This work was supported by DARPA grant number 84389.01.44908, an NSF CAREER award (DBI-1253293), an NIH exploratory/developmental research grant (CA195709), and NIH New Innovator Awards (HD080351, DP2-AR068129-01), and a New Directions grant from the UCSF resource allocation program.

1. SU8 Master Fabrication
<ol>
	<li>Design the microfluidic structures for three layer fabrication using design software and have the designs printed by a vendor on circuit board film with 10 &#181;m resolution. The details of device design are given in an attached reference 6 and the channel geometries are shown in Figure 1. The layers should include alignment marks to help collocate features from each fabrication layer 8.</li>
	<li>Place a pre-cleaned 3 inch diameter silicon wafer on a spin coater and turn on the vacuum to affix it to the chuck. Apply 1 ml of SU8-3050 in the center of the wafer and spin for 20 sec at 500 rpm, then 30 sec at 1,750 rpm, providing a layer thickness of 80 &#181;m.</li>
	<li>Remove the wafer and bake on a 135 &#176;C hotplate for 30 min. Allow the wafer to cool to RT before moving on to the next step.</li>
	<li>Expose the coated wafer to the 1st layer mask under a collimated 190 mW, 365 nm LED for 3 min. After exposure, place the wafer on a 135 &#176;C hotplate for 1 min, then cool to RT before proceeding to the next step.</li>
	<li>Place the wafer on the spin coater and turn on the vacuum to affix it to the chuck. Apply 1 ml of SU8-3050 in the center of the wafer and spin for 20 sec at 500 rpm, then 30 sec at 5,000 rpm, resulting in a layer that provides an additional thickness of 40 &#181;m.</li>
	<li>Remove the wafer and bake on a 135 &#176;C hotplate for 30 min, then cool to RT before moving to the next step.</li>
	<li>Align the 2nd layer mask onto the geometry patterned in 1.3 and expose the coated wafer to a collimated 190 mW, 365 nm LED for 3 min. After exposure, place on a 135 &#176;C hotplate for 4 min 30 sec, then cool to RT before proceeding to the next step.</li>
	<li>Place the wafer on the spin coater and turn on the vacuum to affix it to the chuck. Apply 1 ml of SU8-3050 in the center of the wafer and spin for 20 sec at 500 rpm, then 30 sec at 1,000 rpm, resulting in a layer that provides an additional thickness of 100 &#181;m.</li>
	<li>Remove the wafer and bake on a 135 &#176;C hotplate for 30 min, then cool to RT before moving to the next step.</li>
	<li>Align the 3rd layer mask onto the geometry patterned in 1.3 and expose the coated wafer to a collimated 190 mW, 365 nm LED for 3 min. After exposure, place on a 135 &#176;C hotplate for 9 min, then cool to RT before proceeding to the next step.</li>
	<li>Develop the masks by immersing in a stirred bath of propylene glycol monomethyl ether acetate for 30 min. Wash the wafer in isopropanol and bake on a 135 &#176;C hotplate for 1 min. Place the developed master in a 100 mm Petri dish for polydimethylsiloxane (PDMS) molding.</li>
</ol>
2. PDMS Device Fabrication
<ol>
	<li>Prepare 10:1 PDMS by combining 50 g of silicone base with 5 g of curing agent in a plastic cup. Mix the contents with a rotary tool fitted with a stir stick. Degas the mixture inside a desiccator for 30 min, or until all air bubbles are removed.</li>
	<li>Pour the PDMS to give a thickness of 3 mm over the master and place back into the desiccator for further degassing. Once all bubbles are removed, bake the device at 80 &#176;C for 80 min.</li>
	<li>Cut the device from the mold using a scalpel and place on a clean surface with the patterned side up and punch the fluidic inlets and outlets with a 0.75 mm biopsy punch. The device must be cut so that the 120 &#181;m and 220 &#181;m tall geometries are accessible from the side of the device.</li>
	<li>Plasma treat the device, with feature side up, along with a pre-cleaned 2 inches by 3 inches glass slide at 1 mbar O2 plasma for 20 sec in a 300 W plasma cleaner. Bond the PDMS device by placing the patterned side of the PDMS device onto the plasma-treated side of the glass slide. Place the device in a 80 &#176;C oven and bake the assembled device for 40 min.</li>
	<li>Render the channels hydrophobic by using a syringe to flush the device with a fluorinated surface treatment fluid. Immediately bake the device at 80 &#176;C for 10 min to evaporate the solvent.</li>
</ol>
3. Preparation of Optical Components
<ol>
	<li>Prepare the laser excitation fiber by removing the insulation from the last 5 mm of a 105 &#181;m core, 125 &#181;m cladding, NA = 0.22 optical fiber.</li>
	<li>Prepare the 2nd color laser excitation fiber by repeating 3.1 for a 2nd identical fiber.</li>
	<li>Prepare the fiber for collecting the fluorescence signal by repeating 3.1 for a 200 &#181;m core, 225 &#181;m cladding, NA = 0.39 optical fiber.</li>
	<li>Inspect the tips of all the fibers &#8212; if the tips do not end in a flat surface, re-cleave the ends with a fiber scribe.</li>
	<li>Attach a laser fiber coupler to a 50 mW, 405 nm laser and attach one of the 105 &#181;m core fibers to the laser. Direct the stripped end at the sensor of the laser power meter, and use the fine adjustments of the laser coupler to maximize the laser power.</li>
	<li>Repeat 3.5 for a 50 mW, 473 nm laser.</li>
	<li>Mount a 446/510/581/703 nm quad bandpass filters on the PMT using lens tubes to block laser light and transmit emitted fluorescence. Attach fiber coupler so that light travels through the filters before hitting the PMT.</li>
	<li>Attach the fiber from 3.3 to the unit assembled in 3.7.</li>
</ol>
4. Offline Mixed Emulsion Generation
<ol>
	<li>Obtain a flow focus microfluidics dropmaker device with a 60 &#181;m x 60 &#181;m orifice.</li>
	<li>Fill a 1 ml syringe with HFE 7500 oils with 2% ionic fluorosurfactant 9.</li>
	<li>Fill a series of 1 ml syringes with the following combinations of fluorescein (FITC) and dextran-conjugated blue dyes (CB) in PBS: 1 nM FITC/10 nM CB, 10 nM FITC/10 nM CB, 1 nM FITC/100 nM CB, and 10 nM FITC/100 nM CB.</li>
	<li>Mount the dropmaker device on the stage of an inverted microscope and the aqueous and oil syringes on syringe pumps. Couple the syringes to the devices using PE-2 tubing.</li>
	<li>Running syringe pumps at 500 &#181;l/hr for the oil and 250 &#181;l/hr for the aqueous solutions, create a mixture of 80 &#181;m droplets containing the solutions from 4.3. For each type of aqueous sample, use a fresh dropmaker device to eliminate cross-contamination. Collect ~200 &#181;l of emulsion from each dye combination into an empty syringe.</li>
	<li>After the 4 droplets types have been collected in a single collection syringe, mix the emulsion by repeatedly rotating the syringe.</li>
	<li>Repeat steps 4.2-4.6 with aqueous solutions containing 0.1, 1, 10, and 100 nM FITC in PBS.</li>
</ol>
5. Optical Fiber Insertion
<ol>
	<li>Place the microfluidic chip fabricated in step 2 on the stage of an inverted microscope coupled with a digital camera capable of &#60; 100 &#181;sec shutter speeds.</li>
	<li>Working carefully from the side, insert the fiber coupled to the 473 nm laser into the farthest upstream 120 &#181;m channel, taking care not to puncture through to the main flow channel.</li>
	<li>Insert the fiber coupled to the 405 nm laser into the farthest downstream 120 &#181;m high side channel, providing fiber spacing of 300 &#181;m.</li>
	<li>Insert the PMT-coupled fiber into the 220 &#181;m tall channel normal to the two laser excitation fibers.</li>
</ol>
6. Fluorescence Detection of Mixed Emulsions&#160;
<ol>
	<li>Mount a 5 ml syringe filled with HFE 7500 with 2% ionic fluorosurfactant to the spacer oil inlet of the detection device with PE-2 tubing.</li>
	<li>Mount the syringe containing the mixed FITC/CB emulsion on a vertically oriented syringe pump and couple to the device&#39;s droplet reinjection inlet using PE-2 tubing. Connect a length of PE-2 tubing from the device exit to a waste container.</li>
	<li>Prime the device by running each of the pumps at 1,000 &#181;l/hr until both oil and droplets are seen to be regularly combining in the device and flowing downstream.</li>
	<li>Adjust the flow rates such that the spacer oil runs at 6,000 &#181;l/hr and the droplets at 100 &#181;l/hr, providing significant spacing between droplets traveling through the detection region.</li>
	<li>Turn on the lasers, start the data acquisition program, and adjust the PMT gain to provide signals that are more than 100x the baseline noise floor. Adjust the laser power so that all of the doublet peaks are clearly visible on a single, linearly scaled timetrace.</li>
	<li>Acquire 60 sec of the PMT voltage timetrace at at least 20 kHz. Import the data into computing program such as Matlab and measure the heights of peaks of the recorded doublets using the custom computing program script. 
	NOTE: This is provided as supplementary information.</li>
	<li>Repeat steps 6.2-6.7 using the FITC-only mixed emulsion created in 4.7, with the 405 nm laser turned off.</li>
</ol>

