Name: microfluidic mixers for studying protein folding
Uploaded: 2012-04-10T13:00:00
Duration: 12 min 42 s
Description: Watch this Scientific Journal Video about Microfluidic Mixers for Studying Protein Folding at JoVE.com

This work is supported by National Science Foundation FIBR (NSF EF-0623664) and IDBR (NSF DBI-0754570). This work was partially supported by funding from National Science Foundation FIBR Grant 0623664 administered by the Center for Biophotonics, an NSF Science and Technology Center, managed by the University of California, Davis, under Cooperative Agreement PHY 0120999. The research of Lisa Lapidus, Ph.D. is supported in part by a Career Award at the Scientific Interface from the Burroughs Welcome Fund.

1. Fabrication of Microfluidic Mixing Chips
Figure 3 shows the basic fabrication steps.
<ol>
	<li>Clean 4 inch (100 mm) fused silica wafers with fresh Piranha solution (3:1 H2SO4:H2O2): i) Heat the sulfuric acid to 130 &deg;C then pour in the peroxide. Note that the surface roughness and the flatness of the fused silica wafers used play an important role for the final bonding step. ii) Immerse the wafers for at least 30 minutes. iii) Rinse for 5 minutes with water and dry. Apply a polysilicon (poly) coating, ~1 &mu;m thick per every intended 10 &mu;m depth of the final microfluidic channels, using a low pressure chemical vapor deposition (LPCVD) furnace.</li>
	<li>Clean the wafers and mask using the Piranha protocol in step 1.1. Clean the mask the same way, then rinse with acetone, methanol and isopropanol and blow dry immediately. Dehydrate the wafers on a hot plate at 150 &deg;C for ~10 minutes (use an aluminum foil tent to keep particulates away) or in a 120 &deg;C oven for at least 120 min (overnight preferably). To increase photoresist adhesion, immediately expose the warm wafers to HMDS vapor for 5 minutes. Spin coat the wafers with a ~900 nm thick layer of AZ5214 photoresist and soft bake at 90 &deg;C directly on a hotplate for 1 minute or in an oven at 110 &deg;C for 30 min (Figure 3, step 1).</li>
	<li>Align the mask and make a hard and vacuum contact. Expose to UV light for several seconds (total energy: ~103 mJ/cm2) (Figure 3, step 2). Develop with AZ400K developer, diluted 4:1 with DI water for ~50 seconds (until photoresist is fully developed) while gently agitating. Rinse with water for 2 minutes. Inspect features with microscope (Figure 3, step 3). If features are poorly resolved, strip the photoresist and start again from 1.2. Hard bake 115 &deg;C directly on a hotplate for 2 minutes.</li>
	<li>Descum wafers with Asher or O2 plasma for 2.5 minutes at 100 W RF power. Using a deep reactive ion etcher (DRIE), etch the polysilicon layer all the way through to the fused silica below. Strip the remaining photoresist with PRS2000 resist stripper at 75 &deg;C for 10 minutes. Rinse and dry. Measure the etch depth to ensure it is all the way through the poly coating (Figure 3, step 4).</li>
	<li>Using an oxide capable DRIE such as an ULVAC NLD-6000 etcher, etch fused silica to a depth of ~10 &mu;m per micron thickness of the poly coating (Figure 3, step 5). This poly coating will be worn away during etching, so it may be necessary to check on the integrity of the coating periodically during the etching process. If intentionally featureless regions of the wafer surface have become stripped bare of the protective poly coating during etching, the surface has most likely become pitted and will no longer be suitable for bonding in the final steps of production. In this case the wafer is most likely no longer viable. Clean with the Matrix Asher for 4 minutes, 100 W RF power. Strip the polysilicon with a XeF2 etcher (Figure 3, step 6). It may be necessary to repeat step 1.4 and this step a few times to remove all of the photoresist and poly coatings.</li>
	<li>Spin coat wafers with a new ~1 &mu;m layer of photoresist to protect the channels during subsequent handling. Drill entrance and exit holes in each chip using a computer controlled diamond-tipped drill or manually with a sandblaster (Figure 3, step 7). If using a drill, mount the wafer (feature-side down) on a sacrificial glass plate using Aqua Bond 55, taking care to keep the bonder free of bubbles. Cool the drill with micro-90 cleaning solution. This keeps small bits of glass from sticking to the channels which then clog the chip during use.</li>
	<li>Rinse the wafer with DI water using a syringe to inject water into each hole. Soak in soapy water for an hour. Remove photoresist and Aqua Bond with PRS 2000 at 60 &deg;C for several hours until surface is clean. Rinse wafer with DI water and warm soapy water. Rinse each hole again with a syringe, avoiding etched features. Dry wafer with nitrogen and in a vacuum oven set to 120 &deg;C. Inspect holes to ensure they are free of grit and repeat cleaning steps as necessary. Measure the depth of the channels with either an optical or mechanical surface profiler (Figure 3, step 8).</li>
	<li>Clean wafers and an equivalent number of 170 &mu;m thick fused silica cover glasses with 1) Piranha solution (1-2 hours according to the procedure in 1.1), 2) RCA 2 (5:1:1 DI:HCl:H202 ) solution for 10 minutes at 75 C, 3) RCA 1 solution (5:1:1 DI:NH4OH:H202) for 22 minutes at 75C). Rinse with DI water for 5 minutes between each solution.</li>
	<li>Place one wafer features side up on top of a stack of 3-4 cleanroom wipes placed on top of a hard flat surface, such as a small pane of glass. Dry the wafer thoroughly with nitrogen gas for at least 5 minutes. Repeat with cover glass. Pick up cover glass by the edge and invert such that the polished side is face down held over the wafer and align the flat edges. Carefully drop the cover glass onto the wafer and allow it to settle. Nudge horizontally only to adjust alignment. Push down with one finger on the center of the wafer. One should see a bonding front begin to radiate outward. If necessary, apply additional pressure to other positions around the wafer to advance the bonding front (Figure 3, step 9).</li>
	<li>Place the sealed wafers in a high temperature oven set to ramp from room temp to 800 &deg;C over 3 hours, then from 800 - 1100 &deg;C over another 1.2 hours. Hold at 1100 &deg;C for 2 hours, and then ramp down to room temperature over another 1.5 hours.</li>
	<li>Dice with a wafer dicing saw into individual chips (Figure 3, step 10).</li>
	<li>The protocol describes the fabrication of channels in the fairly unusual fused silica substrate which is necessary for UV detection. But this protocol can also be adapted for a silicon substrate which requires only one etching step2. A glass (but not fused silica) cover glass can be bonded to the wafer using anodic bonding.</li>
</ol>
2. Mounting and Loading Chips
<ol>
	<li>The manifold to hold the mixing chip and solutions can be made of plastic, such as Lexan or Plexiglas , or aluminum for temperature control (see Figure 4). If using aluminum, the solution reservoirs should be coated with parylene to prevent the dissolution of aluminum by the salt solutions in the reservoirs. The temperature of the entire manifold, solutions and chip can be controlled with thermoelectric devices mounted on the sides and an electronic controller.</li>
	<li>The chip is mated to the manifold by small o-rings and a held in place with a retaining ring that allows a clear optical view of the center of the chip. The o-ring grooves should be shallower than standard to ensure good mating without applying too much pressure to the chip, i.e., a 002 o-ring should be 0.025" deep. Once the chip is mounted, add solutions to the center and side reservoirs, taking care to release any bubbles trapped at the bottom of the wells.</li>
	<li>Because the volumes used in this mixer are so small, the flow can be controlled with air pressure applied above the solution wells. Figure 4 shows the pressure manifold mated to the top of the chip manifold by o-rings. The pressure manifold is connected to computer-controlled pressure transducers that can maintain pressures from ~2-80 PSI. The chip has been designed such that equal pressures on the center and side channels produce a ~100-fold ratio in the linear flow rates. At 20 PSI, the linear flow rate in the exit channel is ~1 m/s.</li>
	<li>Place manifold on an inverted microscope and examine mixing region with the eyepiece or camera output. An incandescent flashlight placed on top of the manifold will provide enough light. Use a dye or high index of refraction solution (i.e. 6 M guanidine hydrochloride) in the center channel to view of the jet.</li>
	<li>At equal pressure on the center and side channels, the jet will be difficult to see by eye so start with the side channel pressure at 0 PSI and slowly increase to equal pressure. If one side channel is clogged, the jet will be pushed against one of the walls of the exit channel. If a visible clog appears, it may be dislodged by applying pressure or vacuum to the exit channel with a syringe.</li>
	<li>If protein or other organic material clogs the chip, it can be cleaned by soaking in Pirhana solution overnight.</li>
</ol>
3. Data Collection
