Binding of purified phage Mu Ner protein to a series of DNA fragments was investigated in order to determine the length requirements for tight specific binding. Gel retardation experiments with wild-type 307 base pair (bp) Mu DNA and shorter, synthetic oligonucleotides were performed, and apparent dissociation constants (KappD) were determined from the half-saturation point. While Ner formed four complexes with the 307 bp DNA fragment, only one complex was observed with the shorter DNAs. The 50 and 30 bp fragments had KappD values of 5 and 20 nM, respectively. Ner binding was progressively weaker with decreasing size of the DNA fragments, with no binding observed for 12mers. The shortest DNA fragments which bound well were two 18 bp fragments for which KappD values were in the range of 50-100 nM. The stoichiometry of Ner complexes with the 30 and 18 bp fragments was determined using a modified Ferguson method. Ner was found to form a tetramer on the 30 bp DNA and a dimer on the 18 bp DNA, which makes the latter a good candidate for the study of a Ner-DNA complex by NMR. In order to clarify which DNA regions were important for Ner-DNA binding, hydroxyl radical footprinting was performed for a range of Ner concentrations from 30 to 500 nM. The footprint revealed that Ner contacts the DNA backbone every 12-13 bp, on both strands of the DNA. The order in which protected regions appeared with increasing protein concentration indicated that two Ner monomers bound to DNA simultaneously. A model of Ner binding to DNA is proposed on the basis of these results.