This work was supported by the University of Utah office of the Vice President for Research, as well as by funds in conjunction with grant P30 CA042014 awarded to the Huntsman Cancer Institute and to the CRR Program at the Huntsman Cancer Institute.

1. Fabrication of the Polydimethylsiloxane Negative
<ol>
	<li>Preparation of polydimethylsiloxane
	<ol>
		<li>Measure 18 g of PDMS silicone elastomer and 1.8 g of the curing reagent. Pour the curing reagent into a measuring boat containing the elastomer.</li>
		<li>Mix the elastomer and the curing reagent vigorously for 1 min and put the mixture into a vacuum chamber for 30 min.</li>
		<li>Remove the PDMS from the vacuum, pour 15 g onto the negative using a cookie cutter (radius = 3.8 cm) to keep the PDMS from running off the side, and cover the remaining PDMS mixture.</li>
		<li>Put the mold containing the PDMS into the vacuum chamber for 10 min.</li>
		<li>Remove the mold containing the PDMS from the vacuum chamber, place it on a hot plate set to 150 &#176;C for 60 min to cure, and cover it with aluminum foil.</li>
		<li>Pour the remaining PDMS onto an inverted glass Petri dish (10 cm in diameter), using another cookie cutter as a mold, to create a pad of PDMS. Put this into the vacuum chamber until the PDMS negative is completely cured.</li>
		<li>Transfer the Petri dish with the PDMS from the vacuum to the preheated hot plate to cure at 150 &#176;C for 60 min.</li>
	</ol>
	</li>
	<li>Plasma surface treatment of polydimethylsiloxane
	<ol>
		<li>Insert a sharp razor blade underneath the PDMS and the cookie cutter to carefully remove the PDMS from the negative.</li>
		<li>Use a sharp hobby knife to cut out the negative from the larger piece of PDMS; it should be cut with enough buffer area to attach another piece of PDMS on top of it without obstructing the negative.</li>
		<li>Insert a sharp razor blade underneath the cured PDMS with the negative to carefully remove the PDMS from the Petri dish.</li>
		<li>Use a hobby knife to cut out a piece of PDMS from the pad that is the same size as the piece cut for the negative. Then, cut a rectangle out of the new piece of PDMS with enough room to fit it around the microchannel when it is placed on top of the negative.</li>
		<li>Put both pieces of PDMS (the negative and the rectangle) onto a flat substrate (such as a slide made from glass, quartz, polystyrene, or other material) and insert them into a radio frequency (RF) plasma cleaner. Close the door and seal the chamber by evacuating the air using a vacuum pump. Inject air up to a pressure of 400 mTorr using a digital vacuum pressure controller. 
		NOTE: If the substrate used for the plasma cleaning has not been used in the plasma cleaner, put it into the plasma cleaner by itself for 60 s prior to treating the negative, in order to prevent the acrylic base layer from sticking to the substrate.</li>
		<li>Turn the RF setting to high, and a pink air plasma should appear in the viewing window. Plasma-treat the two pieces of PDMS for 30 s, then turn the RF setting off. Allow air to slowly reenter the chamber in order to prevent a turbulent air flow from disturbing the contents of the chamber.</li>
		<li>Remove the PDMS from the plasma cleaner and place it on a benchtop. Then, carefully invert the PDMS rectangle over the negative and press it down with a pair of forceps. Let it rest for 10 min to allow the two pieces of PDMS to adhere to each other.</li>
	</ol>
	</li>
	<li>Fluorosilane surface treatment of the polydimethylsiloxane negative
	<ol>
		<li>Use a dropper to put 2 drops of trichloro(1H,1H,2H,2H-perfluoro-octyl)silane (PFOTS) into a small weigh boat and insert it into a glass vacuum chamber.</li>
		<li>Invert the PDMS onto a piece of aluminum foil (so that the microchannel negative faces upward) and put it into the glass vacuum chamber. Then, evacuate the chamber continuously for 24 h.</li>
	</ol>
	</li>
</ol>
2. Fabrication of the MY133 Microchannel
<ol>
	<li>Forming the MY133-V2000 microchannel structure
	<ol>
		<li>Fill the PDMS negative with 400 &#181;L of MY133-V2000. 
		NOTE: The negative must be slightly overfilled.</li>
		<li>Place the MY133-V2000-filled PDMS negative into the vacuum chamber for 2 h to remove any bubbles.</li>
		<li>Remove MY133-V2000 from the vacuum and slowly press a glass slide against the top of the slightly overfilled negative to create a flat surface of MY133-V2000 and to prevent oxygen from inhibiting the polymerization. 
		NOTE: PDMS, at the channel surface, will partially inhibit the polymerization, enabling bonding in later stages. The glass slide has a slightly reduced UV transparency (35%) compared to quartz, which helps to prevent any yellowing of the cured device.</li>
		<li>Insert MY133-V2000 into a 400 W UV oven and set the UV radiation to 50% of the maximum intensity for 300 s&#160;to cure the MY133-V2000 microchannel. 
		NOTE: The peak wavelength used to cure the device is approximately 375 nm with a bandwidth of about 25 nm. Approximately 4,500 mJ/cm2 of power was used to cure the microchannel, which is slightly more than double the minimum curing power recommended by the manufacturer.</li>
	</ol>
	</li>
	<li>Building the acrylic holder
	<ol>
		<li>Draw the acrylic base layer using a vector drawing software. Make sure the base layer is a rectangle that is 1.5 mm thick, 75 mm long, and 25 mm wide, with a centered rectangle that measures 25 mm x 11 mm.</li>
		<li>Draw the acrylic mid-layer using a vector drawing software. Make sure the mid-layer is a rectangle (1.5 mm thick, 75 mm long, and 25 mm wide) with a centered square (5 mm x 5 mm) and two circles (both with a diameter of 3 mm) separated by 15 mm.</li>
		<li>Draw the acrylic top layer using a vector drawing software. Make sure the top layer is a rectangle (3 mm thick, 30 mm long, and 25 mm wide) with a centered square (5 mm x 5 mm) and two circles (both with a diameter of 3 mm) separated by 15 mm.</li>
		<li>Cut out the device designs created in the vector drawing software with a laser cutter. Use the sample settings of 100% power and 30% speed. Run the program twice to ensure that the acrylic is cut out.</li>
		<li>Tap the holes in the acrylic top layer using a size M5 0.80-mm tapping tool.</li>
