Archive for the ‘Fabrication’ Category

A simple and inexpensive device to remove edge beads

Markus Ludwig and Seth Fraden

Martin A. Fisher School of Physics, Brandeis University, Waltham, Massachusetts 02454, United States

Why is this useful?


To achieve high resolution and high aspect ratio features in contact lithography it is necessary to have the photomask in direct contact with the photoresist during the exposure. This is impeded by the formation of edge beads on the edge of the wafer, which can measure multiples of the nominal resist thickness [1].

Dedicated spin coaters remove edge beads automatically just after spin coating. However, during soft baking, a new edge bead forms due to the coffee ring effect [2]. It is therefore recommended, especially for thicker resist films, to remove the edge bead not before, but after soft baking [3].

Edge bead removal is not critical on single layer devices. However, for multi level designs, edge bead removal can improve the feature resolution significantly. This is especially the case if the first layer of resist is thick and the second layer is thin.

This article presents the fabrication and use of a simple and inexpensive device to reliably remove edge beads by solvent spraying.

What do I need?


Materials:

  • 100 ml round media storage bottle GL45
  • Diba Labware bottle cap Q series GL45 2 x 1/4 -28 ports with valves (00945Q-2V)
  • acrylic sheet, 12” x 6”, 3/16” thick
  • polyethylene sheet, 1/16 “ thick
  • 3x AAA battery pack with on/off switch + batteries
  • Parker micro diaphragm pump T2-05 IC (3/32 ID)
  • 2x ¼-28 to barbed 3/32 ID fitting
  • silicone tubing  0.078” ID, 50 cm length
  • 2x flat head machine screw 4-40 3/8 + nuts
  • 2x socket head machine screw 6-32 3/8 + nuts + washers
  • stainless steel blunt needle with 27 gauge Luer polypropylene hub, 1/2″ length
  • cable tie 4”
  • polypropylene tube fitting, male Luer slip to barbed coupler, 3/32″ tube ID

Tools:

  • hot air gun (Aoyue 968a+)
  • laser cutter (40W/45W CO2 Hobby Laser by Full Spectrum Laser) or scroll saw, drill press and drill bits
  • soldering iron (Aoyue 968a+) and solder

Figure 1: Materials needed

What do I do?


To build the edge bead remover, first prepare all components as shown in figure 1.

Cut the acrylic sheet and the polypropylene sheet and drill the holes as indicated in the schematic1.pdf or schematic1.dwg (AutoCAD) file. Solder battery pack wires to the pump. Then follow the instructions in video 1.

1. The blunt needle has to be bent at a right-angle.

Figure 2: Assembled device

Edge bead removal (follow the instructions in video 2):

  1. Spin coat wafer, soft bake and cool down.
  2. Center the wafer on spin coater and spin at 700 rpm. The optional centering tool used in the video was 3D printed on a formlabs form one 3D printer. (Use centering tool.stl file to reprint)
  3. Turn on pump, open pump valve to pressurize bottle, position nozzle over edge bead.
  4. Open spray valve and dissolve edge bead
  5. Sweep outwards slowly and keep spraying for another 15 seconds.
  6. Turn off spray valve and ramp up to 2000 rpm for 10 sec.
  7. Turn off pump and close both valves. Developer may remain in the bottle and in the tubing.
  8. Wafer does not have to be baked again and can be exposed immediately.

Figure 2: SU8 coated 3” wafer before (left) and after edge bead removal (right).

Figure 3: Comparison of feature resolution

CAD drawing of a multi layer design

SU8 master fabricated with edge bead

SU8 master fabricated without edge bead

Video 1 – click on the picture to play:

Video 2 – click on the picture to play:

References


[1] ‪ Shaurya Prakash, Junghoon Yeom, Nanofluidics and Microfluidics: Systems and Applications ‪Micro and Nano Technologies‪, William Andrew, 2014

[2] Robert D. Deegan, Olgica Bakajin, Todd F. Dupont, Greb Huber, Sidney R. Nagel and Thomas A. Witten, Capillary flow as the cause of ring stains from dried liquid drops, Nature 389, 827-829 (23 October 1997), doi :10.1038/39827

[3] http://www.cns.fas.harvard.edu/facilities/docs/SOP031_r2_6_SU-8%20photolithography%20process.pdf

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A simple and low cost method to fabricate NOA microfluidic chips

Gabriele Pitingolo1, Raffaele Vecchione1,3 and Paolo A. Netti1,2,3

1 Center for Advanced Biomaterials for Healthcare, Istituto Italiano di tecnologia (IIT@CRIB), Largo Barsanti e Matteucci, 53, 80125, Naples, Italy.

