Love:μEn hybridoma screening

From OpenWetWare
Jump to: navigation, search

~~craig story 1/9/07

Eli Papa 15:07, 17 April 2008 (EDT)

Summary of protocol:

Day 0

  1. Coat epoxy treated slides with capture antibody or antigen of interest

Day 1

  1. Prepare grids (plasma treat, block)
  2. Count and dilute cells.
  3. Apply cells to grid
  4. Release grids from slides, place slide in blocking/wash solution
    • optional: save grid for slidescanning (CFSE staining or phase)
  5. Probe slide with fluorescent probes
  6. Wash slides
  7. Scan on slidescanner.

Detailed protocol:

  1. Coat glass slides with capture antibody: Coat Super-Epoxy 2 ™ slides (ArrayIt #SME2) with anti-mouse capture antibody. These slides are stored under vacuum. Be sure to apply to correct side of slide. Use goat anti-mouse IgG (a 1:1 mix of Southern Biotech #1030-01 and Zymax gives best results b/c of slightly different isotype reactivity) to a final total conc of 200 µg/ml. Use LifterSlip (Erie Scientific. 601-2-4733) 75 µl per slide. Dilute antibody in Borate-9 buffer (50 mM borate pH 9, 80 mM Trehalose, 50 mM NaCl).
  2. Incubate room temp ~1hr (30 minutes min).
  3. Prepare Grids: Prepare PDMS grids by plasma treating (see detailed SOP #7): Briefly, place grid face up in center of chamber on glass slide. Apply vacuum for 1 minute (make sure valve is closed). Apply high voltage, violet color should be visible, at this point open valve very slightly until pink color is visible. Wait 20 seconds. Turn off voltage, break vacuum, turn off pump, in that order. Place grids in blocking buffer: PBS+ 0.5% BSA (generally we leave Tween-20 out at this step, but 0.05% Tween should not matter). DON’T increase % BSA at this step! It will cause cells to not adhere to grids properly. Good results with overnight at 4 degrees, but 1 hour room temp blocking step is sufficient.
  4. Prepare slides for cell application. Flood slides with Blocking Buffer (PBS+1.5% BSA + 0.05% Tween 20), and incubate for 30 minutes to block any additional epoxide sites on slide and reduce non-specific binding to slide. After this blocking step, briefly rinse first in PBS, then in dH2O and spin dry with MicroArray high-speed slide centrifuge or swinging bucket slide holder. It is best to keep slides humidified until the printing step.
  5. Load cells. Resuspend cells to single cell suspension by pipetting. Count and resuspend cells of interest to 2 x 105 cells/ml density. Use ~1 ml of this solution per grid. Place cells in microwells. (Do all cell manipulation in TC hood to keep sterile throughout). Remove grid from blocking solution and rinse a couple times in PBS to wash out azide, then briefly in DMEM-10, place in TC 10 cm dish. Remove most liquid from grid by gentle suction at the edge. Apply about 0.5 to 1 ml of cells to top of grid, thus forming a “bubble” of liquid. Gently rock back and forth with 10 second pauses for ~1 to 2 min to encourage cells to drop into wells. Monitor on inverted scope. Shorter times will result in fewer cells/well.
  6. Apply grid to slide: Rinse cells off grid at an angle with 5 ml of DMEM-10 media, allowing the media to collect in dish. Suck away extra media and excess media from angled rubber grid surface. Carefully apply rubber to slide surface avoiding trapping excess liquid or air bubbles. Grid should not be “dry” but as much excess liquid as possible should be removed before applying to grid. As a convention, we put the “upper right” of the grid (for example, sector 7 out of 14) at the “upper right” of the slide held horizontally with number at left. It is helpful to take a bite out of the upper right corner before plasma treating grid.
  7. Hold slide in clamp with steady pressure while using three screws to tighten lid. Turn screws approx 1/8 turn past contact while hand-squeezing, just very gentle pressure is all that is desired. The idea is to get even pressure between rubber grid and slide surface. If too much pressure is applied, the lid deforms, and the contact is uneven.
  8. Incubate in 37 deg for 20 min for 50 µm wells, 90 minutes for 100 µm wells. Separate slide and grid. Place grid in media for continued growth of cells, and slide in blocking buffer (see next step). To make a duplicate stamp, remove grid from media and proceed with step 5 above. I have found that it is easiest for multiple stampings to: submerge entire grid, place grid in empty dish, remove excess media, then reapply grid to next slide.
  9. Place slide in Blocking Buffer (PBS + 1.5% BSA, 0.05% Tween 20) Slide can be probed at a later time if desired.
  10. Rinse slide in blocking buffer, remove excess liquid by leaning on toweling. Mix detection proteins and apply to slide under LifterSlip. Example probe dilution is as follows:

*Ova555 at 0.01 mg/ml (1:200 dilution of 2 mg/ml stock) *HA peptide tetramer (1:75 dilution of 1.0 mg/ml stock) *Fluorescent secondary antibodies use at 100 nM (1:1333 dilution of 2 mg/ml stock)

  1. Apply detection reagent for 30 min in dark.
  2. Wash slide 30 min with 3 changes of PBS + 0.05% Tween 20. Keep covered with foil. Be careful not to allow cross-contamination of slides during wash. Keep different staining types separate during washes to remove this possibility.
  3. Rinse first with PBS to dilute Tween, then briefly with water. Spin dry, scan on slide scanner. Genepix software operation see SOP #10, Genepix capture and analysis.