		<li>Wipe the acrylic pieces with acetone to remove any residual marks or burns.</li>
	</ol>
	</li>
	<li>Bonding of MY133-V2000 in the acrylic holder
	<ol>
		<li>Attach the base layer of the acrylic to the glass substrate using an adhesive, such as cyanoacrylate super glue. Place two small drops on the edges of the acrylic and then use a disposable tool to evenly spread the glue. Place the glass substrate onto the acrylic and allow the glue to dry using the weight of the glass to hold it in place.</li>
		<li>Coat the exposed glass with 100 &#181;L of PDMS using a positive displacement pipette; then, insert the base layer into a vacuum spin coater and set it to 1,500 rpm for 2 min to evenly coat the glass with a PDMS film approximately 10-&#181;m thick.</li>
		<li>Remove the base layer from the spin coater and carefully wipe away any excess PDMS that coated the acrylic with acetone and paper towel. Be careful not to disturb the uncured PDMS.</li>
		<li>Bake the base layer with the PDMS in an oven at 65 &#176;C for 2 h to cure the PDMS.</li>
		<li>Pipette 1 mL of PDMS onto a glass slide and, then, use another glass slide to evenly spread the PDMS until it starts to ooze out the sides. Cure this on a hot plate at 150 &#176;C for 10 min.</li>
		<li>Cut the cured PDMS into a rectangle with the same dimensions as the MY133-V2000 device and, then, using the mid-layer of the acrylic as a mold, punch holes for the reservoir and cut a square viewing window in the PDMS to make the PDMS gasket.</li>
		<li>Place the MY133-V2000, channel side up, on the same or a similar flat substrate as used for plasma-treating the PDMS in step 1.2.5. Place the acrylic base layer containing the cured PDMS substrate alongside the MY133-V2000 channel in the O2 plasma cleaner. Set the vacuum pressure to 200 mTorr and the RF level to high and surface-treat the channel and glass substrate for 30 s.</li>
		<li>Using forceps, immediately place the MY133-V2000 channel side-down in the rectangular cutout of the base layer acrylic such that the MY133-V2000 contacts the PDMS.</li>
		<li>Place the PDMS gasket on top of the MY133-V2000 device, lining up the holes in the gasket with the reservoirs in the device.</li>
		<li>Place the unfinished device onto a raised platform above the bench to provide a clamping surface for device assembly.</li>
		<li>Extract 3 mL of acrylic cement using a syringe. Distribute enough acrylic cement on top of the base layer to provide a thin coating. Make the coat as even as possible and do not let the material seep into the channel.</li>
		<li>Place the mid-layer acrylic piece on top of the base layer acrylic. Make sure that the holes line up with the reservoirs of the MY133-V2000 channel.</li>
		<li>Use 3 small clamps to hold the base layer and mid layer together as tightly as possible for 2 min while the acrylic&#160;cement hardens, to bond the pieces of acrylic together. Ensure that the reservoirs are not obstructed during this step to prevent air from being trapped underneath the MY133-V2000 device.</li>
		<li>Remove the pressure from the device and place the acrylic cement on the mid-layer in the places of anticipated contact with the top layer of the acrylic.</li>
		<li>Place the top layer of the acrylic on the mid-layer piece of the acrylic, ensuring that the holes are aligned with the reservoirs, and allow it rest on the bench for 2 min while the acrylic cement dries.</li>
	</ol>
	</li>
</ol>
3. Testing and Use of the MY133-V2000 Device
<ol>
	<li>Leak testing
	<ol>
		<li>Test the MY133-V2000 device by adding 10 &#181;L of food dye or deionized water into one of the reservoir holes to test for the adhesion and flow.</li>
		<li>Insert a tube (with a 1/8-in outer diameter) into the reservoir and connect the other end to a vacuum trap. Turn the vacuum on to pull the dye or water through the channel to ensure that a successful device is created.</li>
		<li>Check under a microscope to verify that there are no leaks in the channel or reservoir, fill the reservoir with ethanol, and allow the ethanol to sit at room temperature for 10 min. Then, pull the ethanol through using the vacuum, to clean the dye or water out of the channel.</li>
		<li>Spray the channel with ethanol and put it into a sterile polystyrene dish. Wrap it tightly with parafilm and store it in a sterile environment until needed. 
		NOTE: At this point, the device must be handled aseptically in order to prevent contamination. When ready to use the device, the outside of the container should be sterilized using ethanol and brought into a biosafety cabinet prior to opening the container.</li>
	</ol>
	</li>
	<li>Plating MCF7 cells in the MY133-V2000 device
	<ol>
		<li>Grow MCF7 cells on 10-cm Petri dishes in a complete media consisting of Dulbecco&#39;s minimum essential medium (DMEM), supplemented with 10% fetal bovine serum, penicillin-streptomycin, glutamine, and non-essential amino acids, in a cell culture incubator that maintains a humid atmosphere containing 5% CO2 and 20% O2 at a temperature of 37 &#176;C. Grow the cells to approximately 80% confluence before seeding them in the microchannel.</li>
		<li>First, spray the polystyrene dish containing the microchannel with 70% ethanol. Then, bring it into the biosafety cabinet before removing the parafilm protecting the channel from the ambient laboratory environment.</li>
		<li>Immobilize human-plasma-derived fibronectin on the microchannel surface by diluting it in Dulbecco&#39;s phosphate-buffered saline (DPBS) to 10 &#181;g/mL. Then, inject 20 &#181;L of this solution into the channel and incubate it at 20 &#176;C for 45 min.</li>
		<li>Wash the fibronectin solution out of the channel by injecting 20 &#181;L of cell culture media into the channel, and allow it to incubate at 20 &#176;C for 10 min.</li>
		<li>Wash the cells with 10 mL of DPBS, and trypsinize the cells by adding 1 mL of 1x trypsin. Incubate the cells at 37 &#176;C for 7 min. Neutralize the trypsin with 9 mL of complete media.</li>
		<li>Inject 20 &#181;L of the cells into the channel and incubate them at 37 &#176;C for 45 - 120 min to allow the attachment to the substrate prior to imaging.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