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R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Picoinjection+Enables+Digital+Detection+of+RNA+with+Droplet+RT-PCR.">Picoinjection Enables Digital Detection of RNA with Droplet RT-PCR.</a> PLOS ONE. 8 (4), (2013).</li><li>Cole, R. H., de Lange, N., Gartner, Z. J., Abate, A. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Compact+and+modular+multicolour+fluorescence+detector+for+droplet+microfluidics.">Compact and modular multicolour fluorescence detector for droplet microfluidics.</a> Lab Chip. 15 (13), 2754-2758 (2015).</li><li>Guo, F., Lapsley, M. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+droplet-based,+optofluidic+device+for+high-throughput,+quantitative+bioanalysis.">A droplet-based, optofluidic device for high-throughput, quantitative bioanalysis.</a> Anal Chem. 84, 10745-10749 (2012).</li><li>. Lithography Available from: <a target="_blank" href="https://www.memsnet.org/mems/processes/lithography.html">https://www.memsnet.org/mems/processes/lithography.html</a> (2015)</li><li>DeJournette, C. J., Kim, J., Medlen, H., Li, X., Vincent, L. J., Easley, C. 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M., Kiesel, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Time+encoded+multicolor+fluorescence+detection+in+a+microfluidic+flow+cytometer.">Time encoded multicolor fluorescence detection in a microfluidic flow cytometer.</a> Lab Chip. 12 (23), 5057-5062 (2012).</li><li>Bliss, C. L., McMullin, J. N., Backhouse, C. 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The authors have nothing to disclose.

