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

All experiments were performed under ethics agreement EK202-N-56 and approved by ETH Zurich's ethics commission. Handling of human cells was performed in a laminar flow cabinet contained in a biosafety level 2 laboratory.

NOTE: The following sections detail the protocol to measure time-resolved cytokine secretion on a single-cell level. The procedure outlined here is applied to the stimulation of peripheral blood mononuclear cells (PBMC) with lipopolysaccharide (LPS) and the parallel measurement of the cytokines IL-6, TNFα and IL-1β. However, if required, the protocol can be adapted to other cell types, stimulants, and cytokines.

1. Observation chamber fabrication

NOTE: To avoid movement of the droplets during imaging, an observation chamber is prepared with a height that is around 10% smaller than the droplet diameter.

1. Preparation of the cutting double-sided tape and top glass slide
   1. Draw or load the desired design of the chamber cutout in the Design tab of the cutting software. For the specific dimensions used here, see Figure 2E.
   2. Fix the double-sided adhesive tape of 32 µm thickness to the adhesive cutting mat using tape and place the cutting mat in the automatic cutting machine.
   3. Cut the chamber design out of the tape, while paying attention to cutting the long edges of the chamber in the same direction for easier detachment in step 1.3.
   4. Store the tape cutouts at room temperature for long-term storage. For short-term storage, store them at -20 °C and remove them only shortly before step 1.3. for easier handling.
   5. Drill two holes of around 1 mm diameter in the middle of one standard microscope slide (76 mm x 26 mm x 1 mm), with the distance between the two holes of around 3.5 cm.
2. Cleaning and plasma activation of glass slides
   1. Clean one glass slide with holes and one without holes using soap. Rinse well with distilled water and dry using lint-free precision wipes.
   2. Place the glass slides in a plasma cleaner and plasma-treat the top surfaces at 55 W for 10 min. Remove the glass slides and proceed to step 1.3.
3. Chamber assembly
   1. Place the glass slide with holes on a clean surface with the plasma-activated side facing upwards, without touching the activated surface.
   2. Remove the protective layer from one side of the double-sided adhesive tape, in the same direction of cutting. Without touching align the tape cutout with the edges of the glass slide and the drilled holes, and slowly put the tape in contact with the glass slide starting from the short edge.
      NOTE: Pay attention not to generate any stretches or folds in the tape since this will result in incorrect chamber heights. Since this step is error prone and needs some practical experience, it is advisable to prepare several glass slides in parallel.
   3. Remove the second protective layer from the tape, again in the direction of cutting, and place the second glass slide without holes with the activated surface facing downwards. Press the whole surface of the two glass slides together by placing a flat board on top and pressing down with upper body strength for about 10 s.
   4. After assembling the two glass slides, flip the chamber so that the two holes are facing you. Glue the nanoports to the two holes by putting a small amount of UV-curable glue in the ring below the port and placing the port on top of the hole in the glass slide. Add a ring of UV curable glue around the port and cure the glue with a UV lamp. The chamber should now look as depicted in Figure 2E. Proceed immediately to step 1.4.
      NOTE: UV light can damage the eyes and skin. Wear appropriate protective equipment.
4. Fluorophilic coating of the chamber surface
   NOTE: This step should take place within 1 h after plasma treatment of the glass slides (step 1.2.2) to ensure good coating efficiency.
   1. Freshly prepare 1 mL of 1% fluorosilane solution (1H,1H,2H,2H-perfluorodecyltrichlorosilane) in fluorinated oil (HFE-7500) and fill it in a syringe. Push the coating solution through a PTFE syringe filter and a 27G x 0.75 inch needle connected to 0.3 mm x 0.76 mm PTFE microtubing, into the observation chamber.
   2. After 1 min incubation, flush the coating solution out of the chamber using nitrogen pressure under a fume hood. Rinse the chamber with fluorinated oil (HFE-7500 only) using another syringe assembly.
   3. Store the chamber filled with fluorinated oil with closed inlets at room temperature (RT). After each experiment, wash out cells and droplets directly to ensure good preservation of the coating.
      NOTE: The protocol can be paused here, and the chambers can be stored and reused for several months.
5. Chamber holder with magnets
   1. For alignment of the magnetic nanoparticles, apply a static magnetic field to the observation chamber during droplet encapsulation and imaging. For this, place the chamber in a custom 3D-printed microscopy holder (see Figure 2D, and the file found in Bounab et al.¹⁷ Supplementary Data 4) that holds two neodymium magnets along the long sides of the chamber.

