Welcome to Chips & Tips

Welcome to Chips & Tips – a unique and regularly updated forum for scientists in the miniaturisation field from Lab on a ChipChips & Tips aims to provide a place where ideas and solutions can be exchanged on common practical problems encountered in the lab, which are seldom reported in the literature.

Do you

  • have problems with bubble formation when injecting your sample?
  • wish there was a quicker way to make prototypes?
  • find connecting chips to pumps and syringes problematic?

Or do you have your own tricks to overcome problems like these?

If so, then Chips & Tips is the forum to address your requirements!  Read the Tips below or see the author guidelines on how to submit your own today.

Chips & Tips is moderated by Glenn Walker (North Carolina State University).

Please note that Chips & Tips before April 2011 were originally published at www.rsc.org.

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Threadless chip-to-world connections on resin 3D printed microscale devices

Hannah B. Musgrove, Rebecca R. Pompano

Department of Chemistry, University of Virginia

Email: hbm8g@virginia.edu, rpompano@virginia.edu


Why is this useful?

With the continued adoption of 3D printed fabrication for microfluidics, the ability to easily connect non-elastomeric, resin 3D printed devices to fluidic tubing is an ongoing area of optimization. Common strategies to connect tubing to printed chips involve incorporating threading for use with luer lock adaptors, or embedding flanges or O-ring into devices.1–5 While useful for many types of inlets, standard luer adaptors are often > 10 mm in scale and thus too large for smaller chip designs. Additionally, the quality of 3D printed threading depends on the resolution of the printer and resin material, and the threads may wear down with repeated use. Embedding flanges1 or O-rings at connection sites6,7 may alleviate some of these print related limitations and increase the durability of the tubing ports. However, they also require additional manual fabrication steps including gluing or embedding parts and alignment. These extra steps are time-consuming and can increase fabrication inconsistencies. A simple option is to add a tubing-sized hole into a channel design which can be printed easily.6,7 This is similar in concept to dermal punching methods common for polydimethylsiloxane (PDMS)-based chips.1,4 However, while the soft, elastic properties of PDMS allows for a secure connection with tubing-sized holes, most 3D printed materials are more rigid, preventing conformal contact.

To address these issues, we developed a simple, threadless design for a raised port, that produces durable, 3D printed connection sites for tubing. We have used it successfully with both low and high viscosity vat-polymerization 3D printing resins (MiiCraft BV007a and FormLabs Clear, respectively) and compared it with, fused deposition fabrication of the same port using polylactic acid filament. This design is intended to decrease fabrication time and can be used with or without additional commercial adapter parts.

What do I need?

Basic materials are listed first, with the specific tools that we used listed in parenthesis.

  • Vat-polymerization 3D printer (MiiCraft P110Y, 385 nm 3D printer)
  • Resin (MiiCraft BV007a, or FormLabs BioMed Clear)
  • 3D design software (Fusion360 with Education License)
  • Microfluidic tubing, 1/16” OD or smaller (1/32” PTFE, 1/32” PEEK, and 1/16” silicon peristaltic tubing)
  • Optional: Ferrule lock rings (Idex Super Flangeless Ferrule, Cole-Parmer)

Measurement components for design

  • Included 3D software files or blueprint in Figure 1
  • Outer diameter (OD) of tubing being used
  • Estimate of print shrinkage and tolerance needed

What do I do?

  1. With 3D designing software, open the design to which you wish to add tubing connection ports.
  2. Above each port location, create a cylinder 2.75 mm tall with a 2.5 mm outer diameter ( 1). These are the standard dimensions that will fit with the optional ferrule locks in later steps.
  3. A hole in the center of the cylinder should be created to reach from the top of the cylinder down to the channel or port of the chip (“ID”, 1). The diameter of this hole should be customized to best fit the outer diameter of the tubing being used. (See next section for tips.)
  4. Filets should be added at three key locations ( 1).
    1. Filet 1 is at the edge of the inner cylinder hole and is set to 0.1 mm.
    2. Filet 2 is at the top, outer edge of the cylinder and is set to 6 mm.
    3. Filet 3 is at the base of the cylinder where the port meets the chip and is set to 4 mm.
  5. Once the raised port designs are added to the chip, use your printing software to orient and build supports for the print as needed ( 2A)
    1. Ports located on the “top” face of the chip (facing away from the baseplate during printing) do not require supports.
    2. Ports located on the “side” faces of the print do typically require at least one support for both the inner diameter and outer diameter overhangs
  6. Once printed and post-processed, tubing should fit directly into the ports, with close, conformal contact.
  7. Optionally, for further connection support, a ferrule lock can be used:
    1. Start by sliding the ferrule lock over the tubing before inserting the tubing into the raised port.
    2. Once inserted, slide the ferrule lock over the raised port ( 2B) to gently tighten the port around the tubing.

What else should I know?

The raised port designs were compared to a printed chip-to-syringe female thread luer inlet and a simple printed hole in the chip, and tested by driving flow through a simple microfluidic channel. We tested printability, ability to maintain a seal during multi-day fluid flow, and ability to be re-used without wearing down in two different resins (MiiCraft BV007a and FormLabs Clear). We also attempted to print the design using a fused deposition modeling (FDM) printer.

For printability, the raised port and simple hole designs printed successfully more often in resin printed chips than did luer lock ports, likely due to size and lack of threading.

For reuse, we found that the raised ports typically retained a good seal with the tubing through > 20 rounds of inserting, removing, and re-inserting tubing, in both FormLabs and MiiCraft resins. Use of the ferrule lock enabled additional re-use of resin printed raised ports. In comparison, simple embedded holes wore down ~7x reuse on average. This is likely due to faster wear around the edges from mechanical stress, from repeated use in addition to forces from material shrinkage that act on the hole by the surrounding printed shape.8

For durability of the connection, prolonged use of the ports was tested over one week by continually pumping saline solution through single-channel chips at 37°C at a rate of 1 µl min-1. It was found that the quality of the seal was primarily limited by resin stability under these conditions, rather than by mechanical wear at the port. Chips printed with BV007a resin began to leak and deform at both the port site and other channel locations after 48 hours, whereas chips with FormLabs Clear resin were stable for at least 5 to 7 days without any noticeable signs of leaking or wear at the ports. These results are consistent with our prior findings that BV007a prints are sensitive to extended heat treatment.8 We conclude that the port design is likely to yield a durable seal as long as the material is stable under the conditions of the experiment.

For additional stability, ferrule locks can also be added to further support and increase the duration of tubing-to-chip connections, especially for devices experiencing challenges with backpressure.

This design for ports works best with resin/vat-polymerization printers that have high print resolution (i.e. are able to print features around 1 mm in scale or lower) and with materials that are stiff yet slightly pliable, such as the listed resins. This design could in principle be adapted for high resolution fused deposition (FDM) printing but would again fair better with slightly pliable materials; we found that more rigid plastics such as polylactic acid (PLA) were difficult to combine with soft tubing, especially if FDM print tolerance is inconsistent.

Several design files are included for convenience on our dataverse website (https://dataverse.lib.virginia.edu/dataverse/PompanoLab). These files can be used “as is” to test fit with the tubing of interest or can be edited and added to an existing chip design.


The inlet design demonstrated in this work was found to be durable, versatile, and simple to fabricate for microfluidic chips printed with resin 3D printers. Though other systems work well for larger chips, the design shown here can be used when a smaller inlet connection is needed.


Research reported in this publication was supported by the National Institute of Allergy and Infectious Diseases under Award No. R01AI131723 and from the National Institute of Biomedical Imaging and Bioengineering under Award No. R03EB028043 through the National Institute of Health (NIH). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.