Figure 5 shows the basic optical instrument layout.
<ol>
	<li>Bring a collimated laser beam into a research-grade microscope using a dichroic mirror either inside or just outside the microscope. Align the laser so that the focus can be seen in the eyepiece or camera somewhat near the mixing region.</li>
	<li>Fluorescence from the protein in the mixer is collected by the objective and sent through the dichroic mirror, focused by a lens to a pinhole and imaged on a photon counter. Scan the chip using a piezoelectric scanner in x and y to image the mixing region (typical step size is 2 &mu;m). Choose a position on the jet and scan in x and z to find the focus at the center of the channel vertically. The depth of field of the microscope is much less than the channel depth and the flow rate is uniform in the channel except within 1 &mu;m of the walls. Therefore the temporal resolution of the observed fluorescence is determined primarily by the size of the confocal spot.</li>
	<li>Scan horizontally within the exit channel (typical step size is 0.2 &mu;m in across the jet, 2 &mu;m along the jet) in 100 &mu;m increments along the jet, observing for ~1 ms per position (see Figure 6). Move the microscope stage by 80-90 &mu;m along the jet to capture later times.</li>
	<li>Alternatively, light can be detected by a spectrograph and CCD after the dichroic mirror using a lens to focus the light onto the entrance slit. Since data collection is slower than for the photon counter (1 second per position), typically the jet is imaged first with the photon counter and then points along the jet are measured with the spectrograph (see Figure 7).</li>
	<li>Data from the photon counter is analyzed by summing 3-5 pixels around the jet at each position to create a plot of intensity vs. distance. Time is calculated from distance using a calculated velocity based on the applied pressures (see Figure 8).</li>
	<li>Data from the spectrograph can be analyzed a couple of ways. For FRET, channels designated as "donor" and "acceptor" can each be summed and the proximity ratio E=IA/(ID+ID) calculated (see Figure 9). Alternatively, single value decomposition can be performed to look at independent spectral components vs. time, such as the shift in the tryptophan emission spectrum as a protein folds.</li>
	<li>Distance within the exit channel can be converted to time using the constant flow rate determined from the applied pressure to the side channels. Within the first 2 &mu;m of the exit channel, the flow rate of the protein jet accelerates ~100x and the times in that region must be calculated with finite element analysis of the streamlines (producing the times plotted in Figure 8). This acceleration produces an offset of ~1-2 &mu;s after the velocity becomes constant and can usually be ignored in measuring kinetics much slower than mixing.</li>
</ol>
4. Representative Results
Figure 6 shows a contour plot of the intensity in the mixing region measured by the photon counter. The background in the exit channel outside of the jet should be close to the noise floor of the detector and is typically not subtracted. A higher background may indicate poor alignment or that the protein is sticking to the walls of the channel. A jet that looks kinked or crooked, especially near the mixing region, indicates a poor etch of the nozzles.
Figure 7 shows the time resolved spectra from the protein acyl-coenzyme A-binding protein (ACBP) labeled with Alexa 488 and Alexa 647. This data has been background subtracted to remove the dark charge and laser signal. The background signal was taken with the center channel pressure set to 0 PSI.
The mixing time can be measured by measuring a rapid reaction that is much faster than mixing such as Trp fluorescence quenching by KI or collapse of single stranded DNA, labeled with FRET dyes, by the addition of NaCl. Because the jet is smaller than the optical resolution of the microscope, there is a large intensity drop in the mixing region as the jet forms. This instrument response can be removed by making a control measurement in which no solution conditions change. Figure 8 shows the ratio of mixing (into 400 mM KI) and non-mixing experiments of N-acetyl tryptophan amide fluorescence. The line shows the predicted concentration of KI from COMSOL simulations. The mixing time, as measured as the time for the concentration to decrease 80%, is 8 &mu;s.
Since data is collected in 100 &mu;m increments along the 500 &mu;m-long exit channel, data must be pasted together from multiple views. There are occasionally small offsets in signal between these views, likely due to small changes in the focus as the chip is moved by the microscope stage. By moving the stage 80 or 90 &mu;m per view, these offsets can be eliminated by examining overlapping data. Typically data is collected with at least two flow rates and the data is combined to confirm any signal changes are due to real spectroscopic changes in the protein, rather than optical or flow effects. Figure 9 shows the FRET changes in the protein ACBP after dilution of denaturant from 6 M to 0.06 M. This data does not need a control experiment since the FRET is already a ratio and the instrument response is already removed. The rapid jump near T=0 represents a rapid change in FRET signal within the mixing time.
<img alt="Figure 1" src="/files/ftp_upload/3976/3976fig1.jpg" /> 
Figure 1. Layout of mixing region measured by electron microscopy.
<img alt="Figure 2" src="/files/ftp_upload/3976/3976fig2.jpg" /> 
Figure 2. Schematic of hydrodynamic mixing to study protein folding. The protein, dissolved in high denaturant to unfold it, flows down the center channel towards the mixing region where it meets buffer flowing ~100 times faster. Laminar flow forces the protein stream into a narrow jet from which denaturant molecules can rapidly diffuse. As the jet flows down the exit channel, the protein can be observed with high spatial and time resolution using a confocal microscope.
<img alt="Figure 3" src="/files/ftp_upload/3976/3976fig3.jpg" /> 
Figure 3. Fused silica fabrication. 1) The process starts with a highly polished 500 &mu;m thick fused silica substrate (a) with a layer of polysilicon (poly) deposited on the top and bottom surfaces (b) and spin-coated with a layer of photoresist (c). 2) A photomask (e) is brought into hard contact with the wafer and the photoresist is exposed to UV light (d). 3) The wafer is placed in a developer bath and the pattern is revealed in the photoresist. 4) The wafer is placed in a DRIE and the features are etched into the top poly coating. The photoresist is then removed. 5) The poly then acts as a mask when the wafer is placed in an oxide etcher and the features are etched 10 - 40 &mu;m into the fused silica substrate. 6) The poly coating is then removed in a XeF2 etcher. 7) The wafer is bonded to a sacrificial glass plate with a heat-activated sealant (g), features side down, and a diamond drill bit (f) abrasively drills through-holes from the backside. 8) A surface profiler (h) then directly measures the etched feature depths. 9) A 170 &mu;m thick fused silica coverslip wafer is directly bonded on the top surface of the wafer. 10) The wafer is diced into individual microfluidic chips.
<img alt="Figure 4" src="/files/ftp_upload/3976/3976fig4.jpg" /> 
Figure 4. The mixer manifold. The microfluidic-chip is secured to the solution manifold with a machined aluminum faceplate and four o-rings under the ~200 &mu;L reservoirs. A large hole is machined through the center of the manifold directly above the mixing region of the affixed microfluidic-chip, allowing excess UV light to escape without backscattering, inhibiting any auto-fluorescence from the manifold itself and allowing direct illumination of the chip by eye or camera for alignment. The air manifold seals each of the reservoirs such that the air pressure in each can be controlled via an external pressure control box.
<img alt="Figure 5" src="/files/ftp_upload/3976/3976fig5.jpg" /> 
Figure 5. Schematic of optical confocal microscope.
<img alt="Figure 6" src="/files/ftp_upload/3976/3976fig6.jpg" /> 
Figure 6. Contour plot of tryptophan fluorescence in the microfluidic mixer. The mixing region is located at ~90 &mu;m on the y-axis.
<img alt="Figure 7" src="/files/ftp_upload/3976/3976fig7.jpg" /> 
Figure 7. Time dependent fluorescent spectra collected within the mixer. The green and red rectangles show the wavelengths designated as donor and acceptor channels, respectively.
<img alt="Figure 8" src="/files/ftp_upload/3976/3976fig8.jpg" /> 
Figure 8. Mixing time measured by fluorescence quenching of tryptophan by potassium iodide (points and right axis) and calculated by finite element analysis (line and left axis).
<img alt="Figure 9" src="/files/ftp_upload/3976/3976fig9.jpg" /> 
Figure 9. FRET changes measured during the folding of the protein ACBP. The rapid rise near t=0 represents a burst phase within the mixing time and the slower rise represents a gradual accumulation of structure as the protein folds. The data was taken at two different flow rates and overlaid, leading to different densities of measurements at different times.

