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. Author manuscript; available in PMC: 2023 Dec 21.
Published in final edited form as: Microfluid Nanofluidics. 2022 Nov 15;26(12):104. doi: 10.1007/s10404-022-02610-7

Microfluidic Continuous Flow DNA Fragmentation based on a Vibrating Sharp-tip

Xiaojun Li a, Jing Wang a, Kathrine Curtin b, Peng Li a,*
PMCID: PMC10735211  NIHMSID: NIHMS1909355  PMID: 38130602

Abstract

Fragmentation of DNA into short fragments is of great importance for detecting and studying DNAs. Current microfluidic methods of DNA fragmentation are either inefficient for generating small fragments or rely on microbubbles. Here, we report a DNA fragmentation method in a 3D-printed microfluidic device, which allows efficient continuous flow fragmentation of genomic DNAs without the need for microbubbles. This method is enabled by localized acoustic streaming induced by a single vibrating sharp-tip. Genomic DNAs were fragmented into 700 to 3000 bp fragments with a low power consumption of ~140 mW. The system demonstrated successful fragmentation under a wide range of flow rates from 1 to 50 μL/min without the need for air bubbles. Finally, the utility of the continuous DNA fragmentation method was demonstrated to accelerate the DNA hybridization process for biosensing. Due to the small footprint, continuous flow and bubble-free operation, and high fragmentation efficiency, this method demonstrated great potential for coupling with other functional microfluidic units to achieve an integrated DNA analysis platform.

Introduction

Microfluidics has become an important platform for processing and analyzing nucleic acids owing to its capability of handling small volumes of fluid, integrating multiple functional units and simplifying the overall analysis workflow (Vyawahare et al.2010; Zare and Kim 2010). For many applications involving DNAs, such as next-generation sequencing (NGS), microarray analysis, epigenetic studies, and DNA hybridization, fragmentation is an important step in generating short DNA fragments to ensure efficient amplification and binding (Knierim et.al 2011; Morey et al. 2013).

Typical strategies for fragmenting DNAs include enzymatic digestion, sonication, nebulization, and hydrodynamic shearing. Enzymatic digestion utilizes restriction enzymes that cut the DNAs at specific sites. The process is efficient and can target specific sequences. However, it is not desired for applications requiring fragments without bias (Anderson 1981; Thorstenson et al.1998; Kasoji et al. 2015; Roe 2004; Seed et al. 1982; Vaxelarie 1994; Sarokin et al. 1984; Wong et al. 1997). Physical fragmentation methods, including sonication, nebulization, and hydrodynamic shearing, generate fragments that are not dependent on specific sequences. Nebulization cuts DNA into small pieces through the mechanical shear force generated from forcing DNA through the orifice of a nebulizer (Deininger 1983; Burger et al.2007). This process is straightforward to perform, but it is not compatible with microfluidic systems (Lentz et al.2005). Hydrodynamic shearing breaks DNAs by creating a sudden change of flow cross-section to exert excess shear stress on the DNAs (Paul 1997; Davison 1959). Sonication-based fragmentation is based on the ultrasonic cavitation effect (Oefner 1996; Schoppee and Wamhoff 2011). When the small gas bubbles implode, the local high temperature and pressure field lead to the generation of DNA fragments.

Both hydrodynamic shearing and sonication have been applied in microfluidic systems. Berg et al. utilized the extensional strain forces generated by acceleration of the DNA solution near constriction entrances for breaking DNA into fragments with a sample consumption of 10 μL and short fragmentation time (a few minutes). However, the size of the fragments is limited to 6–10 kbp with a pressure requirement from 1–10 bar, and the maximum flow rate allowed by this system is 18 μL/min (Shui et al. 2011). Sonication-based fragmentation methods have been reported to generate much smaller fragments. Merten et al. utilized the strong acoustic energy generated from piezoelectric transducer coupled with cavitation bubbles to shear genomic DNA into fragments (Tseng et al. 2012). Lu and coworkers fabricated crescent-shaped structures in a microfluidic chamber to enhance cavitation and acoustic streaming (Cao and Lu, 2016). This method achieved fragments smaller than 500 bp, which is optimal for chromatin immunoprecipitation (ChIP) assay. Recently, Sun et al. reported bubble enhanced ultrasonic DNA fragmentation under stop flow conditions, which generated 300 bp λDNA fragment in 30 min (Sun et al. 2022). While sonication-based methods can generate very small fragments, they generate microbubbles inside microchannels, which is detrimental to many microfluidic applications (Lochovsky et al. 2012, He et al. 2021, Kang et al. 2008, Liu et al. 2011). In addition, existing sonication methods are not performed under continuous flow. Therefore, they have not been employed in integrated microdevices with multiple functional units. To better integrate microfluidic DNA fragmentation with downstream analysis, there is still a need for a continuous flow and bubble-free microfluidic fragmentation method with good fragmentation efficiency.