  1. Price, A. J. N., Capel, A. J., Lee, R. J., Pradel, P. & Christie, S. D. R. An open source toolkit for 3D printed fluidics. J. Flow Chem. 11, 37–51 (2021).
  2. van den Driesche, S., Lucklum, F., Bunge, F. & Vellekoop, M. 3D Printing Solutions for Microfluidic Chip-To-World Connections. Micromachines 9, 71 (2018).
  3. Au, A. K., Huynh, W., Horowitz, L. F. & Folch, A. 3D-Printed Microfluidics. Angew. Chem. Int. Ed. 55, 3862–3881 (2016).
  4. Anderson, K. B., Lockwood, S. Y., Martin, R. S. & Spence, D. M. A 3D Printed Fluidic Device that Enables Integrated Features. Anal. Chem. 85, 5622–5626 (2013).
  5. Weisgrab, G., Ovsianikov, A. & Costa, P. F. Functional 3D Printing for Microfluidic Chips. Adv. Mater. Technol. 4, 1900275 (2019).
  6. Bhargava, K. C., Thompson, B. & Malmstadt, N. Discrete elements for 3D microfluidics. Proc. Natl. Acad. Sci. 111, 15013–15018 (2014).
  7. Ji, Q. et al. A Modular Microfluidic Device via Multimaterial 3D Printing for Emulsion Generation. Sci. Rep. 8, 4791 (2018).
  8. Musgrove, Hannah. B., Catterton, Megan. A. & Pompano, Rebecca. R. Applied tutorial for the design and fabrication of biomicrofluidic devices by resin 3D printing. Anal. Chim. Acta 1209, 339842 (2022).
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VGA-inspired tubing connectors to operate microfluidic chips

Imre Banlaki, Horst Henseling, Henrike Niederholtmeyer

Max Planck Institute for Terrestrial Microbiology, Marburg

Why is this useful?

As microfluidic chips increase in complexity, more and more tubes need to be connected to a chip for operation. That is especially the case where many pneumatic valves (“Quake valves”) are included in the design of two-layer devices [1,2]. In many cases, these valves are actuated by a central, electronic switch board, driving solenoid valves [3]. The central switchboard makes each individual valve easily programmable for multiple applications.

The drawback of this versatile set up is the need to rewire all connectors to a new chip to initiate the experiment. This is time consuming and increases the likelihood of human error, especially if the setup is used by multiple researchers, who use different chips.

A plug and play solution would allow each researcher to have their personal end pieces connected to their chips and interfacing with the switchboard. This allows preparation on the lab bench and facilitates the change from one experiment to another.

To that end, we designed a 3D printable mould to cast PDMS trapezoids with 8 integrated tubes (Fig. 1). The shape and tube-pattern make the connector unambiguous, similar to a VGA or D-sub electronics connector. A D-sub microfluidic connector of similar design has been published before, however it relies on actual, disassembled D-sub connectors which have gone out of fashion [4].

What do I need?

PDMS Sylgard, Dow Corning
Resin 3D printer or other means of fabrication
Tygon tubing (0.51 x 1.52 mm) or other as preferred
Blunt dispensing needles 23G (0.43 x 0.64 mm)
Glass petri dish
4x M3x20 screws and nuts

Figure 1 Blueprint of the pieces to fabricate and assemble the mould. Measurements in mm.

How do I do it?

  1. Fabricate the mould

3D print or mill the mould from a thermally stable resin, polymer or noncorrosive metal. We have shared our design on Metafluidics.org. (https://metafluidics.org/devices/vga-inspired-tubing-connector/)

Drill the holes in the trapezoid plate. Make sure the hole-array is axisymmetric by measuring each placement from the middle.

  1. Prepare the mould

Remove 8 metal pins from the plastic of the blunt dispensing needle tips. Cut 8 approx. 3-5cm Tygon tubing pieces and push them onto the end of the metal pins that was inserted in the plastic.

Insert the pins + tubing into the holes of the trapezoid plate (Fig. 2A).

Place the plate into the notch of the mould with the pins sticking out towards the shorter end, and screw the top plate on (Fig. 2C+D).

Figure 2 Assembly steps of the mould. A) Pins and tubing inserted in the trapezoid plate to fabricate a plug. B) Pins and tubing inserted in the plate to fabricate a socket. C) Plate, with pins and tubing, placed into the grove before adding the top plate. D) Fully assembled mould with flush pins sticking out. E) Vertical placement of assembled mould ready to be filled.

  1. Cast the PDMS

Mix 10g PDMS resin (1:10 as per supplier instruction).

Place the mould vertically so the hole is on top (Fig. 2E). Make sure the pins are flush with the bottom (use the petri dish). Arrange the bend of the tubing in a way that makes it easy to identify which tube belongs to which pin.

Cast the PDMS until at least 50% of the mould is filled.

Cure in the oven at 80°C for 30-60 mins.

  1. Remove the connector

After curing, unscrew and remove the top plate. Carefully remove the PDMS piece including the trapezoid plate from the mould. Now, you should be able to slide the plate off the pins.

The tubing on the back end is now ready to be connected to tubing from a control board by individual bridging with metal pins. The front end forms the quick connector for 8 lines.

  1. Setup for receiver socket

To cast a receiver socket, insert 8 metal pins without tubing into the trapezoid plate. Now connect some tubing to the pins so that the plate is sandwiched between pin and tubing (Fig. 2B). Insert the assembly into the mould as before and cast the PDMS. After curing and removal of the piece, the metal pins can be pulled out leaving only the tubing embedded within the PDMS.

Figure 3 Plugs and socket A) after removal from the mould and B) connected to the switch board. C) Switch board with 24 valves clustered into 3×8 (A, B, C) control lines. D) Three plugs (A, B, C) connected to sockets.

What else should I know?

Curing without degassing may create bubbles, and depending on the resin for 3D printing the surface may remain somewhat sticky. This will not affect functionality. The stickiness can be avoided by placing the mould in isopropanol for a few days. Adding pigment to the PDMS could be used to colour code each plug. Expanding the design for more lines should be possible but connecting will be increasingly difficult.

Uncareful connection, puncturing the PDMS, may clog a pin.

When milling the device, instead of 3D printing, it is advised to change the plate shape and notch to a semi-circle and mill on the face to guarantee a flush fit of the plate.


This work was supported by Deutsche Forschungsgemeinschaft grant NI 2040/1-1.


  1. Unger MA, Chou H-P, Thorsen T, Scherer A, Quake SR. Monolithic Microfabricated Valves and Pumps by Multilayer Soft Lithography. Science. 2000;288: 113–116. doi:10.1126/science.288.5463.113
  2. Niederholtmeyer H, Stepanova V, Maerkl SJ. Implementation of cell-free biological networks at steady state. Proc Natl Acad Sci. 2013;110: 15985–15990. doi:10.1073/pnas.1311166110
  3. Brower K, Puccinelli RR, Markin CJ, Shimko TC, Longwell SA, Cruz B, et al. An open-source, programmable pneumatic setup for operation and automated control of single- and multi-layer microfluidic devices. HardwareX. 2018;3: 117–134. doi:10.1016/j.ohx.2017.10.001
  4. Scott A, Au AK, Vinckenbosch E, Folch A. A microfluidic D-subminiature connector. Lab Chip. 2013;13: 2036–2039. doi:10.1039/C3LC50201E
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A simple Matrigel cooling setup for optimal cell seeding of microfluidic devices

Torben Roy

MeBioS biomimetics group, KU Leuven, Willem de Croylaan 42, 3001 Leuven, Belgium

Why is this useful?

Some organ-on-a-chip models require seeding of cells suspended in an extracellular matrix (ECM) such as Matrigel.1-3 When seeded inside microfluidic channels, cells experience shear stresses due to mechanical forces of the fluid which can have a negative effect on the viability of the cells.

