Welcome to Chips & Tips

Welcome to Chips & Tips – a unique and regularly updated forum for scientists in the miniaturisation field from Lab on a ChipChips & 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 that Chips & Tips before April 2011 were originally published at www.rsc.org.

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Manual Razor Patterned Tape Based Prototyping for Droplet Microfluidics

Saifullah Loneab* I. W. Cheongb and S. T. Thoroddsena

aDivision of physical Sciences and Engineering, King Abdullah University of Science & Technology, (KAUST), Thuwal, 23955-6900, Saudi Arabia.
bInstitute of Advanced Energy Technology, Kyungpook National University, Daegu, South Korea,
Phone: +821053165673, Office: +82-53-950-7590, FAX: +82-53-950-6594. Email: saifullah.lone@gmail.com, inwoocheong@gmail.com, and sigurdur.thoroddsen@kaust.edu.sa

Why is it Useful?

The subject of droplet microfluidics has grown in importance among researchers in chemistry, physics and biology, hence it has found applications in drug delivery, encapsulation, single-cell analysis, pickering-emulsion and phase-separation. For generating monodisperse droplets, various methods have been employed in constructing microfluidic devices. Emulsions with a coefficient of variation ≤ 5% have been previously reported in T-junction, flow focusing, co-axial, as well as other types of microfluidic devices. Microdroplets with ≤100 µm size offer attractive applications in industry and biology.  Small channel-diameters attained by clean-room soft lithography is the most precise technique for fabricating microfluidic devices.1, 2 This technique is widely used to make master molds for PDMS-based devices.3 However, regarding the cost and complexity, it is difficult to install clean-room soft lithography in financially challenged countries and laboratories. Therefore, the cost and special clean-room training restricts its wide-spread application. To develop low cost robust technologies; inkjet printing, controlled numerical machining, xurography or razor-writing, printed circuit technology, print-and-peel (PAP) microfabrication and 3-D printing have been tested to fabricate microfluidic devices without clean-room technology.  However, creating droplets under 100 µm size ranges by non-cleanroom technologies is challenging and open for upgradation. Recently, a rapid prototyping technique for microfluidics has been reported by employing laser-patterned tape4 This technique relies on computer-controlled CO2 laser beam. This work was further simplified by manual razor patterned tape-based prototyping for patterning mammalian cells.5 Building on this prototyping concept, we extended the idea to produce monodisperse droplets under 100 µm size rages by overlapping the razor patterned tape strips (at right angles) on a flat glass surface. The production of monodisperse emulsion under 100 µm size ranges are greatly useful in pharmaceutical and cosmetic industries. Hence, our approach may well serve as one of the simplest approaches to fabricate droplet microfluidic generators.

What do I need?   

  1. One-sided adhesive tape (Temflex 1500 electrical, thickness 150 µm)
  2. Flat glass slides, such as a microscope slide
  3. 30cm stainless steel metal ruler 
  4. Sharp razor blade
  5. Uncured mixture of PDMS base and curing agent (10:1 w/w)
  6. Oxygen plasma
  7. Oven or hot plate
  8. A microfluidic PDMS puncher for drilling holes
  9. Deionized water (D.I. water) and 20 cSt and 10 cSt silicone oil

What do I do?

Figure 1 outlines the prototyping procedure. Prototyping begins by attaching adhesive tape on a flat glass substrate. With a sharp razor-blade, the tape is cut into fine parallel strips. The thickness of the tape (150 µm) determines the depth of the microchannel, but this can be increased by attaching multiple layers of tape on top of each other. Next the tape is removed from the regions outside the fine strips.

To construct a cross-junction, one strip of the tape is lifted and horizontally placed on top of another at an angle of 90ᴼ. The junction is pressed gently to ensure the strips are well attached. These adhering strips of tape serve as a master for PDMS-based replica casting.

A mixture of PDMS silicone elastomer base and a curing agent (in 10:1 ratio) is poured on top of the master within a plastic petri dish. The mixture is degassed under vacuum for 1 h and cured for 4 hrs at 65°C. Cured PDMS replica is then cut and peeled-off from the master. The master can be used repeatedly to fabricate multiple copies of the PDMS replica by following the afore-mentioned steps. Inlet and outlet holes are drilled through PDMS replica, which is then bonded on a glass substrate, after both replica and glass has been exposed to oxygen plasma.  Figure 1(g) shows the resulting PDMS-device for generating monodisperse water-in-oil (W/O) emulsion. The technique is easily extended to fabricate T-junction or double T-junction prototypes (Figure 1h and i).

Figure 1. (a) Manual Razor Patterned Tape Based Prototyping for Droplet Microfluidics (b) Strips of adhesive tape on flat glass-substrate cut by a sharp razor-blade and a ruler, (c) The PDMS-casting, by pouring a mixture of PDMS silicone elastomer base and curing agent on top of the master in a plastic container. (d) Cut and peeled-off replica after curing. (e) Final assembled cross-junction microfluidic PDMS device. (f) Microfluidic device connected to syringe pumps under an optical microscope. (g)  Video frame showing water-in-oil droplet formation in a flow-focusing prototype. Panels (h) and (i) show razor patterned tape-based T-junction and double T-junction prototypes, respectively.
Figure 2. (a) Image sequence from a video recorded at 10 kfps showing water droplet formation at a cross-junction.  Time between subsequent frames is 200 µs.  (b) Droplet size as a function of capillary number based on the viscosity and flow-rate of the continuous-phase (20 cSt silicone oil), for the channel in panels (c) for 300 µm wide channel above the horizontal line and (d) for 150 µm channel below the horizontal line with 10 cSt silicone oil.  (c) Images extracted from video, showing flow regimes and droplet sizes as a function of flow-rate, with 300 µm channel width and 150 µm channel depth. (d) Video frame from smaller square channel with 150 µm width and depth.

Figure 2(a), demonstrates the droplet formation at a cross junction in our tape-based microfluidic device, with a channel width and depth of 300 µm and 150 µm respectively. In Figure 2(c), we kept the flow rate of aqueous phase fixed at 25 µl/min, while systematically increasing the flow-rate of outer continuous oil-phase (20 cSt silicone oil).  As the outer flow-rate is increased, the regime is found to shift from dripping at lower flow-rate to jetting at higher flow-rate (Figure 2(c)).  For lowest flow-rate, the aqueous-phase breaks into elongated plugs, while at higher flow-rates regular drops are pinched off.  Various factors affect the size of the droplets, but it is primarily determined by competition between viscous stress in the continuous phase Fv~ µU/d which tends to rip off the drop and the interfacial tension Fs ~ s/d which try to keep the drop attached. Here µ is the dynamic viscosity of the outer phase and U is its velocity; while s  is the interfacial tension between the water and the oil, s=0.040 N/m.  For small channels, the characteristic length-scale d is the same for the two forces and it therefore drops out of the balance and when Fv~ Fs then the non-dimensional capillary number Ca=µU/s characterizes their relative strength.  Figure 2b shows the droplet-size as a function of Ca.  When the flow-rate of oil-phase reaches 65 µl/min, the size of the droplets reaches ~100 µm and the droplet breakup occurs at a large distance from the cross-junction. Figure 2(d) shows drop formation with a channel width of 150 µm and a channel depth of 150 µm. In this case, the droplet size reaches down to ~73 µm, when the oil-phase (10 cSt) is flowing at 25 µl/min and aqueous-phase at 10 µl/min.

 

Acknowledgement: This work was jointly funded by King Abdullah university of Science & Technology (KAUST), Thuwal, Saudi Arabia , and the Ministry of Trade, Industry and Energy, Korea (Grants No. 10067082 and 10070241).

 

Reference

[1] Qin, D.; Xia, Y.; Whitesides, G. M. Rapid prototyping of complex structures with feature sizes larger than 20 μm. Adv. Mater. 1996, 8, 917-919.

[2] Xia, Y.; Whitesides, G. M. Soft Lithography. Angew. Chem. Int. Ed. 1998, 37 550- 575.

[3] Duffy, D. C.; McDonald, J. C.; Schueller, O. J.; Whitesides, G. M. Rapid Prototyping of Microfluidic Systems in Poly (dimethyl siloxane). Anal. Chem., 199870, 4974–4984.

