Name: fluorescence detection methods for microfluidic droplet platforms
Uploaded: 2011-12-10T14:00:00
Duration: 14 min 16 s
Description: Watch this Scientific Journal Video about Fluorescence detection methods for microfluidic droplet platforms at JoVE.com

The authors would like to acknowledge the support of the following research councils and grants: EPSRC, HFSP, National Research Foundation of Korea (Grant Number R11-2009-044-1002-0K20904000004-09A050000410).

1. Microchip fabrication
<ol>
	<li>SU8 master fabrication. 
	To obtain microfluidic channels of 40 &mu;m height, spin coat SU8-50 (MicroChem Corp.) on a Si wafer at 3000 rpm for 30 s. Soft bake on a hot-plate at 65&deg;C for 5 minutes and 95&deg;C for 30 minutes to evaporate the solvent and densify the film. After cooling down, expose the SU8 resist with 360-440 nm radiation at 300 mJ/2 under the appropriate mask. Following exposure, bake the wafer at 65&deg;C for 2 minutes and 95&deg;C for 4 minutes. After cooling, develop the unexposed areas (Micrpoposit EC solvent, Chestech) for 5 minutes, with stirring.5 Use fresh EC solvent to rinse the wafer and then dry it under gas to generate a clean surface. Treat the wafer with hexane and methanol washes. Complete the SU8 master fabrication process with a final step of evaporating trichloro(1,1,2,2-perfluoocytl)silane onto the surface (in a desiccator for 40 minutes).</li>
	<li>PDMS curing, dicing and hole-punching. 
	To form structured microfluidic substrates, we mix Sylgard 184 (Dow Corning) in a 10:1 (w/w) ratio of resin to crosslinker and then pour over the master. Degas in a desiccator, and then cure the device (top layer) for one hour at 65&deg;C. Cut the cured PDMS and peel it off the master. Create access holes for fluidic tubing using a biopsy punch. Cure another layer of PDMS (8 g of the resin evenly spread in a 10x10 cm Petri dish) for 20 minutes at 65&deg;C. Place the prepared top layer on the bottom layer and then cure overnight at 65&deg;C. Peel the chips from the Petri dish and dice for use.</li>
</ol>
	
