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Juyue Chen^a, Rui Zhang^b and Wei Wang^*a
a National Key Laboratory of Science and Technology on Micro/Nano Fabrication, Institute of Microelectronics, Peking University, Beijing 10871, P. R. China
^b School of Pharmaceutical Sciences, Peking University, Beijing 10871, P. R. China

Why is this useful?

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Microporous PDMS has been proposed as a functional PDMS material for cell culture related microfluidics applications where high gas perfusion is required to improve cell survival and functions. The phase separation micro-molding (PSμM) technique, which is widely used in microporous polymer preparation¹, creates difficulties in fabricating microporous PDMS as there are many restrictions on the solvent – including high boiling point, low volatility, stability and appropriate compatibility with nonsolvent. Yuen reported that microporous PDMS can be simply prepared by curing PDMS pre-polymer with a porogen, such as salt or sugar particles, and then dissolving and washing away the porogen.² However, it is difficult to obtain a microporous PDMS with pore size of micrometer scale by using this solid porogen. The small porogen makes the soaking and washing time cumbersome for most applications. As mentioned by Yuen, the porogen should be dissolved and washed away by soaking the PDMS and washing it in ethanol solution in an ultrasonic cleaner for at least 3 hours (longer may be required for smaller particles of porogen).

Here, we propose a simple way to fabricate microporous PDMS using an emulsion of PDMS and water, as illustrated in  Figure 1. After manually blending the PDMS pre-polymer and water (1% SDS inside), water droplets were dispersed inside the PDMS pre-polymer. When the mixture is heated at a relatively low temperature (80^oC) and in a relatively highly humid environment, the pre-polymer partially cures with the water droplets, keeping their original state. Through further curing at a higher temperature (120^oC), the water trapped in the PDMS evaporates and leaves pores inside the PDMS matrix. Pore density is determined by the ratio of the water volume to the PDMS pre-polymer volume.

What do I need?

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• PDMS (Sylgard 184, Dow Corning Co.)
• SDS (dodecyl sulfate sodium salt)
• Deionised water
• Petri dish
• High temperature durable container
• Oven

What do I do?

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1. Mix the PDMS according to the manufacturer’s instructions, with a mass ratio of base to curing agent of 10:1.
2. Make the SDS solution, with a mass ratio of SDS to DI water of 1:100.
3. Pour the PDMS pre-polymer and water (1% SDS inside) into a Petri dish with a given volume ratio, and manually blend them until a uniform emulsion (milky and opaque) is achieved. The water can be added step by step to facilitate the blending. Pore density is determined by the ratio of water volume to PDMS pre-polymer volume, as shown in Figure 2.
   

   Figure 2. SEM photos of the prepared microporous PDMS. R = Vwater:VPDMS. (a) R = 0.01; (b) R = 0.05; (c) R = 0.3; (d) R = 0.7. All the scale bars represent 20μm

4. Add some DI water in a high temperature durable container, and put the Petri dish with the water-in-PDMS emulsion inside on the water, then cover with the container lid. Put the container into the oven for about 2 hours at 80^oC.
5. Once the PDMS has been partially cured, remove the Petri dish from the container and finish curing it at a relatively high temperature in the oven for about 1 hour. After all the trapped water droplets have evaporated, the finished result is microporous PDMS.

References

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[1] L. Vogelaar, R. G. H. Lammertink, J. N. Barsema, W. Nijdam, L. A. M. Bolhuis-Versteeg, C. J. M. van Rijn and M. Wessling, Phase separation micromolding: a new generic approach for microstructuring various materials, Small, 2005, 1, 645–655.
[2] P. K. Yuen, H. Su, V. N. Goral and K. A. Fink, Three-dimensional interconnected microporous poly(dimethylsiloxane) microfluidic devices, Lab Chip, 2011, 11, 1541-1544.

E. M. Dunfield, Y. Y. Wu, T. P. Remcho, M. T. Koesdjojo and V. T. Remcho
Department of Chemistry, Oregon State University, Corvallis, OR 97331, USA

Why is this useful?

