The aim of this work was to study the transcriptional regulation mechanism of ter operon by OxyR in Yersinia pestis. Total RNAs were extracted from the wild-type (WT) strain and the oxyR null mutant (Delta oxyR) strain. Primer extension assay was employed to detect the promoter activity (the amount of primer extension product) of terZ in WT and Delta oxyR. terZ promoter-proximal region was cloned into the pRW50 plasmid containing a promoterless lacZ gene. The recombinant LacZ reporter plasmid was transformed into WT and Delta oxyR, respectively, to measure the promoter activity (the beta-galactosidase activity) of terZ in WT and Delta oxyR by using the beta-galactosidase enzyme assay system. The entire promoter-proximal region of the terZ gene was amplified by PCR from Y. pestis strain 201, and the over-expressed His-OxyR was also purified under native conditions with nickel loaded HiTrap Chelating Sepharose columns (Amersham). Electrophoretic mobility shift assay was applied to analyze the DNA-binding activity of His-OxyR to terZ promoter region in vitro. Primer extension assay detected only one transcriptional start site located at 50 bp upstream of terZ, whose transcript was directly activated by OxyR in Y. pestis. The EMSA result shows that His-OxyR has the ability to bind to the upstream DNA region of terZ. The transcription of ter operon was found to be directly activated by OxyR in Y. pestis.