Staining of Mitochondria, Nucleus, and Cell Membrane for Fluorescent Microscopy (in vitro adherent cells)

Updated 9/22/2023 11:45 AM

by RIVERA-OMICS (created by Os.)

Introduction

This lab protocol outlines the steps for simultaneous staining of mitochondria, nucleus, and cell membrane in in vitro adherent cells using specific fluorescent dyes. The stained cells will be visualized under a fluorescent microscope, allowing the observation of multiple cellular structures in a single sample.

⚠ Safety Precautions:

Handle paraformaldehyde and methanol with caution, as they are hazardous. Wear appropriate personal protective equipment (PPE) such as lab coats, gloves, and safety goggles. Dispose of all chemical waste according to the established laboratory protocols. Always follow the manufacturer's instructions and ensure proper disposal of chemicals and materials used in this protocol. Adjust incubation times, staining concentrations, and other parameters based on the specific reagents and cell types used.

Author: Osvaldo D. Rivera

References:

Materials

In vitro adherent cells grown on glass coverslips

Phosphate-buffered saline (PBS), pH 7.4

Methanol

4% paraformaldehyde solution

Permeabilization buffer

1% Triton X-100 in PBS

Working Staining Solution (Recipes here)

Mitochondrial stain (Rhodamine-123)
Nucleus stain (Hoechst 33342)
Cell membrane stain (FM 4-64)
Diluted in PBS

Fluorescent mounting medium

Microscope slides

Cover slips

Pipettes and tips

Petri dishes or multi-well plates

Procedure

– DAY 1 –

Prepare Coverslips:

Wash thoroughly a glass petri dish and forceps with soap.

Wash out the soap from the plate and forceps with tap water.

Wash out the tap watter from the plate and forceps with dH₂O water.

Dry thoroughly with a paper towel.

Place a filter paper inside the glass petri dish; enough to cover the inside surface.

Wash coverslips 3 times with dH₂O, dry them with a kimwipe.

Place up to 4 coverslips on the filter paper inside the cleaned glass petri dish.

Repeat the process per set of 4 coverslips.

Sterilize prepared dishes and forceps in the autoclave with a heat-dry cycle.

Heat cycle: 15 minutes at 121°C ~ 15 psi.

Dry cycle: 60°C for 30 minutes

00:45:00

Let the cover slips finish cooling down at room temperature.

Aseptically transfer each coverslip with sterile forceps into individual wells of the a 12-well plate.

Grow adherent cells on glass coverslips:

Harvest cell culture to create cell suspension.

Aspirate medium > wash with PBS > Trypsinization > Add cell culture medium > Pipette up and down

Transfer 200µL of cell suspension solution into each well. Top off the well with 1.3mL of culture medium.

Gently move plate to distribute cells evenly.

PAUSEGrow cells until the wells reach ~80%-90% confluence.

Follow lab guidelines for leftover cell suspension disposal.

– DAY 2 –

Fixation:

Carefully and gently wash the cells with 1mL ice-cold PBS to remove any traces of growth medium.

Pipette the PBS onto the side of the well to prevent disturbing the cells.

Prepare a 4% paraformaldehyde solution in PBS. Keep cold.

Must be done in a chemical hood.

Gently aspirate the PBS from the wells and fix the cells to the coverslips by gently adding 1mL of ice-cold methanol to the side of the well. Incubate for 10 min @ -20°C.

00:10:00

Continue fixing cells by adding 500µL of paraformaldehyde solution.

Incubate for 10 min @ room temp. Must be done in a chemical hood.

00:10:00

Wash the cells with 1mL of ice-cold PBS (2 times) to remove residual fixative. Gently add PBS, tilt the plate back and forth, and carefully aspirate off the PBS via the side of the wells.

Must be done in a chemical hood.

Permeabilization:

Prepare the permeabilization buffer (e.g., 1% Triton X-100 in PBS).

Gently 500µL of permeabilization buffer onto the side of the plate until you cover the cells.

Incubate for 7 min at room temp.

00:07:00

Wash the cells with 1mL ice-cold PBS (2 times) to remove the permeabilization buffer. Gently add PBS, tilt the plate back and forth, and carefully aspirate off the PBS via the side of the wells.

Staining:

Prepare a working solution containing all three fluorescent dyes by diluting each dye in PBS.

Note: Recipe for dye working solutions can be found here.

Add the working solution to the cells simultaneously. Incubate for 30 min @ 37ºC

00:30:00

Incubate the cells with the staining solution for the recommended time and concentration specified by the manufacturer. This step may vary depending on the dyes used.

Wash the cells with 1mL ice-cold PBS (2 times)to remove excess dye and unbound staining solution. Gently add PBS, tilt the plate back and forth, and carefully aspirate off the PBS via the side of the wells.

Mounting:

Apply 25 μL of Vectashield Antifade Mounting Medium over a clean microscope slide and place the coverslip over it. Try to avoid the formation of air bubbles.

Fix the coverslip on the slide by applying permount (inside the chemical hood) around the edges of the coverslip.

Store the slides inside a dark box at 4°C until they are to be evaluated under the fluorescent microscope.

Microscopy:

Place a prepared slide on the stage of the fluorescent microscope.

Use appropriate excitation wavelengths and emission filters for each fluorescent dye to visualize the stained structures (e.g., green for mitochondria, blue for nucleus, and red for cell membrane).

Manual for using our in-house microscope can be found here.

Imaging and Analysis:

Capture images of the stained cells using appropriate microscope settings.

Analyze the images using image processing software to study the localization and distribution of mitochondria, nucleus, and cell membrane within the cells.