2. Experimental setup
<ol>
	<li>Microfluidic setup
		<ol>
			<li>Fill syringes (plastic or glass from Beckton, Dickinson and Company or from SGE Analytical Science) with the required solutions, ensuring no air bubbles remain in any parts of the tubing. For droplet generation two phases are used. The aqueous (dispersed) phase contains the reagents of interest (for example a suspension of cells in growth medium, or a solution of molecular fluorophores). The oil phase, which in this case acts as the continuous phase, is generally formed from a mixture of oil and surfactant, e.g. 2% Raindance surfactant (RainDance Technologies) in fluorinated oil (3M Fluorinert Electronic Liquid FC-40); or 2% Span 80 (Croda) in Mineral oil.</li>
			<li>Fit tubing (Polyethylene tubing, Harvard Apparatus) of the same internal diameter as the syringe needle tips on one end to the needles. The tubing should be cut to the shortest possible length to minimize flow instabilities due to vibrational perturbations.</li>
			<li>Connect the other ends of the tubing directly to the holes punched in the PDMS substrate. More complex solutions, such as the use of silica capillary ports or connectors (e.g Upchurch Nanoports) can also be implemented for a more robust interface between the microfluidic device and the tubing. Connect the outlet port of the chip to tubing and direct into a collector (for waste collection or further analyte processing).</li>
			<li>Mount the syringes on a syringe pump (e.g. PHD 4400 Hpsi programmable syringe pumps, Harvard Apparatus) and define the desired infusing flow-rate for each phase (usually on the order of microliters per minute). A minimum ratio of oil/aqueous flow-rates of 1 is recommended to avoid wetting of the PDMS by the aqueous phase, which would lead to unstable droplet formation (depending on device geometry). For the same reason, it is also recommended to first flush the oil phase alone, through any microfluidic channels, before introduction of the aqueous phase. The final setup used for the experiments herein is illustrated in Figure 2a.</li>
		</ol>
	</li>
	<li>Optical system setup
		<ol>
			<li>To probe small volumes (down to a few picoliters) within the microfluidic channels with high spatial and temporal resolution use a confocal microscope (with automated submicron positioning capacity).</li>
			<li>Use water-soluble fluorescent molecules with ideally high quantum yields. Excite the molecules using a laser operating at a wavelength that matches the absorption of the fluorophore involved. For example, common fluorescent molecules such as fluorescein 5-isothiocyanate (FITC) and Alexa-Fluor 488 can be efficiently excited using a 488 nm diode laser (e.g. PicoQuant Gmbh, LDH-P-C-485). Fluorescent molecules, such as Texas Red or Allophycocyanin (APC) can be efficiently exited at 633 nm using a He/Ne laser (e.g. Newport R-31425).</li>
			<li>Use pulsed lasers operating at repetition rates of several MHz (with several ns pulse separations) and fluorophores with sufficiently differentiated lifetime properties for Fluorescence Lifetime Imaging (FLIM) experiments.</li>
			<li>Use beam-steering mirrors and optics to control the beam height as well as beam direction.</li>
			<li>Use a dichroic mirror to reflect the laser beam into the objective lens. If two lasers are used simultaneously, a dual-band dichroic mirror can be installed to allow reflection of the two laser beams (e.g. for 488 and 633 nm beams a z488/633rdc mirror from Chroma Technology Corporation is ideal).</li>
			<li>Use an infinity-corrected, high numerical aperture (NA) microscope objective (i.e. Olympus 60 x /1.2 NA, water immersion) to bring the laser light to a tight focus within a microfluidic channel. When working with tall microchannels (&gt;100 &mu;m) the objective used must be appropriately selected to ensure that its working distance effectively covers the full height of the microchannel.</li>
			<li>Collect fluorescence emitted by the sample (within the microfluidic channel) using the same high NA objective and transmitted through the same dichroic mirror.</li>
			<li>Remove any residual excitation light using a single or dual-band emission filter (e.g. a z488/633 filter from Chroma Technology Corporation).</li>
			<li>Focus the fluorescence emission onto a 75 &mu;m pinhole using a plano-convex lens (e.g. +50.2 F; Newport Ltd.). Position the pinhole in the confocal plane of the microscope objective.</li>
			<li>Use another dichroic mirror (e.g. 630dcxr, Chroma Technology Corporation) to split the fluorescent signal onto two detectors.</li>
			<li>Filter the fluorescence reflected by the dichroic mirror with an emission filter (e.g. an hq540/80 m filter from Chroma Technology Corporation) and focus it with a plano-convex lens (e.g. f = 30.0, i.d. 25.4 mm, Thorlabs) onto the first ('green') detector. For experiments involving a secondary red fluorophore filter the fluorescence transmitted by the dichroic mirror with another emission filter (e.g. an hq640lp filter from Chroma Technology Corporation) and then focus it with another plano-convex lens onto the second ('red') detector.</li>
			<li>Use avalanche photodiodes (e.g. AQR-141, EG&amp;G, Perkin-Elmer) for detection of fluorescence photons. The final setup is illustrated in Figure 2b.</li>
		</ol>
	</li>
</ol>
3. Droplet generation and data acquisition
<ol>
	<li>Droplet generation methods
		<ol>
			<li>You can use two main configurations of microchannels to generate droplets: a flow focusing or T-junction format (Figure 3a and 3b).6,7 Both methods are equivalent as long as the droplets formed are as wide as the microchannel itself. As a result of the shear forces that arise from using these geometries to bring the two immiscible fluids together and the subsequent capillary instabilities that develop in the interface, monodisperse droplets (as small as a few femtoliters) are generated spontaneously.</li>
			<li>You can encapsulate reagents in different ways: reagents can be prepared and mixed off chip to the desired mixture/concentration, or they can be brought separately into the chip and then combined together prior to droplet formation (Figure 3c). This last method provides for higher flexibility since mixtures/concentrations can be modified by simply tuning the relative flow-rates. It is noted that for cell encapsulation, the cell suspension in the syringe should be stirred to avoid condensation of the cells over the timescale of the experiment.</li>
		</ol>
	</li>
	<li>Fluorescence data acquisition.
		<ol>
			<li>Couple the electronic signal from the avalanche photodiode detector (described in 2.2.12) to a multifunction DAQ device for data logging (e.g. a PCI 6602, National Instruments), running on a personal computer.</li>
			<li>For FLIM experiments connect the detectors to a Time-Correlated Single Photon Counting (TCSPC) card (e.g. TimeHarp 100, PicoQuant GmbH) running on a separate PC. This card allows for fluorescence lifetime data to be obtained using a TCSPC methodology: each detected fluorescent photon is correlated to an incident laser pulse, and therefore each emitted fluorescent photon is assigned a delay time. This data can be fitted to a single or multi-exponential decay curve to obtain an estimation of the fluorophore/s radiative rate coefficient/s. Deconvolute the raw data with the instrument response function (IRF) of the detector: this is obtained using a low concentration Auramine-O solution (which exhibits a fluorescence lifetime of a few picoseconds).8</li>
		</ol>
	</li>
</ol>
4. Cell recognition and mapping of mixing within droplets
<ol>
	<li>Cell recognition and mapping of mixing within droplets
		<ol>
			<li>Use Escherichia coli Top 10 strain for the viability assay experiment. Culture the cells in Luria-Bertani broth overnight and match the optical density to 0.5 prior to the experiments.</li>
			<li>Use 0.4 &mu;M SYTO9 and 1 &mu;M propidium iodide for detecting the viability of the cells. Both are DNA-intercalating dyes and their fluorescence intensity increases by over 20 folds upon binding to DNA. SYTO9 is a green fluorescent dye that is membrane permeable and propidium iodide is a red fluorescent dye which is membrane impermeable. Thus live cells fluoresce 'green' while dead cells exhibit both 'green' and 'red' emissions.</li>