Fiber optic detection requires the alignment of optical fibers with respect to fluid channels. Because our device utilizes guide channels fabricated with multilayer photolithography, placement of masks with respect to each other is of great importance. If the fiber guide channels are too close to the fluid channel, there is a potential for fluid leakage; if the guide channels are located too far away or misaligned, the fluorescence signal gathered by the detection fiber may be significantly diminished. Proper alignment can be aided by designing alignment marks, such as concentric circles into masks to be co-located during photo patterning. Additionally, manual insertion of fiber optics into the device is a delicate process with the potential to break the tips of fibers within the device, rendering them unusable. Restraining fiber against the glass slide bonded to the PDMS device and inserting the fibers slowly while observing with the microscope are keys to successful fiber insertion. Last, reinjection of pre-formed emulsions should occur with a minimum of handling. Ideally, once the emulsion is made and deposited directly into an empty syringe, it should not be transferred until it is used for droplet reinjection.
Multicolor detection requires that the peaks in a signal doublet be distinct from one another and that signals from consecutive droplets do not overlap. The detection setup that we use here uses a pair of 125 &#181;m optical fibers separated at a center-to-center distance of 300 &#181;m, providing a non-illuminated region of ~175 &#181;m between excitation regions. This allows an 80 &#181;m droplet to be excited by only one light source at a time and ensures that each of the peaks in a doublet is solely a result of one type of laser excitation. Designs where the separation between the excitation regions is similar to the droplet diameter give problematic signals with overlapping emission peak doublets. The width of a signal doublet correlates to the time that it takes for the leading edge of a droplet to enter the first excitation regions to the time the trailing edge of the droplet to leave the final excitation region, a distance of ~500 &#181;m in the device we describe. In order to maintain distinctness of adjacent detected droplets, signal doublets need to be spaced by a low signal region at least as wide as the signal doublet. For the configuration described here, 80 &#181;m droplets are spaced at least 1 mm apart.
Although multi-color detection with a single photodetector allows for significant savings in terms of cost and complexity compared to standard techniques, there is some loss of robustness because emitted light is not filtered by wavelength. This can be problematic when detecting droplets that contain multiple fluorophores that excite at the same wavelengths, because the fluorescence source is indistinguishable. These limitations can be overcome by performing experiments with nonoverlapping excitation profiles, or by designing experiments where spectral bleed over does not occur into a &#34;channel&#34; where a sensitive assay is being performed. This is compatible with many droplet microfluidic detection applications, where a bright background dye is used to indicate the presence of a droplet, and a second fluorophore is used to report the result of a molecular biology assay 10,11.
Compared to standard techniques, our approach has two main advantages: 1) spatially registered excitation regions eliminate the need for complicated light filtering, and 2) use of fiber optics removes the needs for a microscope and optical hardware to direct light. Because we address common technical concerns with microfluidic detection systems, previous investigators have developed methods to address similar concerns. For instance, Martini et al. frequency-encode multicolor fluorescence recorded by a single detector by projecting an excitation source through uniquely patterned &#34;bar code&#34; filters 12. Our use of an optical fiber array allows the utilization of a similar temporal encoding scheme using standard, off the shelf components. The literature shows optical numerous optical coupling techniques that are more controlled that the simple fiber insertion that we describe. Bliss et al. use liquid filled optical wave guides for the carefully controlled delivery and collection of light from specific microchannel locations 13, Martinez Vasquez et al. use waveguides laser etched into fused silica substrates to deliver excitation light 14, and Vishnubhatla et al. are able to precisely fabricate fiber insertion channels using a combination of laser irradiation and chemical etching 15. Each of these techniques require additional steps beyond those used to fabricate the base fluidic architecture of a microfluidic chip. While these fabrication procedures may be necessary for highly sensitive detection applications, we have found that fiber insertion into a molded PDMS channel is adequate for most of the everyday applications in our lab. The detection system was able to detect FITC concentrations down to 100 pM in 80 &#181;m droplets, roughly equivalent to the detection of ~17,000 fluorophore molecules. This sensitivity is adequate for most detection applications in our laboratory currently using an epifluorescence-based system, the most common of which being PCR-activated cell sorting of assay positive droplets containing the equivalent of 106 fluorophore molecules 11. The detection sensitivity also compares favorably to recently reported sensitivities of other optical fiber based systems &#8212; for instance, it is 200x more sensitive than the detection limit of 20 nM of Alexa Fluor 488 reported in &#62; 100 &#181;m droplets7.
We have presented a compact and modular droplet microfluidic multi-color detection scheme as an alternative to more costly, complex, and bulky epifluorescence microscope based systems. The necessary detection components used here (lasers, fibers, fiber adapters, filter, and PMT) can be purchased for &#60;$10,000, making the multi-color droplet detection accessible to a great number of investigators, and allowing individual investigators to afford multiple detection stations in a laboratory. The system also remove some expertise-based barriers to entry, as a degree of familiarity is required to align free space optical systems in conjunction with an epifluorescence microscope. Additionally the small footprint of the detections setup makes it ideally suited for portable and diagnostic applications.

