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Detailled procedure for staining with phalloidin dye conjugates, including tips for choosing the most suitable phalloidin conjugate.
Reviewed November 9, 2021
Phalloidin is a highly selective bicyclic peptide used for staining actin filaments (also known as F-actin). It binds to all variants of actin filaments in many different species of animals and plants. Typically, phalloidin is used conjugated to a fluorescent dye, such as FITC, Rhodamine, TRITC or similar dyes, such as Alexa Fluor® 488 or iFluor 488.
Phalloidin can be used with sample types such as formaldehyde-fixed and permeabilized tissue sections, cell cultures and cell-free experiments. It can also be used in paraffin-embedded samples that have been de-paraffinized.
Importantly, phalloidin is also pH sensitive: at elevated pH, a key thioether bridge is cleaved, and the phalloidin loses its affinity for actin. Phalloidin staining can be combined with antibody-based staining by adding the phalloidin conjugate during the primary or secondary antibody incubation step.
2.1 Grow cells in a 96 well black wall/clear bottom plate until they reach confluence (70–80%).
2.2 Cells can also be grown directly on coverslips inside a petri dish.
2.3 Aspirate cell culture medium (with care to avoid dislodging cells).
2.4 Wash once in PBS.
Tip: Avoid fixatives containing methanol or acetone: these disrupt the actin structure and prevent phalloidin staining.
Tip: Suspension cells can be attached to poly-D-lysine microplates or coverslips and then stained using the protocol for adherent cells.
Tip: Avoid fixatives containing methanol or acetone: these disrupt the actin structure and prevent phalloidin staining.
2.1 Grow cells until they reach desired confluence (70–80%).
2.2 Centrifuge cells at 1,000 rpm for 5 minutes and aspirate the supernatant, preserving the cell pellet.
2.3 Resuspend the cell pellets gently in pre-warmed (37°C) growth medium and transfer to microplate or coverslips.
2.4 Aspirate cell culture medium carefully to avoid dislodging cells. Wash once in PBS.
Tip: If you need to save time, suspension cells can be attached to poly-D-lysine microplates or coverslips and then stained using the protocol for adherent cells.
2.1 Grow cells in a 96 well black wall/clear bottom plate until they reach confluence (70–80%).
2. 2 Cells can also be grown directly on coverslips inside a petri dish.
2.3 Aspirate cell culture medium (with care to avoid dislodging cells).
2.4 Wash once in PBS.
Tip 1: Avoid fixatives containing methanol or acetone: these disrupt the actin structure and prevent phalloidin staining. (HowToTip)Tip 2: Suspension cells can be attached to poly-D-lysine microplates or coverslips and then stained using the protocol for adherent cells. (HowToTip)
Tip: Avoid fixatives containing methanol or acetone: these disrupt the actin structure and prevent phalloidin staining.
Alternative: Preparing culture of cells in suspension
1 Grow cells until they reach desired confluence (70–80%).
2 Centrifuge cells at 1,000 rpm for 5 minutes and aspirate the supernatant, preserving the cell pellet.
3 Resuspend the cell pellets gently in pre-warmed (37°C) growth medium and transfer to microplate or coverslips.
4 Aspirate cell culture medium carefully to avoid dislodging cells. Wash once in PBS.
Tip: If you need to save time, suspension cells can be attached to poly-D-lysine microplates or coverslips and then stained using the protocol for adherent cells.
2. Preparing culture of adherent cellsTip: Pre-incubating fixed cells with 1% BSA in PBS for 20–30 minutes may improve staining.
Tip: When staining coverslips, keep them in a covered container to minimize evaporation.
3.1 Fix cells in 3–4% formaldehyde in PBS at room temperature for 10–30 minutes.
3.2 Aspirate fixation solution and wash cells 2–3 times in PBS.
3.3 Add phalloidin-conjugate working solution. Incubate at room temperature for 20–90 minutes.
3.4 Rinse cells 2–3 times with PBS, 5 min per wash.
3.5 Add mounting media to preserve fluorescence (and seal to the slide if using coverslips).
3.6 Observe the cells at Ex/Em 493/517 nm.
Tip: A fast one-step approach to phalloidin staining is effective in some circumstances: a 20-minute incubation at 4ºC in 3.7% formaldehyde and 50–100 µg/mL lysopalmitoylphosphatidylcholine with phalloidin conjugate, followed by three washes and mounting.
Typical staining images:
Image 1 (above): Actin filament staining in HeLa cells. Actin filaments (green) were stained with Phalloidin-iFluor 488 reagent (ab176753); tubulin filaments were stained with a mouse anti-tubulin antibody/goat anti-mouse IgG (red). Nuclei were stained with Hoechst 33342.
Image 2 (above): Actin filament staining in HeLa cells. Actin filaments (red) were stained with Phalloidin-iFluor 555 reagent (ab176755). Nuclei were stained with Nuclear Green DCS1 (ab138905).
Image 3 (above): Actin filament staining in HeLa cells. Actin filaments (blue) were stained with Phalloidin-iFluor 350 reagent (ab176751).
We recommend varying the fixation time and formaldehyde concentration across a range to optimize fixation conditions for staining.
For paraffin-embedded sections, we recommend deparaffinization following our protocol. Antigen retrieval is not necessary for phalloidin staining.
Slides that have been fixed in 4% formalin may not preserve cytoskeletal structures effectively.
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