structured matrix to separate molecules based on the molecule-structure interactions. Usually a large separation matrix patterned over several millimeters is required for accurate separation, which in turn requires a large chip size and long operation time. 
     According to an embodiment, the separation matrix (e.g., shown in  FIG. 25 ) in one example may be 35 μm by 400 μm, which is tens of times smaller than the previous approaches of the state-of-the-art. Therefore, embodiments allow high-density chip integration, less expensive fabrication, and high-throughput analysis. 
     According to an embodiment, nanofluidic devices may use nanopillar arrays  320  to deflect biopolymers according to their sizes.  FIGS. 21A and 21B  are schematics of a top view of the nanopillar array  320  illustrating the principle of biopolymer separation according to an embodiment. 
       FIG. 21A  illustrates that the biopolymers flow downward through the nanopillar arrays, which are tilted to an angle α relative to the fluid flow direction.  FIG. 21A  shows separation of long biopolymers  2105  and short biopolymers  2110  through nanopillar arrays  320 . 
       FIG. 21B  is an enlarged view of  FIG. 21A , and  FIG. 21B  illustrates that long biopolymers  2105  have a larger radius of gyration R, which depends on the polymer length L and the pillar gap confinement, and the long polymer  2105  bumps into the pillars  314  to travel along the slanted path (i.e., travel along the critical angle α). For example, the long polymer  2105 , with a radius of gyration R, bounces off the pillars (pillar diameter size and gap size G) and travels along the slanted arrays, which are tilted at angle α. Short biopolymers  2110 , which have a smaller radius of gyration r, in contrast move through the pillars  314  in a zigzag fashion following the fluid flow direction. Note that the gap size G can be the same in both the x-axis and z-axis, and that the pitch λ can be the same in the x-axis and z-axis. The row shift fraction ε is 0.1, where ε=δ/λ. The critical bumping diameter/size is designated C, where C=γ·G. The critical bumping size divided by the gap size G is the γ. 
       FIG. 22A  is a schematic of an example nanopillar array  320 .  FIG. 22A  shows the nanopillar array  320  with the tilted angle as 5.7° relative to the fluid flow direction. 
     The nanopillar array  320  has a pillar array wall  2210 , and the nanopillar array  320  is utilized in fluorescence microscope images in  FIGS. 22B, 22C, 22D, and 22E . 
     Experimental results show the deflection of T4 bacterial phage DNA by the nanopillar arrays  320 .  FIGS. 22B, 22C, 22D, and 22E  show fluorescence microscope images of the deflection and focusing of T4 bacterial phage DNA (166 kilo base pairs (kb)) in the nanopillar array  320  with critical angle α=5.7°.  FIGS. 22B, 22C, 22D, and 22E  are consecutive fluorescence images of the travel of the T4 DNA, with two preventative DNA flowing at the tilted angle. 
     The DNA molecules, e.g. DNA 1 A and DNA 2 A as marked in the  FIGS. 22B, 22C, 22D, and 22E , flow through the channels from the bottom to the top at about 750 μm/sec (driven by capillary force). The DNA molecules are stretched and deflected to the left side of the arrays  320  toward the left pillar array wall  2210 .  FIG. 22B  shows the DNA 1 A and DNA 2 A focused to the left, as DNA 2 A enters the field of view.  FIG. 22C  shows both DNA 1 A and DNA 2 A being deflected toward the left wall  2210  by the nanopillar array  320 . The traces of DNA 1 A and DNA 2 A are measured as 5.7° relative to the fluid flow direction, exactly the same as the designed pillar array tilting angle; this shows perfect control of DNA flow direction using such pillar arrays  320 . The deflected DNA (DNA 1 A and DNA 2 A) eventually merge into the single-lane trace along the wall  2210 , as shown in  FIGS. 22D and 22E , which shows the nanopillar array&#39;s ( 320 ) capability of focusing biopolymer samples for easy collection. In  FIGS. 22B, 22C, 22D , and  22 E, the length of the T4 DNA is 166,000 base pairs (bp). 