2 Dipartimento di Ingegneria Chimica, dei Materiali e della Produzione Industriale D.I.C.MA.P.I, Università di  Napoli  Federico II, Naples 80125, Italy.

3 Centro di Ricerca Interdipartimentale sui Biomateriali (CRIB), Università di Napoli Federico II, p.le Tecchio 80, Naples, 80125, Italy

Why is this useful?


Microfluidic chips are often made of silicon or glass which presents the drawbacks of being relatively expensive, time consuming and has limitations to the geometries that can be realized. PMMA is an optimal solution to overcome these aspects but it presents a low chemical resistance to organic solvents and aggressive chemicals

Norland Optical Adhesive 60 (“NOA60″) is a clear, colorless, liquid photopolymer that cures when exposed to ultraviolet light1. Surface bonding can be activated with light therefore monolithic and transparent devices especially useful for optical elements can be realized. In particular, the use of NOA 60 eliminates premixing, drying, or heat curing operations common to other optical adhesive systems. Curing time is a matter of minutes and is dependent upon the thickness applied and the energy of ultraviolet light available. Dupont and colleagues have recently developed a NOA microfluidic channel via a photolithography multistep method that presents a long time process2.

Here, we demonstrate the possibility to micromachine already cured NOA substrates by micromilling that is much easier and cheaper than photolithographic techniques for fabrication of microchannels or microstructures in general.  In addition, by micromilling it is possible to easily drill and make open channels in NOA substrates if needed. Also, in the case of microstructures on the two layers to be bonded if one layer presents microstructures with feature sizes below 25 micron and one substrate with feature sizes above this value then it is possible to prepare one substrate by photolithographic techniques and the other substrate by micromilling with following bonding, saving time and money.

What do I need?


  • Fully cured PDMS mold
  • Norland Optical Adhesive 60
  • UV light (E-Series Ultraviolet Hand Lamps)
  • Micromachining machine
  • Oxygen plasma machine
  • Clamp

What do I do?


1. Pour liquid photopolymer NOA into a preformed PDMS mold covering the entire surface (figure 1A). After a few minutes to stabilize the liquid polymer put the PDMS mold under UV light for 30 minutes at a  365 nm wavelength (fig.1B).

2. After the curing time, NOA substrate is ready to use; to fabricate a microfluidic chip two NOA substrates are prepared. NOA substrates are replicated onto flat PDMS surfaces exploiting the flexibility of the PDMS mold as shown in figures 2A and 2B.

3. Take the NOA substrate and mill a channel and related inlet/outlet holes using a micromilling machine (Minitech CNC Mini-Mill) (fig. 3A-3B), the certified positioning accuracy of the three-axis are 12″ / 300mm in x-axis, 9″ / 228mm in y-axis, and 9″ / 228mm in z-axis. To minimize the experimental uncertainty, the NOA substrates preformed in point 2 are smoothed before milling.

4. Prepare the NOA channel (clean with water and dry with an absorbent cloth) and  treat the channel and top layer by exposing to oxygen plasma for 60s, at a pressure less than 0.1 Torr and with a plasma power of 20 W (fig 4A). Clamp the two substrates and put the clamped channel under UV-light for 1 h finalizes the bonding process (fig 4B).

5. Your well-bonded NOA microfluidic chip (Figure 5) is now ready to use.


References

  1. https://www.norlandprod.com/literature/60tds.pdf
  2. E. P. Dupont, R. Luisier and M. A. M. Gijs, Microelectronic Engineering, 2010, 87, 1253-1255.
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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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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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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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