Droplet microfluidics provide a platform for high throughput biology by compartmentalizing experiments in a large number of aqueous droplets suspended in a carrier oil 1. Droplets have been used for applications as varied as single cell analysis 2, digital polymerase chain reaction (PCR) 3, and enzyme evolution 4. Fluorescent assays are the standard mode of detection for droplet microfluidics, as their bright signals and fast time response are compatible with detecting sub-nanoliter droplet volumes at kilohertz rates. Many applications require fluorescence detection for at least two colors simultaneously. For instance, our lab commonly performs PCR-activated droplet sorting experiments that use one detection channel for the result of an assay, and uses a secondary background dye to make assay-negative droplet countable 5.
Typical detection stations for droplet microfluidics are based on epifluorescence microscopes, and require complicated light manipulations schemes to introduce excitation light from free space lasers into microscope to be focused on the sample. After fluorescence is emitted from a droplet, the emitted fluoresced light is filtered so that each detection channel utilizes one photomultiplier tube (PMT) centered on a wavelength band. Epifluorescence microscope-based optical detection systems provide a barrier to entry due to their expense, complexity, and required maintenance. Optical fibers provide the means to construct a simplified and robust detection scheme, since fibers can be manually inserted into microfluidic devices, removing the need for mirror-based light routing, and allowing light paths to be interfaced using optical fiber connectors.
In this paper, we describe the assembly and validation of a compact and modular scheme to perform multicolor fluorescence detection by utilizing an array of optical fibers and a single photodetector 6. Optical fibers are coupled to individual lasers and are inserted normal to an L shaped flow channel at regular spatial offsets. A fluorescence collection fiber is oriented parallel to the excitation regions and is connected to a single PMT. Because a droplet passes through the laser beams at different times, data recorded by the PMT shows a temporal offset that allows the user to distinguish between the fluorescence emitted after the droplet is excited by each distinct laser beam. This temporal shift eliminates the need to separate emitted light to separate PMTs using a series of dichroic mirrors and bandpass filters. To validate the efficacy of the detector, we quantitate fluorescence in droplet populations encapsulating dyes of different color and concentration. The sensitivity of the system is investigated for single color fluorescein detection, and shows the ability to detect droplets with concentrations down to 0.1 nM, a 200x sensitivity improvement as compared to recent fiber based approaches reported in the literature 7.

<table><tbody><tr><td>Photomasks</td><td>CadArt Servcies</td><td></td><td></td></tr><tr><td>3&quot; silicon wafers, P type, virgin test grade</td><td>University Wafers</td><td>447</td><td></td></tr><tr><td>SU-8 3035</td><td>Microchem</td><td>Y311074</td><td></td></tr><tr><td>SU-8 2050</td><td>Microchem</td><td>Y111072</td><td></td></tr><tr><td>Sylgard 184 silicone elastomer kit</td><td>Krayden</td><td>4019862</td><td></td></tr><tr><td>1 ml syringes</td><td>BD</td><td>309628</td><td></td></tr><tr><td>10 ml syringes</td><td>BD</td><td>309604</td><td></td></tr><tr><td>27 gaugue needles</td><td>BD</td><td>305109</td><td></td></tr><tr><td>PE 2 polyethylene tubing</td><td>Scientific Commodities, Inc.</td><td>B31695-PE/2</td><td></td></tr><tr><td>Novec 7500</td><td>Fisher Scientific</td><td>98-0212-2928-5</td><td>Commonly knowns as HFE 7500</td></tr><tr><td>Ionic Krytox Surfactant</td><td></td><td></td><td>Synthesis instructions in ref #10</td></tr><tr><td>Dextran-conjugated cascade blue dye</td><td>Life Technologies</td><td>D-1976</td><td></td></tr><tr><td>Fluorescein sodium salt</td><td>Sigma</td><td>28803</td><td></td></tr><tr><td>Quad bandpass filter</td><td>Semrock</td><td>FF01-446/510/581/703-25</td><td></td></tr><tr><td>PMT</td><td>Thorlabs</td><td>PMM02</td><td></td></tr><tr><td>Fiber port</td><td>Thorlabs</td><td>PAFA-X-4-A</td><td></td></tr><tr><td>lens tube</td><td>Thorlabs</td><td>SM1L05</td><td></td></tr><tr><td>Patch cable with 200 &amp;#956;m core / 225 &amp;#956;m cladding optical fiber with one stripped end and one FC/PC connector</td><td>Thorlabs</td><td>Custom</td><td></td></tr><tr><td>Patch cable with 105 &amp;#956;m core / 125 &amp;#956;m cladding optical fiber with one stripped end and one FC/PC connector</td><td>Thorlabs</td><td>Custom</td><td></td></tr><tr><td>125 &amp;#956;m fiber stripping tool</td><td>Thorlabs</td><td>T08S13</td><td></td></tr><tr><td>225 &amp;#956;m fiber stripping tool</td><td>Thorlabs</td><td>T10S13</td><td></td></tr><tr><td>laser fiber adapter</td><td>OptoEngine</td><td>FC/PC Adapter</td><td></td></tr><tr><td>405 nm CW laser at 50&amp;nbsp;mW</td><td>OptoEngine</td><td>MDL-III-405</td><td>Distributor for CNI lasers</td></tr><tr><td>473 nm CW laser at 50&amp;nbsp;mW</td><td>OptoEngine</td><td>MLL-FN-473-50</td><td></td></tr></tbody></table>

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.

