Immunofluorescence Microscopy (Pf, fixed)

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Current revision (15:20, 7 October 2009) (view source)
(Reagents)
 
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**1.5 g in 50 mL, make fresh, sterile filter, and keep on ice during use.
**1.5 g in 50 mL, make fresh, sterile filter, and keep on ice during use.
**Alternative blocking solutions: 5% animal serum in PBS, or 5% nonfat dried milk powder in PBS.
**Alternative blocking solutions: 5% animal serum in PBS, or 5% nonfat dried milk powder in PBS.
 +
*Antibodies and Nuclear Dye
*1% (v/v) Polyethyleneimine (PEI) in water
*1% (v/v) Polyethyleneimine (PEI) in water
**PEI is very viscous
**PEI is very viscous
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*Mounting Medium
*Mounting Medium
**Right before using, place 2-3 crystals p-phenylenediamine (~1 mm diameter) in a microcentrifuge tube and add 100 uL water.  Vortex and let sit in dark for 5 minutes.  Centrifuge briefly and use the supernatant as antifade solution.  Make fresh every time.
**Right before using, place 2-3 crystals p-phenylenediamine (~1 mm diameter) in a microcentrifuge tube and add 100 uL water.  Vortex and let sit in dark for 5 minutes.  Centrifuge briefly and use the supernatant as antifade solution.  Make fresh every time.
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===Equipment===
===Equipment===

Current revision

Contents

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Abstract

Use antibodies or sera to localize proteins to the surface of fixed, permeabilized Plasmodium falciparum infected red blood cells.

Reagents

  • 1% gelatin (w/v) in RPMI
  • RPMI Media
    • Store at 4C, warm up to 37 C right before use.
  • PBS
    • Phosphate buffered saline, keep on ice.
  • Fixative Solution
    • 4% formaldehyde + 0.0075% gluteraldehyde in PBS
    • Make fresh for every use. Thaw a fresh frozen aliquot of 25% gluteraldehyde every time you make this.
  • Permeabilization Solution
    • PBS + 0.1% Triton X-100
  • Blocking Buffer
    • 3% Bovine Serum Albumin (BSA) (w/v) in PBS
    • 1.5 g in 50 mL, make fresh, sterile filter, and keep on ice during use.
    • Alternative blocking solutions: 5% animal serum in PBS, or 5% nonfat dried milk powder in PBS.
  • Antibodies and Nuclear Dye
  • 1% (v/v) Polyethyleneimine (PEI) in water
    • PEI is very viscous
    • Sigma Cat. No. 408727-100mL
  • Glass Coverslip
    • No. 1.5 (0.17 mm, 0.16 - 0.19 mm) thickness is best for DeltaVision Deconvolution microscope
  • Microscope Slide
  • Nail Polish
    • To seal coverslips on microscope slide
  • Mounting Medium
    • Right before using, place 2-3 crystals p-phenylenediamine (~1 mm diameter) in a microcentrifuge tube and add 100 uL water. Vortex and let sit in dark for 5 minutes. Centrifuge briefly and use the supernatant as antifade solution. Make fresh every time.

Equipment

  • Heat block set to 37 C
  • Refrigerated microcentrifuge, or a microcentrifuge placed in a cold room (4C)
  • Bunsen burner

Notes

  1. In this protocol, examples assume 12 mL (1 plate) of culture that is gelatin purified, which will be suspended in a 1 mL volume after purification.
  2. All centrifuge steps are at 2000 rpm in the microcentrifuge for 2 minutes.
  3. NEVER vortex to resuspend cells. If need be, gently flick the tube or pipet slowly up and down with a 1 mL pipet tip.
  4. Keep all solutions with fluorescent dye in the dark whenever possible!

