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1. Preparing Human CD4+ Th1 Effector/Memory T Cells
- Apply a tourniquet to the arm of the donor, wipe the vein with alcohol, and insert a needle. Slowly draw 15 ml of blood into a vacutainer using EDTA as an anticoagulant. When the blood has been drawn, untie the tourniquet before removing the needle. Immediately apply pressure to the wound with sterile gauze when the needle is removed.
- Transfer blood to a 50 ml tube. Add RPMI-1640 at RT at a 1:1 dilution (final volume 30 ml). Carefully overlay the diluted blood onto two 50 ml tubes containing 15 ml of pre-filtered lymphocyte isolation medium such as Ficoll-Paque at RT.
- Centrifuge the gradient at RT for 30 min at 1,200 x g in a swinging bucket rotor. While centrifuging prepare T cell medium (500 ml of RPMI-1640, 50 ml FCS, 5 ml penicillin/streptomycin).
- Upon removal of the 50 ml tube from the centrifuge, observe four layers: a pellet of red blood cells in the bottom, the paque, a layer of cells that contains white blood cells (including lymphocytes), and the plasma. Carefully remove the white blood cell layer with a Pasteur pipette and transfer it to a 50 ml Falcon tube.
- Wash the white blood cell layer by adding RT RPMI-1460 (up to 20 ml) and centrifuging at RT for 5 min at 1,200 x g. Resuspend the white blood cells in 1 ml of T cell medium. Add 5 µl of the 1 ml cell suspension to 250 µl T cell medium in a 1.5 ml centrifuge tube and mix gently up and down by pipette.
- Add 25 µl diluted cell suspension to 25 µl 0.4% Trypan blue. Add 10 µl of mixture to each side of a standard hemocytometer.
- Place the hemocytometer on a low-power light microscope. Using a 10X objective count the number of living cells that have excluded the Trypan blue dye and are present in the middle square of both sides of the hemocytometer.
- To calculate the cell concentration, multiply the average of the 2 squares by 100 (dilution factor) and then multiply by 104 to give the number of cells/ml.
- Adjust the final concentration to 0.5 x 106 cells/ml in T cell medium. Add a final concentration of 1 µg/ml each of bacterial superantigens staphylococcal enterotoxin B (SEB) and toxic shock syndrome toxin 1 (TSST) to the cells. Culture for 72 hr (37 °C and 5% CO2) to expand the CD4+ T cell population.
- Pellet T cells (1,200 x g, 5 min) and resuspend at 0.5 x 106 cells/ml in T cell medium with the addition of human IL-15 (20 ng/ml). Transfer lymphocytes to a T150 flask. Continue to expand/split cells in full medium-IL-15 every 24-48 hr as needed (based on media color; i.e., whenever media turns from pink to slightly yellow) thereafter. Maintain the resulting lymphocyte population for up to 15 days.
NOTE: By design, this protocol will activate and expand specifically a subset of CD4+ T cells that are reactive to SEB and TSST and then drive them toward a Th1-like effector/memory phenotype. Other white blood cells fail to survive and grow under these conditions, such that by step 1.10 the cells will be at least 95% CD4+, CD45RO+ T cells, as can readily be assessed by flow cytometry. If desired, further purification can readily be achieved through commercially available antibody/magnetic-bead-based positive or negative selection kits.
2. Starting Primary Human Endothelial Cell Culture
- Coat a T25 flask with fibronectin (FN) 20 µg/ml in PBS in sterile conditions. Leave at RT for 30-60 min. Remove FN and add 5 ml complete medium (Endothelial Basal Medium (EBM-2) medium supplemented with Endothelial Growth Medium (EGM-2) singlequots). Pre-incubate in a 37 °C cell culture incubator for at least 30 min.
- Thaw a vial of frozen human lung or dermal microvascular endothelial cells (HLMVECs or HDMVECs) in a 37 °C water bath with occasional gentle agitation for ~2-3 min. Immediately transfer cells to the T25 flask containing pre-warmed media. Gently swirl and place in an incubator at 37 °C.
- Change the media after ~4-6 hr. Continue to change media approximately every 48 hr (or when media becomes slightly yellow) until the plate reaches ~90-95% confluency.
3. General Splitting and Expansion of Endothelial Cells
- Grow cells to ~90-95 confluency. This may take 2-5 days. For splitting, remove the media and rinse with PBS. Remove PBS and replace with a minimum volume of fresh 1x trypsin (0.5 ml for T25 or 1.5 ml for T75). Gently swirl to cover all surfaces with trypsin. Incubate at 37 °C for ~5 min. Monitor the detachment of the cells from the plate using a low-power light microscope.