Matrigel starts to polymerize at a temperature above 10 °C, which hinders the injection of the ECM inside microfluidic channels due to an increase in viscosity. An increase in pressure from the pipette or pump is then required and if not met, the microfluidic channel will not be fully deposited as intended (Figure 1). Precautions including the cooling of the ECM solution, pipette tips and microfluidic chip need to be taken to ensure proper deposition.

A study by Kane et al. found that a temperature between 8 °C and 10 °C is ideal for cell seeding as in this temperature range a minimal shear stress is observed.4 Below 8 °C an increase in shear stress is observed due to temperature induced liquefaction, while above 10 °C polymerization induces a higher rate of shear stress. Thus to ensure maximum cell viability, it is recommended to seed a microfluidic device with a cell-Matrigel solution in the 8 – 10 °C range. The use of an ice bath or current commercial coolers does not allow for stable cooling in that temperature range.

A new method is presented that allows for cooling of a cell-Matrigel solution in the optimal temperature window (8 – 10 °C). The cooling setup is composed out of a heating block (from a dry block heater) (or CoolRack™ from Corning), ice packs and a thermometer. Ice packs are less efficient at cooling compared to the ice bath and can be added and removed until a desired stable temperature is obtained. While the metal heating block (common lab equipment) allows for a homogeneous spread of the temperature.

Figure 1 – Unsuccessful deposition of a microfluidic chip with Matrigel. The Matrigel solution did not fill the entire main channel as intended due to premature polymerization.


What do I need?

  • Heating block (e.g. modular heating block for vials, VWR) or CoolRack™ (Corning)
  • Ice packs
  • Cooling element
  • Thermometer
  • Matrigel® Basement Membrane Matrix, Phenol Red-Free, LDEV-Free (356237, Corning)
  • Eppendorf tube
  • Pipette
  • Pipette tips


How do I do it?

  1. Prepare the cell-Matrigel solution in a sterile environment, aliquot in an Eppendorf tube and place on ice.
  2. Place the heating block, (autoclaved) pipette tips and (autoclaved) microfluidic chip inside the fridge to cool to 4°C.
  3. Remove the heating block from the fridge, place it on a cooling element (stable surface) and surround the heating block with ice packs.
  4. Place the Matrigel-cell solution and thermometer inside the heating block holes.
  5. Add or remove ice packs until a stable temperature in the 8 – 10°C range is achieved.
  6. Remove the microfluidic chip from the autoclave insert and place it on a cooling element. Inject the Matrigel-cell solution using cooled pipette tips.


Figure 2 – Cooling setup. A heating block is surrounded by ice packs to reach a stable temperature in the 8 – 10°C temperature range.


  1. Wang Y, Wang L, Guo Y, Zhu Y, Qin J. Engineering stem cell-derived 3D brain organoids in a perfusable organ-on-a-chip system. RSC Advances. 2018;8(3):1677-1685.
  2. Trietsch S, Israëls G, Joore J, Hankemeier T, Vulto P. Microfluidic titer plate for stratified 3D cell culture. Lab on a Chip. 2013;13(18):3548.
  3. Moreno E, Hachi S, Hemmer K, Trietsch S, Baumuratov A, Hankemeier T et al. Differentiation of neuroepithelial stem cells into functional dopaminergic neurons in 3D microfluidic cell culture. Lab on a Chip. 2015;15(11):2419-2428.
  4. Kane K, Moreno E, Lehr C, Hachi S, Dannert R, Sanctuary R et al. Determination of the rheological properties of Matrigel for optimum seeding conditions in microfluidic cell cultures. AIP Advances. 2018;8(12):125332.
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A simple, efficient, and cost-effective spin coater: A waste to wealth approach

S.J.Samuel Justin1, P.Wilson1

1Department of Chemistry, Madras Christian College

Email: sam310just@gmail.com, wilson@mcc.edu.in


Why is this useful?

Spin-coating is a highly reproducible, simple, time-efficient, and cost-effective coating technique is widely employed technique for the fabrication of thin-film coatings over large areas with smooth and homogeneous surfaces [1]. It has been widely used for the production of monolayer- and multilayer-thin coatings, freestanding (FS) nanosheets and membranes, for various industrial and biomedical applications, e.g. mitigation of corrosion [2], wound dressings, cell culture substrates, and as drug delivery devices.

A tip to develop portable spin coaters by recycling computer fans and mobile phone wall chargers was previously presented [3]. Using adhesive strips to secure a metal alloy sample on the center of the computer fan often slips and knocked down while trying to remove the sample after the coating process. This affects the coating and thereby a proper substitute to hold the sample tight during the spin and at the same time easier to remove it after the coating is essential. In addition, the sample spinning at high rpm levels expels the excess coating substrate away from the system which has to be addressed in order to prevent the contamination of the surroundings. In this regard an improved form of portable spin coater is proposed and presented.


What do I need?

Parts for the spin coater

  1. Polycarbonate container, Dimensions 155 X 155 X 60 mm
  2. Panel cooling fan 120 x 120 x 38 mm with 2800 rpm
  3. Acrylic support rod of length 15 mm and diameter of 12 mm
  4. Faucet water tap adapter
  5. Black cap of 5mL clear round glass bottle – 4 pieces
  6. 8 pieces set of 3mm diameter round head 20 mm bolt and nut
  7. Power cable

Parts and chemicals for the specific examples

  • Nitro cellulose lacquer
  • Aluminium alloy 15mm x15mm x6mm


What do I do?

Assembling of spin coater

  1. The acrylic rod is fixed at the centre of the fan using instant cyanoacrylate ester adhesive (Fig.a)


  1. A hole of 15 mm dia. is drilled at the centre of the container base and 2.8 mm hole at the four corners of the container (Fig.b).
  2. The container is placed over the fan and the corners are fixed using bolt and nut (Fig.c)
  3. The stem of the faucet water tap adapter was cut and the hole is enlarged to 12mm dia. It is then fixed using instant adhesive on the top of the acrylic rod (Fig.d, e & f)

  1. The black caps were drilled at the center to create a hole of 2.8mm dia. The bolt is inserted from below each cap and then tightened with the base of the cooling fan at the four corners (Fig. g and h).
  2. The power cable is attached to the fan (Fig. i)

  1. The lid of the container is drilled at the center to 15mm dia. which acts as a space for dropping the coating material (Fig. j and k) and the final product is displayed in (Fig. l).
  2. The liquid containing the coating material (pigmented nitrocellulose lacquer) is placed on top of                 the substrate (Aluminium alloy) (Fig. m) by using a (micro) pipette (Fig. n.)
  3. The fan is turned on and the substrate is spin coated for about 30 seconds (Fig. o) (time can vary    depending on the substrate viscosity and coating thickness required). The coated substrate (Fig. p) is subjected to further evaluation.




[1] Moreira, Joana, A. Catarina Vale, and Natália M. Alves. “Spin-coated freestanding films for biomedical applications.” Journal of materials chemistry B 9, no. 18 (2021): 3778-3799.

[2] Telmenbayar, Lkhagvaa, Adam Gopal Ramu, Daejeong Yang, Minjung Song, Tumur-Ochir Erdenebat, and Dongjin Choi. “Corrosion resistance of the anodization/glycidoxypropyltrimethoxysilane composite coating on 6061 aluminum alloy.” Surface and Coatings Technology 403 (2020): 126433.

[3] A second life for old electronic parts: a spin coater for microfluidic applications, https://blogs.rsc.org/chipsandtips/2018/04/25/a-second-life-for-old-electronic-parts-a-spin-coater-for-microfluidic-applications/ (accessed August 2020)

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The Name is Bond – Heat Bond: Using a Heated Lamination Press for Thermoplastic Thin Film Bonding

Travis Swaggard1, Johanna Bobrow1, Peter Carr, Isabel Smokelin1, Todd Thorsen1, Christina Zook1, David Walsh1

1MIT-Lincoln Laboratory, 242 Wood St, Lexington, MA 02421


Why is this useful?