Here, we report a microfluidic DNA fragmentation method based on a single vibrating sharp-tip (Fig. 1). The present method leverages the strong acoustic streaming generated around a vibrating sharp-tip to shear the DNAs into small fragments. A vibrating sharp-tip is an efficient way of generating acoustic streaming, achieving orders of magnitude higher streaming velocity than conventional boundary-driven acoustic streaming. It has been reported for a wide range of applications, including fluid mixing, pumping, cell lysis, particle trapping, microfluidic ELISA, and enzyme kinetics measurement (Po-Hsun et al. 2015; Zhen et al. 2021; Po-Hsun et al.2018; Po-Hsun et al. 2015; Xioajun et al. 2021; Xiaojun et al. 2019). However, it has not been demonstrated as an effective method for DNA fragmentation to date. In this work, we developed a 3D printed microfluidic device to anchor a pulled glass capillary for acoustic streaming. Strong acoustic streaming is generated in the microfluidic channel by vibrating the pulled glass capillary with a piezoelectric transducer. When genomic DNA is continuously flowed through the fragmentation chamber, the DNA is fragmented by the strong acoustic streaming. Genomic DNAs were efficiently fragmented to a size range of 700 to 3000 bp without the need of generating any microbubbles. The present method demonstrated successful fragmentation of DNA samples under a continuous flow with flow rates up to 50 μL/min. The small fragment size, bubble-free, and continuous flow operation, and small footprint of this method make it a promising method to be coupled with other functional units for integrated microfluidic DNA analysis.

Figure 1.

Figure 1.

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Schematic of the vibrating sharp-tip DNA fragmentation platform. (a) A Pulled-tip glass capillary is inserted into a 3D printed microfluidic channel through a side anchoring channel. The sharp-tip is vibrated by applying radio frequency signals to the piezoelectric transducer. The DNA fragmentation occurs in the flow chamber. (b) A photograph representation of the assembled DNA fragmentation device.

Materials and Methods

Reagents

Fluoresbrite plain YG 10.0-micron microspheres were purchased from PolyScience (Warrington, PA, US). Tris-borate-EDTA (TBE) 10X solution, agarose (BP1356-100), and Isopropanol (IPA) were purchased from Fisher Scientific (Pittsburgh, PA, USA). Poly (ethylene glycol) diacrylate (PEGDA, MW 250), phenylbis (2,4,6-trimethylbenzoyl) phosphine oxide (Irgacure 819), human male genomic DNA were purchased from Sigma–Aldrich (St. Louis, MO, USA). 2-nitrophenyl phenyl sulfide (NPS) was purchased from TCI (Tokyo, Japan). Ultra-Ever Dry super-hydrophobic coating was purchased from Ultratech. Water was purified using a Millipore purification system (Bedford, MA, USA). Generuler 1kb plus DNA ladder was purchased from Thermo Fisher Scientific. Customized DNA sequences were purchased from IDT, Indiana, USA.