			<li>Use the setup introduced in 2.2) for green/red fluorescence detection.</li>
			<li>Use a portable, mini-magnetic stirrer (Utah Biodiesel Supply, Utah) to stir the cell suspensions within a 3mL BD plastipak syringe fitted with a 7mm magnetic stir bar to prevent cell sedimentation .</li>
		</ol>
	</li>
	<li>Mapping of mixing events using FLIM
		<ol>
			<li>Focus the optical probe volume at half the height of a microchannel along which droplets are flowing.</li>
			<li>Form droplets from two aqueous solutions (as in Figure 3c), each containing a (non-interacting) fluorophore with different characteristic fluorescent lifetimes.</li>
			<li>Beginning from one side of the channel, carry out each experiment along the entire width of the channel at 1 &mu;m intervals. The channel edges can be easily identified as the fluorescence intensity drops drastically once the laser beam is focused in their proximity.</li>
			<li>Implement an algorithm to differentiate signal bursts (associated with droplets) from the noise background of the oil phase and to establish the duration of each burst.</li>
			<li>Implement a second algorithm to extract the delay time and intensity values along the length of each droplet at the particular width where the experiment has been carried out.9</li>
			<li>Then use a Maximum Likelihood Estimator (MLE) algorithm to evaluate the fluorescence lifetime for each droplet in the experiment.8 Averaging the lifetime values for all the droplets in the experiment, reduces the final error of the MLE calculation (the more droplets probed, the smaller the error).</li>
			<li>Once a lifetime trajectory has been obtained for each width, combine all the trajectories in a 2D map. Since each lifetime value is associated with a particular mixture of the two fluorophores, a concentration (or mixing) map can thus be obtained.</li>
			<li>Optionally, a 3D map of droplet mixing could be easily obtained by repeating this protocol at different positions along the height of the microfluidic channel. </li>
		</ol>
	</li>
</ol>
5. Data analysis
<ol>
	<li>Reinterpret the raw experimental data files into the appropriate computing language so that they can be further analyzed (e.g. text files that contain the variation of photon counts over time, can be readily extracted into Matlab for data processing and analysis). You might have to write specific codes in order to access the data contained in files of more complex experiments (such as those for FLIM) or in order to run MLE algorithms or build 2D maps of mixing patters from lifetime trajectories.</li>
</ol>
6. Representative Results
Cell recognition
Typical examples of the variation of photon counts as a function of time for the cell-based experiments are shown in Figures 4a and c, over a period of two hundred milliseconds. Importantly the Luria&#x2013;Bertani broth (LB medium) acts as a weakly fluorescing background defining the aqueous droplet boundaries, whilst distinct photon bursts, corresponding to the presence of individual cells, can be distinguished on top of this background. The LB fluorescent background is characterised by an approximately rectangular cross-section (marked by the red and green dotted lines).
The full width half maximum (FWHM) of fluorescent bursts arising from E. coli cells is 30-fold shorter than the background droplet events, which is consistent with the relative length of droplets (40 microns, corresponding to a volume of 30 picolitres) and E. coli cells (1.5&#x2013;2.0 microns).10
With the dual channel detection system, the existence of live cells or dead cells can be distinguished from analysis of the green and red channels. Figures 4 b and d show the probability distribution describing the number of cells per droplet. 
Mapping of mixing events using FLIM
The molecular fluorescence lifetime is an intrinsic property of an individual fluorophore that is affected only by its chemical environment. Accordingly its measurements can be used to obtain environmental information (such as solution pH, temperature, polarity and viscosity) with superior resolution and precision than time integrated fluorescence measurements, which are dependent on experimental factors (such as concentration, excitation intensity, and optical collection efficiency).9,11
As presented elsewhere,8 it is possible to map the mixing patterns inside droplets by using two fluorophores with different lifetime values. In a mixture containing two (non-interacting) fluorescent species the decay will take on a biexponential form as is shown in equation 1: 
<img src="/files/ftp_upload/3437/3437eq1.jpg" alt="Equation 1" />
The individual lifetimes are defined as &tau;1 and &tau;2; and the amplitude of each component is given by &alpha;1 and &alpha;2 respectively. The average lifetime can then be defined as follows: 
<img src="/files/ftp_upload/3437/3437eq2.jpg" alt="Equation 2" />
Where &beta;1 and &beta;2 are the fraction of each component and are defined as follows: 
<img src="/files/ftp_upload/3437/3437eq3.jpg" alt="Equation 3" />
Since the probe volume is fixed and droplets are flowing across that position, a trajectory of the lifetime values along the length of the droplets at that particular width can be thus obtained. A characteristic trajectory for a particular width, averaged among 6000 droplets, can be observed in Figure 5a.
Combining the trajectories throughout the entire width of the channel leads to the formation of a 2D concentration (or mixing) map. A typical result is presented in Figure 5b.
By averaging the results among a large number of droplets, the standard error of the results obtained can be greatly reduced (1% of standard error with a 95% confidence interval for Figure 5a). The determination method of the lifetime trajectory assumes that all droplets are practically identical. This is proven to be largely correct for two main reasons: if droplets were significantly different, the averaged lifetime trajectories obtained would have flatter, less diverging profiles (and sharp differences in lifetime along the length of the averaged droplets are consistently detected). Furthermore, the results are reproducible regardless of the individual droplets analyzed or the amount of droplets used in the calculations.8 
<img src="/files/ftp_upload/3437/3437fig1.jpg" alt="Figure 1" /> 
Figure 1. process flow on the microfluidic chip fabrication.
<img src="/files/ftp_upload/3437/3437fig2.jpg" alt="Figure 2" /> 
Figure 2. a) Diagram representing the syringe pumps and syringes, connected to the microfluidic chip for droplet formation; b) Schematic representation of a typical optical setup for a two laser &#x2013; two fluorophore (green and red) excitation and detection. DC = Dichroic mirror, EM = Emission filter, L = Lens, PH = Pin hole, APD = Avalanche photodiode detector.
<img src="/files/ftp_upload/3437/3437fig3.jpg" alt="Figure 3" /> 
Figure 3. On-chip droplet formation strategies: a) flow-focusing configuration; b) T-junction configuration. c) On chip encapsulation in droplets of two originally separate aqueous reagents.
<img src="/files/ftp_upload/3437/3437fig4.jpg" alt="Figure 4" /> 
Figure 4. Optical readout of 0.2 s traces recorded for (a) dead and (c) live cells (flow rates for cell suspension: 1 &mu;l min-1, oil: 2 &mu;l min-1). Each arch-shaped signal corresponds to the weakly fluorescent LB medium that forms the aqueous droplet, with droplet boundaries marked with dashed lines. Green signals represent readout under 633nm and red signals over 633 nm wavelengths. Cells are distinguished by the vertical spike arising from the DNA intercalating dyes. Probability distribution for cellular occupancy within single droplets for (b) dead cells and (d) live cells. 
<img src="/files/ftp_upload/3437/3437fig5.jpg" alt="Figure 5" /> 
Figure 5. a) Typical averaged lifetime trajectory along the length of a droplet at a particular width; b) Typical representation of the combination of all the averaged trajectories probed along the full width of the droplets into a 2D lifetime (or concentration, expressed as % FITC / % Str-AF488) map.