<ol>
	<li>Eaton, W. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fast+kinetics+and+mechanisms+in+protein+folding.">Fast kinetics and mechanisms in protein folding.</a> Annual Review of Biophysics and Biomolecular Structure. 29, 327-359 (2000).</li><li>Hertzog, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Femtomole+mixer+for+microsecond+kinetic+studies+of+protein+folding.">Femtomole mixer for microsecond kinetic studies of protein folding.</a> Analytical Chemistry. 76, 7169-7178 (2004).</li><li>Hertzog, D. E., Ivorra, B., Mohammadi, B., Bakajin, O., Santiago, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optimization+of+a+microfluidic+mixer+for+studying+protein+folding+kinetics.">Optimization of a microfluidic mixer for studying protein folding kinetics.</a> Analytical Chemistry. 78, 4299-4306 (2006).</li><li>Yao, S., Bakajin, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improvements+in+Mixing+Time+and+Mixing+Uniformity+in+Devices+Designed+for+Studies+of+Protein+Folding+Kinetics.">Improvements in Mixing Time and Mixing Uniformity in Devices Designed for Studies of Protein Folding Kinetics.</a> Analytical Chemistry. 79, 5753-5759 (2007).</li><li>Lapidus, L. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein+Hydrophobic+Collapse+and+Early+Folding+Steps+Observed+in+a+Microfluidic+Mixer.">Protein Hydrophobic Collapse and Early Folding Steps Observed in a Microfluidic Mixer.</a> Biophysical Journal. 93, 218-224 (2007).</li><li>Bilsel, O., Kayatekin, C., Wallace, L. A., Matthews, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+microchannel+solution+mixer+for+studying+microsecond+protein+folding+reactions.">A microchannel solution mixer for studying microsecond protein folding reactions.</a> Review of Scientific Instruments. 76, (2005).</li><li>Chan, C. -. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Submillisecond+protein+folding+kinetics+studied+by+ultrarapid+mixing.">Submillisecond protein folding kinetics studied by ultrarapid mixing.</a> PNAS. 94, 1779-1784 (1997).</li><li>Shastry, M. C. R., Luck, S. D., Roder, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+continuous-flow+capillary+mixing+method+to+monitor+reactions+on+the+microsecond+time+scale.">A continuous-flow capillary mixing method to monitor reactions on the microsecond time scale.</a> Biophysical Journal. 74, 2714-2721 (1998).</li><li>Pollack, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Compactness+of+the+denatured+state+of+a+fast-folding+protein+measured+by+submillisecond+small-angle+x-ray+scattering.">Compactness of the denatured state of a fast-folding protein measured by submillisecond small-angle x-ray scattering.</a> Proceedings of the National Academy of Sciences of the United States of America. 96, 10115-10117 (1999).</li><li>Takahashi, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Folding+of+cytochrome+c+initiated+by+submillisecond+mixing.">Folding of cytochrome c initiated by submillisecond mixing.</a> Nature Structural Molecular Biology. 4, 44-50 (1997).</li><li>Knight, J. B., Vishwanath, A., Brody, J. P., Austin, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hydrodynamic+focusing+on+a+silicon+chip:+Mixing+nanoliters+in+microseconds.">Hydrodynamic focusing on a silicon chip: Mixing nanoliters in microseconds.</a> Physical Review Letters. 80, 3863-3866 (1998).</li><li>DeCamp, S. J., Naganathan, A. N., Waldauer, S. A., Bakajin, O., Lapidus, L. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+Observation+of+Downhill+Folding+of+lambda-Repressor+in+a+Microfluidic+Mixer.">Direct Observation of Downhill Folding of lambda-Repressor in a Microfluidic Mixer.</a> Biophysical Journal. 97, 1772-1777 (2009).</li><li>Waldauer, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ruggedness+in+the+folding+landscape+of+protein+L.">Ruggedness in the folding landscape of protein L.</a> Hfsp Journal. 2, 388-395 (2008).</li><li>Shastry, M. C. R., Roder, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evidence+for+barrier-limited+protein+folding+kinetics+on+the+microsecond+time+scale.">Evidence for barrier-limited protein folding kinetics on the microsecond time scale.</a> Nature Structural Biology. 5, 385-392 (1998).</li><li>Uzawa, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Collapse+and+search+dynamics+of+apomyoglobin+folding+revealed+by+submillisecond+observations+of+alpha-helical+content+and+compactness.">Collapse and search dynamics of apomyoglobin folding revealed by submillisecond observations of alpha-helical content and compactness.</a> Proceedings of the National Academy of Sciences of the United States of America. 101, 1171-1176 (2004).</li><li>Zhu, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evidence+of+Multiple+Folding+Pathways+for+the+Villin+Headpiece+Subdomain.">Evidence of Multiple Folding Pathways for the Villin Headpiece Subdomain.</a> The Journal of Physical Chemistry B. 115, 12632-12637 (2011).</li></ol>

