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Fixation and Staining

Millicell cell culture inserts (single-well, 24-well plates and 96-well plates) are designed to support all fixation, staining and immunostaining procedures in a single device. The 24- and 96-well plates are also automation compatible. The variety of membranes offered in the inserts allow the fixed and stained cells to be visualized by stereoscopic microscopy, phase contrast microscopy, or fluorescent methods.

The following are examples of typical fixing and staining protocols. Immunocytochemical staining is a technique employing fluorescently labeled antibody, by which cells can be localized, labeled, and examined via fluorescent microscopy.

The majority of staining procedures employ a fixation step first. Fixation is required to stabilize sub-cellular morphology and prevent degradation of antigens during subsequent staining procedures. Consult the following table for chemical compatibility information with common fixative chemicals.

Chemical Compatibility

	PS	HA	CM	PCF	PET
Acetic Acid	NR NR	NR NR	R R	R NR	NR Acetone R
Acetronite	NR	NR	R	NR	ND
Ammonium Hydroxide	TST R TST	NR NR NR	R R R	TST NR R	ND DMSO ND Alcohols R
Formaldehyde	NR	NR	R	R	R
Glutaraldehyde	RS R	ND R	R R	ND R	ND Glycerol R
Hydrochloric Acid, IN	R RS	R NR	R R	R R	R Methanol ND Sodium
Dodecyl Sulfate	ND	R	ND	TST	ND Sodium
Hydroxide, 3N	R	NR	R	NR	TST
TCA (aqueous solution)	ND	NR	R	TST	NR
Triton® x-100 Surfactant	R	R	R	R	R

General Considerations

• Compatibility key:
• R=recommended
• NR=not recommended
• RS=recommended for short term use
• TST=testing recommended
• ND=no data available
• If the chemicals are compatible with the membrane but not the polystyrene housing, remove the membrane from the housing before adding the chemical.
• Unless otherwise stated, the chemicals listed are at maximum concentration. If the plastic housing and/or membranes are not compatible with the maximum concentration, they might be compatible at lower concentration.

Toluidine Blue Staining

Materials

• Millicell inserts or insert plates
• Milli-Q® water
• Millex-GP filter unit — Millipore cat. no. SLGP 033 RS
• Toluidine blue (Sigma)
• 3% glutaraldehyde in Phosphate Buffered Saline (PBS)
• Triton X-100 (Sigma), 0.5% Method

Method

1. Prepare a 0.3 % solution (gram percent) of toluidine blue in Milli-Q water, stir, and filter through a Millex-GP filter unit.
2. Remove the Millicell insert from the plate and wash gently with PBS to remove growth media.
3. Fix the cells for 15 minutes with 3% glutaraldehyde in PBS.
4. Rinse gently with Milli-Q water. Repeat twice
5. Permeabilize the cells with 0.5% Triton X-100 for 5 minutes.
6. Rinse gently with Milli-Q water. Repeat twice.
7. Apply stain to the apical cell side of membrane for 30–60 seconds.
8. Observe as a wet mount.

Hema-3 Quick Stain

The Hema-3® stain kit is a quick (less than 15 minutes) nuclear staining procedure that can be used with all Millicell cell culture products. The kit is available through Fisher (cat. no. 22–122911).

Wright's Staining

Materials

• Millicell-CM cell culture plate inserts — Millipore cat. nos. PICM 012 50, PICM 030 50
• 3% glutaraldehyde in Phosphate Buffered Saline (PBS)
• Milli-Q water
• 100% methanol
• Wright’s stain

Method

1. Remove the Millicell-CM insert from the plate and wash gently with PBS to remove growth media.
2. Fix the cells for 15 minutes with 3% glutaraldehyde in PBS.
3. Rinse gently with Milli-Q water. Repeat twice
4. Rinse once with methanol and incubate in fresh methanol for 5 minutes.
5. Aspirate the methanol. Add Wright’s stain to cover the inside membrane.
6. Incubate for 30 seconds.
7. Rinse gently with Milli-Q water. Repeat three times.
8. Observe as a wet mount.

Hematoxylin Staining for Millicell-HA

Note: Standard histological hematoxylin and eosin staining techniques can be performed on thin sections.