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Paper-based microfluidics offers several advantages over conventional microfluidics, and has great potential to generate inexpensive, easy-to-use, rapid and disposable diagnostic devices. Unlike traditional microfluidics, which often requires pumps to move fluid through the microfluidic channels, paper microfluidics can be performed without such instrumentation due to the flow of fluid being driven by capillary action through the paper. Hence, paper-microfluidics is well-suited for use in point-of-care diagnostics and in developing countries where expensive instrumentation is not available. There have been several advancements in the fabrication of paper microfluidic chips. Fabrication of paper-based chips can be done by photolithography [1], wax printing [2, 3], plasma etching [4], inkjet etching [5], and use of a cutting plotter [6]. However, such techniques may require the use of organic solvents during the fabrication process, they can be labor-intensive and expensive, and may impose limits on the types of materials that may be used in the chip. A novel technique is presented that utilizes a hydrophobic film material, Parafilm® “M”, which is heated to above its melting temperature of 60^οC, and pressed into a piece of paper. The channel mask is cut from polycarbonate (PC) film, and sandwiched between the paper and the Parafilm®. The PC film mask prevents the melted Parafilm® from penetrating into the paper in the channel region, and therefore defines the hydrophobic boundaries of the paper channel. This technique has been tested on a wide variety of paper materials including Whatman Grade 1 filter paper, VWR light-duty tissue wipes and Kimtec Kimwipes, among other papers. While in some cases the paper can be cut directly into the desired channels, thin paper can easily tear and complex patterns proved to be challenging when cut by a cutting plotter. In contrast, the more rigid PC film can easily be cut into simple or complex patterns with a cutting plotter. Furthermore, the process is inexpensive, rapid, and does not require the use of organic solvents during fabrication.

What do I need?

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• Paper, such as Whatman filter paper, Kimtech Kimwipe or VWR light-duty tissue paper
• Polycarbonate film, thickness approximately 100 µm
• Parafilm® “M”
• Aluminium foil
• Scissors or x-y cutting plotter (for more complex channel patterns)
• Hot press

What do I do?

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1. Cut out the desired channel patterns in polycarbonate film. For better cutting precision and accuracy, an x-y cutting plotter can be used.
2. Create an assembly consisting of paper, polycarbonate (PC) cutout, and Parafilm® “M” as shown in figure 1. The paper can be Whatman filter paper, Kimtech Kimwipes, VWR light-duty tissue wipes or other paper of desired properties.
   

   Figure 1. Paper, polycarbonate and Parafilm® stack set-up before heating and pressing.

3. Cover both sides of the paper, PC, Parafilm® stack with aluminium foil to prevent sticking of the Parafilm® to the hot press plates, and place the whole assembly into the hot press.
4. Heat the hot press to above 60^οC, and apply ~200 psi of pressure for 1 minute. Note: 200 psi is necessary when using Whatman Grade 1 filter paper. Applied pressure varies depending on the thickness and porosity of the paper used.
5. Allow the aluminium packet to cool, and then remove the foil from the paper microfluidic chip.
   

   Figure 2. Completed paper microfluidic chips made using Parafilm®. (a.) Spiral design has a channel width of 1 mm. (b.) Design has circles of 4mm diameter and 8 straight channels of 2mm width and 10mm length. (c.) Blue dye added to the paper microfluidics chip shown in b.

What else should I know?

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For heavier weight paper such as Whatman Grade 1 filter paper, it is necessary to apply higher pressure (~200 psi) such as in a hot press to produce the microchips. However, for lighter weight paper such as Kimtech Kimwipes and VWR light-duty tissue wipes, microchips can simply be made by heating the plates of a heating element such as a hair straightener, and then applying gentle pressure to the paper, PC, Parafilm® stack to produce the paper-based chips.

References

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[1]  A. W. Martinez, S. T. Phillips, B. J. Wiley, M. Gupta, and G. M. Whitesides, Lab Chip, 2008, 8, 2146-2150.
[2] Y. Lu, W. Shi, L. Jiang, J. Qin, B. Lin, Electrophoresis, 2009, 30, 1497-1500.
[3] E. Carrilho, A. W. Martinez, G. M. Whitesides, Anal. Chem., 2009, 81, 7091-7095.
[4] X. Li, J. Tian, T. Nguyen, W. Shen, Anal. Chem., 2008, 80, 9131-9134.
[5] K. Abe, K. Suzuki, D. Citterio, Anal. Chem., 2008, 80, 6928-6934.
[6] E. M. Fenton, M. R. Mascareñas, G. P. Lopez, S. S. Sibbett, ACS Appl. Mater. Interfaces, 2009, 1, 124-129.

Gurucharan V. Karnad^*a,b, R. N. Ninad^*b and V. Venkataraman ^a
a Dept. of Physics, Indian Institute  of Science, Bangalore, India
^b Dept. of Electronics and Communication Engineering, Amrita School of Engineering, Bangalore, India
* Corresponding authors: Gurucharan V. Karnad, R. N. Ninad
Email: gvkarnad[at]physics.iisc.ernet.in, rnninad[at]gmail.com

Why is this useful?

