A simple, effective and cheap means to reduce bubbles in microchannels

Brian Miller*, Dr. Helen Bridle, Dr Stewart Smith
School of Engineering, The University of Edinburgh, G1 John Muir Building,, Kings Buildings ,Edinburgh, EH9 3JL
Email: B.miller[at]ed.ac.uk
Tel: +44 131 650 7860

Why is this useful?


A common issue that arises when attempting to fill microfluidic channels is that of air bubbles becoming trapped against surfaces such as walls and features within the channel. Many different techniques have been used to minimise/eliminate these including pre-filling with surfactant dosed water combined with soaking in an ultra-sonicated bath1, attempting to fill very shortly after O2 plasma treatment while surfaces are hydrophilic2 and bubble traps integrated into the design3-5.

Presented in this tip is a cheap post-manufacture solution for the reduction/elimination of bubbles when filling devices. This method takes advantage of the 10-fold increase in solubility of CO2 gas when compared to O2 and N2. By pre-filling your device with pure CO2 the trapped gas is dissolved away rapidly in comparison to air.

What do I need?


• CO2 “ Cornelius keg charger” and CO2 canister (Amazon B000NV9CE6)
• DI/RO water
• 0.01” ID, 1/16” OD PEEK tubing (Sigma-Aldritch Z226661)
• PEEK Tubing to Luer Lock Adaptor + Ferrule (LS-T116-100 and LS-T116-300 Mengel Engineering)
• AN-4 to 1/8” Female NPT adaptor (Aeroquip#023-FCM2721)
• Male Luer Lock x 1/8″ NPT male adaptor (Cole Parmer PN: OU-31507-84)

What do I do?


Assemble the metal adaptors onto the keg charger as illustrated in Fig.1. Cut a ~3ft to 3 ½ ft length of PEEK tubing and seal into the luer-lock adaptor. This acts as a flow resistor that will reduce the output pressure from ~60 bars to ~ 1-2 bars. Connect the PEEK adaptor to the Luer lock on the charger (Fig.2).

Place PEEK tubing into a device input (Fig.3). Ours mount directly into the device having used a 1.2mm hole punch to create input ports on the PDMS devices. Depending on your device ports you may need to develop a unique solution suitable for your preferred porting method.

Use your fingers to block off any other inputs and use the trigger of the keg charger to release CO2 into your device. Use your fingers to block off the output ports and depress keg charger trigger to fill the other input channels.

Immediately connect DI/RO water filled tubing to your device and begin filling. Remember to pre-fill the tubing with water so that air trapped in the tubing is eliminated.

 Please see the time-lapsed videos showing a bubble completely dissolved (SI.1) in ~10 mins (flow rate ~50ul/min in a 50um high device). Also note that for very large bubbles this technique slowly decreases in efficacy until it appears a saturation point is reached (SI.2) after roughly 30 mins. It is recommended to “massage” any very large bubbles towards the outputs and ideally break them up into smaller pockets of trapped gas. Compare these to a video of a medium sized (1mm radius) bubble of normal air to see little to no reduction at a similar flow rate over 10 mins SI.3)

Figure 1

Fig.1: Charger Adaptor Assembly

Figure 2

Fig.2: Assembled CO2 filling instrument with PEEK tubing flow resistor

Figure 3

Fig.3: Filling a device with pure CO2

References


[1] D. W. Inglis, N. Herman and G. Vesey, Biomicrofluidics, 2010, 4.
[2] I. Wong and C.-M. Ho, Microfluidics and Nanofluidics, 2009, 7, 291-306.
[3] A. M. Skelley and J. Voldman, Lab on a Chip, 2008, 8, 1733-1737.
[4] W. Zheng, Z. Wang, W. Zhang and X. Jiang, Lab on a Chip, 10, 2906-2910.
[5] C. Lochovsky, S. Yasotharan and A. Gunther, Lab on a Chip, 12, 595-601.

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Flexible and customized male Luer adapters with low dead-volume requirements

Bretton Fletcher*, Miguel Z. Rosales and Bruno F.B. Silva
Department of Physics, Department of Materials, and Molecular, Cellular & Developmental Biology Department, University of California, Santa Barbara, California 93106, United States
*email: brettonfletcher[at]gmail.com  

Why is this useful? 


Microfluidic chips are used for several different purposes in the lab. Experiments so diverse as nanoparticle synthesis; cell sorting; analytical determination of chemical and/or biological species; are all widely performed nowadays in many laboratories. Due to these so diverse applications, it is often hard to find parts (e.g. adaptors, tubing, etc) that fit the researcher’s needs for a particular project. Among these is the difficulty in finding suitable adapters to connect the microfluidic chip to tubing and syringes. A common difficulty is to connect small diameter tubing (inner diameter of 0.01 or 0.02”) to Luer adapters, which are normally designed to fit larger ID tubing sizes. A second common difficulty is the large dead volume normally associated with common Luer connectors. This can be a major setback when working with expensive and rare materials. 

Nanoports from Upchurch® are a valid solution for these two problems, but they are expensive, and somewhat long and bulky, which may restrict some experiments. For instance, if the connector is long, it may restrict access of a microscope lens to the area around it.
Here we show a suitable way of preparing customized Luer adapters that (i) fit a wide range of microfluidic tubing; (ii) have very low dead volume requirements, and (iii) are smaller than most of the market alternatives. 
  

What do I need? 


  • Natural Polypropylene Male Luer Coupler (Value Plastics, MLRLC-6). This design has the advantage of providing two custom connecters per original connecter (Figure 1). Other designs may be used, as long as one end has a Male Luer fitting. Other materials may also be used, provided that they bond with PDMS.
  • Tygon Microbore Tubing, S-54-HL ID: .02in and ID: .04. Other sizes can be used.
  • Polydimethylsiloxane (PDMS). We mix a 10 to 1 ration of silicone elastomer base and silicone elastomer curing agent.            
  • Devcon 5 Minute Epoxy
  • Petri Dish, Tape, & Scissors

  

What do I do? 