Fabrication of a PDMS device that allows for the insertion of optical fibers requires a multistep photolithography procedure to create channels of varying height (Figure 1). First, an 80 &#181;m tall layer of SU-8 is spun onto a silicon wafer and patterned using a mask to create the fluid handling geometry. Next, an additional 40 &#181;m layer of SU-8 is spun onto the wafer, and patterned using a second mask to create features that will form 120 &#181;m tall laser fiber insertion channels. Last, 100 &#181;m more SU-8 is spun onto the wafer, and patterned to give 220 &#181;m tall detection fiber insertion channels. The master is developed and used to mold a PDMS device, which is in turn bonded to a glass slide. The final fabrication contains 80 &#181;m tall flow geometry, accessed via holes punched in the top of the device and 120 &#181;m and 220 &#181;m tall optical fiber channels accessed through the side of the device.
The geometry of the device used for detection is shown in Figure 2. Externally generated 80 &#181;m emulsions are re-injected into a drop reinjection port and spaced by spacer oil before they flow into a L shape detection channel. Two laser-coupled fibers are inserted into 120 &#181;m tall channels provide excitation at discrete spatial locations in the flow channel. Because the multimode fibers inserted into these channels have a 125 &#181;m external diameter and a 105 &#181;m core diameter, the entire cross section of the flow channel is illuminated by laser light. In our device the detection flow channel makes an abrupt 90 degree turn after the last laser excitation region to create close optical access for a 225 &#181;m OD fiber used to detect emitted fluorescence. The larger fiber is used because of its high numerical aperture (NA = 0.39), which is similar to a standard microscope objective.
A droplet flowing through the detection channel encounters discrete excitation regions in front of each of the laser-coupled optical fibers (Figure 3A). The single PMT coupled to the detection fiber records emitted fluorescence with a temporal shift the maps to the excitation source. For a droplet that contains dyes that excite at 473 nm (region &#34;1&#34; in Figure 3A) and at 405 nm (region &#34;2&#34; in Figure 3B), the resulting PMT signal contains a signal doublet with a peak separation that correlates with the time it takes a droplet to move from one excitation region to the next. By temporally encoding spectral data in this manner, the need for further light filtering and separate PMTs for each fluorophore color is eliminated. Figure 3B shows signal doublets for a train of 4 droplets with different dye combinations. The first peak in the double corresponds to the fluorescence emitted after excitation by the 473 nm laser in region &#34;1&#34; and the second peak corresponds to fluorescence emitted after excitation by the 405 nm laser in region &#34;2&#34;.
The efficacy of the detection scheme was tested by detecting a mixed emulsion of droplets containing dyes that excite at 405 nm (dextran-conjugated blue, CB) and at 473 nm (fluorescein, FITC). Droplets were flowed through the detection regions at roughly 50 Hz and data was recorded for 60 sec, then post-processed with a computing program script. The data is plotted in Figure 4 and shows clear separation between the four re-injected droplet types. Because the detection system does not differentiate between different colors of emitted light, each droplet is described by its maximum total fluorescence after being excited in region &#34;1&#34; (peak 1) and in region 2 (peak 2).
The dynamic range and absolute sensitivity of the detector is investigated by turning off the 405 nm laser and measuring the fluorescence of an emulsion containing fluorescein-only droplets (Figure 5). The data shows the ability to detect dye concentration that range from 0.1 to 100 nM FITC on a single detector with the same detection settings.
<img alt="Figure 1" src="/files/ftp_upload/54010/54010fig1.jpg" /> 
Figure 1. Multi-layer photolithography. The microfluidic device is fabricated by creating a master with three different feature heights. First, a 80 &#181;m tall layer of SU-8 is spun on and patterned using a mask for the fluid channels. Next, 40 &#181;m more SU-8 is spun on and 120 &#181;m tall features are patterned with a 2nd mask through both of the SU-8 layers to yield geometry to accommodate 125 &#181;m tall optical fibers. After this, 100 &#181;m more SU-8 is spun on and patterned with a 3rd mask to yield 225 &#181;m tall features. After developing, PDMS molding, and plasma bonding to glass, the resultant device contains large channels accessible from the side for fiber insertion, in addition to standard enclosed fluid channels. <a href="https://www.jove.com/files/ftp_upload/54010/54010fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54010/54010fig2.jpg" /> 
Figure 2. Device design and optical layout. The fluidic portions of the device are comprised of a droplet reinjector and an oil spacer, coupled to an L-shaped channel. Fibers coupled to individual lasers are inserted normal to the direction of flow. A detection fiber is inserted perpendicular to the laser fibers in order to collect fluorescent light. This light is filtered through a bandpass filter, before being detected by a single PMT. <a href="https://www.jove.com/files/ftp_upload/54010/54010fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54010/54010fig3.jpg" /> 
Figure 3. Temporal encoding of multicolor information. (A) Droplets flow through excitation regions &#34;1&#34; and &#34;2&#34; at different times, providing time shift that allows the identification of the emitted light. (B) Data from a train of 4 different droplets flowing through the detection channel. The first peak of each doublet corresponds to the fluorescence emitted after excitation in region &#34;1&#34; and the second peak in each double corresponds to the fluorescence emitted after excitation in region &#34;2&#34;. The height of droplet peaks are proportional the amount of dye excited at a given wavelength, except when spectral bleed occurs due to the use of dyes with overlapping excitation spectra. The width of the peaks are determined by the amount of time that it takes a droplet to traverse an excitation region. <a href="https://www.jove.com/files/ftp_upload/54010/54010fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/54010/54010fig4.jpg" /> 
Figure 4. Multicolor detection of a mixed droplet population. Scatter plot showing maximum droplet intensities for a mixed droplet population after excitation and emission by a 473 nm laser (peak 1) and a 405 nm laser (peak 2). The droplets contain combinations of fluorescein (FITC) and cascade blue (CB) at nM concentrations. <a href="https://www.jove.com/files/ftp_upload/54010/54010fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/54010/54010fig5.jpg" /> 
Figure 5. Single color detection on highly varied droplet population. Histogram of emissions by a mixed droplet population containing 0.1-100 nM of fluorescein after excitation by a 473 nm laser. <a href="https://www.jove.com/files/ftp_upload/54010/54010fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Multicolor Fluorescence Detection for Droplet Microfluidics Using Optical Fibers at JoVE.com

