Rapid Customization of 3D Integrated Microfluidic Chips via Modular

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Rapid customization of 3D integrated microfluidic chips via modular structure-based design Jingjiang Qiu, Qing Gao, Haiming Zhao, Jianzhong Fu, and Yong He ACS Biomater. Sci. Eng., Just Accepted Manuscript • DOI: 10.1021/acsbiomaterials.7b00401 • Publication Date (Web): 02 Sep 2017 Downloaded from http://pubs.acs.org on September 5, 2017

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ACS Biomaterials Science & Engineering

Rapid customization of 3D integrated microfluidic chips via modular structure-based design Jingjiang Qiua,b, Qing Gaoa,b, Haiming Zhaoa,b, Jianzhong Fua,b, Yong Hea,b, ‡ a. State Key Laboratory of Fluid Power and Mechatronic Systems, School of Mechanical Engineering, Zhejiang University, Hangzhou 310027, China b. Key Laboratory of 3D Printing Process and Equipment of Zhejiang Province, School of Mechanical Engineering, Zhejiang University, Hangzhou 310027, China

Corresponding Author: Yong He ‡E-mail: [email protected].

ACS Paragon Plus Environment


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ABSTRACT: In recent years, 3D integrated microfluidic systems have become increasingly more popular because of their ability to incorporate multifunctional components, including porous membranes and biological scaffolds. Due to limitations in resolution, fabrication efficiency and materials, it is hard to develop complex integrated microfluidic systems with low cost and high efficiency. In this paper, we present a novel method that utilizes modular structure-based design, which could greatly reduce the time and cost for customization of complete integrated chips, compared to traditional techniques. By printing sacrificial patterns on the substrate using the 3D printing approach and subsequently covering them with PDMS pre-polymer, PDMS slices with modular structures were obtained, each with specific functions. By combining different PDMS slices with specific modular structures and other functional components, such as membranes and scaffolds, the conceptual design was efficiently converted into complete integrated microfluidic chips. As proof-of-concept, customized 3D microfluidic chips were generated and successfully used for cell culture and biological analysis. Furthermore, the flexible combination with biofabrication of hydrogel beads was also presented, revealing the potential use of this technique in the fabrication of organ-on-a-chip. KEYWORDS: modular structure, rapid customization, 3D printing, microfluidic chip

1. INTRODUCTION

In recent years, the study of microfluidic technologies has become an important aspect of many research fields, including chemistry, medicine and bioengineering.1-3 As the merits of microfluidics are being highlighted in research, considerable interest is sparked by the development of microfluidic devices that enable integrated functional features, which could potentially solve problems in


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chemical or biological research. Available studies report that integrated microfluidic chips often contain necessary functional components used for magnetic, electronic, chemical and biological detection, which can support comprehensive on-chip analysis.4 The electronic part is essential for the microfluidic electronic chemical biosensing area5 and was successfully integrated into various analysis systems, such as automated cancer markers detection microsystem6 and flexible polyethylene terephthalate (PET) microfluidic systems,7 for cell culture and disease diagnostics. Furthermore, to better understand the mechanism of cell co-culture, cell metabolic activity, cell–cell interactions and drug metabolism, membrane-based microfluidic devices were introduced by integrating different semipermeable membranes and microfluidic channels.8-14 The typical fabrication methods of integrated microfluidic systems, which include soft lithography, hot embossing and femtosecond laser writing, are time-consuming and often expensive, limiting their applications. Recently, 3D printing has been suggested for the production of integrated microfluidic systems as it is automatic, low-cost and has relatively high throughput.15, 16 The fused deposition modeling (FDM) technique was described to obtain microfluidic devices integrating polylactic acid (PLA) microchannels and other materials. Using FDM, Gaal et al. successfully built a 3D-printed electronic tongue (e-tongue),17 while Tsuda et al. developed millifluidic devices with droplet generator and 3D-printed valves.18 Another 3D printing technique, known as stereolithography (SLA), was utilized to produce microfluidic systems featuring directly embedded optical fibers.19 Corbel et al. investigated the photocatalytic degradation of salicylic acid (SA) using a microfluidic reactor that was manufactured by SLA in epoxy resin.20 By incorporating a porous polycarbonate membrane above channels, a 3D-printed fluidic device was produced for studying drug transport, with a commercially available 3D printer.21 Moreover, bioprinting can be used to