  

  1. Cut the cylindrical ends off either side of a Luer coupler to get two connector pieces (figure 1).
  2. Push a few centimeters of tubing into the rough (cut) side of one connector and out of the clean side (figure 2).
  3. Fix the connector and tubing vertically by attaching to a petri dish with tape so that the rough end faces up (figure 3).
  4. Apply epoxy to the top of the connector so that it completely plugs the opening and fixes the tubing in place. Wait for it to dry (figure 4). One of the main purposes of the epoxy is to fix the tubing to the connector. Since it is very viscous and dries quickly, this is very suitable.
  5. Once the epoxy dries, reattach the connector to the petri dish with the opposite end facing up, and fill the rest of the connector with PDMS. Be sure not to leave any air bubbles in the connector (figure 5).
  6. Place connector and tubing on aluminum foil in an oven for two hours at 60°C. Be sure not to let the tubing touch hot surfaces in the oven. At this temperature the PDMS cures adequately and the Tygon tubing does not degrade.
  7. Once the PDMS is completely hardened, cut off the excess tubing from the clean end of the connector for a smooth face (figure 6).

  

Figures 1-4

1) Male Luer coupler after cutting. Both cylinders can be used to make connectors. 2) Small length of tubing pushed through connector. 3) Vertical attachment to petri dish. This makes it convenient to make many at one time. 4) Epoxy covering top of connector.

Figures 5-7

5) Epoxy on bottom, PDMS filling rest of connector. 6) Finished connector. 7) Connector attached to device.

  

 What else should I know? 


This concept is very simple and can be extended to other types of tubing (material, diameter, etc) and connectors. By fixing one tube to one connector, we avoid all compatibility problems, and minimize possible leakage problems. The tubing and Luer connector become one single unit. With this comes the slight disadvantage that each connector will be associated permanently with one tube. We don’t find this discouraging. Since both tubing and connectors are relatively cheap, we prepare a whole range of connectors at a time that fit our needs. The total amount of work required is also not more than one hour, and does not require a high level of skill. 

One advantage of this method, compared to other nice solutions shown here in “Chips and Tips”, is the lack of needles in the connector, which decreases the chance of leakages and needle-related injuries. 

We have tested the connectors with water and protein solutions of intermediate viscosity at several flow rates and leaks from the Tygon tubing-connector junction were never detected. 

The main purpose of the epoxy resin is to fix the tubing to the connector. Since it is very viscous and dries quickly, this is very suitable. A second advantage is that indeed, epoxy creates a rigid and robust junction between the Tygon tubing and the polypropylene material, making it very robust. The PDMS is then poured on top of the epoxy. Since the epoxy is porous, it would not be suitable to have in contact with the liquids under study. By isolating it with PDMS we fix this problem.

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Parallel fabrication of an array of holes in PDMS

D. Qi, C. K. Chan, and A. C. Rowat*
Department of Integrative Biology and Physiology, University of California, Los Angeles, 610 Charles E. Young Drive East, Los Angeles, CA 90095, USA
* e-mail: rowat[at]ucla.edu

Why is this useful?


A major challenge in microfluidics is interfacing micron-scale flow channels with fluid samples and peripheral equipment of the macroscopic world, such as plate readers and liquid-handling robots. To exchange fluids in and out of the device, inlet and outlet holes are essential. In polydimethylsiloxane (PDMS) microfluidic devices, small <1 mm inlet- and outlet-holes are typically used to interface fluidic samples and micron-scale channels. These holes are typically fabricated by manually excising PDMS slabs to produce holes, for example using a needle or razor-sharp biopsy punch. However, to achieve scale-up of microfluidic devices can require ~10^2 inlets and outlets required for parallelization of multiple samples.

Manually punching holes is tedious due to the following factors: (1) it is often difficult to determine where to punch holes due to the micron-scale dimensions of the channels and transparency of PDMS; (2) the resulting holes do not always have smooth edges, and resultant chunks of PDMS can clog micron-scale channels; (3) biopsy punches used to excise holes become dull after multiple uses and thus frequently need to be replaced; (4) it is difficult to manually punch holes that are vertically straight and have reproducible spatial position; this makes it challenging to achieve the precision required for fabricating devices that robustly interface with existing high throughput equipment, such as liquid-handling robots. Here we present a method to easily and reproducibly fabricate smooth, vertically straight arrays of interfacing holes in PDMS using standard arrays of 96 or 384 pipette tips.

What do I need?


  • Common items to prepare PDMS devices: petri dish, master wafer or device mold, PDMS silicone elastomer (base and curing agent), vacuum desiccator, and oven.
  • Micropipette tip racks. Depending on microfluidic device designs, users can build their own tip racks; for example, we machined a 96-tip holding rack out of Perspex (polymethylmethacrylate).
  • Micropipette tips, e.g., 10-, 20- or 200- μl volume.
  • Four 3-inch steel bolts with nuts.

What do I do?


  1. Assemble tip racks using four steel bolts with nuts (see Figure 1.); the bottom rack is fixed and the top one is adjustable vertically along the bolts.
  2. Put the petri dish (with the master wafer inside) on the bottom rack.
  3. Position pipette tips in the top rack.
  4. Align tips to the inlet- and outlet-hole positions of the desired microfluidic channels; move the top rack down until the tips physically touch the master wafer.
  5. After the tip alignment, put a flat light weight object, e.g., a Perspex block ~155g, on top of the tips to hold them in position, while also making sure that tips and the master wafer are in physical contact.
  6. Prepare PDMS silicone elastomer (base-to-curing agent ratio-10:1) and mix well by vigorous stirring.
  7. Carefully pour the silicone elastomer mixture into the petri dish.
  8. Put the entire setup in the vacuum desiccator and degas for 20-30 minutes.
  9. Move the setup into an oven and thermally cure silicone elastomer at 65 degree Celsius for >1 hour.
  10. After curing, remove the setup from the oven, and let cool to room temperature.
  11. Using tweezers, gently pull out tips from the thermally cured PDMS.
  12. Using a razor, cut out the PDMS device with molded channels and holes from the petri dish.
  13. Bond PDMS to glass coverslips or other substrates, as required for your application.

