We thank the Eindhoven University of Technology for generous support.

1. 3-Inlet Polydimethylsiloxane (PDMS) Device Fabrication
<ol>
	<li>Measure 40 g of PDMS base in a conditioning mixer cup and add 4 g of PDMS curing agent to the base reagent in the cup, carefully, using a dropper.</li>
	<li>Place the cup in the holder of the conditioning mixer and measure the total weight of the cup with the holder. Set the value of the centrifuge balance weight on the conditioning mixer accordingly.</li>
	<li>Mix the base and curing agent in the conditioning mixer at 2,000 rpm for 2 min followed by de-foaming at 2,000 rpm for 2 min.</li>
	<li>Prepare an aluminum boat, with a diameter approximately the same size as that of a 100 mm silicon wafer. Place the silicon wafer, fabricated for the replica molding process, in the aluminum boat and put this setup in a Petri dish (diameter = 120 mm, height = 20 mm). 
	NOTE: The size of the Petri dish depends on the size of the silicon wafer.</li>
	<li>Remove the cup from the holder and pour the pre-cured PDMS mixture (contents of the cup), carefully, on the silicon wafer.</li>
	<li>Place the Petri dish, containing the silicon wafer with the pre-cured PDMS mixture, in a desiccator for about 20 min to remove all the air bubbles.</li>
	<li>Remove the Petri dish after 20 min and check for any remaining air bubbles that can be removed.</li>
	<li>Place the Petri dish in an oven, set at 65 &#176;C, for at least 3 h.</li>
	<li>Remove the Petri dish from the oven after 3 h and carefully peel the cured PDMS from the silicon wafer.</li>
	<li>Cut PDMS devices along the cut lines, using a knife or a scalpel. Punch holes at the inlets and outlet of each device using a 1.2 mm puncher. Clean each PDMS device with scotch tape to remove any dust or residual pieces of PDMS.
	<ol>
		<li>Optionally, blow with nitrogen to remove residual PDMS pieces.</li>
	</ol>
	</li>
	<li>Prepare glass slides by cleaning them with soap-water, followed by isopropanol and dry with nitrogen.</li>
	<li>Bond a clean PDMS device with a clean glass slide in a plasma asher to close the flow lines. Use the following settings: Power: 50 W, Time: 45 s, Bleed delay time: 2 s, Process gas: Gas 1 (Air), Vent: Both valves, Restricted vent time: 60 s, Pump spin down time: 10 s, Vent hold time: 0 s, Gas shutoff time: 1 s, Turbo pumping enabled: 0. Disconnect all the other gas lines. 
	NOTE: The settings used for the plasma asher can vary according to the brand of the plasma asher used.</li>
	<li>Prepare the silane solution by adding 50 &#181;L of silane (1H,1H,2H,2H-Perfluorooctyltriethoxysilane) to 950 &#181;L of fluorinated oil. 
	NOTE: Silane is toxic. Please operate under fume hood.</li>
	<li>Draw the prepared silane solution in a syringe, which is connected to a Teflon tubing.</li>
	<li>Salinize the device by flushing the prepared silane solution through the outlet of the device.</li>
	<li>Place the device in an oven, set at 65 &#176;C, for 30 min.</li>
	<li>Remove the salinized device from the oven and flush excess silane out of the device with fluorinated oil.</li>
	<li>Place the device back in an oven, set at 65 &#176;C, for at least 1 h to complete the bonding process. 
	Note: The Protocol can be paused here.</li>
</ol>
2. Loss-Less Cell Encapsulation
<ol>
	<li>Cell harvesting
	<ol>
		<li>Re-suspend Jurkat T cells in Roswell Park Memorial Institute (RPMI) medium at concentrations of 1.0x106 cells/mL, 2.0x106 cells/mL, 4.0x106 cells/mL, and 8.0x106 cells/mL; pDCs in hematopoietic serum-free culture media (e.g., X-VIVO 15) at concentrations of 1.3x106 cells/mL, 3.0x106 cells/mL, and 13.0x106 cells/mL; A549 cells in Dulbecco&#39;s Modified Eagle&#39;s Medium (DMEM) at a concentration of 1.0x106 cells/mL. 
		NOTE: The type of cells and concentration of cells can vary based on the experiment. Labelling of cells can also be done based on the experiment.</li>
	</ol>
	</li>
	<li>Tip-loading for aqueous droplet generation
	<ol>
		<li>Prepare fluorinated oil with 3% biocompatible surfactant mixture by adding 3&#160;mL of surfactant to 2&#160;mL of fluorinated oil. 
		NOTE: The concentration of the surfactant added to the fluorinated oil determines the stability of the emulsion for different incubation periods. The concentration of the surfactant varies depending on the media used for specific cell-types.</li>
		<li>Draw the oil phase mixture in a syringe (1 mL). Remove air bubbles from the syringe and connect it to a Teflon tubing of appropriate length.</li>
		<li>Prepare a sample syringe by drawing biocompatible mineral oil in a syringe. Remove air bubbles and connect the syringe to a Teflon tubing of appropriate length.</li>
		<li>Punch a PDMS plug with a diameter of 5 mm from a cured PDMS slab. 
		NOTE: The cured PDMS slab can be prepared using steps 1.1 to 1.9. Use a plain silicon wafer instead of a fabricated silicon wafer.</li>
		<li>Punch another hole in the center of the plug with a 1 mm puncher.</li>
		<li>Insert the plug in a 200 &#181;L pipette tip, from the larger end, such that it fits tightly. 
		NOTE: Use a 1,000 &#181;L pipette tip for larger sample volume and larger cells. For 1,000 &#181;L pipette tip, plugs of diameter ranging between 5 mm and 7 mm can be used. With a plug of diameter 5 mm, a sample volume of around 400 &#181;L can be aspirated in the pipette tip. If a plug of larger diameter is used (7 mm), more sample volume can be aspirated (around 900 &#181;L).</li>
		<li>Insert the tubing, which is connected to the syringe, in the PDMS plug, which has been inserted in the pipette tip. Push the syringe plunger slowly to fill the connected pipette tip with mineral oil. Push out all the residual air from the pipette tip.</li>
		<li>Lower the pipette tip, connected to the syringe, in the sample solution and aspirate about 150 &#181;L of sample in the tip.</li>
		<li>Repeat steps 2.2.4 to 2.2.8 to prepare a second sample syringe.</li>
		<li>Carefully place all the three prepared syringes on the syringe pump.</li>
		<li>Insert both the pipette tips, containing the sample, in the two inner inlets of the PDMS chip. Insert the tube containing the oil phase mixture in the outer inlet.</li>
		<li>Set the value of the flow rates on the syringe pump as follows: continuous phase solution: 600 &#181;L/h, cell samples: 100 &#181;L/h, each. Enter and set the dimensions of the syringe. 