MY133-V2000 can be used as an alternative to traditional soft lithography fabrication materials such as PDMS. Previous work has shown that materials with a high index of refraction, such as PDMS, introduce significant artifacts near the channel walls due to the mismatching indices of refraction between the fabrication material and the aqueous solution inside the channel13. MY133-V2000 enables matching the refractive index of the microfluidic device to the aqueous solutions commonly used in biomedi...

The development of microfluidic technology has enabled a wide range of new biomedical techniques that leverage the unique physics of microscopic-scale flows1,2. This includes the diagnostic techniques built on microfluidic platforms that quantify clinically relevant biomarkers, including cell stiffness3, surface markers4, and growth5. By manipulating single cells, microfluidic devices can also be used to measure biomarker heterogeneity, for example as an indicator of malignancy6. The ability to combine microfluidic applications with microscopy has further increased the utility of these platforms by allowing for devices that measure multiple biomarkers simultaneously7.
QPM is a microscopy technique that measures the phase shift as light passes through and interacts with the matter inside transparent samples. The mass of individual cells can be calculated from QPM measurements, by using the known relationship between the refractive index and the biomass density8,9. Previous work has shown that QPM is capable of measuring clinically relevant parameters such as cell growth10,11 and cell mechanical properties via disorder strength12. When combined with microfluidics, QPM can potentially be used to measure cell behavior in a highly controlled environment in vitro. One of the primary challenges facing combining QPM with microfluidics is the high refractive index of most polymers used to construct microfluidic channels via soft lithography13.
An important challenge facing the combination of microfluidics with various microscopy techniques is the mismatch between the refractive index of the device material relative to the refractive index of water14,15. One method to address this is through the use of a low refractive index polymer such as CYTOP16 or MY133-V200013. The latter is a fluorinated ultraviolet (UV)-curable acrylate polymer that has a refractive index similar to water (n = 1.33) and that is compatible with soft lithography techniques, allowing for a smooth integration into many established microfluidic device fabrication workflows. This makes MY133-V2000 not only suitable for microfluidic device fabrication, but also allows it to be readily combined with QPM and other microscopy approaches, to measure cell behavior both at the colony and on a single-cell scale. MY133-V2000 eliminates artifacts due to phase unwrapping by producing little, if any, phase shift as light passes through the water-MY133 interface.
Although eliminating the mismatch in refractive index, one major challenge associated with the devices fabricated from fluorinated polymers, such as MY133-V2000, is the low adherence to other materials such as glass or PDMS. The present work demonstrates the fabrication of an MY133-V2000 microfluidic device using soft lithography. Using O2 plasma to treat the surface of both the channel and the PDMS substrate combined with a custom-fabricated acrylic holder ensures that the device adheres to the substrate, creating a sealed channel. This device is suitable for cell culture and QPM to measure the mass of cells in the channel, which has important applications for measuring the growth of live cells and the intracellular transport of cell biomass, both of which have clinical relevance in diagnostic medicine and drug discovery.

