Chips and Tips

DIY peristaltic pump

26 Jun 2015

By RSC Internet Services.

Shannon Faley, Bradly Baer, Matthew Richardson, Taylor Larsen, and Leon M. Bellan*

Vanderbilt University, Department of Mechanical Engineering, Nashville TN, 37235, USA

*leon.bellan@vanderbilt.edu

Why is this useful?

Figure 1: Fully assembled peristaltic pump

The majority of microfluidic applications require an external pumping mechanism. Multi-channel, individually addressable pumps are expensive, often large, and prone to failure when operated inside cell culture incubators at 95% humidity. The number of experiments that can be run at a given time is limited by the availability and expense of pumps. Perfusing artificial tissue scaffolds containing engineered vasculature requires long-term (days to weeks) continuous flow at low rates. We designed an inexpensive (~$100 for 2 pumps, ~$70 for each additional set of 2 pumps) peristaltic pumping system using an Arduino- controlled stepper motor fitted with a custom 3D-printed pump head and laser-cut mounting bracket. Each pump has a footprint roughly that of the NEMA 17 stepper motor and is easily controlled individually using open source software. Up to 64 motor shields can be stacked for a given Arduino Uno R3, each capable of supporting two stepper motors, and thus has the expansion potential to control 128 pumps in parallel. We have successfully implemented two stacked motor shields driving four independent stepper motors. Flow rate is dependent upon both tubing diameter and step rate. We found flow rates to range between ~50-250 μl/min for 1/16” tubing and ~500-1500 μl/min for 1/4″ tubing. We anticipate that this pump design will likely prove more resilient to incubator humidity compared to standard peristaltic pump powered by DC motors. Since implementation, these pumps have functioned without fail for 3 months (intermittent) under humid conditions. In the event of failure, however, cost of motor replacement is an economical $14.

Figure 2

What do I need?

Materials:

Nema 17 stepper motor ($14, spec, vendor)
Arduino Uno R3 Controller ($25, spec, vendor)
Arduino Motor Shield ($20, spec, vendor)
M3 machine screws (4) & hex bolts (4) ($1, McMaster-Carr)
DB9 Male & Female Solder Connectors ($9, StarTech)
18AWG 4C speaker cable ($10, Monoprice)
Spring steel
ABS Filament
6-32 machine screws & square nuts (3) ($1, McMaster-Carr)

Equipment:

3D printer
Laser/Metal cutter
Soldering iron & solder
Butane torch

What do I do?

Pump head fabrication:

Using ABS filament, 3D print pump head from file pumphead.crt.9
Cut three 15 mm (length) sections from rigid ¼” tubing to serve as rollers.
Use the three 6-32 machine screws and square nuts to assemble the tubing to pump head as shown in Figure 3.

Figure 3

Mounting bracket fabrication:

Using bracket template file (2000 Pump Mount v4) and laser cutting facilities, produce a mounting bracket from spring steel, or other appropriate metal. Note that the score line bisecting the bracket is intended to be cut at a lower power. This line is just a marker to show where to bend the bracket in the following step.
Using handheld butane torch, heat mounting bracket along score line and bend with pliers. Repeat until mounting bracket forms a right angle (see Figure 1).

Motor Electrical Wiring: (see figure 4 for example orientation)

Solder motor wires to DB9 Male Connector
Solder one end of speaker wire to DB9 Female Connector
Connect opposite end of speaker wire to Arduino Motor shield

Figure 4 Example of connection scheme by wire color

Pump Assembly:

Use M3 machine screws to attach mounting bracket to stepper motor, with corresponding hex nuts as spacers between motor and bracket.
Press fit pump head onto rotor shaft.
Connect motor to Arduino using DB9 connectors

Arduino/Motor Shield Assembly:

Follow assembly instructions provided by adafruit.com (https://learn.adafruit.com/adafruit-motor-shield-v2-for-arduino/stacking-shields). See also Figure 5.

Figure 5

Computer Control:

1. See online resources for easy starter code (https://learn.adafruit.com/adafruit-motor-shield-v2-for-arduino/install-software)

2. Load example code to control 2 stacked motor shields running four independent pumps simultaneously. (foursteppers_v2.ino)

3. Start pumping! See video clip for multi-pump demonstration:

Please click here to download the DIY Peristaltic Pump Files

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Low Temperature Melting Metal Solders For Electrical Interconnects On Plastics

25 Jun 2015

By Liz Bowley.

Abdul Wasay, Dan Sameoto
Department of Mechanical Engineering, University of Alberta, Edmonton, T6G 2R3, CANADA.
Email: sameoto@ualberta.ca

Why is it useful?

