The uptake of microfluidics by the wider scientific community has been limited by the fabrication barrier created by the skills and equipment required for the production of traditional microfluidic devices. Here we present simple 3D printed microfluidic devices using an inexpensive and readily accessible printer with commercially available printer materials. We demonstrate that previously reported limitations of transparency and fidelity have been overcome, whilst devices capable of operating at pressures in excess of 2000 kPa illustrate that leakage issues have also been resolved. The utility of the 3D printed microfluidic devices is illustrated by encapsulating dental pulp stem cells within alginate droplets; cell viability assays show the vast majority of cells remain live, and device transparency is sufficient for single cell imaging. The accessibility of these devices is further enhanced through fabrication of integrated ports and by the introduction of a Lego ® -like modular system facilitating rapid prototyping whilst offering the potential for novices to build microfluidic systems from a database of microfluidic components.

The 3D printed devices presented here were designed to allow easy integration with traditional fluid handling systems. As such, threaded ports at the inlets and outlets allow for the use of simple PEEK finger-tight fittings to connect devices to tubing and pumps. An example design is shown in Fig 1 . For further accessibility connectable fluidic modules were also designed. The fluidic modules were based on the Lego ® blocks that are familiar around the world and can be simply clipped together creating leak-free re-configurable microfluidic systems.

Until recently, 3D printing has been limited by resolution or cost of the printers [ 24 , 25 ]. However, recent advances mean that microfabrication is now possible off-the-shelf, without sophisticated manufacturing centres [ 26 ] and advances continue apace meaning the possibilities are likely to increase further still. 3D printing has been applied to micro- and millifluidics in combination with other techniques [ 27 ] or as a fabrication process in its own right [ 22 , 24 , 26 , 28 – 31 ]. Stereolithography (SL) printing, which has been more commonly used for microfluidics, involves the layer-by-layer photocuring of a polymer resin enabling the manufacture of three-dimensional objects from a reservoir of liquid resin. The more prevalent and accessible printing technology, fused filament fabrication (FFF), has often been discounted for the production of microfluidic devices [ 22 ]. This type of printing has often been dismissed due to leaking between extruded layers, along with the lack of transparency and a perceived lack of resolution and accuracy [ 32 ]. FFF printing (also known as fused deposition modelling (FDM) or extrusion printing) is an additive manufacturing process where the extrusion of molten polymer layer-by-layer enables the construction of 3D objects. By this method, proof-of-concept 800μm diameter microfluidic channels have been produced in polypropylene [ 24 ]. Whilst SL printing can provide greater channel resolution than FFF, it has rarely been demonstrated with channel dimensions below 500μm. Significantly, the increased complexity and cost of the SL printing process means that fabrication of such devices has often been outsourced to specialists [ 22 , 29 , 33 , 34 ] creating comparable barriers to uptake as with traditional manufacturing methods. With printers currently available at prices below £1000, FFF offers much higher levels of accessibility meaning most laboratories should be capable of fabricating their own devices in-house, facilitating cheap production and fast prototyping for a wide range of applications.

Recently, extrusion based 3D printing (fused filament fabrication (FFF)) has developed into a readily available consumer technology. As ease of use and resolutions have improved, and prices fallen, the ability to create custom objects quickly and easily has become available to all, causing a paradigm shift in small scale manufacturing. The ease of creating custom objects and devices cheaply and easily is democratising previously specialist areas of manufacturing, greatly increasing the capability of non-experts, and allowing cheap and rapid prototyping and production.

The barriers to wider uptake are clear if the fabrication technique of soft lithography with polydimethylsiloxane (PDMS) is taken as just one example. Despite the advantages to this fabrication technique, the creation of a PDMS device requires knowledge of- and access to- photolithographic and plasma bonding equipment. Similar know-how and equipment requirements are true of other common fabrication techniques such as laser fabrication, micro milling, ion-etching and photolithographic fabrication. Berthier et al summarise the problem stating “adoption of microscale technologies by biologists hinges on… successful collaborations between engineers and biologists… and the establishment of standard platforms that… are widely accessible and available to the biology community at large”[ 23 ]. Here we present a microfluidic platform capable of addressing this challenge, taking versatile microfluidics into the laboratories of the wider scientific community at low cost and without the need for specialist equipment or expertise.