<table><tbody><tr><td>MY133-V2000</td><td>MY Polymers</td><td>MY133-V2000</td><td></td></tr><tr><td>Sylgard 184</td><td>Ellsworth Adhesives</td><td>184 SIL ELAST KIT 0.5KG</td><td></td></tr><tr><td>Fisher Premium microscope slides</td><td>Fisher Scientific</td><td>12-544-4</td><td></td></tr><tr><td>.118&quot;(3.0mm) x 12&quot; x 12&quot; Acrylic Sheet</td><td>United States Plastic Corp</td><td>44290</td><td></td></tr><tr><td>.060&quot;(1.5mm) x 12&quot; x 12&quot; Acrylic Sheet</td><td>United States Plastic Corp</td><td>44200</td><td></td></tr><tr><td>SCIGRIP 3 Very Fast Set Acrylic Cement</td><td>United States Plastic Corp</td><td>45735</td><td></td></tr><tr><td>Standard Aluminum Foil (.6 mm thick)</td><td>VWR</td><td>89107-726</td><td></td></tr><tr><td>Kim Wipes</td><td>Fisher Scientific</td><td>06-666</td><td></td></tr><tr><td>Insta-Cure+ Super Glue</td><td>Bob Smith Industries</td><td>BSI-109</td><td></td></tr><tr><td>1/8&quot; PVC tubing</td><td>McMaster Carr</td><td>5231K55</td><td></td></tr><tr><td>McCormick Food Coloring</td><td>Target</td><td>13353207</td><td></td></tr><tr><td>X-Acto #1 Precision Knife</td><td>X-Acto</td><td>X3201</td><td></td></tr><tr><td>X-Acto #18 Heavyweight wood chiseling blade</td><td>X-Acto</td><td>X218</td><td></td></tr><tr><td>VWR Razor Blades</td><td>VWR</td><td>55411-055</td><td></td></tr><tr><td>Surface Treated Cell Culture Dishes</td><td>Fisher Scientific</td><td>FBO12922</td><td></td></tr><tr><td>Fibronectin Human Plasma</td><td>Sigma-Aldritch</td><td>F0895-1MG</td><td></td></tr><tr><td>Trypsin-EDTA 10x</td><td>Fisher Scientific</td><td>15-400-054</td><td></td></tr><tr><td>Corning Dulbecco&#039;s Phosphate Buffered Saline</td><td>Fisher Scientific</td><td>MT21030CM</td><td></td></tr><tr><td>Gibco Penicillin-Streptomycin</td><td>Fisher Scientific</td><td>15-140-148</td><td></td></tr><tr><td>HyClone Nonessential Amino Acids 100x</td><td>Fisher Scientific</td><td>SH3023801</td><td></td></tr><tr><td>Fetal Bovine Serum</td><td>Omega Scientific</td><td>FB-12</td><td></td></tr><tr><td>Corning DMEM with L-glutamine and glucose</td><td>Fisher Scientific</td><td>MT10013CV</td><td></td></tr><tr><td>Trichloro(1H,1H,2H,2H-perfluorooctyl)silane</td><td>Sigma-Aldritch</td><td>448931</td><td>Reacts violently with water</td></tr><tr><td>Ethanol, 200 proof Decon Labs</td><td>Fisher Scientific</td><td>04-355-223</td><td></td></tr><tr><td>Acetone</td><td>Fisher Scientific</td><td>A18P-4</td><td></td></tr><tr><td>Bel-Art 42025 Plastic Dessicator</td><td>Cole-Parmer</td><td>EW-06514-30</td><td></td></tr><tr><td>Epilog Fusion Laser Cutter, 120 W</td><td>Epilog Laser</td><td>Epilog Fusion M2 32 Laser</td><td></td></tr><tr><td>Isotemp Stirring Hotplate</td><td>Fisher Scientific</td><td>SP88850200</td><td></td></tr><tr><td>Ateco 14111 1.5 inch stainless steel cutter</td><td>Ateco</td><td>14111</td><td></td></tr><tr><td>Pyrex Glass Cell Culture Dish</td><td>Fisher Scientific</td><td>08-747B</td><td></td></tr><tr><td>Radio Frequency Plasma Cleaner</td><td>Harrick Plasma</td><td>PDC-32G</td><td>Used with Oxygen gas</td></tr><tr><td>Black Hole Laboratories Digivac</td><td>Black Hole Laboratories</td><td>Model 215</td><td></td></tr><tr><td>Intelli-Ray Ultraviolet Oven</td><td>Uvitron</td><td>UVO338</td><td></td></tr><tr><td>Compact Spin Coater</td><td>MTI Corporation</td><td>VTC-100A</td><td></td></tr><tr><td>Fisher Brand Isotemp Oven</td><td>Fisher Scientific</td><td>15-103-0510</td><td>Forced Air Convection</td></tr><tr><td>Gilson Positive Displacement Pipette P1000</td><td>Fisher Scientific</td><td>FD10006G</td><td></td></tr><tr><td>HeraCell VIOS 160i</td><td>Fisher Scientific</td><td>13 998 212PM</td><td></td></tr></tbody></table>

fabrication of refractive-index-matched devices for biomedical microfluidics

The use of microfluidic devices has emerged as a defining tool for biomedical applications. When combined with modern microscopy techniques, these devices can be implemented as part of a robust platform capable of making simultaneous complementary measurements. The primary challenge created by the combination of these two techniques is the mismatch in refractive index between the materials traditionally used to make microfluidic devices and the aqueous solutions typically used in biomedicine. This mismatch can create optical artifacts near the channel or device edges. One solution is to reduce the refractive index of the material used to fabricate the device by using a fluorinated polymer such as MY133-V2000 whose refractive index is similar to that of water (n = 1.33). Here, the construction of a microfluidic device made out of MY133-V2000 using soft lithography techniques is demonstrated, using O2 plasma in conjunction with an acrylic holder to increase the adhesion between the MY133-V2000 fabricated device and the polydimethylsiloxane (PDMS) substrate. The device is then tested by incubating it filled with cell culture media for 24 h to demonstrate the ability of the device to maintain cell culture conditions during the course of a typical imaging experiment. Finally, quantitative phase microscopy (QPM) is used to measure the distribution of mass within the live adherent cells in the microchannel. This way, the increased precision, enabled by fabricating the device from a low index of refraction polymer such as MY133-V2000 in lieu of traditional soft lithography materials such as PDMS, is demonstrated. Overall, this approach for fabricating microfluidic devices can be readily integrated into existing soft lithography workflows in order to reduce optical artifacts and increase measurement precision.