       FIGS. 23A, 23B, 23C, and 23D  show consecutive fluorescence images illustrating 5 kb DNA flowing through nanopillar arrays  320  according to an embodiment. In  FIGS. 23A, 23B, 23C, and 23D , the 5 kb DNA (DNA 1 B, DNA 2 B, DNA 3 B) is deflected and focused using the same array  320 . As can be seen in  FIGS. 23A-23D , the shorter  5   k  DNA molecules (DNA 1 B, DNA 2 B, DNA 3 B) are also deflected, and eventually exit the array only from the left side of the array  320  near the wall  2210  as shown in  FIG. 23C .  FIGS. 23A, 23B, and 23C  show the progression of the 5 kb DNA (DNA 1 B, DNA 2 B, DNA 3 B).  FIG. 23D  shows that the DNA 1 B, DNA 2 B, and DNA 3 B have exited the nanopillar array  320 . The 5 k DNA molecules that flow at a speed of 500 μm/sec and 350 μm/sec are effectively bumped to the designed array angle 5.7°. 
     In comparison, even shorter DNA, e.g., 1 kb DNA and 2 kb DNA are not deflected in the same array  320 . Another example is provided below for the even shorter DNA which exhibits the zigzag mode instead of the bump mode.  FIGS. 24A, 24B, 24C, 24D, 24E, 24F, 24G, and 24H  show consecutive fluorescence images illustrating that 2 kb DNA molecules are not deflected. In other words, the 2 kb DNA molecules exhibit the zigzag mode. The 2 kb DNA is denoted DNA 1 C and DNA 1 , and the flow rate is approximately 100 μm/sec. According to embodiments, the results show that the nanopillar arrays  320  are capable of continuously separating biopolymers based on their dimensions. In the example design, the approximately 250 nm pillar gaps (G) can separate biomolecules (e.g., DNA, RNA, etc.) equal to and bigger than 5 kb from the biomolecules smaller than 5 kb. 
     Furthermore, by designing different pillar dimensions (e.g., different gap sizes) on the same chip, embodiments can separate biopolymers with all different dimensions in a cascaded fashion for fast sample purification as shown in  FIG. 25 . By building nanopillar stages with gradually changed sizes, the biopolymer samples can be separated and accurately retrieved according to their specific sizes. 
     According to an embodiment,  FIG. 25  is a schematic of a cascaded pillar array design to separate biopolymers (e.g., entities) according to their lengths/dimension.  FIG. 25  demonstrates a cascading array concept that can be used to increase the resolution power of the pillar arrays  320 . The cascaded pillar array structure  2500  consists of multiple (N) stages of pillar arrays. In this example, there are 3 stages of nanopillar pillar arrays  320  which show array  2 A output to array  3 A and array  3 B, array  3 A output to array  4 A and array  4 B, and array  3 B output to array  4 C and array  4 D. 
     Given properly designed array lengths and widths, each array  2 A,  3 A,  3 B,  4 A,  4 B,  4 C, and  4 D has its own specially designed critical bumping diameter D X , where X denotes the array number such as  2 A,  3 A,  3 B,  4 A,  4 B,  4 C,  4 D. In this case, the cascaded array structure  2500  is designed so that D 4A &lt;D 3A &lt;D 4B &lt;D 2A &lt;D 4C &lt;D 3B &lt;D 4D . 
     Each pillar array is capable of binary separation of biopolymers into two groups: one group that is larger than the array critical (bumping) diameter and thus bumped to the side into the next stage, and one group that is smaller than the critical (bumping) diameter and sent directly down to the next stage. When the one group is bumped to the subsequent stage, this group is bumped along the critical angle into the next stage. The critical angle or bumping direction is generally shown to the right side in  FIG. 25 , but the bumping direction could be to the left direction. However, when the other group is sent directly down to the next stage, this group travels (e.g., zigzag mode) along the fluid flow direction. As a result, the N-stage cascaded array is cable of separating the biopolymers into 2 N  groups, as shown in the  FIG. 25 , where N=3 and there are accordingly 8 separated groups. 