2. Nanoparticle functionalization

NOTE: The process for the nanoparticle functionalization is similar for each cytokine, the only difference being the addition of cytokine-specific capture antibodies. The functionalization for each cytokine is performed in different, individual reaction tubes in parallel. Prior to this protocol, the TNFα capture antibody and IL-1β detection antibody were labeled in-house with biotin and Alexa Fluor 647, respectively. Conjugation was performed according to the manufacturer's protocol found on the vendor's website (see links in Table of Materials) and the antibodies were aliquoted and stored at -20 °C.

1. Add 50 µL of streptavidin-functionalized nanoparticles (diameter (Ø) 300 nm) into the tube destined for TNFα detection, 50 µL for IL-1β, and 100 µL for IL-6. Dilute the nanoparticle solution 1:1 (v/v) in phosphate-buffered saline (PBS).
2. Add to each tube 1/20 (v/v) of the respective volume of the biotinylated capture antibodies (stock concentrations at 0.5 mg/mL) and incubate for 30 min at RT.
   NOTE: When adding small volumes to the nanoparticle solution, deposit the volume at the top of the tube and wash it down multiple times with the bulk of the solution. This ensures proper mixing and prevents the formation of aggregates.
3. Add 1/100 (v/v) of 1 mM D-biotin solution to the tube and incubate for 5 min at RT. This results in a final biotin concentration of 10 µM.
   NOTE: Excess biotin blocks free binding sides on the nanoparticles and reduces unwanted aggregate formation.
4. Collect the particles by holding a neodymium magnet close to the tube. Wait until the supernatant is clear and discard the supernatant.
   NOTE: The magnets used throughout the assay exhibit very strong attractive forces, which can cause physical harm if two magnets snap together accidentally.
5. To reduce non-specific adsorption to the nanoparticle surface, immediately resuspend the nanoparticles in 0.5x the final volume of step 2.1 of Pluronic F-127 (10%) and 0.5x the volume PBS. Incubate the solution for 30 min at RT.
6. Collect the particles using the magnet, discard the supernatant, and resuspend in 1x the volume of storage buffer (RPMI 1640, 5% knockout serum replacement, 1% Pen/Strep, 1% recombinant human serum albumin (HSA), 25 mM HEPES, 0.1% Pluronic F-127). Incubate the solution for 30 min at RT.
   NOTE: The protocol can be paused here, and the particles can now be stored for up to 1 week at 4 °C.
7. Immediately before encapsulation, resuspend the particles by pipetting and mix the conjugated nanoparticles at a ratio of 2:1:1 (v/v) for IL-6:TNFα:IL-1β, respectively.
   NOTE: The different ratios of functionalized nanoparticles depend on the used antibody pair for each cytokine and have been determined experimentally through calibration samples to obtain an optimal dynamic range.
8. Wash with complete media (RPMI 1640, 10% FBS, 1% Pen/Strep, 25 mM HEPES) by collecting the particles with the magnet, discarding the supernatant and resuspending. Repeat this step but resuspend only in 0.5x the volume of complete media from step 2.7.
9. Add the differently labeled IL-6, TNFα, and IL-1β detection antibodies to the solution to reach a final concentration of 10 nM each. The solution is now ready to be used for the droplet experiments.

3. Cell preparation

NOTE: PBMC were isolated from a buffy coat received from the Zürich blood bank. The cells were frozen and stored in cryovials (1 x 10⁷ cells/vial) in liquid nitrogen for several months.

