Welcome to Chips & Tips

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

Do you

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

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

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

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


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

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The use of polymer filaments or metal wires to pattern arbitrary-shaped micro-sized electrodes for microfluidic applications

Bruno Rafael Becker1, 2, Tristan Sun1, Shengjie Zhai1 and Hui Zhao1

1. Department of Mechanical Engineering, University of Nevada, Las Vegas NV, US

2. Department of Mechanical Engineering, Universidade Tecnologica Federal do Parana, Brazil

Why is this useful?


Electric fields are widely used to manipulate particles or fluid in lab-on-a-chip systems, for example to separate or assemble particles and pump or mix liquids, due to their favorable scaling with miniaturization. In order to exploit the advantages of electric fields, electrodes have to be fabricated and integrated with lab-on-a-chip devices. Patterned electrodes can also serve as biosensing by measuring the electric current change due to biomolecular binding. Traditional lithography has been used for such electrode fabrication, but particularly for arbitrary shaped electrodes, it is time consuming, requires expensive equipment and needs to be conducted in a cleanroom environment.

Hence, here we developed a simple fabrication method using commercial polymer fishing lines or metal bonding wires as a mask that does not require sophisticated procedures, expensive specialized instruments, and can be done outside a cleanroom environment. The gap size between the electrodes can be controlled by the commercial polymer fishing lines or metal bonding wires (from 1 mm to 1 mm). Due to the flexibility of fishing lines or metal wires, electrodes with arbitrary shapes can be readily fabricated by manipulating flexible lines. The fabrication method is particularly useful for demonstration of proof-of-concept or quick prototyping in terms of searching for the optimal shape, or researchers who lack the access to cleanroom or expensive lithographic equipment.

What do I need?


  • Glass slides
  • 3M double-sided tape
  • Deionized water (DI water)
  • Isopropyl alcohol
  • Acetone
  • Ultrasonic cleaner
  • Plasma cleaner
  • Sputter coating machine
  • Commercial fishing lines or metal bonding wires

What do I do?


1. Use DI water to clean the glass slide in the ultrasonic cleaner for 240 seconds and then dry the slide with the pressurized air (Fig. 1).


Fig 1. Using the ultrasonic cleaner to clean the glass slide.

2. Repeat the step 1 with isopropyl alcohol.

3. Repeat the step 1 with acetone. Acetone not only dissolves remaining contaminants but also provides negative surface charges to the glass surface to prevent adhesion of colloidal particles.

4. Put the glass slide into the oxygen plasma cleaner for 2 minutes (Fig. 2).

Fig 2. Using the plasma cleaner to clean the glass slide.

5. Use the double-sided tape to fix the fishing line on the glass slide.

6. Cover the glass slide with a cut paper. The cut paper along with the fishing line serves as a mask. The double-sided tape also keeps the paper fixed on the glass slide (Fig. 3).

Fig 3. The fishing line along with the cut paper serves as the mask.


7. Place the glass slide covered with the mask in the sputter coating machine (Fig. 4).

Fig 4. Put the glass slide into the sputter coating machine.

8. Sputter coating for 50-60 seconds to get 50-60 nm thickness metal electrodes.

9. Remove the mask from the glass slide (Fig. 5).

Fig 5. Patterned electrodes with the fishing line ready for use.

What else should I know?


Copper wires with a diameter of 140 µm were also tested. But because of their rigidity, they could not be aligned securely with the glass surface; consequently, the gold was sputtered around the wire and contaminated the channel. A 25 µm gold bonding wire was tested to determine the applicability of the method to fabrication with even smaller feature sizes. Electrodes with a 25 µm gap size are successfully fabricated, showing the robustness of our method (Fig. 6).


Fig 6. Patterned electrodes with the 25 µm gold bonding wire.

In the end, we connected the electrodes with a 100 ohms resistor with the electrodes, applied a voltage, measured the current, and plotted the I-V curve (Fig. 7). The I-V curve shows that the resistance of the electrodes is around 10 ohms, demonstrating the effectiveness of our method in patterning electrodes.

Fig7. Measured currents as a function of the applied voltages.

Experiments with the fabricated gold electrodes


We used the fabricated electrodes to assemble colloidal particles into functional structures (Fig. 8). Due to the dipole-dipole interactions induced by the applied electric fields, 5 micrometer latex particles are assembled into ordered structures with an AC electric field of 100 KHz and 10 V, which can find applications in photonics or biosensing.


Fig.8 Assembly of 5 m colloidal particles using the electric field.

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Different strategies for the fabrication of cell culture chambers for live-cell imaging studies

David Caballero1,2, Josep Samitier1,2

1 Nanobioengineering group, Institute for Bioengineering of Catalonia (IBEC), Barcelona, Spain

2 Centro de Investigación Biomédica en Red de Bioingeniería, Biomateriales y Nanomedicina (CIBER-BBN), Zaragoza, Spain.

Why is this useful?