     Assume that the effective diameter of the biopolymer is D, and the effective diameter D determines which direction the biopolymer flows through the cascaded array structure  2500 . For instance, the process of bumping a biopolymer with an effective diameter of D 3A &lt;D&lt;D 4B  can be explained as follows. First, the biopolymer travels through array  2 A without bumping, because D&lt;D 4B &lt;D 2A . Therefore, the biopolymer goes down to the next array  3 A. Second, as the biopolymer is larger than D 3A , the polymer is bumped to the right side of array  3 A to end up in array  4 B. Third, since the biopolymer is smaller than the critical diameter D 4B , the biopolymer goes down (i.e., zigzag mode) in the array  4 B to end up in the 3 rd  separation group (outlet). 
     As can be seen, the cascaded pillar array structure  2500  is binary and yields 2 N  separations through outlets. In  FIG. 25 , there are 8 outlets to receive the 8 separation (i.e., 2 3 =8). It is understood that additional stages can be added where each additional stage has twice as many nanopillar arrays as the previous stage, and such that each additional stage has a nanopillar array to receive both the zigzag mode output and bumped mode output of the nanopillar arrays of the previous stage. Therefore, higher resolution can be provided by increasing the number of stages. Also, there can be fewer stages than 3 in one implementation. 
     Each of the nanopillar arrays  320  in  FIG. 25  can be fabricated as discussed herein, such that the respective arrays, arrays  2 A through  4 D, each has its desired gap size (G) according to its own critical bumping diameter D X . For example, array  3 A has a smaller gap size than array  2 A because D 3A &lt;D 2A . Additionally, array  4 A has a smaller gap size than array  3 A because D 4A &lt;D 3A . 
     A separation resolution estimation is provided below, and the separation resolution can be applied to the cascaded nanopillar array structure  2500  based on the number of stages (N). The critical size of the entity (e.g., biopolymer) that can be bumped in an array is designated C (critical size). The DNA size R that can be separated in an array with a gap G in terms of base pairs is roughly dependent on the gap size by C=γ·G, where γ is approximately 0.5 using the current design. 
     By sending DNA into the N-staged cascaded pillar arrays with different sizes, the cascaded nanopillar array structure  2500  can separate 2 N  DNA sizes. If the critical gap size is changed from g to G, the resolution (Res) is as follows: 
     
       
         
           
             Res 
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                 Δ 
                 ⁢ 
                 
                     
                 
                 ⁢ 
                 C 
               
               = 
               
                 
                   
                     γ 
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                       ( 
                       
                         G 
                         - 
                         g 
                       
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                     2 
                     N 
                   
                 
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     Accordingly, this resolution (Res) can be tuned by array geometries. For example, for g=100 nm, G=110 nm, N=2 (step of gap size as 10 nm), γ=0.5, this results in Res=1.25 nm, or approximately 17 base pairs (DNA as blobs, using De Genes model). The resolution is estimated as 11 bp (base pairs) and 45 bp for g/G=50/60 or 150/160 nm. 
       FIG. 26  is a method  2600  of sorting biopolymers according to an embodiment. At block  2605 , the biopolymers are introduced into the nanopillar array  320 , and the biopolymers include a first population and a second population. The nanopillar array  320  includes nanopillars  314  arranged to have a gap (G) separating one from another. 
     At block  2610 , the biopolymers are sorted through the nanopillar array  320  by transporting the first population of the biopolymers less than a predetermined bumping size (e.g., less than the critical bumping size C) according to a fluid flow direction and by transporting the second population of the biopolymers at least the predetermined bumping size (e.g., equal to or greater than the critical bumping size C) according to a bumped direction different from the fluid flow direction. 
     At block  2615 , the nanopillar array  320  configured to employ the gap with a gap size less than 300 nanometers in order to sort the biopolymers. 
     The biopolymers include DNA and RNA. The first population of the biopolymers when uncoiled (and/or coiled) is shorter than the second population of biopolymers when uncoiled (and/or coiled). The first population of the biopolymers includes lengths of at least one of 1 kilo base pairs and/or 2 kilo base pairs. 
     The second population of the biopolymers includes lengths of at least 5 kilo base pairs or greater. The second population of the biopolymers includes lengths in a range of about 5 kilo base pairs to about 166 kilo base pairs. 
