The primary approach for the characterization of the insulin receptor has been through the study of its interaction with 125I-labeled insulin. Recently, we demonstrated that insulin receptors can also be identified by flow cytometry using antibodies to the receptor. In the present study, we characterized the insulin receptor on human lymphoblastoid cells (IM-9) and studied its regulation using insulin and antiinsulin antibodies as a probe for flow cytometry. The mean peak fluorescence of the cells treated with insulin followed by antiinsulin serum was 30-50 U above the control value. There was a close correlation between [125I]insulin binding and peak fluorescence. Fish insulin, which has about 50% the affinity of porcine insulin for the insulin receptor but does not bind to antiinsulin antibodies, did not enhance antiinsulin antibody binding, but competed for the pork insulin-antiinsulin antibody complexes in a dose-dependent manner. Exposure of IM-9 cells to insulin or antireceptor antibodies resulted in reduction in the number of insulin receptors. Cells down-regulated with 10(-6) M insulin or a monoclonal antibody to the insulin receptor had 40% of the [125I]insulin binding of the control cells and 40-50% of the peak fluorescence when insulin-antiinsulin was the probe for the immunofluorescence studies. Cells down-regulated with human autoantibodies to the receptor had 4% [125I]insulin binding and 10% peak fluorescence. In all cases, receptors were lost proportionally from all cells, yielding a single symmetrical fluorescent peak. These date indicate that flow cytometry with insulin-antiinsulin antibody complexes provides a new approach to the measurement of insulin receptors, since it provides direct measurement of the occupied receptor.