Have you ever tried to bond a thin clear bottom to your custom thermoplastic microfluidic device for high magnification microscopy? The bonding process can be cumbersome and relies on the heating temperature, the pressure placed on the device assembly, surface chemistry, as well as many other factors.1 We have developed a method that utilizes a heated laminator press and a UVO oven, which together cost less than hiring an external vendor who will likely charge per design and item manufactured. Currently, manufacturing bonded devices in-house can be difficult, expensive, and time consuming.

The new paradigm in microfluidic device prototyping using high-resolution 3D-printing has significantly reduced the barrier to participate in microfluidics, which traditionally required dedicated infrastructure and specialized expertise.2 However, once the design-build-test prototyping cycle has gone beyond the 3D-printing stage, additional prototyping in thermoplastics (e.g. CNC milling) is typically required for eventual scale-up using traditional hot embossing or injection molding manufacturing techniques. One of the critical challenges at this stage is placing optically-clear bottoms below the thermoplastic microfluidic device for optical interrogation, while withstanding fluidic flow and pressure. For example, adhesive bonding such as using double-sided tape can be tricky and tedious to align, and solvent-based adhesives can be messy or challenging to avoid impacting thin-film clarity, and even pose environmental safety concerns.

We have developed an inexpensive bonding protocol which can utilize almost any commercially available heated machine press to mitigate the aforementioned drawbacks, and bond layers of thin (125 micrometer) acrylic film to any flat acrylic microfluidic device surface. This allows for a strong bond that can provide a clear bottom for high-resolution optical interrogation, such as using a confocal microscope, for example. New low-cost commercial UVO ovens (Jelight Model 18 – $3,000) and heated laminated presses (Nugsmasher $500 [Mini] – $7,000 [Pro Touch]) have helped enable democratization of the thermoplastic film bonding process.

Here is a detailed walkthrough for the preparation and assembly of these devices. We have included photos and figures from each step for reference and convenience.

What do I need?

  • Thin, optically clear acrylic film (e.g. Röhm GmbH Acrylite Film 99524 or similar)
  • Microfluidic device from cast acrylic (e.g. Protolabs – CNC Machined)
  • Scissors
  • Oven mitts (or equivalent)
  • Multipurpose Neoprene rubber sheet 6”x 6” x 1/16” (or equivalent heat-safe separator)
  • Heated lamination press (Premiere Manufacturing SKU: 714343996295 or similar)
    • Must apply pressures between 30 and 50 PSI (roughly finger-tight, part won’t move if pushed by a finger)
    • Must be able to heat up to 118 °C

How do I do it?

  1. Using scissors, cut piece(s) of thin, optically clear acrylic film slightly larger than your acrylic device (this will eliminate any alignment steps, you can cut the remaining excess after the device has been pressed). Your film should be about half an inch oversized on all edges of the device to ensure that it is completely covered
  2. Place the cutout of acrylic film(s) and plastic device(s) into the UVO oven so they are not touching and run for eighty minutes (see Fig.1). You can also perform oxygen plasma bonding alternatively; however, as that bond weakens over time, it’s important to perform the bonding immediately before the treatment. (Note: Be sure to put the bonding surfaces face-up in the UVO machine for best results.)
  3. Preheat laminated press to 118 °C (or about 250 °F). It may take up to twenty minutes to get up to temperature and an hour total to stabilize at the set point (see Fig.2).
  4. Very carefully (press will be very hot) place thin film on bottom plate and plastic device on top of film (use oven mitts or tongs for safety).
  5. Place a neoprene cover sheet over plastic coupon/film assembly. This will provide cushioning between the top press and your device and prevent any possible cracking or pitting from the top press (see Fig.3).
  6. Using the lever crank in the back, apply manual pressure until the machine registers a reading of 30-50 PSI (or roughly finger-tight, so the part won’t move if pushed by a finger or tong).
  7. Let device sit at steady pressure and heat, hold for ten minutes.
  8. Turn off heat but keep the device under pressure.
  9. Once heat returns to near room temperature, release pressure and remove device (Note: This may be overnight). Once complete, the devices should be effectively bonded and show optically clear bottoms with no aberrations (see Fig.4).

What Else Should I Know?

Before running this protocol, check your heated laminator press for scratches or pitting that could imprint on your film. If there are any significant scratches, these might become imprinted on your film and cause uneven bonding later in the process. A steel stainless steel sheet with a polished finish can be added on top of the bottom plate using thermal paste.

UVO treatment of plastics will create an oxide layer on the surface, rendering it more hydrophilic. The generation of oxygen species on the surface of plastics will reach a plateau after about sixty minutes.3

Almost any thermoplastic material4 can be bonded to the same type of material using the rule-of-thumb of heat being 5 °C over glass temperature, however it is worth noting that identical materials will bond most effectively, for example cyclic olefin polymer (COC) bonds to COC material straightforwardly, whereas COC will not bond as well to dislike material such as polycarbonate.

Do not overexpose your film and device to UVO (>sixty min) as this may cause significant yellowing and irreversible changes to the chemical makeup of the acrylic. Apply UVO as directed in this protocol. Some yellowing may happen, but in our hands, this does not create any noticeable background shading or autofluorescence.

Make sure not to put too much pressure on the film and device during the bonding process to ensure that devices are not warped, deformed or cracked, use only the pressure recommended in this protocol.

Always allow the heat from the laminated press to return back to room temperature, do not try to remove the device from the press before returning to room temperature as this will potentially produce an incomplete bonding.

If the pressure required for this bond causes deformation of critical features, consider applying ethanol to reduce surface glass temperature for a lighter pressure bond.5


While this publication discusses bonding to CNC-milled devices, there is potential for devices made from alternative fabrication techniques such as xurography or laser cutting to be bonded using a modified version of this protocol as well.6



We would like to acknowledge Chris Phaneuf, Ph.D. from Sandia National Laboratories as well as Edge Embossing (Charlestown, MA) for their assistance and expertise with helping us create this protocol. This work was funded by NIH/NCATS through an Interagency Agreement with MIT-Lincoln Laboratory as well as by NIH/NIBIB via grant award number 1R01EB025256-01A1.

© 2022 Massachusetts Institute of Technology.
Delivered to the U.S. Government with Unlimited Rights, as defined in DFARS Part 252.227-7013 or 7014 (Feb 2014). Notwithstanding any copyright notice, U.S. Government rights in this work are defined by DFARS 252.227-7013 or DFARS 252.227-7014 as detailed above. Use of this work other than as specifically authorized by the U.S. Government may violate any copyrights that exist in this work.




  1. Putting the Lid on Microfluidics [Internet]. Microfluidics. 2017 [cited 2022 Jan 7]. Available from: https://microfluidics.technicolor.com/wp-content/uploads/2021/12/Microfluidic-Chip-Bonding-Considerations-and-Case-Studies.pdf
  2. Bhattacharjee N, Urrios A, Kang S, Folch A. The upcoming 3D-printing revolution in microfluidics. Lab on a Chip. 2016;16(10):1720-42.
  3. Lin TY, Pfeiffer TT, Lillehoj PB. Stability of UV/ozone-treated thermoplastics under different storage conditions for microfluidic analytical devices. RSC advances. 2017;7(59):37374-9.
  4. Jiang J, Zhan J, Yue W, Yang M, Yi C, Li CW. A single low-cost microfabrication approach for polymethylmethacrylate, polystyrene, polycarbonate and polysulfone based microdevices. RSC Advances. 2015;5(45):36036-43.
  5. Harriet Riley, Development Editor. (2017, November 1). A solvent-based method to fabricate PMMA microfluidic devices – Chips and Tips. https://blogs.rsc.org/chipsandtips/2017/11/01/a-solvent-based-method-to-fabricate-pmma-microfluidic-devises/?doing_wp_cron=1625578271.8969628810882568359375 (Accessed November 2021)
  6. Burgoyne, F. (2010, May 17). Fast-iteration prototyping and bonding of complex plastic microfluidic devices – Chips and Tips. https://blogs.rsc.org/chipsandtips/2010/05/17/fast-iteration-prototyping-and-bonding-of-complex-plastic-microfluidic-devices/?doing_wp_cron=1625578517.8055989742279052734375 (Accessed November 2021)
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Daniel Alcaide Martín, Jean Cacheux, Sergio Dávila & Isabel Rodríguez

Madrid Institute for Advanced Studies in Nanoscience (IMDEA Nanoscience), Ciudad Universitaria de Cantoblanco, C/Faraday 9, Madrid 28049, Spain

1- Why is this useful?