<ol>
	<li>Whitesides, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+origins+and+the+future+of+microfluidics.">The origins and the future of microfluidics.</a> Nature. 442, 368-368 (2006).</li><li>Zheng, B., Angew, I. s. m. a. g. i. l. o. v., F, 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+Microfluidic+Approach+for+Screening+Submicroliter+Volumes+against+Multiple+Reagents+by+Using+Preformed+Arrays+of+Nanoliter+Plugs+in+a+Three-Phase+Liquid/Liquid/Gas+Flow.">A Microfluidic Approach for Screening Submicroliter Volumes against Multiple Reagents by Using Preformed Arrays of Nanoliter Plugs in a Three-Phase Liquid/Liquid/Gas Flow.</a> Chem. Int. Ed. 44, 2520-2520 (2005).</li><li>Huebner, A., Sharma, S., Srisa-Art, M., Hollfelder, F., Edel, J. B., Demello, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microdroplets:+A+sea+of+applications.">Microdroplets: A sea of applications.</a> Lab on a Chip. 8, 1244-1244 (2008).</li><li>deMello, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Control+and+detection+of+chemical+reactions+in+microfluidic+systems.">Control and detection of chemical reactions in microfluidic systems.</a> Nature. 442, 394-394 (2006).</li><li>Thorsen, T., Roberts, R. W., Arnold, F. H., Quake, S. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+pattern+formation+in+a+vesicle-generating+microfluidic+device.">Dynamic pattern formation in a vesicle-generating microfluidic device.</a> Phys. Rev. Lett. 86, 4163-4163 (2001).</li><li>Anna, S. L., Bontoux, N., Stone, H. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Formation+of+dispersions+using+"flow+focusing"+in+microchannels.">Formation of dispersions using "flow focusing" in microchannels.</a> Appl. Phys. Lett. 82, 364-364 (2003).</li><li>Casadevall i Solvas, X., Srisa-Art, M., deMello, A. J., Edel, B. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mapping+of+Fluidic+Mixing+in+Microdroplets+with+1+&#x03BC;s+Time+Resolution+Using+Fluorescence+Lifetime+Imaging.">Mapping of Fluidic Mixing in Microdroplets with 1 &#x03BC;s Time Resolution Using Fluorescence Lifetime Imaging.</a> Anal. Chem. 82, 3950-3950 (2010).</li><li>Robinson, T. A., Schaerli, Y., Wootton, R., Hollfelder, F., Dunsby, C., Baldwin, G., Neil, M., French, P., deMello, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Removal+of+background+signals+from+fluorescence+thermometry+measurements+in+PDMS+microchannels+using+fluorescence+lifetime+imaging.">Removal of background signals from fluorescence thermometry measurements in PDMS microchannels using fluorescence lifetime imaging.</a> Lab. Chip. 9, 3437-3441 (2009).</li><li>Huebner, A., Srisa-Art, M., Holt, D., Abell, C., Hollfelder, F., deMello, A. J., Edel, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+detection+of+protein+expression+in+single+cells+using+droplet+microfluidics.">Quantitative detection of protein expression in single cells using droplet microfluidics.</a> Chem. Commun. 1218, (2007).</li><li>Edel, J. B., Eid, J. S., Meller, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Accurate+single+molecule+FRET+efficiency+determination+for+surface+immobilized+DNA+using+maximum+likelihood+calculated+lifetimes.">Accurate single molecule FRET efficiency determination for surface immobilized DNA using maximum likelihood calculated lifetimes.</a> J. Phys. Chem. B. 73, 2986-2990 (2007).</li></ol>