[4] Luo, L. W.; Teo, C. Y T.; Ong, W. L.; Tang, K. C.; Cheow, L. F.; Yobas, L. Rapid prototyping of microfluidic systems using a laser-patterned tape J. Micromech. Microeng. 2007, 17 N107–N111

[5] Anil, B. S.; Ali, H.; Cheul, H. C.; and Raquel, P-C. Adhesive-tape soft lithography for patterning mammalian cells: application to wound-healing assays. BioTechniques, 2012, 53 315–318.

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Rapid Inoculation and Recovery of Microbes in a Microfluidic Device

Greiner, A.1,2, Tekwa, E.W.1,3, Gonzalez, A.1, Nguyen, D.4

1Department of Biology, McGill University, 1205 Dr. Penfield, Montreal, QC, H3A 1B1, Canada.
2Department of Ecology and Evolution, University of Toronto, 25 Willcocks Street, Toronto, ON, M5S 3B2, Canada
3Department of Ecology, Evolution, and Natural Resources, Rutgers University, 14 College Farm Road, New Brunswick, NJ, 08901, USA.
4Meakins Christie Laboratories, Research Institute of the McGill University Health Centre, and Department of Medicine, McGill University, 1001 Decarie Blvd, Montreal, QC, H4A 3J1, Canada.

 

Why is this useful?

Microfluidic devices are used for many different types of experiments across the medical, ecological and evolutionary disciplines (Park et al., 2003; Keymer et al., 2008; Connell et al., 2013; Hol & Dekker, 2014). For example, microfluidic devices for microbial experiments require inoculation into smaller chambers that simulate natural microbial environments such as porous soils (Or et al., 2007) and biological hosts (Folkesson et al., 2012). These devices often involve complicated pump setups and irreversible seals. We developed a technique that requires only common lab equipment and makes the device reusable while also allowing the microbes to grow undisturbed (based on Tekwa et al., 2015; Tekwa et al., in review). Here, we provide a detailed guide for the assembly and the previously undocumented non-destructive disassembly of polydimethylsiloxane (PDMS) experimental devices to recover microbes in situ, which can then be plated for relative counts and further molecular analyses of population changes. This is complemented by videos for each step.

Figure 1: Microfluidic device containing 14 habitats on an elastomer (PDMS) layer pressed onto a 60mm x 24mm glass cover slip. The habitats are 10 or 20µm in depth, range from 1400 µm to 2670 µm in diameter and take the shape of a ring or network of patches. This device is used to test the effects of habitat patchiness on microbe dynamics. Habitats were dyed blue for visualization. For more information see Tekwa et al. (2015).

 


What do I need?

  • Single-layer PDMS devices with habitats on one side
  • Pipette + sterile pipette tips
  • Sterile petri dishes (1/device)
  • Sterile tweezers
  • Inoculum
  • Filtered water
  • Kimwipes
  • Autoclavable plastic container
  • Ethanol
  • Tinfoil
  • Sterile 1μL inoculating loop (1/habitat)
  • Sterile eppendorfs
  • Phosphate-buffered saline (PBS)
  • Biological safety cabinet (BSC)

How do I do it?

  1. Clean the PDMS devices: The PDMS device should be pre-treated once with 0.01N HCl for one hour and plasma-treated to keep it hydrophilic and amenable to bonding to glass or plastic substrates (Cho et al., 2007; Tekwa et al., 2015). Fill autoclavable plastic container 1/3rd full of 70% ethanol and PDMS devices and then cover with tinfoil, let them sit in this for >30 minutes before carefully disposing of the ethanol down the sink. Fill and empty the container with water 10 times in order to rinse the devices. Lastly, fill the container with filtered water, seal with tin foil and autoclave in order to sterilize the devices.
  2. Inoculating the devices (perform in a Biological Safety Cabinet, BSC): Set the devices to dry in the BSC for 30 minutes. Place the devices with features facing up on the lid of a petri dish and place a small amount (i.e. 0.7 µL) of inoculum onto each (of 14 in sample device Fig. 1) habitat. The amount of liquid must be sufficient to fill the habitats, but not too much as to prevent bonding between PDMS and the cover glass/petri dish (Fig. 2). Using sterile tweezers, pick up the device and place face down into the centre of a petri dish or cover glass, sealing it to the surface by pushing on the back with gloved fingers repeatedly, using a kimwipe to wick excess liquid away from the side. Then surround, but not touch, the device with kimwipes soaked in filtered water (Fig. 3) to ensure that the device does not dry out in the incubator, before closing the petri dish. Place upright in incubator for the desired amount of time. The experiment can now proceed untouched for up to 24 hours (see Supplementary video).

Figure 2. Device with bacteria droplets, 1 droplet per habitat.

Figure 3. An ‘upright’ petri dish + kimwipes + device + coverslip ready to be incubated.

 

  1. Recovering from the devices (perform in a BSC): Open petri dish, carefully remove and discard kimwipes and then use sterile tweezers to gently unseal the device and place face up in the lid of the petri dish. By around 12 hours, the spaces between the habitats will be void of liquid from PDMS absorption, preventing microbes from being mixed across chambers during disassembly. Habitats that have dried out will appear white (Fig. 4) and cannot be used.  Dip sterile inoculating loop into an eppendorf with PBS, then use that loop to scrape one of the habitats (Fig. 5).  Dip that inoculating loop back into the eppendorf media again, which can then be grown overnight for further analyses such as plating for relative cell count (if there are different strains) and other molecular analyses. Repeat for the rest of the habitats that you are interested in, using new inoculating loops. The PDMS device can now be cleaned as in Step 1 and reused again.

Figure 4. View of an inoculated and incubated device, looking through the bottom of a petri dish.

 

Figure 5. Recovering bacteria from a habitat in the disassembled PDMS device.

 

What else should I know?

The recovery technique can be used to estimate relative proportions of different types of microbes (e.g. morph frequencies) which is useful when performing competition assays and evolutionary experiments. Unlike in Tekwa et al. (2015), this technique forgoes the use of a confocal microscope; assessment of the contents of the device is instead performed through direct microbe recovery and standard plating procedures.

 

Links to Videos

These videos go through the specific procedure that we used to perform experiments on competition and cooperation in Pseudomonas aeruginosa and may be useful in determining specific amounts of media, growth times, etc. that may be utilized for experiments with similar PDMS microfluidic devices.

Part 1 – Intro + Washing the Devices https://youtu.be/Bne3ZN3wU4Q

Part 2 – Inoculating the Devices https://youtu.be/p-UYIRreYyM

Part 3 – Recovering from the Devices https://youtu.be/YLnmGxqMYgE

Supplementary – Fluorescent Bacteria Experiment https://youtu.be/lgMtryS62pA

 

Acknowledgements

AGr was supported by an NSERC Undergraduate Student Research Award and by an NSERC Discovery Grant. EWT was supported by the Fonds Québécois de la Recherche sur la Nature et les Technologies and the Québec Centre for Biodiversity Science. AGo was supported by the Canada Research Chair program and NSERC Discovery grants. DN was supported by a CFI Leaders Opportunity Fund (25636), a Burroughs Wellcome Fund CAMS award (1006827.01) and a CIHR salary award.

 

References

Cho, H., Jönsson, H., Campbell, K., Melke, P., Williams, J. W., Jedynak, B., … & Levchenko, A. (2007). Self-organization in high-density bacterial colonies: efficient crowd control. PLoS biology, 5(11), e302.

Connell, J. L., Ritschdorff, E. T., Whiteley, M., & Shear, J. B. (2013). 3D printing of microscopic bacterial communities. Proceedings of the National Academy of Sciences, 110(46), 18380-18385.

Folkesson, A., Jelsbak, L., Yang, L., Johansen, H. K., Ciofu, O., Høiby, N., & Molin, S. (2012). Adaptation of Pseudomonas aeruginosa to the cystic fibrosis airway: an evolutionary perspective. Nature reviews. Microbiology, 10(12), 841.

Hol, F. J., & Dekker, C. (2014). Zooming in to see the bigger picture: Microfluidic and nanofabrication tools to study bacteria. Science, 346(6208), 1251821.

