In the Laboratory
Using Infrared Spectroscopy to Investigate Protein Structure
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Janet C. Olchowicz, Deidra R. Coles, Lori E. Kain, and Gina MacDonald* Department of Chemistry, James Madison University, Harrisonburg, VA 22807; *
[email protected] Biochemistry laboratories include numerous molecular and biophysical methods to reflect the broad range of techniques currently used in the study of proteins (1–5). Concurrently, many project-oriented laboratories have been implemented to allow undergraduates to participate in experiments that more accurately reflect the environment of a research laboratory. These project-oriented laboratories introduce students to a multitude of techniques that are used to solve a single problem; for some examples, see refs 3 and 6–8. Our upper-level biochemistry laboratory enrolls both chemistry and biology majors. Although the chemistry majors have had hands-on experience with UV–vis, NMR, and infrared spectrometers in the integrated organic–inorganic, projectoriented laboratory (6 ), most of the biology majors have had little exposure to these techniques. Furthermore, the chemistry majors do not know that these techniques are commonly used to study large molecules such as proteins. To emphasize the importance of these techniques in the study of biological molecules, we have introduced a variety of spectroscopic techniques into the upper-level biochemistry laboratory and moved toward a more project-oriented environment. This report describes a laboratory that utilizes infrared spectroscopy to observe a protein’s dominant form of secondary structure. This technique has also been used to monitor the unfolding of myoglobin and to determine its melting temperature. Infrared spectroscopy could easily be used to monitor the unfolding of any protein that has a substantial amount of α-helical structure and whose characterization is of interest in an inquiry-based laboratory.
relatively small shifts observed in amide I upon deuteration, secondary structures absorb in similar ranges in both H2O and D2O. Typically, α-helices show amide I frequencies in H2O around 1648–1660 cm᎑1 (1651–1657 cm᎑1 in D2O), whereas β structure is observed in the 1625–1640 cm᎑1 and 1675–1695 cm᎑1 regions in H2O and the 1620–1640 cm᎑1 and 1670–1680 cm᎑1 regions in D2O (9, 10). Unordered structure can be observed in the 1640–1648 cm᎑1 region in H2O. Aggregated, denatured proteins exhibit an amide I frequency around 1610–1628 cm᎑1 in H 2O. The use of deuterium oxide facilitates the distinction between helical and unordered secondary structure, as the latter has a larger shift (about 10 cm᎑1) upon deuteration than the approximately 5-cm᎑1 shift typical of helical structures (9). Myoglobin, a well-characterized α-helical protein composed of a single polypeptide chain and a heme group, has a molecular weight of about 18 kDa (11). It is an excellent choice for a project-oriented laboratory because it is stable and several spectroscopic techniques can be utilized to characterize the purified protein under a variety of conditions. Bylkas and Anderson developed a laboratory that uses protein purification and electronic absorption techniques to study the different redox states of myoglobin (12). Our experiment provides a solid addition to a project-oriented laboratory that incorporates biochemical and spectroscopic studies of myoglobin or other proteins that are isolated by the students during the course of the semester. The purification methods utilized by our students include those of Bylkas and Anderson as well as those of Shimada and Caughey (11, 12).
Background
Summary of Materials and Methods
Infrared spectroscopy is useful for the characterization of protein secondary structure and the identification of protein components involved in events such as ligand binding and electron-transfer reactions (9). The amide I frequencies were used to determine primary structural components and to monitor unfolding reactions in the experiments described in this report. The two dominant bands in any protein’s absorbance spectrum are the amide I and amide II absorptions. These peaks are sensitive to hydrogen-bonding interactions, and therefore they are sensitive to differences in protein secondary structure. The amide I band absorbs around 1655 cm᎑1 and is predominately due to contributions from the C=O stretching vibration of the peptide backbone. The amide II (around 1550 cm᎑1) vibration arises from the N–H bending and C–N stretching vibrations of the backbone. Because water has a strong absorbance in the region of interest, infrared studies are facilitated by the use of deuterium oxide as the solvent. Under these conditions, the O–H vibration of water is shifted 400 cm᎑1 (9). However, the protein’s amide II vibration downshifts approximately 100 cm᎑1, while the amide I frequency remains in the same region (only 5–10 cm᎑1 shifts) (10). Owing to the broad absorption range and the
Equipment and Materials Any standard commercial Fourier transform infrared spectrometer (FTIR) can be used to perform these experiments. We have used both a Midac FTIR and a Nicolet Impact 410 FTIR. Calcium fluoride windows (25 × 4 mm), 15-µm Teflon spacers, and sample holders can be obtained from Spectra Tech (Shelton, CT). Vacuum grease is used to help seal the infrared cell. Proteins and deuterium oxide can be purchased from a variety of vendors. Methods Proteins were dissolved in deuterium oxide to a concentration of approximately 60 mg/mL. If protein unfolding is being monitored, separate tubes containing about 25 µL of the myoglobin solution were incubated at the appropriate temperature (30–90 °C) for approximately 15 min before the infrared spectra were obtained. Infrared spectra were obtained at room temperature. About 10–20 µ L of protein solution was deposited on a calcium fluoride window having a 15-µm spacer. The cell was sealed with the second calcium fluoride window. If spacers are not available, very small dabs of vacuum grease can be used around the window edges to seal the IR cell. Data were
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In the Laboratory 0.30 1641
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myoglobin 78 °C myoglobin 62 °C
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Figure 1. Absorption spectra of ~60 mg/mL solutions of cytochrome c and trypsinogen dissolved in D2O. Ten microliters of each protein sample was placed between calcium fluoride windows separated by a 15-µm spacer. The data were recorded at 4-cm᎑1 resolution, and each spectrum is the result of 50 scans.