		NOTE: The diameter settings will vary based on the type of syringe.</li>
		<li>Start the pump to flush sample solution through the inner channels of the device and oil phase through the outer channel of the device.</li>
		<li>Plug in a tubing of appropriate length to the outlet to start collecting the droplets when the droplet formation is stable. The time of collection varies based on the experiment.</li>
		<li>Collect droplets in a lock tube. Add 200 &#181;L of RPMI medium (without serum) on top of the collected droplets and incubate the sample. 
		NOTE: Incubation time of the collected droplets varies based on the experiment. Droplets are collected in a lock tube when flow cytometry-based analysis or isolation is performed after retrieving the cells from droplets by breaking the emulsion. It is possible to collect the droplets in a glass chamber if the experiment requires in-droplet microscopic analysis.</li>
	</ol>
	</li>
	<li>Emulsion breaking and cell retrieval for flow cytometric analysis
	<ol>
		<li>Prepare 20% 1H,1H,2H,2H-Perfluoro-1-octanol (PFO) solution (v/v) in fluorinated oil by adding 2 mL of PFO in 10 mL of flourinated oil.</li>
		<li>Remove excess oil from the bottom of the collection tube, containing the droplets, using a syringe.</li>
		<li>Add 100 &#181;L of 20% PFO solution to the emulsion to break the emulsion and release the encapsulated cells into the aqueous phase. Tap and mix briefly. Do not vortex at this point. Incubate for 1-2 min. 
		NOTE: The amount of PFO added depends on the quantity of droplets produced. Keep adding additional PFO until the oil layer is completely dissolved. Keep in mind that PFO is toxic for the cells and that too high PFO concentrations or too long incubation in PFO might lead to increased cell death.</li>
		<li>Spin the solution shortly at the lowest possible rcf for 30 s.</li>
		<li>Prepare 100 mL cold Phosphate-Buffered Saline (PBS) solution supplemented with 2% Fetal Calf Serum (FCS) (2 mL of FCS in 98 mL of PBS).</li>
		<li>Pipette 550 &#181;L of the aqueous fraction, immediately after centrifugation and transfer it to a new lock tube containing 500 &#181;L of cold PBS solution supplemented with 2% FCS, as prepared in step 2.3.5. Let any residual oil sink to the bottom of the new lock tube.</li>
		<li>Aspirate 950 &#181;L of the aqueous phase containing the cells from this lock tube, carefully, without aspirating any residual oil and transfer the solution to a new lock tube.</li>
		<li>Spin down the cells in the new lock tube for 10 min.</li>
		<li>Re-suspend the cells in 300 &#181;L of cold PBS solution supplemented with 2% FCS, as prepared in step 2.3.5. 
		NOTE: The cells can be also re-suspended in any other suitable solution such as media depending on the experiment.Stain the cells, based on the experiment, for analysis using flow cytometry.</li>
	</ol>
	</li>
</ol>
3. Cell Pairing
<ol>
	<li>Cell harvesting and staining
	<ol>
		<li>Count Jurkat T cells, from culture flask, and spin down the cells at 1,500 rpm for 5 min.</li>
		<li>Remove the supernatant and re-suspend 1.0x106 cells in 1 mL PBS to get a concentration of 1.0x106 cells/mL. The amount of PBS added depends on the cell count.</li>
		<li>Repeat steps 3.1.1 to 3.1.2 to prepare a second sample of Jurkat T-cells with the same cell concentration.</li>
		<li>Wash both samples twice with 1 mL of PBS at 1,500 rpm for 5 min.</li>
		<li>Re-suspend one cell sample with 1.25 &#181;M carboxyfluorescein succinimidyl ester (CFSE) dye and the other cell sample with 1.25 &#181;M far red dye or 1.25 &#181;M cell proliferation dye at a cell concentration of 1.0x106 cells/mL. The total staining solution is 1 mL for 1.0x106 cells. 
		NOTE: Cells can be labelled with different dyes depending on the filters available in the flow cytometer or in the fluorescence microscope.</li>
		<li>Incubate the cell samples with dyes for 10 min at 37 &#176;C.</li>
		<li>Stop the staining reaction by adding 1 mL of ice cold FCS after 10 min.</li>
		<li>Wash the cell samples twice with 1 mL of PBS at 1,500 rpm for 5 min.</li>
		<li>Re-suspend the cell samples in RPMI media at a concentration of 10.0x106 cells/mL, for each color.</li>
	</ol>
	</li>
	<li>Tip-loading for production of agarose hydrogel droplets for cell pairing 
	Note: For cell pairing using agarose hydrogel droplets, maintain the temperature of the system between 27 &#176;C and 37 &#176;C throughout the droplet generation and collection process to prevent gelling of the hydrogels and to warrant cellular viability9.
	<ol>
		<li>Dissolve ultra-low gelling temperature agarose by heating it up to 75 &#176;C in PBS at a concentration of 4% (w/v) and stir the mixture for 20 min.</li>
		<li>Mix the agarose solution with labelled Jurkat T cells to yield an agarose concentration of 2% (w/v). Repeat this for the other sample with differently labelled cells.</li>
		<li>Prepare fluorinated oil with 2% surfactant mixture by adding 20 mL of surfactant to 30 mL of fluorinated oil (oil phase mixture).</li>
		<li>Follow the steps 2.2.2 - 2.2.14. 
		NOTE: Because of the viscous nature of low melting agarose and to ensure stable droplet production, set the value of the flow rates on the syringe pumps as follows: oil phase mixture: 2,000 &#181;L/h, cell samples: 200 &#181;L/h. Enter and set the dimensions of the syringe.</li>
		<li>Collect the droplets in a lock tube and incubate the droplets at 4 &#176;C for 60 min.</li>
	</ol>
	</li>
	<li>Emulsion breaking and agarose bead retrieval for FACS analysis
	<ol>
		<li>After incubation of droplets for 60 min, remove excess oil from the lock tube, containing the droplets, using a syringe.</li>
		<li>Add 200 &#181;L of PFO to remove the oil interphase from the droplets. 
		NOTE: The amount of PFO added to the tube depends on the quantity of droplets produced. Keep adding additional PFO until the oil layer is completely dissolved.</li>
		<li>Wash the collected agarose beads twice with 1 mL of cold PBS to remove oil completely by centrifugation at 1,500 rpm for 10 min.</li>
		<li>Analyze collected agarose beads using flow cytometry. 
		NOTE: It is also possible to observe the beads under a fluorescence microscope.</li>
	</ol>
	</li>
</ol>