Device design and fabrication

To fabricate the microfluidic chip, 3D structures were first designed using SolidWorks. Then, the device was printed using an Asiga Pico2 HD 3D printer with an X-Y plane resolution (pixel size) of 37 μm and a Z-axis control of 1 μm. The light source wavelength is 385 nm. A mixed PEGDA with 0.5% (w/w) Irgacure 819 and 0.5% (w/w) NPS were prepared and stirred for 30 min as the printing resin. An absorber NPS was used to control the penetration depth of the incident light and print the microchannels inside the chip (Paquin et al. 2015). After printing was complete, the device was removed immediately from the printing tray and washed by IPA for dissolving unpolymerized material inside the channels. After a post-exposure process under UV (365 nm) for 5 min, the device was glued onto a glass slide (VWR, Radnor, PA, USA) using epoxy glue (5-minute epoxy, Devcon) for enhancing the transparency of the device. The device design includes two inlets for reagent infusion, a side-channel for glass capillary insertion, and a main chamber with a dimension of 5×6×0.5 mm for DNA fragmentation activation.

The acoustic-activated glass capillary was fabricated with a piezoelectric transducer (7BB-27–4L0, Murata, Kyoto, Japan) and a pulled-tip glass capillary fixed to a glass slide. The pulled-tip glass capillary can be made by pulling capillary tubes (Drummond Scientific, Broomall, PA, USA) using a laser-based micropipette puller (P-2000, Sutter Instrument, Novato, CA, USA) or purchased from Tritech Research for customized tip size requirement. To maximize the streaming efficiency, the pulled-tip glass capillary was fixed on the edge of one end of the glass slide using glass glue (Loctite, Rocky Hill, CT, USA) with a 30-degree angle between the capillary and the shorter side of the glass slide with a distance to the corner ~5 mm based on our previous studies (Ranganathan et al. 2019, Xiaojun et al. 2021). The capillary used in this work has a total length of 50 mm with an I.D. of 0.5 mm and O.D. of 0.7 mm. The I.D. of the capillary tip is ~20 μm, and the O.D. is ~25 μm. The other end of the glass slide was attached with a piezoelectric transducer using epoxy glue.

To measure the streaming velocity, fluorescent particles (10.2 μm) in water were loaded to a microfluidic chamber with a vibrating sharp-tip capillary. The particle movement was then recorded using an inverted fluorescence microscope with a sCMOS camera. Since the streaming patter around the vibrating capillary tip is double vortex, we analyzed the velocity of particles ~200 μm above the tip end. Based on the recorded videos, the streaming velocity (ν) can be calculated from the particle displacement (L) between frame and the frame time (t).

A two-step superhydrophobic coating on the side channel’s inner surface and the glass capillary’s outer surface was applied to avoid any liquid leakage from the side channel. The coating process is done by loading the reagents to the side channel wall and glass capillary with a micropipette. After coating, the glass capillary was carefully inserted into the side channel. A Tektronix function generator (AFG1062) connected with an amplifier (LZY-22+, Mini-Circuits) was used to drive the piezoelectric transducer for acoustic signal generation.

DNA sample preparation, collection, and gel electrophoresis

The human male genomic DNA was diluted in TBE buffer and loaded to the microfluidic device inlet by a syringe pump (Fusion 200, Chemyx Inc., Stafford, TX) for further detection. The acoustic signal generator was turned on to initiate DNA fragmentation. After the sample fragmentation process, a fixed volume of the sample determined by the gel electrophoresis testing requirement was collected from the outlet of the device by using a pipette and analyzed using 1% agarose gel electrophoresis. Gel electrophoresis was conducted using a MyGel Mini Electrophoresis System from Accuris Instruments, and the DNA samples were separated under 50 V for 1 hr. The result was read by the ChemiDoc MP system. The size of the bands was determined using 75 – 20,000 bp DNA ladders.

Characterization of DNA fragmentation performance

A syringe pump was used to infuse the human male genomic DNA sample through the device’s inlet. The acoustic streaming at the sharp-tip was generated by the vibration of the glass slide by piezoelectric transducer. To characterize the influence of different input voltages on the fragmentation performance, a series voltage of 4.00, 5.33, 6.67, 8.00, 9.33, 10.67, and 12.00 Vpp were applied to the system with a flow rate of 1 μL/min and 10 μL/min. With the acoustic signal supplied from the generator, the fragmentation process began along with the acoustic streaming generated from vibrating sharp-tip. After the fragmentation process finished, the fragmented samples were collected using a pipette for further gel electrophoresis detection.