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fabricate tissue constructs within microfluidic systems and form tumor-on-a-chip platforms that can mimic physiological, mechanical and chemical cues, which is very appealing for cancer research.22 Microfluidic chips can be obtained either by direct 3D printing, or by using 3D-printed templates covered with polydimethylsiloxane (PDMS), which is then peeled off in slices from masters, and finally sealed into chips .23,

24

This method takes advantage of the merits of PDMS. However,

3D-printed masters require complex and time-consuming post-treatment, due to the impact of the curing reaction of PDMS on the printed master material. In terms of resolution, accuracy and repeatability, 3D printing is very promising for the rapid customization of integrated microfluidic chips, but limitations posed by materials, hardware and cost should still be taken into consideration and continue to be improved, when applied in biological research.25, 26 Another important problem is that 3D printing usually generates monolithic microfluidic systems. To avoid this, all functional components should compromise to form the system, leading to extra optimization for the functionality and integration of individual components. Furthermore, changes in the operation surroundings or poor design can affect the performance of microfluidic systems, resulting in unnecessary costs. The modular approach, which involves assembling functional and modular elements into complete systems, can act as a low-cost and efficient method of fabrication, allowing the rapid customization of complex microfluidic systems. When using this method, any extra specific functional component can be added to the system, without influencing its overall performance. One option is using PDMS, or thermoplastic modules made with conventional methods, to customize the microfluidic architecture and form a multiple functional system for chemical synthesis27 and biochemical analysis.28-30 Organ-on-a-chip systems could also be realized to monitor organ behaviors in situ, using a modular design with integrated multi-sensors.31 Another fast


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and convenient way is utilizing different 3D-printed modules to build either an integrated “plug-n-play” modular microfluidic system32 or assembled with metal pins to build an entire integrated device for biosensing.33 Moreover, a “stick-n-play” modular microfluidic system, presented by Yuen, could be easily reconfigured for various integrated microfluidic systems, using magnetic interconnects to stick different modules.34 A more effective method connected high-precision SLA-printed discrete elements in a self-aligned manner and reduced the complexity of the 3D microfluidic system by applying the circuit theory.35 In addition, the microfluidic building blocks were attained via PDMS casting with 3D-printed master molds, thus assembling into a Lego®-like modular microfluidic platform.36 Although combining the 3D printing technique with the modular approach is promising, efforts still need to be devoted to the development of suitable materials, such as resins and plastic filaments. Available materials are not as water-impermeable, biocompatible, transparent, gas-permeable and elastomeric as PDMS. Furthermore, the printing resolution, accuracy, surface roughness and biocompatibility partially limit the application of 3D printing in microfluidics for biological research. In this paper, we propose a new method for the rapid customization of integrated microfluidic systems, with PDMS slices possessing specific modular structures, generated using the 3D-printed sacrificial template obtained via the 3D printing approach previously reported by our group.37 The 3D modular structures were directly fabricated on the substrate and did not require any post-treatment. Therefore, the PDMS slices with specific structures were directly obtained after PDMS was cured, which is fast and efficient. Furthermore, smooth channel surface and high resolution could be guaranteed with this method. In our method, the use of PDMS generated chips with high biocompatibility, whereas the 3D printing contributed to the rapid customization and low



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cost of the microfluidic chips. Through this approach, PDMS slices with modular structures were produced in less than one hour, and could be assembled with electronic circuits, membranes or biological scaffolds in approximately 10 minutes, using the adhesive-free packaging method. Compared to other methods, PDMS slices could be reused and rapidly fabricated, allowing for rapid optimal design and customization of complex integrated microfluidic devices, with relatively low cost. Combined with the bioprinting technique, this method also showed its potential for integration of microfluidic chips and bioprinted hydrogel constructs, which is very promising in fabricating organ-on-a-chip.