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Spin coating is one  of  the  coating  techniques  used  to  apply  thin uniform  films  to  flat  substrates. A Spin Coater is a machine used for spin coating, and is one of the most ubiquitous instruments in any laboratory dealing with microfluidic devices.

Most commercial spin coaters are extremely expensive (≈ > US $1,995 [1]) and come with features and specifications not necessarily needed for fabricating and experimenting with polymer based microfluidic devices.

The cost of this instrument should not act as a deterrent for groups intending to venture into fabricating and experimenting with simple devices, hence the effort in this direction.

We have designed an inexpensive spin coater (with a touch-screen interface) costing less than US $350. The user input is through the touch-screen interface, where parameters such as spin duration and speed can be entered. Real time speed is also displayed alongside. A microcontroller forms the intelligence of the system and manages the inputs, display, and speed and duration control. The real time speed is sensed by the microcontroller using an optical encoder, and a control loop keeps it within acceptable error limits.

The substrate is mounted either using a double-side tape or a set of clamps. Improvements with regard to vacuum chuck, computer interfacing etc., can be done as per necessity.

What do I need?

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• PIC Microcontroller Programmer (with support for 18F4550)
• PIC 18F4550 (≈ US $04.47 [2])
• Graphic LCD 128×64( JHD 12864E or equivalent) (≈ US $16.80 [3])
• TouchPanel (≈ US $07.00 [4])
• TouchPanel Connector Board (≈ US $04.00 [5])
• Infrared Distance Sensor (equivalent – Cytron IR01A Medium Range Infrared Sensor) – Optical Encoder-Sensor (≈ US $08.10 [6])
• Crystal oscillator circuit (11 Mhz Crystal Oscillator, 22pf Capactitor (2) – connection as given in PIC 18F4550 Datasheet) (≈ US $01.00 [7])
• IR 2110 (≈ US $07.25 [8])
• IRFP 150N (≈ US $02.77 [9])
• 1N 4744A (≈ US $00.21 [10])
• BYQ 28E200 (≈ US $00.98 [11])
• Brushed DC Motor (Como Drills 719RE850 or equivalent) (≈ UK £21.69 [12] ≈ US $35)
• Any PC CPU Case (for Power Supply-SMPS and as instrument control box) (≈ US $50.00 [13])
• PC (to program the microcontroller initially)
• Software – MPLAB IDE (Free), HI TECH C Compiler for PIC 18 MCU (Free Version)
• Machining, raw material, workshop access ( for chuck, optical encoder mount and motor cabinet) (≈ US $200.00)

What do I do?

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1. Interface and connect the PIC Microcontroller Programmer to the PC as per the instructions given in its manual
2. Burn the Program01.hex file into the microcontroller with the help of the programmer
3. Connect the microcontroller and other components as given in the circuit diagram (Figure 1). The circuit and display can be appropriately mounted in a PC CPU case
4. Power up the circuit
5. The Graphic LCD should display data similar to Figure 2. Figure 3 and Figure 4 will appear on the display if appropriate icons on the Menu “Selected Values” screen are touched
6. There can be dissimilarities with each touch panel and hence it may not respond to input due to change in touch co-ordinates. Hence, there would be a need to modify the Program01.c. If there are no problems, jump to instruction 11
7. Install the software mentioned above
8. Modify the program, by finding out the new co-ordinates of the icons by following the instructions given in readme.txt
9. Compile the modified Program01.c and program the microcontroller as per the instructions given in the programmer manual
10. Repeat instruction 2 to 5
11. Make an appropriate box to mount the motor
12. An aluminium ( light and easy to clean) chuck (Figure 6, Figure 7) to fit the motor shaft  has to be machined
13. A simple optical encoder set up  which includes mounting of a slim white acrylic piece on the bottom of the motor shaft and the IR optical sensor has to be made ( Figure  5)
14. Put together all the components appropriately. The spin coater (Figure 8 ) should now be ready ( appropriate modification of  the chuck may result in a centrifuge too)
15. Mount the wafer samples for spin coating either with a double-sided tape or a set of adjustable clamps

What else should I know?

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• Basic knowledge of C is essential to modify the code to suit individual requirements or specification
• The spin coater has been set to receive input of speeds from 1000-7000 RPM ( limited only due the motor used, can be modified easily)
• The maximum spin time is 999 seconds (increments of 1)
• Due to nonlinear response of the brushed DC motors to voltage ( and hence varying PWM values),  the speed response of the motor to variation in PWM values has to be plotted and an appropriate equation has to be estimated. The equation in Program01.C has to be modified (see the readme.txt).  Note: This needs to be done only if a Brushed DC motor other than Como Drills 719RE850 is used, or if further fine tuning of response is required.