We have nothing to disclose.

Rapid mixing has been a development goal for the field of protein folding for many years because it has long been recognized that molecular processes of biomolecules occur over time scales ranging from picoseconds to seconds. Conventional stopped-flow mixers have dead times of 1-5 ms which is primarily limited by turbulence. Turbulent continuous flow mixers with mixing times of 30-300 &mu;s have been developed over the past 15 years by a small number of groups but generally require high flow rates to achieve these mixing times and therefore use a lot of sample6-8.
This protocol describes the use of microfluidic mixing chips to achieve rapid dilution in the laminar flow regime. There are multiple advantages to working within this regime, but a primary one is the ability to simulate the entire mixing process with high precision which allows one to modify the mixing response. T-mixers developed by other groups with simple geometries have demonstrated mixing times of 100-200 &mu;s that were primarily limited by the size of the channels9-10. By scaling the size of the channels to 10 &mu;m Knight reduced the mixing time to ~10 &mu;s11. Yao and Bakajin showed that small changes in geometry could lead to significant improvements in the mixing time4. However, that work showed that acceleration of mixing can also lead to slower phases as well. The design used in this work12-13 is a compromise between the two best designs in reference4 to minimize the slower exponential decay in denaturant concentration and also to make the feature more reproducible in DRIE etching.
There has been some controversy in the field of ultrarapid mixing over the definition of the mixing time. It is important to recognize the difference between "mixing time" and "dead time". The dead time is defined in a turbulent mixer as the time during which a measurement cannot be made and may in fact be significantly longer than the actual mixing time. It is typically determined by making several measurements of a pseudo-first order bimolecular reaction (such as quenching of tryptophan fluorescence by N bromosuccinate) at various concentrations. The observed exponential decays are fitted to converge at a single value assumed to be t=0 s and the time between that point and the first measured point is the dead time. For laminar flow mixers, measurements can be made during the mixing process so there is no dead time, only a mixing time. The mixing time is simply the time to combine two solutions until sufficient uniformity is reached. We have previously defined the mixing time to be a 90/10 time, the time for the concentration of denaturant to decrease from 90% to 10% of the unmixed value. However, the shape of the mixing curve is not perfectly symmetric with the tail of the decay having a small exponential component. Furthermore, protein folding has a highly non-linear dependence on denaturant concentration so that folding can often commence even if the denaturant is only reduced by two fold. Therefore we have more recently defined the mixing time as the time to reduce the denaturant concentration by 80%. Certainly other definitions can be used depending on the requirements of the experiment.
The use of this mixer to study protein folding has consistently revealed surprising results. Folding of cytochrome c and apomyoglobin have long been studied to have multiple folding steps and early results with continuous flow mixers showed collapse to occur on the ~100 &mu;s timescale10,14-15. Our measurements with this mixer showed there to be at least 2 steps on this timescale, a very fast, likely non-specific, collapse (as measured by Trp fluorescence spectral shift) within the mixing time of the mixer followed by a slower phase (as measured by the quenching of total Trp emission) that is likely the first formation of native structure5. We have also observed a 50 &mu;s process in the B1 domain of protein L, overturning the conventional interpretation that this protein is a 2-state folder13.
An advantage of this rapid mixer is that it can probe the folding of the fastest folding proteins that have usually only been measureable by nanosecond T-jump instruments. T-jump typically requires observation of folding/unfolding relaxation near the melting temperature of the protein and therefore never follows the folding of the entire population. With this mixer we have looked at the folding of &gamma;-repressor (&lambda;6-86) with both Trp total emission and spectral shift and have found evidence that the protein has no barrier to folding under strong folding conditions that were not accessible to previous T-jump measurements12. Finally we have measured the folding of the villin headpiece HP-35, one of the fastest folders yet measured and found that the folding rate measured after mixing is ~5 times slower than measured by T-jump under the same conditions16. This suggests that folding kinetics depends on the starting conditions, overturning a major assumption in the field.