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We have nothing to disclose.

In this protocol, we have demonstrated an efficient and straightforward technique to load and encapsulate cells in droplets for high-throughput, single-cell analysis and to perform controlled cell pairing for cellular interaction studies. Furthermore, we have compared several conventional approaches to load cells to microfluidic devices and showed that our tip loading approach is a more efficient technique in comparison to other methods.
Studying clinical specimens or rare cell types scarce in number by droplet-based microfluidics possess some inherent challenges. Like we have also demonstrated, cells tend to sediment in syringes and surface of the tubing, thereby, preventing cellular encapsulation to conform to the predicted values. To evade this problem, some groups use stirring bars in the syringes. However, when using rare and limited cell populations, the total cell volume is also limited, thereby, limiting the use of large syringes and stirring bars. Furthermore, we also replaced more commonly used tubing with Teflon coated tubing to prevent cell attachment but this method did not improve the results and if the tubing is too long, the problem of cell attachment aggravates (data not shown). Alternatively, we used a vertical tube loading approach where the cells were loaded in the tubing and not in the syringe to prevent the loss of cells in large syringe volumes. Using this technique, cells with small sample volume can be loaded, e.g., pDCs which are rare and limited. Also, the sample from the tubing is loaded to the device vertically to prevent cell sedimentation. The tubing used for cell seeding has small dimensions and can be compared to microchannels. The flow in the tubing is pressure driven and follows a parabolic velocity profile26. This implies that the maximum flow velocity is at the center of the tubing and minimum velocity is at the edges of the tubing27. When flushing a population of cells through the tubing, the velocity gradient causes the cells to be pushed towards the edges where they settle down because the velocity at the boundary is close to zero. The sedimentation or settlement of cells in the tubing, thereby, reduces the encapsulation efficiency as shown in the representative results where the experimental data did not match with the predicted model.
Another commonly adapted solution used by scientists, working with droplet microfluidics, is to increase the density of the cell culture media by addition of density matching reagents such as Iodinaxol to prevent cell sedimentation in syringes19. However, density matching reagents can influence cellular behavior and adversely affect the cytokine secretion by cells (data not shown)28.
Even though several small and big modifications in conventional cell loading techniques showed slight improvements in encapsulation efficiencies, the obtained experimental results still did not match the theoretical calculations. However, with the tip loading approach we could the overcome the limitations of previous methods and encapsulation efficiency governed by Poisson statistics. This technique is not only advantageous for loading suspension cells but can also be applied for loading adherent cells, such as primary keratinocytes and A549 to microfluidic chips. When using abundant cell lines, for example A549, K562, etc., larger sample volume can be used. Therefore, depending on the volume of the sample, different sized pipette-tips can also be used and this simple technique can be adapted for both single-cell encapsulation and multiple cell encapsulation.
While low cell concentration is required to ensure the encapsulation of single cells in droplets, higher cells concentrations are desired to increase the average number of cells encapsulated in each droplet for studies related to cell pairing. There are several single-cell methods that have been previously described to pair immune cells on microfluidic chips or microfabricated nanowells29,30,31. In droplet microfluidics, Poisson statistics dictates that 1:1 cell pairing for two different cell types can be achieved at optimal cell concentrations. Based on the Poisson prediction, there is also a probability that droplets might contain other combinations. While 1:1 cell pairing can be desirable to study cellular interactions at single cell level and results in increased cellular understanding, multiple cell pairing also has major advantages. It allows to comprehend the influence of multiple cells of one cell type on the other cell type. Cross talk between different immune cells help to generate an effective immune response against several infections and pathogens and also adds robustness to our immune system32. As such, cellular communication can be interrogated with high precision in distinct contexts, e.g., 1:1, 2:1, 1:2, 2:2, 3:1, etc. yielding increased understanding on how single or pairs of cells control the induction of immune responses. This is particularly interesting to study for example the capacity of natural killer cells or cytotoxic T cells to serially kill their respective target cells.
As discussed, for multiple encapsulation of cells in droplets, higher cell concentrations are desired. However, when loading cells from one inlet for cell encapsulation, higher concentrations of cell sample can cause cells to aggregate at the inlet. This results in lower encapsulation rates and higher deviation from the theoretical values. To evade this problem, the cells can be loaded from two separate inlets as well. Theoretically, it would be possible to develop other microfluidic devices with multiple inlets to achieve even higher levels of cell encapsulation where an average on x number of cells is warranted. In this study we investigated the encapsulation efficiency of Jurkat T cells when loaded from both one inlet and two inlets using the same total concentration and obtained similar encapsulation efficiency. This modification allows researchers to pair different cell types on chip.
While this method aids in loading cells to microfluidic devices without significant loss of cells, there are certain precautions that need to be kept in mind. When filling the syringes with mineral oil and aspirating the cell sample in pipette tips, incorporation of air bubbles should be avoided and the entire system should be air-free. It is also important to keep in mind that the mineral oil should not mix with the sample. Pipette tips, containing samples, should be inserted firmly in the inlets of the microfluidic device, with utmost precaution, to prevent leakage and further incorporation of air bubbles. To summarize, tip-loading is a straightforward, yet, robust technique that allows for high-throughput analysis of cellular behavior through cell encapsulation without significant loss of cells in a cost-effective manner. When used with optimal sample concentrations at the inlet, this approach of loading cells with pipette-tips is very flexible and can be adapted for different cell types, especially for rare primary immune cells, to obtain higher encapsulation efficiency, close to predicted models.