Long-term imaging of cells is typically performed using standard Petri dishes. Frequently, these ´chambers´ are not convenient when sample manipulation and treatment (e.g. functionalization, immunostaining…) is needed, both before and after the experiment1. To overcome this ´problem´, glass coverslips are used. They can be easily manipulated when following multistep protocols prior to cell deposition. For live-cell microscope imaging, the customized coverslips are secured into holders (chambers) containing the adequate cell culture medium2, 3. These holders are either supplied by the microscope manufacturers or fabricated in a mechanical workshop; this implies time for the design and money.

In this work we show two different strategies for the simple, fast and cheap fabrication of chambers for live-cell imaging, using materials and simple tools typically available in a bioengineering laboratory. These materials include Petri dishes, polydymethylsiloxane (PDMS), Falcon tube cap and glass coverslips. In the first strategy (A) we drill a hole in a Petri dish where a customized glass coverslip is adhered at the bottom using wax. In the second strategy (B), a PDMS frame is used to hold the coverslip inside a Petri dish. Depending on the user final application one strategy is recommended over the other (see below). All the material is biocompatible and simple to obtain. These two methods provide several advantages: (i) they are easy and cheap; (ii) the chambers can be fabricated in a short-time period; (iii) this approach avoids the purchase of commercially-available holders or ordering the fabrication to a mechanical workshop.

What do I need?



    Figure 1

Figure 1. Material needed for the fabrication of the home-made chambers.

(1) PDMS
(2) Glass coverslip #1, 25 mm in diameter
(3) Cap of a 15 mL Falcon (any brand)
(4) Syringe 5mL
(5) Polishing paper
(6) Wax
(7) P35 plastic Petri dish (any brand)
(8) Sharp Tweezers
(9)Glass Pasteur pipette

You will also need:

  • Hot plate
  • Bunsen burner (optional)
  • Soldering iron (or drill)
  • EtOH 70%
  • Oven
  • SYLGARD 184 PDMS and crosslinker agent (Dow Corning)


What do I do?


Figure 1 (above) shows all the material needed for the fabrication of the home-made chambers using both strategies A and B. The steps describing both strategies are detailed next.

STRATEGY A:

1. Make a circular hole of around 2 cm in diameter in the middle of the lower part of a P35 Petri dish using a pre-heated, sharp-tip soldering iron. Alternatively, a drill can be used. Ensure a perfect circular hole by using a 15 mL Falcon cap as a template (see Fig.2a-c).

Figure 2. Fabrication of the cell culture chamber using Strategy A. Drilling a hole on the lower part of a Petri. (a) First, draw a circumference of about 2 cm in diameter in the center of the lower part of a Petri dish. Use a 15 mL Falcon cap as template. (b-c) Next, a soldering iron is used to drill a hole. (d) Finally, the edge of the hole is polished using thin polishing paper.

2. Polish the hole using polishing paper (see Fig.2d). Check that the Petri is free of debris. Rinse the sample with etOH 70%. (Optional: Sonicate).

3. Use the tweezers to place and secure the glass coverslip on the back side of the holey dish with its customized side (e.g. functionalized) facing the inner part of the Petri (see Fig.3a-b).

Figure 3. Adhering the glass coverslip to the lower part of the drilled Petri dish using wax. (a) The drilled Petri dish and glass coverslip #1, 25 mm in diameter are (b) placed together and secured using the tweezers. (c) Next, the wax is melted using a heat plate. A small volume is absorbed by capillarity using a glass Pasteur pipette. (d) Following the edge defined by the coverslip and the dish, the chamber is sealed. This step is critical to ensure a good sealing. (e) Check that no open remaining points are left. (f) A Bunsen burner or equivalent can be used to melt the solidified wax inside the pipette.

4. Melt the wax using the hot plate and fill the Pasteur pipette with a small volume (see Fig.3c). NOTE: Capillarity will make the liquid wax to flow inside the pipette.

5. Gently, put in contact the tip of the pipette (filled with wax) with the coverslip. Follow the edge formed by the coverslip and the Petri (see Fig.3d). Cover it completely with wax until the entire contour is sealed (see Fig.3e). Refill the pipette if necessary. NOTE 1: The wax may solidify inside the pipette quickly. If so, melt it again using the Bunsen burner (or equivalent) (see Fig.3f). NOTE 2: Ensure that no empty spaces are left; cell medium will flow through them.

6. Culture the cells of interest (see Fig.4). Place the chamber inside the microscope and start the live-cell imaging experiment. NOTE: Manipulate gently the sample.

Figure 4. Finished chamber for live-cell imaging experiments. (a) Front view of the chamber filled with cell culture medium. (b) Back side view showing the wax-sealed region. No leakage is observed. The coverslip can be easily recovered after the experiment by pushing it down gently with the tweezers.

7. At the end of the experiment, the medium can be removed and the coverslip detached by pushing it down gently with the tweezers. Treat the sample as desired (e.g. immunostaining).

