Matthew J. Hinton and Dr. Roger G. Harrison, Chemistry and Biochemistry
An exciting area of research is based on the idea that small molecules can be used to perform the same chemical reactions as enzymes. In living organisms, enzymes play important roles in facilitating the vast majority of biochemical reactions necessary for life. One particular class of these biomolecules, non-heme iron containing enzymes, are of particular interest because they have structural characteristics that may allow their chemistry to be modeled by small molecules. In the active site of these enzymes there is often one or more iron atoms bound by two or three histidine side chains. I was interested in determining whether a small, five amino acid peptide with two histidine residues would be able to act in a similar way to these enzymes.
One challenge to developing a small molecular model from a five residue peptide is conformational stability. Large proteins benefit from having numerous amino acid residues that interact to fold them into relatively stable three dimensional shapes, or conformations. In contrast, a five residue polypeptide chain is far more likely to exist in numerous random conformations in solution. Yet effective modeling of enzyme active sites requires that the small peptide model have a consistent structure. For this purpose I employed the use of an unnatural amino acid, -aminoisobutyric acid (Aib, U). Aib is similar to alanine except that it has a methyl group replacing the chiral proton. Previous researchers have found that when incorporated into peptides, Aib has a tendency to cause them to fold into helical conformations.1,2 Even some small, four residue peptides can fold into 3,10 helices when Aib is present. My research project involved synthesizing the small pentapeptide HULHV and characterizing it to determine if it adopts a stable helical conformation in solution. If found to adopt a consistent structure, the peptide may then be studied as a potential model for non-heme iron proteins.
To synthesize the peptide, I used the technique of solid phase peptide synthesis.3 This process began by chemically attaching the first amino acid residue in the chain (valine) to an insoluble resin bead. This support was then conveniently used to keep track of the growing chain as I added each subsequent residue until the chain was finished. Because amino acids do not normally react with each other at an appreciable rate in solution, it was necessary to activate each amino acid at every step in the chain building process. I wanted the carboxylate group of the incoming amino acid to react with the amino group of the amino acid at the end of the chain. To achieve this purpose, I used an activated ester compound known as PyBOP which greatly facilitated the linking of the amino acid residues in the proper orientation.4 Furthermore, to prevent unwanted reactions between two activated amino acids, I selected reagents that had Fmoc protecting groups already in place on the amino end of the residues. A deprotecting agent (piperidine) was used to remove the Fmoc protecting group after each addition cycle to prepare the terminal amino acid for the next round of attachment. Chemical cleavage using tri-fluoroacetic acid was used at the end of synthesis to remove the peptide from the resin support.3 I synthesized two forms of the peptide: the peptide acid, which has an amino group at one terminus and a carboxylate group at the other, and a peptide amide which replaces the carboxylate group with an amide group.
Once the peptide had been synthesized and cleaved from its support, I used mass spectrometry to determine if I had actually obtained my desired product and judge the relative purity of the peptide. The peptide acid (molecular weight of 589 daltons) gave a strong peak at 590, which was the predicted M+l peak. The spectrum showed high purity for the peptide which I roughly estimated at about 90%. The peptide amide, which has an amide group at the C terminus also showed high yield with a strong M+l peak at 589, which was expected due to its molecular weight of 588 daltons. Analysis of the nuclear magnetic resonance spectrum of the peptide also showed that the peptide had been produced in high purity.
The primary tool that I used to characterize the peptide was nuclear magnetic resonance spectroscopy (NMR). I employed a particular technique called water-labile deuterium exchange to understand how the peptide was folding in solution. Intramolecularly bound hydrogen atoms in the molecule will be less likely to exchange with deuterium solvent than will unbound hydrogen atoms. The fact that deuterium is invisible on the NMR spectrum makes it possible to add small quantities of deuterium oxide (D20) and observe the corresponding decrease in peak integration of the more labile protons on NMR spectrum. At the same time, the less labile protons (such as those that are intramolecularly hydrogen bound) will show less of a decrease on the same spectrum. Thus, successive spectra taken after sequential additions of D20 will show the relative lability of the various amide protons in the molecule. Once these are identified, a hydrogen bonding pattern can be deduced which will indicate the presence or absence of helix formation.
The water labile study was moderately successful in determining the presence of helix formation. The relative rate at which the peaks diminished on the NMR spectra seemed to indicate a hydrogen bonding pattern consistent with a 310 helix. I attempted to verify these results using several other methods. For example, X-ray crystallography is an excellent way to identify three dimensional structure of polypeptides, but I experienced difficulty in growing uniform crystals. Circular dichroism also showed promise of producing reliable results, however I was not able to obtain adequate data by publication time. Further work using two dimensional NMR and the aforementioned techniques should be able to confirm the 310 helical nature of the peptide.
This research project gave me excellent experience with the use of modem analytical techniques such as NMR, high performance liquid chromatography (HPLC), circular dichroism (CD), and X-ray crystallography. It also gave me an appreciation of the challenges that research presents and the use of the scientific method in dealing with those challenges.
References
- C. Tonolio et al., Biopolymers 1983, 22, 205-215.
- G.R. Marshall et al., Proc. Natl. Acad. Sci. U.S.A. 1990, 87,487-491.
- NovaBiochem 1997/98 Catalog & Peptide Synthesis Handbook 1996, S38
- E. Frerot et al., Tetrahedron 1991, 47, 259-270.
- Many thanks to the Department of Chemistry and Biochemistry for their support of this research. Also, the assistance of Dr. Paul Savage with HPLC and Dr. Du Li with NMR was greatly appreciated, as well as the constant mentorship of Dr. Roger Harrison.