In recent years, the use of microfluidics as a robust and versatile platform for cellular analysis at the single cell level has rapidly increased1. These platforms provide high-throughput screening of single cells and biological molecules with high precision and sensitivity using very small sample volume2,3,4. Among different types of microfluidic designs, droplet-based platforms enable high-throughput analysis of single cells by isolating them in an aqueous phase droplet surrounded by an immiscible phase that allows precise and accurate control over the cellular microenvironment5,6. Droplet-based microfluidics gives the flexibility to isolate single or multiple-cells in, both, aqueous and hydrogel droplets and is valuable in probing complex cellular behavior, such as protein secretion or cellular interactions7,8,9. Signaling and cross-talk amongst immune cells can be influenced by interactions with other cells in the microenvironment10. Isolation of single cells in droplets provides an effective noise-free analytical laboratory, free from the influence of external environmental factors for more efficient and accurate results11,12. Modifying the design of a droplet-microfluidic platform with multiple inlets allows the encapsulation of multiple cell types to study cellular interactions via cell-pairing12,13.
The process of encapsulation of cells in droplets is random and the rate of encapsulation of cells can be statistically determined using the formula for the Poisson distribution14,15. This rate of encapsulation can be estimated by considering the average rate of the arrival of cells at the droplet junction and assuming that the arrival of each cell is independent from the arrival of other cells16. Even though independent cell arrival cannot be guaranteed, in cases of sparsely distributed cells, the assumption of independence can be considered and the probability of a droplet containing one or more cells can be predicted as a function of the number of cells present in each droplet and the average number of cells per droplet16,17. Since this estimation of cellular encapsulation in droplets is dependent on the number of cells present in each droplet, one can suggest that increasing the concentration of the cells at the inlet will increase the average number of cells present in each droplet16. Therefore, to ensure single cell encapsulation, the cell concentrations must be reduced but this often leads to a large number of empty droplets18.
Loss of cells during loading by either attachment, sedimentation, and/or clumping in the syringe, tubing, or production device is a common drawback responsible for the deviation of actual encapsulation values from the predicted encapsulation values19. This problem gets further exaggerated when seeding rare immune cells or clinical specimens as they are already scarce in population and the encapsulation of only a few cells, much lower than expected, does not provide sufficient data for experimental analysis. Plasmacytoid dendritic cells (pDCs) are a rare subset of immune cells that only constitutes approximately 0.2 - 0.6 percent of the entire white blood cell population20. These cells secrete massive amounts of type I interferons upon activation and thereby play a critical role in immune responses21. When studying the cellular behavior of such rare cells in droplets, it is imperative to prevent cell loss during cell seeding and encapsulation22. There are several design related developments that have ensured the encapsulation of single cells in droplets using active encapsulation methods that utilize different physical forces such as acoustic or electrical forces for generation of droplets containing single-cells23,24. However, these methods have their own limitations in terms of droplet production16.
In this study, we established a robust and straightforward method that circumvents the shortcomings of traditional methods for loading single or multiple cells to microfluidic devices. Our method, inspired by Rho et al., utilizes differently-sized pipette tips for seeding small volumes of rare immune cells to droplet microfluidic platforms without significant sample loss and yielded results that are coherent with theoretical predictions25. This methodology can be easily and successfully adapted for several applications involving droplet-based microfluidics and applied for a wide variety of cell types or even microparticles.

<table><tbody><tr><td>1H,1H,2H,2H-Perfluoro-1-octanol&amp;nbsp;</td><td>Sigma-Aldrich</td><td>171468-5G</td><td></td></tr><tr><td>1H,1H,2H,2H-Perfluorooctyltriethoxysilane</td><td>Fluorochem/UK</td><td>S13150</td><td>Silane (toxic)</td></tr><tr><td>Agarose (Ultra-low Gelling Temperature)</td><td>Sigma-Aldrich</td><td>&amp;nbsp;9012-36-6</td><td></td></tr><tr><td>BD&amp;nbsp;Wegwerpspuiten met Luer-Lok-punten</td><td>Fisher Scientific</td><td>10630694</td><td>Syringe</td></tr><tr><td>Biopsy Punch 1.2 mm</td><td>Harris Uni-Core</td><td></td><td></td></tr><tr><td>Cell Proliferation Dye eFluro 670</td><td>eBioscience</td><td>65-0840-85</td><td></td></tr><tr><td>CellTrace CFSE</td><td>Invitrogen</td><td>C34554</td><td></td></tr><tr><td>CellTrace Far Red Cell</td><td>Invitrogen</td><td>C34564</td><td></td></tr><tr><td>Eppendorf Tubes</td><td>Eppendrof Tubes</td><td></td><td>Safe-Lok tubes 1 mL and 2 mL</td></tr><tr><td>Glass Slide</td><td>Sigma Aldrich</td><td>CLS294775X38-72EA</td><td>Corning microscope slides, plain L &amp;times; W 75&amp;nbsp;mm &amp;times; 38&amp;nbsp;mm</td></tr><tr><td>Harvard Pumps</td><td>Harvard Apparatus</td><td>C-400750; C-400727</td><td>Syringe pumps</td></tr><tr><td>HFE-7500 3M Novec Engineered fluid</td><td>Fluorochem/UK</td><td>51243</td><td>Flourinated oil</td></tr><tr><td>Kai Biopsy Punch 5 mm</td><td>Amstel Medical</td><td>1980130</td><td></td></tr><tr><td>Luer stub</td><td>Instechlabs/USA</td><td>LS20S</td><td>Luer stub, 20ga (pink) x 0.5in (12mm), non-sterile</td></tr><tr><td>Mineral oil (Light)</td><td>Sigma Aldrich</td><td>M8410-1L</td><td></td></tr><tr><td>Phosphate buffered saline</td><td>Sigma-Aldrich</td><td>P4417-50TAB</td><td>Tablets</td></tr><tr><td>Pico-Surf 1 (5%in Novec 7500)</td><td>Sphere Fluidics</td><td>020317-09</td><td>Surfactant</td></tr><tr><td>Plasma Asher</td><td>Emitech</td><td>K1050X&amp;nbsp;</td><td>Plasma asher</td></tr><tr><td>RPMI 1640 Medium</td><td>Gibco</td><td>11875093</td><td></td></tr><tr><td>Silicone Elastomer Base 184</td><td>Sylgard</td><td>9355218</td><td>PDMS base</td></tr><tr><td>Silicone Elastomer Curing Agent&amp;nbsp;</td><td>Sylgard</td><td>9355218</td><td>Curing Agent&amp;nbsp;</td></tr><tr><td>Stainless steel catheter coupler</td><td>Instechlab/USA</td><td>SC20/15</td><td>20ga x 15mm, non-sterile</td></tr><tr><td>TFE Teflon Tubing</td><td>Sigma-Aldrich</td><td>58696-U</td><td>PTFE Tubing&amp;nbsp; L &amp;times; O.D. &amp;times; I.D. 50 ft &amp;times; 1/16 in. &amp;times; 0.031 in.</td></tr><tr><td>Thinky mixer ARE-250</td><td></td><td>EX-4025F</td><td>Conditioning mixture</td></tr></tbody></table>