<table border="1">
	<tbody>
		<tr>
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalogue number</td>
			<td>Comments </td>
		</tr>
		<tr>
			<td>AZ P4110 Photoresist</td>
			<td>AZ Electronic Materials</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>AZ 400 K Developer</td>
			<td>AZ Electronic Materials</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Baker PRS 2000 photoresist stripper</td>
			<td>Avantor Performance Materials</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>AquaBond</td>
			<td>AquaBond Technologies</td>
			<td>AquaBond 55</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>500 &mu;m cover wafer</td>
			<td>SENSOR Prep Services</td>
			<td>&nbsp;</td>
			<td>&Oslash;: 100 mm &plusmn; 0.5 mm 
			Thickness: 0.5 mm &plusmn; 0.025 mm 
			Standard Tolerances (Unless Noted) 
			Surface Finish: SI: 4-8&Aring;; SII: 8-12&Aring; 
			Flats: One Primary SEMI-STD; One Scribe 
			Clean and Polished, Both Sides</td>
		</tr>
		<tr>
			<td>170 &mu;m cover wafer</td>
			<td>SENSOR Prep Services</td>
			<td>7980 2G Wafers</td>
			<td>&Oslash;: 100 mm &plusmn; 0.5 mm 
			Thickness: 0.17 mm &plusmn; 0.025 mm 
			Standard Tolerances (Unless Noted) 
			Surface Finish: SI: 4-8&Aring;; SII: 8-12&Aring; 
			Flats: One Primary SEMI-STD; One Scribe 
			Clean and Polished, Both Sides</td>
		</tr>
		<tr>
			<td>Deep reactive ion etcher for oxides</td>
			<td>ULVAC</td>
			<td>ULVAC NL-6000</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Argon Ion Laser</td>
			<td>Cambridge Laser Laboratories</td>
			<td>Lexel 95-SHG</td>
			<td>&lambda;=258 nm</td>
		</tr>
		<tr>
			<td>Argon Ion Laser</td>
			<td>Melles Griot</td>
			<td>&nbsp;</td>
			<td>&lambda;=488 nm</td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Olympus</td>
			<td>IX-51</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Microscope Objective</td>
			<td>Thor Labs</td>
			<td>OFR LMU-40X-UVB 0.5 NA</td>
			<td>&lambda;= 258 nm</td>
		</tr>
		<tr>
			<td>Microscope Objective</td>
			<td>Olympus</td>
			<td>UPLSAPO 60XW 1.2 NA</td>
			<td>&lambda;=488 nm</td>
		</tr>
		<tr>
			<td>Nanopositioner</td>
			<td>Mad City Labs</td>
			<td>Nano-LP100</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Motorized Translation Stage</td>
			<td>Semprex Corp</td>
			<td>KL-Series 12-6436</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>GaAsP Photon Counter</td>
			<td>Hamamatsu</td>
			<td>H7421-40</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Monochrometer</td>
			<td>Horiba Instruments Inc</td>
			<td>MicroHR</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>CCD camera</td>
			<td>Andor Technology</td>
			<td>iDus 420A-BU</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Guanidine hydrochloride (GuHCl)</td>
			<td>Sigma-Aldrich</td>
			<td>G4505</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>

microfluidic mixers for studying protein folding

The process by which a protein folds into its native conformation is highly relevant to biology and human health yet still poorly understood. One reason for this is that folding takes place over a wide range of timescales, from nanoseconds to seconds or longer, depending on the protein1. Conventional stopped-flow mixers have allowed measurement of folding kinetics starting at about 1 ms. We have recently developed a microfluidic mixer that dilutes denaturant ~100-fold in ~8 &mu;s2. Unlike a stopped-flow mixer, this mixer operates in the laminar flow regime in which turbulence does not occur. The absence of turbulence allows precise numeric simulation of all flows within the mixer with excellent agreement to experiment3-4.
Laminar flow is achieved for Reynolds numbers Re &le;100. For aqueous solutions, this requires micron scale geometries. We use a hard substrate, such as silicon or fused silica, to make channels 5-10 &mu;m wide and 10 &mu;m deep (See Figure 1). The smallest dimensions, at the entrance to the mixing region, are on the order of 1 &mu;m in size. The chip is sealed with a thin glass or fused silica coverslip for optical access. Typical total linear flow rates are ~1 m/s, yielding Re~10, but the protein consumption is only ~0.5 nL/s or 1.8 &mu;L/hr. Protein concentration depends on the detection method: For tryptophan fluorescence the typical concentration is 100 &mu;M (for 1 Trp/protein) and for FRET the typical concentration is ~100 nM.
The folding process is initiated by rapid dilution of denaturant from 6 M to 0.06 M guanidine hydrochloride. The protein in high denaturant flows down a central channel and is met on either side at the mixing region by buffer without denaturant moving ~100 times faster (see Figure 2). This geometry causes rapid constriction of the protein flow into a narrow jet ~100 nm wide. Diffusion of the light denaturant molecules is very rapid, while diffusion of the heavy protein molecules is much slower, diffusing less than 1 &mu;m in 1 ms. The difference in diffusion constant of the denaturant and the protein results in rapid dilution of the denaturant from the protein stream, reducing the effective concentration of the denaturant around the protein. The protein jet flows at a constant rate down the observation channel and fluorescence of the protein during folding can be observed using a scanning confocal microscope5.