This protocol describes the fabrication of MY133-V2000, a fluorinated polymer with a low refractive index matching that of water. A key feature of this protocol is how to overcome the lack of adhesion that is characteristic of fluorinated polymers by using oxygen plasma and by fabricating the device within an acrylic holder to provide the extra mechanical force required to seal the channel against the PDMS substrate (Figure 1). The low refractive index of the...

Watch this Scientific Journal Video about Fabrication of Refractive-index-matched Devices for Biomedical Microfluidics at JoVE.com

Fabrication of Refractive-index-matched Devices for Biomedical Microfluidics

This protocol describes the fabrication of microfluidic devices from MY133-V2000 to eliminate artifacts that often arise in microchannels due to the mismatching refractive indices between microchannel structures and an aqueous solution. This protocol uses an acrylic holder to compress the encapsulated device, improving adhesion both chemically and mechanically.

The use of microfluidic devices has emerged as a defining tool for biomedical applications. When combined with modern microscopy techniques, these devices ...

fabrication-refractive-index-matched-devices-for-biomedical

Universidad San Sebastian

Research

JoVE Journal

Engineering

7.4K Views.  University of Utah. This protocol describes the fabrication of microfluidic devices from MY133-V2000 to eliminate artifacts that often arise in microchannels due to the mismatching refractive indices between microchannel structures and an aqueous solution. This protocol uses an acrylic holder to compress the encapsulated device, improving adhesion both chemically and mechanically.

Video: Fabrication of Refractive-index-matched Devices for Biomedical Microfluidics

Fabrication of Silica Ultra High Quality Factor Microresonators

Whispering gallery resonant cavities confine light in circular orbits at their periphery.1-2 The photon storage lifetime in the cavity, quantified by the quality factor (Q) of the cavity, can be in excess of 500ns for cavities with Q factors above 100 million. As a result of their low material losses, silica microcavities have demonstrated some of the longest photon lifetimes to date1-2. Since a portion of the circulating light extends outside the resonator, these devices can also be used to probe the surroundings. This interaction has enabled numerous experiments in biology, such as single molecule biodetection and antibody-antigen kinetics, as well as discoveries in other fields, such as development of ultra-low-threshold microlasers, characterization of thin films, and cavity quantum electrodynamics studies.3-7
The two primary silica resonant cavity geometries are the microsphere and the microtoroid. Both devices rely on a carbon dioxide laser reflow step to achieve their ultra-high-Q factors (Q&gt;100 million).1-2,8-9 However, there are several notable differences between the two structures. Silica microspheres are free-standing, supported by a single optical fiber, whereas silica microtoroids can be fabricated on a silicon wafer in large arrays using a combination of lithography and etching steps. These differences influence which device is optimal for a given experiment.
Here, we present detailed fabrication protocols for both types of resonant cavities. While the fabrication of microsphere resonant cavities is fairly straightforward, the fabrication of microtoroid resonant cavities requires additional specialized equipment and facilities (cleanroom). Therefore, this additional requirement may also influence which device is selected for a given experiment.
Introduction
An optical resonator efficiently confines light at specific wavelengths, known as the resonant wavelengths of the device. 1-2 The common figure of merit for these optical resonators is the quality factor or Q. This term describes the photon lifetime (&tau;o) within the resonator, which is directly related to the resonator's optical losses. Therefore, an optical resonator with a high Q factor has low optical losses, long photon lifetimes, and very low photon decay rates (1/&tau;o). As a result of the long photon lifetimes, it is possible to build-up extremely large circulating optical field intensities in these devices. This very unique property has allowed these devices to be used as laser sources and integrated biosensors.10
A unique sub-class of resonators is the whispering gallery mode optical microcavity. In these devices, the light is confined in circular orbits at the periphery. Therefore, the field is not completely confined within the device, but evanesces into the environment. Whispering gallery mode optical cavities have demonstrated some of the highest quality factors of any optical resonant cavity to date.9,11 Therefore, these devices are used throughout science and engineering, including in fundamental physics studies and in telecommunications as well as in biodetection experiments. 3-7,12
Optical microcavities can be fabricated from a wide range of materials and in a wide variety of geometries. A few examples include silica and silicon microtoroids, silicon, silicon nitride, and silica microdisks, micropillars, and silica and polymer microrings.13-17 The range in quality factor (Q) varies as dramatically as the geometry. Although both geometry and high Q are important considerations in any field, in many applications, there is far greater leverage in boosting device performance through Q enhancement. Among the numerous options detailed previously, the silica microsphere and the silica microtoroid resonator have achieved some of the highest Q factors to date.1,9 Additionally, as a result of the extremely low optical loss of silica from the visible through the near-IR, both microspheres and microtoroids are able to maintain their Q factors over a wide range of testing wavelengths.18 Finally, because silica is inherently biocompatible, it is routinely used in biodetection experiments.
In addition to high material absorption, there are several other potential loss mechanisms, including surface roughness, radiation loss, and contamination loss.2 Through an optimization of the device size, it is possible to eliminate radiation losses, which arise from poor optical field confinement within the device. Similarly, by storing a device in an appropriately clean environment, contamination of the surface can be minimized. Therefore, in addition to material loss, surface scattering is the primary loss mechanism of concern.2,8
In silica devices, surface scattering is minimized by using a laser reflow technique, which melts the silica through surface tension induced reflow. While spherical optical resonators have been studied for many years, it is only with recent advances in fabrication technologies that researchers been able to fabricate high quality silica optical toroidal microresonators (Q&gt;100 million) on a silicon substrate, thus paving the way for integration with microfluidics.1
The present series of protocols details how to fabricate both silica microsphere and microtoroid resonant cavities. While silica microsphere resonant cavities are well-established, microtoroid resonant cavities were only recently invented.1 As many of the fundamental methods used to fabricate the microsphere are also used in the more complex microtoroid fabrication procedure, by including both in a single protocol it will enable researchers to more easily trouble-shoot their experiments.