1. Cell thawing
   1. At 1 h before starting the experiment, leave the complete media and MACS buffer (2 mM EDTA, 0.5% BSA in DPBS, sterile filtered) to warm up at RT. Prepare the tube containing the cells by adding 9 mL of complete media into a 15 mL tube and keeping it in the water bath at 37 °C.
   2. Retrieve a PBMC cryovial (containing ~1 x 10⁷ cells) from its storage in liquid nitrogen. Swirl the cryotube in the water bath at 37 °C until only a small amount of ice remains.
   3. Wipe the tube with 70% EtOH and transfer it to the laminar flow cabinet. Add 1 mL of pre-warmed complete media to the cryovial, mix gently and transfer all the cells into the tube containing warm complete media. The cryovial can be washed with 1 mL of warm complete media to recover the maximum number of cells.
   4. Spin the cells at 500 x g for 5 min at RT, discard the supernatant and resuspend gently with a pipette the cell pellet with 1 mL of complete media. Add 9 mL of complete media.
   5. Spin the cells at 500 x g for 5 min at RT. Discard the supernatant and resuspend as previously into 1 mL of complete media.
   6. Count the cells using the available cell counter. In this case, an automated cell counter was used. Cells were counted by mixing 10 µL of cell suspension with 10 µL of Trypan blue and transferring 10 µL of the mixture into the cell counting slide.
2. Staining and FcR blocking
   1. Calculate the total number of cells and the volume needed to resuspend the cells at 2 x 10⁶ living cells/mL. Prepare the cell staining solution (CellTrace Violet) by diluting the stock (5mM) 1000x in PBS (working concentration of 5 µM).
   2. Spin the cells at 500 x g for 5 min at RT. Discard the supernatant and resuspend the cells in the calculated volume of cell staining solution prepared in step 3.2.1. Incubate the cells at 37 °C for 5 min.
   3. At the end of the incubation, quench the remaining dye in the solution by adding complete media (at least 2x the volume of dye solution). Spin the cells at 500 x g for 5 min at RT.
   4. Discard the supernatant, resuspend the cell pellet into 60 µL of MACS buffer and add 20 µL of human FcR block per 1 x 10⁷ cells. Incubate the cells for 10 min at RT.
   5. Fill the tube to 10 mL with MACS Buffer and spin the cells at 500 x g for 5 min at RT.
   6. Discard the supernatant and resuspend the cells in 1mL of complete media Count the cells as described in step 3.1.6.
3. Cell stimulation with LPS
   1. Using the cell count, dilute the cells at 1 x 10⁶ cells/mL and transfer 2 mL of cells into each well in an ultra-low binding 6-well plate.
   2. Dilute LPS in complete media and add it into the well containing the cells for a final concentration of LPS of 1 µg/mL. Incubate the cells for 6 h at 37 °C.
4. Preparation for encapsulation
   1. At the end of the stimulation time, transfer the cell suspension into a new 15 mL tube.
   2. Add 1 mL of complete media into the empty well. With a cell scraper, detach the remaining cells. Transfer the cells into a new 15 mL tube. Wash the well with 1 mL of complete media and transfer to another 15 mL tube.
   3. Spin the two tubes at 500 x g for 5 min at RT and transfer 1 mL of the undiluted supernatant solution (from the first tube containing the non-washed cells) to a new tube for further analysis if needed (e.g., ELISA).
   4. Discard the rest of the supernatants.
   5. Resuspend the pellets in 0.5 mL of complete media, combine the cells from the same well, and transfer them into a centrifuge tube. Count the cells as described in step 3.1.6.
   6. Spin the cells at 500 x g for 5 min at RT and discard most of the supernatant (leaving approximately 100 µL). Without resuspending the pellet, add very carefully 200 µL of complete media.
   7. Discard the supernatant. Resuspend the cells in complete media at a concentration of 6.6 to 13.3 x 10⁶ cells/mL to achieve an average cell number per droplet of λ = 0.2-0.4 for encapsulation, as defined in step 8.6.
      NOTE: Steps 3.4.6 and 3.4.7 should be performed immediately before the encapsulation to avoid cytokine secretion into the supernatant. The number of cells per droplet follows a Poisson distribution: , where P shows the fraction of droplets containing X cells and λ is the mean number of cells per droplet.