STRATEGY B:

1. Insert upside down the cap of a 15 mL Falcon in the middle of a P35 Petri dish (see Fig.5a).

Figure 5. Fabrication of the cell culture chamber using Strategy B. (a) A 15 mL Falcon cap is placed in the center of a P35 Petri dish. The empty region is filled with PDMS using a syringe. (b) The sample is degassed and cured. (c) The cap is removed and the PDMS frame released.

2. Fill the empty space with a syringe (or equivalent) with PDMS in a 10:1 ratio (pre-polymer:cross-linker). Degass and cure it at 65ºC for 4h (see Fig. 5b). NOTE 1: Holding the cap with adhesive tape will ensure that it remains in the center during curing. In this case, curing must be performed at RT overnight. NOTE 2: A very thin PDMS layer may appear after the removal of the cap. Remove it manually to ensure a through-hole in the PDMS. Alternatively, a weight can be applied on top of the cap.

3. Remove the cap using the tweezers and release the PDMS frame (see Fig. 5c).

4. Sterilize the PDMS frame. Rinse it with etOH 70% and UV-irradiate for 15 min.

5. Working in the cell culture room, deposit a drop (~50 uL) of culture medium in the center of a new P35 Petri dish (see Fig. 6a).

Figure 6. Finished chamber for live-cell imaging experiments. (a) A small drop of cell culture medium is deposited in the center of a new P35 Petri dish. (b) The customized coverslip is placed on top of it. (c) The PDMS frame is introduced inside the Petri and pushed down to hold the coverslip forming the chamber. Finally, the chamber is filled with cells.

6. Place the (customized) glass coverslip facing-up on top of the drop (see Fig. 6b). NOTE: This will ensure that no air bubbles are formed. Hold it with the PDMS frame.

7. Culture the cells of interest (see Fig. 6c). Place the sample inside the microscope and start the experiment.

8. At the end of the experiment, the medium can be removed and the coverslip released by removing the PDMS frame with the tweezers. Treat the sample as desired (e.g. immunostaining).

Figure 7. Microfabricated coverslips for live-cell imaging studies. (a) Glass coverslip covered with PDMS microstructures. (b) The modified coverslip can be used for the fabrication of chambers using both strategies. (c) Zoomed image of the microstructures (parallel grooves). The dimensions are: 1 um x 1 um x 1 cm (HxWxL); the separation between grooves is 1 um. Scale bar: 10 um. (d) NIH3T3 fibroblasts aligned parallel to the grooves. The arrow shows the direction of the structures. Scale bar: 50 um.


What else should I know?


By using these two approaches, chambers can be easily fabricated in the lab. Most importantly, this approach allows the manipulation of coverslips before and after the experiment with cells. If the coverslips are (bio)chemically modified (e.g. micropatterned with proteins of the extracellular matrix), manipulation must be performed carefully and fast to avoid sample degradation. Similarly, for cell guidance assays, coverslips can be easily modified with microfabricated structures to orient cell growth and motility (see Fig. 7), following the same steps described in this protocol.

For Strategy A, users must be aware of the melting temperature of wax (around 45ºC). This implies that the sealing may be fragile when performing the experiments at 37ºC and manipulation must be performed gently to avoid liquid leakage. Other materials and shapes, besides circular glass coverslips, could be used. This will depend on the experimental requirements of the user. For Strategy B, the PDMS frame and the coverslip could be used without the Petri in some applications. However, for live-cell imaging, a perfect fit between the chamber and the microscope stage is needed and the use of the Petri is therefore strongly recommended.

Finally, the user must consider the magnification needed for the experiment and the thickness of the sample on each strategy. Strategy A is recommended for high magnification microscopy (40X – 100X) and Strategy B for low magnification (4X – 20X). If needed, thinner coverslips (#0) could be used.


Acknowledgements

Dr. Daniel Riveline, Dr. Jordi Comelles (Laboratory of Cell Physics ISIS/IGBMC, Strasbourg, France) and David Izquierdo (Nanobioengineering group – IBEC, Barcelona, Spain) are acknowledged for technical help and discussions.

References

1.      Zanella F, Lorens JB, Link W. High content screening: seeing is believing. Trends Biotech 2010; 28:237-45.

2.      Caballero D, Voituriez R, Riveline D. Protrusion Fluctuations Direct Cell Motion. Biophysical Journal 2014; 107:34-42.

3.      Riveline D, Buguin A. Devices and methods for observing the cell division WO/2010/092116, 2009.

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Reservoir Poly(dimethylsiloxane) Cap Fabrication

Luis G. Rigat-Brugarolas1,2, Antoni Homs-Corbera1,2 and Josep Samitier1,2,3

1 Nanobioengineering group, Institute for Bioengineering of Catalonia (IBEC), Barcelona, Spain
2 Centro de Investigación Biomédica en Red de Bioingeniería, Biomateriales y Nanomedicina (CIBER-BBN), Zaragoza, Spain.
3 Department of Electronics, Barcelona University (UB), Martí I Franques, 1, Barcelona, 08028, Spain.

Why is this useful?