Microfluidic devices need to be connected to fluidic pumps for regulation of the flow during the device operation1. Connecting and disconnecting devices is a tedious and time-consuming operation that often causes air bubbles which are detrimental for the fluidic experiment and a real nuisance for the time it takes to eliminate them.

Furthermore, in microfluidic experiments dealing with biomolecules or cell cultures, volumes are of concern as these materials are typically limited and/or costly. Hence, it will be very useful if the reagent filling or replacement process and the connecting and disconnecting operations to microchips are minimized to avoid both bubbles and reagents waste. Reducing the reservoir volume to the volumes needed for the experiment minimizing dead volumes will also allow saving expensive reagents.

With this aim, we have designed a fixture to make practical fluidic connections to a microchip from a pressure controller for fluidic control and device operations. It allows for easy opening and closing operations and for easy re-filling or replacement of the reagents into the microchannels without moving any tubing connection.

Here, we present a dual connector cum reservoir fixture as a practical and effective means to making fluidic connections onto polydimethylsiloxane (PDMS) microfluidic based chips. The fixture is completely built by stereolithography (SLA) 3D printing and includes two components: a piece including a reservoir with an O-ring slit and a cone shaped outlet as chip connector and, another piece that closes the reservoir and has a cone shaped inlet or air pressure connector.

This Chips and Tips builds on a previous approach,2 dealing with microchip fluidic fixtures using magnets. However, in this case, the reservoirs and release connection are moved off the chip which would be more practical to work with the chip on the microscope stage for real time observations. Moreover, it is adaptable for any chip design and particularly for microchips made in soft PDMS.


2- What do I need?

  • 3D design software.
  • SLA 3D printer.
  • Cubic magnets.
  • O-rings.


3 – What do I do?

We first digitally design the parts of the dual connector: the reservoir-chip connector and the reservoir-seal air pressure adaptor. See the drawing in Figure 1. The corresponding F3D files can be here downloaded.

In our design, the reservoir sits aside from the PDMS chip and it is connected to it through a standard tube fitted on to the connecting outlet. The reservoir-seal encloses the reservoir. Four permanent magnets are inserted in each of the two components to produce a magnetic compressive force onto the O-ring and a tight seal to allow for applying a controlled pressure to cause the reagent to flow at the desired rate into the microfluidic chip.

Once the components are printed, the magnets are inserted into the lateral slots. Then, the plastic tubes are connected to the chip inlets and outlets and to the reservoir chip connector. The reservoir is filled with the correct volume of reagent (in this design is 0.2 ml) and the reservoir- seal piece is placed on top. Finally, through the air pressure inlet, tubing is connected to a pressure controller. When air pressure is applied, the reagent flows through the chip. The closing system formed by the O-ring seal and magnet force is able to handle at least 1 bar working pressure without any leakage.

Figure 2 – Fluidic system set-up showing .two dual connector-reservoir fixtures connected to a microchip and to a pressure controller.


This work was performed within the framework of the EVONANO project funded by the European Union’s Horizon 2020 FET Open programme under grant agreement No. 800983.


  1. Interfacing of microfluidic devices – Chips and Tips. https://blogs.rsc.org/chipsandtips/2009/02/27/interfacing-of-microfluidic-devices/.
  2. Reusable magnetic connector for easy microchip interconnects – Chips and Tips. https://blogs.rsc.org/chipsandtips/2011/06/27/reusable-magnetic-connector-for-easy-microchip-interconnects/.


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Solvent Extraction of 3D Printed Molds for Soft Lithography

Jonathan Tjong1, Alyne G. Teixeira1 and John P. Frampton1,2

1School of Biomedical Engineering, Dalhousie University, Halifax, NS, Canada

2Department of Biochemistry and Molecular Biology, Dalhousie University, Halifax, NS, Canada

Why Is This Tip Useful?

Casting polydimethylsiloxane (PDMS) in molds produced from additive manufacturing (i.e., 3D printing) enables rapid prototyping of parts with microscale features without the need for conventional photolithography. Whereas conventional photolithography followed by soft lithography involves the use of silicon substrates and photomasks, which can be costly and may require special preparation and processing (e.g., application and removal of photoresist in a clean room environment), the emergence of stereolithographic 3D printers allow for the rapid manufacture of masters for PDMS casting in almost any laboratory space. Stereolithographic 3D printers use photopolymer resins that when cured can withstand temperatures as a high as 200 °C without plastic deformation, which occurs with thermoplastics such as polystyrene that have glass transition temperature around 95-105 °C (Lerman et al.). The ability to withstand such high temperatures opens the possibility for PDMS and other silicone elastomers to be cured quickly and also provides the possibility for greater control of the mechanical properties of the elastomers (Johnston et al.). However, the components within the 3D printed resin mold such as residual photoinitiators and unreacted oligomers may interfere with the curing of PDMS, resulting in incomplete curing at the interface of the printed mold and the PDMS part. Here, we demonstrate a simple treatment to remove these unwanted materials through solvent extraction.

What Do I Need?

  • Stereolithographic 3D printer and appropriate resin
  • Leak-proof, sealable container large enough to hold the 3D-printed mold
  • Dishwashing detergent
  • 95% ethanol or isopropanol
  • Orbital shaker table
  • Uncured PDMS base and curing agent (10:1 w/w)
  • Post-curing UV lightbox

What Do I Do?

  1. After cleaning and post-curing of the 3D-printed mold in the UV lightbox, place the mold in the container and add enough solvent to submerge the part.
  2. Seal the container and leave on a shaker table for 24 hours.
  3. Discard the old solvent and add new solvent. Seal and agitate for another 24 hours.
  4. Remove the part from the solvent and allow to air dry at room temperature.

What else should I know?

The exact composition of photopolymer resins for stereolithography may vary significantly between different manufacturers; therefore, it may be necessary to adjust the protocol (e.g., the type of solvent). In addition, larger prints will likely require more solvent and a longer duration of solvent extraction to account for the increased migration time of unwanted components from the print to the free solvent.

To demonstrate our procedure, we printed two sets of 5 identical molds with basic geometric features. One set used 1 mm thick outer walls, while the other set used 3 mm thick outer walls (Figure 1). The molds were designed using Onshape (Onshape, Cambridge, MA) CAD software and printed using a B9Creator v1.2 (B9Creations, Rapid City, SD) stereolithographic 3D printer. For all prints, we used B9-R2-Black resin from B9Creations. This resin is documented by the manufacturer as having a heat deflection temperature of 65 °C at 0.45 MPa determined through ISO 75-1/2:2013 standards (B9Creations). As no significant mechanical load would be placed on the resin molds (with a maximum depth of 6 mm for the PDMS chamber), we decided this resin was suitable for our test molds. After printing, excess resin was removed from the molds by submerging and agitating in an approximately 1:10 mixture of Dawn dishwashing detergent and water in a 1 L container. This was followed with additional cleaning by rinsing with excess isopropanol using a wash bottle until no visible evidence of uncured resin was present on the surface (approximately 10-20 mL per part). The molds were then post-cured in a UV lightbox for 20 minutes.