Watch this Scientific Journal Video about Microfluidic Mixers for Studying Protein Folding at JoVE.com

Microfluidic Mixers for Studying Protein Folding

In this work we explain the fabrication and use of a microfluidic mixer capable of mixing two solutions in ~8 &mu;s. We also demonstrate the use of these mixers with spectroscopic detection using UV fluorescence and fluorescence resonance energy transfer (FRET).

The process by which a protein folds into its native conformation is highly relevant to biology and human health yet still poorly understood. One reason ...

microfluidic-mixers-for-studying-protein-folding

Research

JoVE Journal

Bioengineering

A Microfluidic-based Hydrodynamic Trap for Single Particles

The ability to confine and manipulate single particles in free solution is a key enabling technology for fundamental and applied science. Methods for particle trapping based on optical, magnetic, electrokinetic, and acoustic techniques have led to major advancements in physics and biology ranging from the molecular to cellular level. In this article, we introduce a new microfluidic-based technique for particle trapping and manipulation based solely on hydrodynamic fluid flow. Using this method, we demonstrate trapping of micro- and nano-scale particles in aqueous solutions for long time scales. The hydrodynamic trap consists of an integrated microfluidic device with a cross-slot channel geometry where two opposing laminar streams converge, thereby generating a planar extensional flow with a fluid stagnation point (zero-velocity point). In this device, particles are confined at the trap center by active control of the flow field to maintain particle position at the fluid stagnation point. In this manner, particles are effectively trapped in free solution using a feedback control algorithm implemented with a custom-built LabVIEW code. The control algorithm consists of image acquisition for a particle in the microfluidic device, followed by particle tracking, determination of particle centroid position, and active adjustment of fluid flow by regulating the pressure applied to an on-chip pneumatic valve using a pressure regulator. In this way, the on-chip dynamic metering valve functions to regulate the relative flow rates in the outlet channels, thereby enabling fine-scale control of stagnation point position and particle trapping. The microfluidic-based hydrodynamic trap exhibits several advantages as a method for particle trapping. Hydrodynamic trapping is possible for any arbitrary particle without specific requirements on the physical or chemical properties of the trapped object. In addition, hydrodynamic trapping enables confinement of a "single" target object in concentrated or crowded particle suspensions, which is difficult using alternative force field-based trapping methods. The hydrodynamic trap is user-friendly, straightforward to implement and may be added to existing microfluidic devices to facilitate trapping and long-time analysis of particles. Overall, the hydrodynamic trap is a new platform for confinement, micromanipulation, and observation of particles without surface immobilization and eliminates the need for potentially perturbative optical, magnetic, and electric fields in the free-solution trapping of small particles.

In this article, we present a microfluidic-based method for particle confinement based on hydrodynamic flow. We demonstrate stable particle trapping at a fluid stagnation point using a feedback control mechanism, thereby enabling confinement and micromanipulation of arbitrary particles in an integrated microdevice.

The ability to confine and manipulate single particles in free solution is a key enabling technology for fundamental and applied science.  Methods for ...

Thermodynamics of Membrane Protein Folding Measured by Fluorescence Spectroscopy

Membrane protein folding is an emerging topic with both fundamental and health-related significance. The abundance of membrane proteins in cells underlies the need for comprehensive study of the folding of this ubiquitous family of proteins. Additionally, advances in our ability to characterize diseases associated with misfolded proteins have motivated significant experimental and theoretical efforts in the field of protein folding. Rapid progress in this important field is unfortunately hindered by the inherent challenges associated with membrane proteins and the complexity of the folding mechanism. Here, we outline an experimental procedure for measuring the thermodynamic property of the Gibbs free energy of unfolding in the absence of denaturant, &Delta;G&deg;H2O, for a representative integral membrane protein from E. coli. This protocol focuses on the application of fluorescence spectroscopy to determine equilibrium populations of folded and unfolded states as a function of denaturant concentration. Experimental considerations for the preparation of synthetic lipid vesicles as well as key steps in the data analysis procedure are highlighted. This technique is versatile and may be pursued with different types of denaturant, including temperature and pH, as well as in various folding environments of lipids and micelles. The current protocol is one that can be generalized to any membrane or soluble protein that meets the set of criteria discussed below.

This video article details the experimental procedure for obtaining the Gibbs free energy of membrane protein folding by tryptophan fluorescence.

Membrane protein folding is an emerging topic with both fundamental and health-related significance.  The abundance of membrane proteins in cells ...