Multicolor Fluorescence Detection for Droplet Microfluidics Using Optical Fibers

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 ...

multicolor-fluorescence-detection-for-droplet-microfluidics-using

Universidad San Sebastian

Research

JoVE Journal

Bioengineering

10.4K Views.  University of California, San Francisco. 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.

Video: Multicolor Fluorescence Detection for Droplet Microfluidics Using Optical Fibers

The Visual Colorimetric Detection of Multi-nucleotide Polymorphisms on a Pneumatic Droplet Manipulation Platform

A simple and visual method to detect multi-nucleotide polymorphism (MNP) was performed on a pneumatic droplet manipulation platform on an open surface. This approach to colorimetric DNA detection was based on the hybridization-mediated growth of gold nanoparticle probes (AuNP probes). The growth size and configuration of the AuNP are dominated by the number of DNA samples hybridized with the probes. Based on the specific size- and shape-dependent optical properties of the nanoparticles, the number of mismatches in a sample DNA fragment to the probes is able to be discriminated. The tests were conducted via droplets containing reagents and DNA samples respectively, and were transported and mixed on the pneumatic platform with the controlled pneumatic suction of the flexible PDMS-based superhydrophobic membrane. Droplets can be delivered simultaneously and precisely on an open-surface on the proposed pneumatic platform that is highly biocompatible with no side effect of DNA samples inside the droplets. Combining the two proposed methods, the multi-nucleotide polymorphism can be detected at sight on the pneumatic droplet manipulation platform; no additional instrument is required. The procedure from installing the droplets on the platform to the final result takes less than 5 min, much less than with existing methods. Moreover, this combined MNP detection approach requires a sample volume of only 10 &#181;l in each operation, which is remarkably less than that of a macro system.

This work presents a simple and visual method to detect multi-nucleotide polymorphisms on a pneumatic droplet manipulation platform. With the proposed method, the entire experiment, including droplet manipulation and detection of multi-nucleotide polymorphisms, can be performed near 23 &#176;C without the aid of advanced instruments.

A simple and visual method to detect multi-nucleotide polymorphism (MNP) was performed on a pneumatic droplet manipulation platform on an open surface. ...

Genetics

Synthesis and Operation of Fluorescent-core Microcavities for Refractometric Sensing

This paper discusses fluorescent core microcavity-based sensors that can operate in a microfluidic analysis setup. These structures are based on the formation of a fluorescent quantum-dot (QD) coating on the channel surface of a conventional microcapillary. Silicon QDs are especially attractive for this application, owing in part to their negligible toxicity compared to the II-VI and II-VI compound QDs, which are legislatively controlled substances in many countries. While the ensemble emission spectrum is broad and featureless, an Si-QD film on the channel wall of a capillary features a set of sharp, narrow peaks in the fluorescence spectrum, corresponding to the electromagnetic resonances for light trapped within the film. The peak wavelength of these resonances is sensitive to the external medium, thus permitting the device to function as a refractometric sensor in which the QDs never come into physical contact with the analyte. The experimental methods associated with the fabrication of the fluorescent-core microcapillaries are discussed in detail, as well as the analysis methods. Finally, a comparison is made between these structures and the more widely investigated liquid-core optical ring resonators, in terms of microfluidic sensing capabilities.

Fluorescent-core microcavity sensors employ a high-index quantum-dot coating in the channel of silica microcapillaries. Changes in the refractive index of fluids pumped into the capillary channel cause shifts in the microcavity fluorescence spectrum that can be used to analyze the channel medium.

This paper discusses  fluorescent core microcavity-based sensors that can operate in a microfluidic  analysis setup. These structures are based on the ...

Engineering

Lensless Fluorescent Microscopy on a Chip

On-chip lensless imaging in general aims to replace bulky lens-based optical microscopes with simpler and more compact designs, especially for high-throughput screening applications. This emerging technology platform has the potential to eliminate the need for bulky and/or costly optical components through the help of novel theories and digital reconstruction algorithms. Along the same lines, here we demonstrate an on-chip fluorescent microscopy modality that can achieve e.g., &lt;4&mu;m spatial resolution over an ultra-wide field-of-view (FOV) of &gt;0.6-8 cm2 without the use of any lenses, mechanical-scanning or thin-film based interference filters. In this technique, fluorescent excitation is achieved through a prism or hemispherical-glass interface illuminated by an incoherent source. After interacting with the entire object volume, this excitation light is rejected by total-internal-reflection (TIR) process that is occurring at the bottom of the sample micro-fluidic chip. The fluorescent emission from the excited objects is then collected by a fiber-optic faceplate or a taper and is delivered to an optoelectronic sensor array such as a charge-coupled-device (CCD). By using a compressive-sampling based decoding algorithm, the acquired lensfree raw fluorescent images of the sample can be rapidly processed to yield e.g., &lt;4&mu;m resolution over an FOV of &gt;0.6-8 cm2. Moreover, vertically stacked micro-channels that are separated by e.g., 50-100 &mu;m can also be successfully imaged using the same lensfree on-chip microscopy platform, which further increases the overall throughput of this modality. This compact on-chip fluorescent imaging platform, with a rapid compressive decoder behind it, could be rather valuable for high-throughput cytometry, rare-cell research and microarray-analysis.