a pipette-tip based method for seeding cells to droplet microfluidic platforms

Amongst various microfluidic platform designs frequently used for cellular analysis, droplet-microfluidics provides a robust tool for isolating and analyzing cells at the single-cell level by eliminating the influence of external factors on the cellular microenvironment. Encapsulation of cells in droplets is dictated by the Poisson distribution as a function of the number of cells present in each droplet and the average number of cells per volume of droplet. Primary cells, especially immune cells, or clinical specimens can be scarce and loss-less encapsulation of cells remains challenging. In this paper, we present a new methodology that uses pipette-tips to load cells to droplet-based microfluidic devices without the significant loss of cells. With various cell types , we demonstrate efficient cell encapsulation in droplets that closely corresponds to the encapsulation efficiency predicted by the Poisson distribution. Our method ensures loss-less loading of cells to microfluidic platforms and can be easily adapted for downstream single cell analysis, e.g., to decode cellular interactions between different cell types.

For our experiments, we used a 3-inlet PDMS based microfluidic device with the height of 25 microns (Figure 1). In this device setup, we used the outer inlet for flushing the oil with surfactant and the two inner inlets for flushing the aqueous phases with cell suspensions or media. After generation and collection, the droplets are incubated for a couple of hours off chip before downstream analysis using flow cytometry. During the incubation period, serum components present in the media can interact with the surfactant and cause droplets to become unstable and disintegrate. It is therefore important to add an optimized concentration of surfactant to the fluorinated oil. We tested the stability of the monodispersed droplets containing hematopoietic serum-free culture media supplemented with 2% human serum with different concentrations of surfactant in fluorinated oil. It can be inferred from Figure 2 that these monodispersed droplets are highly stable for up to 24 hours when at least 3% surfactant is added to the oil phase. Similar results were obtained with RPMI media with and without the addition of 10% FCS (data not shown). Therefore, droplet stability is highly dependent on optimal surfactant concentrations when working with different sources of culture media and serum components.
To demonstrate the encapsulation efficiency of our approach we first seeded the cells using tubing connected to syringes, which is the most conventional approach for seeding cells (Figure 3A). We harvested Jurkat T cells at different concentrations of 1.0x106 cells/mL, 2.0x106 cells/mL, and 4.0x106 cells/mL and obtained an encapsulation efficiency that was lower than predicted values (Figure 3B). At 1.0x106 cells/mL, the fraction of droplets that contained a single cell was 2.5%, which did not increase even upon using higher cell concentrations. To increase the cell-loading efficiency, we modified our previous approach and mounted the tubing at half the length to an elevated tripod and loaded the cell suspension in the half that was attached to the PDMS device (Figure 4A). Using this approach, we encapsulated Jurkat T cells at different concentrations of 1.0x106 cells/mL, 2.0x106 cells/mL, and 4.0x106 cells/mL, and also rare pDCs at different concentrations of 1.0x106 cells/mL, 2.0x106 cells/mL, and 12.0x106 cells/mL. We expected improved encapsulation rates by preventing cell sedimentation with this method. However, at all the concentrations tested, the experimental results were much lower that the predicted Poisson values (Figure 4B and Figure 4C).
Using our tip-loading approach we optimized our cell encapsulation rates to obtain experimental results coherent with the statistically predicted values (Figure 5A). For different concentrations of Jurkat T cells, the obtained encapsulation efficiency matched our calculated values at all concentrations (Figure 5B). Remarkably, even with adherent cells like A549 tumor cells, which tend to clump, we observed a slightly improved encapsulation efficiency at a cellular concentration of 1.0x106 cells/mL (Figure 5C). We also assessed the efficacy of our system to encapsulate less available and scarce pDCs at different cell concentrations of 1.0x106 cells/mL, 3.0x106 cells/mL, and 13.0x106 cells/mL (Figure 5D). To facilitate the loading of possibly larger volumes exceeding 200 &#181;L, e.g., when working with cell lines or more abundant primary immune cells, we also investigated the cell encapsulation efficiency by using 1000 &#181;L tips (blue). We demonstrated that these 1,000 &#181;L tips gave a similar encapsulation efficiency in comparison to the 200 &#181;L tips (yellow) (Figure 5E).
Dependent on the chip design and research question at hand, our tip loading technique can be used to load cells through either one inlet, for probing into cellular heterogeneity, or multiple inlets in parallel, for decoding cellular interactions. We compared the loading of Jurkat T cells (at a concentration of 10.0x106 cells/mL) from one inlet to two differently labelled populations of Jurkat T cells (at a combined concentration of 10.0 x106 cells/mL) from two inlets (Figure 6A and Figure 6B). During encapsulation, the droplets were generated using ultra-low gelling temperature agarose and gelled after production to form agarose hydrogel beads that allowed subsequent downstream analysis via microscopy and flow cytometry (Figure 6C and Figure 6D). Microscopic analysis revealed that cell pairing was achieved at different combinations indicating for high throughput cell pairing (Figure 6C). Furthermore, analysis of the same population of hydrogel beads by flow cytometry revealed that beads without cells could be separated from beads with cells based on the distinct forward (FSC, size) and sideward (SSC, granularity) scatter pattern (Figure 6D). Gating on the population of beads without cells confirmed a lack of cell encapsulation by the absence of fluorescent signals. Additionally, gating on the bead population with cells revealed the existence of multiple sub-populations indicative for the encapsulation of differently labeled Jurkat T cells. Our results demonstrate that efficient cell pairing can be achieved, based on both microscopic and flow cytometric analysis, and showed a slightly increased encapsulation efficiency compared to the Poisson prediction (Figure 6E).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57848/57848fig1.jpg" /> 