Fluorescence detection methods for microfluidic droplet platforms

The development of microfluidic platforms for performing chemistry and biology has in large part been driven by a range of potential benefits that accompany system miniaturisation. Advantages include the ability to efficiently process nano- to femoto- liter volumes of sample, facile integration of functional components, an intrinsic predisposition towards large-scale multiplexing, enhanced analytical throughput, improved control and reduced instrumental footprints.1
In recent years much interest has focussed on the development of droplet-based (or segmented flow) microfluidic systems and their potential as platforms in high-throughput experimentation.2-4 Here water-in-oil emulsions are made to spontaneously form in microfluidic channels as a result of capillary instabilities between the two immiscible phases. Importantly, microdroplets of precisely defined volumes and compositions can be generated at frequencies of several kHz. Furthermore, by encapsulating reagents of interest within isolated compartments separated by a continuous immiscible phase, both sample cross-talk and dispersion (diffusion- and Taylor-based) can be eliminated, which leads to minimal cross-contamination and the ability to time analytical processes with great accuracy. Additionally, since there is no contact between the contents of the droplets and the channel walls (which are wetted by the continuous phase) absorption and loss of reagents on the channel walls is prevented.
Once droplets of this kind have been generated and processed, it is necessary to extract the required analytical information. In this respect the detection method of choice should be rapid, provide high-sensitivity and low limits of detection, be applicable to a range of molecular species, be non-destructive and be able to be integrated with microfluidic devices in a facile manner. To address this need we have developed a suite of experimental tools and protocols that enable the extraction of large amounts of photophysical information from small-volume environments, and are applicable to the analysis of a wide range of physical, chemical and biological parameters. Herein two examples of these methods are presented and applied to the detection of single cells and the mapping of mixing processes inside picoliter-volume droplets. We report the entire experimental process including microfluidic chip fabrication, the optical setup and the process of droplet generation and detection.

Droplet-based microfluidic platforms are promising candidates for high throughput experimentation since they are able to generate picoliter, self-compartmentalized vessels inexpensively at kHz rates. Through integration with fast, sensitive and high resolution fluorescence spectroscopic methods, the large amounts of information generated within these systems can be efficiently extracted, harnessed and utilized.

The development of microfluidic platforms for performing chemistry and biology has in large part been driven by a range of potential benefits that ...

High-throughput Protein Expression Generator Using a Microfluidic Platform

Rapidly increasing fields, such as systems biology, require the development and implementation of new technologies, enabling high-throughput and high-fidelity measurements of large systems. Microfluidics promises to fulfill many of these requirements, such as performing high-throughput screening experiments on-chip, encompassing biochemical, biophysical, and cell-based assays1. Since the early days of microfluidics devices, this field has drastically evolved, leading to the development of microfluidic large-scale integration2,3. This technology allows for the integration of thousands of micromechanical valves on a single device with a postage-sized footprint (Figure 1). We have developed a high-throughput microfluidic platform for generating in vitro expression of protein arrays (Figure 2) named PING (Protein Interaction Network Generator). These arrays can serve as a template for many experiments such as protein-protein 4, protein-RNA5 or protein-DNA6 interactions.
The device consist of thousands of reaction chambers, which are individually programmed using a microarrayer. Aligning of these printed microarrays to microfluidics devices programs each chamber with a single spot eliminating potential contamination or cross-reactivity Moreover, generating microarrays using standard microarray spotting techniques is also very modular, allowing for the arraying of proteins7, DNA8, small molecules, and even colloidal suspensions. The potential impact of microfluidics on biological sciences is significant. A number of microfluidics based assays have already provided novel insights into the structure and function of biological systems, and the field of microfluidics will continue to impact biology.

We present a microfluidic approach for the expression of protein arrays. The device consists of thousands of reaction chambers controlled by micro-mechanical valves. The microfluidic device is mated to a microarray-printed gene library. These genes are then transcribed and translated on-chip, resulting in a protein array ready for experimental use.

Rapidly increasing fields, such as systems biology, require the development and implementation of new technologies, enabling high-throughput and ...

A Microfluidic Device for Studying Multiple Distinct Strains

The study of cell responses to environmental changes poses many experimental challenges: cells need to be imaged under changing conditions, often in a comparative manner. Multiwell plates are routinely used to compare many different strains or cell lines, but allow limited control over the environment dynamics. Microfluidic devices, on the other hand, allow exquisite dynamic control over the surrounding conditions, but it is challenging to image and distinguish more than a few strains in them. Here we describe a method to easily and rapidly manufacture a microfluidic device capable of applying dynamically changing conditions to multiple distinct yeast strains in one channel. The device is designed and manufactured by simple means without the need for soft lithography. It is composed of a Y-shaped flow channel attached to a second layer harboring microwells. The strains are placed in separate microwells, and imaged under the exact same dynamic conditions. We demonstrate the use of the device for measuring protein localization responses to pulses of nutrient changes in different yeast strains.

We present a simple method to produce microfluidic devices capable of applying similar dynamic conditions to multiple distinct strains, without the need for a clean room or soft lithography.

The study of cell responses to environmental changes poses many experimental challenges: cells need to be imaged under changing conditions, often in a ...

A Microfluidic Chip for ICPMS Sample Introduction

This protocol discusses the fabrication and usage of a disposable low cost microfluidic chip as sample introduction system for inductively coupled plasma mass spectrometry (ICPMS). The chip produces monodisperse aqueous sample droplets in perfluorohexane (PFH). Size and frequency of the aqueous droplets can be varied in the range of 40 to 60 &#181;m and from 90 to 7,000&#160;Hz, respectively. The droplets are ejected from the chip with a second flow of PFH and remain intact during the ejection. A custom-built desolvation system removes the PFH and transports the droplets into the ICPMS. Here, very stable signals with a narrow intensity distribution can be measured, showing the monodispersity of the droplets. We show that the introduction system can be used to quantitatively determine iron in single bovine red blood cells. In the future, the capabilities of the introduction device can easily be extended by the integration of additional microfluidic modules.

We present a discrete droplet sample introduction system for inductively coupled plasma mass spectrometry (ICPMS). It is based on a cheap and disposable microfluidic chip that generates highly monodisperse droplets in a size range of 40&#8722;60 &#181;m at frequencies from 90 to 7,000 Hz.

This protocol discusses the fabrication and usage of a disposable low cost microfluidic chip as sample introduction system for inductively coupled plasma ...

Studying Protein Function and the Role of Altered Protein Expression by Antibody Interference and Three-dimensional Reconstructions

A strict management of protein expression is not only essential to every organism alive, but also an important strategy to investigate protein functions in cellular models. Therefore, recent research invented different tools to target protein expression in mammalian cell lines or even animal models, including RNA and antibody interference. While the first strategy has gathered much attention during the past two decades, peptides &#160;mediating a translocation of antibody cargos across cellular membranes and into cells, obtained much less interest. In this publication, we provide a detailed protocol how to utilize a peptide carrier named Chariot in human embryonic kidney cells as well as in primary hippocampal neurons to perform antibody interference experiments and further illustrate the application of three-dimensional reconstructions in analyzing protein function. Our findings suggest that Chariot is, probably due to its nuclear localization signal, particularly well-suited to target proteins residing in the soma and the nucleus. Remarkably, when applying Chariot to primary hippocampal cultures, the reagent turned out to be surprisingly well accepted by dissociated neurons.