Figure 1
Figure 1. Images of hole-molding setup with (a) standard pipette tip rack, and (b) custom fabricated Perspex holder. Images of the molded holes in PDMS are shown in (c) and (d); inset in (d) shows one micropipette tip with tiny PDMS plug at its tip.

Figure 2
Figure 2. Typical inlet/outlet holes in a PDMS slab (a) molded by our molding method using a 200 μl micropipette tip; and (b) punched by a 0.75mm Harris Uni-Core (TED PELLA) punch. The PMDS slab is patterned with 50 μm cylindered posts.

What else should I know?


  • The arrangement of tips can be tuned for specific applications by using a custom-fabricated pipette tip rack to accommodate unique inlet- and outlet-hole arrangements; such as the Perspex rack as shown in Figure 1. (b).
  • During molding, a small amount of silicone elastomer is taken up into the micropipette tip ends during degassing in the vacuum chamber (Fig. 1d, inset); this occurs due to the imperfect sealing between the micropipette tips and the master wafer. This tiny plug of PDMS is easily removed from the device together with the pipette tip, resulting in holes that are free of any obstructions.
  • When using tubing to interface microfluidic devices with macroscopic equipment, the outside diameter (OD) of the tubing should be larger than the diameter of such mold holes in PDMS to avoid leakage. We have successfully used both 20- and 200- μl micropipette tips to mold holes, which are ~0.7 mm in diameter at the end; 10 µL pipette tips have similar diameter. Both 1⁄32″ and 1⁄16″ OD tubing (VICI Valco Instruments) interfaces well with holes of these dimensions.
  • By our proposed molding method, the size of the holes closely follows the outer diameter of the pipette tips; the holes do not contract after the tips are removed since they are formed during PDMS thermal curing. As shown in inset of Figure 1. (d), the PDMS plug in the micropipette tip is tapered, which helps to produce clean holes when the micropipette tip is pulled out of PDMS.
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Facile and plasma-free bonding of PDMS

J. Waynelovicha, Da’Kandryia Petersb, P. Salamona,  A.M. Segallb, Nicholas Sam-Soonc*
aDepartment of Mathematics & Statistics, San Diego State University
San Diego, CA 92182
bDepartment of Biology, San Diego State University
San Diego, CA 92182
cDepartment of Engineering, San Diego State University
San Diego, CA 92182
Email: nsamsoon[at]gmail.com

Why is this useful?


Microfluidic-based approaches have become widely adopted for analysis of cells, molecules, reactions, and processes. While plasma bonding has become the de facto way of bonding PDMS to other substrates, this method is expensive and, does not in fact work (without further additional steps) on plastics [1]. Here we show a simple way of using cheap and readily available materials to irreversibly reseal leaking plastic connections and bond PDMS to itself and other plastics.

What do I need?


  • PDMS (Polydimethylsiloxane) arts / microdevices
  • Loctite Plastic Bonding System (two part cyanoacrylate mix)
  • Plastic / silicon rubber based tubing

What do I do?


The procedure can be broken down into two steps; the priming of the substrates, and the application of the glue.

1) Priming
The end of the tube which contacts the leaking part is taken out and roughened up with sand paper. The activator is then applied on the around the surface as well as on the top of the device and left to dry for 30 seconds. (Figures 2A & B).

2) Glue application
The tube is reinserted into the hole and glue is then quickly smeared around the tubing and hole. (Figure 3).  Excess glue can be removed with an exacto knife. Allow at least 10 minutes to dry.

Figure 1

Figure 1, A) Loctite Plastic Bonding System B) Droplet formation due to poor seal at tube.

Figure 2

Figure 2. A) Applying the activator to the end of the tubing and B) to the top of the device.

Figure 3

Figure 3. Application of glue and bonding of the tubing to the part.

Figure 4

Figure 4. Bonding PDMS to A) PDMS B) Glass

What else should I know?


Loctite Plastic Bonding System can also be used to bond PDMS to PDMS (Figure 4A). This allows the creation of thick mounting blocks on thin devices providing a more durable friction fit.
If the device is to be used in a biological capacity, adequate time should be allowed to ensure complete polymerization of the glue. This along with flushing the device with buffer prior to using should minimize the possibility of any toxicity from unpolymerized material.
This system can also be used to bond PDMS to glass, PET and possibly other plastics. (Figure 4B)

References


[1]   Lee, Kevin S, and Rajeev J Ram., Plastic-PDMS bonding for high pressure hydrolytically stable active microfluidics. Lab Chip, 2009, 9, 1618-1624.

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Quick fluid interface to rigid microfluidic chips with single-sided adhesive rubber ports

Stuart Williams presents an alternative interconnection to plasma bonded PDMS-based ports for microfluidic chips.

Stuart J. Williams
University of Louisville, Louisville, KY, USA

Why is this useful?


This work demonstrates the use of a commercially-available single-sided rubber adhesive sheet to interface a rigid microfluidic chip for fluid access. This interconnect is an alternative to PDMS-based ports, whose plasma bonding characteristics may not be applicable for all rigid materials. The adhesive component of the port is incorporated with the rubber sheet; hence no additional curing time is needed. This technique provides a quick and inexpensive method to interface rigid microfluidic chips.

