A Vaccine-Pulsed Dendritic Cell-Based In Vitro Assay to Induce Lymphocyte Proliferation

1. Production of Naïve MoDCs

NOTE: Whole blood samples were obtained from a single pathogen-free calf by jugular vein puncture with heparinized vacutainers (eight 9 mL blood tubes were used for this study). Transport the blood in an ice box. Store the samples at 2-4 °C for later use, or process them immediately. Keep the blood rotating to avoid blood clotting. Sterilize the vacutainers with 70% ethanol. All the following experiments were performed with one biological sample and six technical replicates.

1. Density gradient centrifugation to isolate PBMCs from the heparinized blood
   1. Mix the blood well by inverting it 10x.
   2. Using a 10 mL pipette, transfer 20 mL of heparinized blood to a sterile 50 mL tube, and dilute it with 10 mL of phosphate-buffered saline (PBS).
   3. Pipette 15 mL of lymphocyte isolation medium to a sterile 50 mL tube.
      NOTE: Allow the lymphocyte-isolating medium to reach room temperature prior to pipetting.
   4. Tilt the 50 mL tube containing the lymphocyte isolation medium to a 45° position. Point the tip of a 25 mL pipette perpendicularly, and carefully layer the 30 mL of blood-PBS on top of the lymphocyte isolation medium, without mixing the two. Very slowly, bring the 50 mL tube back to a vertical position.
   5. Centrifuge at 800 × g for 35 min at 20 °C with maximum acceleration and without braking (deceleration function turned off).
   6. Use a Pasteur pipette to collect the PBMC layer (the thin white layer right after the first layer of plasma) and transfer it into a new 50 mL tube.
      NOTE: Density gradient centrifugation separates whole blood into different layers. As a result, the erythrocytes settle at the bottom as a pellet, the mononuclear cells (PBMCs) settle at the interface layer, and the plasma forms the top layer. Granulocytes are found between the pellet and the PBMC layer.
   7. Wash the harvested PBMCs 2x by adding PBS up to a volume of 40 mL and mixing thoroughly by pipetting up and down. Then, centrifuge at 500 × g at 4 °C for 7 min with maximum acceleration and maximum deceleration.
   8. Discard the supernatant by decantation, resuspend the pellet in 15 mL of 1x ammonium-chloride-potassium (ACK) buffer, and incubate at room temperature for 10-15 min.
      NOTE: Do not incubate for more than 15 minutes. Commercially available ACK buffer is used to ensure the lysis of the residual red blood cells (RBCs) present in the PBMC fraction.
   9. Add PBS up to a volume of 40 mL, and then centrifuge at 500 × g and 4 °C for 7 min with maximum acceleration and deceleration.
   10. Discard the supernatant by decanting, and repeat step 1.1.9.
   11. Discard the supernatant, and resuspend the pellet in 10 mL of complete culture medium (at room temperature).
       NOTE: The complete culture medium is composed of RPMI 1640 cell culture medium supplemented with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin.
2. Magnetic separation of the CD14^+/− cells from the PBMC population using CD14-conjugated magnetic beads
   1. Count the cells with a clean hemocytometer cell counter using 10 µL of cell suspension mixed with 10 µL of trypan blue (0.4%) solution.
      ​NOTE: Trypan blue dye is a toxic chemical and potential carcinogen. It should be handled while wearing personal protective equipment (PPE) to avoid contact, and waste should be discarded in an airtight specialized toxic waste container. Dead cells will appear blue, whereas live cells will appear clear under the microscope.
   2. Centrifuge for 7 min at 500 × g.