Keymer, J. E., Galajda, P., Lambert, G., Liao, D., & Austin, R. H. (2008). Computation of mutual fitness by competing bacteria. Proceedings of the National Academy of Sciences, 105(51), 20269-20273.

Or, D., Smets, B. F., Wraith, J. M., Dechesne, A., Friedman, S. P. (2007). Physical constraints affecting bacterial habitats and activity in unsaturated porous media – a review. Advances in Water Resources, 30(6), 1505-1527.

Park, S., Wolanin, P. M., Yuzbashyan, E. A., Silberzan, P., Stock, J. B., & Austin, R. H. (2003). Motion to form a quorum. Science, 301(5630), 188-188.

Tekwa, E. W., Nguyen, D., Juncker, D., Loreau, M., & Gonzalez, A. (2015). Patchiness in a microhabitat chip affects evolutionary dynamics of bacterial cooperation. Lab on a Chip, 15(18), 3723-3729.

Tekwa, E.W., Nguyen, D., Loreau, M., Gonzalez, A. Defector clustering is linked to cooperation in a pathogenic bacterium. In review.

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Rapid and easy fabrication of glass-bottom culture dishes for long-term live cell imaging

Ayako Yamada123, Jean-Louis Viovy123, Catherine Villard123 and Stéphanie Descroix123

1 Laboratoire Physico Chimie Curie, Institut Curie, PSL Research University, CNRS UMR168, Paris, France.

2 Sorbonne Universités, UMPC Univ. Paris 06, Paris, France

3 Institut Pierre-Gilles de Gennes, Paris, France

Email: ayako.yamada@curie.fr

 

Why is this useful?

Glass is a versatile surface for chemical treatments, and it still is by far the most used substrate for surface engineering (e.g. micropatterning, surface chemistry) or plasma-bonding of PDMS microfluidic devices. For cell culture on such substrates, glass-bottom culture dishes are desired to keep over the cells well defined medium volumes, and to protect the cells from contamination and medium evaporation. Moreover, they are optically better suited for microscopy observation than polystyrene dishes routinely used for cell culture. Although glass-bottom culture dishes are commercially available (e.g. Fluorodish from WPI), the presence of plastic walls limits the treatments that can be performed onto the glass bottom surface and those are more expensive ( 5 € per dish; φ 50 mm) than polystyrene dishes (e.g. φ 40 mm dish from TPP, 0.5 € per dish). In this Tip, we describe an easier way than a previous Tip1 to transform a polystyrene culture dish into a glass-bottom one, while preserving the possibility to apply to the glass any treatment before its assembly into a dish. Note that in this method, the body of the culture dish will be upside down and the lid is thus no longer lifted above the dish opening by lid stoppers. However, the gas exchange through the gap between the body and its lid seems to be enough to culture cells healthily in this dish. In summary, this Tip provides a low-cost and rapid solution for cell culture in a microfluidic device or on an engineered surface directly in a culture dish, suited for a long-term live cell imaging.

 


What do I need?      

  • φ 40 mm polystyrene tissue culture dish (e.g. TPP #93040)
  • φ 40 mm cover slide (e.g. Thermo Fisher Scientific #11757065, ∼0.2 € per slide)
  • Large screw driver (or a similar tool)
  • Uncured mixture of PDMS base and curing agent (10:1 w/w)
  • Oven or hotplate
  • Ferromagnetic metal plate (e.g. lid of a PDMS container, optional)
  • Cylindrical magnets (optional)

 


What do I do?

  1. Place a polystyrene culture dish upside down on a surface and hit a few times the center of the dish bottom with the grip of a large screw driver (Fig. 1a) until the dish bottom falls apart from the dish wall (Fig. 1b). The bottom should fall easily with the success rate around 9 over 10. Avoid breaking the dish wall by hitting the bottom too strongly.
  2. Spread uncured PDMS mixture on a flat substrate (e.g. a larger plastic Petri dish) and coat the edge of the dish (broken part up) with PDMS (Fig. 1c).
  3. Place the dish (broken part up) on a cover glass slide (Fig. 1d) and cure PDMS in an oven or on a hotplate e.g. at 80 °C for 10 min (Fig. 1e).
  4. Surface treatment (e.g. micro-contact printing) or plasma-bonding of a PDMS chip to the glass surface can be performed after or before the dish assembly (Fig. 1f).

  1. To keep humidity for on-chip cell culture, the dish can be filled with e.g. phosphate buffered saline (Fig. 2a). Dishes with chips or micropatterns loaded with cells can be placed in a CO2 incubator with or without further protection (Fig. 2b).
  2. Long-term live cell imaging can be performed using a stage top incubator (Fig. 2c).

 


What else should I know?

  1. Depending on the support type of microscopes, it might be necessary to well align the contours of the dish and the glass slide. This can be done using cylindrical magnets (3 per dish) and a ferromagnetic metal plate (Fig. 3a) during PDMS curing in an oven or on a hotplate (Fig. 3b).

 

 


Acknowledgement

This work is supported by the French National Research Agency (ANR) as part of the “Investissements d’Avenir” program (reference: ANR 10-NANO 0207) and ERC Advanced Grant CellO (FP7-IDEAS-ERC-321107).

 

Reference

[1] Caballero D, Samitier J, Different strategies for the fabrication of cell culture chambers for live-cell imaging studies. Chips and Tips, 02 Dec 2014 (http://blogs.rsc.org/chipsandtips/2014/12/02/different-strategies-for-the-fabrication-of-cell-culture-chambers-for-live-cell-imaging-studies/)

 

 

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Multilayer photolithography with manual photomask alignment

Frank Benesch-Lee, Jose M. Lazaro Guevara, and Dirk R. Albrecht

Worcester Polytechnic Institute, Worcester, MA 01609 USA

Why is it useful?

Modern microfluidic devices can incorporate channels of different heights to fulfill their designed function. Examples include hydrodynamic focusing [1], cell traps [2], and chambers that isolate cellular components [3]. These devices are fabricated from a multilayer SU-8 photoresist master mold. Each layer height requires a separate set of photolithographic steps, including photoresist spin, photomask alignment, exposure, and bakes, followed by a development step at the end to reveal the 3D resist pattern.

Mask aligners have microscopes and stage micrometers for precise, micron-scale alignment of each layer’s photomask with visible marks on the substrate wafer.  They are indispensable tools for creating multilayer patterns with accurate registration, but while available in cleanrooms at many research universities, their substantial expense may place them out of reach of teaching institutions and individual laboratories.

In contrast, single-layer microfluidics can be prepared using an inexpensive UV light source, or even a self-made one [4]. In principle, manual photomask alignment could be made under a microscope, then brought to the UV source, yet this poses several complications. First, alignment features can be very difficult to see using inexpensive microscopes or stereoscopes, especially in thin SU8 layers, due to poor contrast between exposed and unexposed regions before development. Second, misalignment can occur during movement to the exposure system.

Here we present a manual photomask positioning method that yields a 50 µm accuracy, without the aid of a mask aligner.

 

What do I need?

  • Equipment and supplies for photolithography:
    • Spin coater, and UV exposure system
    • Substrate wafer and SU-8 photoresist
  • Small microscope (e.g. USB) or stereo microscope
  • Photomask transparencies for each layer
  • Scotch tape
  • Fine-tip permanent marker
  • Straight razor blade
  • Cutting mat
  • 4 small (3/4”) or mini (1/2”) binder clips
  • Glass plate, approx. 4 x 5”, compatible with exposure system

 

What do I do?

  • Cut the photomasks from the transparency sheet, leaving 4 corner tabs. Align the two masks relative to each other under the microscope (Figure 1a) and clip them together with a binder clip. Ensure correct mask orientation and check alignment accuracy at multiple alignment marks across the mask. (Note that horizontal alignment accuracy with a stereomicroscope is low, because each eye’s optical path is angled 5 – 8 degrees, whereas vertical alignment is unaffected. Align in the vertical direction first then rotate the masks 90 degrees to ensure accurate alignment in both horizontal and vertical directions.) Add binder clips to each corner (Figure 1b), and verify alignment. Next, remove one binder clip at a time and use a straight razor blade to cut a sharp V-notch into each tab, through both masks. Press the blade straight down to avoid shifting the alignment. Replace the binder clip, and proceed to the next corner until all 4 notches are cut (Figure 1c).