2. MATERIALS and METHODS

2.1 Materials and reagents

Materials used for the device fabrication: PDMS (Sylgard 184, Dow Corning, USA), maltitol (Aladdin Industrial Corporation, China) and PET porous membrane (pore size: 0.4 µm, transwell insert, Corning Inc, USA). Reagents for cell culture: cell culture medium contained Minimal Essential Medium (MEM; GIBCO, USA) supplemented with 10% fetal bovine serum (FBS) (Sijiqing, China). The matrigel matrix was purchased from Corning (USA). The Live/Dead assay kit was obtained from Key-GEN BioTECH Co., Ltd (Calcein-AM/PI, China). Sodium alginate (Na-Alg) and

calcium

chloride

4',6-diamidino-2-phenylindole

(CaCl2)

were

dihydrochloride

purchased

from

(DAPI)

(TRITC)-phalloidin were purchased from Yeasen Co., Ltd (China).


and

Sigma-Aldrich

(USA).

tetramethylrhodamine

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2.2 Method of fabrication

Several steps were involved in customizing the 3D integrated microfluidic system (Figure 1). Before printing, each modular pattern was designed with computer aided design (CAD) software (Solidworks, Dassault Systems SolidWorks Corp). All the CAD drawings of modular patterns were then exported as STL files and further processed by 3D printing software, following the usual procedure of 3D printing. Fused maltitol was directly printed on the smooth substrate (glass or polymer) by the 3D printer (Figure S1) and the movement of the printhead followed the designed path, resulting from the processed CAD designs. The printing process for each PDMS slice lasted only 5 minutes. After that, the printed maltitol patterns were prepared for the next treatment. First, PDMS was prepared by mixing the base and curing agents at 10:1 (w/w) ratio and then placed in a vacuum drying oven for degassing. After obtaining the printed maltitol patterns, rectangular molds cut from acrylic (PMMA) plates were stacked on the smooth substrate, to precisely control the shape and layer height of PDMS slices. PDMS was then poured on the printed modular structures and excess PDMS was removed. Subsequently, PDMS was cured in an oven at 85°C, for 25 minutes. Next, the PDMS slices with modular structures were obtained by slightly peeling off cured PDMS slices from the substrate. For this, a sharp scalpel was first used to cut edges which could separate the PDMS slices from the acrylic mold walls. Secondly, the PDMS was gradually lifted from the substrate. As the printed maltitol patterns are firmly attached to the substrate, it was then easy to separate PDMS from both maltitol patterns and substrate. The printed maltitol patterns were left on the substrate and could be reused if undamaged. A second option, which allows for the direct removal of PDMS slices, is placing the slices containing maltitol patterns in boiling water (10 seconds at 100°C) and then drying them. This could result in a smoother microchannel



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surface when compared with the procedure where the cured PDMS slices were just directly peeled off from the maltitol pattern. However, in this case, the acrylic mold is removed, then both PDMS and maltitol structures are directly detached from the substrate using an engraving knife. Consequently, the printed maltitol patterns can only be used once.

Figure 1. Schematic of fabrication process: Sugar filaments are printed on the substrate according to the pattern, then PDMS is poured and cured. Complete PDMS slices with modular structures can be obtained by peeling off the cured PDMS. Using several PDMS slices with different functions and other functional components, a customized 3D microfluidic chip is rapidly assembled. After final test, the chip can be reused several times.

After obtaining the chip slices via modular structure-based design, several different PDMS slices and other modular components (membrane or electronic circuits) were assembled into different functional microfluidic chips, according to the design of the integrated system. The various components used for device assembly are shown in Figure 1. Custom-made PMMA frame and 3D-printed PLA frame form the main frame of the chip, while the PDMS slices become the core part


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of the device. Other components include screws, metal tubes and punched holes. PDMS slices are punched to generate fluidic inlets and outlets, using the Harris Uni-Core (0.5 mm). The device components were thoroughly cleaned and then rapidly assembled with adhesive-free packaging technology. M2×16 mm screws were first inserted into the bottom PMMA frame. PDMS slices and the PLA frame were then placed on top of the PMMA frame. PLA frame and the shape and size of the PDMS slices were controlled for precise and fast alignment with some alignment marks, premade on the PDMS slices. Next, the top PMMA frame was placed in close contact with the PDMS slices and metal tubes were slowly inserted. After that, all nuts were tightened to obtain a complete microfluidic chip. The duration of the entire process, from fabrication of chip slices to the final assembly of the microchip, was approximately 1 hour. Thus, the rapid customization of integrated microfluidic systems can be achieved with low cost and high throughput.