   3. Discard the supernatant using a pipette, and resuspend the pellet in fluorescence-activated cell sorting buffer (FACS) using a volume of 40 µL for every 1 × 10⁷ cells. Mix thoroughly using a pipette.
      NOTE: FACS buffer is composed of 2% FBS and 2 mM ethylenediaminetetraacetic acid (EDTA) dissolved in PBS.
   4. Add 5 µL of CD14 microbead buffer per 1 × 10⁷ cells, and mix thoroughly using a pipette.
   5. Incubate for 30 min at 4-8 °C for the positive selection of bovine CD14⁺ monocytes. Vortex every 15 min during the incubation period.
   6. Optional step (for flow cytometry analysis): Add 10 µL of CD14-FITC (fluorescein isothiocyanate) staining antibody with subsequent incubation for 15 min at 4-8 °C. Refer to the flow cytometry analysis in step 5 for details.
   7. Add 1 mL of FACS buffer, and centrifuge for 7 min at 500 × g.
   8. Discard the supernatant, and resuspend the pellet in 500 µL of FACS buffer per 1 × 10⁸ cells.
   9. Insert the immunomagnetic cell separation column into the separator (magnetic column holder), and place a 15 mL collection tube under the column outlet for collecting the flowthrough.
   10. Wash the column with 1 mL of degassed FACS buffer. After rinsing, replace the used 15 mL tube with a new one.
   11. Allow the cell suspension (from step 1.2.8) to pass through the column by pipetting 1 × 10⁸ cells in 500 µL of buffer at a time.
       NOTE: Pay attention not to generate air bubbles while mixing the cells before allowing them through the column.
   12. Rinse the column 3x with 3 mL of degassed FACS buffer each time.
   13. Collect the flowthrough, and label this first eluted fraction as the CD14⁻ cell fraction (PBMCs minus CD14⁺ monocytes); this is also termed the naïve lymphocyte fraction.
       NOTE: The CD14⁻ cells are maintained in a complete culture medium until antigen-pulsed MoDCs are produced.
   14. Remove the column from the separator.
   15. Place a new sterile 15 mL tube under the column for the collection of the effluent.
   16. Pipette 5 mL of FACS buffer into the column, and immediately push it through using a plunger.
   17. Collect the flowthrough, and label this second eluted fraction as the CD14⁺ cell fraction; this is also called the naïve monocyte fraction.
   18. Add complete culture medium to the harvested CD14⁺ monocytes to attain 1 × 10⁶ cells/mL.
       NOTE: Count the live cells in the eluted CD14⁺ cell fraction to estimate the volume of the complete culture medium required to attain the desired cell count.
   19. Take a sterile 24-well plate, and add 1 mL of the cell suspension (1 × 10⁶ cells/mL) to each well.
3. Differentiation of the CD14⁺ naïve monocytes into naïve MoDCs by the addition of 3% w/v cytokine cocktail (GM-CSF + IL-4) with a total of 5 days of incubation
   1. Supplement each well containing CD14⁺ monocytes with 40 µL (3% w/v) of the cytokine cocktail provided in the kit.
   2. Incubate the plate in a humidified incubator with 5% CO₂ and at 37 °C for 48 h.
   3. On day 2, transfer half of each well's content (500 µL) using a pipette to individual 1.5 mL tubes and centrifuge at 500 × g at 4 °C for 7 min.
   4. Discard the supernatant and resuspend the pellet in 500 µL of fresh complete culture medium.
   5. Transfer 500 µL of this cell suspension from step 1.3.4 back to each designated well so that the final volume is 1 mL.
   6. Enrich each well with 20 µL of cytokine cocktail.
   7. Incubate the culture for 72 h in a humidified incubator with 5% CO₂ and at 37 °C.
   8. After the incubation on day 5, remove the plate from the incubator.
      NOTE: Each well will contain 1 mL of a cell suspension of naïve MoDCs. The number of MoDCs will be a percentage (~20%) of the original monocytes plated (1 × 10⁶/mL).