No conflicts of interest declared.

We have described the fabrication of a microfluidic device, an experimental setup and associated protocols for microdroplet formation and reagent encapsulation and optical detection of the processed droplets. The two examples selected (the detection of single cells in droplets and the mapping of mixing processes inside flowing droplets) represent common applications that are currently being investigated with droplet microfluidics.
As the presented experiments illustrate, the use of droplet microfluidic platforms present certain attractive characteristics, such as high throughput, quasi-perfect confinement, high reproducibility&#x2026; The large amount of information produced and the high speed at which this information is being generated, though, require the use of fast detection methods with high spatiotemporal resolution if the full advantage is to be gained from these miniaturized platforms. In this case we demonstrate that this is possible using highly precise fluorescent spectroscopic techniques. In particular, the FLIM detection technique (which makes use of a pulsed laser with repetition periods as short as a few ns) reveals the capacity to obtain very fast information from rapidly flowing droplets (of around 200 per second). In this case the time resolution of detected events was as low as 1 &mu;s. In terms of limits in droplet size and concentration, the use of larger numerical aperture lenses and larger power lasers would easily permit the detection of nM concentrations within droplets as small as 5 &mu;m (the limitations in droplet size being imposed by the resolutions of the photolithographic fabrication process and of the submicron positioning stage). 
Microfluidic droplets are ideal candidates to carry out large scale experimentation involving small amounts of reagents, down to single target/event. Current limitations of this technology involve the difficulty to generate droplets from a large variety (tens or hundreds) of different sources in a high-throughput manner. Additionally, the difficulty to target and manipulate each single droplet generated imposes severe limitations to the practical applicability of these techniques. Therefore, these issues are at the center of important research efforts aimed at developing the necessary technological solutions that will enable microfluidic droplet platforms to become a standard reference method of research and analysis in high throughput applications.

