2009 – Chips and Tips

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A simple and economical holder for casting PDMS chips

20 Nov 2009

Gang Li*, Qiang Chen and Jianlong Zhao

Shanghai Institute of Microsystem and Information Technology, Chinese academic of Sciences, Shanghai, China

Why is this useful?

To fabricate PDMS chips, one usually pours an un-cured PDMS mixture into a Petri dish or an aluminum foil tray containing the patterned master, and then cures for 2 h at 85°C. However, these casting dishes or trays present some cost problems. Often a relatively high consumption of PDMS is demanded for these casting holders because quite a part of PDMS will seep into the gap between the wafer and trays due to capillary forces during PDMS pouring. On the other hand, the seepage of PDMS under the master wafer may tilt the master causing the thickness of the PDMS mold to vary from one side to the other. Furthermore these casting dishes or trays are generally throw-away holders for casting PDMS chips, which also increases the fabrication cost of PDMS chips.

This tip presents a new casting holder, made of a hollow plate whose bottom is enveloped by an adhesive tape. This casting holder can simplify the operation of releasing the cured PDMS casting, and minimize the consumption of PDMS. In addition, this holder can be used to fabricate double-side flat PDMS chips by placing a flat substrate on its top.

What do I need?

Circular hollow plate (home-made of PMMA)
Adhesive tape (Nitto Denko Corp., SPV-224)
PDMS (Dow Corning Sylgard 184)
Hotplate (Chemat Technilogy Inc., KW-4AH hotplate)
PET (Polyethylene terephthalate) film and flat glass slide (for optional double-side flat PDMS chips)

What do I do?

1. Seal the bottom opening of the hollow plate with adhesive tape to form a casting holder (Figure 1), whose diameter is a little bigger than that of master wafer.

: Figure 1

2. Place the master in the holder, and carefully apply a pressure to attach it to the tape, preventing any gaps from forming at the interface (Figure 2).

: Figure 2

3. Mix the PDMS according to the manufacturer’s procedure [1].

4. Pour the PDMS mixture in the casting holder, and then place the holder on a hotplate at 85ºC for 2 hours.

5. Once the PDMS is cured, remove the holder from the hotplate and allow it to cool to room temperature.

6. Gently peel off tape from the bottom of holder (Figure 3).

: Figure 3

7. Carefully cut around the edge of the PDMS mold using a sharp scalpel, and separate the PDMS from the holder (Figure 4).

: Figure 4

8. Finally, peel the PDMS mold from the master wafer (Figure 5).

: Figure 5

9. (Optional, for double-side flat PDMS chips) Prepare the casting holder as described above, pour the PDMS mixture in the casting holder until the level of PDMS is a little higher than the top of holder, and then carefully drop a PET film onto the prepolymer mixture (Figure 6), which provides an easy way to remove the cover plates from the PDMS molds after curing.

: Figure 6

10. Apply a pressure on the top of the film with a stack of glass slides and steel block to planarize the surface of the PDMS chips, and then cure the PDMS by placing the holder on a hotplate (Figure 7).

: Figure 7

11. After curing, remove the glass slide and steel block from the top of holder, and then gently peel off the tape and PET film from the PDMS block (Figure 8).

: Figure 8

12. Finally, separate the PDMS block from the holder and peel the PDMS mold from the master.

Acknowledgements

This material is based upon work supported by the Major State Basic Research Development Program of China (No. 2005CB724305), and the National High Technology Research and Development Program of China (No.2006AA02Z136).

References

[1] Dow Corning Product Information, “Information about Dow Corning® brand Silicone Encapsulants,” 2005.

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“Custom made” production of cheap Luer lock adapters for chip-to-syringe interfacing

20 Nov 2009

Stefania Mazzitelli, Stefano Focaroli and Claudio Nastruzzi
Department of Chemistry and Technology of Drugs, University of Perugia, Perugia, Italy

Why is this useful?

One of the standard procedure to pump solutions, emulsions or suspensions, into microfluidic chips is based on the use of syringes, through peristaltic pumps. Syringe pumps are usually preferred over peristaltic ones, for their ease of use, for the accurate and stable control of the flow rate and, finally, for the possibility to employ sterile conditions.

Luer taper is a standardized system of small-scale fluid fittings used for making leak-free connections between a male-taper fitting and its mating female part on medical and laboratory instruments, including hypodermic syringe tips and needles or stopcocks and needles. The fitting is named after the 19th century German medical instrument maker Hermann Wülfing Luer.

There are two varieties of Luer Taper connections: Luer-Lok and Luer-Slip; Luer-Slip fittings simply conform to Luer taper dimensions and are pressed together and held by friction (they have no threads). Luer components are manufactured either from metal or plastic and are available from many companies worldwide but they are usually sold in few standard dimensions and are relatively expensive.

In this Tip we present a way to easily produce a variety of “on demand” Luer connectors, including: (a) female Luer x female Luer adapter, (b) female Luer x female Luer elbow, (c) male Luer x male Luer adapter, (d) male Luer x male Luer elbow, (e) male Luer x female Luer coupler and finally (f) male Luer x female Luer elbow.

What do I need?

