Our rail-based open microfluidic platform consists of a 3D printed structure of photo curable resin and an underlying pressure sensitive adhesive (PSA) film. 3D-printed rail structures consisted of high rails (HRs) surrounded by low rail (LRs) and reservoir walls which supports rail structures and are bonded to the PSA film. HRs and LRs have different heights from the PSA film to retain liquid with different strengths (Fig. 1a). Figure 1b illustrates the basic working principle of the novel patterning process of hydrogel, which allows us to form hollow microchannels surrounded by precisely defined hydrogel patterns. The length of the rail, l, is 5 mm, and the width of low and high rail, ({w}_{l}) and ({w}_{h}), is uniformly 1 mm. The heights of low and high rails are respectively h = 100 μm and H = 300 μm. Two through-holes at the ends of a high rail function as fluid ports for liquid injection and aspiration. Air plasma treatment activates the surface of the device by increasing hydrophilicity. We first inject a hydrogel precursor solution through one of the fluid ports, which fills the area under all rails (the second column of Fig. 1b). Upon aspirating through the same port, the liquid is sucked into the pipette only from the region below high rail (HR), while the low rail (LR) strongly retains the liquid (the third column of Fig. 1b). The result is the hollow microchannel defined by the empty region under the high rail with the hydrogel precursor solution remaining pinned by the low rail (the fourth column of Fig. 1b). In the following, we delineate the physical principle behind this selective removal and controlled pinning of hydrogel patterns, which leads to various hydrogel-based channel structures and facile fabrication of cell co-culture systems.
Schematic diagram of aspiration-mediated patterning within an open microfluidic device. (a) Schematic of a rail-based open microfluidic device, which is designed to yield a single channel beneath the high rail. (b) Illustration of steps in the aspiration-mediated microchannel formation process. The bottom row features optical images taken through the clear PSA film from the underside of the device during filling and aspiration of green dyed water.
Theoretical analysis of microchannel formation
As the pressure at a port is lowered by aspiration with a micropipette, the interior pressure of the hydrogel precursor solution is also lowered. The Young–Laplace equation states that the liquid–gas interface of the hydrogel precursor solution originally filling the gaps caves inward (Fig. 2a(i—iii)) due to the pressure difference between the atmosphere (pa) and the liquid (pl): ∆p = pa − pl = γ(1/R1 + 1/R2), where γ is the gas–liquid interfacial tension and R1 and R2 are two radii of curvature in orthogonal planes intersecting the interface30. The contact line where the gas–liquid interface meets the solid surface is initially immobile while resisting the pulling force, because it can recede only when the contact angle is reduced below the critical receding contact angle, θR,c. The patterning of hydrogel that originally fills the gaps under both HRs and LRs critically depends on which interface starts to recede first by reaching θR,c.
Theoretical considerations for designing rail structures capable of aspiration-mediated liquid patterning. (a) Schematic diagrams indicating key device dimensions and the Laplace pressures of menisci (i) immediately after aspiration, (ii) as the port interface deforms, and (iii) as the receding interface travels along the HR. (b) Time-lapse images from top to bottom illustrating meniscus dynamics during aspiration under three different combinations of critical capillary pressures. The dimensions of the structures are such that [H, h, D] = [300, 100, 300] μm, (Left column), [300, 200, 300] μm (middle column), [200, 175, 500] μm (Right column). Lengths and widths of all rails are fixed as 5 and 1 mm, respectively. (c) Plots of the effects of port diameter and LR height on bursting at the open port. (d) Plots of the effects of HR height and LR height on formation of microchannel. (c,d) Shaded regions demarcate dimensions suitable (green) and unsuitable (red) for microchannel formation based on the theoretical calculations. Closed markers and open markers indicate successful and unsuccessful microchannel formation in experiments, respectively. Deionized water (filled circle), 2.5 mg/ml of fibrinogen solution (filled traingle), and 3 mg/ml of collagen type I solution (filled square) were used for experimental verification. Experimental results were derived through three consecutive successes or failures for each condition.