To test the minimum voltage requirement for different flow rates ranging from 2 μL/min to 50 μL/min, the DNA samples were fragmented and collected under input voltages ranging from 6.67 Vpp to 24 Vpp for each flow rate. Then, the fragments for each flow rate and voltage were electrophoresed, and the voltage achieving the fragment size below 1000 bp was chosen as the minimum voltage requirement for each flow rate.

DNA hybridization experiments

Cy5 labeled human MTHFR gene sequence was used as the detection probe (5’-/5cy3/AAA AAA AAA AAA AAA AAA AAT GAT GAA ATC GGCT-3’) and was prepared at a series of concentrations of 1, 1.3, 2, 4, 100 nM. Biotin-labeled complementary DNA sequence (5’-/5cy3/CCC GCA GAC ACC TTC TCC TTC-3’) was used to capture DNA. After the fragmented DNA samples were prepared, capture and detection probes were added to the solution of target DNA. After a 10 min denaturation step at 95 °C, the hybridization of the probes to the DNA target was achieved under gentle agitation at 55 °C, and hybridization time was respectively controlled at 5, 10, 20, and 30 min. Finally, streptavidin-coated polystyrene magnetic beads (2.9 μm, Spherotech Inc., Lake Forest, IL) were added to the solution under gentle agitation for 30 min at room temperature. Thus, the unbound DNAs were washed out using a magnet. The hybridized DNAs on the magnetic beads were detected and analyzed using an Olympus IX73 fluorescence microscope.

Results and Discussion

DNA fragmentation by a vibrating sharp-tip

Existing studies have shown that vibrating sharp-edges can generate strong acoustic streaming with high energy efficiency (Doinikov et al. 2020, Zhang et al. 2020, Nama et al. 2016). Ovchinnikov et al. performed numerical simulation of a vibrating sharp tip using perturbation theory, which showed good agreement with experiments (Ovchinnikov et al. 2014). The strong streaming could be a result of the centrifugal force induced by the high-speed oscillation of the sharp tip. We hypothesized that the strong acoustic streaming induced by a vibrating sharp-tip could be utilized to break DNA into small fragments in microfluidic channels. With the rapid development of 3D printing technologies, using a 3D printer to fabricate microfluidic devices has become increasingly accessible to laboratories that do not have the expertise or resources to perform conventional microfabrication procedures. Therefore, in this work, we aimed to achieve DNA fragmentation using 3D printed microdevices for better adoption of this method by the broader community. Conventional microfabricated sharp-edge structures cannot be fabricated with current 3D printing methods due to the limitation of printing resolution. Here, we combined a pulled capillary with a 3D printed microdevice to generate strong acoustic streaming (Fig 1a). The pulled capillary was glued onto a glass slide on which a piezoelectric transducer was attached. The pulled capillary device was coupled to the main microfluidic channel via a 3D printed side anchor channel. The inner wall of the side channel and the pulled capillary’s outer surface were coated with a superhydrophobic coating to prevent fluid leakage from the main channel.

The streaming pattern generated by the vibrating sharp-tip was studied using 10 μm diameter fluorescent beads (Fig 2a). After loading the fluorescent beads into the main fluidic channel, an RF signal of 95 kHz and 2.67 Vpp was applied to the piezoelectric transducer to vibrate the pulled capillary with a tip OD ~ 20 μm. As shown in Fig 2b, strong acoustic streaming can be generated with two symmetrical counter-rotating vortices. The streaming velocity increases as the increase of input power. To test if the acoustic streaming induced by the vibrating sharp-tip can cause DNA fragmentation, 100 nM genomic DNA solution was introduced into the microfluidic channel at a flow rate of 1 μL/min (Fig 3a). The input power was set at 6.67 Vpp. The collected DNA fragment solution was examined using gel electrophoresis. As shown in Fig 3b, after passing through the acoustic streaming chamber, the genomic DNAs were cut into small pieces ranging from 700–5000 bp, demonstrating the successful fragmentation by the vibrating sharp-tip induced acoustic streaming. When the long-chain DNAs entered the vortex region, they were exposed to excessive hydrodynamic shear because of the acceleration of the DNA molecule in the vortex. Due to the molecule stretching from this extensional strain forces created by the acceleration around the streaming region, the DNA could break into fragments.