Figure 1. PDMS based droplet microfluidic device with three inlets and one outlet. The device consists of three inlets for continuous oil phase, cell culture media, and cell suspension, respectively. The generated droplets are collected at the outlet. The samples flow laminarly to the flow-focusing junction where they are encapsulated in droplets. At the inlets, filter structures hold large particles like protein or cell aggregates back. The diameter of the gaps in the filter structure are indicated by blue lines. The diameter of the channels at the production nozzle are indicated by red lines. The channel height on the entire chip was 25 &#181;m. <a href="https://www.jove.com/files/ftp_upload/57848/57848fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57848/57848fig2.jpg" /> 
Figure 2. Droplet stability over 24 hours. The graphs show the area of droplets containing hematopoietic serum-free culture media + 2% human serum, over time for three different concentrations of surfactant A) 0.5% B) 3% C) 5%. <a href="https://www.jove.com/files/ftp_upload/57848/57848fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57848/57848fig3.jpg" /> 
Figure 3. Tubing based cell loading approach. Jurkat-T cells are loaded at different concentrations to the device using a syringe connected to tubing. A) The illustration shows the experimental setup B) The cell encapsulation rate as determined by light microscopy. Dots: experimentally determined values; Closed lines: Poisson distribution. Error bar represents standard error of mean. <a href="https://www.jove.com/files/ftp_upload/57848/57848fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57848/57848fig4.jpg" /> 
Figure 4. Encapsulation of various cell types at different concentrations using a vertical tube loading approach. Jurkat T cells and pDCs (of different concentrations) were encapsulated to determine the efficiency of cell encapsulation. A) The illustration shows the experimental setup for the vertical tube loading approach. B) The graph shows the encapsulation efficiency of Jurkat T cells. C) The graph shows the encapsulation efficiency of pDCs. Dots: experimentally determined values; Closed lines: Poisson distribution. Error bar represents standard error of mean. <a href="https://www.jove.com/files/ftp_upload/57848/57848fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/57848/57848fig5.jpg" /> 
Figure 5. Tip loading approach to encapsulate different cells types. A) Schematic illustration of the tip loading technique. B) The graph shows the Encapsulation efficiency of Jurkat cells. C) The graph shows the encapsulation efficiency of A549 cells. D) The graph shows the encapsulation efficiency of pDCs. E) The graph shows the encapsulation efficiency of Jurkat T cells using 200 &#181;L pipette tips (yellow) and 1,000 &#181;L pipette tips (blue). Dots: experimentally determined values; Closed lines: Poisson distribution. Error bar represents standard error of mean. <a href="https://www.jove.com/files/ftp_upload/57848/57848fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/57848/57848fig6.jpg" /> 
Figure 6. Cell pairing in droplets. A) Schematic illustration of the tip loading approach for pairing distinct cells from 2 inlets in droplets. B) The graph shows the encapsulation of Jurkat T cells using one inlet or two inlets in parallel. The cell concentration for one tip is 2.0x106 cells/mL and the cell concentration for two tips both 1.0x106 cells/mL. Dots: experimentally determined values; Closed lines: Poisson distribution. C) The Fluorescence microscopic overlays of hydrogel beads and labelled Jurkat T cells. D)The graph shows the flow cytometric analysis of paired cells in agarose hydrogel beads. The plot demonstrates both forward scatter and sideward scatter. E) Comparison of cell numbers in agarose hydrogel beads as detected by fluorescence microscopy and flow cytometry. Bars: mean value; Whiskers: standard error of mean, n &#8805; 4. Error bar represents standard error of mean. <a href="https://www.jove.com/files/ftp_upload/57848/57848fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Pipette-Tip Based Method for Seeding Cells to Droplet Microfluidic Platforms at JoVE.com

A Pipette-Tip Based Method for Seeding Cells to Droplet Microfluidic Platforms

This article presents a protocol for seeding scarce population of cells using pipette-tips to droplet microfluidic devices in order to provide higher encapsulation efficiency of cells in droplets.

Amongst various microfluidic platform designs frequently used for cellular analysis, droplet-microfluidics provides a robust tool for isolating and ...

a-pipette-tip-based-method-for-seeding-cells-to-droplet-microfluidic

Research

JoVE Journal

Engineering

11.1K Views.  Eindhoven University of Technology. This article presents a protocol for seeding scarce population of cells using pipette-tips to droplet microfluidic devices in order to provide higher encapsulation efficiency of cells in droplets.

Video: A Pipette-Tip Based Method for Seeding Cells to Droplet Microfluidic Platforms

Picoinjection of Microfluidic Drops Without Metal Electrodes

Existing methods for picoinjecting reagents into microfluidic drops require metal electrodes integrated into the microfluidic chip. The integration of these electrodes adds cumbersome and error-prone steps to the device fabrication process. We have developed a technique that obviates the needs for metal electrodes during picoinjection. Instead, it uses the injection fluid itself as an electrode, since most biological reagents contain dissolved electrolytes and are conductive. By eliminating the electrodes, we reduce device fabrication time and complexity, and make the devices more robust. In addition, with our approach, the injection volume depends on the voltage applied to the picoinjection solution; this allows us to rapidly adjust the volume injected by modulating the applied voltage. We demonstrate that our technique is compatible with reagents incorporating common biological compounds, including buffers, enzymes, and nucleic acids.

We have developed a technique for picoinjecting microfluidic drops that does not require metal electrodes. As such, devices incorporating our technique are simpler to fabricate and to use.

Existing methods for picoinjecting reagents into microfluidic drops require metal electrodes integrated into the microfluidic chip. The integration of ...

