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 these or similar problems?

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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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 Xua
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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Simple visualization of a microfluidic acoustic pump’s sound path

R. Rambacha, C. Schecka, V. Skowroneka, L. Schmida and T. Frankea,b,*
aMathematisch-Naturwissenschaftliche Fakultät, Lehrstuhl für Experimentalphysik I, Soft matter and Biological physics, Universität Augsburg, Germany
b Department of Physics and School of Engineering and Applied Science, Harvard University, USA.
*E-mail: tfranke[at]seas.harvard.edu; Thomas.Franke[at]physik.uni-augsburg.de

Why is this useful?


Microfluidic PDMS-chips are widely used in many labs. Recently, the use of acoustics in combination with PDMS devices has attracted much attention in the field because it is simple to use and allows for unique control of minute amounts of fluid, cells and particles on a microfluidic chip. This trend is also reflected by a series of tutorial papers1 and front covers2,3 in Lab on a Chip that are dedicated to this topic.

The core component of these chips is a versatile interdigital transducer (IDT), consisting of a pair of interlocked electrodes on a piezoelectric substrate. It can be tailored to meet the demands of a specific application, e.g. on microfluidic chips the IDT is often used as an acoustic pump. The overall shape of the electrode arrangement determines the fluid actuating sound-path on the chip and makes the difference between specific IDTs. For better aligning the IDT with other fluidic components, such as a PDMS channel, testing the functionality of an IDT and probing completely new IDT-designs, it is often necessary to visualize IDT’s acoustic path or the focal point of a focused IDT. However this visualization still remains elusive and requires expensive equipment such an AFM, SEM or a vibrometer. These methods are not available in every lab and very time-consuming. We show a simple and quick way to visualize the IDT’s sound path. The only components needed are isopropanol and a microscope, which are both available in almost every microfluidic lab.

What do I need?


  • Sample microfluidic chip with IDT and frequency generator
  • Isopropanol
  • Microscope

What do I do?


1. Place the chip with the IDT on the microscope.

2. Put a few drops of isopropanol onto the chip to wet its surface.

3. Turn on the IDT and observe with microscope.

4. Then, knowing the position of the acoustic wave path, place the PDMS onto the chip.

5. Because there is still alcohol on the chip, firm bonding of the plasma treated PDMS to the chip is delayed and there is still some time (10min or so) to align the PDMS precisely under the microscope. When the alcohol has evaporated the PDMS eventually bonds firmly to the chip and is ready to use.

Covering the chip with an isopropanol film before turning on the IDT.

Covering the chip with an isopropanol film before turning on the IDT.

Optical micrograph of the chip with a straight IDT. An interference pattern of the SAW can be seen.

Optical micrograph of the chip with a straight IDT. An interference pattern of the SAW can be seen.

Focused tapered IDT excited with two different frequencies (left: 120MHz, right: 240MHz). The acoustic path shifts with higher frequency towards the center of the IDT.

Focused tapered IDT excited with two different frequencies (left: 120MHz, right: 240MHz). The acoustic path shifts with higher frequency towards the center of the IDT.

Two opposed tapered IDTs at (from top to down) 161MHz, 163MHz and 165MHz.

Two opposed tapered IDTs at (from top to down) 161MHz, 163MHz and 165MHz.

Visualization of a focused tapered IDT’s acoustic paths at 90MHz.

Visualization of a focused tapered IDT’s acoustic paths at 90MHz.

Demonstrating the visualization of the focused IDT’s acoustic path. Depending on the material’s anisotropy the focal point is shifted towards or away from the IDT.

Demonstrating the visualization of the focused IDT’s acoustic path. Depending on the material’s anisotropy the focal point is shifted towards or away from the IDT.

What else should I know?


  • With other fluids such as water, glycerin and ethanol, the visualization effect was less pronounced. The effect was most evident with isopropanol.
  • By knowing the applied frequency and the distance between the electrodes of the tapered IDT at the excited spot (it is equal to the wavelength), this is also a very simple and quick method to calculate the material’s surface speed of sound with an error below 5%.
  • One can determine the material’s anisotropy by measuring the difference between experimental and geometrical distance of the focal points of a focused IDT4.

References


[1] H. Bruus, J. Dual, J. Hawkes, M. Hill, T. Laurell, J. Nilsson, S. Radel, S. Sadhal and M. Wiklund. Lab on a Chip, 2011, 11, 3579
[2] J. Shi, S. Yazdi, S. Lin, X. Ding, I. Chiang, K. Sharp and T. Huang. Lab on a Chip, 2011, 11, 2319
[3] L. Schmid and T. Franke. Lab on a Chip, 2013, 13, 1691
[4] J.B. Gree, G.S. Kino and B.T. Khuri-Yakub. IEEE 1980 Ultrasonnics Symposium, 69

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A simple trick to open up clogged microfluidic chip

Stefania Mazzitelli and Claudio Nastruzzi,
Department of Life Sciences and Biotechnology, University of Ferrara, Ferrara, Italy
Email: mzzsfn[at]unife.it

Why is this useful?


An event any microfluidic researcher never wants to occur is channel clogging. Unfortunately this drawback is not unusual, especially when working with narrow channel microchips and polymeric solutions or cell suspensions. On many occasions, microchips perfectly fitting the experiment appear to be irremediably lost and on the way to the rubbish bin.

Presented in this tip is a very simple and inexpensive solution that can revitalize a clogged microfluidic chip. We have tested this simple protocol both on glass or PDMS microchips with positive results, moreover we solved severe clogging (as evidenced by optical microscopy) caused by cell clustering as well as by the frequent occurrence of polymer precipitation within the microchannels. Our method solves the issues reported above using an extremely simple approach and a microwave oven.

What do I need?


  • 21 gauge hypodermic needle [1]
  • Plastic tube (ETEF, FEP or PTFE) 1/16” OD, 0.75mm ID [2]
  • A 50 mL syringe [3]
  • A microwave oven

What do I do?


1–5 Using the 21 gauge hypodermic needle and the FEP (fluorinated ethylene-propylene) tube, build up the plastic port for microfluidic interfacing (the needle should fit perfectly in the FEP tube assuring a tight, fluid proof inlet, see panel 4).

6–9 Insert the FEP tube into a microfluidic port. Depending on the position of the clog (determined by microscopic observation) insert the tube in the port as far away from the clog as possible.

10–12 Pump, by hand held syringe, filtered distilled water into the chip, applying as much pressure as possible. In the clog is constituted of lipophilic materials or hydrophobic polymers, replace the water with ethanol, isopropanol, acetone or their mixtures with water.

13 Put the microfluidic chip into a standard kitchen microwave oven for 5 min at 500-700 watts.

NOTE: before treating the chip in microwave, REMOVE the metallic needle or the entire port.

14–15 Remove the microfluidic chip from the microwave oven, reinstall the port and flush, as soon as possible, water (or solvent) into the channels. In case one treatment in the microwave does not open up the channels, repeat the entire procedure.

16 Your beloved chip is now open and ready for a new set of experiments.

Pictures 1-8

Images 9-16

References


[1] http://www.picsolution.com/
[2] http://www.upchurch.com
[3] http://www.artsana.com

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

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

Why is this useful?


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

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

What do I need?


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

What do I do?


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

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

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

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

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

Figure 1

Fig.1: Charger Adaptor Assembly

Figure 2

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

Figure 3

Fig.3: Filling a device with pure CO2

References


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

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

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

Why is this useful? 


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

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

What do I need? 


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

  

What do I do? 


  

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

  

Figures 1-4

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

Figures 5-7

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

  

 What else should I know? 


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

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

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

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

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