- When the majority of cells appear rounded or detached, add 5 volumes (i.e., compared to the trypsin volume added) of pre-warmed complete EGM-2 medium and gently pipette over the surface of the flask to detach all cells.
- Count endothelial cells with a hemocytometer as described in 1.6-1.7. Pellet the cells by centrifugation (5 min, 1,200 x g). Remove the supernatant. Adjust concentration to 0.5 million cells per ml by addition of pre-warmed complete EGM-2 MV media.
- Transfer aliquots of cells to the appropriate FN-coated dishes or flasks for maintenance. Gently swirl and place in the incubator. Change the media within 6-12 hr of plating. Media should be changed approximately every 48 hr thereafter.
4. Endothelial Cell Transfection
NOTE: Primary endothelial cells are refractory to transfection by the most common chemical and electroporation methods. The nuclear transfection-based method described below allows for relatively high transfection efficiency (~50-70%). An effective alternative method is the use of infection by appropriate viral vectors (see comments in Materials Table).
- Prepare T25 or T75 flasks (as needed) of HLMVECs or HDMVECs to a final density of 90-95% confluency. Coat with fibronectin (FN) 20ug/ml in PBS in sterile conditions either microscope culture plates such as Delta-T plates (for step 5) or 12 mm circular glass coverslips placed inside a well of a 24-well cell culture plate (for step 6) with as described above (2.1).
- Add 1 ml of complete EGM-2 culture media to microscope culture plates or 0.5 ml to each 24 well and equilibrate plates in a humidified 37 °C/5% CO2 incubator.
- Harvest and count endothelial cells as in steps 3.1-3.3. Centrifuge the required volume of cells (0.5 million cells per sample) at 1,200 x g for 5 min at RT. Resuspend the cell pellet carefully in 100 µl RT nuclear transfection solution per sample.
- Combine 100 µl of cell suspension with 1-5 µg DNA. Transfer cell/DNA suspension into a certified cuvette; the sample must cover the bottom of the cuvette without air bubbles.
NOTE: Constructs targeting YFP or DsRed to the cell membrane (through the N-terminal 20 amino acids of neuromodulin that contains a signal for posttranslational palmitoylation) were used alone ( membrane-YFP alone or membrane-DsRed alone) or co-transfected with a cytoplasmic volumetric marker (e.g., membrane-YFP and soluble DsRed). Many permutations of fluorescent protein markers can be used.
- Close the cuvette with the cap. Insert the cuvette with cell/DNA suspension into the cuvette holder of the electroporator and apply electroporation program S-005. Take the cuvette out of the holder once the program is finished.
- Add ~500 µl of the pre-equilibrated culture media to the cuvette and gently remove the cell suspension from the cuvette using the plastic transfer pipettes provided in the nuclear transfection kit.
- For experiments using a microscope culture plates partition the cell suspension from one reaction equally between two dishes containing pre-warmed media (Steps 4.2-4.3). For experiments using 24 wells/plates, partition one reaction equally between 3 wells.
- Incubate the cells in a humidified 37 °C/5% CO2 incubator and change media 4-6 hr, and again at 12-16 hr post-transfection.
5. Live Cell Imaging and Analysis
- Preparing Endothelium
- Day 0: Co-transfect primary HLMVECs with membrane-YFP and soluble DsRed via a nucleofection technology as described in step 4 and plate onto live-cell imaging culture plates.
- Day 1: Replace medium with fresh medium containing IFN-γ (100 ng/ml) to induce MHC-II expression. On Day 2. Stimulate transfected cells by the addition of 20 ng/ml TNF-α to the existing media.
- On Day 3, incubate the endothelium with 1 µg/ml each of bacterial superantigens staphylococcal enterotoxin B (SEB) and toxic shock syndrome toxin 1 (TSST) at 37 °C for 30-60 min immediately prior to experiments. Omit this step for '-Ag' control conditions.
- Preparing Lymphocytes
- In parallel with step 5.1.3, prepare Buffer A (phenol red-free HBSS) supplemented with 20 mM HEPES, pH 7.4, and 0.5% v/v human serum albumin pre-warmed to 37 °C. Take a sample of cultured lymphocytes and determine the density by counting with a hemocytometer (Step 1.12-1.17).
- Centrifuge 2 million cells per sample at 1,200 x g for 5 min at RT in a 15 ml conical tube. Aspirate media and gently resuspend the cell pellet in 2 ml of Buffer A such that no cell clusters remain.
- Remove a fresh aliquot of Fura-2 calcium dye and resuspend in DMSO to make a stock concentration of 1 mM.
- Add 2 μl of Fura-2 stock solution to the T cell suspension (2 μM final concentration), cap the tube, and mix immediately by inverting the tube to ensure even dispersion of dye. Incubate at 37 °C for 30 min.