 

 

 

 

  1. Spin the first layer of SU-8 onto the wafer to the desired thickness and prebake. Attach 4 pieces of scotch tape onto the bottom of the wafer so that the sticky side faces up (Figure 2a). Position the first mask on the wafer, pressing gently to adhere it to the tape tabs. Use a fine-tip marker to trace the alignment notches (Figure 2b) onto the scotch tape (Figure 2c).  Transfer to the UV exposure system and expose.  Carefully remove the mask without detaching the scotch tape from the wafer and postbake.  Scotch tape is compatible with 95 °C baking.  Apply an additional piece of tape to cover the sticky tape tabs to protect the marker from smearing and allow smooth alignment of the next mask.

 

 

 

  1. Spin coat the next photoresist layer and prebake (Figure 3a). Mount the wafer onto a glass plate with a loop of scotch tape to keep it in place. Position the second mask onto the wafer, ensuring that alignment “V” markings are centered within each alignment notch and across all 4 corners (Figure 3b). Affix the mask to the glass plate with thin (2-3 mm wide) pieces of tape, and adjust alignment as necessary.  Carefully transfer the glass plate with wafer and aligned photomask for exposure (Figure 3c).

 

 

  1. Repeat step 3 for any additional layers. Remove the tape tabs and develop the photoresist. Evaluate alignment accuracy under a microscope (Figure 4).

 

 

Conclusions:

In this tip, we present a method for manual alignment of multiple transparency photomasks.  We achieved repeatable accuracy of <100 µm and as good as 50 µm (Figure 4a). These accuracies are within required tolerances of many multilayer designs (Figure 4b).  In many cases, minor design alternatives can relax alignment tolerances, such as in a trap design containing a thin horizontal channel that allows fluid bypass but captures larger objects (Figure 4c). In this example, a 100 µm wide bypass channel only partially covered the trap indentations, whereas widening the bypass channel to 400 µm enabled a functional device despite slight misalignment.  Overall, this simple method allows fabrication of microfluidic device molds containing multiple layer heights, without expensive mask alignment equipment, to an accuracy of at least 50 µm.  Furthermore, after alignment marks are cut, no microscope is needed at all during the photolithography process, speeding the fabrication of multiple masters.

 

Acknowledgments:
Funding provided by NSF IGERT DGE 1144804 (FBL), Fulbright LASPAU (JMLG), University of San Carlos of Guatemala (JMLG), NSF CBET 1605679 (DRA), NIH R01DC016058 (DRA), and Burroughs Wellcome CASI (DRA).Acknowledgments:

 

References:

  1. Chih-Chang, C., H. Zhi-Xiong, and Y. Ruey-Jen, Three-dimensional hydrodynamic focusing in two-layer polydimethylsiloxane (PDMS) microchannels. Journal of Micromechanics and Microengineering, 2007. 17(8): p. 1479.
  2. Erickson, J., et al., Caged neuron MEA: A system for long-term investigation of cultured neural network connectivity. Journal of Neuroscience Methods, 2008. 175(1): p. 1-16.
  3. Taylor, A. M., et al., A microfluidic culture platform for CNS axonal injury, regeneration and transport. Nature Methods, 2005. 2(8): p. 599-605.
  4. Erickstad, M., E. Gutierrez, and A. Groisman, A low-cost low-maintenance ultraviolet lithography light source based on light-emitting diodes. Lab on a Chip, 2015. 15(1): p. 57-61.

 

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A simple, bubble-free cell loading technique for culturing mammalian cells on lab-on-a-chip devices

Sahl Sadeghi1­* and Meltem Elitas1

1 Faculty of Engineering and Natural Sciences, Sabanci University, 34956, Istanbul, Turkey

* Sahl Sadeghi wrote the paper.

 

Purpose

Lab-on-a-chip (LOC) devices significantly contribute different disciplines of science. Polydimethylsiloxane (PDMS) is one of the main materials, which is widely used for the fabrication of biological LOCs, due to its biocompatibility and ease of use. However, PDMS and some other polymeric materials are intrinsically water repellant (or hydrophobic), which results in difficulties in loading and operating LOCs. The eminent consequence of hydrophobicity in LOCs for biological systems is the entrapment of air bubbles in microfluidic channels. Although the oxygen plasma treatment of PDMS reduces the surface hydrophobicity for a certain period of time, the hydrophilic property of PDMS vanishes over time1. The persistent problem of bubbles in the microfluidics led to several studies conducted to overcome it. Some of these solutions suggested implementing bubble traps2,3, surface treatment of LOCs through hydrophilic coatings4, and using actively controlled bubble removal systems 5,6.

Although the aforementioned design complexities are introduced to LOCs in order to reduce the clogging problem caused by the bubbles, these modifications also result in higher production cost, complex operation, and long device preparation time. In many single-cell experiments without losing or damaging the rare cells, these cells needs to be introduce into the LOCs. Here, we present a simple method that enables loading a small number of cells without introducing bubbles in the microfluidics channels.


Materials

·         PDMS (Dow Corning Sylgard 184 Silicon Elastomer Kit)
·         Pipette tips (20-200 ul, Eppendorf, # 3120000917)
·         Pipetman (Gilson, P200, #69989-5)
·         Aqueous ethanol 70% (ZAG Chemistry)
·         Cell culture medium (DMEM, PAN Biotech, #P04-01548)
·         Mammalian cells (MCF7, ATCC-HTB-22)
·         Sterile syringe (BD 10 ml Syringe, Luer-Lok Tip, #300912)
·         Sterile Hamilton syringe (Hamilton, 100 ul SYR, #84884)

 


Procedure

Step 1: Insert two 200-ul pipet tips at the inlet and outlet ports of the PDMS device as illustrated in Figure 1.

Step 2: Introduce a 70% aqueous ethanol into the inlet-pipet tip using a pipetman. Thus, the inner surface of microfluidic channels will be disinfected and the fluid flow will be tested within the micro channels as it is applied in many other protocols for LOCs7,8.

Step 3: Gently apply pressure pressing the pipetman to force ethanol solution flow through the micro channels and cavities of the PDMS device. Take care to avoid applying negative pressure from the outlet-pipet tip, which might create air leakage through the pipet connections. Besides, applying a negative pressure will directly affect the amount of a gas dissolved in the liquid according to Henry’s law9 that might contribute formation of more bubbles inside the micro-channels. The positive pressure will facilitate removal of the air bubbles via dissolving them.

Step 4: After flushing the chip with ethanol solution, inspect the chip to ensure bubble removal. In case of air bubbles, repeat the steps 2 and 3.

Step 5: Fill the syringe (10 ml) with medium or phosphate buffered saline (PBS). Take care to remove the air bubbles inside the syringe, mount and lock the needle on the syringe. Then, flow medium through the needle too make sure the needle is full of medium without any bubble. Insert the needle in the inlet-pipet tip; gently apply positive pressure to replace the ethanol with medium. Next, collect the excess medium from the outlet-pipet tip.

Step 6: Fill the inlet pipet with fresh medium in such a way that due to certain height (h) between the levels of the medium in the inlet and outlet pipet tips, very slow medium flow will be established inside the micro-channels.

Step 7: Load your cells into the Hamilton syringe and take care to ensure that there is no air bubble inside its needle and syringe. Insert the needle of the Hamilton syringe into the inlet-pipet tip as explained in Step 5, Figure 1. Introduce the cells via applying gentle positive pressure to the syringe. Established flow streams in the PDMS chip will deliver the released cells to the desired positions in the chip. Flow rate can be arranged adjusting the applied positive pressure and amount of medium collected in the inlet and outlet pipet tips. The excess supernatant from the outlet-pipet tip can be collected, and fresh medium can be supplied through the inlet-pipet tip during the experiment.