Microfluidic devices are often connected to external reservoirs in order to perform experiments with large samples or to generate some types of closed loop systems.1 When containing biological content, once sealed and fluidically connected, these reservoirs should allow proper gas exchange and facilitate their placement in a controlled environment such as an incubator.2 On the other hand, often the connections used for this kind of assays may also suffer from sample leakages.

In this work we present a simple and cheap way of facing these issues by introducing a fabrication methodology for a sealing cap made of poly(dimethylsiloxane) (PDMS) that can be used for any kind of reservoir or bottle. The cap can be customized in order to allow multiple tubing connections depending on the specific user needs. PDMS is easy to manipulate, it’s biocompatible and permits gas exchange.3 This solution, consisting on fabricating a flexible PDMS cap covering the reservoir, should become a versatile alternative to expensive or unreliable other strategies and could be used in several microfluidic applications.

What do I need?


  • PDMS and crosslinker agent.
  • Plastic or glass reservoir.
  • Scalpel.
  • Tubing.
  • Oven.
  • Harris Uni-Core punchers or any kind of punchers.
  • Petri dishes.
  • Plasma cleaner.

What do I do?


Figure 1 shows the basic utensils and products needed to fabricate the PDMS cap.


Fig 1: Basic utensils and products needed to fabricate the PDMS cap

The steps are detailed next:

  1. Place reservoir/bottle in an upside down position inside a Petri dish. Fill the plate with PDMS as shown in Figure 2. In addition, fill another empty Petri dish with a thin layer of PDMS.
  2. Place both plates/samples inside an oven in a 65-90 ºC temperature range (depending on the reservoir’s material) for about 1.5 hours (Figure 3).
  3. Once the samples have polymerized, they have to be taken from the dishes (Figure 4), and the one corresponding to the reservoir cap replica can be cut, if needed, with a scalpel, as shown in Figure 5.
  4. After cleaning both samples with ethanol, they have to be chemically modified in O2 plasma (1 minute at high frequency, Figure 6) and immediately pressed together to form a permanent bond. It is important that the flat piece is sealed to the correct side of the cap replica.
  5. Once the final piece is achieved (Figure 7 left), it is useful to use Harris Uni-Core punchers in order to pierce the sample with the diameter of the tubing that is going to be used. It is recommended to make the holes with a slighter smaller diameter than the outer diameter of the tubing to form a pressure seal between the tubing and the hole (Figure 7 right); this will prevent leakage of the sample or undesired particles entering to the reservoir.
  6. Finally it is possible to screw the PDMS cap into the reservoir (an example is shown in Figure 8).

Fig 2: Fill two Petri dishes with PDMS, one with the reservoir (upside down) and the other just has to be covered with a thin polymer layer

Fig 3: Dispose the dishes inside an oven for 1-2 hours at 65-90ºC

Fig 4: An example of two PDMS samples. The left one corresponds to the reservoir cap replica. The right one to the PDMS thin layer

Fig 5: PDMS cap replica

Fig 6: Dispose both samples in a plasma cleaner in order to chemically modify the PDMS surfaces and join the cap with the PDMS plane piece

Fig 7: In order to pierce the access holes, a puncher is needed; afterwards the tubing can be connected to the PDMS cap

Fig 8: Example of a PDMS cap screwed into a plastic reservoir


References

[1] Herricks T., Seydel KB., Turner G., Molyneux M., Heyderman TT., Rathod PK.  (2011) A microfluidic system to study cytoadhesion of Plasmodium falciparum infected erythrocytes to primary brain microvascularendothelial cells. Lab Chip, 11, 2994.

[2] Rochow N., Manan A., Wi W., Fusch G., Monkman S., Leung J., Chan E., Nagpal D., Predescu D., Brash J., Selvaganapathy PR., Fusch C. (2014) An integrated array of microfluidic oxygenators as a neonatal lung assist device: in vitro characterization and in vivo demostration. Artif Organs, Doi: 10.1111/aor.12269.

[3] Regehr KJ., Domenech M., Koepsel JT., Carver KC., Ellison-Zelski SJ., Murphy WL., Schuler L., Alarid ET., Beebe DJ. (2009) Biomedical implications of polydimethylsiloxane-based microfluidic cell culture. Lab Chip, 9, 2132–2139.

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Simple fabrication of three-dimensional ramped microstructures using SU-8 negative photoresist

Luis G. Rigat-Brugarolas1,2, Antoni Homs-Corbera1,2 and Josep Samitier1,2,3

1 Nanobioengineering group, Institute for Bioengineering of Catalonia (IBEC), Barcelona, Spain 2 Centro de Investigación Biomédica en Red de Bioingeniería, Biomateriales y Nanomedicina (CIBER-BBN), Zaragoza, Spain.

3 Department of Electronics, Barcelona University (UB), Martí I Franques, 1, Barcelona, 08028, Spain.

Why is this useful?