Despite these unique properties and diverse application areas, microfluidics has largely remained a somewhat specialist research area with limited uptake by those who could benefit most from the technology. Two key factors have limited the take-up by wider disciplines; manufacturing and versatility. Firstly, traditional microfluidic manufacturing methods, such as soft lithography, require skills and equipment that is often not readily available in a typical biology, chemistry or pharmacy laboratory [ 21 , 22 ]. Secondly, the fixed nature of the fabricated devices limits iterative process optimisation or flexible application.

For many years microfluidics has been hailed as a field that could transform the way research is performed in the biological and chemical communities [ 1 – 3 ]. Microfluidics enables the precise manipulation of fluids on the small scale, enabling the use of small sample volumes, reduced costs, increased throughput, and parallel and sequential processing easily amenable to automation and portability on a scale beyond that achievable by manual or traditional robotic manipulation [ 4 ]. Microfluidics exploits the unique behaviour of fluids on the micro-scale where surface and viscous forces dominate over gravity and inertia, giving rise to laminar flow in single phase systems, and reproducible and programmable droplet flow in multiphase systems [ 5 ]. These advantageous fluid dynamics have been used in diverse fields with applications in cell encapsulation [ 6 ], DNA analysis [ 7 , 8 ], drug prototyping [ 9 ], high throughput screening [ 10 , 11 ], cell and droplet sorting and separation [ 12 – 15 ], chemical synthesis [ 16 ], chemical separations [ 17 ] radiopharmaceutical production [ 18 ], proteomics [ 19 ] and diagnostic technologies [ 20 ] amongst others.

Cells were centrifuged at 400g for 5 min. The supernatant was discarded and cells were resuspended in 1 mL of sterile PBS containing CellMask Orange (C10045, Molecular Probes) at a concentration of 5 ng/mL. The cells were incubated with this membrane stain for 10 min at room temperature before being centrifuged at 900g for 2 min. The supernatant was discarded and the cell pellet was washed by resuspension in sterile PBS and incubation at room temperature for 1 min. The cells were then recentrifuged (900g, 2 min) and resuspended once more in 1 mL of PBS for downstream analysis. Droplets containing sulforhodamine b (250 nM and 25 nM) and aqueous suspension of CellMask Orange labelled stem cells were imaged under flow conditions in the transparent PLA device equivalent to those used in the device transparency tests. Imaging was conducted at 4x or 10x magnification with 532 nm illumination via a TIRF fibre couple interfaced in epi-fluorescence mode on a Nikon Eclipse Ti-U inverted microscope. A 590/50 nm fluorescence emission filter (Chroma, USA) filtered emitted light prior to imaging on an Andor iXon camera at an acquisition frame time of 18 ms.