For the initial hydrogel solution-loaded configuration, liquid–gas interfaces arise at the remaining port opposite to the port of aspiration, and lateral faces of the fluid filling the gaps below the LRs. We will refer to the opposing port as an “open port” in the following analysis. For the open port of diameter D, the maximum pressure difference that the interface can withstand before the contact angle on the inner wall of HRs reaches zero, ∆pO, corresponds to the pressure difference for a hemispherical interface with the maximum curvature of 4/D: ∆pO = 4γ/D (Fig. 2a(ii)). The capillary pressure at the air–liquid interface beneath LRs is given by ∆pL = γ(2cosθ/h + 1/η), where θ is the instantaneous contact angle of liquid, h is the height of the rail, and η is the radius of curvature on a plane parallel to the substrate. During aspiration, θ decreases until reaching θR,c when the capillary pressure is maximized. We experimentally found that θR,c is nearly zero for hydrogel precursor solution on the hydrophilized 3D printed part of resin and PSA film surfaces. Hence, we estimate cosθR,c to be 1 due to the small receding contact angle, θR,c. Furthermore, η scales with l, the length of rails, and h < < l for most rail designs, thus the 1/η term is negligible. Hence, we estimate the maximum pressure difference for the side area below LRs as ∆pL = 2γ/h.
If ∆pO < ∆pL, the critical liquid pressure for interface recession is reached at the open port before the side area. Once burst at the open port, the interface spontaneously bulges accompanying the decrease of interface curvature to meet the bottom substrate. Then the interface curvature is newly defined by the height (H) and width (wH) of HR (Fig. 2a(iii)). The critical capillary pressure that can induce interface recession beneath HRs is given by ∆pH = 2γ(1/wH + 1/H). If ∆pL > ∆pO > ∆pH, the liquid pressure is already low enough to ensure continuous recession of the liquid beneath HR. If ∆pL > ∆pH > ∆pO, further decrease of the liquid pressure following bursting of the open port can cause the liquid to recede along a lane beneath the HR. In short, both cases of ∆pL > ∆pO > ∆pH and ∆pL > ∆pH > ∆pO result in successful patterning. However, if ∆pH > ∆pL, the liquid will stop upon bursting at the open port and recede beneath LRs, rather than HRs, resulting in patterning failure.
Figure 2b displays the experimental results of hydrogel patterning for different geometries of microdevices. In the left column, ∆pL is greater than both ∆pO and ∆pH, and thus the interface recession occurs from the open port, continues beneath HR, to result in a straight empty lane flanked by the hydrogel beneath LR. In the middle column, the side areas beneath LRs burst before the open port because ∆pL < ∆pO. In the right column, the open port bursts first because ∆pL > ∆pO, but further decrease of the liquid pressure bursts the side areas beneath LRs before the interface from the open port starts to recede beneath HR.
Figure 2c,d display the experimental results of microchannel formation as a function of the geometry of device, the port diameter and the heights of low and high rails, while fixing the length and width of all rails to 5 mm and 1 mm, respectively. We tested three types of liquids: deionized water, 2.5 mg/ml of fibrin gel pre-solution, and 3 mg/ml of collagen type I pre-solution. After fully filling the area under a rail structure, we immediately aspirated the liquid with a manual micropipette by gradually increasing the suction pressure until one of the interfaces starts receding. In Fig. 2c, we can predict where the interface recession onsets upon aspiration, whose criterion is given by ∆pL = ∆pO, or D = 2 h. For D > 2 h, we get ∆pL > ∆pO, and thus the port bursts first, leading to successful channel formation in the beginning. Otherwise, the interface moves at the side area of LR before at the port, preventing the formation of the microchannel under HR. Figure 2d gives the condition of the height of HR to ensure that the interface formed from the open port keeps receding while the interface beneath LR remains pinned. The corresponding boundary is given by ∆pL = ∆pH, or H = h/(1 − h). For H > h/(1 − h), we get ∆pL > ∆pH, leading to continuous recession of the interface under the HR from the open port. Otherwise, the interface under LR starts to recede, eventually merging with the interface originated from the open port. We see in Fig. 2c,d that our theoretical predictions agree well with the experimental results regardless of the liquid type. As all interfaces formed by a liquid under a rail structure share the same interfacial tension, γ, the moving interface is determined only by the geometry of the rail structure. The physical properties of the liquids with surface tension and viscosity were measured with Smartdrop (FEMTOBIOMED.inc, Korea) and AR-G2 (TA instruments, USA) respectively as listed in Table 1.