Electrodes have been used for driving electrochemical reaction on various Lab on chip applications, running electroosmotic pumping or simple sensing of current or voltages. With the recent thrust to move to thermoplastics, simple electrodes have been patterned at low temperatures using vacuum plasma sputtering via intermediate masks¹ or ink electrodes². While, wire-bonding or soldering have been the standard protocols for providing interconnections to the bond pads on silicon or glass chips, the temperatures in these cases are often beyond the glass transition or even melting temperatures of most thermoplastic substrates, which leads to defects and ineffective connection. Other techniques such as conductive epoxies can be problematic due to longer cure times, permanent fixtures and potentially low uniformity of conductive properties which are affected by sintering times and temperatures.

We present a simple soldering technique using low temperature melting metals that is compatible with nearly any thermoplastic substrate and thin-film electrodes.

Fig.1. Basic Apparatus

Field’s metal (eutectic alloy of 32.5% Bi, 51% In, 16.5% Sn) preferred for low toxicity, although lead based alternatives exist.
Syringe connected to tygon tube
Copper wire
Aluminum foil dish
Electrodes patterned on thermoplastic substrate (here, polystyrene with ~ 20 nm thick Au electrodes)

Tools

Hot Plate
Shearing scissor
Hot glue gun (in this case a Mastercraft dual temperature glue gun, operated at low temperature setting)

What do I do?

To extrude the Fields metal into a thin filament, (Fig.2)

melt the Fields metal (ROTO144³; T_m=62.2^oC, typical resistivity ~520nΩ-m) in an aluminum foil dish. The metal is very soft, so bolt cutters can easily remove smaller portions for melting if desired.
use a suitable diameter and length tygon tube and pull the molten metal into it using a syringe – it will freeze within a few inches (1/32” inner diameter tube shown here).
allow it to cool to room temperature (less than a minute) and then pull out the filament.

Soldering Process,(Fig.3.)

Hold the Field’s metal filament with a hot glue gun used for lower melting point glues (measured tip temperature ̴120 C) over the electrode patterned petri dish and melt it over the electrodes.
Solder the copper wire with the Field’s metal. After soldering, excess metal can be removed from the hot glue gun tip with paper towel.

The resultant solder is strong enough to be used for mainstream lab on chip applications. They can be removed by a strong tug, under which case the thin electrodes on plastics could be stripped.

Fig.2. Fields metal extrusion process

Fig.3. Soldering process

Fig.4. Gecko adhesives based reversibly bonded4 capillary electrophoresis device with integrated electrodes

Fig.5. a) Demonstration of solder strength b) small electrode stripping as observed when the solder is removed by a strong tug

What else should I know?

While there are quite a few options of low melting temperature metals to choose from, most of them contain heavy metal like lead or cadmium, which may ideally be avoided. While Field’s metal is relatively expensive due to the indium content, the solders can be recovered for reuse in the lab and in general is a better option for electrically connecting thin-film electrodes to temperature sensitive substrates in a quick, reliable fashion.

Watch the video here: Low Temperature Melting Metal Solders For Electrical Interconnects On Plastics

References:

1 A. Toossi, M.Sc, University of Alberta, 2012.
2 C. E. Walker, Z. Xia, Z. S. Foster, B. J. Lutz and Z. H. Fan, ELECTROANALYSIS, 2008, 20, 663-670.
3 http://www.rotometals.com/product-p/lowmeltingpoint144.htm
4 A. Wasay and D. Sameoto, Lab Chip, 2015, (DOI:10.1039/C5LC00342C).

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Micromilling Techniques II: Tool Alignment

10 Jun 2015

By Kate Bandoo, Publishing Assistant.

David J. Guckenberger¹, Theodorus E. de Groot¹, Alwin M.D. Wan², David J. Beebe¹ and Edmond W. K. Young^2,*

¹Department of Biomedical Engineering, Wisconsin Institutes for Medical Research, University of Wisconsin-Madison, Madison, WI, USA

²Department of Mechanical and Industrial Engineering, University of Toronto, 5 King’s College Road, MC313, Toronto, ON, Canada.

* Corresponding Author, E-mail: eyoung@mie.toronto.ca; Tel: +1(416) 978 1521

Why is this useful?