Human dental pulp stem cells (hDPSCs) were encapsulated in alginate capsules (1x10 6 cells/mL). The cells were obtained from third molars (donors aged 17–20) with all patient’s informed written consent in accordance with the Research and Human Tissue Act 2004. Ethical approval was granted by South East Wales Research Ethics Committee of the National Research Ethics Service (permission number: 07/WESE04/84. Ethical documentation can be found in S1 – S3 Figs) and cultured in α-modification minimum essential medium (αMEM) containing 2mM glutamine, ribonucleosides and deoxyribonucleosides (Life Technologies, UK). The medium was supplemented with 1% (v/v) penicillin/streptomycin, 10% (v/v) heat-inactivated foetal bovine serum (FBS) (Life Technologies, UK) and 100μM l-ascorbic acid 2-phosphate (Sigma-Aldrich, UK). The medium was changed every 2–3 days until cells reached 80–90% confluence. Upon reaching confluence, culture medium was removed by aspiration and the cells washed with phosphate buffered saline (PBS) (Sigma-Aldrich, UK). Cells were dissociated by adding trypsin-EDTA 0.25% (v/v) (Sigma-Aldrich, UK) and returned to the incubator for 3–5 minutes until they became rounded and detached. The trypsin was neutralised by adding culture medium. The medium and cell solution were then transferred to 15ml falcon tubes and centrifuged at 1500rpm for 5 minutes. After discarding the supernatant, pellets were resuspended in medium and cell counts performed using a haemocytometer. Finally, cells were centrifuged again and resuspended at a density of 1 million cells per ml of alginate in 2% low viscosity alginate solution (AO682, Sigma Aldrich, UK). The alginate solution was created by adding 5mg/ml of calcium carbonate (Sigma-Aldrich) to αMEM supplemented with 1% (v/v) penicillin/streptomycin before adding 20mg/ml of alginate and stirring for 2 hours at 50°C. Monodisperse alginate droplets containing stem cells were created on the 3D printed devices within sunflower oil continuous phase. Alginate droplets were gelled for approximately 10 minutes on exiting the chip in a bath solution of sunflower oil containing glacial acetic acid (0.3% v/v). After gelling the capsules were washed in culture medium and, after varying periods of time in culture, viability assessed using a LIVE/DEAD ® Viability/Cytotoxicity assay kit for mammalian cells (Invitrogen), with calcein-AM (green) indicating intracellular esterase activity in live cells, and ethidium homodimer-1 (red) fluorescence indicating loss of plasma membrane integrity in dead cells. Laser scanning confocal imaging of encapsulated cells was performed using a Leica SP5 Confocal Microscope and LAS AF imaging software (Leica Microsystems, Germany). Images of encapsulated cells were acquired from confocal Z scan over a depth of 600 μm.

To assess the quality and accuracy of the 3D printing, a series of channels were printed with diameters ranging from 400μm-1.5mm in increments of 100μm. The channel dimensions were then measured using a Nikon AZ100 microscope with NIS Elements 3.2 software. The channels were printed using both FFF printing and SL printing. FFF printed channels were printed in both horizontal and vertical orientations. Horizontally printed channels were measured in two directions to assess both the width and height of the channels. 25μm, 50μm and 100μm layer thickness prints were assessed. The measured channel dimensions were compared to the original dimensions specified in the models created in Solidworks.

Flow-focusing junctions were tested using mineral oil (Sigma-Aldrich) dyed with Oil Blue N (Sigma-Aldrich). Further tests used an alginate solution that contained 2% alginate in distilled water with 7.5mg/ml of calcium carbonate, CaCO 3 , (Sigma-Aldrich) (with SilverSpoon red food colouring for visualisation). The solution was stirred magnetically for 1h at 50°C to dissolve the alginate. To create oil in water emulsions, a 10mM aqueous oleic acid (Sigma-Aldrich) solution at pH13 was used with pH adjustment achieved with sodium hydroxide (Sigma-Aldrich) in deionised water (4g/L). The solution was then sonicated for 1 minute after the oleic acid was added to ensure that the solution was monophasic. Measurements of the droplets created on the flow-focusing junctions were taken using a high speed camera (Megaspeed) in combination with NIS Elements 3.2.

Initial tests were performed to investigate the suitability of a number of commercially available printer filaments. T-junction devices for droplet generation were created with 1mm diameter channels and the output of the device was observed. Sunflower oil and Water (containing Red Silverspoon dye) were delivered via syringe pumps (Legato 210, KD Scientific) to create water in oil droplet emulsions. Devices were fabricated using PLA filament (Ultimaker PLA 2.85mm), polyethylene terephthalate (PET) transparent filament (Taulman t-glase 3mm), a modified acrylonitrile butadiene styrene (ABS) transparent filament (Bendlay 3mm), and transparent PLA (Faberdashery, Crystal Clear 3mm). SL printed devices were produced using DETAX Luxaprint clear resin.

To characterise the fluidic interfacing and connections between modules, the maximum pressure that the connections could withstand before leaking was measured. This was achieved by blocking the outlet of one module while pumping water into the inlet of a connected module at a constant flow rate of 12ml/hr (Rheos 2000, Flux Instruments). The pressure required to drive the water was monitored throughout the experiment. Leaks were detected by visual inspection. Pressure data was recorded using the pump control software (Janeiro II 2.6).