2. Generation of antigen-pulsed MoDCs

NOTE: A commercially available and clinically approved vaccine against rabies virus (RV) can induce the differentiation of naïve MoDCs into mature antigen-presenting MoDCs. Use 0.1% (~1 µL) of a single RV vaccination dose to generate antigen-pulsed MoDCs. Furthermore, it is preferred to produce RV-pulsed MoDCs in the same culture plate used to generate naïve MoDCs because transferring the naïve MoDCs to a new 24-well plate will negatively affect them.

1. Add 1 µL/mL of RV suspension to 1 mL of naïve MoDC culture in the 24-well plate, and incubate for 48 h at 5% CO₂ and 37 °C.
   NOTE: naïve MoDCs cultured without RV stimulation are used as a background control (and as non-specific stimulation for co-culture with lymphocytes in the next steps).
2. After incubation, on day 7, keep the 24-well plate containing the antigen-pulsed MoDCs on ice for 10 min.
3. Add 1 mL of ice-cold PBS per well. Mix each well thoroughly by pipetting, and transfer the suspension to a 15 mL tube.
4. Wash the wells with 2 mL of ice-cold PBS to collect the residual cells left within each well.
5. Transfer the contents into their respective tubes, and centrifuge the cell suspension at 500 × g for 7 min.
6. Discard the supernatant, resuspend the cell pellet (antigen-pulsed MoDCs) in a complete culture medium, and adjust the volume to a final cell concentration of 1 × 10⁵ cells/mL.
   NOTE: Count the live cells to estimate the volume of medium required to attain the desired cell count. Use RV-pulsed MoDCs from day 7 for the MoDC-lymphocyte co-culture system.

3. MoDC-lymphocyte co-culture

NOTE: The in vitro MoDC-lymphocyte co-culture system determines the ability of MoDCs to prime antigen-specific lymphocytes. The different treatment groups of cells after 16 days of co-culture include specific, non-specific, and control. The specific group is defined as lymphocytes co-cultured with RV-pulsed MoDCs; the non-specific group is defined as lymphocytes co-cultured with non-antigen-pulsed MoDCs; and the control group is defined as lymphocytes cultured without MoDCs.

1. Add complete culture medium to the eluted naïve lymphocyte fraction (CD14⁻ cells) to attain a cell concentration of 2 × 10⁶ cells/mL.
   NOTE: Count the live cells in the CD14⁻ cell culture that have been maintained in complete culture medium in a 24-well plate since their collection to estimate the volume of medium required to attain the desired cell count.
2. On day 7, take a sterile 24-well plate, and seed the wells with 1 mL of naïve lymphocyte cell suspension (2 × 10⁶ cell/mL) plus 1 mL of antigen-pulsed or non-antigen-pulsed MoDC suspension (1 × 10⁵ cell/mL).
   NOTE: The total volume in each well will be 2 mL with a ratio of 1:20 MoDCs to lymphocytes per well.
3. Incubate the plate for 48 h at 37 °C and 5% CO₂.
4. Culture enrichment and restimulation with antigen-pulsed MoDCs
   1. After the incubation of the antigen-pulsed MoDC-lymphocyte co-culture, on day 9, supplement each well with 20 ng/mL recombinant IL-2, and continue to incubate for another 120 h (5 days incubation).
   2. On day 14, transfer 1 mL of the co-culture to a sterile 1.5 mL tube, and centrifuge at 500 × g for 7 min.
   3. Discard the supernatant, and resuspend the cell pellet with 1 mL of antigen-pulsed or non-pulsed MoDCs (1 × 10⁵ cell/mL). Mix gently by pipetting, and transfer the cell suspensions back to their designated wells.
      NOTE: At this step, the total volume in each well is 2 mL.
   4. Continue with incubation at 5% CO₂ and 37 ˚C for 48 h. After incubation on day 16, the co-culture is ready for analysis using flow cytometry, qPCR, and ELISA.
      NOTE: For every 2 mL in each well of the MoDC-lymphocyte co-culture, 1 mL of resuspended cells is stained for flow cytometry, 1 mL is used for RNA extraction, and the supernatants from both are used for ELISA.

4. Flow cytometric analysis

NOTE: Stain the cells with appropriate markers/mAb prior to running the samples on a flow cytometer. Refer to the Table of Materials for details on the reagents (staining mAb and isotype controls), kit, instrument, and software used for the flow cytometry analysis.