A lensless on-chip fluorescent microscopy platform is demonstrated that can image fluorescent objects over an ultra-wide field-of-view of e.g., &#62;0.6-8 cm2 with &#60;4&#956;m resolution using a compressive sampling based decoding algorithm. Such a compact and wide-field fluorescent on-chip imaging modality could be valuable for high-throughput cytometry, rare-cell research and microarray-analysis.

On-chip lensless imaging in general aims to replace bulky lens-based optical microscopes with simpler and more compact designs, especially for ...

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 ...

Using Three-color Single-molecule FRET to Study the Correlation of Protein Interactions

Single-molecule F&#246;rster resonance energy transfer (smFRET) has become a widely used biophysical technique to study the dynamics of biomolecules. For many molecular machines in a cell proteins have to act together&#160;with interaction partners in a functional cycle to fulfill their task. The extension of two-color to multi-color smFRET makes it possible to simultaneously probe more than one interaction or conformational change. This not only adds a new dimension to smFRET experiments but it also offers the unique possibility to directly study the sequence of events and to detect correlated interactions when using an immobilized sample and a total internal reflection fluorescence microscope (TIRFM). Therefore, multi-color smFRET is a versatile tool for studying biomolecular complexes in a quantitative manner and in a previously unachievable detail.
Here, we demonstrate how to overcome the special challenges of multi-color smFRET experiments on proteins. We present detailed protocols for obtaining the data and for extracting kinetic information. This includes trace selection criteria, state separation, and the recovery of state trajectories from the noisy data using a 3D ensemble Hidden Markov Model (HMM). Compared to other methods, the kinetic information is not recovered from dwell time histograms but directly from the HMM. The maximum likelihood framework allows us to critically evaluate the kinetic model and to provide meaningful uncertainties for the rates.
By applying our method to the heat shock protein 90 (Hsp90), we are able to disentangle the nucleotide binding and the global conformational changes of the protein. This allows us to directly observe the cooperativity between the two nucleotide binding pockets of the Hsp90 dimer.

Here, we present a protocol to obtain three-color smFRET data and its analysis with a 3D ensemble Hidden Markov Model. With this approach, scientists can extract kinetic information from complex protein systems, including cooperativity or correlated interactions.

Single-molecule F&#246;rster resonance energy transfer (smFRET) has become a widely used biophysical technique to study the dynamics of biomolecules. For ...

Biochemistry

Microfluidic Imaging Flow Cytometry by Asymmetric-detection Time-stretch Optical Microscopy (ATOM)

Scaling the number of measurable parameters, which allows for multidimensional data analysis and thus higher-confidence statistical results, has been the main trend in the advanced development of flow cytometry. Notably, adding high-resolution imaging capabilities allows for the complex morphological analysis of cellular/sub-cellular structures. This is not possible with standard flow cytometers. However, it is valuable for advancing our knowledge of cellular functions and can benefit life science research, clinical diagnostics, and environmental monitoring. Incorporating imaging capabilities into flow cytometry compromises the assay throughput, primarily due to the limitations on speed and sensitivity in the camera technologies. To overcome this speed or throughput challenge facing imaging flow cytometry while preserving the image quality, asymmetric-detection time-stretch optical microscopy (ATOM) has been demonstrated to enable high-contrast, single-cell imaging with sub-cellular resolution, at an imaging throughput as high as 100,000 cells/s. Based on the imaging concept of conventional time-stretch imaging, which relies on all-optical image encoding and retrieval through the use of ultrafast broadband laser pulses, ATOM further advances imaging performance by enhancing the image contrast of unlabeled/unstained cells. This is achieved by accessing the phase-gradient information of the cells, which is spectrally encoded into single-shot broadband pulses. Hence, ATOM is particularly advantageous in high-throughput measurements of single-cell morphology and texture &#8211; information indicative of cell types, states, and even functions. Ultimately, this could become a powerful imaging flow cytometry platform for the biophysical phenotyping of cells, complementing the current state-of-the-art biochemical-marker-based cellular assay. This work describes a protocol to establish the key modules of an ATOM system (from optical frontend to data processing and visualization backend), as well as the workflow of imaging flow cytometry based on ATOM, using human cells and micro-algae as the examples.

Microfluidic Imaging Flow Cytometry by Asymmetric-detection Time-stretch Optical Microscopy ATOM

This protocol describes the implementation of an asymmetric-detection time-stretch optical microscopy system for single-cell imaging in ultrafast microfluidic flow and its applications in imaging flow cytometry.

Scaling the number of measurable parameters, which allows for multidimensional data analysis and thus higher-confidence statistical results, has been the ...