4. Encapsulation and droplet production

NOTE: The encapsulation of cells in droplets is enabled by a microfluidic droplet generator chip, for which the fabrication is described in great detail elsewhere¹⁷. Alternatives are commercially available (see example in Table of Materials). A suitable droplet generator chip design has two inlets for aqueous phases, one inlet for the oil phase, and one outlet for the generated droplets. Furthermore, a suitable commercial droplet generator chip should enable the production of water in fluorinated oil droplets of 40-60 pL volume. The protocol described here results in water/oil emulsions (droplets) with a diameter of 50 µm. Using various options for altering the protocol can result in bigger or smaller droplets.

1. Preparation of the syringe pump (Figure 2A)
   1. Fill a 1 mL syringe with 500 µL of continuous phase consisting of 2% 008-Fluorosurfactant in HFE-7500 fluorinated oil. Connect a 27G x 0.75 inch needle to 0.30 mm x 0.76 mm PTFE microtubing and mount the assembly on the syringe and subsequently onto the syringe pump.
      NOTE: Ensure no air remains in the syringe or cannula as this prevents consistent flow rates.
   2. Prepare two custom-made pipette tip connectors for the aqueous phases (Figure 2B): Punch a hole with a Ø0.75 mm biopsy punch into the middle of a ~5 mm high PDMS cutout with Ø6 mm. Pull ~3 cm of PTFE tubing (0.56 mm inner diameter, 1.07 mm outer diameter) through the hole in the PDMS cutout and push the assembly into the top of a 200 µL pipette tip. Connect the other side of the tube to a 23Gx 1.25 inch needle. Seal the connector by spreading UV-curable glue on top of the pipette and cure it with UV light.
      NOTE: Since UV light is harmful to the eye, wear UV-blocking goggles for protection.
   3. Fill two 1 mL syringes with 500 µL of light mineral oil, attach two 23G needles with the custom-made attachments, and mount both onto the syringe pump.
   4. Aspirate 30 µL of nanoparticle and 30 µL of cell solution into the pipette tips of the aqueous phases using the syringe pump control software.
   5. Prepare an observation chamber by cleaning the surface with water to remove dirt and dust and dry it with precision wipes. Clamp the chamber into the printed chamber holder equipped with two neodymium magnets.
      NOTE: Make sure the magnets point in the right direction (attracting each other) to form an elongated aggregate.
   6. Slightly angle the chamber (30°). Open both ports and plug a paper towel in the upper port to absorb the excess outer phase during filling.
2. Droplet production and chamber filling
   1. Connect the continuous phase via tubing to the top inlet of the microfluidic chip (Figure 2A,C,F). Flush the chip for around 30 s with continuous phase using a flow rate of 1800 µL/h.
   2. Connect the pipette tips of the aqueous solutions to the two middle inlets (Figure 2A,C,F).
   3. Start the flow of the aqueous solution with 200 µL/h each and let the channels and the outlet be filled with liquid. When using magnetic nanoparticles, a homogenous, brown-red colored solution should flow out of the chip outlet.
   4. Once the liquid appears at the outlet, start the fluorinated oil phase flow at 800 µL/h and wait until a stable droplet production is established, confirmed by the outflow of a homogenous, grey, shiny solution at the outlet.
   5. Once a stable production of droplets is established, collect the produced droplets by connecting PTFE microtubing (0.3 mm inner diameter x 0.76 mm outer diameter) to the outlet port and direct them into an observation chamber by passing the microtubing through the ferrule module of a finger-tight one-piece fitting (Figure 2A).
   6. If proper droplet production occurs, a homogenous, shiny liquid should fill the chamber with a straight front from bottom to top.
   7. Once the chamber is filled, stop the flow and close off the ports with port plugs using finger-tight pressure.
      ​NOTE: Pay attention to not close the chamber too tightly. Trapping or influx of air can lead to movements of the droplets and thus compromise tracking during the measurement.
   8. After droplet production, flush the chip with fluorinated oil and blow out any reminders of fluid with nitrogen to preserve its function. Chips can be reused multiple times and stored for months as long as they are not clogged.