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 ...

Bioengineering

Micropipette Aspiration of Substrate-attached Cells to Estimate Cell Stiffness

Growing number of studies show that biomechanical properties of individual cells play major roles in multiple cellular functions, including cell proliferation, differentiation, migration and cell-cell interactions. The two key parameters of cellular biomechanics are cellular deformability or stiffness and the ability of the cells to contract and generate force. Here we describe a quick and simple method to estimate cell stiffness by measuring the degree of membrane deformation in response to negative pressure applied by a glass micropipette to the cell surface, a technique that is called Micropipette Aspiration or Microaspiration.
Microaspiration is performed by pulling a glass capillary to create a micropipette with a very small tip (2-50 &mu;m diameter depending on the size of a cell or a tissue sample), which is then connected to a pneumatic pressure transducer and brought to a close vicinity of a cell under a microscope. When the tip of the pipette touches a cell, a step of negative pressure is applied to the pipette by the pneumatic pressure transducer generating well-defined pressure on the cell membrane. In response to pressure, the membrane is aspirated into the pipette and progressive membrane deformation or "membrane projection" into the pipette is measured as a function of time. The basic principle of this experimental approach is that the degree of membrane deformation in response to a defined mechanical force is a function of membrane stiffness. The stiffer the membrane is, the slower the rate of membrane deformation and the shorter the steady-state aspiration length.The technique can be performed on isolated cells, both in suspension and substrate-attached, large organelles, and liposomes.
Analysis is performed by comparing maximal membrane deformations achieved under a given pressure for different cell populations or experimental conditions. A "stiffness coefficient" is estimated by plotting the aspirated length of membrane deformation as a function of the applied pressure. Furthermore, the data can be further analyzed to estimate the Young's modulus of the cells (E), the most common parameter to characterize stiffness of materials. It is important to note that plasma membranes of eukaryotic cells can be viewed as a bi-component system where membrane lipid bilayer is underlied by the sub-membrane cytoskeleton and that it is the cytoskeleton that constitutes the mechanical scaffold of the membrane and dominates the deformability of the cellular envelope. This approach, therefore, allows probing the biomechanical properties of the sub-membrane cytoskeleton.

Here we describe a quick and simple method to measure cell stiffness. The general principle of this approach is to measure membrane deformation in response to well-defined negative pressure applied through a micropipette to the cell surface. This method provides a powerful tool to study biomechanical properties of substrate-attached cells.

Growing number of studies show that biomechanical properties of individual cells play major roles in multiple cellular functions, including cell ...

Transcriptome Analysis of Single Cells

Many gene expression analysis techniques rely on material isolated from heterogeneous populations of cells from tissue homogenates or cells in culture.1,2,3 In the case of the brain, regions such as the hippocampus contain a complex arrangement of different cell types, each with distinct mRNA profiles. The ability to harvest single cells allows for a more in depth investigation into the molecular differences between and within cell populations. We describe a simple and rapid method for harvesting cells for further processing. Pipettes often used in electrophysiology are utilized to isolate (using aspiration) a cell of interest and conveniently deposit it into an Eppendorf tube for further processing with any number of molecular biology techniques. Our protocol can be modified for the harvest of dendrites from cell culture or even individual cells from acute slices.
We also describe the aRNA amplification method as a major downstream application of single cell isolations. This method was developed previously by our lab as an alternative to other gene expression analysis techniques such as reverse-transcription or real-time polymerase chain reaction (PCR).4,5,6,7,8 This technique provides for linear amplification of the polyadenylated RNA beginning with only femtograms of material and resulting in microgram amounts of antisense RNA. The linearly amplified material provides a more accurate estimation than PCR exponential amplification of the relative abundance of components of the transcriptome of the isolated cell. The basic procedure consists of two rounds of amplification. Briefly, a T7 RNA polymerase promoter site is incorporated into double stranded cDNA created from the mRNA transcripts. An overnight in vitro transcription (IVT) reaction is then performed in which T7 RNA polymerase produces many antisense transcripts from the double stranded cDNA. The second round repeats this process but with some technical differences since the starting material is antisense RNA. It is standard to repeat the second round, resulting in three rounds of amplification. Often, the third round in vitro transcription reaction is performed using biotinylated nucleoside triphosphates so that the antisense RNA produced can be hybridized and detected on a microarray.7,8

In this article we describe a simple method for the harvesting of single cells from rat primary neuronal cultures and subsequent transcriptome analysis using aRNA amplification. This approach is generalizable to any cell type.

Many gene expression analysis techniques rely on material isolated from heterogeneous populations of cells from tissue homogenates or cells in ...

Neuroscience

Capillary-based Centrifugal Microfluidic Device for Size-controllable Formation of Monodisperse Microdroplets

Here, we demonstrate a simple method for the rapid production of size-controllable, monodisperse, W/O microdroplets using a capillary-based centrifugal microfluidic device. W/O microdroplets have recently been used in powerful methods that enable miniaturized chemical experiments. Therefore, developing a versatile method to yield monodisperse W/O microdroplets is needed. We have developed a method for generating monodisperse W/O microdroplets based on a capillary-based centrifugal axisymmetric co-flowing microfluidic device. We succeeded in controlling the size of microdroplets by adjusting the capillary orifice. Our method requires equipment that is easier-to-use than with other microfluidic techniques, requires only a small volume (0.1-1 &#181;l) of sample solution for encapsulation, and enables the production of hundreds of thousands number of W/O microdroplets per second. We expect this method will assist biological studies that require precious biological samples by conserving the volume of the samples for rapid quantitative analysis biochemical and biological studies.

Here, we demonstrate a simple production method for size-controllable, monodisperse, water-in-oil (W/O) microdroplets using a capillary-based centrifugal microfluidic device. This method requires only a small sample volume and enables high-yield production. We expect this method will be useful for rapid biochemical and cellular analyses.

Here, we demonstrate a simple method for the rapid production of size-controllable, monodisperse, W/O microdroplets using a capillary-based centrifugal ...