At present, normal photolithographic techniques constitutes binary image transfer methodologies, where the developed pattern consists of regions with or without photoresist depending whether the UV Light has been in contact with the sample or not during the exposure process.1 Complex 3D patterns construction is of increasing importance in the miniaturization of fluidic devices.2 In the following work we introduce a photoresist-based technique to produce three-dimensional ramped microstructures for lab-on-a-chip applications.

We present a new technique that can be used to form multilevel features in SU-8 or any other negative photoresist using a single photolitograpy step, thus minimizing stages in the fabrication process in a simple and cheap way. This method thereby allows using a normal photomask without needing to add a complementary grayscale pattern, enabling complex microchannel structures.

What do I need?


  • Common items and devices used in photolitographic processes (mask aligner, hot plates, chemical baths, negative photoresist and transparent substrate)
  • Step Variable Metallic Neutral Density Filters (Thorlabs, Inc., NJ, USA).

What do I do?


  1. Dispose the sample in the mask aligner with the SU-8 photoresist on the bottom side as depicted in Figure 1. This will force UV light to cross through the transparent substrate and to first polymerize those photoresist regions in contact with the substrate.
  2. Place the photomask in the aligner standard position, normal to the light beam.
  3. Select the filter (continuous, step, etc.) according to your needs (see an example in Figure 2).
  4. Place the filter in the position between the photomask and the UV light source, with the filter’s design in contact with the photomask (see Figure 3).
  5. Enter a correct UV exposure time; since another element is going to be added in the UV light trajectory, this value has to be adjusted (final result in Figure 4).

Fig 1: Scheme of disposal of the SU-8 photoresist in the mask aligner for achieving relief structures.

Fig 2: Example of rectangular step filter available at Thorlabs, Inc.

Fig 3: Step filter placed between the photomask and the UV light source.

Fig 4: Example of three-dimensional ramped structure constructed using SU-8. Relief characterization obtained with a profilometer. The white line represents the structure obtained after the development process (with an angle value of 30º), showing a slope from 0 µm to 12 µm (in this case, a rectangular step filter was used). The red line shows what it would look like the profile if no filter had been dispose between the photomask and the UV light source.

What else should I know?


As with any negative photoresist, grayscale exposure in conventional processes will lead to hardening the surface, removing the substrate if unattached during the development, in a methodology normally used to create cantilever structures. This is why it is important, when trying to create relief structures, to turn the sample and expose it from the glass substrate side leaving the SU-8 or any other negative photoresist on the bottom side.

Acknowledgments


We thank David Izquierdo and Juan Pablo Agusil for their technical help and for providing the material.


Reference

[1] S.D. Minteer. Microfluidic techniques: reviews and protocols. Humana Press, 2006. ISSN: 1064-3745.

[2] C. Chen, D. Hirdes, A. Folch. Gray-scale photolithography using microfluidic photomasks. PNAS, 2003. DOI:10.1073

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Simple alignment marks patterning for multilayered master fabrication

Luis G. Rigat-Brugarolas1,2, Antoni Homs-Corbera1,2 and Josep Samitier1,2,3

1 Nanobioengineering group, Institute for Bioengineering of Catalonia (IBEC), Barcelona, Spain

2 Centro de Investigación Biomédica en Red de Bioingeniería, Biomateriales y Nanomedicina (CIBER-BBN), Zaragoza, Spain.

3 Department of Electronics, Barcelona University (UB), Martí I Franques, 1, Barcelona, 08028, Spain.

Why is this useful?


Nowadays it is common to fabricate multi-layered microfluidic microdevices by means of photolithographic techniques to create sophisticated structures allowing novel functionalities.1,2 Without any doubt, one of the critical steps in this manufacturing process is the alignment of the different transparent layers to perform the final device.

Several approaches have been made and studied to obtain a correct structuring of the three-dimensional device like, for example, the use of gold deposition, by means of sputtering techniques, for drawing the alignment marks in the substrate,3 or using expensive mask aligners with integrated alignment protocols. Those methods are expensive and laborious. In this work we present a novel, simple and non-time-consuming methodology for drawing alignment marks using Ordyl SY330 negative photofilm, a photoresist that can be easily displayed in the substrate and, thanks to its green color, it can be readily seen in a standard microscope.

What do I need?


  • Common items and devices used in photolithographic processes (mask aligner, hot plates and transparent substrate)
  • Sheets of Ordyl SY330 negative photofilm.
  • Ordyl Developer, SU-8 Developer or acetone.
  • Photomask with the alignment marks details.
  • Hot laminator.

What do I do?


A scheme of the alignment marks’ fabrication process can be seen in Figure 1. The steps are as follows:

  1. Dispose the Ordyl photofim over the substrate (Figures 1A-B and 2).
  2. Introduce both the substrate and the Ordyl film in a hot laminator in order to attach it firmly (see Figure 3).
  3. For the exposure, use an acetate or chrome-on-glass photomask (an example can be seen in Figure 4) with the design of the alignment marks. Because Ordyl is a negative photoresist, the design should have the marks in transparent in order to polymerize them and draw them in the substrate (Figure 1C).
  4. After exposure to UV Light (Figures 1D-E), an Ordyl Developer is needed (or, failing this, SU-8 developer or acetone), for having the final marks drawn in the substrate (an example can be seen in Figure 5).