We describe the use of a carbon dioxide laser reflow technique to fabricate silica resonant cavities, including free-standing microspheres and on-chip microtoroids. The reflow method removes surface imperfections, allowing long photon lifetimes within both devices. The resulting devices have ultra high quality factors, enabling applications ranging from telecommunications to biodetection.

Whispering gallery resonant cavities confine light in circular orbits at their periphery.1-2 The photon storage lifetime in the cavity, quantified by the ...

Fabrication and Testing of Microfluidic Optomechanical Oscillators

Cavity optomechanics experiments that parametrically couple the phonon modes and photon modes have been investigated in various optical systems including microresonators. However, because of the increased acoustic radiative losses during direct liquid immersion of optomechanical devices, almost all published optomechanical experiments have been performed in solid phase. This paper discusses a recently introduced hollow microfluidic optomechanical resonator. Detailed methodology is provided to fabricate these ultra-high-Q microfluidic resonators, perform optomechanical testing, and measure radiation pressure-driven breathing mode and SBS-driven whispering gallery mode parametric vibrations. By confining liquids inside the capillary resonator, high mechanical- and optical- quality factors are simultaneously maintained.

Parametric optomechanical excitations have recently been experimentally demonstrated in microfluidic optomechanical resonators by means of optical radiation pressure and stimulated Brillouin scattering. This paper describes the fabrication of these microfluidic resonators along with methodologies for generating and verifying optomechanical oscillations.

Cavity optomechanics experiments that parametrically couple the phonon modes and photon modes have been investigated in various optical systems including ...

Endothelialized Microfluidics for Studying Microvascular Interactions in Hematologic Diseases

Advances in microfabrication techniques have enabled the production of inexpensive and reproducible microfluidic systems for conducting biological and biochemical experiments at the micro- and nanoscales 1,2. In addition, microfluidics have also been specifically used to quantitatively analyze hematologic and microvascular processes, because of their ability to easily control the dynamic fluidic environment and biological conditions3-6. As such, researchers have more recently used microfluidic systems to study blood cell deformability, blood cell aggregation, microvascular blood flow, and blood cell-endothelial cell interactions6-13.However, these microfluidic systems either did not include cultured endothelial cells or were larger than the sizescale relevant to microvascular pathologic processes. A microfluidic platform with cultured endothelial cells that accurately recapitulates the cellular, physical, and hemodynamic environment of the microcirculation is needed to further our understanding of the underlying biophysical pathophysiology of hematologic diseases that involve the microvasculature.
Here, we report a method to create an "endothelialized" in vitro model of the microvasculature, using a simple, single mask microfabrication process in conjunction with standard endothelial cell culture techniques, to study pathologic biophysical microvascular interactions that occur in hematologic disease. This "microvasculature-on-a-chip" provides the researcher with a robust assay that tightly controls biological as well as biophysical conditions and is operated using a standard syringe pump and brightfield/fluorescence microscopy. Parameters such as microcirculatory hemodynamic conditions, endothelial cell type, blood cell type(s) and concentration(s), drug/inhibitory concentration etc., can all be easily controlled. As such, our microsystem provides a method to quantitatively investigate disease processes in which microvascular flow is impaired due to alterations in cell adhesion, aggregation, and deformability, a capability unavailable with existing assays.

A method to culture an endothelial cell monolayer throughout the entire inner 3D surface of a microfluidic device with microvascular-sized channels (&lt;30 &mu;m) is described. This in vitro microvasculature model enables the study of biophysical interactions between blood cells, endothelial cells, and soluble factors in hematologic diseases.

Advances in  microfabrication techniques have enabled the production of inexpensive and  reproducible microfluidic systems for conducting biological and ...

Bioengineering

Fabrication and Operation of Acoustofluidic Devices Supporting Bulk Acoustic Standing Waves for Sheathless Focusing of Particles

Acoustophoresis refers to the displacement of suspended objects in response to directional forces from sound energy. Given that the suspended objects must be smaller than the incident wavelength of sound and the width of the fluidic channels are typically tens to hundreds of micrometers across, acoustofluidic devices typically use ultrasonic waves generated from a piezoelectric transducer pulsating at high frequencies (in the megahertz range). At characteristic frequencies that depend on the geometry of the device, it is possible to induce the formation of standing waves that can focus particles along desired fluidic streamlines within a bulk flow. Here, we describe a method for the fabrication of acoustophoretic devices from common materials and clean room equipment. We show representative results for the focusing of particles with positive or negative acoustic contrast factors, which move towards the pressure nodes or antinodes of the standing waves, respectively. These devices offer enormous practical utility for precisely positioning large numbers of microscopic entities (e.g., cells) in stationary or flowing fluids for applications ranging from cytometry to assembly.