To determine the effect of aspiration pressure on patterning, we conducted patterning tests by aspirating deionized water that filled the region under a 3D-printed basic rail structure whose D, h, and H are 600 μm, 100 μm, and 300 μm, respectively, with various aspirating pressures. For the designed parameters, maximum capillary pressures at each interface are ∆pL = 1439 Pa, ∆pO = 479.8 Pa, and ∆pH = 311.9 Pa. To apply different aspiration pressures, we applied maximum aspiration pressures of four different types of micropipettes whose maximum handling volumes are 10, 20, 100, and 200 μl, respectively. Maximum aspiration pressures of the four micropipettes were measured by tracking the velocity of receding fluid interfaces in straight microfluidic channels as detailed in the supplementary information and Table S1. When we aspirated the water with 10 μl and 20 μl pipettes, whose maximum aspiration pressures are below 2000 Pa, the water remained only under LRs as predicted. However, when using 100 μl and 200 μl pipette, whose maximum aspiration pressures are above 3000 Pa, the water under LRs was removed simultaneously with the water under HR, resulting in patterning failure. The results show that aspiration pressures (~ 1950 Pa) slightly greater than ∆PL under the LR also allow for successful patterning. We expect that further studies of meniscus kinetics using pressure-controlled pumps may explain how successful patterning can still result from aspiration pressures greater than the capillary pressure at interfaces under LRs.
Throughput and uniformity of the patterning method
The unique nature of this hydrogel patterning method is advantageous in filling multiple devices. To fill a series of devices using conventional pipetting methods, the operator must repeatedly uptake the exact volume of a single device from the stock container before loading the next device. In contrast, the present technique can uptake the combined volume of all desired devices at the onset of loading. Each device is initially overfilled by arbitrarily depressing the pipette prior to aspiration of the excess fluid by releasing the pipette piston. Video S1 shows a comparison of the two patterning procedures to fill a dozen devices that were used for 3D cytotoxicity assay in our previous study26. The reduced uptake procedure can enhance experimental throughput when loading hydrogel precursors in microfluidic devices. This is especially beneficial when handling gels with a short pot life, such as fibrin gels whose cross-linking time is approximately 1 min16. Using aspiration-mediated patterning, it took around 20 s to fill a dozen devices, while the conventional method that injects the exact amount of hydrogel solution for each device took approximately two-fold longer.
In this manner, the aspiration-mediated patterning method achieves improvements in throughput using a manual micropipette that typically necessitate costlier electronic pipette equipment with automated dispensing capabilities. Furthermore, our experiments show that such electronic pipettes become less reliable with certain chip designs that require small fluid volumes (~ 1 µL) and produce high capillary action. Video S2 provides a side-by-side comparison of patterning eight devices identical to those in Video S1 using three methods: automated injection with an electronic micropipette, manual injection with a standard micropipette, and aspiration-mediated patterning with a standard micropipette. The video provides a bottom view through the transparent PSA film to better visualize the patterning of fluid in the chips. Even though the electronic micropipette in Video S2 was programmed to dispense a uniform volume (1 µL) into each device, the micropipette exhibits a pattern of overfilling one device, underfilling the next, and then again overfilling the subsequent device. We believe that this is due to the high capillary action of the plasma treated microscale structures wicking excess fluid from the pipette into the device. Visual inspection of the micropipette tip after loading each overfilled device consistently revealed an air gap at the tip. Attempting to fill the next well results in ejection of the air into the channel rather than liquid. This ejection of the air primes the micropipette for overfilling of the subsequent device by excess wicking. On the other hand, both manual pipetteing methods successfully pattern all devices, although the aspiration-mediated method provides superior speed.
To further characterize patterning uniformity, we conducted aspiration-mediated patterning with dyed water using a basic 3D-printed device, whose D, h, and H are 600 μm, 100 μl, and 300 μm, respectively. We measured the area of dyed water that remained after aspiration and compared it with the area of 1.2 μl of dyed water injected through one of two ports in the HR. Even though water is injected through a port, the water fills the area under LRs instead of the HR due to large capillary force. Consequently, both methods result in similar positioning of dyed water. The uniformities of the remained dyed water were 92.5% and 89.0% for the aspiration method and manual injection through a port, respectively, and they showed no statistical difference as shown in Figure S2.