Once post-cured, the molds were each placed in new, 15 mL polypropylene centrifuge tubes (Falcon® Corning, Corning, NY) with 10 mL of the test solvents (reverse osmosis-treated (RO) water, isopropanol, 95% ethanol, or methanol), and exposed to the two 24-hour extraction procedures listed in the “What Do I Do” section. After extraction, the molds were briefly rinsed with RO water and allowed to air dry for 1 hour. Then, approximately 0.4 mL or 1 mL of premixed 10:1 uncured PDMS and curing agent were added to the 1 mm and 3 mm molds, respectively. The PDMS parts were heat-cured in a dry oven at 65 °C overnight. The cured PDMS parts were then carefully removed from the mold using a stainless-steel spatula. Images were taken using the default settings on an iPhone 7 camera.

Resin molds that did not undergo extraction or that underwent extraction in water or methanol produced PDMS parts with major defects. For these extraction conditions, fragments of semi-cured and traces of uncured PDMS remained at the PDMS-mold interface (Figure 2A-C). Extraction with methanol appeared to weaken the cured resin, with significant softening and the appearance of cracks on these molds (Figure 2C), and while the cured PDMS easily released from the methanol-extracted molds, this left artifacts on the PDMS surface. We found that resin parts pretreated with either isopropanol or 95% ethanol performed well as molds for PDMS. The cured PDMS parts easily released from the substrates, with no visible traces of uncured PDMS (Figure 2D-E), and the PDMS parts we retrieved cleanly replicated the features of the resin mold (Figure 3). In addition to producing defects in the mold itself, extraction with methanol also led to bubbles forming in the PDMS as it cured (Figure 3C). Overall, PDMS casts were most easily released from the 1 mm-thick molds compared to the 3 mm-thick molds, but this may simply be due to the lower aspect ratio (h/l) of the 1 mm-thick molds.

Take Home Message

When casting PDMS parts from molds produced by stereolithography, incomplete curing and defects in the PDMS part can be minimized by extracting residual photo-initiators and oligomers present in the mold using either isopropanol or 95% ethanol.





B9Creations. Black Resin Material Properties. 2018, pp. 9–11, https://cdn2.hubspot.net/hubfs/4018395/Material Data Sheets/B9Creations Black Material Properties.pdf.

Johnston, I. D., et al. “Mechanical Characterization of Bulk Sylgard 184 for Microfluidics and Microengineering.” Journal of Micromechanics and Microengineering, vol. 24, no. 3, 2014, doi:10.1088/0960-1317/24/3/035017.

Lerman, Max J., et al. “The Evolution of Polystyrene as a Cell Culture Material.” Tissue Engineering – Part B: Reviews, vol. 24, no. 5, 2018, pp. 359–72, doi:10.1089/ten.teb.2018.0056.




Figures and Legends


Figure 1. Design features of molds produced by stereolithography. Top panels in (A) and (B) are the Onshape renderings. Bottom panels in (A) and (B) are parts printed in the B9Creations B9-R2-Black resin.

Figure 2. Molds produced by stereolithography following extraction in various solvents. Fragments of partially cured PDMS and uncured PDMS remain on the surface of molds that have not undergone extraction, as well as those extracted in RO-water and methanol. Isopropanol and 95% ethanol extraction produce molds that can be re-used numerous times for PDMS curing.

Figure 3. PDMS parts obtained from curing in resin molds extracted using various solvents. Extraction of residual photo-initiators and oligomers present in the mold prior to soft lithography using either isopropanol or 95% ethanol results in clean PDMS parts that are free of defects.

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Cutting the cords: Two paths to well-plate microfluidics

Sara E. Parker1 and Peter G. Shankles2, Maddie Evans1, Scott T. Retterer1,2,3

1 Biosciences Division, Oak Ridge National Laboratory, Oak Ridge, TN

2 The Bredesen Center, The University of Tennessee, Knoxville, TN

3 The Center for Nanophase Materials Sciences

Why is this useful?

Even simple microfluidic devices often require complex and expensive pumping and valving systems for accurately metering and controlling fluid flow. This often necessitates substantial and time-consuming set-up, and sometimes make these chips unwieldly and difficult to image. It can also represent a significant departure from the rather straight forward process of pipetting fluids from one small volume to another, making adoption by non-microfluidic experts unlikely. However, the development of well-plate microfluidics1,2 provides a high throughput, simplified method for studying fluid exchange and shear flow, while minimizing the set-up and need for multiple fluid connections. Creating an interface between the polystyrene (PS) plate and the polydimethylsiloxane (PDMS) fluidics presents the largest obstacle in creating these hybrid devices. Khine et al.1 utilized pressure to create a tight interface while Conant et al.2 adhered the surfaces using glue, but neither elaborated on their techniques and in practice small changes in the process can result in failed devices. Here, two techniques are detailed on consistently creating an effective interface between the well-plate and microfluidics. Resulting in individual wells that are interconnected via custom microchannels in a PDMS device attached to the bottom of the well-plate. Reagents are then added to wells and, driven through the underlying channel network into an outlet well via hydrostatic pressure or a pressure control system3,4.

With the use of this platform, flow can be introduced into traditional well-plate studies allowing various physiological conditions to be more closely mimicked. Further, the compatibility of these custom devices with well-plate microfluidic control systems provides the opportunity to precisely and dynamically control experimental conditions including temperature, pressure, and gas environment3,4. The use of multi-well plates also allows for multiple devices to be bonded in parallel to the same plate, increasing throughput without increasing the complexity of the control system5. Additionally, the familiarity and ubiquity of the well-plate platform provides a familiar platform for technical professionals within the lab and is automatically compatible with the host of microscope stage attachments already available for use with conventional well-plates.

Well-plate microfluidic fabrication has been shown with a pressure seal between the microfluidics and well-plate1 as well as gluing the two together2. This work builds off these ideas by detailing bonding with a liquid adhesive or chemical activation and bonding. The process of bonding customized PDMS devices to well-plates for well-plate microfluidics has only been vaguely described previously5,6. Herein, we present two approaches that utilize either (3-Aminopropyl)triethoxysilane (ATPES) to modify the surface of the PS well-plate to bond with plasma treated PDMS, or uncured PDMS to act as a glue between the PS and PDMS surfaces7. While the APTES modification provides a stronger bond without adding additional material, the uncured PDMS bonding procedure requires less pressure, avoiding any distortion of nanoscale features. An overview of the process is shown in Figure 1:

Figure 1 – Diagram of the fabrication process with the APTES process above and PDMS glue below.

What do I need?


  • PDMS device replica with inlets and outlets designed to align with a well-plate
  • 48 well-plate; Flat-bottomed, non-tissue culture treated
  • Isopropyl alcohol (IPA)
  • Coverslip or slide large enough to cover channels
  • X-Acto knife
  • Scotch tape

APTES bonding only

  • Deionized water
  • Hard rubber brayer
  • Sealable plastic container

PDMS bonding only

  • Tapered tip plastic syringe (Nichiryo 6mL syringe with tips)
  • Uncured PDMS (10:1 w/w elastomer base to curing agent)


  • Drill press
  • Plasma cleaner (Harrick Plasma, Basic plasma cleaner PDC-32G)
  • Hot plate
  • Oven (75°C)

What do I do?

Well-plate preparation (for both bonding methods)

  1. Prepare the well-plate by drilling a hole in the center of each well corresponding to an inlet or outlet on the PDMS replica (Figure 2).
  2. Using an X-Acto knife, clean the edges of the drilled holes such that the bottom surface of the well-plate is smooth and any lips that may have formed from drilling have been removed.