Acoustofluidic devices use ultrasonic waves within microfluidic channels to manipulate, concentrate and isolate suspended micro and nanoscopic entities. This protocol describes the fabrication and operation of such a device supporting bulk acoustic standing waves to focus particles in a central streamline without the aid of sheath fluids.

Acoustophoresis refers to the displacement of suspended objects in response to directional forces from sound energy. Given that the suspended objects must ...

One-Step Approach to Fabricating Polydimethylsiloxane Microfluidic Channels of Different Geometric Sections by Sequential Wet Etching Processes

Polydimethylsiloxane (PDMS) materials are substantially exploited to fabricate microfluidic devices by using soft lithography replica molding techniques. Customized channel layout designs are necessary for specific functions and integrated performance of microfluidic devices in numerous biomedical and chemical applications (e.g., cell culture, biosensing, chemical synthesis, and liquid handling). Owing to the nature of molding approaches using silicon wafers with photoresist layers patterned by photolithography as master molds, the microfluidic channels commonly have regular cross sections of rectangular shapes with identical heights. Typically, channels with multiple heights or different geometric sections are designed to possess particular functions and to perform in various microfluidic applications (e.g., hydrophoresis is used for sorting particles and in continuous flows for separating blood cells6,7,8,9). Therefore, a great deal of effort has been made in constructing channels with various sections through multiple-step approaches like photolithography using several photoresist layers and assembly of different PDMS thin sheets. Nevertheless, such multiple-step approaches usually involve tedious procedures and extensive instrumentation. Furthermore, the fabricated devices may not perform consistently and the resulted experimental data may be unpredictable. Here, a one-step approach is developed for the straightforward fabrication of microfluidic channels with different geometric cross sections through PDMS sequential wet etching processes, that introduces etchant into channels of planned single-layer layouts embedded in PDMS materials. Compared to the existing methods for manufacturing PDMS microfluidic channels with different geometries, the developed one-step approach can significantly simplify the process to fabricate channels with non-rectangular sections or various heights. Consequently, the technique is a way of constructing complex microfluidic channels, which provides a fabrication solution for the advancement of innovative microfluidic systems.

Several methods are available for the fabrication of channels of non-rectangular sections embedded in polydimethylsiloxane microfluidic devices. Most of them involve multistep manufacturing and extensive alignment. In this paper, a one-step approach is reported for fabricating microfluidic channels of different geometric cross sections by polydimethylsiloxane sequential wet etching.

Polydimethylsiloxane (PDMS) materials are substantially exploited to fabricate microfluidic devices by using soft lithography replica molding techniques. ...

Design and Development of a Three-Dimensionally Printed Microscope Mask Alignment Adapter for the Fabrication of Multilayer Microfluidic Devices

This project aims to develop an easy-to-use and cost-effective platform for the fabrication of precise, multilayer microfluidic devices, which typically can only be achieved using costly equipment in a clean room setting. The key part of the platform is a three dimensionally (3D) printed microscope mask alignment adapter (MMAA) compatible with regular optical microscopes and ultraviolet (UV) light exposure systems. The overall process of creating the device has been vastly simplified because of the work done to optimize the device design. The process entails finding the proper dimensions for the equipment available in the laboratory and 3D-printing the MMAA with the optimized specifications. Experimental results show that the optimized MMAA designed and manufactured by 3D printing performs well with a common microscope and light exposure system. Using a master mold prepared by the 3D-printed MMAA, the resulting microfluidic devices with multilayered structures contain alignment errors of &lt;10 &#181;m, which is sufficient for common microchips. Although human error through transportation of the device to the UV light exposure system can cause larger fabrication errors, the minimal errors achieved in this study are attainable with practice and care. Furthermore, the MMAA can be customized to fit any microscope and UV exposure system by making changes to the modeling file in the 3D printing system. This project provides smaller laboratories with a useful research tool as it only requires the use of equipment that is typically already available to laboratories that produce and use microfluidic devices. The following detailed protocol outlines the design and 3D printing process for the MMAA. In addition, the steps for procuring a multilayer master mold using the MMAA and producing poly(dimethylsiloxane) (PDMS) microfluidic chips is also described herein.

This project allows small laboratories to develop an easy-to-use platform for the fabrication of precise multilayer microfluidic devices. The platform consists of a three-dimensionally printed microscope mask alignment adapter using which multilayer microfluidic devices with alignment errors of &lt;10 &#181;m were achieved.

This project aims to develop an easy-to-use and cost-effective platform for the fabrication of precise, multilayer microfluidic devices, which typically ...

