In the Laboratory
Advanced Chemistry Classroom and Laboratory
edited by
Joseph J. BelBruno Dartmouth College Hanover, NH 03755
Circular Dichroism of Globular Proteins
W
Brenda A Bondesen and Merlyn D. Schuh* Department of Chemistry, Davidson College, Davidson, NC 28036-1719; *
[email protected] Rationale for Experiments For decades circular dichroism spectroscopy (CD) has been an important method used by biochemists to analyze structure in globular proteins (1). As a method of learning about protein structure, CD spectroscopy is pedagogically more significant than techniques such as routine ultraviolet (UV) absorption and fluorescence spectroscopies, because it can reveal greater detail. Most routine UV absorption and fluorescence studies of proteins provide only qualitative information about tertiary structural changes in the microenvironments of the three aromatic amino acid residues phenylalanine (Phe), tyrosine (Tyr), and tryptophan (Trp). In contrast, CD spectra provide separate information about both tertiary structure and secondary structure. Also, subtle changes in CD spectra can be more readily observed and interpreted, and unlike UV absorption, particular CD spectral features characterize the specific types of secondary structure present in proteins. Despite the need for chemistry students to receive instruction in the use of CD spectroscopy, a search of the chemistry and biochemistry literature yielded no experimental descriptions of CD that are suitable for undergraduate instruction. To help meet this need, we have developed experiments, based on research results described in the biochemistry literature, that teach students some important applications of CD to protein biochemistry during a 3- to 4-hour junior- or senior-level laboratory session. The goal is to present an overview of the biochemical applications of CD rather than an in-depth understanding of a single application, requiring more thorough understanding of CD than is warranted in a one-afternoon experiment. The experiments are straightforward and can be readily adopted into a junior/senior laboratory course. Concepts and Experimental Topics
Protein Structure and Protein Folding/Unfolding Four levels of structure provide the overall three-dimensional shape or conformational state of a globular protein. 1. Primary structure (1°) refers to the sequence of amino acid residues along the covalently bonded backbone of the protein molecule. 2. Secondary structure (2°) refers to local repetitive structures within proteins that are maintained by hydrogen bonds and are found in most globular proteins. Two common secondary structures are α-helices and β-sheets. 3. Tertiary structure (3°) refers to the overall three-dimensional shape into which the polypeptide backbone folds in creating a unique shape (native state) for each globular protein.
1244
4. Quaternary structure (4°) refers to the specific spatial relationship between individual polypeptide chains in protein molecules that consist of more than one polypeptide.
Characterization of these four levels of structure and the changes that occur in them when a protein passes between the chemically active native state (N) and unfolded (denatured) state (D) is fundamental in understanding proteins (2, 3). A variety of experimental approaches have been developed to examine the 2°, 3°, and 4° structures, and the transition between native and unfolded states can be induced by agents such as chemical denaturants and temperature. During protein folding, the tertiary structure is believed to pass through a partially folded molten globule conformation, which is generally assumed to closely resemble the native-state conformation but to have weaker internal interactions and greater internal flexibility. Characterizing the molten globule conformation is important because it is believed to be the last essential conformation through which a protein folds in achieving the native state. Experimental studies of protein structures and transitions between the native and unfolded states are important in revealing how functional proteins are translated from DNA and in understanding the amyloid diseases (such as cystic fibrosis, scurvy, and possibly Alzheimer’s disease), which are believed to be causally related to the misfolding of proteins (4–6 ). Studies of protein misfolding have been aided by the ability of 2,2,2trifluoroethanol (TFE) to convert a predominantly β-sheet protein structure into a mostly α-helix structure (termed helicogenesis). The helicogenicity of TFE is believed to arise because it stabilizes local hydrogen bonding, as occurs in an α-helix, more than nonlocal hydrogen bonding, as occurs between nonadjacent strands of a β-sheet structure (7 ).