What do I need?


  • FDA-compliant silicone rubber adhesive back, 1/16″ thick (#8991K523 through McMaster-Carr, ~US$20 for one square foot, or ~US$0.02 per square centimeter)
  • A rigid microfluidic chip, with access ports prepared or predrilled. For demonstration purposes, glass used and drilled with a 0.9mm diamond rounded tip bit (diamondburs.net #HP801-009)
  • A rubber punching tool. Here, a sharpened 20G needle was used (Small Parts NE-201PL-C). This is the same punching method we have used to create ports in PDMS devices, hence other similar methods may also work
  • Tubing to insert into the port. Here, Tygon tubing (Small Parts TGY-010-C) was used with 0.010″ ID and 0.030″ OD
  • A razor
  • Tweezers (optional)

What do I do?


  1. Prepare and clean the surface of the rigid chip where the rubber port is to be applied.
  2. Manually cut a small square piece of rubber from the larger sheet. This is the size of your port and should be larger than the hole on the chip. For example, a 4mm x 4mm piece was cut for a 0.9mm diameter drilled hole.
  3. Sharpen the 20G needle with a metal file or similar tool (Fig. 1). This provides easier punching of the sheet itself.
  4. Place the small rubber piece, adhesive side down, on a flat, flexible surface. We recommend using a larger excess rubber piece as a punching surface. Using the sharpened needle, manually punch through the small rubber piece. Be sure to punch through the piece completely, including the paper backing. If done successfully, the punched port will contain the bulk rubber, adhesive and backing (Fig. 2). There will not be any adhesive around the immediate vicinity of the punched hole (Fig. 3).
  5. Remove the paper backing on the port, exposing the adhesive.
  6. Align the punched hole with the microfluidic chip. Manually apply pressure to adhere the port against the chip. Note: Alignment through a transparent chip can be accomplished through visual observation through the back side of the chip. Alignment with opaque chips can be done by using a smaller gauge needle (e.g. 30G) as an alignment guide through the punched hole with the chip access hole.
  7. The port is ready to be used. Insert the tubing into the port with tweezers.
  8. A completed chip is shown in Fig. 4.
Original and sharpened 20G needle tips

Fig. 1: Magnification of an original and sharpened 20G needle.

Image of rubber plug, adhesive and backing

Fig. 2: Magnification of a plug after being punched from the rubber adhesive sheet. The punching process simultaneously removes the rubber bulk, adhesive, and backing.

Punched hole in rubber sheet

Fig. 3: Magnification of a punched hole through the rubber sheet (bottom view). There is an absence of adhesive around the punched hole, preventing potential clogging of the port.

Completed chip

Fig. 4: Image of a completed chip that has two rubber microfluidic ports.

What else should I know?


Pressure limitations using this technique have not been quantified. However, no leaks were observed using manual injection methods, even after multiple uses.

Alternatively, pipette tips, syringe needles, or other items can be inserted into the rubber port for direct fluid injection.

Other adhesive rubber pads and thicknesses may work. However, other single-sided adhesive rubber pads were tested (Buna-N Rubber #8635K262, Neoprene #8583K162, Vinyl Rubber #8513K32, EPDM Rubber #8610K91, and Butyl Rubber #8609K12) and FDA Silicone performed the best overall in terms of ease of punching, bonding strength, and absence of adhesive clogs.

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Identifying multi layers or direction of flow with colored dots using Elmer’s Glue (polyvinyl acetate) and food coloring

Penny Burke and Teresa Porri
Cornell Nanobiotechnology Center, Cornell University, Ithaca, NY, USA

Why is this useful?


When making devices that are direction-specific and very small, you need to check them in the microscope each time to see which side to start your flow.  With this technique you can mark the PDMS with a color marker that does not interfere with the device.  When working with a multilayer device that has multiple valves and channels it is convenient to have identification markers.

What do I need?


  • PDMS
  • Elmer’s Glue (polyvinyl acetate)
  • Food coloring
  • Applicator stick
  • 1ml syringe
  • 27ga blunt tip needle

What do I do?


  1. Put 2-3 ml of Elmer’s Glue in a small container and mix 1-3 drops of food coloring, depending on how bright you want the color to be. Mix a large enough amount of colored glue so that you can draw it up into the syringe without adding bubbles.  Larger volumes are easier to draw into the syringe.
  2. Express some of the glue out of the syringe so that you do not introduce any bubbles into the PDMS.
  3. Mix PDMS in the usual 10:1 ratio and pour over your wafer, checking for bubbles.
  4. Gently insert the syringe needle into the PDMS and inject a small amount of glue. Injected glue tends to stay where it is injected (Fig. 1).
  5. Carefully remove the syringe from the PDMS.
  6. Cure the PDMS as normal (Fig. 2).
Image of colored glue injected into PDMS

Fig.1: Injected glue tends to stay where it is injected

Cured PDMS

Fig. 2: Cure the PDMS as normal

What else should I know?


An applicator stick may be used instead of a syringe. Also, this procedure will work on top of the PDMS, but it will change its surface.

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A stacked microfluidic device for improving experiment throughput

Jiandong Wuab, Xun Wubc and Francis Linabcd*
a Department of Biosystems Engineering, University of Manitoba, Canada
b Department of Physics and Astronomy, University of Manitoba, Canada
c Department of Immunology, University of Manitoba, Canada
d Department of Biological Science, University of Manitoba, Canada
Email: flin[at]physics.manitoba.ca

Why is this useful?