Fabrication of Three-dimensional Paper-based Microfluidic Devices for Immunoassays

Paper wicks fluids autonomously due to capillary action. By patterning paper with hydrophobic barriers, the transport of fluids can be controlled and directed within a layer of paper. Moreover, stacking multiple layers of patterned paper creates sophisticated three-dimensional microfluidic networks that can support the development of analytical and bioanalytical assays. Paper-based microfluidic devices are inexpensive, portable, easy to use, and require no external equipment to operate. As a result, they hold great promise as a platform for point-of-care diagnostics. In order to properly evaluate the utility and analytical performance of paper-based devices, suitable methods must be developed to ensure their manufacture is reproducible and at a scale that is appropriate for laboratory settings. In this manuscript, a method to fabricate a general device architecture that can be used for paper-based immunoassays is described. We use a form of additive manufacturing (multi-layer lamination) to prepare devices that comprise multiple layers of patterned paper and patterned adhesive. In addition to demonstrating the proper use of these three-dimensional paper-based microfluidic devices with an immunoassay for human chorionic gonadotropin (hCG), errors in the manufacturing process that may result in device failures are discussed. We expect this approach to manufacturing paper-based devices will find broad utility in the development of analytical applications designed specifically for limited-resource settings.

We detail a method to fabricate three-dimensional paper-based microfluidic devices for use in the development of immunoassays. Our approach to device assembly is a type of multilayer, additive manufacturing. We demonstrate a sandwich immunoassay to provide representative results for these types of paper-based devices.

Paper wicks fluids autonomously due to capillary action. By patterning paper with hydrophobic barriers, the transport of fluids can be controlled and ...

Computer Numerical Control Micromilling of a Microfluidic Acrylic Device with a Staggered Restriction for Magnetic Nanoparticle-Based Immunoassays

Microfluidic systems have greatly improved immunoassay techniques. However, many microfabrication techniques require specialized, expensive, or complicated equipment, making fabrication costly and incompatible with mass production, which is one of the most important preconditions for point-of-care tests (POCT) to be adopted in low-resource settings. This work describes the fabrication process of an acrylic (polymethylmethacrylate, PMMA) device for nanoparticle-conjugated enzymatic immunoassay testing using the computer numerical control (CNC) micromilling technique. The functioning of the microfluidic device is shown by performing an immunoassay to detect a commercial antibody using lysozyme as a model antigen conjugated to 100 nm magnetic nanoparticles. This device integrates a physical staggered restriction of only 5 &#181;m in height, used to capture magnetic microparticles that make up a magnetic trap by placing an external magnet. In this way, the magnetic force on the immunosupport of conjugated nanoparticles is enough to capture them and resist flow drag. This microfluidic device is particularly suitable for low-cost mass production without the loss of precision for immunoassay performance.

Microfluidics is a powerful tool for the development of diagnostic tests. However, expensive equipment and materials, as well as laborious fabrication and handling techniques, are often required. Here, we detail the fabrication protocol of an acrylic microfluidic device for magnetic micro- and nanoparticle-based immunoassays in a low-cost and simple-to-use setting.

Microfluidic systems have greatly improved immunoassay techniques. However, many microfabrication techniques require specialized, expensive, or ...

Soft Lithographic Procedure for Producing Plastic Microfluidic Devices with View-ports Transparent to Visible and Infrared Light

Infrared (IR) spectro-microscopy of living biological samples is hampered by the absorption of water in the mid-IR range and by the lack of suitable microfluidic devices. Here, a protocol for the fabrication of plastic microfluidic devices is demonstrated, where soft lithographic techniques are used to embed transparent Calcium Fluoride (CaF2) view-ports in connection with observation chamber(s). The method is based on a replica casting approach, where a polydimethylsiloxane (PDMS) mold is produced through standard lithographic procedures and then used as the template to produce a plastic device. The plastic device features ultraviolet/visible/infrared (UV/Vis/IR) -transparent windows made of CaF2 to allow for direct observation with visible and IR light. The advantages of the proposed method include: a reduced need for accessing a clean room micro-fabrication facility, multiple view-ports, an easy and versatile connection to an external pumping system through the plastic body, flexibility of the design, e.g., open/closed channels configuration, and the possibility to add sophisticated features such as nanoporous membranes.

A protocol for the fabrication of plastic microfluidic devices with transparent view-ports for visible and infrared light imaging is described.

Infrared (IR) spectro-microscopy of living biological samples is hampered by the absorption of water in the mid-IR range and by the lack of suitable ...

Rapid Fabrication of Custom Microfluidic Devices for Research and Educational Applications

Microfluidic devices allow for the manipulation of fluids, particles, cells, micro-sized organs or organisms in channels ranging from the nano to submillimeter scales. A rapid increase in the use of this technology in the biological sciences has prompted a need for methods that are accessible to a wide range of research groups. Current fabrication standards, such as PDMS bonding, require expensive and time consuming lithographic and bonding techniques. A viable alternative is the use of equipment and materials that are easily affordable, require minimal expertise and allow for the rapid iteration of designs. In this work we describe a protocol for designing and producing PET-laminates (PETLs), microfluidic devices that are inexpensive, easy to fabricate, and consume significantly less time to generate than other approaches to microfluidics technology. They consist of thermally bonded film sheets, in which channels and other features are defined using a craft cutter. PETLs solve field-specific technical challenges while dramatically reducing obstacles to adoption. This approach facilitates the accessibility of microfluidics devices in both research and educational settings, providing a reliable platform for new methods of inquiry.

Here we present a protocol to design and fabricate custom microfluidic devices with minimal financial and time investment. The aim is to facilitate the adoption of microfluidic technologies in biomedical research laboratories and educational settings.