References

  1. Tan, S.H., N.T. Nguyen, Y.C. Chua, and T.G. Kang,. Biomicrofluidics, 2010. 4(3).
  2. Zheng, W.F., Z. Wang, W. Zhang, and X.Y. Jiang,. Lab on a Chip, 2010. 10(21): p. 2906-2910.
  3. Wang, Y., D. Lee, L.S. Zhang, H. Jeon, J.E. Mendoza-Elias, T.A. Harvat, S.Z. Hassan, A. Zhou, D.T. Eddington, and J. Oberholzer, . Biomedical Microdevices, 2012. 14(2): p. 419-426.
  4. Wang, Y.L., C.E. Sims, and N.L. Allbritton,. Lab on a Chip, 2012. 12(17): p. 3036-3039.
  5. Karlsson, J.M., M. Gazin, S. Laakso, T. Haraldsson, S. Malhotra-Kumar, M. Maki, H. Goossens, and W. van der Wijngaart,. Lab on a Chip, 2013. 13(22): p. 4366-4373.
  6. Cortes, D.F., T.-X. Tang, D.G.S. Capelluto, and I.M. Lazar,. Sensors and Actuators B: Chemical, 2017. 243: p. 650-657.
  7. Benavente-Babace, A., D. Gallego-Perez, D.J. Hansford, S. Arana, E. Perez-Lorenzo, and M. Mujika,. Biosensors & Bioelectronics, 2014. 61: p. 298-305.
  8. Yesilkoy, F., R. Ueno, B.X.E. Desbiolles, M. Grisi, Y. Sakai, B.J. Kim, and J. Brugger,. Biomicrofluidics, 2016. 10(1).
  9. Henry, W., Phil. Trans. R. Soc. Lond., 1803. 93: 29–274.

Figure 1 – Schematic of cell loading procedure in a microfluidic PDMS device.

 

 

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A novel low cost method to prepare a cross-linked gelatin membrane for potential biological applications

Gabriele Pitingolo1 and Valerie Taly1

 1 INSERM UMRS1147, CNRS SNC 5014, Université Paris Descartes, Equipe labellisée Ligue Nationale contre le cancer 2016. Paris, France.

email: gabriele.pitingolo@parisdescartes.fr

 

Why is this useful?

In recent years, several gelatin types (i.e. GelMA, A or B) have been used in pharmaceutical formulation and tissue engineering due to their excellent biocompatibility, propensity to cell differentiation and availability at low cost.1 The use of gelatin as biomaterial is also advantageous for the possibility to tune the mechanical properties of the substrate, changing the concentration in water and the degree of cross-linking. However, Transwell® is the most used permeable support with microporous membranes and is a standard method for culturing cells.2 This commercial type of support has been widely used to study the molecular secretion by different cell types and also to reproduce several in vitro physiological barriers (i.e. blood brain barrier).3 Transwell® cell culture inserts are convenient, because they are sterile and easy-to-use, but they are very expensive (~ 300 $ for a 12 pack) and possess a limited range of biomaterial properties because they are made from polyester or polycarbonate. Furthermore, as shown by Falanga and colleagues, these porous membranes are often integrated onto microfluidic chip for permeability studies.4

Recently, Yong X. Chen et al. proposed an alternative method to prepare a suspended hydrogel membrane platform for cell culture, designing a complicated protocol to synthesize the GelMA and to fabricate an open-grid structure made of polylactic acid (PLA) polymer using a commercial printer.5 Our tip shows a novel low-cost method for preparing a cross-liked gelatin membrane as a permeable support useful for potential biological applications. In addition, the proposed protocol doesn’t require the use of sophisticated fabrication technologies or expensive materials. It uses gelatin from porcine skin and the formaldehyde vapor technique to cross-link the gelatin membrane. As proof of concept, we integrate the gelatin membrane into a microfluidic chip, to show the possibility to develop a platform for comparable studies in static and dynamic conditions.

Furthermore, to demonstrate the resistance to culture temperature (around 37° C) of the prepared gelatin membrane, we tested the mechanical properties (Young’s modulus) before and after the incubation time (2 days). Finally, we observed the preservation of the mechanical properties and structural integrity that makes the membrane usable for studies with cell culture.


 

What do I need?

  • Transwell insert
  • Porcine gelatin type A
  • Formaldehyde solution
  • Scalpel
  • PMMA milled chamber or similar

 

What do I do?

  1. Remove the porous membrane from the Transwell® insert (Fig. 1a-1c) or alternatively use a similar homemade support. To facilitate this step is convenient to use a scalpel to incise the membrane along the entire diameter.

 

  1. After the removal of the membrane, the Transwell® support is ready to use. Position the structure at the center of the PMMA chamber (depth 2 mm at least) (Fig 2a) and pour liquid 10% w/v gelatin, without bubbles, onto the PMMA chamber and inside the Transwell® support (Fig 2b). After 2 min of stabilization, put the system in the fridge, for at least 10 minutes.

 

  1. After the gelation time (10 minutes at 4°C) it is possible to remove the formed gelatin membrane from the PMMA chamber, with the aid of a scalpel to facilitate the detachment (Fig 3a-3b). As shown in Figure 3c the gelatin membrane it appears very flat, an ideal characteristic for cell cultivation (Fig 3c). To guarantee the preservation of the mechanical properties during the cell culture step, cross-link the gelatin membrane using the classical protocol “cells-biocompatible”, such as glyceraldehyde6, formaldehyde7 and glutaraldehyde8 methods or natural products such as genipin.9

 

  1. In this case, we used the vapor formaldehyde method to cross-link the prepared gelatin membrane and to obtain a system with lower aqueous solubility, higher mechanical strength and stability against enzymatic degradation. We exposed the gelatin membrane to formaldehyde vapors for 1 day. In Figure 4a we show the difference, after 48 h of culture conditions, between a sample cross-linked (left) and not (right). The final depth of the cross-linked gelatin membrane is around 1 mm, however, it is possible to change the depth tuning the liquid gelatin amount. Finally, we calculated, before and after the incubation time, the young’s modulus of the cross-linked gelatin membrane, observing a similar value of 40 kPa (compression test by using hydraulic testing system Instron DX).

 

  1. Integration of the gelatin membrane into a microchip. In this section, we detail the integration of the gelatin membrane into a microfluidic chip. As example, we used the same geometry proposed in our previous work4, to make a device for a permeability experiment (Figure 5a). As shown in Figure 5b, just pour the liquid gelatin into the smaller drilled microchannel, using a pipette to form a thin uniform layer of gelatin (Figure 5c). After gelation at 4° C, cross-link the formed gelatin membrane using the previously described method, the result is shown in Figure 5d. To bond the different PMMA-PDMS substrates we propose here a magnetic approach recently developed by our group10, to preserve the gelatin membrane by physical-chemical stress as in the case of solvent evaporation and plasma bonding. Figure 5e shows the final chip.

 

 


Conclusions: In this tip, a novel biocompatible gelatin permeable support was obtained by using a simple and low cost fabrication method. Vapor formaldehyde method or other chemical crosslinking approach can be applied to crosslink the integrated gelatin membrane for the use as potential scaffolds for cell culture. Furthermore, the Young’s modulus and the thickness of the permeable membrane can be adjusted by changing the initial concentration of the gelatin, the degree of cross linking and the amount of liquid gelatin. Finally, we showed the integration of the gelatin membrane into a modular microchip. Therefore, we propose an easy and low cost method to prepare a permeable gelatin membrane for cell biology and for other applications.

 

 

Acknowledgements

This work was carried out with the support of the Pierre-Gilles de Gennes Institute equipment (“Investissements d’Avenir” program, reference: ANR 10-NANO 0207).