Figure 2.

Figure 2.

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(a) An illustration of acoustic streaming vortex with inserted sharp-tip. (b) Generation of acoustic streaming by the vibrating sharp-tip in a 3D-printed microchamber. Streaming was generated under a frequency of 95 kHz with voltage of 2.67 Vpp. Fluorescent microbeads (10 μm) are used for tracing the streaming patterns. Ɩ represented the length of the microbeads movement along with streaming trace at different streaming layers in a specific time period. The detection window for measuring streaming velocity is labeled by the red box.

Figure 3.

Figure 3.

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(a) An illustration of the DNA fragmentation workflow. DNA sample was loaded from top inlet and collected at bottom outlet after passing through the acoustic fragmentation chamber. (b) Gel electrophoresis of DNA fragments after fragmentation under flow rate of 1 μL/min, voltage of 6.67 Vpp. A fragments distribution at 700–5000 bp was observed after fragmentation process.

Notably, the present method achieved effective DNA fragmentation without the presence of microbubbles or the cavitation effect, whereas existing sonication-based methods all rely on microbubbles. We further examined if microbubbles would affect the performance of the present device. We prepared two identical microdevices (device a and device b) with the same device dimension, geometry, and sharp-tip size. The only difference between the two devices is whether the bubbles are present or not. For device a, the DNAs were fragmented under different voltage inputs under a flow rate of 1 μL/min without bubbles present. For device b, air bubbles were induced to the microchamber before the experiment using a pipet. Two separate bubbles of different sizes were induced at two corners of the chamber. The bubbles’ radius was about 1.5 mm and 0.5 mm, respectively and were introduced at the upper left and lower right corners of the chamber. Three input voltages of 2.67, 6.67, and 13.33 Vpp were selected to be applied to each device for distinguishing the effects of bubbles and no bubbles on the system. After the same experiment duration, the samples were collected in the same volume for electrophoresis detection. The intensity of large fragments (~5000 bp) under low voltages (2.67, and 6.67 Vpp) for the bubble free device is slightly higher than the intensity generated by the device with bubbles (Fig. 4). This difference may be caused by the enhanced shearing effect with the presence of bubble in the device. For the high voltage group (13.33 Vpp), the fragmentation results from the two devices were similar regardless of the presence of bubbles. This result confirmed that the performance of the vibrating sharp tip without any bubbles, and indicated that the presence of bubbles may further enhance the fragmentation efficiency. However, introducing gas bubbles is highly undesirable for many microfluidics applications, especially when coupled with additional functional units. Compared with existing sonication methods, the vibrating sharp-tip is more advantageous for an integrated microfluidic system as it allows bubble-free operation with similar performance.

Figure 4.

Figure 4.

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Influence of bubbles on DNA fragmentation working mechanism. Gel electrophoresis of DNA fragments after fragmentation without bubbles (device a in the left) and with bubble (device b in the right) induced to the microchamber. The voltage applied to the fragmentation system are 2.67, 6.67, and 13.33 Vpp for both of the devices with/without bubbles.