Figure 2 – The prepared PDMS device is shown in a. and the prepared well-plate is shown in b.

APTES bonding Procedure

Well-plate APTES modification

1.      Clean the bottom surface of the well-plate with IPA and expose to oxygen plasma on high setting for 2 minutes, with the bottom surface of the plate facing up (Figure 3a).

2.      In a fume hood, prepare a 100 mL aqueous solution of 1% v/v APTES and pour the solution into a shallow, resealable container.

3.      Place the plasma treated well-plate in the APTES container so that the bottom surface of the plate is completely submerged. Seal the container and let soak for 30 minutes (Figure 3b)

4.      Remove the plate from the APTES bath and rinse the top and bottom with water. Dry the well-plate using compressed air and place it on a 50°C hot plate to ensure thorough drying.

Figure 3 – The well-plate was exposed to air plasma and submerged in a water/APTES solution to modify the surface chemistry and enable bonding between PS and PDMS. A coverslip was then plasma bonded to the PDMS surface.


1.      Clean the top of the PDMS replica (opposite to the channels) using scotch tape and plasma clean on high for 1 minute.

2.      With the channeled side of the PDMS replica facing up, align the inlets/outlets of the replica with the holes of the APTES-modified well-plate and press the layers together. Roll a brayer over the surfaces to remove any bubbles and ensure an even, uniform bond. Bake at 75°C for 20 minutes (Figure 3c).

3.      Remove the well-plate with bonded device from the oven and use scotch tape to remove debris from the channel-exposed PDMS. Clean a glass coverslip with IPA and expose the coverslip and well-plate to oxygen plasma on high for 1 minute. Bond the coverslip to the PDMS replica, thus enclosing the channels and bake at 75°C for 20 minutes.

Uncured PDMS procedure

1.      Remove any dust from the bottom (channel-exposed) side of the PDMS replica using Scotch tape and clean a glass coverslip with IPA. Expose both to oxygen plasma for 1 minute on high setting and bond them together, enclosing the channels. Bake at 75°C for 1 hour (Figure 4a).

2.      Clean the bottom surface of the prepared well-plate with IPA. Using the tapered tip syringe, place small droplets of uncured PDMS onto the bottom surface of the well-plate where the PDMS device will be bonded (Figure 4b).

3.      Using scotch tape, remove any dust from the top (opposite to the channels) of the coverslip-bonded PDMS replica. Align the inlets/outlets of the device with the holes of the well-plate and lightly press the device onto the well-plate (Figure 4c). Remove any uncured PDMS that may have leaked into the wells or inlets/outlets of the device. Bake at 75°C for 1 hour.

Figure 4 – The PDMS device was first bonded to a coverslip (a) and then bonded to a well-plate using uncured PDMS (b). c shows the completed device from the top and side view.


We present two methods for attaching PDMS microfluidic devices to polystyrene well-plates, providing the opportunity to utilize customized channels for well-plate microfluidics. Assays using these devices can be run in conjunction with well-plate microfluidic controllers or using simple pipetting methods by adding the desired reagent or media to the inlet wells (Figure 9). While the fabrication process is more involved than typical PDMS processing, well-plate microfluidics removes the need for complicated tubing connections by working with a single manifold controller, or hydrostatic flow using the well height to produce pressure.


1         M. Khine, C. Ionescu-Zanetti, A. Blatz, L. P. Wang and L. P. Lee, Lab Chip, , DOI:10.1039/b614356c.

2         C. G. Conant, M. A. Schwartz, J. E. Beecher, R. C. Rudoff, C. Ionescu-Zanetti and J. T. Nevill, Biotechnol. Bioeng., , DOI:10.1002/bit.23243.

3         Fluxion, White Pap., 2008, 1–6.

4         2012, US00825796.

5         C. G. Conant, J. T. Nevill, M. Schwartz and C. Ionescu-Zanetti, J. Lab. Autom., 2010, 15, 52–57.

6         P. J. Lee, N. Ghorashian, T. A. Gaige and P. J. Hung, J. Lab. Autom., , DOI:10.1016/j.jala.2007.07.001.

7         V. Sunkara, D.-K. Park, H. Hwang, R. Chantiwas, S. a. Soper and Y.-K. Cho, Lab Chip, 2011, 11, 962–965.


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A second life for old electronic parts: a spin coater for microfluidic applications

Gabriele Pitingolo1, Valerie Taly1 and Claudio Nastruzzi2

1INSERM UMR-S1147, CNRS SNC5014; Paris Descartes University, Paris, France. Equipe labellisée Ligue Nationale contre le cancer.

2Dipartimento di Scienze della Vita e Biotecnologie, Università di Ferrara, Ferrara, Italia

*Email: gabriele.pitingolo@parisdescartes.fr, nas@unife.it

Why is this useful?

It is well known that the rapid proliferation of information and communications technologies (ICT) has resulted in a global mountain of high-tech trash (e-waste). The problem with e-waste is not only the accumulation of electronic products and therefore the high disposal costs, but rather the hazardous substances present in their various components. Therefore, the importance of recycling is evident in the area of resource and energy conservation, finding a new, second life for electronic components.

Spin coaters are widely used instruments useful to deposit uniform thin films to flat substrates [1]. In microfluidics, the spin coating is used to coat a photoresist layer (such as SU-8) or to bond separate substrates by using the adhesive properties of PDMS. The spin coating technology is also used to fabricate thin polymer membranes. PDMS membranes are, for example, employed for a wide range of applications due to their several advantages. For instance, being PDMS membrane permeable, they can be used to exchange gas (in cell culture application for example) or small molecule (in filtration application) [2]. In addition, as recently reported, spin coating is suitable to fabricate microchannels with a circular section [3].

Unfortunately, most commercial spin coaters are expensive (£2,000-6,000) and possess some unwanted or redundant specifications, not necessarily needed for the fabrication/modification of microfluidic devices.

In this respect, we present here a tip to develop portable spin coaters by recycling computer fans and mobile phone wall chargers. The most common fans in personal computers have a size of 80 mm, but the size can range from 40 to 230 mm. It’s also known that the fans of different size show also a different rotational speed. Typically, the 80 mm fans have a rotational speed of 2000 rpm (that represent a suitable speed for common thin layering in microfluidics).

What do I need?

Parts for the spin coater

  • Personal computer fan
  • Insulated male/female wire pin connectors
  • Tesa power strip
  • Wall chargers from (old) mobile phones

Parts and chemicals for the specific examples

  • Milled poly(methyl methacrylate) (PMMA) microchannel
  • Glass slide
  • Sylgard® 184 silicone elastomer kit
  • Clumps

What do I do?

Assembling of spin coater

1.Remove the fan from an old pc (or mac, if are particularly posh) (Fig.1).

2. Connect the wall charger and the fan wires with insulated female and male wire pins. Afterwards, to turn on the fan, connect the female and male pins.

3. Using the tesa power strips, secure the substrate (i.e. glass slide or PMMA microchannel) to the central part of the fan (left picture). For devices larger than the fan, use an adeguate plastic stopper to elevate the device (right picture).

4. Drip, by a (micro)pipette, the liquid containing the coating material on top of the substrate.

5. Turn on the fan and spin coat the substrate for about 30 seconds (time can vary depending on the substrate viscosity and coating thickness required).

6. Verify the coating by peeling off the PDMS membrane from the glass slide by tweezer (left picture) or analyze the microchannel profile by microscopy (right panels).

What else should I know?

In this tip a portable spin coater for microfluidic applications was developed using old electronic parts. A single fan can be re-used many times (up to hundreds in our experience). The amount of PDMS (in form of droplets) falling on the fan is quite limited. If necessary the fan can be cleaned after any use by simply rubbing it with a wipe soaked with some petroleum ether (aka liquid paraffin or white petroleum). In the worst cases (very rarely occurring) the fan can be easily replaced, since they are available for free by any old unused PC.