2.3 Device leakage test

An experimental system was set up to measure the leakage pressure that this method could reach, which was determined by the maximal internal pressure under which the device could work normally, without leakage (Figure 2). A programmable syringe pump (TJP-3A, LongerPump, China) was connected to the chip inlet and dyed water was perfused to gradually increase the internal pressure. The pressure was monitored using a pressure gauge (PTL503S, LONGLV, China), which was connected to the chip outlet. When leakage occurred, the maximal internal pressure was recorded and regarded as leakage pressure.



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Figure 2. Device leakage test: (a) Experimental setup for the measurement of leakage pressure. (b) Leakage pressure measurement system.

2.4 Device design and microfluidic cell culture of L929 cells

In our work, a 3D microfluidic chip was fabricated for proof-of-concept, as shown in Figure 3. Using the stack design, and a porous PET membrane between the two PDMS slices, two separate chambers were created. The PET membrane (pore size: 0.4 µm, 10 µm thickness) was obtained by directly cutting from Transwell. Top and bottom chambers could be perfused via individual inlets/outlets and separate fluid channels. The integrated PET membrane permits fluid perfusion and mass transport of soluble factors through the PET membrane, thus, the top chamber could be used for cell culture, while the bottom chamber served as a tool for drug and nutrient transport. Therefore, this microchip incorporated different modular structures into one system, enabling both cell culture and further cellular cytotoxicity analysis. The L929 cell line was cultured to investigate the feasibility of cell cultures in the microchip. The L929 cells were obtained from the Shanghai Cell Bank of the Chinese Academy of Sciences. The cell culture medium contained MEM supplemented with 10 % FBS. Matrigel matrix was inserted in the device to coat the PET membrane. As a control, a second device was not coated with


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Matrigel matrix. The cells were seeded in the culture chamber with culture medium at a concentration of 1 × 106 cells/ml. Cells were then incubated at 37°C in a humidified atmosphere containing 5% CO2. The cell culture medium was changed every 12 hours by injecting new cell culture medium through the microchannel underneath the PET membrane. Pores in membrane could enable the fluid perfusion and mass transport of soluble factors through the PET membrane, which led to the refreshment of cell culture medium in both the cell culture chamber above and in the microchannel underneath the PET membrane.

Figure 3. Rapid customization of an integrated microfluidic chip. (a) Structure of the microfluidic chip: Modular PDMS slices for cell culture and fluid perfusion are assembled with PET membrane. (b) Schematic of the microfluidic chip integrated with porous membrane. (c) Image of the chip.

2.5 Evaluation of cells under oxidative stress

The customized microfluidic chip was also used to evaluate cell viability under oxidative stress.



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According to the previous experiment results, cells grew better in coating condition, compared to no coating condition. After coating the PET membrane with Matrigel matrix, it was easier for processing cell seeding and cell culture. We followed the suggested procedure of microfluidic cell culture in Kim’s work,38 which preferred the coating condition. All PET membranes were coated with Matrigel matrix first and after 60 h cell culture, 1 mM of hydrogen peroxide (H2O2) solution, which caused the oxidative stress, was injected into the micro-channels, underneath the PET membrane, to treat the L929 cells. The live/dead cells were counted using the Calcein-AM/PI kit. Initial cell number was obtained by staining the cells at 60 h and counting using a fluorescent inverted microscope (OLYMPUS IX81, Olympus Optical Co. Ltd., Japan). After being treated with H2O2 and incubated for 12 h, cells were stained and counted with the fluorescence microscope.

2.6 Microfluidic chips with bioprinted hydrogel constructs

To further demonstrate the feasibility of combining both customized microfluidic chips and bioprinted hydrogel constructs, the rapid customized microfluidic chip was utilized as platform for the culture of bioprinted alginate beads. There are various ways to fabricate living constructs with specific architectures, including cell seeding on microscaffolds39, bioprinting of cell-laden microcarriers40 or hybrid approach with both microscaffolds and extrusion-based bioprinting41. In this case, extrusion-based bioprinting was used to deposite bioink and alginate hydrogel was selected as the bioink to bioprint beads following the common protocol. And cell line of L929 was used to display the feasibility of the idea instead of using primary culture cell. First, L929 cells were used to prepare the cell-laden alginate solution, by mixing the cell culture medium with concentration of 3 × 106 cells/mL and 3% sodium alginate solution at 1:2 (v/v) ratio. Second, the hydrogel constructs


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were directly printed onto the culture chambers of the bottom PDMS slice with the help of a customized bioprinter (Figure 9a). Third, after completion of the bioprinting process, the chip with bioprinted beads was quickly assembled and connected to a syringe pump (TJP-3A, LongerPump, China). The chip was then continuously perfused, through the upper microchannel, with cell culture medium at a flow rate of 100 µL/hr. The hydrogel constructs were also cultured in petri dishes as control group and the cell culture medium was changed every day.