The relationship between input power voltage and the size of DNA fragments

Upon demonstrating that the acoustic streaming effectively fragments the genomic DNA, we explored the influence of input voltage on the fragmentation size distribution. As shown in Fig 5, the size of fragments changed as the input voltage increases, indicating that the present method could offer some tunability of the fragment size. The direct result of increasing the input voltage is the increased streaming velocity, which is expected to increase the hydrodynamic shear for DNA fragmentation. First, we studied the relationship between input voltage and the streaming velocity. 10 μm fluorescent particles were used to visualize the acoustic streaming. Videos were taken using an inverted microscope with a frame rate of 30 fps. The streaming velocity was then determined by measuring the displacement of the fluorescent particles between frames. For different input voltages, the particles were all selected at a fixed location. As shown in Fig 5a, the streaming velocity increased as the increase of input voltage from 4.00 to 10.67 Vpp. Within the full voltage range, the relationship between the streaming velocity is not linear. At low voltages, the increase of streaming velocity with respect the increase of voltage was slow. For voltages above 6.67 Vpp, a near linear relationship was observed. Based on this result, we further examined DNA fragment size under various input voltages using gel electrophoresis (Fig 5b–c). In this experiment, DNA samples were introduced the vibrating sharp-tip under two different flow rates of 1 μL/min and 10 μL/min, respectively. The input voltage was applied from 2.94 Vpp to 12.00 Vpp. Under the flow rate of 1 μL/min, a clear decrease of fragment size as the increase of voltage was observed. At 2.94 Vpp, the size range of fragments is from 2000 to 7000 bp, while some original genome DNAs can still be seen. As the voltage increased to 6.67 Vpp, the size range shifted to 1000 to 5000 bp, and all the genome DNAs were fragmented. Further increasing the voltage to 10.67 Vpp lead to the smallest fragment sizes from 700 to 3000 bp. Increasing the flow rate to 10 μL/min lead to the decrease of fragmentation efficiency. This is expected as the convective flow in the main fluidic channel suppresses the acoustic streaming. With same voltage at 10.67 Vpp, the fragment size was observed 700 – 3000 bp for 1 μL/min and 1500–5000 bp for 10 μL/min, respectively. The typical size range for the present device is from 700 to 5000 bp. This range is particularly important for PCR based or direct DNA analysis methods. In PCR-based DNA specificity studies, the primers locate target DNA regions at the fragment distributions of 100–2500 bp, thus the fragments generated from this report are applicable for these primer sizes. For example, Leishmania kDNA used for monitoring new leishmaniasis treatments are 700 bp (Ravel et al. 1995); new primers N524 used for 12S ribosomal DNA sequencing are 1000 bp (Goebel et al. 1999); and single ALU primer for the amplification of sequences range from 150 – 3000 bp (Aslanidis and Jong 1990). In addition, the size tunability of this method also facilitates the integration of this method with different sequencing technologies. For sequencing methods with short read length, high input voltage is preferred to generate smaller fragments. While low voltage will lead to large fragments, which can meet the need of emerging long read length sequencing platforms (e.g., nanopore sequencing).

Figure 5.

Figure 5.

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Fragment size characterization in DNA fragmentation performance based on input voltage. (a) Relationship between acoustic streaming velocity and input voltage induced to the fragmentation system. (b) Gel electrophoresis for fragment size difference with voltage range 2.94 – 9.33 Vpp under flow rate of 1 μL/min. (c) Gel electrophoresis for fragment size difference with voltage range 2.94 – 12.00 Vpp under flow rate of 10 μL/min.

Study of flow rates for a single vibrating sharp-tip device

As shown in Figure 5 b and c, the DNA sample flow rate affects the fragmentation performance as the main sample flow suppresses the acoustic streaming. Practically, higher flow rates are preferred for high throughput applications, whereas low flow rates are for samples with limited volume. Here, we studied the range of working flow rates for the present method. Generally, a higher DNA flow rate requires an increase in voltage to achieve the same fragmentation effect. Therefore, we expected an upper limit flow rate for the present system before the breakdown of the piezoelectric transducer and/or the generation of gas bubbles. Fig 6 shows the relationship between the DNA sample flow rate and minimum input voltage. The minimum input voltage is defined as the voltage that can generate DNA fragments below 1000 bp (Fig S1). This fragment length was selected because a wide range of applications, including DNA hybridization and sequencing, are suitable. As the flow rate increased from 2 μL/min to 50 μL/min, the minimum required voltage increased from 6.67 Vpp to 21.33 Vpp. For flow rates higher than 50 μL/min, higher input voltages are needed. However, as the voltage approached the limit of the transducer (30 V), the system became unstable for long-term operation. Therefore, the highest throughput for the present device is 50 μL/min. For applications that require even higher throughput, employing multiple sharp-tips may be necessary.

Figure 6.

Figure 6.

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Relationship between the minimum required voltage and the flow rates for acoustic-based DNA fragmentation.