     The first population and the second population of biopolymers are sorted when coiled and uncoiled through the nanopillar array. 
     The pillar array has the gap size less than 200 nanometers. In another case, the gap has a gap size less than 100 nanometers. In additional cases, the gap size may be less than 80 nm, 60 nm, 50 nm, etc. 
       FIG. 27  is a method  2700  for configuring the cascaded array structure  2500  according to an embodiment. 
     At block  2705 , a first stage (in  FIG. 25 ) is configured to output entities to a second stage according to a fluid flow direction and is configured to bump the entities to the second stage according to a bumped direction. 
     At block  2710 , the second stage is configured to output the entities to a third stage according to the fluid flow direction and is configured to bump the entities to the third stage according to the bumped direction; 
     At block  2715 , the third stage is configured to output the entities according to the fluid flow direction and is configured to bump the entities according to the bumped direction; 
     At block  2720 , separate collection outlets (e.g., for the 1 st  separation group through the 8 th  separation group) are provided in order to collect the entities from the third stage according to sorting through the first stage to the third stage. 
     The cascaded array structure  2500  is configured to yield a specific number of separations (e.g., 2 N ) for the entities based on a number of stages. The fluid flow direction is different from the bumped direction. The bumped direction can be different for different stages. For example, some arrays  320  in the cascaded array structure  2500  may be configured to bump entities to the left while others may bump entities to the right. 
     In this case, the bumped direction is the same for different stages. For example, the arrays  320  in the cascaded array structure  2500  may be configured to generally bump entities to the right, when the entities meet the critical bumping size. However, the pillar array can be designed to bump the biopolymers to either the left or the right, depending on the relative direction of shifted pillars to the flow direction. Therefore, it is possible to combine different pillar arrays designs on the same chip for best chip layout and performance considerations. 
     The entities include biopolymers, such as DNA, RNA, etc. 
     The first stage includes one nanopillar array (e.g., array  2 A), the second stage includes two nanopillar arrays (e.g., arrays  3 A and  3 B), and the third stage includes four nanopillar arrays (e.g., arrays  4 A,  4 B,  4 C,  4 D). The one nanopillar array (e.g., array  2 A) in the first stage outputs to the two nanopillar arrays (arrays  3 A and  3 B) of the second stage. One (e.g., array  3 A) of the two nanopillar arrays outputs to two (e.g., arrays  4 A and  4 B) of the four nanopillar arrays of the third stage and another (e.g., array  3 B) of the two nanopillar arrays outputs to another two (e.g., array  4 C and  4 D) of the four nanopillar arrays. 
     Each of the one nanopillar array, the two nanopillar arrays, and the four nanopillar arrays is configured to output the entities according to the fluid flow direction and/or to bump the entities according to the bumped direction. 
     As discussed in  FIG. 25 , the cascaded array structure  2500  is configured to sorting entities according to an embodiment. The first stage includes a first nanopillar array corresponding to a first critical bumping dimension (e.g., D 2A ). The second stage includes a second nanopillar array and a third nanopillar array, where the second nanopillar has a second bumping dimension (e.g., D 3A ) and the third nanopillar array has a third critical bumping dimension (e.g., D 3B ). 
     The first nanopillar array (e.g., array  2 A) is coupled to the second nanopillar array (e.g., array  3 A) and is configured to output the entities less than the first critical bumping dimension (e.g., less than D 2A ) to the second nanopillar array. The first nanopillar array is coupled to the third nanopillar array (e.g., array  3 B) and is configured to output the entities meeting the first critical bumping dimension (e.g., equal or greater than D 2A ) to the third nanopillar array. The third stage includes a fourth nanopillar array having a fourth critical bumping dimension (D 4A ), a fifth nanopillar array having a fifth critical bumping dimension (D 4B ), a sixth nanopillar array having a sixth critical bumping dimension (D 4C ), and a seventh nanopillar array having a seventh critical bumping dimension (D 4D ). 
     The second nanopillar array (e.g., array  3 A) is configured to output the entities less than the second critical bumping dimension (less than D 3A ) to the fourth nanopillar array (e.g., array  4 A) and is configured to output the entities meeting the second critical bumping dimension to the fifth nanopillar array (e.g., array  4 B). 