References

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[1] http://www.mtixtl.com/DeskTopSpinCoater0-5100rpmwithCompleteAccessoriesVTC-100P.aspx

[2] http://us.element14.com/microchip/pic18f4550-i-p/ic-8bit-mcu-pic18f-48mhz-dip-40/dp/74K8623?in_merch=Popular%20Products

[3] http://www.ebay.com/itm/Graphic-Matrix-LCD-Module-LCM-JHD-12864-E-128X64-    /200392901987?pt=LH_DefaultDomain_0&hash=item2ea8590563#ht_4043wt_906

[4] http://www.mikroe.com/eng/products/view/277/various-components/

[5] http://www.mikroe.com/eng/products/view/207/touchpanel-connector-board/

[6] http://www.robotshop.com/2cm-10cm-infrared-distance-sensor-2.html

[7] http://us.element14.com/

[8] http://us.element14.com/international-rectifier/ir2110-1pbf/ic-mosfet-driver-high-low-side/dp/19K8414?in_merch=Popular%20Products

[9] http://us.element14.com/international-rectifier/irfp150npbf/n-ch-mosfet-100v-42a-to-247ac/dp/63J6848?Ntt=irfp+150n

[10] http://us.element14.com/multicomp/1n4744a/zener-diode-1w-15v-do-41/dp/90R9431?in_merch=Popular%20Products

[11] http://in.mouser.com/Search/ProductDetail.aspx?qs=kop3EGWU%252bIt67Q6mQhs7oA%3D%3D

[12] http://uk.rs-online.com/web/p/dc-motors/3213192/

[13] http://www.amazon.com/Computer-Cases-Add-Ons-Computers/b?ie=UTF8&node=572238#/ref=sr_nr_p_n_feature_keywords_0?rh=n%3A172282%2Cn%3A%21493964%2Cn%3A541966%2Cn%3A193870011%2Cn%3A572238%2Cp_n_feature_keywords_browse-bin%3A3606145011&bbn=572238&sort=price&ie=

R.J. Shilton, L. Y. Yeo, and J. R. Friend
MicroNanophysics Research Laboratory, Department of Mechanical and Aerospace Engineering, Monash University, Clayton, Victoria, 3800, Australia

Purpose

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It is often necessary to form millimeter order holes in glass (and similar) substrates to form inlet or outlet ports in microfluidic devices. The easiest way to do this is to simply drill a hole in the required position, however owing to the brittle nature of most materials used in these devices, this can often lead to a high failure rate where the devices crack during the drilling process. Outlined here is a simple procedure for drilling ports in microfluidic devices, which has been tested with glass, silicon, and lithium niobate, with a very high success rate.

Materials

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• ~ 1mm diamond drill bit (UKAM Industrial Superhard Tools, Valencia, CA)
• Drill press
• Double sided tape
• Disposable plastic petri dish
• Small piece of alumina (or other hard flat material)
• Substrate to drill ports into

Procedure

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1. Attach microfluidic device to alumina with double sided tape. We have used this method reliably to drill ports in glass, silicon, and lithium niobate, however in principle it should work on a range of similar substrates. Silicon is shown in all images.

2.  Stick alumina to bottom of petri dish with a small amount of double sided tape.

3. Fill petri dish with a generous amount of water to cool down the drill site, and to remove particles into the fluid while drilling the hole.

4. Attach diamond drill bit to drill press. We successfully used drill bits of diameter 0.75 and 1 mm, however others should work equally as well.

5. Drill at a high speed (~10,000 RPM). Quite a bit of force can be applied without cracking the substrate, as it is stuck to a rigid backing. Drilling through a 0.5 mm thick substrate should take about ten to fifteen seconds, if it is attached firmly.

What else should I know?

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• Release the downward force a little near the hole exit, to avoid a rough hole on the other side.
• Keep device immersed in water after drilling until it is ready to be cleaned to avoid particles becoming stuck in device channels etc…
• Change water regularly to remove particle build up

Tiama Hamkins-Indik, Sandra Lam, Megan E. Dueck and Luke P. Lee
Department of Bioengineering, Berkeley Sensor and Actuator Center, Biomolecular Nanotechnology Center, University of California, Berkeley, CA 94720, US

Why is this useful?

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Currently, there is no simple method for aligning multiple layers of PDMS microfluidic devices onto a glass slide.  We report a method for alignment that is easy, inexpensive, and has many relevant applications including printing proteins adjacently , flowing cells over previously printed proteins, and aligning two PDMS devices on top of each other for complex 3D geometries .  Currently, glass etching can be used to permanently mark glass, but this process is labor intensive and costly .  As a proof-of-concept, a 3-layered, 3 mm wide replica of Van Gogh’s Starry Night was created (Figure 5).

