ABSTRACT
Flow cytometric detection of human immunodeficiency virus (HIV)-infected lymphoid cells at low frequencies is described. Infected cells from human T lymphoid cell lines H9 and A3.01 were detected at frequencies as low as 10(-4) following indirect immunofluorescence labeling. For labeling, cells were treated with an HIV-inactivating, permeabilizing fixative followed by binding of a monoclonal antibody specific for the HIV major core protein p24, and then by binding of fluorescein isothiocyanate-conjugated F(ab')2 fragments of goat anti-mouse immunoglobulin antibody. We compared two fixation procedures, one using a mixture of methanol and acetone, the other a three-step fixation using methanol, paraformaldehyde and Triton X-100. The latter fixation protocol was found to be superior in its ability to resolve mixtures of infected and uninfected cells. The method allowed determination of the percentage of the cell population that was infected and the relative amount of p24 antigen per cell. At analysis rates of several thousand cells/s, detection of HIV-infected cells as rare events was possible. Excellent agreement was obtained between flow cytometric evaluation and reverse transcriptase (RT) assay of infected H9 cells cocultured with uninfected H9 cells in various proportions for 7 days. In time course of infection experiments, cultures infected by small numbers of viral particles were positive by flow cytometry up to 3 days earlier than by RT assay.