3. RESULTS and DISCUSSION

3.1 Analysis and comparison of the merits and drawbacks of the proposed fabrication method

The proposed method is aimed at fabricating PDMS slices with different functions, according to a modular structure-based design, and assemble them quickly, using the adhesive-free packaging method. In our previous work,37 a new 3D sugar printer was developed and maltitol was used as melted material to obtain smooth microchannels and 3D-printed PDMS chips. Based on this, different reusable PDMS slices can be fabricated in less than one hour. Instead of traditional packaging technologies, including reversibly or irreversibly bonding, an adhesive-free packaging technology42, 43 was used, to form an integrated microfluidic chip consisting of stiff polymeric frames, soft PDMS functional slices and other functional parts. The merits of this technology include the quick assembly of the PDMS based microfluidic system, with no demand for any clean room facilities, and easy reuse of the multifunctional PDMS slices. The 3D printing technique enables the rapid fabrication of modular PDMS slices, with relatively low cost and high throughput. The diameter of printed microchannels could vary from 600 µm to less than 100 µm, and the channel size is not limited to this scale, which could satisfy the demand for



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complex microfluidic chips. Microchannel cross sections were observed by field scanning electron microscopy (FESEM, SU8010, Hitachi) (Figure 4). The used 3D printing parameters are detailed in Figure S2. From a to f, the printing speeds were a: 400 mm/min, b: 500 mm/min, c: 700 mm/min, d: 1000 mm/min, e: 2000 mm/min, f: 2200 mm/min. The images illustrate a change in channel geometry, from ellipse to near-circular, with the decrement of channel size. As the diameter of the channel becomes smaller, the cross section of the microchannel becomes increasingly circular. The surface tension and gravitational effect contribute to the formation of different channel shapes during sacrificial pattern printing, when different printing velocities are applied. More details about the dimensional changes of the microchannels replicated in the PDMS slices compared with the 3D printed structures were described in Figure S3. Moreover, the round microchannel is very promising for studies mimicking the blood vessel system, which has been investigated by many researchers. For example, Borenstein et al. successfully developed a method for fabricating channels with circular cross-sections, and then used it to develop functional endothelialized microvascular networks.44 As previously mentioned, an important advantage of the proposed method is the smoothness of obtained microchannels. Using a coherence scanning interferometer (NewView 8200, Zygo Corporation), the surface roughness of the inner microchannel was measured at 0.549 µm (Figure 4i), which guarantees proper fluid flow properties in the microfluidic chips.


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Figure 4. (a-f) SEM images of cross sections of PDMS microchannels, showing the channel shape and inner smooth surface. (a-c) scale bar = 200 µm, (d-f) scale bar = 100 µm. (g-i) Analysis of fabricated PDMS channels with a coherence scanning interferometer. Images of cross sections of PDMS channels (g, h) also illustrate the shape of printed microchannels. Measurements of channel surface roughness (i) indicate that the inner surface of the channels was very smooth (Ra = 0.549 µm).

However, there are still some limitations of this method. One critical drawback is that it is restricted to the used material, i.e. PDMS, which needs a secondary curing step of the final chip after



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printing. In several other 3D printing approaches, photosensitive resins are selected and microfluidic chips can be generated in one step, whereas when using the proposed method, the sugar structure must be printed first and later the poured PDMS can be cured. This translates into extra time and requires adequate operation, meaning that reliability is not as high as that of the automated one-step fabrication. Another limitation is that it can be tedious to generate very complicated structures, as this method is modified from FDM (fused deposition modeling) and sometimes requires additional support. Considering the merits of PDMS, including biocompatiblity, transparency, gas-permeablility and elastomericity, this method is still very helpful in biomedical research and especially suitable for experiments that require microscopic imaging, biocompatibility and mechanical stimuli.