DNA hybridization performance

Finally, we studied if DNAs fragmented by the present method can be used to improve the DNA hybridization experiments. We selected a common gene in the human genome, MTHFR, as the target. A Cy5 labeled detection probe and biotin-labeled capture probe were designed based on Storhoff et al.’s report (Storhoff et al. 2014). After denaturation, the detection probe and capture probe were first mixed with fragmented DNA samples, and then captured by streptavidin coated magnetic beads from fluorescence detection (Fig. 7a).

Figure 7.

Figure 7.

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DNA hybridization experiments. (a) Schematic of the DNA hybridization assay. (b) Comparison of hybridization fluorescent intensity between fragmented and non-fragmented DNA samples (2 nM) at 5 min and 20 min, respectively. (c) The relationship between fluorescent intensity fragmented DNA sample concentration with a hybridization time of 20 min. Error bars represent the standard deviation of 3 replicate trials.

We compared the performance of hybridization assay between DNA sample fragmented at 1 μL/min and the intact DNAs. Compared with the intact genome, fragmented DNAs should allow faster hybridization processes due to their smaller size. We first studied the detection of MTHFR gene in the 2 nM DNA sample at 5 min and 20 min. The fluorescence intensity on magnetic beads was measured as the signal of detection probes captured. By recording the fluorescence intensity on the microbeads using a sCMOS camera, we compared the performance of hybridization with target DNA at two time points (Fig. 7b). At 5 min, the fragmented sample group already showed a significant increase in signal intensity compared to the negative control group (p < 0.05), indicating that the MTHFR gene in 2 nM DNA sample can be detected at 5 min after fragmentation. In contrast, for the intact DNA, no significant difference in signal intensity is seem between the sample group and the negative control group at 5 minutes. At 20 min, both the fragmented DNA and intact DNA samples showed increased intensity, but the signal intensity for the fragmented group is still significantly higher than the intact DNA group. After 30 min of incubation, the hybridization efficiency for the intact DNA group is ~22% of the fragmentation group based on the fluorescence signals detected from the beads. These results indicate that DNA fragmentation facilitates the detection of positive signals faster than intact DNA.

We further examined whether the signal intensity for DNA fragments is concentration-dependent (Fig. 7c). We prepared a series of DNA concentrations including 1, 5, 20, and 50 nM. In the range of 1 nM to 50 nM, the relationship between the concentration and the fluorescence intensity is linear (R2 = 0.9888). We also tested concentrations higher than 50 nM. We observed that the fluorescence intensity started to reach a plateau, which could be related to the saturation of the beads. Collectively, the utility of the present DNA fragmentation method for DNA hybridization experiments has been demonstrated. The time for detecting a hybridization signal greatly was shortened greatly compared to intact DNA samples and the fragmentation method is concentration dependent.

Conclusion

The fragmentation of DNA is a critical step for many biological applications such as next generation sequencing and microarray analysis. We have designed and fabricated an efficient acoustic streaming-based method for DNA fragmentation inside a micro channel. The hydrodynamic shearing force for efficient DNA fragmentation was generated from acoustic streaming induced by the vibrating sharp-tip protruding in the flow chamber. The size distribution of the DNA fragments can be easily controlled by adjusting the input voltage under continuous flow, where the DNA fragments decrease in size with the increase of input voltage. The microdevice is the first to achieve DNA fragmentation in 3D printed microfluidics with simple fabrication and bubble-free operation. Compared to the existing microfluidic DNA fragmentation methods, this system is more amenable to be integrated with other functional units of a microfluidic system. We successfully demonstrated the DNA fragmentation method can be coupled with DNA capture, which shows that fragments generated can be used in downstream analysis. Future work will focus on the integration of this fragmentation method with on-chip DNA analysis units in 3D printed microdevices.

Supplementary Material

Supporting Information

Acknowledgement

This work was supported in part by National Institute of Health (R01GM135432) and National Science Foundation (ECCS-2144216). KC acknowledges the support from National Science Foundation (DGE1102689).

Footnotes

Conflicts of interests

The authors declare no conflicts of interests that are relevant to the content of this article.

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Supporting Information
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