Fig 1: Scheme of the fabrication process of the Ordyl alignment marks.

Fig 2: Placement of Ordyl photofilm on a microscope slide.

Fig 3: The hot laminator is used to attach the Ordyl film to the substrate.

Fig 4: Photomask with the design of the alignment marks.

Fig 5: Example of Ordyl alignment marks on a glass substrate.


Reference

[1] Dongeun Huh, Hyun Jung Kim, Jacob P Fraser, Daniel E Shea, Mohammed Khan, Anthony Bahinski, Geraldine A Hamilton and Donald E Ingber. Microfabrication of human organs-on-chips. Nature protocols. 2013 Nov. 11, vol.8. Doi:10.1038/nprot.2013.137.

[2] Michael P Cuchiara, Alicia CB Allen, Theodore M Chen, Jordan S Miller, Jennifer L West. Multilayer microfluidic PEGDA hydrogels. Biomaterials. 2010 31 5491e5497

[3] Eugene JH Wee, Sakandar Rauf, Kevin MS Koo, Muhammad JA Shiddiky, and Matt Trau. µ-eLCR: A Microfabricated Device for Electrochemical Detection of DNA Base Changes in Breast Cancer Cell Lines. LabChip. 2013 Nov 21;13(22):4385-91. DOI: 10.1039/c3lc50528f.

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Rapid fabrication of in/outlets for PDMS microfluidic devices

Ali Hashmi, Jie Xu
Washington State University

Thomas Foster
University of Washington

Why is this useful?

Previously we presented a method for connecting inlets and outlets to an external source that involved tubing and needles [1]. However, the process involves the use of needles which could be a safety concern. The process is also somewhat time-consuming. We have now developed a more convenient and rapid method for fabricating inlets/outlets in a PDMS chip without the need for needles.

What do I need?


  • A puncher, for example, we use a Schmidt punch press (Syneo, LLC)
  • Connectors with barbs and corresponding tubings, for example, we use elbow tube fitting with classic series barbs for 1/16” (1.6 mm) ID tubing (Valueplastic.com)

What do I do?


1. When the PDMS device has been cured, punch inlets and outlets from the top of the device.

(a)

(b)

(c)

Figure 1: (a) Schmidt Press; (b) punching through holes at desired locations on the device; (c) device with a set of three punched holes.

2. After sealing the device, insert the connectors (“Elbow Tube Fitting with classic series Barbs, 1/16”, (1.6 mm) ID Tubing, White Nylon”) into the inlets and outlet.

3.Tubing can then be connected to the connectors at one end, and to a syringe pump at the other end.

What else should I know?


The diameter of the punched holes is specific to the nominal cutting edge diameter of the punch. The punch, connectors, and tubing can be any size as long as they correspond to each other so that the connection does not leak. The connectors and tubing can be ordered from Value Plastics, INC.

The height of the Schmidt press can be adjusted according to the thickness of the PDMS device to ensure a through hole.

The maximum pressure we have tried with this type of connection is about 240 kPa, beyond which other parts of the chip fail (such as the PDMS/glass).

bonding.


Reference

[1]  P. Li., W. Xue, and J. Xu, The fabrication of PDMS interconnecting interface assisted by tubing fixationLab Chip, Chips and Tips, 10 June 2011

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An easy and fast System for bonding UPCHURCH® NanoPorts to PMMA

Gabriele Pitingolo, Enza Torino and  Raffaele Vecchione
Center for Advanced Biomaterials for Healthcare, Istituto Italiano di tecnologia (IIT@CRIB), Largo Barsanti e Matteucci, 53, 80125  – Napoli  – Italy.

Why is this useful?


Common systems to connect microfluidics devices with classic fluidic equipment (such as syringe  or peristaltic pumps) are based on the use of commercial connectors which are not always compatible with the device material.

Upchurch (Oak Harbor, WA, USA) NanoPorts™ assemblies are the first commercially available products to provide consistent fluid connectors for microfluidic chips. These products bond easily to some chip surfaces such as glass and polydimethylsiloxane (PDMS) with the provided preformed adhesive rings. All NanoPort™ components are made of inert, biocompatible PEEK™ polymer (nuts and ports) and Perlast® perfluoroelastomer (ferrules and gaskets). However, many microfluidic devices are made of polymethylmethacrilate (PMMA) and in this case the preformed adhesive rings are not suitable.

Here, we demonstrate an easy and effective way to bond NanoPorts to PMMA microdevices. Our approach is a hybrid system which glues the commercial nanoports with an alternative epoxy adhesive. Also remarkably this is a reusable system, in fact the Flat Bottom Port and the Flat Bottom Port Gasket may be removed and re bonded on another device as explained in the procedure.

What do I need?