Owing to the advantages in miniaturization and cellular microenvironmental control, microfluidic devices have been increasingly applied to cell biology research [1]. Particularly, microfluidic devices can precisely configure chemical concentration gradients and flexibly manipulate the gradient conditions in space and in time [2, 3]. Various microfluidic gradient-generating devices have been used for studying cell migration and chemotaxis [2, 3]. These studies rely on live cell microscopy and usually only one experiment can be performed at a time. Previously, a double gradient device was demonstrated for parallel cell migration experiment with a motorized stage to image cells in different gradient channels [4]. However, the full XYZ motorized stage is expensive and thus often times limits the practical use of high-throughput microfluidic devices.

To overcome this limitation, here we report a stacked microfluidic device that allows parallel live cell imaging experiments on a single chip with only a Z motorized stage. This device is fabricated with multiple stacked layers of PDMS devices, and the cell imaging channels in each layer are aligned so they all fit into a single microscope viewing field. Thus, by only adjusting the vertical focus using a Z motorized stage, multiple cell channels can be imaged repeatedly over time. If a full XYZ motorized stage is available, the throughput of the stacked device can be further increased along horizontal dimensions. Making a stacked device is straightforward and this strategy can be useful for improving experiment throughput, especially in a limited microscopy facility.

What do I need?


  • Two identical PDMS chips and a glass coverslide
  • An oxygen plasma cleaner
  • A fluorescent microscope equipped with a programmable motorized Z stage or a full XYZ motorized stage, and a CCD camera

What do I do?


  1. Fabricate your SU-8 mask using standard photolithography. We used a simple ‘Y’ shape design with a 350µm wide main channel.
  2. Make two PDMS replicas of the same design from your SU-8 masks using a standard soft-lithography method.
  3. Bond the two PDMS chips using O2 plasma treatment. The channels should all face down and the main channels in the two chips should be vertically aligned. To avoid overlap of the inlets, we align the two layers so that their inlets are separated by some distance, while the main channels of the two layers are still within the same viewing field in the microscope. We used a 10X objective in our experiment, and this strategy works well with a 350µm channel. However, this method may not work if higher magnification is required, and it may also cause shifted gradients in the two layers. As an alternative strategy, we can align the two layers perfectly along the vertical direction, but punch inlet holes for the two layers with different distance relative to the ‘Y’ junction so the inlets of the two layers can be separated (this will require punching inlet holes for the top layer before plasma bonding). Two masters with different inlet designs can also be used to separate inlets of the stacked device.
  4. Punch the holes for making fluidic inlets and outlets.
  5. Bond the double-layer PDMS chips to a glass coverslide using O2 plasma or air plasma treatment to complete the simple stacked device (Figs. 1A & 1B).
  6. Image the channels and cells using a microscope equipped with a Z motorized stage.
  7. We used food coloring to show the channels in the two layers of the stacked device (Fig. 1B). Furthermore, we show that we can generate chemical gradients in each layer of the stacked device by mixing buffer and FITC-Dextran as well as imaging cells loaded to the main channel of each layer only with a Z motorized stage (Fig. 2).
  8. Finally, using a different design that consists of two gradient channels in each layer and a full XYZ motorized stage, we demonstrate that four individual channels can be imaged in the stacked double-layer device. Again, food coloring is used to show the channels in the upper layer and the bottom layer (Fig. 1C).

Figure 1
Fig. 1. Illustration of the stacked microfluidic device. (A) The schematic drawing of the stacked double-layer device that consists of 2 identical ‘Y’ shape channel; (B) A real picture of the stacked device of the same design. Food coloring is used to show the channels in the 2 layers. (C) A real picture of the stacked device of the design that consists of 2 gradient-generating channels in each layer. Again, food coloring is used to show the channels in the 2 layers.

Figure 2
Fig. 2. Gradient generation and cell images in the double-layer stacked device. (A) Gradient of FITC-Dextran 10kDa generated in the bottom layer channel using the ‘Y’ shape design, and images of Jurkat cells in the same channel. (B) Gradient of FITC-Dextran 10kDa generated in the upper layer channel using the ‘Y’ shape design, and images of Jurkat cells in the same channel. Only a Z motorized stage is used for the imaging.

What else should I know?


Here we demonstrate the simple double-layer stacked device. More layers of PDMS can be stacked to further increase the throughput. In addition, in the current demonstration, the channel in the bottom layer is formed between PDMS and a coverslide while the channel in the top layer is formed between PDMS. If needed, the double-layer chip can be first bonded to another piece of PDMS before bonding to the glass substrate. This way, both layers of channels are in PDMS for consistency.

References


[1] G. B. Salieb-Beugelaar et al., Latest developments in microfluidic cell biology and analysis systems. Anal. Chem., 2010, 82, 4848-4864.
[2] S. Kim, H. J. Kim, and N. L. Jeon, Biological applications of microfluidic gradient devices. Integr. Biol., 2010, 2, 584-603.
[3] J. Li and F. Lin, Microfluidic devices for studying chemotaxis and electrotaxis. Trends Cell Biol., 2011, 21, 489-497.
[4] W. Saadi et al., A parallel-gradient microfluidic chamber for quantitative analysis of breast cancer cell chemotaxis. Biomed. Microdevices, 2006, 8, 109-118.

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A refinement of a method to prevent sagging during the bonding or lamination of chips with high aspect ratio chambers

Brian Miller, Stewart Smith and Helen Bridle refine a method to prevent the sagging of high aspect ratio channels during bonding, which allows the technique to be applied to shallower channels for devices in which performance is sensitive to channel height

Brian Miller, Stewart Smith and Helen Bridle*
School of Engineering, University of Edinburgh, 3.17 William Rankine Building, Kings Buildings, Edinburgh, EH9 3JL, UK
Email: h.bridle[at]ed.ac.uk

Why is this useful?


Jie Xu and Daniel Attinger previously described a method to prevent the sagging of high aspect ratio channels during bonding [1]. The method involves careful placement of salt crystals into the channel prior to bonding to create a supporting structure. A limitation of the technique is the depth of channels this method can be used on, which must be within the size range of the salt crystals (~100 μm).

