Strains of Enterobacter cloacae were selected on the basis of resistance to aztreonam, ceftazidime, moxalactam, or imipenem. All strains produced the same E2 beta-lactamase, with an isoelectric point greater than 9.5 and with high hydrolytic activity in the presence of cephaloridine. Resistance to beta-lactams could not be correlated with the amount of beta-lactamase present in the various strains. beta-Lactamase activity was induced strongly by moxalactam and imipenem in the wild-type and moxalactam-resistant strains, with beta-lactamase representing as much as 4% of the total cellular protein after induction (2 X 10(5) molecules per cell). Ceftazidime and aztreonam were poor inducers. None of the antibiotics studied was readily hydrolyzed by the E2 beta-lactamase; aztreonam and moxalactam inhibited the enzyme with apparent Ki values of 1.2 and 100 nM, respectively. Aztreonam, which bound covalently to the E2 beta-lactamase with a half-life of 2.3 h at 25 degrees C, was used to measure penetrability of beta-lactam into the periplasmic space of the resistant E. cloacae strains. In all of the E2-producing organisms studied, a significant permeability barrier existed. A maximum concentration of 0.02 microgram of aztreonam per ml should have saturated the periplasmic beta-lactamase in the highest enzyme producers studied. However, fully active beta-lactamase was observed in the periplasm of cells exposed to aztreonam at concentrations at least 1,000-fold higher than that theoretically necessary to inhibit the total enzyme within the cell. Thus, the major cause for resistance to beta-lactam antibiotics in these E. cloacae strains was lack of penetration across the outer membrane.