Micromilling is a highly efficient method for fabricating microfluidic devices directly in polymeric materials like thermoplastics. Please see the review article by Guckenberger and co-workers for a primer on micromilling.¹ After securing your workpiece to the milling table,² the next step in the milling process is to align the tool to the workpiece, thereby defining the coordinate origin. Many high-end mills designed for micromilling have automated tool alignment systems. However, lower cost mills that are also capable of milling microdevices may not have tool alignment systems, and the user must therefore manually align the tool to the workpiece to the desired accuracy. Here we present several alignment techniques that are low cost and can be performed by minimally trained users. We divide tool alignment into two separate directions: the vertical z-axis direction, and the planar x-y plane direction.

What do I need?

CNC Mill (PCNC 770, Tormach)
Tool, e.g. endmill of choice
Workpiece (properly secured to milling table, see ref. [2])

Specific tooling or materials required for each technique are referenced separately below, under the “Tooling Note”.

What do I do?

Z-axis Direction

Fig. 1. Four techniques for aligning the tool to the workpiece in the z-axis (from left to right): (i) reflection, (ii) chip, (iii) paper, and (iv) collet technique.

(i) Reflection Technique

Tooling Note: This technique requires a reflective surface (e.g., transparent materials such as PS or PMMA), but otherwise does not require any specific tooling.

Step 1: Start with the tip of the tool slightly above the workpiece, with the spindle turned off.
Step 2: Looking from a near planar location with respect to the workpiece, lower the tool until the tool itself comes in contact with its reflection.
Step 3: Set this location as z = 0.

Tip: Placing a piece of paper behind the tool will improve contrast, making it easier to identify when the tool contacts the reflection. A magnifying glass can be used to improve visibility.

(ii) Chip Technique

Tooling Note: This technique does not require any specific tooling.

Step 1: Start with the tool slightly above the surface with the spindle running (i.e., the tool should be rotating).
Step 2: Lower the tool towards the surface until either (a) a chip is observed, (b) a mark is made on the surface, or (c) a sound is made from the tool cutting the material.
Step 3: Set this location as z = 0.

Tip: This method works best for large and flat endmills, and can be more difficult with small endmills or any tool that is pointed or round at the tip. Note that this is a physical contact method, and will blemish the surface.

(iii) Paper Technique

Tooling Note: This technique requires a small piece of paper of known thickness.

Step 1: Start with the tool above the surface with the spindle turned off.
Step 2: Place a piece of paper (with known thickness) between the tool and the workpiece.
Step 3: While moving the piece of paper back and forth, lower the tool in stepwise increments.
Step 4: Continue lowering the tool until it causes resistance to the sliding piece of paper.
Step 5: Set this location as z = the thickness of the paper (e.g., .003”)

Tip: Practice more to become comfortable with identifying when the endmill comes in contact with the surface.

(iv) Collet Technique

Tooling Note: This technique requires an ER20 tool holder (#31829, Tormach) from the Tormach Tooling System (TTS) and an ER20 1/8” collet (#30112, Tormach).

Step 1: Place tool in collet and secure using the setscrew on the side.
Step 2: Without the spindle running, lower the tool until it is just above the surface.
Step 3: Loosen the setscrew and allow the tool to fall into contact with the workpiece.
Step 4: Tighten the setscrew.
Step 5: Set this location as z = 0.

X-Y Plane Direction

Fig 2. Techniques for aligning the tool in the x–y plane. (A) Illustrations of the (i) edgefinder, (ii) chip, and (iii) paper techniques. (B) Guide to offsetting a tool (left) and finding the center of an object (right).

(i) Edgefinder Technique

Tooling Note:
This technique requires an edgefinder (e.g., #02035186, MSC Industrial Supply).

Step 1: Place edgefinder in collet and start spindle (1000 rpm works well with the edgefinder).
Step 2: Deflect the tip of the edgefinder so that it lies eccentric to its initial axis.
Step 3: Starting with the x-axis, move the edgefinder toward a perpendicular surface. The tip of the edgefinder will become concentric upon contact with surface, and then return to an eccentric position immediately afterward. This sudden “jump” in eccentricity marks the edge.
Step 4: Set the current location to either plus or minus the radius of the tip. See Fig. 2B for more details on deciding a positive or negative bias.

Chip Technique

Tooling Note: This technique does not require any specific tooling.

Step 1: Start with the tool near the face of interest with the spindle running (i.e., the tool should be rotating).
Step 2: Step the tool towards the surface, until (a) a chip is observed, (b) a mark is made on the surface, or (c) a sound is made from the tool cutting the material.
Step 3: Set this location as (plus or minus) the radius of the endmill.

Tip: This method works best for large diameter endmills. Note that this is a physical contact method, and will blemish the surface.

Paper Technique

Tooling Note: This technique requires a small piece of paper of known thickness.

