Staining Filters

Please see warnings for additional information on caring for reusable instruments.

Staining Polycarbonate Filters

Implements used in the following procedures are available in an accessory pack (stock # P48AP), or they can be purchased separately (see chemotaxis accessories).

  1. Aspirate fluid from the top wells or empty them by shaking the chamber over a sink or container.
  2. Remove the thumbnuts while holding down the top plate, and invert the entire chamber onto a paper towel. Grasp the four corners of the top plate (now on the bottom) and push down evenly so that it stays level as it drops to the table. If the gasket hangs up on the post hardware, carefully push it down evenly onto the plate. Take care not to touch the filter, which should be stuck to the gasket. Immerse the remaining plate (with stud hardware in place) in cool distilled water.
  3. The migrated cells are now facing up on the filter—this side of the filter is henceforth referred to as the cell side. Lift up one end of the filter with forceps and catch 1mm of the edge in the large filter clamp. Lift the filter and quickly attach the small filter clamp to the edge of the free end. Keeping the cell side up, wet the underside (non-migrated cell side) of the filter in a dish containing PBS. Do not let the PBS wash over the cell side of the filter.
  4. Holding the filter by the large clamp, with the small clamp attached to the other end and hanging free, wipe the cells off the non-migrated cell side of the filter by drawing the filter up over the wiper blade. The blade should first contact the filter just below the jaws of the wide clamp. Use only gentle pressure against the blade, and maintain an angle of about 30° from the vertical for the portion of the filter above the wiper. It is important to complete the wiping carefully and quickly so that the cells will not dry on the filter; drying takes place in 10 to 20 seconds, and will prevent complete removal of the non-migrated cells.
  5. Clean the wiper with a cotton swab, again wet the underside of the filter in PBS, and repeat Step 4. Clean the wiper again, wet the filter a third time in PBS, and repeat Step 4.
  6. For granulocytes and monocytes, carefully immerse the filter in methanol, then place the filter cell-side up on a disposable lint-free towel for air-drying. Rinse all chamber components in cool distilled water. For other kinds of cells, consult the literature for staining techniques.
  7. When the filter is dry, clamp the edge of one end with a large filter clamp, weight the other end with a small filter clamp, and stain in Diff-Quik® (available from VWR Scientific Products – www.vwrsp.com) or equivalent dye, according to the manufacturer’s instructions. To avoid contaminating the chamber components with stain, it is convenient to have two sets of filter clamps, one for removing the filter from the gasket, and one for staining.
  8. Place the wet filter cell-side up on a 50 x 75mm microscope slide to dry. When the filter is dry, center it on the slide and place a drop of immersion oil on it. Rub the oil over the filter with a smooth, blunt instrument to remove all bubbles and wrinkles. The filter is now ready for counting.
  9. If you are using a template to help locate cell sites on the filter (stock # P48TM or C48TM) modify these steps following the P48TM or C48TM protocol.

Suggested Reading

Falk, Goodwin, and Leonard. “A 48 Well Micro Chemotaxis Assembly for Rapid and Accurate Measurement of Leukocyte Migration.” 1980, Journal of Immunological Methods, 33, 239-247.

Harvath, Falk, and Leonard. “Rapid Quantification of Neutrophil Chemotaxis: Use of a Polyvinylpyrrolidone-free Polycarbonate Membrane in a Multiwell Assembly.” 1980, Journal of Immunological Methods, 37, 39-45.

Richards and McCullough. “A Modified Microchamber Method for Chemotaxis and Chemokinesis.” 1984, Immunological Communications, 13 (1), 49-62.

Harvath and Leonard. “Two Neutrophil Populations in Human Blood with Different Chemotactic Activities: Separation and Chemoattractant Binding.” 1982, Infection and Immunity, 36 (2), 443-449.

Staining Cellulose Nitrate Filters

We have found that different laboratories prefer different staining techniques for cellulose nitrate filters. Several staining protocols that have been successfully employed are described here.

Clip filters on the edges so that they bow out, allowing exposure on both sides. When going from dish to dish of reagents the slides should be plunged up and down to assure clearing behind the filter. After staining but before adding Permount and the cover slip, the filters are flattened with finger or Pasteur pipette to clear any reagent trapped under the filter and to assure that the filter is flat for easier reading under microscope.

(If you are using a template (stock # C48TM) to help locate cell sites on the filter, follow the C48TM protocol.)

Please note that cellulose nitrate dissolves in solutions of ethanol or methanol above 20%; dissolution time depends on the temperature and concentration of the solution. The following protocols use these reagents for fixing and drying, so some dissolving of the membrane matrix will occur. If you experience an excess of this, we recommend that you shorten the exposure or change reagents. For additional technical information, contact the filter manufacturer, Sartorius.

Sample protocol 1: Courtesy of Bill Cruikshank, Boston University

  1. Fix filters in 95% EtOH (1:1 with H2O) for a minimum of 10-15 minutes, maximum of 48 hours.
  2. Wash in running water bath for 10 minutes.
  3. Stain in modified Harris Hematoxylin for 8-10 minutes. If staining is weak, add a few drops of glacial acetic acid and repeat staining.
  4. Wash in running water bath 10 minutes.
  5. Five dips (approximately 5 seconds total) in 0.5% HCl.
  6. Wash in running water bath for 10 minutes.
  7. This step is for neutrophils only: Counter-stain in chromotrope 2R for 30 seconds.
  8. This step is for neutrophils only: Wash in water bath for 6 minutes.
  9. Fifteen dips (approximately 8-10 seconds) in 95% EtOH.
  10. Fifteen dips (approximately 8-10 seconds) in 95% EtOH in another dish.
  11. In this step, time is crucial. Two minutes in 1-Propanol. (Fisher grade is fine.)
  12. Two minutes in 1-Propanol in another dish.
  13. In this step, time is crucial. Three minutes in 1-Propanol, 1:1 with xylene.
  14. Approximately three minutes in xylene.
  15. Approximately three minutes in xylene in another dish.
  16. Approximately three minutes in xylene in another dish.
  17. Approximately three minutes in xylene in another dish.
  18. Approximately three minutes in xylene in another dish.