Materials

• Millicell-HA cell culture inserts — Millipore cat. no. PIHA 012 50, PIHA 030 50
• Millex-GP filter unit — Millipore cat. no. SLGP 033 RS
• 3% glutaraldehyde in PBS, store at 4°C
• Phosphate Buffered Saline (PBS)
• 0.5% Triton X-100 (Sigma) in Milli-Q water
• Hematoxylin solution, HHS-1, 7.5 g/L
• 0.5% Triton X-100 (Sigma) in Milli-Q water
• Dilute ammonium hydroxide (8–10 drops concentrated ammonium hydroxide in 100 mL of water)
• 0.5% hydrochloric acid in 70% ethanol
• Milli-Q water
• Cork borer

Method

1. Filter hematoxylin solution through a Millex-GP unit and cover the cell layer.
2. Incubate for 15 minutes at room temperature.
3. Rinse the Millicell-HA insert with Milli-Q water to remove stain. Repeat twice.
4. Destain by adding 0.5% hydrochloric acid in 70% ethanol for 2–3 minutes.
5. Rinse with Milli-Q water. Repeat twice.
6. Add dilute ammonium hydroxide into the Millicell-HA insert to cover the membrane. Incubate for 3 minutes or until a uniform blue color is observed on the membrane.
7. Rinse with Milli-Q water. Repeat twice.
8. Using a cork borer remove membrane. Mount membrane on a slide in a commercial mounting medium. Make sure the cell layer is facing the microscope objective.

Hemtoxylin Staining for Millicell-CM

Materials

• Millicell-CM Culture Plate Inserts — Millipore cat. nos. PICM 012 50, PICM 030 50
• Millex-GP Filter Unit — Millipore cat. no. SLGP 033 RS
• Milli-Q water
• Sterile Phosphate Buffered Saline (PBS)
• 3% glutaraldehyde in PBS, store at 4°C
• Methanol
• 0.5% Triton X-100 in Milli-Q water
• Hematoxylin solution (Gill No. 1), filter sterilized with a Millex-GP unit (Millipore cat. no. SLGP 033 RS)
• 0.5% hydrochloric acid in 70% ethanol
• Dilute ammonium hydroxide (8–10 drops concentrated ammonium hydroxide in 100 mL of water)
• Mounting media

Method

1. Remove the Millicell-CM insert from the plate and wash gently with PBS to remove growth media.
2. Add 3% glutaraldehyde (at 4°C) in PBS (pH 7.3) to the inside and outside (cell culture plate well) of the Millicell-CM device. Leave for 15 minutes.
3. Carefully remove glutaraldehyde. Add methanol to the inside and outside (cell culture plate well) of the Millicell-CM unit. Leave for 10 minutes.
4. Carefully remove methanol. Add 0.5% Triton X-100 to the inside and outside (cell culture plate well) of the Millicell-CM unit. Leave for 5 minutes.
5. Carefully remove Triton X-100. Add hematoxylin solution (Gill No. 1, Sigma Chemical, filtered with a Millex unit). Leave for 15 minutes.
6. Wash the Millicell-CM insert gently with Milli-Q water. Repeat twice.
7. Add 0.5% hydrochloric acid in 70% ethanol for 45 seconds to remove excess stain.
8. Rinse with Milli-Q water. Repeat twice.
9. Add diluted ammonium hydroxide and leave for approximately 45 seconds.
10. Rinse with Milli-Q water. Repeat twice.
11. Store wet or mount for microscopy.

General Immunofluorescent Protocol for Millicell Products

Materials and Reagents

• Millicell Cell Culture Inserts and Insert Plates
• Sterile Phosphate Buffer Saline (PBS)
• Methanol, 100%
• Glycerol
• FITC-conjugated antibody
• 1% BSA in PBS
• Glass slides

Method

1. Aspirate cell culture media and wash Millicell inserts or plates gently on both sides with PBS. Incubate for 5 minutes and repeat 2–3X. Consult the recommended working volumes table for appropriate volumes.
2. Add fixative solution (e.g. methanol) for 5–10 minutes. It is not required to treat the underside of the membrane with fixative. Incubate according to protocol instructions.
3. After treatment, aspirate fixative and fill filter wells with washing buffer. Repeat 2–3X in order to fully remove the fixative solution from both sides of the filter membrane. Do not allow cells to dry.
4. Dilute primary antibody according to vendor recommendations. In order to obtain best results, it is recommended that optimal working dilutions be determined by the user. If permeabilization is required (such as for cytoplasmic or nuclear antigens), saponin can be added to the solution at a concentration of 0.1%
5. Add antibody solution to each well then incubate at recommended temperature (typically room temperature or 4°C) with mild shaking or rocking to assure that solution wets out entire filter surface. If antibody is fluorescently labeled (direct labeling), cover plate with foil to protect from light.
6. Aspirate antibody solution and wash both sides of membrane as indicated in Step 1 to remove all unbound antibody.
7. If performing indirect labeling with a secondary antibody, repeat steps 4 through 6 with the secondary antibody. For visualization using fluorescent antibodies, continue to Visualization and Microscopy procedures. For enzyme linked assays (HRP, etc.), follow vendor procedures for developing.

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