1-30 mL polypropylene syringes (Artsana, Italy) [1]
tubing: Upchurch Scientific® FEP (fluorinated ethylene-propylene); Tub FEP Nat 3/16 x .125 x 20ft (Upchurch Scientific, UK; No.: 1524) [2]
tubing: Timmer-Pneumatik GmbH; H-PTFE-4/2 mm (OD/ID)-blue, catalog Timmer 2001 [3]
Aesculap scalpel blades fig. 23, carbon steel, package of 100 pieces in dispenser package [4]
Aesculap scalpel handle fitting no. 4 for blades 18-36, 135 mm, 5 ¼, [5]
Black & Decker heat gun, model kx1693, [6]

What do I do?

1. Tubing (A, B) and template syringes (C) are used for the production of Luer connectors. A: FEP (fluorinated ethylene-propylene) tubing; B: H-PTFE tubing.

: Figure 1

2. Tubing cutting by scalpel. The length of the Luer connector can be adjusted depending on the specific needs of the researcher (for instance, the distance between the syringe pump and the chip). Note that in the case of H-PTFE tube (B), the cutting is made with an angle of 45° with respect to the tube major axis.

: Figure 2

3. Preparation of a female Luer adapter: A. Heating of a FEP tube end by heat gun for 1-3 min; B-D. Press and insert the male Luer of a polypropylene syringe into the heated tube. E. Cool down the tube by tap water. F. Permanent deformation of the tube end.

: Figure 3

4. Preparation of a male Luer adapter: A. Heating of a FEP tube end by heat gun for 1-3 min; B-C. Press and insert the H-PTFE tube end (cut at 45°) into the heated FEP tube, cool down the tube connection by tap water until a permanent deformation is reached.

: Figure 4

5. Examples of female Luer X female Luer adapter (A) and female Luer X male Luer adapter (B).

: Figure 5

6. Examples of female Luer X female Luer elbow (A) and female Luer X male Luer elbow (B).

: Figure 6

7. Examples of the use of female Luer X female Luer adapter to connect syringes to a commercial chip.

: Figure 7

8. Examples of the use of female Luer X male Luer elbow to connect syringes to a homemade chip.

: Figure 8

References

[1] http://www.artsana.com
[2] http://www.upchurch.com
[3] http://www.pneumatica-it.timmer-pneumatik.de
[4] http://www.chirurgische-instrumente.info/en/search.html?kw=bb523
[5] http://www.chirurgische-instrumente.info/en/search.html?kw=bb084r

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A novel method for fabrication of reusable microfluidic interconnects

28 Oct 2009

Koesdjojo, M.T., Mandrell, D.T., Tennico, Y.H., Remcho, V.T.*
Department of Chemistry, Oregon State University, Corvallis, Oregon, USA

Why is this useful?

The lack of an efficient interface, or interconnect, between microfluidic devices and the macroscale world is a major impediment to the broader application of micro total analysis system (µTAS). There are a number of current products and solutions available that seek to address this issue. An example of a simple approach is the direct integration of tubing or a syringe needle into the microchip inlet using epoxy glue.[1] However, direct connection to the inlet reservoir using an epoxy often leads to clogging of the microchannels. Commercially available connections to microfluidic chips and devices, such as the NanoPort from Upchurch Scientific (Oak Harbor, WA, 98277, USA), accommodate these needs by use of a threaded nut and ferrule system. These fittings can be integrated onto chip reservoirs by means of adhesive rings or epoxy glue.[2] The drawback to this method is that NanoPorts require the use of an epoxy glue that takes time to cure and is non-removable, or an adhesive ring that is not compatible with many solvents. Another drawback is that the epoxy requires high temperatures for complete curing, making it incompatible with low glass transition (Tg) polymers.[3] Commercial edge connectors provide efficient interfacing and allow rapid connections and reusability.[4] However, they are designed to be interfaced to a standard microchip format; which limits the compatibility of other chips with different geometries layout or sizes.

This tip presents an alternative approach to making simple, cost-effective universal interconnects. The device was developed to allow for standard compression tubing connectors, such as those available from Upchurch, to be interfaced with microfluidic chips. A standard breadboard fixture was used to compress the ports against the chip to apply sealing pressure, preventing leaks at the interface. This method allowed for connections to be made without glue, epoxy, or any form of bonding, enabling the components to be easily and quickly reused or reconfigured without the need to machine new fixtures or re-bond ports. The working pressure of these ports was tested with a syringe pump, and withstood pressures in excess of 1000 psi without any signs of leakage or failure.

What do I need?

0.25 and 0.5 inch thick plastic substrates such as polycarbonate (PC)
#21, #9 drill bits
10-32 bottoming tap
1mm drill bit
1/16 inch PEEK or Teflon tubing
stainless steel 10-32 socket head cap screws
1/32 inch Silicon o-rings
Upchurch 10-32 ferruled fittings (F-333)

: Figure 1. Picture of system components

What do I do?

1. The first component was a port which allowed a standard ferruled connector to be attached perpendicular to the microfluidic device. It contained a threaded portion for the fitting and a fluid path with small port on the bottom for the interface (Figure 2). The port was manufactured using 0.25 inch PC, but can be made of any material thick enough to allow room for the ferruled connector between the compression plates. It was machined to size using a knee mill. The threaded port was drilled with a #21 drill bit, then tapped with a 10-32 bottoming tap. The fluid paths were drilled using a 1mm drill bit (substitute any drill to achieve the desired internal volume). An o-ring was then placed in the threaded hole to ensure a tight seal against the ferruled fitting.