Controlling protein expression is not only essential to every organism alive, but also an important strategy to investigate protein functions in cellular models. The protocol presented shows the application of antibody interference in mammalian cells including primary hippocampal neurons and demonstrates the use of three-dimensional reconstructions in studying protein function.

A strict management of protein expression is not only essential to every organism alive, but also an important strategy to investigate protein functions ...

Fully Automated Centrifugal Microfluidic Device for Ultrasensitive Protein Detection from Whole Blood

Enzyme-linked immunosorbent assay (ELISA) is a promising method to detect small amount of proteins in biological samples. The devices providing a platform for reduced sample volume and assay time as well as full automation are required for potential use in point-of-care-diagnostics. Recently, we have demonstrated ultrasensitive detection of serum proteins, C-reactive protein (CRP) and cardiac troponin I (cTnI), utilizing a lab-on-a-disc composed of TiO2 nanofibrous (NF) mats. It showed a large dynamic range with femto molar (fM) detection sensitivity, from a small volume of whole blood in 30 min. The device consists of several components for blood separation, metering, mixing, and washing that are automated for improved sensitivity from low sample volumes. Here, in the video demonstration, we show the experimental protocols and know-how for the fabrication of NFs as well as the disc, their integration and the operation in the following order: processes for preparing TiO2 NF mat; transfer-printing of TiO2 NF mat onto the disc; surface modification for immune-reactions, disc assembly and operation; on-disc detection and representative results for immunoassay. Use of this device enables multiplexed analysis with minimal consumption of samples and reagents. Given the advantages, the device should find use in a wide variety of applications, and prove beneficial in facilitating the analysis of low abundant proteins.

This protocol demonstrates how to achieve femto molar detection sensitivity of proteins in 10 &#181;L of whole blood within 30 min. This can be achieved by using electrospun nanofibrous mats integrated in a lab-on-a-disc, which offers high surface area as well as effective mixing and washing for enhanced signal-to-noise ratio.

Enzyme-linked immunosorbent assay (ELISA) is a promising method to detect small amount of proteins in biological samples. The devices providing a platform ...

Designing Microfluidic Devices for Studying Cellular Responses Under Single or Coexisting Chemical/Electrical/Shear Stress Stimuli

Microfluidic devices are capable of creating a precise and controllable cellular micro-environment of pH, temperature, salt concentration, and other physical or chemical stimuli. They have been commonly used for in vitro cell studies by providing in vivo like surroundings. Especially, how cells response to chemical gradients, electrical fields, and shear stresses has drawn many interests since these phenomena are important in understanding cellular properties and functions. These microfluidic chips can be made of glass substrates, silicon wafers, polydimethylsiloxane (PDMS) polymers, polymethylmethacrylate (PMMA) substrates, or polyethyleneterephthalate (PET) substrates. Out of these materials, PMMA substrates are cheap and can be easily processed using laser ablation and writing. Although a few microfluidic devices have been designed and fabricated for generating multiple, coexisting chemical and electrical stimuli, none of them was considered efficient enough in reducing experimental repeats, particular for screening purposes. In this report, we describe our design and fabrication of two PMMA-based microfluidic chips for investigating cellular responses, in the production of reactive oxygen species and the migration, under single or coexisting chemical/electrical/shear stress stimuli. The first chip generates five relative concentrations of 0, 1/8, 1/2, 7/8, and 1 in the culture regions, together with a shear stress gradient produced inside each of these areas. The second chip generates the same relative concentrations, but with five different electric field strengths created within each culture area. These devices not only provide cells with a precise, controllable micro-environment but also greatly increase the experimental throughput.

Micro-fabricated devices integrated with fluidic components provide an in vitro platform for cell studies mimicking the in vivo micro-environment. We developed polymethylmethacrylate-based microfluidic chips for studying cellular responses under single or coexisting chemical/electrical/shear stress stimuli.

Microfluidic devices are capable of creating a precise and controllable cellular micro-environment of pH, temperature, salt concentration, and other ...

Multi-step Variable Height Photolithography for Valved Multilayer Microfluidic Devices

Microfluidic systems have enabled powerful new approaches to high-throughput biochemical and biological analysis. However, there remains a barrier to entry for non-specialists who would benefit greatly from the ability to develop their own microfluidic devices to address research questions. Particularly lacking has been the open dissemination of protocols related to photolithography, a key step in the development of a replica mold for the manufacture of polydimethylsiloxane (PDMS) devices. While the fabrication of single height silicon masters has been explored extensively in literature, fabrication steps for more complicated photolithography features necessary for many interesting device functionalities (such as feature rounding to make valve structures, multi-height single-mold patterning, or high aspect ratio definition) are often not explicitly outlined. 
Here, we provide a complete protocol for making multilayer microfluidic devices with valves and complex multi-height geometries, tunable for any application. These fabrication procedures are presented in the context of a microfluidic hydrogel bead synthesizer and demonstrate the production of droplets containing polyethylene glycol (PEG diacrylate) and a photoinitiator that can be polymerized into solid beads. This protocol and accompanying discussion provide a foundation of design principles and fabrication methods that enables development of a wide variety of microfluidic devices. The details included here should allow non-specialists to design and fabricate novel devices, thereby bringing a host of recently developed technologies to their most exciting applications in biological laboratories.

Multilayer microfluidic devices often involve the fabrication of master molds with complex geometries for functionality. This article presents a complete protocol for multi-step photolithography with valves and variable height features tunable to any application. As a demonstration, we fabricate a microfluidic droplet generator capable of producing hydrogel beads.

Microfluidic systems have enabled powerful new approaches to high-throughput biochemical and biological analysis. However, there remains a barrier to ...