Figure 2. Absorption spectra of ~60 mg/mL myoglobin solutions in D2O preincubated at 62 and 78 °C for 15 min. Spectral conditions are identical to those described in Figure 1.
collected at 4-cm᎑1 resolution, and each spectrum is the result of 50 scans. The protein absorbance spectra were constructed by taking the ratio of the sample to a background spectrum obtained under identical conditions. Both the background and the sample spectra were recorded after a 5–15-min purge with dry air or nitrogen. To reduce the effect of water vapor on the absorbance spectra, purge times for the background and the samples should be identical. After the absorbance spectra were obtained, the center of the amide I vibration was determined by inspection.
As part of the laboratory students obtain absorbance spectra of a set of control protein samples and then identify the predominant secondary structure of their unknown. They have been extremely successful in identifying the larger percentage of secondary structure present in their unknown. Figure 2 shows absorbance spectra of myoglobin incubated at 62 and 78 °C. Comparison of the absorbance spectra of samples incubated at 62 °C and 78 °C reveals the unfolding of myoglobin at the higher temperature. Incubation at higher temperatures results in the downshift of the amide I peak and the appearance of a new peak around 1620 cm᎑1. Absorption in the 1610–1628 cm᎑1 region is due to denatured, aggregated protein (9). If students plot the maximum amide I absorbance versus temperature of incubation they generate a sigmoidal curve whose inflection point allows them to estimate the unfolding temperature of myoglobin. The derivative should be used to verify the melting temperature of myoglobin obtained from these experiments. Most students found this to be around 75 °C (72–80 °C), similar to the midpoint near 80 °C reported in the literature (14). The absorption due to denatured aggregated protein is particularly evident in some samples incubated at 95–100 °C. Therefore, care must be taken when using these data to generate the unfolding curve, and incubation temperatures may be limited to the 30–90 °C range unless the instructor wishes to address the distinction between unordered and aggregated structures.
Hazards There are no significant hazards associated with the materials utilized in this laboratory. However, the standard laboratory safety practices should be observed. Results and Discussion Figure 1 shows the absorbance spectra of cytochrome c and trypsinogen. The proteins used in this laboratory contained at least two times the amount of one secondary structure (αhelix) as the other (β-sheet), so that the prominent secondary structure could be ascertained by the infrared spectra. All spectra presented were obtained in D2O. The α-helical cytochrome has an amide I frequency centered around 1650 cm᎑1, whereas trypsinogen (containing a larger percentage of β-sheet structure) has an amide I frequency centered around 1640 cm᎑1 (10, 13). The spectra shown here are representative of those obtained on other proteins that contain at least twice the amount of one type of secondary structure. Other commercially available proteins that can be used in this experiment are myoglobin and lysozyme (α-helical) and α-chymotrypsin, β-lactoglobulin, and ribonuclease A (β structure) (10, 13).
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Conclusions This laboratory allows students to see how infrared spectroscopy can be used to observe secondary structure of proteins. It would be appropriate for upper-level biochemistry or biophysical chemistry laboratories. The experiment enables students to differentiate between proteins that are composed primarily of either α-helix or β-sheet secondary structures. It
Journal of Chemical Education • Vol. 79 No. 3 March 2002 • JChemEd.chem.wisc.edu
In the Laboratory
also allows students to experimentally estimate the melting temperature of myoglobin and could be applied to monitor the unfolding of any other protein that is largely α-helical. The techniques described could be incorporated into a project-oriented laboratory as a means to characterize a protein that the students have purified during the semester. If time does not permit students to purify their own myoglobin for the protein-unfolding experiments described in this report, the commercially available protein can readily be used. Extended projects may include more elaborate protein folding experiments such as those that monitor unfolding as a function of pH, concentration of chemical denaturant, or effect of added detergents. The spectroscopic characterization of proteins provides a means of integrating important chemical, spectroscopic, and biochemical concepts and reinforces the benefits of an interdisciplinary means of addressing biological problems. Acknowledgments We gratefully acknowledge support from the National Science Foundation (MCB-9733566). We also thank James Madison University and all of the students involved in testing this experiment over the course of its development. Supplemental Material The student handout for this experiment is available in this issue of JCE Online. W
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