A monoclonal antibody, raised initially against pure bovine low-density lipoprotein (LDL) receptor, was found to bind saturably to LDL receptors on the surface of cultured human fibroblasts. It was apparently internalized by the cells but rapidly returned to the surface and was not significantly degraded. LDL did not affect the binding of antibody. However, the antibody reduced both the apparent affinity of the receptor for LDL and the amount of LDL bound. In the presence of antibody, the concentration of LDL required for one-half maximum binding increased from 14.5 nM to 22.3 nM at 37 degrees C and from 2.8 nM to 11.5 nM at 4 degrees C. Maximum heparin-releasable binding of LDL was reduced by 56% at 37 degrees C and by 36% at 4 degrees C. Fab fragments halved the affinity at both temperatures without affecting the maximum amount of LDL bound. At 4 degrees C, maximum heparin-releasable LDL binding was the same as that of antibody and of Fab fragments. At 37 degrees C, heparin-releasable binding of LDL and binding of Fab fragments were double that at 4 degrees C, whereas binding of antibody remained unchanged. The results suggest that the antibody bound to a site on the receptor remote from the LDL binding region and that intact antibody recycled with the receptor. It halved LDL binding and degradation at 37 degrees C and reduced binding at 4 degrees C, probably through a different mechanism. The possibility that both cooling and antibody led to receptor aggregation is discussed.