Step 1: Start with the tool near the face of interest with the spindle running
Step 2: Gripping gently with thumb and fore-finger, or by pressing it against the surface place a piece of paper between the tool and the surface.
Step 3: Step the tool towards the surface until the tool pulls the paper.
Step 4: Set this location as (plus or minus) the sum of the endmill radius and paper thickness.

Tip: This method works best for large endmills.
Caution: Be sure to keep fingers clear of the cutting tool.

References

1. Guckenberger DJ, de Groot T, Wan AMD, Beebe DJ, Young EWK, “Micromilling: A method for ultra-rapid prototyping of plastic microfluidic devices”, Lab on a Chip, DOI:10.1039/c5lc00234f (2015).

2. Guckenberger DJ, de Groot T, Wan AMD, Beebe DJ, Young EWK, “Micromilling Techniques I: Securing Thin Plastic Workpieces for Precise Milling”, Chips & Tips (2015).

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Micromilling Techniques I: Securing Thin Plastic Workpieces for Precise Milling

10 Jun 2015

By Kate Bandoo, Publishing Assistant.

David J. Guckenberger¹, Theodorus E. de Groot¹, Alwin M.D. Wan², David J. Beebe¹ and Edmond W. K. Young^2,*

¹Department of Biomedical Engineering, Wisconsin Institutes for Medical Research, University of Wisconsin-Madison, Madison, WI, USA

²Department of Mechanical and Industrial Engineering, University of Toronto, 5 King’s College Road, MC313, Toronto, ON, Canada.

* Corresponding Author, E-mail: eyoung@mie.toronto.ca; Tel: +1(416) 978 1521

Why is this useful?

Micromilling is a highly efficient method for fabricating microfluidic devices directly in polymeric materials like thermoplastics. A recent review article highlights the use and relevance of micromilling in the field of microfluidics.¹While milling is most popular among machinists, the technique is becoming cheaper and more accessible to many others. Thus, there is a need for disseminating technical know-how for achieving quality micromilled parts. One common technical issue is ensuring flat work surfaces, and eliminating warping and bending of thin plastics during micromilling, which can lead to uneven milling and large errors in feature dimensions. Here we present a simple technique to achieve flat work surfaces – leveled to within 40 μin / in (.04 μm / mm)) – and properly secure thin plastic workpieces to minimize warping. Please refer to the review article by Guckenberger and co-workers for a primer on micromilling

What do I need?

Tools:

CNC mill (PCNC 770, Tormach)
3/8” set screw holder (#31820, Tormach)
Granite block (6” x 12” x 2” Grade AA, Standridge)
T-slot clamps (#32580, Tormach)
Drop test indicator (#24-315-4, SPI)
Wrench (choose appropriate size for clamps)

Materials:

Plastic workpiece (e.g., polystyrene, cyclo-olefin copolymer (COC), PMMA)
Surface protection tape (#6092A24, McMaster-Carr)
Two-sided or transfer tape (#9472LE, 3M)
Silicone rubber, 1/16” thick (#8632K41 – Durometer Hardness: 40A, McMaster-Carr)

Fig. 1 Tooling and Materials. (A) Hardware necessary for setting up the granite block. Assembly of the T-slot clamps is shown in Fig. 2. (B) (Top) Components needed for the drop test indicator: (i) collet/tool holder, (ii) aluminum adapter, (iii) drop test indicator, (iv) screw to attach indicator to the adapter. (Bottom) Side view of the assembled components. (C) Materials needed for adhering workpiece to the block: (i) surface protection tape, (ii) silicone rubber, (iii) transfer tape, (iv) polystyrene sheet.

What do I do?

Preparation: The dial indicator must be affixed to the head of the mill. To do so, mounting brackets can be purchased or custom-made, as we have done using a block of aluminum. This aluminum serves as a simple adapter between the dial indicator and the collet.

Step 1: Place granite block on the milling table. Note that a stiff yet compliant material is necessary between the block and the milling table to allow for minor adjustments. The particle-board feet affixed to the granite block by the manufacturer (Standridge) are sufficient.

Step 2: Assemble the clamps (Fig. 2A), and position the clamps in either a 3-clamp (Fig. 2B) or 4-clamp (Fig. 2C) configuration. The front of the clamp (i.e., the portion that contacts the granite block) should be the same height as or lower than the rear of the clamp.

Step 3: Hand-tighten each clamp, then tighten an addition 1/8^th turn using a wrench.