: Figure 2. Interconnect section view (left) and the solid model (right).

2. The second component was a breadboard clamp which applied pressure between the microfluidic device and interconnects (Figure 3). This breadboard approach was particularly useful for prototyping devices having different dimensions and port locations, as interconnects could easily be relocated. A wide variety of bolt locations in the clamping plates allowed for a wide variety of component sizes and configurations. The breadboard clamp assembly was manufactured by cutting 0.5 inch thick PC into two five inch squares. Using the knee mill, a one inch grid of holes was drilled and taped on one plate with the #21 drill and 10-32 tap. Clearance holes were drilled on the same grid, using a #9 drill bit, in the second clamping plate.

: Figure 3. Side view of a chip, interconnects, and the compression plates.

3. The 90° interconnect ports were placed on each of the reservoir holes of a microchip. The ferrules were connected to their respective tubing. The top plate of the breadboard clamp was installed and using the holes closest to the sides of the chip, 10-32 stainless steel screws were inserted to apply even pressure by clamping the entire fixture. Figure 4 shows the setup used to pressure-test the manufactured ports.

: Figure 4. Setup used for pressure testing (left) and the close up view of a chip in the device (right).

References

[1] T. Das, Interfacing of microfluidic devices, Chips & Tips (Lab on a Chip), 27 February 2009.
[2] http://www.upchurch.com/
[3] J. Greener, W. Li, D. Voicu, E. Kumacheva, Reusable, robust NanoPort connections to PDMS chips, Chips & Tips (Lab on a Chip), 8 October 2008.
[4] http://www.dolomite-microfluidics.com/

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Plug-and-play reservoirs for microfluidics

28 Oct 2009

Christopher Y. Wong*, Georgian Ionut Ciobanu*, Mohammad A. Qasaimeh, and David Juncker
Micro and NanoBioEngineering Laboratory, Biomedical Engineering Department, Faculty of Medicine, McGill University, Montreal, Canada.
* Authors contributed equally

Why is this useful?

Microfluidic and lab-on-a-chip (LOC) devices are typically connected to external sample reservoirs. A significant issue for LOC devices is interfacing multiple sample reservoirs and connecting them both to external pressure controllers or syringe pumps and to the microfluidic chip [1,2]. Researchers typically use home-made reservoirs and interfacing them to the chip and the controllers requires specialized skills and training and/or additional fabrication steps.[3,4] Often the connections are unreliable and leaky, and as a consequence much time is lost in setting them up and it is difficult to add or remove connections from the reservoirs.

Here we introduce a “plug-and-play” reservoir that allows for multiple connections to be simply plugged in. The reservoirs are made using off-the-shelf threaded microcentrifuge tubes and screw-on septum caps. Needles, capillaries and steel tubing can be inserted one at a time – or in combination – through the cap’s septum and used to connect the reservoir to a pressure source and to one or several inlets of a microfluidic chip. The connectivity between reservoir and pressure source(s) and chip(s) can be adapted to specific needs, for example to form a reservoir with a single input and a single output, or with multiple inputs and outputs. Reservoirs with single or multiple input/output lines can be switched easily by by unscrewing the microcentrifuge tube and screwing the cap with the capillaries on another microcentrifuge tube containing a different solution.

The reservoirs are cheap, disposable, easy to assemble, and robust enough to support pressures up to 345 kPa (and even higher pressures with the variant described in the supplementary material) while having almost no dead volume. The connection of reservoirs to microfluidic chips – or to other reservoirs so as to form complex circuits – can be performed in a matter of seconds. Arrays of off-chip reservoirs can be assembled within racks and many lines can be connected at high density to a microfluidic chip. These plug-and-play reservoirs are a versatile solution to the world-to-chip interface and should find widespread application for microfluidic experimentation.

What do I need?

For the reservoir:

threaded screw cap with membrane/septum [5]
threaded microcentrifuge tube (any volume, as long as it has the same thread type as the cap) [5]

For the connections:

sharp hypodermic needle with Luer-Lock (any size from 23 to 16 gauge) [6]
1/16″ ID Tygon tubing or other preferred tubing [7]
male Luer-lock to 1/16″ ID hose barbed tubing adapters [7]
glass capillary (Note: OD of glass capillary needs to be smaller than ID of needle) [8]

Optional for other connection types (shown in supplementary materials):

0.0250″ OD hollow steel tubing [9]
0.0200″ ID microbore Tygon Tubing [10]

Fig. 1 shows the required parts. Fig. S1 in the supplementary materials shows the alternative options in our laboratory of microcentrifuge tube sizes and input/output connection types. (A) Threaded screw cap with membrane/septum [5]. (B) Threaded microcentrifuge tube [5]. (C) Sharp 20G1½ hypodermic needle with Luer-Lock [6]. (D) 1/16″ ID Tygon tubing [7]. (E) Male Luer-lock to 1/16″ ID hose barbed tubing adapter [7]. (F) Glass capillary of 350 µm OD [8].

: Figure 1. Different parts used to make the reservoir.

What do I do?

1. In order to insert a glass capillary to serve as an output, use a sharp hypodermic needle with an inner diameter (ID) that is just larger than the capillary’s outer diameter (OD). Insert the needle into the membrane screw cap and then slide the capillary through the needle, as shown in Fig. 2.

: Figure 2. Insert a needle into the membrane screw cap and pass the glass capillary through the needle.

