Rapid amplification of transposon ends for the isolation, cloning and sequencing of transposon-disrupted chromosomal genes.

Transposon mutagenesis is a commonly used method for identifying genes of interest in a bacterial genome because the insertion of a transposon into chromosomal or plasmid DNA often results in the disruption of a coding or regulatory region. However, identification and molecular analysis of the disrupted gene can often be difficult and time-consuming. We describe a simple and efficient protocol for the rapid identification and isolation of the transposon insertion site in the bacterial chromosome. This protocol is referred to as rapid amplification of transposon ends (RATE). RATE combines methodologies such as the rapid amplification of cDNA ends (RACE) (1,2,5), solidphase DNA separation technology (3,4) and DNA sequencing. Sequences at either or both ends of the transposon are used as templates in the design of specific oligonucleotide primers for the amplification by polymerase chain reaction (PCR) (9,11,14) of adjacent chromosomal DNA. The strategy used in the amplification and isolation of the adjacent unknown genomic DNA sequences of the transposon-mutated strains is described below and summarized in Figure 1. Construction of the Neisseria meningitidis and Listeria monocytogenes transposon Tn916-mutated strains used in this study is described elsewhere (7,8). In the first step, chromosomal DNA is isolated from the transposon-mutated bacterial strain and 3–5 μg digested with a restriction endonuclease. The restriction enzyme used must cleave at one or both termini of the transposon as near to the chromosomal junction as possible, while leaving the anchor primer sites intact. For our experiments, Sau3AI proved to be the most convenient restriction enzyme. There are seven Sau3AI sites within Tn916, two of which meet the criteria described above (near the transposon/chromosome junction) (Figure 1). Restriction of the bacterial chromosome with Sau3AI (New England Biolabs, Beverly, MA, USA) was performed following the manufacturer’s recommendations. After digestion, the samples were phenol/chloroform-extracted, ethanol-precipitated and vacuum-desiccated using standard methods (15). The DNA pellet was resuspended in 20 μL of TE buffer (0.01 M Tris-HCl, 50 mM EDTA, pH 8.0) and 2 μL of the appropriate linkers (250 μM), 10 U of T4 DNA ligase (New England Biolabs) and 2.5 μL of 10× T4 DNA ligase buffer added and the ligation reaction (final volume of 25 μL) allowed to proceed for at least 3 h at room temperature or at 17°C overnight. The sequences of the linker primers (AUS1 and AUS2) used in this experiment are presented in Table 1. The construction of this linker was performed as described by Utt et al. (19). Unligated linkers were removed using Amicon Microcon 100 Microconcentrators (Millipore, Beverly, MA, USA) and the volume adjusted to 20–25 μL with TE buffer. This step is very important because the presence of unligated linkers might interfere with subsequent steps, such as PCR amplification of the target fragment. Unidirectional amplification of a single-stranded PCR product was performed using 5′ biotin-labeled primers specific for the known sequences of the right or left arm of the transposon, P1 and P2, respectively. Two to five microliters of the ligated mixture were subjected to 15 cycles of PCR (95°C for 1 min, 42°C for 1 min, 72°C for 11⁄2 min in 25-μL volumes) using a thermostable Taq DNA polymerase (Boehringer Mannheim, Indianapolis, IN, USA) following the manufacturer’s recommendations. All amplification reactions were carried out with the Model 480 DNA Thermal Cycler (PE Applied Biosystems, Foster City, CA, USA). This reaction yields single-stranded DNA molecules containing sequences corresponding to the remaining portion of the transposon, the adjacent chromosomal DNA and the ligated linker. Unincorporated biotin-labeled oligonucleotides were removed using Micro-