Procedure

  1. Gelatin purify 12 mL of parasite culture to enrich for troph and schizont stage parasites
    1. Thaw an aliquot of frozen 1% gelatin at 37C. Use two tubes for 12 mL of culture.
    2. Spin culture at 1400 rpm (394 g) for 5 minutes with a brake of 1 (low) and discard supernatant.
    3. Add a volume of RPMI that is equal to the volume of the pellet. (12 mL culture at 4% hematocrit should be 480 uL pellet)
    4. Add four times the org. pellet volume of 1% gelatin and aliquot 1 mL volumes to 1.5 mL eppendorf tubes.
      1. For example, if pellet is 480 uL, add 480 uL RPMI and mix. Aliquot 240 uL of this mixture into each of four tubes. Add 960 uL of 1% gelatin to each of the tubes.
    5. Incubate at 37C for 10 minutes (Use different times for different applications, e.g. 20 minutes to get really pure trophs). Uninfected blood and ring stage infected red blood cells (iRBCs) will sediment in the pellet and mature stage iRBCs remain in the suspension.
    6. Transfer upper layer to another tube.
    7. Wash the pellet and upper suspension three times with PBS.
      1. Washing can take place in multiple tubes, but in the end, the 12 mL of culture after gelatin purification should yield roughly 40 uL of pellet. Resuspend this in a final volume of 1 mL to roughly recreate 4% hematocrit.
  2. Smear the iRBCs from the pellet and upper suspension to determine percentage of parasites.
  3. Wash cells: Gently centrifuge cells and exchange media with room-temp PBS.
  4. Fix cells: Centrifuge cells and exchange media with fixative solution. Incubate for 30 min @ RT
  5. Wash cells: To remove fixative, centrifuge and wash cells in PBS twice.
  6. Permeabilise cells: To gently permeabilize cells, incubate in 0.1% triton/PBS for 10 min. Check for permeabilization – supernatant should be pink from released hemoglobin.
  7. Wash cells: Centrifuge and wash cells in PBS twice to remove detergent.
  8. Blocking step: Resuspend cells in blocking buffer for a few hours at RT or overnight at 4C on a rotator or rocker.
    1. You can keep the cells at 4C for up to a week.
  9. Primary Antibody Step: Resuspend cells in antibody diluted 1:50 to 1:100 in blocking buffer for 1 hour at RT on a rotating or rocking platform.
    1. If you started with 12 mL culture, you don’t need to use all of it for one microscope slide. Use 125 uL suspension (5 uL pellet) per slide.
  10. Wash cells: Wash cells two times in PBS.
  11. Secondary Antibody Step: Resuspend cells in secondary antibody diluted 1:200 and DAPI (10mg/mL) diluted 1:500 in blocking buffer for 30 min at RT on a rotating or rocking platform. If you have a tertiary antibody, don’t add the DAPI until the tertiary antibody incubation step.
  12. Wash cells: Wash cells two times in PBS.
  13. (Optional) Tertiary Antibody Step: Resuspend cells in tertiary antibody diluted 1:200 and DAPI (10mg/mL) diluted 1:500 in blocking buffer for 30 min at RT on a rotating or rocking platform.
  14. Wash cells: Wash cells two times in PBS.
  15. Resuspend cells in PBS, wrap in foil, and leave to sit while you prepare the slides.
  16. Coat Coverslip with 1% PEI in water: Flame a coverslip over a Bunsen burner. With a glass capillary or pipet tip, drop a spot of PEI solution and spread it over the coverslip with the side of the capillary. Leave to dry.
  17. Add cells: Put a small drop (5-10 uL) of cells on the center of the coverslip and let sit for a few minutes.
  18. Add mounting media: Add 1.5 uL to the middle of the coverslip and mix by pipetting
  19. Add Slide: Flame a slide, let it cool, then place over the coverslip. Keep slides in dark as much as possible to prevent photobleaching of dyes.
  20. Seal slide by painting the edges of the coverslip with nailpolish or Valap (1:1:1 vaseline:lanolin:paraffin wax)
  21. Image!


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