Devices were produced via fused filament fabrication using an Ultimaker 2 printer. Devices were printed at a print speed of 30mm/s at a nozzle temperature of 215°C. The modules were printed with 100% fill density on a build plate that was heated to 70°C. Device schematics were designed in Solidworks 2013 before being converted to a print pattern using Simplify3D 2.2.2 software. Alternating 50μm thick layers were printed such that the pattern ran perpendicular and then parallel to the length of the device, enabling leak-free and transparent devices to be printed with clear Polylactic acid (PLA). 3D printed microfluidic devices were also produced stereolithographically using a Miicraft printer with 100μm layers and 25s curing time. Full print settings can be found in S1 Table .

Results & Discussion

To demonstrate the capabilities of FFF printed microfluidic devices, flow focusing junctions were created to form droplets in a controlled manner (Fig 2). Droplets of water in oil are readily formed. In addition, alginate droplets in oil, and oil droplets in a continuous phase of water and oleic acid were also demonstrated, illustrating the feasibility of consistent droplet formation in 3D printed microfluidic devices. Importantly, Fig 2 also shows that it is possible to observe the fluid flow within 3D printed PLA devices, either through the use of an embedded glass observation window for microscopy applications (Fig 2A), or through the use of semi-transparent PLA with sufficiently thin walls (Fig 2B and 2C).

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larger image TIFF original image Download: Fig 2. Extrusion 3D printed flow focusing junctions. A) Water in oil droplets formed in a PLA module that has a glass observation window embedded within it. B) Alginate solution droplets in sunflower oil generated in a semi-transparent PLA module. Oil flow rate: 3ml/hr, alginate flow rate: 1ml/hr C) Mineral oil droplets in water and 10mM oleic acid carrier phase. Generated on a semi transparent PLA device. Water flow rate: 4ml/hr, oil flow rate: 1.5ml/hr. Junctions A and C have inlet channel widths of 1mm with a 1.4mm wide outlet. Junction B has 600μm wide channels with a 900μm wide outlet. https://doi.org/10.1371/journal.pone.0152023.g002

The PLA flow-focusing junction with glass observation window (Fig 2A) was used to assess the reproducibility and control of water in oil droplet formation (Fig 3). Droplet size can be controlled by varying the ratio of the water and oil flow rates with a higher oil flow rate, relative to the water flow rate, leading to smaller droplets. The channel geometry and fluid properties define the minimum droplet size. The droplet generation frequency was investigated by increasing the total flow rate at a fixed oil:water flow ratio. This results in an increase in droplet frequency without altering the diameter. Droplets of a consistent diameter of 504μm (±18μm) were produced throughout the frequency test, as droplet production frequency increased from 1 to 10.4Hz. Frequency variation was very small as evidenced by the small fluctuation range of 0.2–1.2% (95% confidence interval), indicative of high stability operation. However, at the lowest flow rate of (1.5 ml/hr) 13% variation in production frequency was measured, this is due to this data being collected at a lower frame rate using a different camera than the other data points. The behaviour of this 3D printed flow focusing junction is consistent with previous studies of flow focusing junctions [35].

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larger image TIFF original image Download: Fig 3. Droplet data from a 3D printed flow-focusing junction ( Droplet data from a 3D printed flow-focusing junction ( Fig 2A ). Water droplets formed in mineral oil. A) Droplet diameter as the flow rate into each oil inlet is increased. The oil flow rate was increased as multiples of the water flow rate. Curve added to illustrate the trend. The error ranged from 1.3–3.3% based on 95% confidence level. B) Droplet frequency as the oil and water flow rates into each inlet are increased (fixed oil:water flow rate ratio of 2:1). For example, at the 10ml/hr data point both oil inlets are set to 10ml/hr whilst the water inlet is set to 5ml/hr. The error the error for the 1.5ml/hr data pointwas 13% due to data gathering using an alternative camera with a lower frame rate, the error for all the other data points ranged from 0.2–1.2% based on 95% confidence. In both A) and B) 10 droplets were measured for each data point (n = 10). Error bars not plotted as they are obscured by data points. https://doi.org/10.1371/journal.pone.0152023.g003