2. Screw the cap onto a reservoir of the desired size and then remove the needle while holding the capillary in place, as shown in Fig.3. Repeat steps 1 & 2 for every additional glass capillary connection required.

: Figure 3. Pulling the needle out but keeping the capillary in place fixed by the membrane.

3. Insert the input pressure needle to appropriate depth (see next section for details) and connect to the Tygon tubing to the needle using the Luer-lock to tubing adapters, as shown in Fig.4. Repeat this step for every additional Tygon tubing connection required.

: Figure 4. Reservoir with a needle connection interfaced to a glass capillary (typically connected to the microfluidic chip) and Tygon tubing with a Luer-lock adaptor (typically connected to the pressure source) which can be used to control the flow from the reservoir to the chip.

4. The reservoir is completed. Unscrew the cap from the container and fill it with your reagent and screw the cap back on.

5. (Optional step) Instead of using a glass capillary, a steel tube connected to Tygon microbore tubing can be used. We used 0.0250″ OD steel tubing, inserted into 0.0200″ ID Tygon tubing and then forced the pin through the septum, as seen in Fig. 5. Multiple steel-Tygon tubing connections can be connected to one septum as shown in Fig. S2.

: Figure 5. Steel tubing and Tygon microbore tubing connected to a reservoir.

6. Connect the 1/16″ Tygon tubing to the pressure source and the capillary (or the microbore Tygon tubing) to the chip. Make sure to create slightly smaller sized holes for the input connections on the chip to create a tight seal. When using multiple reservoirs, they can be placed into microcentrifuge tube racks for easy handling (Microtube Storage Racks), as shown in Fig. 6.

: Figure 6. A picture of a chip connected to three reservoirs placed on a storage rack, each with an independent input line and multiple outputs.

7. Everything is now ready to use!

What else should I know?

This setup is simple, easy to build and can be done in less than a minute. Depending on the cap, the membrane/septum provides an airtight seal, but after inserting and removing needles more than a few times, especially with larger needles, leakage becomes apparent. Since the system is pressure controlled, a small leakage rate relative to the supply flow rate can be tolerated. Only a limited number of connections, up to 10 in our experience, can be connected to a single cap or else leakage becomes excessive. The reservoirs work well up to 345 kPa, and for higher pressure applications solid caps can be used as described in the supplementary material.

It is important to maintain the needle connected to the pressure source above the liquid surface in order to prevent the formation of bubbles. Be careful of the sharp points and remember to dispose of the needles properly.

Supplementary materials

Figure S1. The different available options (excluding glass capillaries) in our laboratory of microcentrifuge tubes sizes, membrane caps, steel tubing, and input/output connections.

: Figure S2. Multiple tubing outputs through multiple inserted steel tubing.

The presented setup above was not tested at pressures higher than 345 kPa, and at pressures above 140 kPa, there was minimal gas leakage. However, we have developed another setup that works without any leakage, even at 345 kPa, but requires several minutes to make and a precision micro-drill. This setup does not use a membrane cap, but a conventional screw cap without a membrane. Using suitable micro-sized drill bits, different sized holes can be made in the micro tube or the cap. After inserting the output capillaries or steel tubing and the input syringe needle, Instant Krazy glue [10] is applied to affix the inputs and outputs at the precise place. Dual melt glue [11] is then applied to seal the terminals from air/liquid leaks. The final setup is shown in Fig. S3.

We mention here that this setup is permanent and should be robust at high pressure applications. At the same time, it requires a precision micro drill, more time to make, and it is less flexible in the sense that it is harder to modify the interface.

: Figure S3. High pressure off-chip containers with one Luer-Lock input and one glass capillary output at the bottom.

References

[1] S. Li and S. Chen, IEEE Trans. Adv. Packag., 2003, 26, 242-247.
[2] R. Lo and E. Meng, Sens. Actuators, B, 2008, 132, 531-539.
[3] T. Liu, E. V. Moiseeva and C. K. Harnett, Integrated reservoirs for PDMS microfluidic chips, Chips & Tips (Lab on a Chip), 22 April 2008.
[4] T. Das et al., Interfacing of microfluidic devices, Chips & Tips (Lab on a Chip), 27 February 2009.
[5] Sarstedt AG & Co.: http://www.sarstedt.com/
[6] BD: http://www.bd.com/
[7] McMaster-Carr Supply Company: http://www.mcmaster.com/
[8] Polymicro Technologies: http://www.polymicro.com/
[9] New England Small Tube: http://www.nesmalltube.com/
[10] Small Parts, Inc.: http://www.smallparts.com/
[11] Krazy Glue: http://www.krazyglue.com/
[12] The Stanley Works: http://www.stanleyworks.com/

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A remote syringe for cells, beads and particle injection in microfluidic channels

20 Aug 2009

Jean-Christophe Baret
Institut de Science et d’Ingénierie Supramoléculaires (ISIS),
Université de Strasbourg, CNRS UMR 7006, Strasbourg, France
jc.baret[at]unistra.fr

Why is this useful?

In suspension, cells, beads and particles tend to sediment at a speed given by the balance of viscous drag and gravity force. In classical biological experiments, the most straight-forward solution to keep cells in suspension is to shake large volumes. This type of system is relevant at high Reynolds number but is not very compatible with the classical systems used to inject small amounts of liquid in confined geometries, such as microfluidic systems.