This work was supported by the Ministère de l’Enseignement Supérieur et de la Recherche, the Université Paris-Descartes, the Centre National de la Recherche Scientifique (CNRS), the Institut National de la Santé et de la Recherche Médicale (INSERM). This work was founded by CAMPUS FRANCE (n° 39525QJ) and carried out with the support of the Pierre-Gilles de Gennes Institute equipment (“Investissements d’Avenir” program, reference: ANR 10-NANO 0207). Financial support from the Università italo-francese grant G18-208 is gratefully acknowledged.


[1]          D. B. Hall, P. Underhill, and J. M. Torkelson, “Spin coating of thin and ultrathin polymer films,” Polymer Engineering & Science, vol. 38, no. 12, pp. 2039-2045, 1998.

[2]          S. Halldorsson, E. Lucumi, R. Gómez-Sjöberg, and R. M. Fleming, “Advantages and challenges of microfluidic cell culture in polydimethylsiloxane devices,” Biosensors and Bioelectronics, vol. 63, pp. 218-231, 2015.

[3]          R. Vecchione, G. Pitingolo, D. Guarnieri, A. P. Falanga, and P. A. Netti, “From square to circular polymeric microchannels by spin coating technology: a low cost platform for endothelial cell culture,” Biofabrication, vol. 8, no. 2, pp. 025005-025005, 2016 May 2016.

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Development of a cell culture microdevice with a detachable channel for clear observation

Eriko Kamata, Momoko Maeda, Kanako Yanagisawa, Kae Sato*

Department of Chemical and Biological Sciences, Faculty of Science, Japan Women’s University, Bunkyo, Tokyo 112-8681, Japan

*E-mail: satouk@fc.jwu.ac.jp

Why is this useful?

There have been many reports on microfluidic devices for cell culture having upper and lower microchannels separated by a thin PDMS membrane. In these devices, the lower channel often interferes with the microscopic observation of cells cultured in the upper channel. To avoid interference, a microdevice with a detachable lower channel was developed.

What do I need?


PDMS (SILPOT 184w/c, Dow Corning Toray)


PMMA sheet (56 × 76 × 2 mm)

Glass slides (52 × 76 mm and 26 × 76 mm)

Cover slip (24 × 60 mm)

1 kg weight

PTFE tubing (1 × 2 mm and 0.46 × 0.92 mm)

Tygon tubing (1.59 × 3.18 mm)

Biopsy Punches (1 mm and 2 mm, Kai corporation)


Vacuum desiccator

Oven (65 ˚C and 100˚C)

Spin coater

Plasma generator

Vacuum pump

What do I do?

  1. PDMS molding

Mix the elastomer and curing agent at a 10:1 mass ratio. De-gas the mixture under vacuum until no bubbles remain (20 min). Pour the degassed PDMS mixture onto a master, which has an upper channel (1 × 1 × 10 mm) or a lower channel (0.5 × 2 × 15 mm) structure, and then place it in an oven at 65˚C for 1 h. Peel off the PDMS replica from the master and adhere it to a glass slide (26 × 76 mm). Place it in an oven at 100˚C for 1 h (Fig.1).

  1. Preparation of a thin PDMS membrane

Spin coat 600 µL of the PDMS prepolymer (at a 10:1 mass ratio) on a PMMA sheet at 500 rpm for 20 s followed by 2,400 rpm for 600 s. Bake it at 65˚C for 1.5 h.

Fig.1 PDMS sheet with the upper channel, that with the lower channel, PDMS membrane, and tubing.

  1. Permanent bonding of the PDMS membrane and the sheet with the upper channel

Punch the inlet and outlet holes at both the ends of the upper channel with a 2-mm biopsy punch. Expose both the bonding surfaces of the PDMS membrane and the PDMS sheet with the upper channel (upper sheet) to plasma at 100 W, 35 s (Fig.2a and 2b). Laminate and bake them at 65˚C for 1 h (Fig.2c). Remove the PMMA sheet, and punch a hole to connect the sheet with the lower channel by using a 1-mm biopsy punch from the membrane side.


Fig.2 (a) Schematic diagram of bonding the PDMS membrane and the upper sheet. (b) Plasma treatment. (c) Bonded PDMS membrane and sheet.

  1. Detachable bonding of the PDMS membrane and the PDMS sheet with the lower channel

The PDMS sheet with the lower channel (lower sheet) is bonded to the PDMS membrane by using a PDMS prepolymer diluted with hexane (a dilution ratio of 1:3) as a glue1 (Fig.3a). Spin coat the diluted PDMS prepolymer (600 µL) at 2,000 rpm for 30 s on the surface of a glass slide (52 × 76 mm) to cover the slide with a thin layer of the glue and incubate it for 10 min to dry the solvent (Fig.3b). Place the lower sheet on the coated glass slide (Fig.3c). Apply glue at the four corners of the PDMS membrane bonded with the upper sheet (Fig.3d). Peel off the lower sheet from the glass slide and place the glue-coated surface of the sheet on the PDMS membrane (Fig.3e). After 30 min of incubation, bond the lower sheet to a cover slip by plasma bonding. Place a 1-kg weight and a glass slide on the device and bake it at 100˚C for 1 h (Fig.3f).

Fig.3 (a) Schematic diagram of bonding of the PDMS membrane and lower sheet. (b) Spin coat the diluted PDMS prepolymer (glue). (c) Lower sheet is placed on the thin film of the diluted PDMS prepolymer. (d) The diluted PDMS prepolymer is applied on the four corners of the PDMS membrane. (e)The glue-coated surface of the lower sheet is placed on the PDMS membrane. (f) A weight and a glass slide are placed on the device to bake it at 100˚C.

  1. Tubing

Connect polytetrafluoroethylene (PTFE) tubes (1 × 2 × 100 mm) with Tygon tubes (1.59 × 3.18 × 10 mm) to the holes present at both ends of the upper microchannel. Connect a PTFE tube (46 × 0.92 × 150 mm) to the hole of the lower channel. Apply PDMS prepolymer at the root of the tubes, and then bake it at 100˚C for 1 h for firm connection (Fig.4a and b).

  1. Cell culture

Introduce a cell suspension into the upper microchannel, which is manually precoated with 0.1 mg/mL of fibronectin. Incubate the device at 37˚C with 5% CO2 for 16 h to allow cells to adhere to the bottom of the upper channel (surface of the PDMS membrane).

  1. Detachment of the lower sheet for cell observation

Remove the lower sheet from the device carefully (Fig.4c and d). Place the rest of the device on a cover slip for observation with an inverted microscope (Fig.4 f and h).

Fig.4 (a) The complete microdevice. (b) Side view of the microdevice. The cell culture channel (upper) is filled with water containing a red food color, while the lower channel is filled with water containing a blue food color. (c) and (d) The lower sheet is peeled off from the microdevice carefully. Phase contrast images of cells (e) before and (f) after detachment of the lower sheet. Fluorescent images of cells stained with CellTracker Red CMTPX (g) before and (h) after detachment.


We developed a microfluidic device with a detachable lower microchannel. It is important that different bonding techniques be used for each side of the PDMS membrane. If the lower channel is filled with air and the device is incubated in a CO2 incubator, dew condensation is often observed in the lower channel when the device is taken out from the incubator. The condensation in the lower channel makes observation difficult (Fig.4e and g). This problem was solved with the detachable device.


This work was supported in part by the Japan Society for the Promotion of Science (JSPS) KAKENHI Grant Number JP16H04170.


  1. Chueh, B.H., Huh, D., Kyrtsos, C.R., Houssin, T., Futai, N., Takayama, S. (2007). Leakage-free bonding of porous membranes into layered microfluidic array systems. Analytical Chemistry 79 (9), 3504–3508.
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