Griffin:Immunofluorescence Cell Staining

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Immunofluorescence Cell Staining

Below is a simple and clear approach to immunofluorescence. There are 3 common fixation methods listed below that are proven and effective for a multitude of adherent and suspension cells. 1% formalin PBS is a good starting fixation method. For a typical slide, it takes ~300 microliters to cover an area equal to 2 coverslips.

I) Culture cells to 50-70% confluency. Some cells (endothelial/epithelial) may require a higher level of confluency. For suspension cells in a regular petri dish or flask, transfer enough cells suspended in medium to each well of a poly-D-lysine-coated microslide. Then Incubate microslide in CO2 incubator from 1 hour to overnight to allow cells to attach to the bottom of the slide. The incubation time varies depending on the suspension cell used. Check for cell attachment using a microscope.

Gently aspirate medium from the wells, wash the attached cells briefly with 1x PBS and proceed with the Cell Staining Procedure of choice (i.e., protocol 6: Immunoperoxidase Cell Staining, or protocol 7: Immunofluorescence Cell Staining)

II) Fix the subconfluent cells appropriately, followed by 2 PBS washes (dipping the slides gently into separate beakers of PBS is one way to wash). Optional step: Antigen retireval (see below).

III) Blocking: Gently pipet 10% BSA, PBS onto the slide and incubate for 30 minutes rt.

IV) Primary Antibody Incubation: 1:50 in 2% serum/PBS for 60 minutes at room temperature followed by 3 separate and gentle PBS washes.

V) Secondary Antibody Incubation: 1:100 in 2% serum/PBS for 60 minutes at room temperature followed by 3 PBS washes and 1 dH2O wash.

At this point prop the slides at an angle to allow them to partially dry. (You will see liquid evaporate slowly off slide when faced up. Wait for all liquid to evaporate and then add the mounting medium) approximately 5 ul drop (no air bubbles) to each area (8 areas for 8 well plates) Gently drop the coverslip onto the slide and align. Allow it to settle. Place mounted slide in light proof box and place in 4 degree overnight priot to performing the visualization.

The mounting agent is two component, one powder and one liquid. When you add the liquid make sure all the powder is dissolved. The powder tends to be in chunks on the bottom of the the jar vial. Vortexing may not loosen up this chunk and into its dissolved state. Also let the soultion set for 5 minutes before pipeting to allow air bubbles to rise out of the glycerol. Molecular Probes Prolong Antifade Kit

Visualize under the appropriate fluorochrome filter.

Common fixatives to consider in protocol optimization

Methanol fixation (for cytoskeletal components)

The methanol fixation is an easy method; however, it frequently solubilizes and removes membrane bound antigens. By a simple precipitation of the protein, methanol only provides low structural preservation.

  • Rinse the cover glass with PBS.
  • Fix cells by incubating the cells in pre-cooled 100% methanol at -20 oC for 10 min.
  • Wash cells with PBS.

Formaldehyde fixation (for membrane associated components)

Higher polymers, which are initially insoluble, are sold as a white powder known as paraformaldehyde.

  • Rinse cells with PBS at room temperature.
  • Fix in 3-4 % paraformaldehyde in PBS for 15 min at room temperature.
  • Wash 3-times 5 min each with PBS containing 100 mM glycine.
  • Permeabilize cells with 0.1% Triton X-100 in PBS for 1 to 4 min.
  • Rinsed with PBS.

Formalin fixation (for membrane associated components)

The liquid known as formalin contains 37-40% of formaldehyde and 60-63% of water (by weight), with most of the formaldehyde existing as low polymers.

  • Rinse cells with PBS at room temperature.
  • Fix in 1 % Formalin in PBS (prepared from 10% buffered formalin PBS) for 10 min at room temperature.
  • Wash 3-times with PBS
  • Permeabilize cells with 0.1% Triton X-100 in PBS for 1 to 4 min.
  • Rinsed with PBS.

DAPI Counterstain

4,6-Diamidino-2-phenylindole, 2HCl for DNA labeling. DAPI is a popular nuclear counterstain for use in multicolor fluorescent techniques. Its blue fluorescence stands out in vivid contrast to green, yellow or red fluorescent probes of other structures. DAPI stains nuclei specifically, with little or no cytoplasmic labeling. The following protocol is used for staining of tissue sections or for staining cultured cells on slides.

  • Working concentration: 0.1 to 1 µg/mL DAPI.
  • Staining Pattern: Nuclei will be stained bright blue.
  • Suggested Use: Counterstain for immunofluorescence when green (FITC) or red (Texas Red) fluorescent marker is used.
  • Excitation/Emission: ~359 nm/~461 nm

DAPI Counterstain Solution

DAPI Stock Solution (5mg/ml or 14.3 mM):

DAPI -------------------------------------------------------- 10 mg

Dimethylformamide (DMF) ------------------------------ 2 ml

Mix to dissolve; it may take some time to completely dissolve. Aliquot and store in –20 ºC.

DAPI Working Solution (100ng/ml or 300 nM in PBS):

DAPI stock solution --------------------------------------- 2 ul

PBS -------------------------------------------------------- 100 ml

Store this solution at 4ºC in brown bottle or wrapped with aluminum foil to protect from light. Incubate sections in dark for 30 minutes at room temperature.

Following reconstitution, aliquot and freeze (-20C) for long term storage or refrigerate (+4C) for short term storage. Stable for 1 years as supplied. Stock solutions stable for several weeks at +4C or for several months at -20C

Antigen retrieval for cryostat tissue sections or cultured cells

This is a simple method for antigen retrieval on aldehyde-fixed cryostat tissue sections or cultured cells. In many case, a brief 5 minutes pretreatment with 1% sodium dodecyl sulfate (SDS) produced a dramatic increase in staining intensity by immunohistochemistry and immunofluorescence.

Solutions and Reagents

1% Sodium Dodecyl Sulfate (SDS) in PBS:

  • SDS ----------------------------- 1 g
  • 0.01M PBS (pH 7.4) ---------------- 100 ml

Mix to dissolve.

Procedure

  • Rinse sections three times for 5 min each in PBS.
  • Cover sections with 1% SDS solution and incubate for 5 minutes at room temperature.
  • Rinse sections three times for 5 min each in PBS. It is important to wash sections well, otherwise residual SDS will denature the antibodies subsequently applied to sections.
  • Incubate sections in serum blocking solution.
  • Incubate in the primary antibody and complete immunohistochemical staining steps as desired.
  1. Brown D, Lydon J, McLaughlin M, Stuart-Tilley A, Tyszkowski R, and Alper S. . pmid:9072183. PubMed HubMed [Paper1]
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