Phalloidin staining protocol

Detailled procedure for staining with phalloidin dye conjugates, including tips for choosing the most suitable phalloidin conjugate.

Reviewed November 9, 2021

Introduction

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.

Contents

• Reagents
• Choosing the most suitable phalloidin conjugate
• Preparing culture of adherent cells
• Stain cultured cells with phalloidin conjugates

Reagents

• Phalloidin conjugate – prepare following manufacturers guidelines
  • Tip: We recommend using 1% BSA solution to minimize the amount of phalloidin that binds to the tube.
  • Tip: The optimal concentration of phalloidin conjugate and incubation time will vary depending on the particular cell type, fixation/sample prep conditions and/or permeability of the cells/ tissues to the probe.
• PBS
• Methanol-free formaldehyde
• Optional: Triton X-100
• Mounting media – we recommend Fluoroshield to reduce bleaching. Alternatively, use PBS-buffered 50% glycerol with an anti-bleaching agent: p-phenylenediamine or N-propyl gallate
• Optional: BSA, ethanolamine, glycine, DNA staining dye, lysopalmitoylphosphatidylcholine  

Method

1. Choosing the most suitable phalloidin conjugate

2. Preparing culture of 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: 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.

Alternative step: Preparing culture of cells in suspension

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. 

3. Stain cultured cells with phalloidin conjugates

Tip: 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.

• Tip: Quench excess formaldehyde with 10 mM ethanolamine in PBS (or 0.1 M glycine in PBS) for 5 min.
• Tip: Add 0.1% Triton X-100 in PBS into the fixed cells for 3–5 minutes to increase permeability. Then wash cells 2–3 times in PBS.
• Tip: If cells do not appear healthy, add serum (2–10% range) to stain and wash solutions.

3.3 Add phalloidin-conjugate working solution. Incubate at room temperature for 20–90 minutes.

• Tip: add DNA staining dye at this point. 
• Tip: for vertebrate cells, it may be possible to add phalloidin-conjugate to the final PBS wash and mount it in that medium.

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).

Alternative step: staining with other sample types

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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