     The third nanopillar array (e.g., array  3 B) is configured to output the entities less that the third critical bumping dimension (e.g., less than D 3B ) to the sixth nanopillar array (e.g., array  4 C) and is configured to output the entities meeting the third critical bumping dimension (e.g., equal to or greater than D 3B ) to the seventh nanopillar array (e.g., array  4 D). 
     The nanopillar arrays  320  discussed herein can be utilized to identify/sort biopolymers and/or biomolecules with respect to biopolymer-protein binding. Biomolecules are different types of proteins, such as but not limited to transcription factors, histones, translocases, helicases, polymerases, DNA repair enzymes, nucleases, zinc finger proteins, leucine zipper proteins, helix-turn-helix proteins, topoisomerases, ligases. A biopolymer-protein complex is when a biopolymer such as DNA or RNA binds to a protein, thus resulting in the biopolymer-protein complex. Using the nanopillar array  320 , the can be separation and identification of the unbound proteins (e.g., proteins that travel in a first direction in the array  320  (i.e., zigzag mode)), of unbound biopolymers (e.g., biopolymers that travel in the first direction in the array  320  (i.e., zigzag mode), and of the biopolymer-protein complexes (e.g., the complexes are bumped in the second direction according to the critical angle of the array  320 ). 
     For example, a biopolymer fragment (such as DNA or RNA) of a given size and a mixture of biomolecules (e.g., proteins) can travel through the nanopillar array  320  in a zigzag mode. For example, the biopolymer fragment (unbound) traveling in zigzag mode has not combined with any of the proteins to form a biopolymer-protein complex in the nanopillar array  320 . Similarly, the proteins (unbound) traveling in zigzag mode have not combined with any of the DNA or RNA to form a biopolymer-protein complex in the nanopillar array  320 . 
     However, when one or several biomolecules form a complex with the biopolymer, the resulting biopolymer-protein complex travel down the array in a bumping mode corresponding to the critical angle of the nanopillar array. The resulting complexes are thus isolated/sorted from any unbound proteins and unbound biopolymers, and the components of the complexes can be identified as the biopolymer-protein complexes exit from a different outlet than the unbound proteins and unbound biopolymers. For example, the biopolymer-protein complexes may travel to outlet  940  while the unbound proteins and unbound biopolymers travel to outlet  945 . 
     This method of identifying/sorting biopolymers (DNA, RNA) and biomolecules (proteins) with respect to biopolymer-protein complexes can be used when a biopolymer sequence is known to promote complexing (i.e., binding) with biomolecules: (1) The known biopolymer sequence (DNA and/or RNA known to bind with proteins) is exposed to a mix of biomolecules (e.g., unknown proteins because it is not known which sequence of DNA or RNA that the unknown proteins bind with) in the nanopillar array  320 . (2) The biopolymer-biomolecule complexes are isolated via the sorting process of the nanopillar array  320 . This allows for the unknown biomolecules (proteins) involved in the biopolymer-protein complex formation to be identified because of the sorting. The previously unknown proteins are now known to bind with the known biopolymer sequence (DNA or RNA) because the newly formed biopolymer-protein complex has been bumped in the nanopillar array  320 . 
     Alternatively or additionally, when biomolecules are known to have the ability of complexing (i.e., binding) to biopolymers: (1) The known biomolecules (proteins known to bind with specific sequence of DNA and/or RNA) are exposed to a mix of unknown biopolymer sequences (sequence of DNA or RNA that is not known) in the nanopillar array  320 . (2) The biopolymer-biomolecule complexes are isolated via the sorting process of the nanopillar array  320 . This allows for the exact sequence of the biopolymer responsible for forming the biopolymer-protein complex formation to be identified. The previously unknown DNA sequence (or RNA sequence) is now known to bind with the known protein that binds with a particular DNA sequence (or RNA sequence) because the newly formed biopolymer-protein complex has been bumped in the nanopillar array  320 . 
     These applications are particularly useful to identify gene regulatory complexes, and their binding sequence. Also, these applications are very useful to isolate RNA/biomolecule complexes. 