 

References

  1. Geckil, Hikmet, et al. “Engineering hydrogels as extracellular matrix mimics.” Nanomedicine 5.3 (2010): 469-484.
  2. https://www.corning.com/worldwide/en/products/life-sciences/products/permeable-supports/transwell-guidelines.html
  3. Guarnieri, Daniela, et al. “Shuttle‐Mediated Nanoparticle Delivery to the Blood–Brain Barrier.” Small 9.6 (2013): 853-862.
  4. Falanga, A. P., Pitingolo, G., Celentano, M., Cosentino, A., Melone, P., Vecchione, R. & Netti, P. A. (2016). Shuttle‐mediated nanoparticle transport across an in vitro brain endothelium under flow conditions. Biotechnology and Bioengineering.
  5. Chen, Yong X., et al. “A Novel Suspended Hydrogel Membrane Platform for Cell Culture.” Journal of Nanotechnology in Engineering and Medicine 6.2 (2015): 021002.
  6. Sisson, Kristin, et al. “Evaluation of cross-linking methods for electrospun gelatin on cell growth and viability.” Biomacromolecules 10.7 (2009): 1675-1680.
  7. Usta, M., et al. “Behavior and properties of neat and filled gelatins.” Biomaterials 24.1 (2003): 165-172.
  8. Talebian, A., et al. “The effect of glutaraldehyde on the properties of gelatin films.” Kemija u industriji 56.11 (2007): 537-541.
  9. Bigi, A., et al. “Stabilization of gelatin films by crosslinking with genipin.” Biomaterials 23.24 (2002): 4827-4832.
  10. Pitingolo Gabriele, et al. “Fabrication of a modular hybrid chip to mimic endothelial-lined microvessels in flow conditions.” Journal of Micromechanics and Microengineering” (Accepted manuscript)
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Electric drill/driver as centrifuge with 3D-printed custom holders for non-conventional containers

Minkyu Kima, Guanya Shib, Ming Panc, Lucas R. Blaucha, and Sindy K.Y. Tanga*

aDepartment of Mechanical Engineering, Stanford University, Stanford, CA 94305, USA

bUndergradute Visiting Research Program, School of Engineering, Stanford University, Stanford, CA 94305, USA

cDepartment of Materials Science and Engineering, Stanford University, Stanford, CA 94305, USA

*sindy@stanford.edu


Why Is This Useful?

Many processes in biological and chemical preparation require centrifugation steps. The transfer of samples between the original sample container and tubes required by commercial centrifuges increases the risk of sample contamination, and often leads to the loss of samples. Commercial centrifuges are also not readily available outside laboratory settings. Here we show the design of a simple 3D-printed holder for attaching to the chuck of an electric drill/driver which we use as a centrifuge. The advantages of this method include: 1) The holder can be designed to hold non-conventional containers (e.g., syringes, glass vials, capillaries). 2) Electric drill/drivers are more widely available than centrifuges. We show that a variety of samples (e.g., water-in-oil emulsions, cell suspensions, food and drinks, wet soil) in various containers can be centrifuged with our method. This method should be useful for field work outside of the laboratory, and for the wider DIY community interested in home-based applications that require centrifugation, such as blood separation and related diagnostics, separation of interstitial water from wet soil for pollution detection, extraction and identification of allergens in food samples, and fluid clarification (e.g., olive oil, wine) by accelerating sedimentation.

What do I need?

Figure 1. Photo of components needed.

  1. Electric drill/driver (DeWalt DC742KA Cordless Compact Drill/Driver Kit [1]).
  2. 3D-printed custom holder.
  3. 3D-printed custom support for the drill/driver.
  4. Four 1 mL-syringes (NORM-JECT, Part No.: 4010-200V0) as an example of non-conventional sample containers.
  5. One long bolt (Pan Head Machine Screw, Zinc, #8 x 1-1/4”) and one matching nut (Hex Nut, Zinc, #8-32). The dimensions of the bolt should match the chuck of the drill/driver.
  6. Four short bolts (Pan Head Machine Screw, Zinc, #6 x 1/2”) to secure the 1 mL-syringes to the 3D-printed custom holder.

What do I do?

  1. Design a custom holder using Solidworks or other CAD software.
    1. Measure the outer diameter (w1 = 6.5 mm) of the 1 mL-syringe (Fig. 2a). We included a 0.5 mm-tolerance in deciding the width of the slot (w2 = 7 mm) into which the 1 mL-syringe will be secured (Fig. 2b).
    2. Decide the angle (q = 60°) at which the 1 mL-syringe will be tilted relative to the plane of rotation.
    3. Decide the length (w3 = 80 mm) and the thickness (w4 = 15 mm) of the holder.
    4. Measure or identify the outer diameters of the short and long bolts, and use these values for f3 and f5 respectively.
  2. Design a custom support for the drill/driver (Fig. 2c). The dimensions of this support are not critical so long as the drill/driver is stable and does not topple during operation.
  3. 3D-print the holder and the support. We used a 3D-printer by ROBO 3D [2]. The resolution of the 3D-printer in xyz direction is 100 mm. The material we used was polylactic acid (PLA).
  4. Assemble the centrifuge (Fig. 2d).
    1. Put one long bolt through the center of the holder and tighten with the nut.
    2. Insert the bolt into the chuck of the drill/driver and tighten the bolt by pushing the trigger of the drill/driver a few times.
    3. Make sure the bolt is fixed in the chuck and aligned to the drill/driver.
    4. Place the drill/driver in the 3D-printed support.
    5. Secure four 1 mL-syringes to the holder using the four short bolts.
  5. Start the centrifuge by pushing the trigger of the drill/driver for 5-10 minutes. The plane of rotation should be parallel to the floor.
  6. Unscrew the short bolts to remove the syringes.
  7. If desired, measure the rotational speed of the drill/driver before inserting the real samples. We used the SLO-MO mode in iPhone to calibrate the rotational speed of the drill/driver [3] (Fig. 2e).
  8. Results (Fig. 3).
    1. Water-in-oil emulsion: Micron-sized uniform water-in-oil droplets were collected in a 1 mL-syringe. After a needle was connected to the syringe, the syringe was secured to the 3D-printed holder with needle pointing up. The holder was balanced before centrifugation by adding another syringe containing an equal weight of fluids to the opposite side of the holder. We centrifuged the sample at a speed of 374 rpm for 10 minutes. Droplets were then injected into a microchannel to measure the change of volume fraction before and after centrifugation. The volume fraction was defined as the ratio of the total volume of water droplets to the total volume of fluids filling up the channel. After centrifugation, the volume fraction of the emulsion increased from 60% to 86% without a change in the size of the droplets. Neither break-up nor coalescence of the droplets was observed.
    2. Stentor coeruleus: To demonstrate the concentration of cell suspensions, we used Stentor coeruleus (www.carolina.com) as a model. We filled a 3 mL-syringe with about 20 Stentor cells suspended in 2 mL of aqueous culture media (concentration ~ 10 cells/mL). A needle was connected to the syringe which was then secured in the 3D-printed holder with the needle pointing down. The cell suspension was centrifuged at a speed of 374 rpm for 5 min. The cells were concentrated at the bottom, close to the entrance into the needle. It was then possible to inject this concentrated cell suspension through the needle into a polyethylene tubing. Fig. 2-i) shows the microscopic image of the cells in about 15 mL of aqueous culture media in the tubing (concentration ~ 400 cells/mL).
    3. Korean rice wine (Makgeolli): A separate holder was designed to hold a 20 mL-glass vial. The vial was filled with 12 mL of Korean rice wine and centrifuged at a speed of 1252 rpm for 10 minutes. The sediment was clearly observed after centrifugation. On the other hand, the sediment was not observed for more than 20 minutes without centrifugation.
    4. Wet soil: 10 mL of wet soil in a 20 mL-glass vial was centrifuged at a speed of 1252 rpm for 10 minutes. Interstitial water was separated from the soil.

Figure 2. a) A 1-mL syringe as non-conventional container. b) Drawing of 3D-printed holder generated by SolidWorks. c) Drawing of 3D-printed support generated by SolidWorks. d) Photograph of experimental setup. The 3D-printed holder was connected to the drill/driver placed on the 3D-printed support. Four 1 mL-syringes were then tightened using the short bolts. e) Calibration plot of the rotational speed versus the trigger levels on different modes of the drill/driver. Mode 1 and Mode 2 indicate different gear settings in the transmission of the drill/driver. The numbers 1 to 3 in each mode indicate user-defined trigger levels. The rotational speed ranges from 120 rpm to 1252 rpm. The rotational speeds were measured using SLO-MO function in iPhone 6.