- Centrifuge as in step 5.2.2. Aspirate Buffer-A and gently but thoroughly resuspend lymphocytes in 20-40 μl of fresh Buffer-A.
- Live-cell Imaging Setup and Acquisition
NOTE: A wide variety of systems can be employed for live-cell fluorescence imaging on upright and inverted light microscopes. Basic requirements include a fluorescence light source and filters, a CCD camera, motorized filter switching, and shutters, a heated stage (or microscope-mountable heated chamber), and software for automated image acquisition. For this protocol high numerical aperture, and high magnification (i.e., 40X, 63X) oil immersion lenses are required to achieve the necessary spatial resolution. Special care must be taken in choosing the appropriate fluorescence source and lenses for Fura-2-based calcium imaging as not all are compatible with the requisite 340/380 nm excitation wavelengths. An alternative approach (compatible with standard green and red fluorescence filter sets) can be used with non-ratiometric calcium-sensitive dyes (e.g., Fluo-4, Rhod-3), though these cannot accurately quantify calcium flux and only provide a relative/quantitative readout.
- Turn on the microscope system (PC for operation, microscope, CCD camera, filter wheel, and xenon lamp).
- Open the designated software.
- Set up microscope/software for automated multichannel time-lapse imaging. Include sequential acquisition of a differential interference contrast (DIC), standard green fluorescence, standard red fluorescence, and standard 340 and 380 nm excitation Fura-2 images. Set the interval for acquisition for 10-30 sec and a total duration of ~20-60 min.
- Set exposure times for Fura-2 imaging.
- Add fresh objective oil and mount a microscope dish containing only 0.5 ml of Buffer-A onto the heating stage adaptor and immediately turn on to equilibrate to 37 °C (will take ~2-3 min).
- Add resting Fura2-loaded lymphocytes to the mounted microscope dish chamber using a 20 μl pipette.
- Turn on bright field imaging. Select the light path to the eyepieces. Use the coarse focus knob to bring the objective into contact with the bottom of the microscope dish. Use the eyepiece and the fine focus knob to focus on the T cells settled at the bottom of the dish.
- Use the x-y stage controls to select a field containing at least 10 cells. Avoid over-crowded fields and cell clumps as these will create imaging artifacts.
- Switch from a bright field to a fluorescent light source. Switch from eyepiece imaging to CCD camera. Set acquisition parameters (e.g., exposure time, detector gain, and binning). Using the software acquire resting Fura2-340 and Fura2-380 images (starting with identical exposure time for each, usually in the range of ~200-1,000 msec).
- Use the methods described in Step 5.4.1 to calculate the Fura2-340/Fura2-380 for each lymphocyte. Perform repeated iterations of adjusting the Fura2-340 and Fura2-380 exposure times, acquiring images, and calculating ratios until the average values are close to 1.
- Set exposure times for mem-YFP and DsRed.
- Replace the microscope dish used in step 5.3.3.1 with a microscope dish containing transfected, activated and SAg treated (or untreated; control) HLMVECs or HDMVECs from the cell culture incubator (Steps 4.5.1). Use a disposable transfer pipette to rapidly remove media, rinse one time the addition of ~1 ml of pre-warmed Buffer-A. Aspirate and then add 0.5 ml of Buffer-A.
- Identify fields in which brightly fluorescent positive transfectant endothelial cells are present and appear healthy with well-formed intercellular junctions.
- Adjust acquisition parameters (e.g., exposure time, detector gain, and binning) for mem-YFP and DsRed. Be sure that the mean fluorescence signal intensity in each channel falls between 25% and 75% of the dynamic range of the detector.
- Conduct live-cell imaging experiments.
- Use automated software to begin image acquisition and capture several intervals of images to establish a baseline.
- During acquisition apply ~5 μl of concentrated Fura-2-loaded lymphocytes (from step 5.2) to the center of the microscope dish imaging field by inserting the tip of a small volume (P-5 or P-20) pipette into the media close to the center of the objective and ejecting slowly.
- As lymphocytes settle into the imaging field, make fine adjustments in the focus to ensure that the T cell-endothelial cell interface (immunological synapses) are maintained in the focal plane. With 40 and 63X objectives, ~10-20 cells per field are optimal. If fewer cells are observed in the imaging field repeat step 5.3.4.2.
- After the desired observation interval of the experiment is completed, continue imaging and immediately pipette ionomycin directly into the microscope dish (using the technique as in 5.3.4.2) to a final concentration of 2 μM to induce maximal calcium flux/Fura-2 signaling signal (i.e., a means of calibration, See analysis 5.4.).