Formation of multiple discrete microchannels via single aspiration
Our fluid dynamic studies using basic rail structures in the previous sections verify that the air–liquid interface with the smallest critical capillary pressure tends to move first along the rail. Leveraging this phenomenon can provide control over the sequential formation of discrete microchannels with a single aspiration. Figure 3 exemplifies multiple channel formation using various critical capillary pressures at air–liquid interface pinned at holes. In this device, a rectangular LR, 75 μm apart from the underlying PSA film surrounds ‘S’, ‘N’, and ‘U’ shaped HRs, 300 μm apart from the PSA film. The diameter of fluid ports decreases in the order of S1 to U1 as listed in the table in Fig. 3a while U2 has the largest port diameter. When aspirating dyed water filling under the rail structure from the port labeled ‘U2’, the air–liquid interface at the largest opposing port (S1) begins traveling along the rail to form an ‘S’ shaped microchannel (at 0.5 s in Fig. 3b). As the suction continues at U2, the air–liquid interface at the next largest port (N1) now begins to recede (at 1.1 s in Fig. 3b). Consequently, microchannels with shapes of ‘S’, ‘N’, and ‘U’ form via a single aspiration process through one port as shown in Fig. 3b within 2 s (see also Video S3). These results are achievable because the body of fluid underneath the chip is continuous. If instead LRs directly contact the PSA to form solid barriers between HRs, rather than fluid filled gaps, aspiration-mediated rendering of discontinuous microchannels would not be supported.
Rendering of multiple discrete microchannels using different port sizes. (a) Image of the underside of a device with three HRs in the shapes of ‘S’, ‘N’, and ‘U’, surrounded by a rectangular LR. Loading and aspiration occur through port U2. Opposing port sizes decrease from S1 to U1 as indicated in the bottom table. Scale bar, 2 mm. (b) Sequential snapshots (top to bottom) viewed from the bottom of the device during aspiration. Air–liquid interfaces preferentially advance from the largest port to the smallest port. Red arrows indicate the aspirating port, U2. Scale bar, 2 mm.
During multiple channel formation, differences in port diameter within the same channel lead to orderly movement of the air–liquid interface. If ports of the same diameters are employed, air can infiltrate into the channel through the identical ports simultaneously, to form a thin film of liquid between the infiltrating air from the two ports, as shown in Video S4. Breaking the thin film to form a continuous microchannel requires additional pressure drop since foam films have larger Laplace pressures than droplets of equal curvature, owing to the presence of two air–liquid interfaces30. In the video, a continuous microchannel forms in the top right HR because it contains the port of aspiration, and thus only a single port for air infiltration. In contrast, a similar device employing asymmetrical port sizes in each HR exhibits unidirectional movement of air–liquid interfaces beneath HRs as shown in Video S5.
This unique liquid patterning method enables versatile rendering of microchannel in hydrogels. Figure 4a,b show devices with multiple hydrogel channels arranged in parallel lines and a lattice of squares, respectively. Figure 4c,d respectively highlight the ability of our scheme to generate a single circular and two concentric channels. Figure 4e shows the generation of 98 microchannels under 13.75 mm × 12.7 mm sized low rail via a single aspiration. Video S6 shows the aspirating process to leave hydrogels under LRs as shown in Fig. 4.
Bottom view images of devices with multiple microchannels with various shapes enclosed by green colored hydrogels. The channels are organized into (a) parallel lines, (b) a square lattice, (c) a circle, and (d) concentric circles. (e) Time-lapse images from left to right showing the generation of 98 channels via a single aspiration. Since the opposing hole directly beneath the aspirating hole (red arrow) has the smallest hole diameter, hydrogel solution under the channel containing the aspirating hole is removed last. All scale bars are 2 mm.