Another common way to prevent sedimentation is to match the densities of the particles and the carrier fluid, or to increase the viscosity of the carrier fluid to increase sedimentation times. However, these solutions are not always suitable due to the biological or physical constraints of a microfluidic experiment. For example, increasing viscosities increases the pressure drop in microchannels, and additives for density matching are not always biocompatible.

Previously in the Chips and Tips section a system has been described for cell injection [1]. With a similar system, we observed that sedimentation along the tubing binding the syringe reservoir to the chip hinders good mixing of the cells. This tubing should therefore be as short as possible, which is sometimes incompatible with the large footprint of the syringe pumps or other equipment around the microfluidic chip. Interestingly, in another Chips and Tips section, a system for temperature control of syringe pumps has been described [2]. The idea there was to use a syringe pump to pump liquid in order to move the piston of another syringe placed in a temperature-controlled bath. This system still showed problems related to the length of the connection tubing to the chip.

Here we combine both systems [1,2] to inject cells, beads and particles efficiently into microfluidic devices using small amounts of liquids (typically ~1 mL). The cells are initially placed in a syringe close to the chip and are automatically shaken inside the syringe using a small motor and a magnet. This syringe is then remotely actuated by another syringe mounted on a syringe pump [2] which reduces the footprint of the injection system in the vicinity of the chip. The tubing length from the syringe containing the cells to the chip can then be as small as 5 cm, which reduces the residence time of cells in the tubing and therefore reduces sedimentation.

What do I need?

Syringes: BBraun Injekt (2 mL or 5 mL)
Syringe needles: Terumo NEOPLUS 23G
Tubing: Fisher Scientific PTFE tubing (i.d. 0.56 mm, o.d. 1.07 mm)
Small magnets: Supermagnete, e.g. S-03-01-N
Teflon magnets: Fisher Scientific, length 6 mm, diameter 3 mm
Motor: Radiospares, e.g. 20GN steel 50:1 gear DC motor, 175rpm 12V
Power supply: Radiospares, e.g. LASCAR – PSU 130
Small equipment: stands and support, steel wire

What do I do?

1) The first part of the procedure deals with preparing the syringes. This system is very close to the one presented in [1]. Cut the plungers of 2 syringes, and insert a magnet in one (Fig 1).

2) Bind them together with the metal wire (Fig 2).

: Figure 1

: Figure 2

3) Fill the syringe that does not contain the magnet with water and hold it vertical (Fig 3).

: Figure 3

4) Insert the needles in the tubing, fill a third syringe with water, place the needle in position, flush the tubing with water and connect the syringe with the free needle (Fig 4). Make sure no air bubbles are present in the water phase: they will increase the response time of the system to flow rate modifications.

: Figure 4

5) Invert the syringe to have the magnet in the upper part and fill this part with the liquid containing the cells, beads or particles (Fig 5) and connect a short needle with tubing (Fig 6).

: Figure 5

: Figure 6

6) Rotate the syringe and mount the motor with the magnets. Turn on the power supply (Fig 7). The speed of the motor is simply controlled by the voltage to guarantee gentle agitation of cells.

: Figure 7

7) The system is now ready to be connected to a chip on one hand and to a syringe pump on the other (see video below). More sophisticated systems can probably be adapted to allow a temperature control of the syringe.

References

[1] R. Cooper and L. Lee, Preventing suspension settling during injection, Chips & Tips (Lab on a Chip), 21 August 2007.
[2] L. Capretto, S. Mazzitelli and C. Nastruzzi, An easy temperature control system for syringe pumps, Chips & Tips (Lab on a Chip), 22 April 2008.

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How to prevent sagging during the bonding or lamination of chips with large aspect ratio chambers

24 Jul 2009

Jie Xu and Daniel Attinger
Laboratory for Microscale Transport Phenomena, Department of Mechanical Engineering, Columbia University, New York, NY 10027, USA

Why is this useful?

Assembling multiple layers by bonding or lamination is a simple way to manufacture complex multilayer microfluidic chips [1, 2]. However, bonding or lamination of chambers with large aspect ratio, i.e. wide and shallow, sometimes fails because of sagging. Figure 1 illustrates a sagging problem, which resulted in the top chamber wall being accidentally bonded to the bottom wall. Here, we describe a tip to prevent sagging by using regular cooking salt.

: Figure 1: Accidental bonding due to sagging

What do I need?

Cooking salt (grain size: normally 300 microns, can be further ground to less than 100 micron; melting temperature: 801 °C, good for lamination)
Precision tweezers
Stereomicroscope

What do I do?

1. Carefully pave the bottom of the chamber with ordinary salt as in Figure 2. Try to perform this action using fine tweezers under a stereomicroscope, if the chamber is too small.

: Figure 2

2. Plasma bond or laminate the top layer. Be careful during handling, so that the salt does not end up in your DRIE machine.

: Figure 3

3. After bonding is done, flush the microfluidic system with deionized water for several minutes to dissolve and remove salt particles. As figure 4 shows, the bonded chamber does not exhibit adhesion between the top and bottom wall.

: Figure 4: No accidental bonding

References

1. J. Xu and D. Attinger, Drop on demand in a microfluidic chip, J. Micromech. Microeng., 2008, 18, 065020.

2. P. J. Hung, P. J. Lee, P. Sabounchi, N. Aghdam, R. Lin, and L. P. Lee, A novel high aspect ratio microfluidic design to provide a stable and uniform microenvironment for cell growth in a high throughput mammalian cell culture array, Lab Chip, 2005, 5, 44-48.