  • Fully cured PMMA microchip with via holes to microchannels
  • UPCHURCH® SCIENTIFIC NanoPort Assemblies [1]
  • Loctite Super Attak Power Flex Gel (5g)
  • Binder Clip Medium 1-1/4in
  • FEP Tubing, 1/16’’ x 0.25 mm [2]
  • Scalpel and tweezers
  • Ethanol
  • Hammer

What do I do?


1. Prepare the PMMA surfaces (clean with water and dry with an absorbent cloth)  and NanoPort™ for bonding (Figure 1). The Inlet and Outlet holes must be of a diameter below the inner diameter of the Nanoport (around 2 mm) to guarantee no leakage at the Nanoport-PMMA interface.

2. Put a few drops of Loctite Super Attak Flex Power Gel (5g) on a surface, in our case we used a piece of PMMA (Figure 2). Take the UPCHURCH® SCIENTIFIC NanoPort, insert the gasket seal into the recess in the bottom of the port (Figure 3) and touch the port to the drop of Loctite Super Attak in order to deposit the right amount of glue on the bonding surface (Figure 4). Eliminate the excess glue with the aid of a scalpel and attach the flat bottom port gasket directly on the bottom of the port (Figure 5).

3. Take the complete Nanoport (flat bottom port and gasket) and touch the drop of Loctite super attack, eliminating the excess glue with the aid of a scalpel. Center and place the complete Nanoport on your final substrate surrounding the access hole (Figure 6).

4. Clamp the port to the substrate (Figure 7) for 3 hours.

5. Your well-bonded NanoPort interconnect to PMMA (Figure 8) is now ready to use.

6. It is possible to remove the Nanoport from the PMMA surface. Use ethanol to weaken the epoxy (Figure 9) and after 30 minutes punch with a hammer to separate the NanoPort™ from the PMMA device.


References

[1] http://www.upchurch.com/PDF/I-Cards/N4.PDF

[2] http://www.idex-hs.com/product-families/5/Tubing-Upchurch-Scientific-Ismatec.aspx

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Periodic degassing of PDMS to create a perfect bubble-free sample

Jonathan C. Chen, Shengjie Zhai and Hui Zhao
Department of Mechanical Engineering, University of Nevada, Las Vegas NV, USA

Why is this useful?

The process of mixing the base and curing agent of PDMS often leads to air bubbles within the prepolymer due to its chemical reaction. The presence of air bubbles significantly decreases the strength and diaphaneity of the PDMS chip. Therefore, removing bubbles from PDMS becomes necessary and important.  The traditional degasing method is at least 2 hours for a 10g PDMS mixed solution, which appears too long. A way to shorten the degasing time is in need.

Here, we develop a simple and robust method to speed up the degasing process. By periodically stopping the pump and pulling out the hose, we can remove the bubbles by forcing them to burst since the bubble cannot withstand the dramatic change in pressure, considering the large difference between the low pressure inside the vacuum and the higher atmospheric pressure outside. Using this process, we can speed up the process by around 40 minutes (10-15 gram PDMS solution) and fabricate a smooth and bubble-free PDMS sample for any purpose, especially for optical applications. In practice, this time reduction may depend on the vacuum itself and the volume of PDMS solution.

What do I need?


  • SYLGARD 184 silicone elastomer base & curing agent (Dow Corning)
  • 1 Disposable polystyrene weighing dish (LxWxH 86mm x 86mm x 25mm, white)
  • Gast Doap 704aa compressor vacuum pump 18 HP 115 Vac
  • Bel-Art vacuum chamber and plate (Interior volume 0.21 cu. ft.)
  • IKA Ceramag Midi magnetic stirrer ceramic hot plate (50-1200RPM)
  • 1 magnetic stirring rod octahedral 1” x 5/16”

What do I do?


  1. Weigh the PDMS base and curing agent (10:1) in the weighing dish.
  2. Mix the base and curing agent together mechanically with a magnetic stirrer. (About 1000RPM to 1200 RPM)
  3. Move the dish to assure that the stir bar is around all sides of the weighing dish for proper mixture for about 10-20 minutes. (Depending on amount of PDMS used)
  4. Place the mixed sample into the vacuum chamber, turn on the pump, and leave for about 10 minutes.
  5. After the 10 minutes, there should be a significant amount of air bubbles appearing on the surface. Turn off the pump, quickly pull the vacuum chamber valve out, and let outside air in. Such action changes the pressure of the vacuum chamber, eliminates most of the big surface bubbles, and pulls out the small bubbles in the solution.
  6. Place the hose back on and turn on the pump again.
  7. Repeat step 5 and step 6 until there are no more bubbles on the surface and in the solution.
  8. Cast the treated PDMS over the desired mold, e.g. a patterned wafer.
  9. Cure at 65°C for 1.5 hours.

What else should I know?


If the PDMS is used as a mold and placed in a petri dish with a microscopic glass slide, air bubbles will be significantly harder to remove due to bubbles trapped under the slide. This generally requires longer time within the vacuum and the occasional displacement of the slide to release any trapped air bubbles.