Figure 2: Overview of the microfluidic setup. (A) Setup for droplet encapsulation with the syringe pump, the droplet generation chip, and the observation chamber and microscope holder. (B) Picture of the punched PDMS plug (top) to form a connector to a 200 µL pipette tip (bottom), as described in protocol step 4.1.2. (C) Images of the connection of tubing ang pipette tips to the droplet generation chip. (D) Picture of the chamber placed inside the custom 3D-printed microscope holder with two magnets on top and bottom. (E) Photo of observation chamber (with white tape for illustration). (F) Layout of the microfluidic chip for droplet creation (scale bar: 750 µm). This figure has been modified from¹⁷. Please click here to view a larger version of this figure.

5. Image acquisition and measurement

NOTE: Image acquisition is performed on a standard inverted epi-fluorescence microscope enclosed in an incubator, allowing measurements at 37 °C. The here described settings are specific for a Nikon Eclipse Ti2 microscope running with the NIS Elements software (V. 5.30.04) equipped with an Orca Fusion camera but are generally adaptable to any other fluorescence microscopes and cameras.

1. Setting measurement parameters
   1. To set the size of the image, select an array size of 10 x 10 images. This array will contain roughly 50,000-70,000 droplets. Use 1% overlap and activate blending for image stitching.
   2. To set the number of measured channels, select the DAPI channel for cell detection, FITC, TRITC, Cy5 channels for cytokine detection (beadlines) and the BF channel for droplet detection. Use pixel binning 2 x 2 and 16-bit for bit depth. Adapt the camera settings to achieve in-droplet intensity pixel values that do not reach the maximum of the camera for every fluorescence channel.
      NOTE: Exact exposure times and lamp intensities for every channel depend on the used model and reagents and are established before generating calibration curves (step 7). Using the same acquisition settings in calibration and cell measurements is important for accurate quantification.
   3. To set up the time-resolved measurement, select a measurement every 30 min for 9 measurements in total.
      NOTE: Measurement parameters might differ for used cells, stimulants, reagents, measured cytokines, incubation temperature, and microscope models.
2. Starting the measurement
   1. Mount the chamber holder onto the microscope with a well-plate format stage (Figure 2D) and switch to the brightfield (BF) channel using the 10x objective.
   2. Focus on the immobilized droplets in BF and ensure that the assembly is mounted in a perfect plane by panning around and adjusting if necessary. Move to the middle of the chamber for the subsequent steps.
   3. Activate the automated focusing system (PFS) and set it to the optimal measurement plane on the BF channel so that droplet edges appear as black, sharp circles that can be easily distinguished from the oil phase and background.
      NOTE: Measurements are also possible without an automated focusing system, but if the microscope is equipped with one, we highly recommend using it. This improves the measurement quality for large and heavily stitched images.
   4. Go through all the fluorescence channels and set the optimal measurement plane for each. For relocation measurements on the FITC, TRITC and Cy5 channels, ensure the nanoparticle aggregate is in perfect focus, for the DAPI channel, ensure the cells are in focus.
      NOTE: Optimal focal planes and z-values might differ for all measured channels. Make sure to save individual PFS offsets for every channel.
   5. Before starting the measurement, go through all channels to double-check the individual foci and wait 5 min to equilibrate since movement might occur initially while the solutions warm up.
   6. Start the measurement. After generating the first image, check for any irregularities (focus, moving droplets, wrong channels, etc.). Restart the acquisition if needed, or in the case of air, refill the chamber (start from step 4.1.4). Leave the assembly to image the droplets over 4 h.