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Quick assessment of the stability of flow generated by a syringe pump in a microfluidic device

16 Jul 2009

Rachel Green and Siva A. Vanapalli
Department of Chemical Engineering, Texas Tech University, Lubbock, TX, USA

Why is this useful?

Syringe pumps are often used in a variety of microfluidic applications because of their portability and the ease with which flow rates can be changed. The syringe pump characteristics (type, age and wear), compliance in the tubing, a mismatch between the size of syringe used and the flow rate desired can generate pulsations in microfluidlic flows. These pulsations could be undesirable for lab-on-a-chip applications in which steady fluid flows are needed. The method described here is a quick means to assess the degree of pulsations present in flows driven by syringe pumps. The basic principle relies on using a microfluidic comparator [1,2] to detect small pressure fluctuations in fluid flows.

What do I need?

Microfluidic device with design as shown in Figure 2.
Ring stand
Ring stand clamp
Syringe pump
Distilled water
Dyed distilled water (refill ink)
Syringe (30 or 60 mL)
Tubing
Stereomicroscope

What do I do?

1. Set up ring stand with a wide-mouth syringe in the ring stand clamp (Figure 1). This is your hydrostatic head. The distilled water will go in this syringe.

: Figure 1: Setup of the ring stand and syringe pump with the microfluidic device.

2. Place a syringe with the dyed distilled water into the syringe pump.

3. As shown in Figure 2, connect the syringe pump to the chip (labeled as flow rate Q). Connect the hydrostatic head, P, from the ring stand to the chip.

: Figure 2: Microfluidic chip integrated with the comparator.

The outlets are open to atmospheric pressure. The device consists of two identical channels connected downstream to form a comparator region. At one inlet we impose a constant hydrostatic pressure (P) to generate steady flow and in the other inlet we use a syringe pump to introduce fluid admixed with a dye at a flow rate, Q.

4. Start the syringe pump at the desired flow rate. Vary the height of the hydrostatic head until the two fluid flows meet at the symmetry line of the comparator (see the white line in Figure 3a).

5. Because the hydrodynamic resistances of the two channels are equal, if any pulsations are present in the syringe-pump driven flow then the dyed fluid will be displaced above or below the symmetry line (shown in white) in the comparator, as shown in Figure 3b and 3c.

: Figure 3: Images of the fluid-fluid interface in the comparator at different times.

(a) The flows are steady and balanced. (b) After a time interval of 2 seconds, the dyed fluid is below the symmetry line indicating the syringe pump flow rate is lower than the flow rate corresponding to the imposed hydrostatic pressure. (c) After 6.8 seconds, the dyed fluid flow is above the symmetry line, indicating the syringe pump flow rate is greater than the flow rate corresponding to the imposed hydrostatic pressure.

What else should I know?

The method described is a quick assessment of the pulsations generated by syringe pump driven flow. For a more in depth study, we recommend using a precision hydrostatic head rather than a ring stand. We built a stand that can hold a syringe on a precision linear translation stage (Edmund Optics, Part # NT56-796). The stage allows the hydrostatic head to be adjusted at 0.1 mm intervals. Using this hydrostatic head and a video camera, one can precisely determine the number of fluctuations per unit time for a specified pump, flow rate, syringe size and tubing.

References

1. M. Abkarian, M. Faivre and H. A. Stone, Proc. Natl. Acad. Sci., 2006, 103, 538-542
2. S. A. Vanapalli, D. van den Ende, M. H. G. Duits and F. Mugele, Appl. Phys. Lett., 2007, 90, 114109

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A method for rapid fabrication of microfluidic devices

30 Jun 2009

Rajan Kumar, Robin L. Smith and Michael G. Pappas
Genome Data Systems, Inc., Hamilton, NJ 08610, USA

This Tip describes a useful variation on Holmes and Goddard’s method for rapid fabrication of microfluidic devices.[1]

Why is this useful?

Often, researchers need simple microfluidic devices with uniform height fabricated at short notice. Researchers also need to form channels to enclose materials deposited on glass surfaces, such as microarrays. A number of rapid prototyping methods are not compatible with these requirements. Holmes and Goddard have described an easy method for such preparing microfluidic flow systems.[1] We present a modification of this method using 3M double-sided permanent tape and heat curing with an inexpensive heat press. The resulting devices have a uniform height of ~80 micrometers. We have used such devices for microparticle assays and fluorescence microarrays in our laboratory.

What do I need?

Glass slide or other bottom plate substrate, e.g. plastic
Top plate substrate
3M double-sided permanent tape 665 [2]
3M double-sided removable tape 667
HobbyLite® T-shirt transfer press (Hix Corp., Pittsburg, KS)
Cutting board, razor blade, tweezers

What do I do?

1. Turn on the HobbyLite press and allow it to reach a temperature of about 100°F.

2. Draw the outline of the bottom plate (glass slide) on the cutting board. Draw the channel outline within the bottom plate outline.

3. Attach the glass slide to the cutting board over the outline using a piece of removable, double-sided tape (Figure 1). This step prevents the glass slide from moving.

: Figure 1

4. Attach a piece of double sided permanent tape to the top of the glass slide, covering the channel outline and leaving enough space on all sides of the channel for bonding (Figure 2).