Use of Dual Optical Tweezers and Microfluidics for Single-Molecule Studies

Visual biochemistry is a powerful technique for observing the stochastic properties of single enzymes or enzyme complexes that are obscured in the averaging that takes place in bulk-phase studies. To achieve visualization, dual optical tweezers, where one trap is fixed and the other is mobile, are focused into one channel of a multi-stream microfluidic chamber positioned on the stage of an inverted fluorescence microscope. The optical tweezers trap single molecules of fluorescently labeled DNA and fluid flow through the chamber and past the trapped beads, stretches the DNA to B-form (under minimal force, i.e., 0 pN) with the nucleic acid being observed as a white string against a black background. DNA molecules are moved from one stream to the next, by translating the stage perpendicular to the flow to enable the initiation of reactions in a controlled manner. To achieve success, microfluidic devices with optically clear channels are mated to glass syringes held in place in a syringe pump. Optimal results use connectors permanently bonded to the flow cell, tubing that is mechanically rigid and chemically resistant, and which is connected to switching valves that eliminate bubbles that prohibit laminar flow.

Visual, single-molecule biochemistry studied through microfluidic chambers is greatly facilitated using glass barrel, gas-tight syringes, stable connections of tubing to flow cells, and elimination of bubbles by placing switching valves between the syringes and tubing. The protocol describes dual optical traps that enable visualization of DNA transactions and intermolecular interactions.

Visual biochemistry is a powerful technique for observing the stochastic properties of single enzymes or enzyme complexes that are obscured in the ...

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

Biology

Microfluidic Droplet Assay for Single-Cell Cytokine Secretion Analysis

Microfluidic Approach to Resolve Simultaneous and Sequential Cytokine Secretion of Individual Polyfunctional Cells

Infections, autoimmune diseases, desired and adverse immunological responses to treatment can lead to a complex and dynamic cytokine response in vivo. This response involves numerous immune cells secreting various cytokines to orchestrate the immune reaction. However, the secretion dynamics, amounts, and co-occurrence of the different cytokines by various cell subtypes remain poorly understood due to a lack of appropriate tools to study them. Here, we describe a protocol using a microfluidic droplet platform that allows the time-resolved quantitative measurement of secretion dynamics for several cytokines in parallel on the single-cell level. This is enabled by the encapsulation of individual cells into microfluidic droplets together with a multiplexed immunoassay for parallel quantification of cytokine concentrations, their immobilization for dynamic fluorescent imaging, and the analysis of the respective images to derive secreted quantities and dynamics. The protocol describes the preparation of functionalized magnetic nanoparticles, calibration experiments, cell preparation, and the encapsulation of the cells and nanoparticles into droplets for fluorescent imaging and subsequent image and data analysis using the example of lipopolysaccharide-stimulated human peripheral blood mononuclear cells. The presented platform identified distinct cytokine secretion behavior for single and co-secreting cells, characterizing the expected phenotypic heterogeneity in the measured cell sample. Furthermore, the modular nature of the assay allows its adaptation and application to study a variety of proteins, cytokines, and cell samples, potentially leading to a deeper understanding of the interplay between different immune cell types and the role of the different cytokines secreted dynamically to shape the tightly regulated immune response. These new insights could be particularly interesting in the studies of immune dysregulations or in identifying target populations in therapy and drug development.

Author Spotlight: Unveiling the Polyfunctionality and Heterogeneity in Immune Responses

The protocol describes an advanced microfluidic platform to quantitatively measure cytokine secretion dynamics of individual human peripheral blood mononuclear cells. The platform measures up to three cytokines in parallel (IL-6, TNF&#945;, and IL-1&#946;) for each individual cell stimulated with lipopolysaccharide as an example.

Infections, autoimmune diseases, desired and adverse immunological responses to treatment can lead to a complex and dynamic cytokine response in vivo. ...

Immunology and Infection

Multi-color Localization Microscopy of Single Membrane Proteins in Organelles of Live Mammalian Cells

Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a super-resolution technique that allows for the determination of the microcompartments that are accessible for proteins by generating localization and tracking maps of these proteins. Moreover, by multi-color localization microscopy, the localization and tracking profiles of proteins in different subcompartments are obtained simultaneously. The technique is specific for live cells and is based on the repetitive imaging of single mobile membrane proteins. Proteins of interest are genetically fused with specific, so-called self-labeling tags. These tags are enzymes that react with a substrate in a covalent manner. Conjugated to these substrates are fluorescent dyes. Reaction of the enzyme-tagged proteins with the fluorescence labeled substrates results in labeled proteins. Here, Tetramethylrhodamine (TMR) and Silicon Rhodamine (SiR) are used as fluorescent dyes attached to the substrates of the enzymes. By using substrate concentrations in the pM to nM range, sub-stoichiometric labeling is achieved that results in distinct signals. These signals are localized with ~15&#8211;27 nm precision. The technique allows for multi-color imaging of single molecules, whereby the number of colors is limited by the available membrane-permeable dyes and the repertoire of self-labeling enzymes. We show the feasibility of the technique by determining the localization of the quality control enzyme (Pten)-induced kinase 1 (PINK1) in different mitochondrial compartments during its processing in relation to other membrane proteins. The test for true physical interactions between differently labeled single proteins by single molecule FRET or co-tracking is restricted, though, because the low labeling degrees decrease the probability for having two adjacent proteins labeled at the same time. While the technique is strong for imaging proteins in membrane compartments, in most cases it is not appropriate to determine the localization of highly mobile soluble proteins.

Here, we present a protocol for multi-color localization of single membrane proteins in organelles of live cells. To attach fluorophores, self-labeling proteins are used. Proteins, located in different membranes compartments of the same organelle, can be localized with a precision of ~18 nm.

Multicolor Fluorescence Detection for Droplet Microfluidics Using Optical Fibers

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