Circular Dichroism Spectroscopy The brief description of CD, including equations, presented here is based primarily on ref 1. More detailed descriptions can be found in many physical chemistry texts and refs 1 and 8–10. Circular dichroism refers to the difference between the absorbance of left- and right-circularly polarized components of radiation. This phenomenon occurs when the chromophore is chiral (optically active) as a result of either its intrinsic structure, its being covalently bonded to a chiral center, or its being located in a chiral microenvironment. The recombined, differentially absorbed components produce radiation that is elliptically polarized (i.e., traces out an ellipse) (Fig. 1A). A CD instrument reports the dichroism as either the difference in absorbance, A, of the two components, ∆A = AL – AR, or as the ellipticity in degrees, θ. As shown in Figure 1B, θ is
Journal of Chemical Education • Vol. 78 No. 9 September 2001 • JChemEd.chem.wisc.edu
In the Laboratory
A
B
L
α
E R
a/2 θ b/2
Figure 1. (A) Elliptically polarized light, E, produced by the vectorial sum of right (R) and left (L) circularly polarized light for which the intensities are unequal owing to the differential absorption by the sample of each component. (B) Ellipticity, θ, is defined as the angle for which the tangent is the ratio of the semiminor axis, b/2, to the semimajor axis, a/2, of the ellipse. Here the elliptically polarized light has been optically rotated through angle α.
defined as the arctan of the angle formed by a right triangle of base (a/2) and height (b/2) equal to the semimajor and semiminor axes of the ellipse, respectively: θ = arctan (b/a)
Since θ is usually very small, tan θ can be approximated by θ in radians, and it can be shown that θ (degrees) = 32.98 ∆A
To remove the linear dependence on path length () and concentration (C), molar ellipticity [θ] is defined as follows:
θ ≡ 100 θ = 3298∆ε C where ∆ε ≡ εL – εR and ε is the molar extinction coefficient. In most biochemistry experiments ellipticities are very small, on the order of 10 millidegrees, corresponding to ∆A on the order of 3 × 104. CD spectroscopy can be used to distinguish various aspects of protein structure because different structural elements of proteins absorb radiation over different wavelength ranges. Aromatic amino acid residues absorb in the near-UV region (250–290 nm) and have finite CD signals when, as is usually the case, the microenvironments are asymmetric. Since the microenvironment for each such amino acid in a protein is different, the CD spectrum in the near-UV can be complicated. Nevertheless, since the asymmetry of the microenvironments is lost when a protein unfolds, the corresponding decrease in near-UV CD signal qualitatively reflects the degree of tertiary structure loss around the aromatic chromophores. The peptide bond is the principal absorber in the far-UV region (typically 180 or 190 nm to 250 nm), which can be used to assess the amount of secondary structure of the entire protein because α-helices and β-sheets are asymmetric structures. Also, since all α-helices are similar and all β-sheets are similar, both classes of structure have characteristic far-UV CD spectra. However, assessments of secondary structure must be made with care because absorption of radiation by aromatic residues also contributes to the far-UV spectrum. From the overall shape of the far-UV spectrum, many proteins can be characterized qualitatively as being predominantly α-helical, predominantly β-sheet or a mixture of α-helical and β-sheet structures.
The specific topics addressed in this paper qualitatively compare CD spectra for different secondary structures, demonstrate the utility of CD to provided information about the structure of a molten globule, use CD to follow protein unfolding, and demonstrate helicogenesis. Experimental Chemicals Chicken egg white lysozyme (CAS: 12650-88-3), concanavalin A (type III) (CAS: 11028-71-0), horse heart cytochrome c (CAS: 9007-43-6), horse heart myoglobin (CAS: 100684-32-0), and guanidine hydrochloride (99+%) (CAS: 50-01-1) were obtained from Sigma. Chemicals used in preparing buffers—potassium dihydrogen phosphate (CAS: 7778-77-0), potassium hydrogen phosphate (CAS: 7758-114), and sodium sulfate (CAS: 7757-82-6)—were obtained from Sigma and Fisher. 2,2,2-Trifluoroethanol (99%) (CAS: 75-89-8) was obtained from Acros.
