CHRIS rOBERTS

Chris finished up his undergraduate degree in the Biology Department at GMU in 2003. He worked in my laboratory during the summer and fall of 2002 after applying for and receiving an Undergraduate Faculty/Student Apprenticeship from the Center for Teaching Excellence.

Chris’ project was to generate and characterize an active site mutant of homoserine transacetylase (HTA), the enzyme that catalyzes the first unique step in methionine biosynthesis in many organisms. The reaction catalyzed by this enzyme is shown in the figure to the right.

Although the reaction is written as occurring in a single step, it can actually be divided into two steps. In the first part of the reaction the enzyme transfers the acetate portion of acetyl-CoA to itself, forming an acetyl-enzyme intermediate. In the second part of the reaction the enzyme transfers the acetate to homoserine and releases the final product, O-acetylhomoserine. Reactions of this type are called ping-pong reactions because the first substrate binds, then the first product is released, then the second substrate binds, and finally the second substrate is released. When diagrammed in a standard enzymologist notation it looks a bit like a ping-pong match:

Homoserine transacetylase has designed a special active site to catalyze this reaction and only this reaction. By comparing the sequence of HTA to other protein sequences we realized that it belongs to a family of proteins that all contain a catalytic triad of serine-histidine-aspartic acid. The serine residue is the nucleophile to which the acetate group is transferred. The histidine and aspartic acid residues deprotonate the serine residue, making it a good nucleophile and allowing the reaction to proceed. Chris’ task was to identify the catalytic histidine residue and prove its involvement in the enzymatic reaction.

The first step was to mutate the histidine residue to glycine, completely removing the functional group of the amino acid. This is illustrated in the figure to the left, where the first tripeptide consists of glycine-histidine-aspartic acid and the second tripeptide has the sequence glycine-glycine-aspartic acid. The removed functional group is shown at the bottom right of the figure. Oxygen is represented by the red atoms, nitrogen by the dark blue atoms, and carbon by the light blue atoms. No hydrogens are shown in this figure. The mutated enzyme is expected to have little or no activity because it will be unable to deprotonate the catalytic serine residue.

Generation of point mutations is simple due to the availability of commercial kits such as the QuikChange® Site-Directed Mutagenesis kit from Stratagene. The process, shown to the right, involves design of primers that contain the mutation of interest, annealing of these primers to a plasmid DNA containing the wild-type enzyme, and PCR-amplification of the entire plasmid. An introduction to PCR can be found at this site, and an animation that nicely illustrates the principle is found at the Dolan DNA Learning Center. Because the original plasmid was purified from a bacterial source that methylates its DNA, in our case E. coli, the parent plasmid will be decorated with methyl groups while the mutant plasmid will have no methyl groups attached to it. We can take advantage of this difference in the two populations of DNA by using an enzyme named DpnI that specifically chews up methylated DNA. If we add this enzyme to our PCR product it will destroy all of the original (wild-type) plasmid and leave only the new mutant plasmid. Sequencing of the mutant plasmid will confirm that the correct mutation has been made. Chris successfully mutated the histidine to glycine using this strategy and purified the mutant protein.

When he first analyzed the mutant for catalytic activity, Chris found no catalytic turnover, suggesting that we had hit an important residue. Chris then attempted to rescue the enzyme by adding back the functional group that had been removed. This is referred to as a chemical rescue experiment and the theory is that the mutated enzyme retains the basic structure at its active site, but there is a “hole” where the amino acid has been removed. If the functional group is added back to the reaction mixture, some of the molecules will find their way to the correct spot in the enzyme and be available to catalyze the reaction. It is unlikely that all of the catalytic activity will be regenerated, but hopefully some will return.

Evidence of chemical rescue is shown in the figure to the left. This figure illustrates the change in absorbance at 232 nm over a 5 minute time period. The thioester bond of acetyl-CoA absorbs strongly at 232 nm, and when it is hydrolyzed as part of the enzymatic reaction there is a decrease in absorbance at this wavelength. When no imidazole is added to the reaction buffer there is no change in absorbance at 232 nm over the time period measured, indicating that the mutant enzyme is inactive. When imidazole is added at a concentration of 25 mM we begin to see a decrease in the absorbance at 232 nm, indicating that catalytic activity is occurring. The activity increases as we add more imidazole, and Chris was able to show a saturation of activity with increasing imidazole concentration.

Chris presented his results at the American Chemical Society National Meeting in New Orleans, LA in March of 2003. He has also presented his work on the George Mason campus. The information learned in this study is important for our long term goal, the design of compounds that inhibit HTA and may be a starting point for the design of novel antibacterial compounds. In order to rationally design inhibitors we will need to know the detailed catalytic mechanism as well as the structure of the enzyme. Completion of this project will bring us one step closer to understanding the mechanism of HTA.