What do I need?

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1.    Permanent marker (e.g. Sharpie™ marker)
2.    Syringe (1mL – 10mL) with needle tip (any gauge)
3.    Vacuum chamber
4.    Light microscope with 4X and 10X objectives
5.    Scotch™ Tape
6.    Alignment marker

The Alignment Marker:
While any alignment marker can be used between layers, we suggest using the one shown in Figure 1.  The teeth act as Vernier scales in the x and y directions, thus, the degree of misalignment can be measured.  This alignment marker was designed with 10 µm wide teeth.  On the first layer (Figure 1, left), there are 22 posts which are 10 µm apart.  On the second layer, there are 20 posts each 11µm apart (Figure 1, right), making both markers 430µm wide.  The interlocking geometry of the markers is shown in Figure 2.  An inlet must be placed on the first layer’s alignment marker so that ink may flow through and stain the glass.

How do I do it?

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1. Incorporate alignment marker

• Add the alignment markers into the device design. On the silicon master that will be used to cast PDMS molds, The the channel height of the alignment marker on the silicon wafer should be in the 5 – 200 µm range.  The PDMS channel height only needs to be tall enough for Sharpie™ ink to flow through.

2. Sharpie™ Ink Extraction

• Put a needle onto a 1 mL – 10 mL syringe.
• Insert the needle into the felt tip of a Sharpie™ marker, and slowly pull the plunger.  Slow extraction is necessary to allow air to diffuse into the marker as the ink escapes into the syringe.  Repeat as necessary.  (Figure 3)
• Dispense the Sharpie™ ink into a microcentrifuge tube.  Sharpie™ extraction should result in 0.5 – 1 mL of Sharpie™ ink.
• Dilute Sharpie™ ink 3X in 100% ethanol.

3. First Layer

• Cut and punch desired PDMS device.
• Clean a glass slide and the device with Scotch™ tape.
• Reversibly bond the PDMS device onto a glass slide, by simply placing the cleaned PDMS onto the glass slide. (Figure 4a)
• Load the Sharpie™ ink into the alignment marker channel.  This can be done by placing the device into a vacuum chamber for 5-10 minutes, removing the device from the vacuum, and placing a ~5 µL drop of Sharpie™ ink over the punch hole.  Only punch one entry hole for this method. (Figure 4b)
• Allow the Sharpie™ ink to dry for 2 hours. (Figure 4c)
• Remove PDMS. (Figure 4d)

4. Second Layer

• Cut and punch desired PDMS device.
• Clean device with Scotch™ tape.
• Place Scotch™ tape onto glass slide a few millimeters away from previous design.  (Figure 4e)
• Under 4X or 10X magnification, bring the Sharpie™ alignment marker into the center of view and focus slightly above it.
• Carefully place the cleaned second layer onto the scotch tape, but do not press down on the device so that the PDMS device is not in contact with the glass slide.
• Still under the microscope, align the second layer with the Sharpie™ alignment marker by gently pushing the device along the Scotch™ tape.  The Scotch™ tape prevents the device from bonding with the glass slide.  (Figure 4f)
• Attach the second layer by reversibly binding it to the glass slide by pressing down on the device.
• Remove the scotch tape by holding the middle of the device down and pulling the scotch tape out from the edges of the PDMS device. (Figure 4g)
• If additional layers are necessary, repeat second layer procedure.

What else should I know?

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When using this technique by hand, the accuracy of the alignment between two layers can be down to 5 µm.  If a more precise alignment is necessary, a six axis alignment machine can be used.  As a proof-of-concept, we have reproduced Van Gogh’s Starry Night (Figure 5).  This design has two layers, and each layer was filled with Sharpie™ ink using vacuum loading.

References

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1. Kane et al., Patterning proteins and cells using soft lithogrpahy, Biomaterials, 1999, 20, 2363-2376.
2. Natarajan et al., Continuous-flow microfluidic printing of proteins for array-based applications including surface plasmon resonance imaging, Anal. Biochem., 2008, 373 (1), 141-146.
3. Chiu et al., Patterned deposition of cells and proteins onto surfaces by using three-dimensional microfluidic systems, Proc. Natl. Acad. Sci. U. S. A., 2000, 97 (6), 2408 -2413.
4. 3-Dimensional Molding for Making Microfluidic Devices, MicroDysis – Instrumentation Company with Micro- and Nano-fabrication, and Lab Automation, http://www.microdysis.com/TechMicrofab.aspx, 2010, accessed 16 April 2011.