     A method of separating and identifying unknown biopolymers is provided. Unknown biopolymers and biopolymer-binding proteins are introduced into a nanopillar array. When the unknown biopolymers bind with the biopolymer-binding proteins, biopolymer-protein complexes are formed. When the unknown biopolymers do not bind with the biopolymer-binding proteins, the unknown biopolymers and biopolymer-binding proteins remain unbound. The unknown biopolymers and the biopolymer-binding proteins are sorted through the nanopillar array by transporting any of the unknown biopolymers and the biopolymer-binding proteins which are unbound in a fluid flow direction, as the unknown biopolymers and the biopolymer-binding proteins which are unbound are less than a predetermined bumping size. The unknown biopolymers and the biopolymer-binding proteins are sorted through the nanopillar array by transporting the biopolymer-protein complexes in a bumped direction, as the biopolymer-protein complexes are at least the predetermined bumping size. The nanopillar array is configured to employ the gap with a gap size less than 300 nanometers for sorting. 
     The biopolymers include DNA and RNA and they can form DNA-protein complex (e.g., one type of biopolymer-protein complex) and RNA-protein complex (e.g., another type biopolymer-protein complex) with specific protein molecules (i.e., the biopolymer-binding proteins). 
     The proteins include transcription factors, histones, DNA damage repair proteins, polymerases, helicases, etc. The method can be used to identify unknown biopolymers using proteins that can selectively bind to particular (unknown) biopolymers. 
     A method of separating and identifying unknown proteins is provided. Unknown proteins and protein-binding biopolymers are introduced into a nanopillar array. When the unknown proteins bind with the protein-binding polymers, biopolymer-protein complexes are formed, and when the unknown proteins do not bind with the protein-binding biopolymers, the unknown proteins and biopolymers remain unbound. The unknown proteins and the protein-binding biopolymers are sorted through the nanopillar array by transporting any of the unknown proteins and the protein-binding biopolymers which are unbound in a fluid flow direction, as the unknown proteins and the protein-binding biopolymers which are unbound are less than a predetermined bumping size. The unknown proteins and the protein-binding biopolymers are sorted through the nanopillar array by transporting the biopolymer-protein complexes in a bumped direction, as the biopolymer-protein complexes are at least the predetermined bumping size. The nanopillar array is configured to employ the gap with a gap size less than 300 nanometers for sorting. 
     Deposition is any process that grows, coats, or otherwise transfers a material onto the wafer. Available technologies include, but are not limited to, thermal oxidation, physical vapor deposition (PVD), chemical vapor deposition (CVD), electrochemical deposition (ECD), molecular beam epitaxy (MBE) and more recently, atomic layer deposition (ALD) among others. 
     Removal is any process that removes material from the wafer: examples include etch processes (either wet or dry), and chemical-mechanical planarization (CMP), etc. 
     Patterning is the shaping or altering of deposited materials, and is generally referred to as lithography. For example, in conventional lithography, the wafer is coated with a chemical called a photoresist; then, a machine called a stepper focuses, aligns, and moves a mask, exposing select portions of the wafer below to short wavelength light; the exposed regions are washed away by a developer solution. After etching or other processing, the remaining photoresist is removed. Patterning also includes electron-beam lithography, nanoimprint lithography, and reactive ion etching. 
     The flowchart and block diagrams in the Figures illustrate the architecture, functionality, and operation of possible implementations of systems, methods, and computer program products according to various embodiments of the present invention. In this regard, each block in the flowchart or block diagrams may represent a module, segment, or portion of instructions, which comprises one or more executable instructions for implementing the specified logical function(s). In some alternative implementations, the functions noted in the block may occur out of the order noted in the figures. For example, two blocks shown in succession may, in fact, be executed substantially concurrently, or the blocks may sometimes be executed in the reverse order, depending upon the functionality involved. It will also be noted that each block of the block diagrams and/or flowchart illustration, and combinations of blocks in the block diagrams and/or flowchart illustration, can be implemented by special purpose hardware-based systems that perform the specified functions or acts or carry out combinations of special purpose hardware and computer instructions.