Described here is a method that builds on this technique to allow salt structures to be created with much smaller surface profiles, down to between 25-35 μm maximum heights of profile. This will allow the technique to be applied to shallower channels for devices in which performance is sensitive to channel height, for example, inertial focusing devices (Fig.1) [2,3].

This refined technique also helps to simplify the handling of the devices after preparation; reducing the risk of contamination of equipment as, once they are applied, the salt crystals are adhered to the surface of the device.

Inertial focusing devices
Figure 1. Inertial focusing device A; with solution applied to high aspect-ratio sections as indicated in pinched-flow segment B and output leg C. Device depth is 30μm (max. aspect ratio 60:1 in PDMS)

What do I need?


  • High purity KCl (potassium chloride salt)
  • De-ionised (DI) or reverse osmosis (RO) water
  • Weighing balances/scales
  • Decon 90 surfactant
  • Hypodermic needle (thin gauge)
  • Magnetic stirrer
  • Beaker and syringe

What do I do?


  1. Prepare a solution of the salt and surfactant by measuring 100ml of DI or RO water into the beaker. Add 1.5g of KCl to the water and stir gently for two minutes. Use the syringe to add 5ml of Decon 90 to the solution and stir very gently for a further two minutes, taking care not to foam the solution. Do not allow the solution to rest for more than a few minutes after stirring.
  2. Bending the sharp point of the hypodermic needle on your workbench or other hard, clean surface can help to ‘grab’ sufficiently small quantities of solution from the beaker. Use the needle or a similar applicator to carefully apply a small quantity of solution to the high aspect ratio section of your channel (Fig. 2). Only a very small volume is required and it is important to not allow the solution to overflow the channel. Dabbing the needle on a dust/lint-free wipe can help to regulate the amount of solution delivered to the surface of your device. If you accidentally over-apply the solution, clean the device with DI/RO water and IPA, and re-attempt application once it has dried.
  3. Allow the solution to evaporate at room temperature without assistance of any kind (no added air-flow or heat). An area of crystal formation should form where the solution was applied. The edges of this area tend to grow larger due to the ‘Marangoni effect’. The edge will typically be a maximum of 30-35μm in height, with the centre of the area typically yielding crystal formations between 10 and 25μm tall (as measured on a surface profiler over 4 repetitions), which should suffice to prevent the unintended bonding of the ceiling to the floor of the channel.
  4. Bond your device to your substrate layer following your normal bonding procedure (we used oxygen plasma bonding of PDMS to a glass microscope slide in this example).
  5. Before using the device, run water or buffer through to dissolve away the crystal formations (Fig. 3). The surfactant helps to quickly remove the salt structures and leave the device clean of any remnants.

Application of solution into channels
Figure 2. Careful application of very small quantities of solution into channels, using a hypodermic needle

Support structures before and after rinsing
Figure 3. Top: water salt formations in the support structure before rinsing with DI/RO water. Bottom: the same area after rinsing through, illustrating that practically no residue is left in the device

Notes


  • Failure to use the surfactant will result in much larger crystal formations, as the crystal structures nucleate in very few locations and form much deeper structures. It is conjectured that the surfactant suspends the salt ions with a larger interspaced distribution throughout the solution, causing a much more distributed nucleation of the crystals and yielding the lower profile crystal formations.

References


[1]  Jie Xu and Daniel Attinger, How to prevent sagging during the bonding or lamination of chips with large aspect ratio chambers, Chips & Tips (Lab on a Chip), 24 July 2009.
[2] D. Di Carlo, D. Irimia, R. G. Tompkins and M. Toner, Continuous inertial focusing, ordering, and separation of particles in microchannels. Proc. Natl. Acad. Sci. U. S. A., 2007, 104, 18892-18897.
[2] D. Di Carlo, J. F. Edd, D. Irimia, R. G. Tompkins and M. Toner, Equilibrium separation and filtration of particles using differential inertial focusing. Anal. Chem., 2008, 80, 2204-2211.

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Robust and easy macrofluidic connections in acrylic

Robert Henderson, Nick Selock and Govind Rao present a simple scheme to connect a standard HPLC flat bottom nut to a PMMA microfluidic device

Robert Henderson, Nick Selock and Dr. Govind Rao*
Center for Advanced Sensor Technology, and Department of Chemical, Biochemical, and Environmental Engineering, University of Maryland, Baltimore County, 1000 Hilltop Circle, Baltimore, MD 21250, USA
Email: grao[at]umbc.edu

Why is this useful?


The use of HPLC fittings and tubing in microfluidics is becoming more commonplace as the use of microfluidics in the sciences increases. Even though acrylic or PMMA is one of the most common plastic substrates used in microfluidics, the ability to strongly and easily connect an HPLC fitting to a PMMA chip has until now been elusive. Although there are some commercially available HPLC-to-chip products from Idex, they require heating the microfluidic chip well above the glass transition temperature of PMMA to create a permanent epoxy bond between attachment and chip. For these reasons, we present a simple scheme to strongly connect a standard HPLC flat bottom nut to a PMMA microfluidic device through the creation and bonding of an easy-to-make ¼-28 PMMA microfluidic union, using common lab equipment.

What do I need?


  • Laser cutter or another implement to cut PMMA, such as a hacksaw or band saw
  • An appropriate piece of equipment to anneal the PMMA (we use a convection toaster oven)
  • 5.3mm thick acrylic sheet for the body of the PMMA microfluidic union
  • A drill and drill bits
  • Vise grips, or another way to hold the PMMA union in place while tapping
  • A ¼-28 tap
  • A few sheets of progressively finer sandpaper
  • A 50-50% mixture of ethanol and water, along with an ultrasonic cleaner and lint-free wipes for cleaning purposes
  • A strong solvent for acrylic, such as chloroform or methylene chloride
  • A glass syringe with a blunt needle or needle blunted through filing

This connection scheme has been tested with good results when using the LT-115 and P-259 ¼-28 flat bottom HPLC nut and ferrule combination for 1/16″ OD tubing (Idex). This method was tested on microfluidic devices that were not bonded with the use of adhesives. This method is adaptable to other flat bottomed nuts as long as an appropriate tap and drill bit are used. Keep in mind that as the wall thickness of the PMMA union decreases, so does the strength of the connection between the HPLC nut and the device.