3.2 Rapid customization of microfluidic chips

Due to the characteristics of PDMS, including gas-permeability and transparency, the PDMS-based microfluidic chips were successfully used in chemical and biological analyses. Several customized microfluidic systems, which combined different modular PDMS slices and other functional components, were fabricated and used to demonstrate the feasibility of building various integrated microfluidic systems with 3D-printed modular structures. As shown in Figure 5a, the rapid customization of the microfluidic chips was very simple. Firstly, different functional PDMS slices with modular structures were prepared. Secondly, modular slices were connected to best suit the intended application. Finally, several complex microfluidic chips were formed by adding extra necessary components. The most important advantages of this method were that the modular PDMS slices could be reused and recombined, the reversible packaging of the microfluidic system, and the rapid integration of functional elements with different thickness, made of various materials. These


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advantages are particularly relevant for biomedical and biochemical analyses. Figure 5b depicts two different microfluidic chips produced using the above-mentioned method, which shared the same, reused PDMS slice. Combined with other modular slices, a 3D microfluidic chip and a 3D mixing chip were made to evidentiate the rapid recombination of this method. Dyed deionized water was pumped through the microchannels, to demonstrate the complete features of these chips. As seen in Figure 5b, the first chip was made with two PDMS slices and formed into a 3D microfluidic chip, with straight and curved channels. Purple solution was perfused into its inlet to clearly display its structure, demonstrating the possibility of fabricating microfluidic chips with 3D complex structures. In contrast, the second chip was designed for mixing. By incorporating several perfused solutions, different mixed samples were obtained for further analysis. This was achieved by perfusing the chip with green, blue and red solutions. Subsequently, mixed solutions were observed form the outlets. Another example was taken as proof of rapid customization of highly integrated microfluidic systems with various functional features (Figure 5c). In the first configuration, two PDMS slices with fluidic networks and a soft electronic module were stacked together and reversibly packaged into a diagnostic microfluidic chip that could serve as a sensor. Meanwhile, the second configuration included three separate PDMS slices, a porous semipermeable polycarbonate membrane and biological scaffolds, and the chip could act as a tool for cell culture and drug assay, which confirmed the versatility of the proposed method. More complex integrated chips, including multilayer microfluidic networks for drug assay, are presented in the supplementary materials (Figure S4).



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Figure 5. Demonstration of rapid customization of 3D integrated microfluidic chips. (a) Customization of microfluidic chips with different modular slices. Reusable PDMS slices (A, B, C) were assembled into different functional microfluidic chips (D & E). (b) Two 3D microfluidic chips shared the same modular slice, and by adding different modular slices, a 3D microfluidic chip (1) and a 3D mixing chip (2) were formed. (c) The customization of complex 3D microfluidic chips was achieved by incorporating modular PDMS slices with different functional


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components, such as soft electronics (1) or a porous membrane and biological scaffolds (2).

3.3 Characterization of leakage pressure

The microfluidic chips were sealed using screws, resulting in tightening force being exerted on the PMMA frame and PDMS slices. A theoretical model could be used to analyze the reversible bonding between PMMA and PDMS, referring to previous studies on reversible PDMS bonding.45, 46 The reversible bonding is related to the van der Waals force and this force (Fv) per unit area is given by the Hamaker model:

= −

(1),

where H is the non-retarded Hamaker constant and D is the distance which separates the contacting surfaces of the two PDMS slices. To better understand this model, an original model was simplified to a PMMA-PDMS-PDMS-PMMA bonding model with single microchannel (Figure 6a). The cross-section of the microchannel is described with its width and height and the shape becomes near-circular when height is approximately equal to width. As changes in the vertical direction are responsible for leakage, only the influence of different forces on the vertical direction are discussed here. When a liquid is perfused into the microchannel, an internal pressure P causes the tensile stress force on the PDMS slices, which is described as Fts. The tightening force from the screws limits the expansion of the PDMS slices through the PMMA cover and serves as the counter-force (Fc). The compressive stress force (Fcs) in the PDMS slice and the forces mentioned above form a balance and can be presented as:

= + + with

= P · A


(2) (3)


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F = · A

(4)

= · A

(5)

where A1 is the equivalent area of the microchannel on the vertical direction, A2 is the contact area between the PDMS slice and the PMMA cover and Pc is the counter pressure, resulting from the contact between PDMS and PMMA . The compressive stress can be expressed as:

= · E

(6)

where is the vertical strain of PDMS and E is Young’s modulus of PDMS. The big difference in Young's modulus between PMMA (approximately 3.2 GPa) and PDMS (approximately 0.75 MPa, mix ratio 10:1) determines the even distribution of pressure on PDMS slices, as PMMA materials deform less than PDMS materials. Consequently,

= ·

(7)

where A3 is the contact area between the two PDMS slices. Then, equation (2) can be written as:

P = ε · E + ·

! "

+ ·

"

(8)

Obviously, the microchannel size is much smaller than the contact areas of PMMA-PDMS and PDMS-PDMS, i.e. A2/A1 ≫1 and A3/A1 ≫1. Therefore, the maximum internal pressure the chip

can withstand is mainly determined by counter pressure and the van der Waals force. Therefore, the external force exerted on the PMMA frame determined the reliability of the sealing of microfluidic chips. Furthermore, the sealing was also related to the contact area between the PMMA frames and the PDMS slices. A strong bonding of the assembled chips could be achieved by designing a large interfacing area between these two parts.


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As adequate force was applied using screws, the fabricated device was capable to withstand a maximum pressure of 200 kPa in the leakage test, which was recorded as the leakage pressure. This was consistent with the leakage pressure detected for irreversibly bonded PDMS-based microfluidic chips. This result was also confirmed by simulation with ANSYS Workbench 15 (Figure 6 and Figure S5). The simulation results showed that the maximum vertical displacement of the PDMS slice was 6.4151×10-5m.This feature enabled the reliable performance of rapidly assembled chips during analysis and made the customized microfluidic chips accessible to most chemical and biological analyses.

Figure 6. Characterization of leakage pressure. (a) Schematic of the simplified model using the adhesive-free packaging method. (b) Distributions of forces distributions in the assembled chip. (c) Simulation results on the



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deformation and vertical displacement of the assembled device.

3.4 Analysis of cell viability in the microfluidic cell culture and under oxidative stress

Figure 7. Microfluidic cell culture of L929 cells. (a) Phase-contrast images of L929 cells in the microfluidic chip after 24, 36, 48 and 60 h of culture with and without coating; scale bar= 200µm. (b) Fluorescent images of L929 cells in the chip after 72 h of culture. By Calcein-AM/PI staining, green fluorescent signal indicated active (live) cells and red indicated permeable (dead) cells; scale bar= 200µm. (c) Quantitative analysis of cell viability under both conditions.

To evaluate the cell culture in the customized chip and compare cell viability with and without coating the PET membrane, the cultured cells were observed by phase-contrast microscope after 24, 36, 48 and 60 h of culture (Figure 7a). The density of the population of cells grown on the microchannels under both conditions increased over time. Formed cells were polygonal and widespread with fine filopodia after 60 h of culture, compared to those cultured after 24, 36 or 48 h.


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Additionally, the L929 cells were cultured for 72 h and then stained with the Calcein-AM/PI kit to determine cell viability. Microfluidic channels were imaged under a fluorescent inverted microscope after a 30-min incubation time (Figure 7b). Image J analysis revealed a cell viability of approximately 94.94 ± 1.47% with no coating and 95.72 ± 0.41% when coating was used. This indicates that the 3D microfluidic device could allow for robust cell culture with porous PET membrane and confirmed that this fabrication method could efficiently generate robust devices and reduce the use of complex microfluidic system designs.



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Figure 8. Cell viability with and without oxidative stress (a) Microscope and fluorescent images of L929 cells after 0 h and 12 h exposure to H2O2 solution (1 mM). Calcein-AM (green) for live cells and PI (red) for dead cells were used to measure the cell viability; scale bar= 200µm. (b) Quantitative analysis of cell viability after 0 h and 12 h exposure to H2O2 solution.

H2O2 is well known for producing reactive oxygen species and can be used to induce cellular cytotoxicity. To compare the cell viability with and without oxidative stress, cell numbers after 0 h and 12 h exposure to H2O2 solution (1 mM) were counted by staining with Calcein-AM/PI and examined