: Figure 2

5. Cut out the double-sided tape from the channel outline using a sharp razor blade (Figure 3).

: Figure 3

6. Lift one corner of the tape cut-out using the edge of the razor blade, then lift the tape cut-out with the tweezers (Figure 4).

: Figure 4

7. Place the top plate on the slide, carefully aligning ports with the channel. Press down to ensure adhesion.

8. Cut excess double-sided permanent tape from each end of cover plate and remove with tweezers (Figure 5).

: Figure 5

9. Gently twist and lift the glass slide off the cutting board (Figure 6).

: Figure 6

10. Place the assembly in the HobbyLite press and gently apply pressure for 5-10 minutes (Figure 7). Be careful not to apply too much pressure or glass materials may crack.

: Figure 7

11. Remove the assembly from the press and allow it to cool to room temperature before use (~15 minutes). A completed device chamber filled with dye is shown in Figure 8.

: Figure 8

Acknowledgements

This research is funded by an NIH grant GM078945 awarded to Genome Data Systems, Inc.

References

1. R. J. Holmes and N. J. Goddard, Rapid prototyping of microfluidics, Chips & Tips (Lab on a Chip), 15 February 2007.
2. 3M Scotch® Double-Coated Tape 665 Permanent.

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Simple and reversible bonding of glass to glass, quartz and sapphire wafers

14 May 2009

Javier Atencia and Laurie E. Locascio
Biochemical Science Division, National Institute of Standards and Technology

Why is this useful?

We present a simple method to bond glass wafers reversibly by recreating the bonding that occurs occasionally among glass slides coming from a new box. Typically, the bond is clear without fringes, indicating that the wafers are in close contact, and the bond is so strong that it is impossible to separate the slides with bare hands, though it is not permanent; the slides can be separated with a razor blade.

This type of bonding is called “optical contact” or “direct-bonding”[1] (Isaac Newton was the first to observe it) and has been used extensively in the past, primarily in the optical field.[2] It has also recently been proposed for microfluidic chips using two polished glass wafers.[3] Different protocols to induce optical bonding differ in the amount of labor and the bonding yield. The main advantages of this type of bonding include the following: the bond can be reversible; the method works with materials of different thermal expansion (it does not require heating); and the method does not require adhesive materials.

Here we introduce a simple, quick and reproducible realization of this type of bonding for microfluidic applications.

What do I need?

Two polished 3″ circular wafers (surface roughness <5Å ), one of them with etched patterns on one side (there are many protocols in the literature) and through holes to access the patterns. (For other wafer geometries see the final section of this Tip.)
A bath for piranha etching or RCA cleaning, and appropriate personal protective equipment.
Cassette for 3″ wafers and a spin-dryer for wafers (if one is not available, see the final section of this Tip), e.g. Semitool-101.
Two neodymium magnets.

What do I do?

1. Place the glass wafers into different cassette slits (as usual), and place the cassette into the bath for RCA cleaning or Piranha cleaning (typically for 10 min.), Fig. 1(a).

2. Remove the cassette from the cleaning bath and place it in a bath of deionized water (for approx. 10 min.).

3. Remove the cassette from the deionized water bath, take one of the wafers out of its slit and place it into the same slit as the other wafer, taking care that the side of the wafer with the etched channels faces the other wafer. The two glass wafers should attach to each other by capillary forces and have a thin layer of water in between, Fig. 1(b). Care should be taken not to touch the sides of the wafers that will be in contact.

4. Carefully place two strong magnets on either side of the wafers that are in contact, thus sandwiching the wafers, Fig. 1(c). The magnets should be centered on the wafers.

5. Run the automatic protocol for spin-drying the wafers. The centrifugal force removes the liquid layer between the wafers, bringing them into close contact.

6. Remove the bonded wafers from the spin-dryer and remove the magnets. The microfluidic device is ready for use. (If the device will be used several hours later it is best to leave the magnets on the device until use.)

: Figure 1. Preparation of glass wafers before spin-drying.

What else should I know?

Gas bubbles. Sometimes small gas bubbles may remain between the glass wafers after spinning. These bubbles can usually be removed by pressing on the glass wafers with your gloved fingers and driving them to the edges.

Reversible bond. If there are impurities such as small particulates between the wafers, the device will not bond well. In this case, a razor blade can be used to separate the wafers. Clean the wafers again and repeat the operation.

Bonding Strength. We tested this bonding method using glass microchannels (500 micrometers wide) with a lid made of glass (borofloat), and found that it withstood pressures of 207 kPa (30 PSI) consistently without rupture 24 h after spinning. We also tested a glass-sapphire and a glass-quartz bond and found that both consistently withstood pressures of 138 kPa (15 PSI).

Repeatability. We have used a device for months with this bonding without failure. We have also repeatedly separated the wafers with a razor blade, cleaned them, and bonded them again successfully for reuse.

Surface quality. For this method to be successful, it is critical that the surface roughness is less than 5Å. If the wafers are not flat at the edges, etching a “crown” around the edges while etching the microchannels ensures proper bonding, see Fig. 2.

: Figure 2. Example of etched crown to eliminate defects at the edges that prevent bonding.

Aspect ratio of etched features. Because there is no thermal bonding, high aspect ratio devices can be fabricated without collapse of the features. For example, we created a circular chamber 500 nm deep and 1.5 mm in diameter, without collapse.