What do I do?


  1. Use a laser cutter to cut a circle with a diameter of 13mm out of the 5.3mm thick acrylic sheet. Make sure that you also either etch a center mark or make a small center hole using the laser cutter. If a laser cutter is not available, make a 13mm PMMA square using one of the other mentioned cutting implements. (Fig.1)
  2. Using a small drill bit, drill a pilot hole for the larger drill bit to follow in the center of the circle that you cut out in Step 1.
  3. Using the larger drill bit (a #3 or 0.213″ drill bit for the ¼-28 tap), drill the center hole in the cutout piece following the pilot hole drilled in Step 2.
  4. Using the ¼-28 tap and appropriate technique, tap the hole you have been creating in Steps 1-3. Do not use oil to lubricate the plastic as it is unnecessary and difficult to clean.
  5. After removing the plastic chips from the threads you just created, use progressively finer sandpaper to lap the surface that will make contact with your microfluidic device. Lapping the surface will ensure a very strong bond between the union and your device. We lap using a figure of eight motion to help guarantee a flat bonding surface. (Fig. 2)
  6. Anneal both your created microfluidic union and your device in an appropriate fashion. Annealing will relieve the thermal stress present in the union due to laser cutting and the drilling/tapping steps. (We use a convection toaster oven, set at 85ºC for 90 minutes, 50ºC for 30 minutes and then off with the oven door closed for 30 minutes. We find it useful to put a smooth metal plate below and above the piece to be annealed, to prevent uneven heating in the oven.)
  7. Thoroughly clean both the union and the microfluidic device to be bonded using a 50-50% mixture of ethanol and water. Clean only the union in an ultrasonic cleaner for 10 minutes. This mixture will not cause the union or the device to crack if they are both appropriately annealed beforehand and if the cleaning is performed within a few days of annealing.
  8. Place the union, lapped side down, onto the microfluidic device’s surface where the connection is desired. It is essential to center the union over the hole which receives the fluid or gas from the HPLC tubing. We use a small jig composed of two different sized plastic dowels to help us in this step (the black object in the bottom right corner of Fig.3).
  9. Pull some solvent into the glass syringe.While holding the union in place using your jig, gently drip some solvent from the glass syringe at the interface between the union and your microfluidic device (start with a single drop). The solvent will creep into the joint between the union you created and your microfluidic device due to capillary action. Add enough solvent to fill the gap between the union and your device, but be careful not to get any solvent into the fluid passages within your device.
  10. Being careful not to disturb the union’s position, gently apply pressure on the union until the surfaces become lightly bonded using your gloved fingers (Fig. 3). Light bond is usually achieved in less than a minute. In this step, also attempt to add enough pressure to force out any bubbles at the interface.
  11. Remove the alignment jig and add more solvent around the interface between the union and your device. This step reinforces the union’s connection to the microfluidic device.
  12. Put a small mass directly on top of the union, and leave the bonding union overnight to achieve the strongest connection between the union and your device.
  13. After the piece has dried overnight, remove the mass and then use the connection as you would any other flat bottom HPLC union. (Figure 4)

Raw PMMA union with laser centered hole
Figure 1. The dimensions of the raw PMMA union
Lapping the surface of the microfluidic device
Figure 2. Lapping the surface of the microfluidic device with a figure-eight motion
Bonding union and microfluidic device
Figure 3. Lightly bonding the union to the microfluidic device using gentle pressure
The finished microfluidic union
Figure 4. The finished microfluidic union

Notes


The following YouTube video ‘Power Tools & Carpentry Skills : How to Use a Tap & Die Set’ by Expert Village helps explain the use of taps (Step 4): http://www.youtube.com/watch?v=6DDhw191MLo

Acknowledgements


The authors would like to acknowledge the work of Dr. Yordan Kostov, Mike Tolosa and Mike Frizzell.

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Easy and inexpensive fabrication of PDMS films of different thicknesses

Rodrigo Martinez-Duarte shows a way to easily fabricate method for PDMS by squeezing it between plates

Rodrigo Martinez-Duarte
Microsystems Laboratory (LMIS4), École Polytechnique Fédérale de Lausanne, Station 17, CH-1015 Lausanne, Switzerland

Why is this useful?


The following tip describes an easy and inexpensive way to fabricate PDMS films of different thicknesses. The main advantages are that no infrastructure (e.g. a spin coater) is needed for fabrication and that the materials needed are readily available. The idea here is to use a film of a specific thickness as spacer between two plates. PDMS is first deposited on one plate and squeezed in between plates to get a PDMS film with thickness similar to that of the film used as spacer. A similar methodology could be used to make films of other materials as well.

This methodology allows for very quick fabrication of a wide range of PDMS films using off the shelf components which are common in a traditional laboratory and office or can be easily purchased. The final goal is to fabricate films which feature different thicknesses, according to the original film used as spacer, and different hardnesses, by changing the ratio of PDMS to cross linker. The film can be stored and used as needed to cut off parts such as gaskets, spacers, etc. Here in the lab we use them to make microfluidic chambers of specific thickness.

What do I need?