6. Image analysis

1. Install image analysis software (DropMap Analyzer App v 4.023) in MatLab (https://github.com/ESPCI-LCMD/MiMB) and transfer the generated .nd2 file from the experiment to an analysis computer.
2. Open the application. Select the settings specified, otherwise leave the default value: CH1: DAPI, WD (whole droplet) selected; CH2: FITC, BL (beadline) selected; CH3: TRITC, BL selected; CH4: Cy5, BL selected; Max Drop Diameter (µm): 70; Drop Detection: Full; Tracking: Yes. Press on the Start button (fruit icon) to select the .nd2 file location and start the analysis.
3. After a few minutes, the program will show an example section of the image (Figure 3A). Press Space until you find one suitable for droplet detection, then press Enter. In the same image section, draw a rectangle in a representative area for finding thresholding parameters to detect droplets.
4. After a few minutes, another window will open showing the intensity distribution of the DAPI channel. Drag and drop the slider to detect only the signal from the stained cells and click Done in the upper right corner.
5. After segmenting the image into single droplets with diameters smaller than Max. Drop Diameter (µm), the program will now perform the following steps for each droplet, timepoint, and fluorescence channel without further user input (see Figure 3).
   1. The software calculates the moved pixels of the droplet between time points (droplets moving more than 40 pixels are excluded automatically).
   2. The software measures the average fluorescence value of the whole droplet, detecting and measuring the mean beadline intensity by finding the brightest pixel on a horizontal line and averaging all pixel intensities on a vertical line from top to bottom of the droplet. This is done automatically and is used to calculate average beadline relocation values (Figure 3B) according to the equation:
      
   3. The software calculates the percentage of total pixels in the droplet area above the threshold set on the DAPI channel.
6. The resulting .xslx file will contain the following columns of interest for further analysis: DropIdX (ID of the droplet tracked over time), TrueCentroid_ t*2-1 and t+2 (x and y coordinates, respectively, of the droplet center for timepoint t), DiameterMicrons (droplet diameter in µm), TrackingMove (number of pixels moved over the whole measurement time), FluoChannel_BL_Ratio_t (relocation value for FluoChannel at timepoint t), DAPI_WD_PosPxlCount_t (number of pixels above threshold in the whole droplet in the DAPI channel at timepoint t).

Figure 3: Image analysis performed by the image analysis software. (A) Droplets are detected in the brightfield (BF) channel using a Hough transformation, marking each droplet with a red circle. Scale bars: 200 µm. (B) Within each droplet, the nanoparticle beadline is identified through the brightest pixels in the horizontal plane and the fluorescence intensities averaged for all pixels spanning from top to bottom of the droplet. Additionally, the cell is identified through a pixel percentage >0 above threshold for the whole droplet area. Scale bar: 20 µm. (C) The analyzer software compares the fluorescence intensity on the nanoparticles to the droplet background for the FITC, TRITC and Cy5 channels over all the measured time points for every individual droplet. Shown are timepoints 0, 4 (120 min) and 9 (240 min). To manually check for correct droplet and cell detection, the DAPI and BF channels are displayed as well. Please click here to view a larger version of this figure.

7. Calibration

NOTE: For a quantitative readout, the calibration of cytokine concentrations to fluorescence relocation values needs to be performed once, as differences between different experimental setups can occur. All required steps are detailed in the previous protocol sections as referenced.

1. Prepare nanoparticles as described in step 2.
2. Reconstitute the human IL-6, TNFα and IL-1β recombinant proteins according to the manufacturer's instructions.
   NOTE: Ensure that frozen aliquots are thawed only once and used promptly.
3. Prepare a 2-fold dilution series for all three proteins together using complete media (10% FBS, 1% Pen/Strep, 25 mM HEPES) with a starting concentration of 80 nM down to 0.625 nM.
4. Perform the encapsulation as described in step 4 with functionalized nanoparticles in the first and RPMI only in the second aqueous phase. This measurement serves as the blank and the measured standard deviation is used for data analysis later.
5. Wait 5 min and image the droplets as described in step 5. Take 3 images with an array size of 2 x 2 in the respective fluorescence channels.
6. Repeat steps 7.4 and 7.5 with all prepared calibration solutions, starting with the lowest and ending with the highest concentration.
7. Analyze the images as described in step 6. Do not use the WD option for the DAPI channel and set Tracking to No.
8. The analysis outputs fluorescence relocation values for every measured droplet in one image. Extract the median and standard deviation for every fluorescence channel. Average the median and standard deviation for every measured image per concentration.
9. Generate a calibration curve by plotting the averaged median relocation against the measured concentrations of each recombinant protein.
10. Fit the curves using a one-phase association: ,
    with Y = relocation at x, Y₀ = relocation of blank measurement and x the used concentration. The obtained calibration curve is used to quantify relocation values as described in step 8.
    NOTE: Only fit values up to the highest measured relocation and exclude values from higher concentrations with lower measured relocation. A decrease in measured relocation values at higher concentrations is expected and occurs due to the Hook effect and limited binding capacity of the nanoparticles.