Another thing to note is that when casting the treated PDMS onto the desired mold; make sure not to pour out the mixture too fast. The slower the mixture is poured in, the less likely there will be air bubbles created during the transfer. If the sample generates air bubbles during the casting step, placing the product in the vacuum again for another 10 minutes will eliminate any unwanted bubbles.

Fig 1

Figure 1: Measure out a 10:1 ration of base to cure.

Fig 2

Figure 2: Mechanically stir the mixture.

Fig 3

Figure 3: Turn off the pump and use the difference in pressure to eliminate the bubbles.

Fig 4

Figure 4: Pour the mixture into the desired mold slowly.

Fig 5

Figure 5: Cure at 65°C for 1.5 hours.

Fig 6

Figure 6: The fabricated PDMS sample without bubbles.

Fig 7

Figure 7: The PDMS sample with a microscopic slide placed in.


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Microchannels and chambers using one step fabrication technique

Vivek Kamat, KM Paknikar and Dhananjay Bodas*
Centre for Nanobioscience, Agharkar Research Institute, GG Agarkar road, Pune 411 004
E-mail: dsbodas[at]aripune.org; Tel: +91-20-25653680      

Why is this useful?       


At present many techniques are employed for fabricating channels and chambers, most of them using photolithography and soft lithography [1]. The fabrication of a circular channel and chamber in a monolithic design is challenging, which can be achieved using copper wires of varying diameters (from 20 µm). This simplistic process also eliminates usage of expensive equipment, can be performed in a normal laboratory environment (doesn’t require clean room facilities) and high fidelity structures could be obtained.      

Fabrication of chambers can be achieved in a simple, fast and novel approach by utilizing agarose gel. Agarose gel is an important component used in molecular biology experiments. Agarose powder is mixed with water and is boiled, after cooling the liquid polymerizes to form a gel. This gel can be utilized to mold the desired chamber (variable size and shape) which can be utilized for making chambers on chip.      

What do I need?      


  1. PDMS (1 part curing agent and 10 part of base)
  2. Agarose (1% in distilled water)
  3. Copper wires of desired diameter.
  4. Square box 5 x 5cm which serves as chip caster.
  5. Used syringes (φ 4 mm in the present case)  

What do I do?      


  1. PDMS is prepared by mixing 1:10 proportion (curing agent to base) and degassing for 30 min in a vacuum dessicator [2, 3].
  2. 1% agarose powder is mixed in distilled water and boiled in microwave for 1 min until a clear solution is obtained. Decreasing the amount of agarose will result in softer gel
  3. Cut the tip off of a 4 mm diameter 1 ml syringe. Pour in the agarose solution.
  4. Allow the solution to cool inside the syringe and push the plunger to obtain gel in cylindrical form (see Fig 1). This cylinder so obtained can be cut into desired heights as per design requirement. In our case we have used 5 mm high cylinder for fabrication of the chamber (see Fig 2).
  5. Micro dimensional copper wire is inserted through the cylinder (see Fig 3) and the whole assembly is placed in a box for molding PDMS (see Fig4) and cured at 70°C for 3 h in a convection oven.
  6. After curing, place the chip in IPA for 5 min for removing the copper wire. Agarose gel can be removed by placing the chip in boiling water for 10 min. or by passing hot water using the microchannel. Repeat the process until agarose is washed completely without any traces.
  7. Thus, what we have achieved is a microchannel and chamber connected together fabricated in a single step (see Figs 5 and 6). This monolith design could be extended for multiple applications such as mixing, as a reaction chamber for carrying nanoparticle synthesis, cell lysis, DNA amplification etc. [3]

  

Fig1: After cooling push the plunger to get a cylindrical agarose gel

Fig 1: After cooling push the plunger to get a cylindrical agarose gel

Fig2: Cut desired height to get small cylindrical gels

Fig 2: Cut desired height to get small cylindrical gels

Fig3: Insert copper wire of desired diameter through the gel

Fig 3: Insert copper wire of desired diameter through the gel

Fig4: Place in a caster box, add PDMS and allow for curing 70°C for 3 h

Fig 4: Place in a caster box, add PDMS and allow for curing 70°C for 3 h

Fig5: Top view of the fabricated chip

Fig 5: Top view of the fabricated chip

Fig6: Fluid inside a monolithically fabricated microchannel and a chamber

Fig 6: Fluid inside a monolithically fabricated microchannel and a chamber

References  


1. SKY. Tang and GM. Whitesides, Basic microfluidic and soft lithographic techniques in Optofluidics: Fundamentals, Devices and Applications, McGraw-Hill Professional, 2010.
2. J Friend and L Yeo, Biomicrofluidics, 2010, 4(2), 26502. DOI 10.1063/1.3259624.
3. S Agrawal, A Morarka, D Bodas and KM Paknikar, Appl Biochem Biotechnol. 2012, 167(6), 1668-77. DOI: 10.1007/s12010-012-9597-8.

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