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Short answer
Fixing cells means stopping biological activity and locking cell structure in place so it can be stained and imaged. Two chemistries do this. Aldehyde fixatives such as paraformaldehyde cross-link proteins and preserve morphology well. Organic solvents such as cold methanol or acetone precipitate proteins and dehydrate the cell. Which one suits a sample depends on the downstream stain, as the checklist below shows.
Definition and scope
Fixation is the sample-preparation step that preserves cultured cells for microscopy before staining. This page covers fixation of cultured mammalian cells for fluorescence microscopy, the setting the cited sources address. It does not cover tissue fixation for pathology; for that context see the guides on H and E staining and immunohistochemistry. Fixation is one stage of a longer workflow, covered in the guide to immunofluorescence protocols, controls and troubleshooting.
How fixatives work in practice
Application notes from the Duke Light Microscopy Core and an Agilent (formerly BioTek) white paper sort fixatives into two categories, aldehydes and alcohols.
Cross-linking fixatives: paraformaldehyde, formaldehyde, formalin, glutaraldehyde
Aldehydes form intermolecular bridges, normally through free amino groups. The Duke core notes that they cross-link proteins and generally preserve cell morphology well. The Agilent white paper adds that cross-linkers preserve structure better than organic solvents but may reduce the antigenicity of some cell components, and that they require a separate permeabilization step. Duke names 4% formaldehyde for 10 minutes as a good starting point for mammalian cells, and Agilent describes 4% paraformaldehyde in PBS for 10 minutes at room temperature followed by PBS washes. Aldehydes are somewhat slower acting than organic solvents, especially for thick specimens.
Glutaraldehyde preserves cellular structure very well but causes strong autofluorescence, and Agilent notes it is preferred for electron microscopy. For that workflow see transmission electron microscopy sample preparation.
Precipitating fixatives: methanol and acetone
Alcohols and acetone remove lipids and dehydrate cells while precipitating proteins. Duke gives 10 minutes in solvent at -20 °C as a starting point; Agilent describes cold methanol for 10 to 20 minutes and notes that methanol is best for preserving structure while acetone improves permeabilization. Both sources warn that methanol denatures proteins and must not be used when a fluorescent protein is to be imaged: Agilent states that the activity of genetically encoded fluorescent proteins will be abolished. Small molecules are lost during solvent fixation.
Permeabilization after fixing
Permeabilization is generally needed after formaldehyde but not after alcohol or acetone fixation. Duke lists the detergents NP-40, Triton X-100, Tween-20 and saponin. Agilent gives Triton X-100 at 0.1 to 1% and the milder, cholesterol-based saponin, and notes that SDS is not for precipitation-fixed samples. Agilent also notes that unreacted aldehydes can cross-link antibodies to inappropriate structures, which is the reason a blocking step follows.
Which fixative suits which experiment?
The table pairs common downstream needs with what the Duke and Agilent documents listed under Sources say about them.
| If the experiment involves | What the sources say |
|---|---|
| A genetically encoded fluorescent protein | Do not use methanol; it denatures the protein and abolishes its activity. |
| Preserving cell morphology | Aldehydes cross-link proteins and preserve morphology generally well; cross-linkers preserve structure better than organic solvents. |
| An antibody that fails after aldehyde fixation | Cross-linking may reduce antigenicity and different methods can destroy or mask different epitopes, so several methods may need to be tested. |
| Intracellular targets after an aldehyde | A permeabilization step is required; it is generally not needed after alcohol or acetone. |
| Low background fluorescence | Glutaraldehyde causes strong autofluorescence; fixative-induced autofluorescence is a known issue for some samples. |
| Thick specimens | Aldehydes act somewhat more slowly than organic solvents. |
| Buffer choice with an aldehyde | Use an isotonic, near-physiological buffer such as PBS and avoid amine buffers such as Tris, which react with the fixative. |
Formalin, formaldehyde and paraformaldehyde: what the label means
The Agilent white paper clears up a common ordering confusion. Paraformaldehyde is a polymer of formaldehyde supplied as dry powder and must be depolymerized before use. Formalin is a commercial solution; a saturated solution holds 37 to 40% formaldehyde and is sometimes called 100% formalin, so 10% formalin corresponds to 3.7 to 4% formaldehyde. Commercial formalin can contain up to 15% methanol as a stabilizer, and that methanol can permeabilize membranes and interfere with staining of membrane proteins. A label that says formalin therefore does not describe the same reagent as a freshly prepared paraformaldehyde solution.
Three label checks follow from the Duke and Agilent documents when stocking fixation reagents: whether the product is formalin, which can contain a methanol stabilizer, or a different formaldehyde preparation, whether paraformaldehyde has been depolymerized for use, and whether the working buffer is amine-free. Formaldehyde is also a regulated workplace hazard; see the guides on formaldehyde exposure limits and monitoring and formaldehyde disposal.
Common misunderstandings
- Formalin is the same as paraformaldehyde. It is not: formalin is a stabilized commercial solution, paraformaldehyde is a polymer that must be depolymerized.
- Methanol works for any sample. It abolishes the activity of genetically encoded fluorescent proteins and removes small molecules.
- Any buffer will do. Amine buffers such as Tris react with aldehyde fixatives.
- One protocol fits every antibody. Different methods can destroy or mask different epitopes, so several methods may need to be tested.
- Fixed cells can be stained without further steps. After an aldehyde, permeabilization and blocking are usually part of the workflow.
FAQ
What is the difference between formalin and paraformaldehyde?
Formalin is a commercial formaldehyde solution; a saturated solution holds 37 to 40% formaldehyde and can contain up to 15% methanol as a stabilizer. Paraformaldehyde is a polymer supplied as dry powder that must be depolymerized before use (Agilent white paper).
Can I use methanol if my cells express a fluorescent protein?
No. Duke advises never using methanol when you want to image a fluorescent protein, and Agilent states that the activity of genetically encoded fluorescent proteins is abolished by methanol fixation.
Do cells need permeabilizing after fixation?
Generally yes after formaldehyde and generally no after alcohol or acetone fixation, according to the Agilent white paper. Duke lists NP-40, Triton X-100, Tween-20 and saponin as detergents that can be used.
Which buffer should I avoid with aldehyde fixatives?
Amine-containing buffers such as Tris, which react with the fixative. An isotonic, near-physiological buffer such as PBS is the usual choice.
Why is glutaraldehyde not the default for fluorescence work?
It preserves structure very well but causes strong autofluorescence, and Duke notes it is used mainly for electron microscopy.
Related reading
Fixation chemistry also affects sample preparation for super-resolution microscopy.
Sources
- Duke Light Microscopy Core, Introduction to Sample Preparation: Immunofluorescence. Date checked: 2026-10-29.
- P. Held, BioTek (now Agilent), Sample Preparation for Fluorescence Microscopy: An Introduction (2015, research use only). Date checked: 2026-10-29.