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Type</td>
 <td>Company</td>
 <td>Catalogue number</td>
 </tr>
 <tr>
 <td>Fluorescein 5-isothiocyanate (FITC)</td>
 <td>Reagent</td>
 <td>Sigma-Aldrich</td>
 <td>F3651</td>
 </tr>
 <tr>
 <td>Streptavidin-Alexa Fluor 488</td>
 <td>Reagent</td>
 <td>Molecular Probes</td>
 <td>S11223</td>
 </tr>
 <tr>
 <td>Perfluorodecalin </td>
 <td>Reagent </td>
 <td>Sigma-Aldrich</td>
 <td>P9900</td>
 </tr>
 <tr>
 <td>1H,1H,2H,2H-perfluorooctanol</td>
 <td>Reagent </td>
 <td>Sigma-Aldrich</td>
 <td>370533</td>
 </tr>
 <tr>
 <td>Sylgard 184 silicone elastomer kit</td>
 <td>Reagent</td>
 <td>Premier Farnell UK LTD</td>
 <td>1673921</td>
 </tr>
 <tr>
 <td>auramine-O</td>
 <td>Reagent</td>
 <td>Sigma-Aldrich</td>
 <td>861030</td>
 </tr>
 <tr>
 <td>485 nm pulsed diode laser</td>
 <td>Equipment</td>
 <td>PicoQuant</td>
 <td>LDH-P-C-485</td>
 </tr>
 <tr>
 <td>TCSPC Card</td>
 <td>Equipment</td>
 <td>PicoQuant</td>
 <td>TimeHarp 100</td>
 </tr>
 <tr>
 <td>dichroic mirror</td>
 <td>Equipment</td>
 <td>Chroma Technology Corp.</td>
 <td>z488rdc,</td>
 </tr>
 <tr>
 <td>Microscope objective</td>
 <td>Equipment</td>
 <td>Olympus</td>
 <td>UPLSAPO 60XW</td>
 </tr>
 <tr>
 <td>Avalanche photodiode</td>
 <td>Equipment</td>
 <td>AQR-141, EG&G, Perkin-Elmer).</td>
 <td>AQR-141</td>
 </tr>
 <tr>
 <td>DMEM</td>
 <td>Reagent</td>
 <td>Invitrogen</td>
 <td>ABCD1234</td>
 </tr>
 <tr>
 <td>SYTO9</td>
 <td>Reagent</td>
 <td>Invitrogen</td>
 <td>S34854</td>
 </tr>
 <tr>
 <td>Propidium Iodide</td>
 <td>Reagent</td>
 <td>Invitrogen</td>
 <td>P3566</td>
 </tr>
 <tr>
 <td>Escherichia coli top 10 strain</td>
 <td>Cells</td>
 <td>Invitrogen</td>
 <td>C4040-10</td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Fluorescence detection methods for microfluidic droplet platforms at JoVE.com

Fluorescence detection methods for microfluidic droplet platforms

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 ...

fluorescence-detection-methods-for-microfluidic-droplet-platforms

Research

JoVE Journal

Bioengineering

Microfabricated Platforms for Mechanically Dynamic Cell Culture

The ability to systematically probe in vitro cellular response to combinations of mechanobiological stimuli for tissue engineering, drug discovery or fundamental cell biology studies is limited by current bioreactor technologies, which cannot simultaneously apply a variety of mechanical stimuli to cultured cells. In order to address this issue, we have developed a series of microfabricated platforms designed to screen for the effects of mechanical stimuli in a high-throughput format. In this protocol, we demonstrate the fabrication of a microactuator array of vertically displaced posts on which the technology is based, and further demonstrate how this base technology can be modified to conduct high-throughput mechanically dynamic cell culture in both two-dimensional and three-dimensional culture paradigms.

In this protocol, we demonstrate the fabrication of a microactuator array of vertically displaced posts on which the technology is based, and how this base technology can be modified to conduct high-throughput mechanically dynamic cell culture in both two-dimensional and three-dimensional culture paradigms.

The ability to systematically probe in vitro cellular response to combinations of mechanobiological stimuli for tissue engineering, drug discovery or ...

High Throughput Microfluidic Rapid and Low Cost Prototyping Packaging Methods

In this work, 3 different packaging and assembly techniques are presented. They can be classified into two categories: one-time use and reusable packaging techniques.
The one-time use packaging technique employs UV-based and temperature curing epoxies to connect microtubes to access holes, wire-bonding for integrated circuit connections, and silver epoxy for electrical connections. This method is based on a robust assembly technique that can support relatively high pressure close to 1 psi and does not need any support to strengthen the microfluidic architecture.
Reusable packaging techniques consist of PDMS-based microtube interconnectors and anisotropic adhesive films for electrical connections. These devices are more sensitive and fragile. Consequently, Plexiglas support is added to the microfluidic structure to improve the electrical contact when anisotropic adhesive films are used, and also to strengthen the microfluidic architecture. In addition, a micromanipulator is needed to maintain tubes while using a thin PDMS layer to connect them to the access holes. Different PDMS layer thicknesses, ranging from 0.45-3 mm, are tested to compare the best adherence versus injection rates. Applied injection rates are varied from 50-300 &#956;l/hr for 0.45-3 mm PDMS layers, respectively. These techniques are mainly applicable for low-pressure applications. However, they can be extended for high-pressure ones through plasma-oxygen process to permanently seal the PDMS to glass substrates. The main advantage of this technique, besides the fact that it is reusable, consists of keeping the device observable when the microchannel length is very short (in the range of 3 mm or lower).