1. Flow cytometer staining protocol
   NOTE: Perform cell surface staining for the PBMCs and naïve lymphocytes and monocytes using anti-CD14 mAb (step 1.2.6). For the cell surface staining of naïve MoDCs, use anti-CD86-specific, anti-CD40-specific, and anti-MHC II-specific mAb. For the cell surface staining of cells from day 16 co-cultures, use anti-CD4, anti-CD8, and anti-CD25 mAbs; for intracellular staining, use anti-Ki-67 mAb. Furthermore, stain the cells either with negative isotype control mAb or without mAb (FMO: fluorescent minus one). The main difference when staining for surface and intracellular markers is that for cell surface markers, the cells must be stained with specific surface mAb prior to live/dead (L/D) staining or any other processes. However, intracellular staining is performed after L/D staining and requires a fixation plus permeabilization step to allow intracellular penetration.
   1. Transfer 1 mL of cell suspension to a sterile 1.5 mL tube using a pipette, and centrifuge for 10 min at 500 × g.
      ​NOTE: After centrifugation, when processing the cells from the MoDC-lymphocyte co-culture, save the supernatant for ELISA analysis.
   2. Resuspend the pellet in 1 mL of PBS, and transfer it to a 15 mL tube. Collect the residual cells using 1 mL of PBS, and combine this with the resuspended pellet.
   3. Add 10 mL of PBS to the 15 mL tube containing the cells, mix by pipetting, and centrifuge for 7 min at 850 x g.
   4. Discard the supernatant, and repeat step 4.1.3.
   5. Discard the supernatant, and carefully resuspend the cell pellet with the residual suspension (approximately 180 µL) left after decanting the supernatant.
   6. Transfer the cell suspension to a v-bottom 96-well plate, and seal the wells to avoid spillage.
   7. Centrifuge the plate at 1,400 × g for 5 min. After centrifugation, flick the plate over a waste container to empty the wells of liquid, and tap the plate on absorbent paper to ensure the removal of excess liquid.
   8. Add 25 µL of surface staining master mix per well, and gently mix using a pipette, followed by incubation at 4 ˚C for 30 min.
      NOTE: To prepare the surface staining master mix for a single well, add 2.5 µL of surface mAb to 20 µL of FACS buffer.
   9. Add 200 µL of FACS buffer per well, and centrifuge at 1,400 × g for 5 min. After centrifugation, empty the wells of liquid by decanting, and tap the plate on absorbent paper to remove excess liquid.
   10. Add 100 µL of L/D staining solution per well. Mix thoroughly using a pipette, followed by incubation at 4 ˚C for 15 min in the dark.
       NOTE: For a single well, dissolve 0.25 µL of L/D dye in 100 µL of FACS buffer. The L/D dye helps distinguish between live and dead cells.
   11. Add 100 µL of FACS buffer. Cover the wells with the lid, and centrifuge the plate for 1,400 × g for 5 min. Discard the supernatant by decanting, and tap the plate on absorbent paper.
   12. Repeat step 4.1.11.
   13. Add 50 µL of fixation solution per well (provided with the kit), and mix with a pipette before incubating the plate at room temperature in the dark for 10 min.
   14. Add 150 µL of the 1x permeabilization-wash buffer (PW) provided with the kit, and cover the wells with the lid.
       NOTE: To prepare 10 mL of 1x PW buffer, dilute 1 mL of 10x perm buffer in 10 mL of dH₂O.
   15. Centrifuge the plate at 1,400 × g for 5 min at 4 ˚C. Discard the supernatant by decanting, and tapping the plate on absorbent paper.
   16. Add 200 µL of 1x PW, followed by centrifugation at 1,400 × g for 5 min at 4 ˚C. Discard the supernatant by decanting, and tap the plate on absorbent paper.
   17. For intracellular staining, add 25 µL of intracellular staining master mix (anti-human Ki-67 mAb) per well. Mix well, and incubate the plate for 30 min at 4 ˚C or on ice in the dark.
       NOTE: To prepare the intracellular staining master mix for a single well, add 1 µL of Ki-67 mAb to 24 µL of 1x PW. The intracellular staining marker is used only when processing cells from day 16 co-culture. Intracellular staining is not done while processing PBMCs, naïve lymphocytes, monocytes, and naïve MoDCs.
   18. Following incubation, add 200 µL of 1x PW to each well. Mix with a pipette and centrifuge for 1,400 × g for 5 min. Discard the supernatant by decanting, and tap the plate on absorbent paper.