Fidelity Tests A criticism of FFF printing has been a lack of fidelity between the printed dimensions and the original CAD model [32]. Here, we investigate the correlation between measured circular cross-section of printed channels compared to the originally designed geometry, under a range of print conditions (50μm layers shown in Fig 4 Further printer fidelity analysis can be found in S4 Fig, showing different layer thicknesses and a comparison with SL printing.). High accuracy is possible with the right print parameters (in this case 50μm layer height) with both the height and width of the channel within 2.5% of the designed channel dimensions on average. Fig 4 illustrates the consistency of dimensions in all directions, although there is some unavoidable roughness resulting from the layer by layer nature of FFF printing as can be seen in Fig 4C. Although surface roughness was not considered for optimisation in this work, typical surface roughness was characterised by interferometeric measurement (S5 Fig) of gold sputter coated devices fabricated at 1200 mm min-1, with 50 μm layer deposition, as employed in the transparency tests. Over a 1.4 x 1.4 mm area material surfaces were found to have peak-to-peak variation of 16.81 μm. The nature of the deposition method leads to peaks at the interface between two extruded ‘strips.’ Further optimisation of the surface roughness may be achievable, either through further optimisation of print parameters or post fabrication solvent treatment, but was deemed beyond the scope of this study and unnecessary given the successful functionality of the printed devices. As printing resolution improves, and fabrication dimensions reduce, surface roughness optimisation may become of greater significance. PPT PowerPoint slide

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larger image TIFF original image Download: Fig 4. 3D printer fidelity measurements. a and b) Comparisons of measured and desired dimensions of 3d printed circular channels. Dotted line indicates perfect fidelity between the CAD model and the printed channel. a) Measured width of a horizontally printed channel using FFF printing with 50μm layers. b) Measured depth of a horizontally printed channel using FFF printing with 50μm layers. c) Image of a 1.4mm diameter channels fabricated using extrusion 3D printing. 3 measurements were taken for each data point. Error bars indicate 95% confidence level. https://doi.org/10.1371/journal.pone.0152023.g004 The fidelity measurements demonstrate that the minimum channel dimension achievable was 500μm. Whilst this is relatively large, it is suitable for many microfluidic applications. Currently, this is defined by the size of current printer nozzles. However, with smaller nozzles that have been released recently and advancements in printer technology anticipated, this limit should come down, further widening the application areas for extrusion printed microfluidics as channel dimensions reduce.

Stem Cell Encapsulation To demonstrate the practical utility of FFF 3D printed microfluidics for bio- and chemical applications, we created a device for the production of monodisperse alginate microspheres for the encapsulation of live stem cells. Such hydrogel cell encapsulation systems enable the three-dimensional culturing of cells and their easy manipulation [38]. In addition, cell encapsulation may provide a means for prolonged bio-therapeutic release in vivo [39]. The physical barrier provided by the encapsulating microsphere means that encapsulated stem cell systems are generating a large amount of interest for their potential application in tissue engineering and regenerative medicine therapies [40–42]. Monodisperse stem cell containing alginate droplets and calcium carbonate were created on the 3D printed devices within a sunflower oil continuous phase. Alginate droplets (800μm diameter) were gelled on entry into an acidified oil chamber. Partitioning of acid causes a reduction in pH within the alginate and liberation of Ca2+ from the calcium carbonate, giving rise to subsequent alginate gelation. After gelling the capsules were cleaned in culture medium and then cell viability assessed using a live/dead cell viability assay, with calcein-AM (green) indicating intracellular esterase activity within live cells, and ethidium homodimer-1 (red) fluorescence indicating loss of plasma membrane integrity in dead cells. Confocal imaging revealed that the vast majority of encapsulated cells remained live (Fig 7C). PPT PowerPoint slide

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larger image TIFF original image Download: Fig 7. Live dental pulp stem cells encapsulated within alginate droplets created on a 3D printed device. A) Monodisperse alginate capsules exiting the microfluidic device B) Image of an alginate capsule containing stem cells. C) Confocal z-projection showing live encapsulated stem cells stained green and dead cells stained red. The image shows the edge of a capsule. https://doi.org/10.1371/journal.pone.0152023.g007