Figure 3. a) Photographs of emulsion in 1 mL-syringe and microscopic images of emulsion injected into a microchannel before and after centrifugation. The scale bar in the photographs is 5 mm. b) Photographs of Stentor cells in 3 mL-syringe before and after centrifugation. The red arrows indicate individual cells. The scale bar is 5 mm. i) Microscopic image of 6 Stentor cells in a polyethylene tubing after centrifugation. The scale bar is 300 mm. c) Photographs of Korean rice wine in 20 mL-glass vial before and after centrifugation. The red box indicates sediments. The scale bar is 10 mm. d) Photographs of wet soil in 20 mL-glass vial before and after centrifugation. The red box shows interstitial water separated from wet soil. The scale bar is 10 mm.


What else should i know?

  1. The centrifugal force can be increased by lengthening the arms (w3) holding the containers.
  2. The rotational speed can be measured using a high-speed camera or a smart phone with slow-motion videotaping capability, so long the frame rate is sufficient for the rotational speed used.
  3. The load in the centrifuge should be balanced.
  4. For safety purposes, safety goggles should be worn. The centrifuge should also be placed inside a safety barrier (e.g., a sturdy laundry basket). The safety instructions for the drill/driver should also be observed.
  5. After centrifugation for 10 minutes, we found that the drill/driver started to heat up. If centrifugation time longer than 10 minutes is needed, it should be possible to perform multiple rounds of 10-min centrifugation steps with breaks in between to cool down the drill/driver.

Conclusion
In this work, we demonstrated that 3D-printed holders attached to an electric drill/driver can be used for the centrifugation of samples in non-conventional containers. As 3D-printers and hand drills are easily accessible, we expect this tip to find immediate use in settings outside laboratories for field work, and also at home for DIY users.

    Acknowledgements

    We acknowledge support from the Stanford Woods Institute for the Environment and the National Science Foundation (Award #1454542 and #1517089).

    References

    1. http://www.dewalt.com/products/power-tools/drills/drills-and-hammer-drills/12v–38-10mm-cordless-compact-drilldriver-kit/dc742ka

    2. http://store.robo3d.com/collections/all/products/r1-plus-3d-printer?variant=6274616835

    3. http://www.imore.com/how-to-record-video-iphone-ipad


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Simple and Low-cost Contact Angle Measurements Using a Smartphone with a PDMS-Lens

Jonas M. Ribea, Nils R. Skovb, Ole-Andreas K. Kavlia, Armend G. Håtia, Henrik Bruusb and Bjørn T. Stokkea

a Department of Physics, Norwegian University of Science and Technology, NO–7491 Trondheim, Norway

b Department of Physics, Technical University of Denmark, DK–2800 Kongens Lyngby, Denmark

jonas.ribe@ntnu.no


Why Is This Useful?

Contact angle measurements are important for characterizing the wettability of a liquid to a solid surface. In microfluidics they are of special interest as they provide insight into the intermolecular interactions between the sample liquid and the microchannel surface. Contact angle measurements are also important when assembling polydimethylsiloxane (PDMS) devices using oxygen plasma bonding. For optimal bond strength the water contact angle of plasma treated PDMS should be minimized as shown by Bhattacharya et al. [1] A current hurdle in measuring contact angles is the requirement of a setup that is expensive and non-portable. Here we show a method for measuring contact angles using materials and equipment found in a typical microfluidics lab.

What do I need?

Consumables:

Equipment:

  • Smartphone
  • Digital scale
  • Desiccator with vacuum pump
  • Oven
  • Syringe pump (optional)
  • Light source

For measurements:

  • Pipette (0.5–3μL)
  • Sample (e.g. deionized (DI) water or other liquid sample)

What do I do?

Prepare PDMS:

  1. Weigh 10:1 PDMS (Sylgard 184) in a plastic cup on the digital scale
  2. Mix the PDMS by hand using a plastic spoon
  3. Degas the PDMS in a desiccator to remove the bubbles

Make PDMS-lens:

  1. Use the tip of the plastic spoon handle (or a pipette) to place a small droplet of uncured PDMS in the center of a glass cover slip. Repeat with various amounts of PDMS to obtain lenses with varying magnification.
  2. Mount the cover slips upside down (e.g. between two glass slides) and cure the PDMS hanging at 70 °C for 15 min. Longer curing times might be necessary, if the drop is relatively large.
  3. Center the PDMS-lens over the camera of your smartphone and fixate it using tape.
  4. Test the focus of your camera. For our camera setup the best images were captured with lenses that focus around 2 cm.

Contact angle measurements:

Smartphone contact-angle setup: (A) Focus test of a PDMS lens. (B-C) The smartphone mounted on a syringe pump. The PDMS-lens is mounted on the front facing camera of an iPhone 6S and the sample is centered in front of the lens. The sample is mounted on the pusher block of a syringe pump which can be moved to adjust the focus.

  1. Make a sample stage preferably using a syringe pump or some other system that you can move. We mounted the smartphone on the syringe holder block with the camera pointing towards the pusher block. Make a sample holder on the pusher block using glass slides or other consumables found in the lab. Align the center of the stage with the center of the camera. Tip: aligning is easier if done using the sample that you want to measure. Put the sample on the block and move it into focus by releasing the pusher block and sliding it away/towards the camera. Increase the height of the stage until the top of the sample is centered in the camera.
  2. Place the light source behind the sample and illuminate the stage evenly. Tip: put the sample stage in front of a white wall and light up the wall for a homogenous background and optimal contrast.
  3. Place a small drop (0.5–3 μL) of DI water on top of the sample using a pipette. Place the drop near the sample edge closest to the camera.
  4. Move the sample edge into focus. Block out ambient light in the room.
  5. Measure the contact angle of the drop in the image e.g. using ImageJ [2] software with a plugin for contact angle measurements [3] or get a rough estimate using an app on your smartphone.[4]

Contact angle measurements of water on PDMS: (A) Raw image from iPhone 6S front-facing camera with PDMS-lens. (B) Direct measurement using app on smartphone (based on θ/2 calculation) (C-E) ImageJ measurements using DropSnake plugin. Unmodified PDMS (C) and PDMS treated with oxygen plasma with increasing intensity (D-E).


What else should I know?

The focal length of the PDMS-lens is determined by the volume of PDMS used as described by Lee et al. [5]. However, it is difficult to control the volume of PDMS using a pipette due to the high viscosity of PDMS. We recommend making a range of lens sizes and testing them on your smartphone camera to see which gives the right focal length. If your digital scale has milligram precision you can measure the amount of PDMS used for each lens. The mass of each PDMS-lens is typically less than 10 mg. You can decrease the focal length further by adding PDMS to an already cured lens. Modern smartphones have both a rear-facing and a front-facing camera and in our experience the drop focusing was easier when using the front facing camera. The images taken here were captured with an iPhone 6S from Apple using the front-facing camera with a 5MP sensor. The weight of the cured PDMS lens was 7 mg.

Tip: you can also remove the PDMS-lens from the cover slip and place it directly on your camera. Although, it might be more difficult to center.

Calculating contact angles from images of sessile drops can be done using a range of techniques.[6] If the drop volume is small and the contact angles are not extreme, we can generally neglect droplet distortion due to gravitational effects. Extrand and Moon [7] calculated that gravitational effects can be neglected for a water droplet sitting on a hydrophilic surface (θ=5°) if its volume is less than 5 μL and less than 2.7 μL on a hydrophobic surface (θ=160°). If we assume the drop to be spherical, the contact angle can be estimated by multiplying the angle between the base and the height of the droplet by 2. This is referred to as the θ/2-method and is implemented by e.g. the Contact Angle Measurement app [4] for iOS. Sessile drop measurements are generally limited by the experimental setup and operator error, but typically has a precision of ±3°.[8] Image-processing algorithms relying on curve fitting of the droplet outline can enhance reproducibility. ImageJ [2] with DropSnake-plugin [3] uses active contours (energy minimization) to track the outline of the drop and calculate contact angles. This increases precision, but is slower and currently requires analysis on a separate computer.

Acknowledgements

The Research Council of Norway is acknowledged for the support to the Norwegian Micro- and Nanofabrication Facility, NorFab (197411/V30).