Temperature. We have used these devices at room temperature only, and expect them to fail if higher temperatures are employed due to thermal stresses – particularly when the device is made of two different materials.
No spin-dryer system or different wafer geometries

If a spin-dryer is not available, or if the wafers are different from each other or are not circular (see examples in Fig. 3), the same protocol can be used without spinning. However, in this case one should wait at least 24 h before using the device to allow the layer of water between the wafers to dry.

: Figure 3. Bonding between flat glass pieces of non-circular geometries.

Acknowledgements

We are thankful to G. Cooksey for the photographs displayed in the figures.

Part of this work was performed at the NIST Center for Nanoscale Science and Technology Nanofab, which is partially sponsored by the NIST Office of Microelectronics.

References

[1] J. Haisma, N. Hattu, J. T. C. M. Pulles, E. Steding and J. C. G. Vervest, Appl. Opt., 2007, 46 , 6793-6803.

[2] V. Greco, F. Marchesini and G. Molesini, J. Opt. A: Pure Appl. Opt., 2001, 3, 85-88.

[3] Z. J. Jia, Q. Fang and Z. L. Fang, Anal. Chem., 2004, 76, 5597-5602.

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An inexpensive and durable epoxy mould for PDMS

22 Apr 2009

André Estévez-Torres, Ayako Yamada and Li Wang
Department of Chemistry, Ecole Normale Superieure, Paris, France.

Why is this useful?

To create microfluidic patterns on PDMS one usually pours PDMS on a mould composed of microfabricated resist features on top of a Si or glass wafer. However, these resist moulds are fragile and usually break after a few uses, making it necessary to perform microfabrication again. Transferring the microfabricated features to an epoxy mould conserves the resolution down to 1 µm[1] and allows: i) multiple and inexpensive copies of the mould, ii) hundreds of uses without significant ageing, iii) easy incorporation of macroreservoirs to the PDMS device without needing to punch them every time, and iv) isopropanol cleaning, which is not possible with AZ resists.

What do I need?

Figure 1. Materials

Resist mould
PDMS
Silicone cake mould (e.g. SF025 financiers) from Silikomart (Mellaredo di Pianiga, Italy)
Epoxy resin (type R123, Bisphenol A/F epoxy resin) and hardener (type R614) from Soloplast-Vosschemie (Saint Egreve, France) or EasyCast Clear Casting Epoxy from Castin’Craft, Environmental Technology[2]
Trichloromethylsilane
Plasma cleaner

How do I do it?

: Figure 2. A) PDMS masters on the bottom of silicone cake molds. B) Pouring epoxy over PDMS masters. C) Peeling the epoxy+PDMS brick off the cake mold. D) Resulting epoxy molds.

1. Pour PDMS on the resist mould and cure it as you usually do.
2. Peel off the PDMS layer from the resist mould. Punch the macroscopic reservoirs if you need them. This PDMS part will be called PDMS master.
3. Put the PDMS master (microfabricated features on top) on the bottom of the silikomart cake mould. You may clean it with 3M tape.
4. Prepare the epoxy mixture as recommended by the manufacturer and remove the bubbles by vacuum pumping or ultrasonication.
5. Pour the epoxy mixture on the cake mould containing the PDMS master until you cover it with 2-3 mm of epoxy. Be careful to avoid making bubbles. If you are moulding small features you probably need to remove bubbles by vacuum pumping.
6. Wait for 24h at room temperature.
7. Remove the epoxy+PDMS brick from the mould and then peel off the PDMS master from the epoxy using a scalpel and tweezers. It peels off very easily.
8. The epoxy mould may be a little soft at this stage. Bake it in an oven at 70°C for one day – it becomes softer – and let it harden at room temperature for another day.
9. Silanize the epoxy mould by keeping it in a closed petri dish with a trichloromethylsilane-saturated atmosphere for 5 min. The first time you use the epoxy mould to make a PDMS device you need to put it into a plasma cleaner before silanization.
10. Pour PDMS on the epoxy mould as you usually do with other moulds. Cure it in the oven following your favourite recipe and peel it off. The first peel off needs to be performed slowly and carefully. Subsequent peelings off are straightforward.

: Figure. 3. Optical microscopy image of A) PDMS master, B) epoxy mold and C) PDMS molded on epoxy. The images are 300 µm wide and the smallest feature is 10 µm.

References

[1] This type of technique, called replica moulding, has been reported to yield features down to 100-50 nm (Y. Xia et al., Replica moulding using polymeric materials: A practical step toward nanomanufacturing, Adv. Mater., 1997, 9, 147-149).
[2] http://www.eti-usa.com

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Chips and Tips

Archive for 2009

A simple and economical holder for casting PDMS chips

“Custom made” production of cheap Luer lock adapters for chip-to-syringe interfacing

A novel method for fabrication of reusable microfluidic interconnects

Plug-and-play reservoirs for microfluidics

A remote syringe for cells, beads and particle injection in microfluidic channels

How to prevent sagging during the bonding or lamination of chips with large aspect ratio chambers

Quick assessment of the stability of flow generated by a syringe pump in a microfluidic device

A method for rapid fabrication of microfluidic devices

Simple and reversible bonding of glass to glass, quartz and sapphire wafers

An inexpensive and durable epoxy mould for PDMS

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