  • 2 rigid plates with at least one flat surface per plate. They can be glass, PMMA (>3-4 mm thickness) or other material as long as they are rigid, preferably a fair thermal conductor and do not soften at temperatures up to 100°C. The use of transparent plates is not necessary but recommended to monitor the squeezing process of the PDMS and minimize the presence of bubbles in the final PDMS film
  • Pieces of a film, which can be tape, pieces of a plastic bag or any film that does not compress or  absorbs PDMS, with similar thickness to that being targeted for the PDMS film
  • Paper clamps, binder clips
  • General purpose soap or Mylar® film, the Mylar® film can be replaced by any film to which PDMS won’t adhere
  • And of course, PDMS and the equipment recommended to process it: balance, degasser and oven. Although this equipment is recommended, it is not necessary, as already suggested by Aung K. Soe and Saeid Nahavandi in a previous Tip [1]

Where do I get it and how much will it cost me?


In principle all the materials required are already available in the lab. The two plates can be two pieces of glass, as simple as two glass slides or two old wafers, or pieces of plastic, polycarbonate or PMMA for example. I have used PMMA plates and old glass wafers. The cost of the plates can be minimal and should not be more than $10. For example, you could go and buy a couple of very cheap picture frames with a glass piece of the size you want and use those. The pieces of film you use depend on the final thickness of the PDMS film and they can be obtained from a variety of sources, use your imagination! The paper clamps or binder clips are usually available in the office, if not you can buy them for several cents apiece.

What do I do?


  1. Wear gloves before starting. PDMS can be messy! The materials needed are detailed above and shown in Fig. 1.
  2. A. Dip your rigid plates in soapy water for a couple of minutes to deposit a layer that prevents PDMS from adhering to the plates. Remove the plates and blow them dry or let them dry naturally, do not wipe dry!
    Or alternatively:
    B. You may want to use a Mylar® film in between the rigid plate and PDMS. Take care not to scratch the Mylar® film. The rationale behind using this Mylar® film is to eliminate the need for surface treatment of your plates to prevent adhesion of the PDMS to them. PDMS does not stick to Mylar® and therefore you can easily just peel the Mylar® film off after you fabricate your PDMS film. A further advantage is that you don’t need to worry about scratching your plates or about cleaning the plates each time you use them. Storing your PDMS film between Mylar® films can also help protect it against scratches.
  3. Prepare your PDMS. Different ratios between the polymer and the cross-linker give different hardnesses. A 20:1 mix results in a gluey, soft film that easily sticks to a variety of materials. A traditional 10:1 results in a harder film. 5 grams of mix should be plenty for a couple of films featuring an area of 8 by 8 cm and thickness of up to 200 µm.
  4. A. If you treated the surface of the plates with soap: after the plates are dry (but still with the dried soap layer on them), you can position your spacer film close to the edges of the plate as shown in figure 2. Do this on only two edges to allow plenty of space for the PDMS to flow out from between the plates during step 7.
    B. If you are using the Mylar® films: cut the film into a couple of pieces similar to your plates. You can then position the spacer film close to the edges of one of the Mylar® films. Leave some space in between the spacer films to allow PDMS to flow out while pressing in step 7.
  5. After the PDMS is well mixed, manually deposit the PDMS mix on one plate treated with soap, or on the Mylar® film positioned on top of the plate, as shown in figure 3.
  6. Degas the previously deposited PDMS until the mix looks homogeneous (around 40 minutes in a general purpose degasser).
  7. Remove the arrangement from the degasser, place it on a flat surface and use the other plate to squeeze the PDMS in between the plates if you are using the soap-treating approach. If you are using the Mylar® films, then first lay the second film on top of the PDMS and then squeeze with the other plate. In both approaches, it is recommended to start applying pressure in one side and work your way towards the other side to avoid introducing bubbles in the PDMS.
  8. Use the paper clamps to clamp A) the plate-PDMS-plate or B) plate-Mylar®-PDMS- Mylar®-plate sandwich together at the location of the spacers as shown in figure 4.
  9. Bake for 1 hour at 80°C.
  10. Remove from oven and let cool for 5 min.
  11. A. If using soap: use a knife to remove the PDMS accumulated on the edges of the plates. This is important because it will facilitate the release of the PDMS film in the next step.
    B. If using Mylar® films: unclamp your sandwich and retrieve your PDMS film in between the Mylar® films. You are done!
  12. If using soap-treated plates: slowly but firmly separate the two plates, the PDMS film is likely to remain on one plate from where you can peel it off as shown in figure 5. Make sure you do it slowly to avoid rupturing the film. You may use fine tweezers to aid you during the release.
  13. You are done! You can store your film and later use a hole puncher, knife, etc. to cut off any feature you want.
  14. Clean the PDMS from your plates and store them clean for the next time. Avoid scratching the surface of your plates!

Notes


  • Tape (175, 140, 70 and 50 µm thick) and plastified paper (50 µm thick) have been used as spacers.
  • This methodology has been used to fabricate films as thin as 50-70 µm. The exact thickness of the PDMS film is difficult to measure due to the soft nature of the film.
  • The use of soap may interfere with further applications of the PDMS film. However, here in the lab we haven’t seen any adverse effects so far.

Materials needed
Fig.1 Materials needed: paper clamps, rigid plates and pieces of film (red arrows)

Film on edges
Figure 2. Pieces of film (red arrows) positioned close to the edges of the rigid plate shown on the left.

PDMS deposition
Figure 3. Manually deposited PDMS (blue arrow) on one of the plates

Squeezed PDMS
Figure 4. Squeezed PDMS between rigid plates using paper clamps

Film release
Figure 5. Release of the PDMS film from one of the plates after it has been cross-linked by heating

References


[1]  Aung K. Soe and Saeid Nahavandi, Degassing a PDMS mixture without a vacuum desiccator or a laboratory centrifuge and curing the PDMS chip in an ordinary kitchen oven, Chips & Tips (Lab on a Chip), 26 May 2011.

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