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under an inverted fluorescence microscope. By analyzing the fluorescent images of 0 h and 12 h exposure with the help of Image J, cell viabilities with and without oxidative stress were achieved. Figure 8 shows that H2O2 induced cell death, resulting in a significant difference in cell viability (0 h 96.57 ± 1.61% vs. 12 h 70.32 ± 3.64%). This highlights the advantages of using the rapidly fabricated microchip in researching cellular cytotoxicity. 3.5 Hydrogel-based Beads in microfluidic chips Extrusion-based bioprinting is a common bioprinting technique, which has greatly evolved over the past 10 years. Combining the extrusion-based bioprinting technique with microfluidic chips could be very helpful in the field of tissue engineering, as it could provide a significant platform for the fabrication of living constructs and cell culture, which would enable the reconstruction of artificial organs on chips.47 To successfully fabricate hydrogel constructs on chips, several factors must be taken into consideration, including the design and printing strategies in 3D bioprinting of soft materials48, usage of support materials during 3D bioprinting49 and post-printing surface modification of the bioprinted device50. Preliminary experimental results, which show great promise in the fabrication of organ-on-a-chip are presented below. As seen in Figure 9a, the cell culture chamber of the modular PDMS slice interfaced with the customized bioprinter and the hydrogel-based beads with cells were then bioprinted on the PDMS slice. Upon the completion of the assembly of different functional PDMS slices, the chip with spherical beads was prepared for further analysis. This process combined the rapidly customized microfluidic chips with bioprinters to generate a system that allows for biofabrication, microfluidic culture and biomedical analysis. It is promising that complex hydrogel-based structures could be directly printed in the microfluidic chips. To examine the advantages of this process, the chip with



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bioprinted beads was supplied with dynamic perfusion continuously. After 6 days of culture, the beads were observed using phase-contrast microscope (Figure 9b). The analysis revealed a homogeneous distribution of L929 cells. Cell viability in the hydrogel-based beads was confirmed using the Calcein-AM/PI staining kit. Images were obtained under a confocal fluorescence microscope (LSM780, ZEISS, Germany) and viable cells (green) were observed in the beads (Figure 9d). The results in Figure 9c revealed that bioprinted hydrogel beads in microfluidic chips showed a higher cell viability than those cultured in petri dishes (Microfluidic culture 90.49 ± 4.24 % vs. Traditional culture 82.53 ± 2.71 %). The beads were also stained with TRITC-Phalloidin and counterstained with DAPI (Figure 9e). The results indicated cell proliferation in alginate beads after 6 days and it can be inferred that cell clusters were formed.

Figure 9. (a) Process of bioprinting hydrogel constructs in the chip and microfluidic dynamic culture after chip assembly. (b) Phase-contrast images of L929 cells in the bioprinted alginate beads after 6 days of culture. (c) Quantitative analysis of cell viability in the bioprinted alginate beads by both microfluidic and traditional ways. (d) Viable cells in the hydrogel-based beads; green and red fluorescent dyes indicate live/dead cells, respectively. (e)


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Blue fluorescence indicates the nuclei (DAPI) and red fluorescence indicates the cytoskeleton (TRITC-Phalloidin).

4. CONCLUSIONS

A rapid method for the customization of integrated microfluidic chips was investigated in this study. Modular structure-based 3D-printed designs could reduce the cost and improve the throughput of microfluidic systems. By selecting different functional PDMS slices and assembling them with other functional components, complex microfluidic chips integrated with sensors and biological scaffolds could be realized. This method exploited the advantages of the 3D printing technique and modular approach, which decreased the optimal iterations of customized microfluidic chips, compared to conventional methods. Thus, this method could play an important role in the design and fabrication of complex integrated microfluidic systems. Additionally, a 3D microfluidic chip incorporated with PET membrane was rapidly customized and utilized for primary cell culture, showing great potential for use in biomedical research. The flexible combination of this method and bioprinting technique was also demonstrated. Improving the accuracy of 3D-printed modular structures is still ongoing, and other chemical and biological applications concerning micro analysis systems will be explored with this method in the future. ASSOCIATED CONTENT Supporting Information The following files are available free of charge. Figures S1. Schematic of 3D printing system. Figures S2. 3D printing parameters for chip fabrication. Figures S3. Analysis of printing process.



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Figures S4. Demonstration of rapid customization of 3D integrated microfluidic chips. Figures S5. Simulation results of leakage test. Notes The authors declare no competing financial interest. ACKNOWLEDGMENT This paper was sponsored by the National Nature Science Foundation of China (No. 51622510), the Science Fund for Creative Research Groups of National Natural Science Foundation of China (No. 51521064) and the Nature Science Foundation of Zhejiang Province, China (No. LR17E050001). REFERENCES 1.

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