Procedures All protein solutions were aqueous. Buffers were 0.1 M potassium phosphate pH 7.0 for native state proteins; 0.01 M potassium phosphate pH 2.1 for acid-denatured cytochrome c; 0.35 M sodium sulfate pH 2.1 for the A state of cytochrome c; 0.10 M potassium phosphate pH 2.6 for denaturation of lysozyme; and 0.004 M potassium phosphate pH 7.5 for helicogenesis (the final buffer was 0.002 M in solutions containing 50% TFE). Complete descriptions of how the experiments were performed are available in the supplemental material.W The A state of cytochrome c is believed to resemble the molten globule conformation. Since the A state is typically stabilized in solutions of high ionic strength and low pH, the 0.35 M sodium sulfate pH 2.1 buffer was used to prepare the A state of cytochrome c. Lysozyme denaturation was studied by rapidly mixing in a beaker a buffered lysozyme solution with a buffered guanidine hydrochloride solution (final guanidine hydrochloride concentration 5.0 M). A portion of the mixture was quickly loaded into a cuvette in the CD spectrophotometer. The estimated dead time was about 13 s. Helicogenesis of concanavalin A was carried out using 50/50 volume % solutions of buffered native protein and TFE, and the CD spectra were recorded within a minute after mixing. Protein concentrations were determined spectrophotometrically using literature values for the molar absorptivities: chicken egg white lysozyme (ε280 = 3.65 × 104 M 1 cm1) (11), concanavalin A (ε280 = 7.98 × 104 M 1 cm1 for the tetrameric form at pH 7) (12), horse heart cytochrome c (ε410 = 1.06 × 105 M 1 cm1) (13), and horse heart myoglobin (ε409 = 1.71 × 105 M 1 cm1) (14 ). Protein concentrations were generally 1–2 × 10-5 M and 1 × 104 M for obtaining CD spectra in the 190–250-nm and 250–300-nm wavelength ranges, respectively. Far-UV spectra were recorded for solutions in a 0.1-cm circular cuvette and near-UV spectra were recorded for solutions in a 1.0-cm rectangular cuvette. Instrumentation CD spectra were recorded with a Jasco model J-715 spectropolarimeter. Absorbance measurements were made with a Perkin-Elmer lambda 6 UV–vis spectrophotometer.
JChemEd.chem.wisc.edu • Vol. 78 No. 9 September 2001 • Journal of Chemical Education
1245
In the Laboratory
Structural Characteristic spectral features Type β-sheet Single minimum at 224 nm
Protein
Ref
Concanavalin A
—
Poly-L-lysine
15
β-sheet
Single minimum at 218 nm
Lysozyme
—
α+β
α-Helical spectral features dominate, but the intensities of the double minimum are roughly reversed relative to an α-helix
Myoglobin
—
α-helical Double minimum at about 209, 222 nm
15
α-helical Double minimum at about 209, 221 nm
Poly-L-lysine
8
[θ] / (106 deg cm2 dmol 1)
Table 1. Features in the CD Spectra of Some Globular Proteins and a Reference Structure
1246
c
0
-4
210
230
250
Figure 2. Far-UV CD spectra for native-state proteins (a) concanavalin A (3.59 µM), (b) lysozyme (12.7 µM), and (c) myoglobin (18.7 µM).
2
b
[θ] / (106 deg cm2 dmol 1)
Figure 2 shows the far-UV CD spectra for myoglobin (predominantly α-helical), concanavalin A (predominantly β-sheet), and lysozyme (α + β protein), and spectral features are given in Table 1. Other proteins with predominantly α-helix, predominantly β-sheet, or α + β structures have similar spectra. This can be seen in Table 1 by comparing the wavelengths of protein spectral features to those for poly-L-lysine (which is often used as a reference compound) when it is contained in solvents that make its structure exclusively α-helical or β-sheet (15). Figure 3 shows the far-UV CD spectra for the native and unfolded states of cytochrome c. The absence of a spectral minimum and reduction in magnitude of the signal shows the loss of secondary structure in the unfolded state. The third spectrum (Fig. 3b) is similar but not identical to that of the A state, which is nearly identical to the native-state spectrum. However, Figure 3b is similar enough to be called the A state spectrum herein. For this A-state spectrum the presence of a double minimum, the magnitude of the signal, and a shape similar to the native-state spectrum reveal the retention of much secondary structure in the molten-globule-like state. The near-UV spectral shapes for the native state and A state of cytochrome c (not shown) are very similar. However, conversion of the native state into the A state involves a larger percentage decrease in the CD signal strength in the near-UV than in the far-UV, revealing the larger relative loss of tertiary structure in the A state. Thus, the observed spectral features for the native and A states of cytochrome c are consistent with the view that molten globules retain much of the native state secondary structure but less of the tertiary structure (17). The stability of the molten globule state is partly explained qualitatively as follows. At a low pH, many protonated amino acid residues (notably lysines and arginines) are positively charged. Electrostatic repulsion between these cationic groups pushes apart regions of the polypeptide that are otherwise in close contact in the native state. If the ionic strength is high at low pH, the charge-stabilizing ionic environment surrounding cationic groups partially offsets the electrostatic repulsions so that regions of the polypeptide are maintained in relatively
b
Wavelength / nm
a 1
0
-1
-2 190
c
200
210
220
230
240
250
Wavelength / nm Figure 3. Far-UV CD spectra for the (a) native state (20.9 µM), (b) A state (20.9 µM), and (c) acid-denatured state (20.0 µM) of cytochrome c. The conformations of the A state and the molten globule of cytochrome c are believed to be similar.