A Versatile Automated Platform for Micro-scale Cell Stimulation Experiments

Study of cells in culture (in vitro analysis) has provided important insight into complex biological systems. Conventional methods and equipment for in vitro analysis are well suited to study of large numbers of cells (&ge;105) in milliliter-scale volumes (&ge;0.1 ml). However, there are many instances in which it is necessary or desirable to scale down culture size to reduce consumption of the cells of interest and/or reagents required for their culture, stimulation, or processing. Unfortunately, conventional approaches do not support precise and reproducible manipulation of micro-scale cultures, and the microfluidics-based automated systems currently available are too complex and specialized for routine use by most laboratories. To address this problem, we have developed a simple and versatile technology platform for automated culture, stimulation, and recovery of small populations of cells (100 - 2,000 cells) in micro-scale volumes (1 - 20 &mu;l). The platform consists of a set of fibronectin-coated microcapillaries ("cell perfusion chambers"), within which micro-scale cultures are established, maintained, and stimulated; a digital microfluidics (DMF) device outfitted with "transfer" microcapillaries ("central hub"), which routes cells and reagents to and from the perfusion chambers; a high-precision syringe pump, which powers transport of materials between the perfusion chambers and the central hub; and an electronic interface that provides control over transport of materials, which is coordinated and automated via pre-determined scripts. As an example, we used the platform to facilitate study of transcriptional responses elicited in immune cells upon challenge with bacteria. Use of the platform enabled us to reduce consumption of cells and reagents, minimize experiment-to-experiment variability, and re-direct hands-on labor. Given the advantages that it confers, as well as its accessibility and versatility, our platform should find use in a wide variety of laboratories and applications, and prove especially useful in facilitating analysis of cells and stimuli that are available in only limited quantities.

We have developed an automated cell culture and interrogation platform for micro-scale cell stimulation experiments. The platform offers simple, versatile, and precise control in cultivating and stimulating small populations of cells, and recovering lysates for molecular analyses. The platform is well suited to studies that use precious cells and/or reagents.

Study of cells in culture (in vitro analysis) has provided important insight into complex biological systems. Conventional methods and equipment for in ...

A Microfluidic Platform for High-throughput Single-cell Isolation and Culture

Studying the heterogeneity of single cells is crucial for many biological questions, but is technically difficult. Thus, there is a need for a simple, yet high-throughput, method to perform single-cell culture experiments. Here, we report a microfluidic chip-based strategy for high-efficiency single-cell isolation (~77%) and demonstrate its capability of performing long-term single-cell culture (up to 7 d) and cellular heterogeneity analysis using clonogenic assay. These applications were demonstrated with KT98 mouse neural stem cells, and A549 and MDA-MB-435 human cancer cells. High single-cell isolation efficiency and long-term culture capability are achieved by using different sizes of microwells on the top and bottom of the microfluidic channel. The small microwell array is designed for precisely isolating single-cells, and the large microwell array is used for single-cell clonal culture in the microfluidic chip. This microfluidic platform constitutes an attractive approach for single-cell culture applications, due to its flexibility of adjustable cell culture spaces for different culture strategies, without decreasing isolation efficiency.

Here, we present a protocol for isolating and culturing single cells with a microfluidic platform, which utilizes a new microwell design concept to allow for high-efficiency single cell isolation and long-term clonal culture.

Studying the heterogeneity of single cells is crucial for many biological questions, but is technically difficult. Thus, there is a need for a simple, yet ...

A Femtoliter Droplet Array for Massively Parallel Protein Synthesis from Single DNA Molecules

Advances in spatial resolution and detection sensitivity of scientific instrumentation make it possible to apply small reactors for biological and chemical research. To meet the demand for high-performance microreactors, we developed a femtoliter droplet array (FemDA) device and exemplified its application in massively parallel cell-free protein synthesis (CFPS) reactions. Over one million uniform droplets were readily generated within a finger-sized area using a two-step oil-sealing protocol. Every droplet was anchored in a femtoliter microchamber composed of a hydrophilic bottom and a hydrophobic sidewall. The hybrid hydrophilic-in-hydrophobic structure and the dedicated sealing oils and surfactants are crucial for stably retaining the femtoliter aqueous solution in the femtoliter space without evaporation loss. The femtoliter configuration and the simple structure of the FemDA device allowed minimal reagent consumption. The uniform dimension of the droplet reactors made large-scale quantitative and time-course measurements convincing and reliable. The FemDA technology correlated the protein yield of the CFPS reaction with the number of DNA molecules in each droplet. We streamlined the procedures about the microfabrication of the device, the formation of the femtoliter droplets, and the acquisition and analysis of the microscopic image data. The detailed protocol with the optimized low running cost makes the FemDA technology accessible to everyone who has standard cleanroom facilities and a conventional fluorescence microscope in their own place.

The overall goal of the protocol is to prepare over one million ordered, uniform, stable, and biocompatible femtoliter droplets on a 1 cm2 planar substrate that can be used for cell-free protein synthesis.

Advances in spatial resolution and detection sensitivity of scientific instrumentation make it possible to apply small reactors for biological and ...

Chemistry

An Ultrahigh-throughput Microfluidic Platform for Single-cell Genome Sequencing

Sequencing technologies have undergone a paradigm shift from bulk to single-cell resolution in response to an evolving understanding of the role of cellular heterogeneity in biological systems. However, single-cell sequencing of large populations has been hampered by limitations in processing genomes for sequencing. In this paper, we describe a method for single-cell genome sequencing (SiC-seq) which uses droplet microfluidics to isolate, amplify, and barcode the genomes of single cells. Cell encapsulation in microgels allows the compartmentalized purification and tagmentation of DNA, while a microfluidic merger efficiently pairs each genome with a unique single-cell oligonucleotide barcode, allowing &gt;50,000 single cells to be sequenced per run. The sequencing data is demultiplexed by barcode, generating groups of reads originating from single cells. As a high-throughput and low-bias method of single-cell sequencing, SiC-seq will enable a broader range of genomic studies targeted at diverse cell populations.

Single-cell sequencing reveals genotypic heterogeneity in biological systems, but current technologies lack the throughput necessary for the deep profiling of community composition and function. Here, we describe a microfluidic workflow for sequencing &gt;50,000 single-cell genomes from diverse cell populations.

Sequencing technologies have undergone a paradigm shift from bulk to single-cell resolution in response to an evolving understanding of the role of ...

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. ...