In this article we describe different techniques for microfluidic rapid prototyping platforms. The proposed techniques are based on ultraviolet (UV) sensitive and temperature curing epoxies, polydimethylsiloxane (PDMS) based tubing, wire-bonding, and anisotropic adhesive films. The assembling procedures presented are developed for both one-time use devices as well as reusable microfluidic systems.

In this work, 3 different packaging and assembly techniques are presented. They can be classified into two categories: one-time use and reusable packaging ...

3D Orbital Tracking in a Modified Two-photon Microscope: An Application to the Tracking of Intracellular Vesicles

The objective of this video protocol is to discuss how to perform and analyze a three-dimensional fluorescent orbital particle tracking experiment using a modified two-photon microscope1. As opposed to conventional approaches (raster scan or wide field based on a stack of frames), the 3D orbital tracking allows to localize and follow with a high spatial (10 nm accuracy) and temporal resolution (50 Hz frequency response) the 3D displacement of a moving fluorescent particle on length-scales of hundreds of microns2. The method is based on a feedback algorithm that controls the hardware of a two-photon laser scanning microscope in order to perform a circular orbit around the object to be tracked: the feedback mechanism will maintain the fluorescent object in the center by controlling the displacement of the scanning beam3-5. To demonstrate the advantages of this technique, we followed a fast moving organelle, the lysosome, within a living cell6,7. Cells were plated according to standard protocols, and stained using a commercially lysosome dye. We discuss briefly the hardware configuration and in more detail the control software, to perform a 3D orbital tracking experiment inside living cells. We discuss in detail the parameters required in order to control the scanning microscope and enable the motion of the beam in a closed orbit around the particle. We conclude by demonstrating how this method can be effectively used to track the fast motion of a labeled lysosome along microtubules in 3D within a live cell. Lysosomes can move with speeds in the range of 0.4-0.5 &#181;m/sec, typically displaying a directed motion along the microtubule network8.

In this video protocol we track - at high speed and in three dimensions - fluorescently labeled lysosomes within living cells, using the orbital tracking method in a modified two-photon microscope.

The objective of this video protocol is to discuss how to perform and analyze a three-dimensional fluorescent orbital particle tracking experiment using a ...

From Fast Fluorescence Imaging to Molecular Diffusion Law on Live Cell Membranes in a Commercial Microscope

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the regulation of many cellular functions. However, due to the fast dynamics and the tiny structures involved, a very high spatio-temporal resolution is required to catch the real behavior of molecules. Here we present the experimental protocol for studying the dynamics of fluorescently-labeled plasma-membrane proteins and lipids in live cells with high spatiotemporal resolution. Notably, this approach doesn&#8217;t need to track each molecule, but it calculates population behavior using all molecules in a given region of the membrane. The starting point is a fast imaging of a given region on the membrane. Afterwards, a complete spatio-temporal autocorrelation function is calculated correlating acquired images at increasing time delays, for example each 2, 3, n repetitions. It is possible to demonstrate that the width of the peak of the spatial autocorrelation function increases at increasing time delay as a function of particle movement due to diffusion. Therefore, fitting of the series of autocorrelation functions enables to extract the actual protein mean square displacement from imaging (iMSD), here presented in the form of apparent diffusivity vs average displacement. This yields a quantitative view of the average dynamics of single molecules with nanometer accuracy. By using a GFP-tagged variant of the Transferrin Receptor (TfR) and an ATTO488 labeled 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphoethanolamine (PPE) it is possible to observe the spatiotemporal regulation of protein and lipid diffusion on &#181;m-sized membrane regions in the micro-to-milli-second time range.

Spatial distribution and temporal dynamics of plasma membrane proteins and lipids is a hot topic in biology. Here this issue is addressed by a spatio-temporal image fluctuation analysis that provides conceptually the same physical quantities of single particle tracking, but it uses small molecular labels and standard microscopy setups.

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the ...