   19. Add 200 µL of FACS buffer, followed by centrifugation at 1,400 × g for 5 min. Discard the supernatant by decanting, and tap the plate on absorbent paper.
   20. Resuspend the cell pellet in 200 µL of FACS buffer, and transfer the contents of each well to separate sterile flow cytometer tubes. Use 300 µL of FACS buffer per well to collect the residual cells. The cells are now ready for flow cytometry analysis.
2. Running the sample on a flow cytometer
   NOTE: The samples were run on a flow cytometer (using 488 nm, 638 nm, and 405 nm lasers) according to the manufacturer's manual. Refer to the manual for details, troubleshooting, and customization of the protocol.
   1. Perform routine cleaning procedures before and after reading the samples.
      NOTE: Do not skip a position while loading the tubes in the instrument.
      1. For the cleaning cycle, use a total of four tubes, with the first tube containing flow cleaning reagent and the remaining three tubes containing dH₂O; each tube will have 3 mL. During the cleaning run, acquire data with a medium flow rate for approximately 1 min by capturing 100,000 events per tube under 330 V for forward scatter (FSC) against 265 V for side scatter (SSC).
         NOTE: No debris or cellular events should be reported during the cleaning; if this happens, perform the cleaning step again. For running the samples, make sure that all the samples are in a homogeneous suspension prior to acquiring the data because this ensures an accurate reading.
   2. To connect and power on the cytometer, click on the Application Button located in the left-hand corner of the title bar. From the application dropdown menu, select the option Cytometry, and click on the option Power On.
      NOTE: From the application dropdown menu, the option Open allows users to browse through the preprogrammed assays, while the option New Protocol allows users to create new protocols.
   3. Initially load the negative control sample in the multi-tube holder. Select New Protocol, and observe the worklist on the left side of the workspace/screen.
   4. On the worklist, define the position of the sample in the multi-tube holder, and give a name to the sample and protocol for identification.
   5. From the Hardware Panel located on the left side of the workspace, adjust and select multiple parameters such as the detectors (FSC, SSC, and 10 fluorescence ranges), the voltages (V), and gains of the photomultipliers, and the area-height-width (AHW) of the signal.
      NOTE: The following settings for the detectors were selected based on the experiment and instrument used: FSC-AHW: AHW, V: 270, G: 5; SSC-AHW: A, V: 350, G: 10; FL1 (CD8/FITC Dextran)-AHW: A, V: 400, G: 1; FL2 (CD25/PE)-AHW: A, V: 500, G: 1; FL6 (CD4/Alexa Fluor 647)-AHW: A, V: 700, G: 1; FL7 (Ki-67/Alexa Fluor 700)-AHW: A, V: 625, G: 1; FL9 (L/D)-AHW: A, V: 425, G: 1.
   6. From the Instrument-Acquisition Control Panel located under the title bar, adjust the options for flow rate (medium), Time (5 min), and Events (see step 5.2.8) desired to be detected. Click on the option Acquire Single to allow the instrument to draw/run the sample and show the real-time preview of the events detected.
   7. From the real-time preview, adjust the threshold of fluorescence (voltage and gain) and cell size to eventually draw gates around the desired cell population while excluding any cellular debris.
      NOTE: FSC helps to distinguish cells on the basis of size, thus allowing users to differentiate between cells of the immune system, such as monocytes and lymphocytes; monocytes are larger and show a higher FSC intensity compared to lymphocytes.
   8. Acquire approximately a total of 5,000 live MoDCs, 50,000 live lymphocytes, and 50,000 live monocyte events.
   9. Once all the parameters are adjusted in reference to the negative control sample, click on Stop, and save the protocol.
   10. Now load all the samples onto the multi-tube loader. Define each sample on the worklist, and apply the parameters adjusted in reference to the negative control to all the samples within the experiment. Click on the option Acquire to allow all the samples in the experiment to be run consecutively.
   11. Once the data from all the samples are acquired, save and transform into readable data using an appropriate data analysis software that allows the generation of sequential bi-parametric and mono-parametric histograms.