An open microfluidic device for screening vasculogenic capacities
Even though it is widely known that tumors recruit vasculature to acquire oxygen and nutrients31, it is not always observed in in vitro models. Previous studies also showed very diverse vasculogenic capacities of cancer cell lines from different organs within an injection-molded device32,33, and required additional co-culturing of endothelial cells or fibroblasts to mimic in tumor vasculature34. These results guided us to develop a microfluidic model to screen vasculogenic capacities of cell lines for better recapitulation of tumor angiogenesis. Modification of the device in Fig. 4b provides a platform for convenient and effective comparison of paracrine signaling induced by co-culture of five cell types. Here, the microfluidic device contains four linear HRs surrounded by a square LR (Fig. 5a–c). Four support structures suspend the rail structures by connecting them to the reservoir walls, which form growth medium reservoirs when bonded to a PSA film. Patterning of a fibrin hydrogel (2.5 mg/ml) containing a mixture of primary human umbilical vein endothelial cells (HUVEC, 4 × 106 cells/ml) and primary human normal lung fibroblasts (LFs, 1 × 106 cells/ml) under the LR, and subsequent seeding of fibrin gels containing different cell types into the rendered microchannels establishes the basic experimental setup (Fig. 5d). The present studies involve eleven devices, containing microchannels underneath HRs filled with cells from a colon fibroblast cell line (CCD-18Co), a liver cancer cell line (HepG2), a glioblastoma cell line (U87MG), and a lung carcinoma cell line (H1299). In each device, two channels containing either LFs or acellular fibrin gel serve as positive and negative controls, respectively. The results include 11 samples of LFs and control, 6 samples of H1299 and U87MG, and 5 samples of CCD-18Co and HepG2. All gels contain cell concentrations of 5 × 106 cells/ml except for an acellular fibrin gel as a control. Figure 5e presents a confocal image of an exemplary device taken after 5 days of cultivation. Regions of interest (ROI) measuring 1 × 1.8 mm2 in area, center around each HR, and encompass the interfaces between the HUVEC-LF gel and the gels containing cells of interest. Analysis of z-projected confocal images yield quantification of the vasculogenic capacities of the cells of interest (Fig. 5f,g). In alignment with previous studies6,35, LFs vigorously promote formation of vasculature. Furthermore, acellular gels and gels containing LFs exhibit angiogenic sprouting into the channels. On the other hand, U87MG and H1299 gels inhibit the growth of surrounding vessels and do not provoke cancer angiogenesis. Gels containing CCD-18Co and HepG2 show no significant difference in vasculogenic capacity compared against the acellular matrix. Since tumor angiogenesis is orchestrated by a variety of activators and inhibitors36, we hypothesize that the selected cell lines secreted insufficient pro-angiogenic factors in our experimental setup. Even though we could not find a cell line that induces formation of blood vessel networks, LFs significantly showed pro-vasculogenic performance. The results of these tests corroborate the ability of the platform for screening candidates for cancer angiogenesis in vitro. Furthermore, the versatility of the channel rendering method supports facile device adaptation to study a broad range of paracrine signaling cues in various conformations.
Application of multiple channel rendering for screening vasculogenic capacities of multiple cell types. (a) Schematic overview of the device. (b) A photo of the device held between two fingers. Edges of a PSA film were not trimmed to highlight that it is bonded to the bottom surface of the 3D printed part. (c) Bottom view image of the microfluidic device with four channels surrounded by a square shaped LR (5 mm × 5 mm). (d) Schematic diagram of multi-cell co-culture experiment for assessing capabilities to form vascular network. (e) Confocal fluorescence image of the vasculogenic co-culture model under the square LR showing CD31 labeled HUVECs with green fluorescence. Red dashed box delineates an example region of interest (ROI) (1 mm × 1.8 mm) centered around the HR, with width equal to three times the HR. (f) Representative confocal micrographs of ROIs in channels containing different cell types. Scale bar, 300 μm. (g) Plot of average area of vasculature within the ROIs for each cell line obtained from two independent experiments (n = 11 for LF and Gel only, n = 6 for H1299 and U87MG, and n = 5 for CCD-18Co and HepG2). Error bars represent SEM. *** denotes p < 0.001, ** denotes p < 0.01, and * denotes p < 0.05 as measured by one-way ANOVA and comparison against the ‘Gel only’ condition with Tukey’s post-hoc test.