Fluorescence Recovery after Merging a Droplet to Measure the Two-dimensional Diffusion of a Phospholipid Monolayer

We introduce a new method to measure the lateral diffusivity of a surfactant monolayer at the fluid-fluid interface, called fluorescence recovery after merging (FRAM). FRAM adopts the same principles as the fluorescence recovery after photobleaching (FRAP) technique, especially for measuring fluorescence recovery after bleaching a specific area, but FRAM uses a drop coalescence instead of photobleaching dye molecules to induce a chemical potential gradient of dye molecules. Our technique has several advantages over FRAP: it only requires a fluorescence microscope rather than a confocal microscope equipped with high power lasers; it is essentially free from the selection of fluorescence dyes; and it has far more freedom to define the measured diffusion area. Furthermore, FRAM potentially provides a route for studying the mixing or inter-diffusion of two different surfactants, when the monolayers at a surface of droplet and at a flat air/water interface are prepared with different species, independently.

We present a new technique to measure the lateral diffusion of a surface active species at the fluid-fluid interface by merging a droplet monolayer onto a flat monolayer.

We introduce a new method to measure the lateral diffusivity of a surfactant monolayer at the fluid-fluid interface, called fluorescence recovery after ...

Multicolor Fluorescence Detection for Droplet Microfluidics Using Optical Fibers

Fluorescence assays are the most common readouts used in droplet microfluidics due to their bright signals and fast time response. Applications such as multiplex assays, enzyme evolution, and molecular biology enhanced cell sorting require the detection of two or more colors of fluorescence. Standard multicolor detection systems that couple free space lasers to epifluorescence microscopes are bulky, expensive, and difficult to maintain. In this paper, we describe a scheme to perform multicolor detection by exciting discrete regions of a microfluidic channel with lasers coupled to optical fibers. Emitted light is collected by an optical fiber coupled to a single photodetector. Because the excitation occurs at different spatial locations, the identity of emitted light can be encoded as a temporal shift, eliminating the need for more complicated light filtering schemes. The system has been used to detect droplet populations containing four unique combinations of dyes and to detect sub-nanomolar concentrations of fluorescein.

Multicolor fluorescence detection in droplet microfluidics typically involves bulky and complex epifluorescence microscope-based detection systems. Here we describe a compact and modular multicolor detection scheme that utilizes an array of optical fibers to temporally encode multicolor data collected by a single photodetector.

Fluorescence assays are the most common readouts used in droplet microfluidics due to their bright signals and fast time response. Applications such as ...

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 ...

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 ...

Microfluidic Platform with Multiplexed Electronic Detection for Spatial Tracking of Particles

Microfluidic processing of biological samples typically involves differential manipulations of suspended particles under various force fields in order to spatially fractionate the sample based on a biological property of interest. For the resultant spatial distribution to be used as the assay readout, microfluidic devices are often subjected to microscopic analysis requiring complex instrumentation with higher cost and reduced portability. To address this limitation, we have developed an integrated electronic sensing technology for multiplexed detection of particles at different locations on a microfluidic chip. Our technology, called Microfluidic CODES, combines Resistive Pulse Sensing with Code Division Multiple Access to compress 2D spatial information into a 1D electrical signal. In this paper, we present a practical demonstration of the Microfluidic CODES technology to detect and size cultured cancer cells distributed over multiple microfluidic channels. As validated by the high-speed microscopy, our technology can accurately analyze dense cell populations all electronically without the need for an external instrument. As such, the Microfluidic CODES can potentially enable low-cost integrated lab-on-a-chip devices that are well suited for the point-of-care testing of biological samples.

We demonstrate a microfluidic platform with an integrated surface electrode network that combines resistive pulse sensing (RPS) with code division multiple access (CDMA), to multiplex detection and sizing of particles in multiple microfluidic channels.

Microfluidic processing of biological samples typically involves differential manipulations of suspended particles under various force fields in order to ...

A Droplet-Based Microfluidic Approach and Microsphere-PCR Amplification for Single-Stranded DNA Amplicons

Droplet-based microfluidics enable the reliable production of homogeneous microspheres in the microfluidic channel, providing controlled size and morphology of the obtained microsphere. A microsphere copolymerized with an acrydite-DNA probe was successfully fabricated. Different methods such as asymmetric PCR, exonuclease digestion, and isolation on streptavidin-coated magnetic beads can be used to synthesize single-stranded DNA (ssDNA). However, these methods cannot efficiently use large amounts of highly purified ssDNA. Here, we describe a microsphere-PCR protocol detailing how ssDNA can be efficiently amplified and separated from dsDNA simply by pipetting from a PCR reaction tube. The amplification of ssDNA can be applied as potential reagents for the DNA microarray and DNA-SELEX (Systematic evolution of ligands by exponential enrichment) processes.

This work provides a method for the fabrication of droplet-based microfluidic platforms and the application of polyacrylamide microspheres for microsphere-PCR amplification. The microsphere-PCR method makes it possible to obtain single-stranded DNA amplicons without separating double-stranded DNA.