8. Data analysis

1. Exclude droplets with a TrackingMove value superior to 10, i.e. which moved more than 10 pixels over the time course of the measurement.
2. Identify droplets containing stained cells (DAPI channel) in the first timepoint by sorting for droplets with values superior to 0 in the column DAPI_WD_PosPxlPercent_1.
3. Identify droplets containing secreting cells by applying the following 3 criteria to the fluorescence relocation of every fluorescence channel (FluoChannel_BL_Ratio_t columns).
   1. Identifying droplets with increasing relocation values, by sorting for a positive slope over measurement time.
   2. Identifying droplets with relocation values reaching the limit of detection (LOD). A droplet is selected when the maximum fluorescence relocation over the measurement time is superior to the LOD, calculated as described elsewhere²⁵: , where μ_{Relocation t0} is the median of all the relocation values at timepoint 0 and σ_BLK the standard deviation of the blank measured during the calibration, each are cytokine specific.
   3. Verifying that the increase in relocation value is significant by checking that the change between maximum and minimum measured fluorescence relocation over the measurement time is superior to: .
4. Identify co-secreting cells by meeting the criteria described in step 8.3. for more than one fluorescence channel simultaneously.
5. Repeat step 8.3. for all droplets containing no cell (DAPI_WD_PosPxlPercent_1 = 0). Use these droplets to calculate the false-positive percentage.
6. Determine the accurate λ value of the measurement by randomly selecting 200 – 500 droplets and inspecting them with the Verify and sort function of the image analysis software. Count the number of cells in these droplets and calculate:
   λ = number of counted cells / number of analyzed droplets
7. Calculate the total number of encapsulated cells for the measurement by:
   Total cell count = λ × number of analyzed droplets
8. Calculate the percentage of secreting cells using the determined cell count. Additionally, calculate the false-positive percentage for every cytokine (usually less than 3%-5% relative to the number of real positives per channel) and use them as an internal control for experimental consistency and reproducibility.
9. To calculate secreted cytokine concentrations, convert relocation values to concentration using the established calibration equations from step 7.10.
10. Calculate secretion rate (SR) between time points by using the following equation.
    
11. Calculate the average secretion rate over the measurement by averaging the individual secretion rates between time points. If the maximum measurable relocation was reached before the end of the measurement, set the concentration to the maximum measurable concentration (this value is cytokine-specific and corresponds to the maximum concentration measured and used in the calibration curve in step 6.10), and do not calculate further concentration. Calculate the secretion rate and average only up to this time point.
    NOTE: If less than 50 secreting cells are detected in one fluorescence channel, the droplets should be examined visually through the verify and sort function and droplets having fluorescence or nanoparticle aggregates can be excluded from the analysis.
12. To extract further parameters from each single cell's secretion curve, perform a least square fit to the time-concentration curve for each cell and cytokine using a custom Python script (available upon request). The fitted function is a sigmoidal curve following the formula described below (Fits with R²<0.95 are excluded from the following steps):
    
    where C corresponds to the concentration plateau [nM], t₅₀ to the shift of the half-maximum [s], and m to the Hill slope [min^-1]. From these parameters, the following curve descriptors are extracted as described below.
    1. C_max [nM]: Maximum measured concentration.
    2. : Time of secretion start, where the fit reaches 10% of C.
    3. : secretion rate as the approximated linear slope of the curve between 10% and 90% of the time-concentration curve.