Step 4: Level the granite block by using the drop test indicator to measure the height across the entire block. The following steps are recommended:

a)      Lower the indicator onto the center of the block.
b)      Move the indicator along x-axis to the two ends of the block, and note the height of both ends.
c)      Position the indicator on the high side of the block and tighten the clamps until the height matches the low side.
d)     Repeat steps (b) and (c) for the y-axis.
e)      Repeat steps (b) to (d) until the block is level to desired tolerance.
f)       Tip: When using the 4-clamp configuration, clamps should be equally tightened in pairs to maintain levelness in the other direction.

Fig. 2 (A) T-slot clamp setup configuration. (B) Granite block configuration using three clamps. (C) Granite block configuration using four clamps.

Step 5: (Optional) Apply a protective adhesive film to the workpiece. This will simplify removal of the transfer tape from the workpiece (see Video).

a)      Peel a piece of protective adhesive film from the roll of film.
b)      Place a piece of silicone rubber on the roll of tape or other cylindrical roll.
c)      Place the protective adhesive film on top of the silicone rubber with the adhesive side face up, assuring the tape is flat.
d)     Roll the workpiece onto the piece of tape. The silicone rubber will provide compliance below the tape to prevent bubbles in the tape.
e)      Press the tape down to make sure it is fully adhered to the workpiece.

Step 6: Apply 2-sided tape or transfer tape to the workpiece

Step 7: Stick the workpiece onto the granite block.

Step 8: (Optional) Use the drop test indicator again to ensure the workpiece is flat.

References

1. Guckenberger DJ, de Groot T, Wan AMD, Beebe DJ, Young EWK, “Micromilling: A method for ultra-rapid prototyping of plastic microfluidic devices”, Lab on a Chip, DOI:10.1039/c5lc00234f (2015).

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A simple and inexpensive device to remove edge beads

02 Mar 2015

By Kate Bandoo, Publishing Assistant.

Markus Ludwig and Seth Fraden

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

Why is this useful?

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

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

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

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

What do I need?

Materials:

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

Tools:

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

What do I do?

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

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

Figure 1: Materials needed

Figure 2: Assembled device

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

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

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

Figure 4: Comparison of feature resolution

CAD drawing of a multi layer design

Figure 5: SU8 master fabricated with edge bead

Figure 6: SU8 master fabricated without edge bead

Video 1:

Video 2:

References

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

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

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

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A simple and low cost method to fabricate NOA microfluidic chips

09 Jan 2015

By Kate Bandoo, Publishing Assistant.

Gabriele Pitingolo¹, Raffaele Vecchione^1,3 and Paolo A. Netti^1,2,3

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

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

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

Why is this useful?

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

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

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

What do I need?

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

What do I do?

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

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

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

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

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

References

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

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

08 Dec 2014

By Liz Bowley.

Bruno Rafael Becker^{1, 2}, Tristan Sun¹, Shengjie Zhai¹ and Hui Zhao¹

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

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

Why is this useful?

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

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

What do I need?

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

What do I do?

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

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

2. Repeat the step 1 with isopropyl alcohol.

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

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

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

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

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

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

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

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

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

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

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

What else should I know?

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

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

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

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

Experiments with the fabricated gold electrodes

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

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

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

02 Dec 2014

By Liz Bowley.

David Caballero^1,2, Josep Samitier^1,2

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

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

Why is this useful?

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

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

What do I need?

Figure 1

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

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

You will also need:

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

What do I do?

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

STRATEGY A:

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

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

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

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

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

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

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

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

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

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

STRATEGY B:

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

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

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

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

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

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

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

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

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

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

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

What else should I know?

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

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

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

Acknowledgements

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

References

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

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

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

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Reservoir Poly(dimethylsiloxane) Cap Fabrication

23 Oct 2014

By Liz Bowley.

Luis G. Rigat-Brugarolas^1,2, Antoni Homs-Corbera^1,2 and Josep Samitier^1,2,3

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

Why is this useful?

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

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

What do I need?

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

What do I do?

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

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

The steps are detailed next:

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

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

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

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

Fig 5: PDMS cap replica

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

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

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

References

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

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

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

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Welcome to Chips & Tips

23 Oct 2014

By Liz Bowley.

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

Do you

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

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

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

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

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

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Chips and Tips

DIY peristaltic pump

Low Temperature Melting Metal Solders For Electrical Interconnects On Plastics

Micromilling Techniques II: Tool Alignment

Micromilling Techniques I: Securing Thin Plastic Workpieces for Precise Milling

A simple and inexpensive device to remove edge beads

A simple and low cost method to fabricate NOA microfluidic chips

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

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

Reservoir Poly(dimethylsiloxane) Cap Fabrication

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