Assessing the limitations of paraformaldehyde fixation for accurate cell surface receptor measurement

Quantifying cell surface receptors, distinct from the total cellular receptor pool, which includes both intracellular and membrane-bound compartments, is imperative in cell biology and pharmacology. Cell surface receptors are pivotal for initiating signaling cascades in response to extracellular stimuli, making their accurate quantification critical for understanding initial cellular responses and signal transduction mechanisms.

Whole-cell Enzyme-Linked Immunosorbent Assays (ELISA) are commonly used to assess receptor expression in both permeabilized and non-permeabilized cells. Using angiotensin II type 1 receptor (AT1-R) and sorting nexin 12 (SNX12) as models for cell surface and intracellular proteins, respectively, we evaluated the impact of paraformaldehyde (PFA) fixation on protein quantification.

We found that a fixation step with 4% paraformaldehyde (PFA), widely employed across biological protocols, significantly permeabilizes cellular membranes. This results in the unintended detection of intracellular proteins in non-permeabilized cells, thereby leading to inaccurate measurements of plasma membrane receptor expression. Reducing the PFA concentration to 1% substantially improved the accuracy of cell surface receptor quantification by limiting membrane permeabilization while maintaining total receptor content. This adjustment was particularly important when assessing the surface expression of certain GPCR mutants that are retained intracellularly. However, the effect was protein-dependent, as 1% PFA caused inadequate protein immobilization and partial loss of some cytosolic proteins.

These findings underscore the necessity for tailored fixation protocols, depending on the protein of interest, in order to preserve cell surface integrity and ensure reliable quantification of proteins in diverse biological studies.