References

  1. S. Bhattacharya, A. Datta, J. M. Berg and S. Gangopadhyay, J. Microelectromech. S., 2005 14, 590–597
  2. ImageJ software
  3. DropSnake ImageJ-plugin for contact angle measurements
  4. Contact Angle Measurement iOS app (Japanese)
  5. W. M. Lee, A. Upadhya, P. J. Reece, and T. G. Phan, Biomed. Opt. Express, 2014, 5, 1626–1635
  6. Y. Yuan and T. R. Lee, Surface Science Techniques, Springer, Berlin/Heidelberg, 2013, 51, 3–34.
  7. C. W. Extrand and S. I. Moon, Langmuir, 2010, 26, 11815–11822.
  8. A.F. Stalder, G. Kulik, D. Sage, L. Barbieri and P. Hoffmann, Colloids and Surfaces A: Physicochem. Eng. Aspects, 2006, 286, 92–103.
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Universal and Multi-material Bonding Method for Rapid and Low-cost Assembly of Microfluidic Devices

Ya-Yu Chiang, Nikolay Dimov, Nicolas Szita

Department of Biochemical Engineering, University College London, London, United Kingdom

E-mail: n.szita@ucl.ac.uk


Why is this useful?

The packaging of micro-systems relies strongly on the capability to bond different types of materials reliably whilst maintaining the microstructures and their dimensions. However, the bonding of different materials each with their specific physical and chemical properties frequently turns into a tedious, thus time consuming operation; often, the choice of materials and microfabrication techniques are limited by the bonding technique. Particularly challenging for bonding can be combinations of quartz, glass or silicon with polymers and metals.

Here we demonstrate a rapid, low-cost, UV-irradiation based bonding method, which is suitable for the bonding and assembly of quartz-to-silicon, quartz-to-metal, quartz-to-polymer, quartz-to-quartz devices.  We demonstrate in detail on the more challenging combinations, namely the bonding of a quartz slide to an aluminum sheet. In our example, the aluminum sheet contains the microfabricated structure. The same procedure is applicable for the other material combinations, i.e. quartz-to-silicon, quartz-to-polymer, quartz-to-quartz or quartz-to-metal for a metal other than aluminium; the main requirement for implementing our method is that at least one material is transparent to UV light.


What do I need?

  • Aluminum sheets, thickness of 1 mm (e.g. AW6082-T6, Smiths Metal Centres, UK)
  • Micro milling machine (e.g. CNC MicroMill GT, Minitech, US)
  • Flat head end-mills 0.25 mm, and 2 mm (e.g. PMT Endmill, US)
  • Plasma Cleaner (e.g. PDC-32G-2, Harrick Plasma, UK)
  • UV-curing adhesive (e.g. NOA 61, Noland Products, UK)
  • UV lamp, 100 W, 365 nm (e.g. B-100 AP, UVP, Cambridge, UK)
  • Quartz microscope slide, fused quartz, 25.4 × 76.2 x 1 mm3 (e.g. 42297, Alfa Aesar, UK)


What do I do?

1. Device design and leveling of the metal substrate

  • Draw your device design in any available computer aided design (CAD) software. As the surface roughness of the metal substrate can vary, a polishing step is recommended prior to the actual fabrication.
  • Generate the G-code for the CNC machine using any CAM/CAD software. Two separate files are required: one for the polishing of the substrate, and one for the actual design.

2. Micro milling

  • Clamp the aluminum substrate on the table of the milling device. Make sure that you do not bend the material.
  • Set the initial coordinates (X0, Y0, Z0) for this work.
  • Polish the aluminum substrate with 2 mm flat head end-mill.
  • Change to the smaller diameter tool (0.25 mm).
  • Mill the designed structure in the aluminum sheet with the 0.25 mm end-mill.

3.  Cleaning the aluminum substrate from residues.  Dust is removed first with water, and then the surface is cleaned  first with ethanol, and then with compressed air. Finally the substrate is dried in an oven (120ᴼC, 30 min).

4. Plasma activation of the quartz microscope slide. Place the quartz microscope slide inside the Plasma Cleaner. The plasma treatment is a ‘surface process’, therefore the surface that is about to be bonded should be facing towards the center of the chamber.

  • Evacuate the chamber until a working pressure of 500 mTorr at a constant inflow of air is established.
  • Switch the plasma on at 27 W, which is the highest intensity available for the specified Plasma Cleaner.
  • ‘Turn off’ the plasma after 90 seconds.
  • Vent the chamber of the Plasma Cleaner by opening the needle valve and allowing air to enter through the flow meter.
  • Remove the activated piece of substrate from the Plasma Cleaner.

5. Bonding

  • Align the substrates (and thus enclose the micro fabricated structures) by firmly pressing the activated quartz surface to the aluminum sheet. A fine interfacial gap is forming between the quartz and aluminum surfaces.
  • In case you have a large chip or thin fragile substrates you may need to carefully clamp the substrates together.
  • Prime the gap with the adhesive while holding the two substrates of your device together. In order to do so, place a small drop of adhesive to one edge, i.e. to the gap between the two substrates. The adhesive will flow into the gap due to capillary action. Thick substrates will be held together sufficiently by the adhesive film. The flow of the adhesive will stop at the edge of the microfabricated structures as a results of surface effects (surface tension and wetting angle). Inspect whether the device is completely filled with the adhesive. Add more of the adhesive if necessary.
  • Cure the completely primed device by exposing it to UV-light, 365nm @ 100 W for 5 to 10 minutes.
  • Place the device into the oven at 50°C. According to the supplier’s specifications, the bond reaches its maximum strength after 12 hours at 50°C. Alternatively, for temperature-sensitive materials, longer incubation times at room temperature are also feasible.

The main advantages of the presented bonding method are as follows:

1. Hybrid microfluidic devices can be easily bonded.

2. The method is relatively simple and does not require clean-room conditions.

3. The method works with any UV transparent material as long as the surfaces are clean, smooth and as long as they can promote the capillary action necessary for the priming with adhesive.

4. It is an economic bonding method. An expected 30 mL of UV-curing adhesive should be enough for the bonding of over hundred microfluidic devices. Each assembly will thus cost less than £0.2 GBP (or approximately $0.3 USD).


What else should I know?

Q1. What processes do you use to create the holes in the quartz slide?

A1. The quartz slides are drilled with diamond drill bit (Eternal tools, UK), 1 mm in diameter, and a bench drill (D-54518, Proxxon , Germany) at 1080 rpm.  This is a slow operation as the process is closer to grinding rather than drilling. To avoid crack formations in the quartz slide and to cool diamond bit a droplet of water is applied on the surface of the quartz. After each cycle grinded quartz debris may be accumulating at the bottom of the hole; it can be removed by using a pipette and cooling liquid.

Q2. Have you ever tried this method with channel geometries that are disconnected? For example, a channel  layout shaped like an “O” that would prevent adhesive wetting from the edge of the slide?

A2. We had bonded successfully channels with complex, serpent geometries. For “O”-shaped channels we use additional feed, a hole, drilled in one of the substrates that allows the adhesive to spread.

Q3. Does the adhesive ever “burst” and enter the channels? If so, what methods do you use to minimize the chances of this happening?

A3. Yes, it happens occasionally that the adhesive fills the channel.

To prevent this: minimum amount of glue is applied at a time, and also the propagation of the front needs to be monitored. We wait until the glue reaches the channel edge, and then we place the assembly under the UV-light for curing.

If the channel is filled with small amount of adhesive, the glue could be washed out with a bit of ethanol or acetone.

Completely filled channel requires disassembly, cleaning with acetone or ethanol of the substrates. Afterwards, the procedure can be repeated with less glue.

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Chips & Tips Moderator – Glenn Walker

Glenn Walker

Glenn M. Walker received B.S. and M.S. degrees in biomedical engineering from Louisiana Tech University in 1996 and 1998 respectively, and his Ph.D. degree in biomedical engineering from the University of Wisconsin-Madison in 2002. From 2002 to 2004 he was a Postdoctoral Fellow in the Department of Molecular Physiology & Biophysics at Vanderbilt University Medical Centre. Since 2004 he has been with the Joint Department of Biomedical Engineering at the University of North Carolina and North Carolina State University, where he is currently an Associate Professor. His current research interests are in the areas of sub-visible particle characterization and microfluidic calorimetry.

Faculty webpage

Lab webpage

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