Notes: Clip filters on two edges so that they bow out allowing exposure on both sides. When going from dish to dish the slides should be plunged up and down to assure clearing the filter. After staining, but before adding Permount and coverslip, flatten the filters with a finger or pasteur pipette to clear any xylene trapped under filter and to assure that the filter is flat for easier reading under the microscope.

Sample protocol 2: Courtesy of Liana Harvath, NIH, Bethesda, MD

Solutions

  • Mayer Hematoxylin: (or obtain from commercial source) in 500mL of dH2O dissolve (with heat) 50 gms aluminum ammonium sulfate. After cooling, add mixture of chloral hydrate – 50 gm, citrate – 1.0 gm, and sodium isolate – 0.2 gm. Then add to this mixture, 1.0 gm Hematoxylin dissolved in 10mL absolute EtOH.
  • Blueing solution: 1,000 mL dH2O (or 400mL in 400mL dH2O plus 1 mL concentrated. ammonium hydroxide.)
  • Phosphate buffered saline (PBS).
  • 70% EtOH, 95% EtOH, and 99% EtOH.
  • Xylene.
  • Formaldehyde: 37% formaldehyde (full strength), in a 1:1 solution with dH2O.

Staining Sequence

  1. PBS – dip.
  2. Formaldehyde – 2 minutes (may leave overnight at this stage).
  3. H2O – dip.
  4. Mayer Hematoxylin 6-15 minutes.
  5. H2O – dip: repeat total of three times.
  6. Blueing solution – 2 minutes.
  7. H2O – dip.
  8. 70% EtOH – 2 minutes; 95% EtOH – 2 minutes.
  9. 99% EtOH – 2 minutes; twice total
  10. Xylene – 20 minutes or until clear.

Notes: Place stained filters on glass slides, cover with immersion oil and cover with a 35 X 50 mm glass coverslip for counting. If slides are stored for extended periods of time, the filters may become opaque because of drying of the filter. Usually additional application of immersion oil to the filter surface will correct the opacity.

Sample protocol 3: Courtesy of David Hyslop, University of Missouri

  1. Fix overnight in 70% ethanol.
  2. 50% ethanol – 10 dips.
  3. 30% ethanol – 10 dips.
  4. 15% ethanol – 10 dips.
  5. Mayer Hematoxylin – 3 minutes.
  6. H2O rinse – approximately 2 minutes with running tap water.
  7. 15% ethanol – 10 dips.
  8. 20% ethanol – 10 dips.
  9. 50% ethanol – 10 dips.
  10. 0.5% HCl in 70% ethanol – 7-10 dips (watch for color change).
  11. 0.3% NH4OH in 70% ethanol – 7-10 dips (watch for color change, can use up
    to 0.6% NH4OH in 70% ethanol if results do not show adequate bluing).
  12. 70% ethanol wash – 10 dips.
  13. 0.5% Eosin-Y in 90% ethanol, pH 5.4 – 5.6, 1 – 2 dips.
  14. 95% ethanol – 10 dips.
  15. 95% ethanol – 10 dips.
  16. 100% ethanol – 10 dips.
  17. 100% ethanol – 10 dips.
  18. Xylene – 30-60 seconds (watch for initial clearing).
  19. Xylene – 8 minutes for final clearing.
  20. Mount cell side toward slide with Permount and cover with cover glass.

Sample protocol 4: Courtesy of Karen Richards, University of Minnesota

  1. Fix in absolute methanol 2 – 4 seconds.
  2. Gill’s Hematoxylin, #1 – 2 – 10 minutes.
  3. Distilled water rinse
  4. Absolute ethanol – HCl (1000:1 v/v) – 30 seconds.
  5. Distilled water rinse.
  6. Blueing reagent (0.17M MgSO4, 0.02M NaHCO3 in water) – two minutes.
  7. Distilled water rinse.
  8. 70% ethanol – 30 seconds.
  9. 95% ethanol – 30 seconds.
  10. Absolute ethanol, 3 times – 2 minutes each.
  11. Xylene, 2 times – 2 minutes each.

Sample protocol 5: From: Wilkinson, Peter C., Chemotaxis and Inflammation (Churchill Livingstone: Edinburgh, London, Melbourne, NY, 1982), p. 201

  1. Fix in 70% ethanol for 5 minutes.
  2. Distilled H2O – 2 minutes.
  3. Harris Hematoxylin 30-60 seconds.
  4. Distilled H2O – 2 minutes.
  5. Tap H2O – 10 minutes.
  6. 70% ethanol – 2 minutes.
  7. 95% ethanol – 3 minutes.
  8. Solution – 80% ethanol: 20% butanol – 5 minutes.
  9. Xylene – 10 minutes to clear.

Notes: Care should be taken to maintain the ethanol/butanol solution in an anhydrous state; this assures clearing during the xylene step.