1
0
-1
ln(Z )
Results and Discussion
4
-8 190
Hazards 2,2,2-Trifluoroethanol (TFE) is a volatile lachrymator that may be harmful if absorbed through the skin or swallowed. It may cause severe eye irritation and skin irritation. It should be used in a hood, handled with gloved hands, and disposed of with halogenated waste.
a
-2
-3
-4
-5 0
50
100
150
200
250
Time / s Figure 4. Plot of ln Z vs time according to eq 1 for the CD signal recorded at 289 nm for the denaturation of lysozyme. The apparent first-order kinetics are consistent with a two-state model for protein folding/unfolding, N D. Similarly, the CD signal at 222 nm (not shown) also decays exponentially.
Journal of Chemical Education • Vol. 78 No. 9 September 2001 • JChemEd.chem.wisc.edu
In the Laboratory
protein structures and folding patterns. As is seen in Figure 5, when TFE is added to the solution, the CD spectrum for native-state concanavalin A changes from one having a single minimum to one having minima at both 208 and 221 nm. The minima at these wavelengths clearly indicate conversion of some of the β-sheet structure into α-helix. However, the fact that the minimum at 221 nm is more shallow than the one at 208 nm reveals that not all of the β-sheet structure undergoes conversion. In fact, the presence of TFE in solution makes the spectrum more closely resemble that for the α + β protein lysozyme.
[θ] / (106 deg cm2 dmol 1)
40
a 20
b 0
-20 190
210
230
250
Acknowledgments
Wavelength / nm Figure 5. CD spectra for concanavalin A (a) in the native state and (b) after conversion of the molecular structure into a predominantly α-helical form.
close proximity, and the protein (molten globule) more closely resembles the native state than a denatured state. For denaturation of lysozyme the CD signal at 222 nm and 289 nm decays exponentially with time, consistent with a two-state denaturation model, N D. For this model the first-order rate equation for [N] can be integrated and put into the form of eq 1:
Z=
CD – CDeq CD0 – CDeq
=
ekt
(1)
where CD = circular dichroism signal at time t, CD0 is the circular dichroism signal extrapolated to time 0, CDeq is the circular dichroism signal at equilibrium, and k = kunfold + kfold, which is equal to the sum of microscopic rate constants for N → D and D → N (18). CD data at 289 nm are plotted according to eq 1 in Figure 4. A nonlinear plot in Figure 4 would show the inadequacy of the simple two-state unfolding model and would imply a more complicated unfolding mechanism, perhaps involving one or more intermediates. Plots of ln Z versus time are linear and have slopes of 1.55 ± 0.02 × 102 s1 for the CD signal at 222 nm and 1.37 ± 0.02 × 102 s1 for the CD signal at 289 nm. These slopes indicate the similarity of folding/unfolding rates for tertiary and secondary structure and suggest that during denaturation both secondary and tertiary structures are lost on a similar time scale. Because of the 13-s dead time mentioned in the Experimental section, a significant portion of the CD signal is not recorded. However, this loss of data is inconsequential because the same slope of the linear plot in Figure 4 will be obtained, and extrapolation of the plot to time zero makes it possible, in effect, to recover the lost data completely if desired. Although a similar linear plot involving change in simple UV absorption by lysozyme could also be used to follow protein denaturation, the utility of CD is in being able to resolve the unfolding of both secondary and tertiary structure. Insights into protein folding/unfolding can be gained by studying the solute–solvent interactions that stabilize various
This work was supported in part by NSF award MCB9417885. Acknowledgment is also made to the donors of the Petroleum Research Fund, administered by the American Chemical Society. W
Supplemental Material
A student handout containing background information and instructions for the experiment and a list of questions (with answers) is available in this issue of JCE Online. Literature Cited 1. Fasman, G. D. Circular Dichroism and the Conformational Analysis of Biomolecules; Plenum: New York, 1996. 2. Fersht, A. Structure and Mechanism in Protein Science; Freeman: New York, 1999; Chapters 17 and 18. 3. Levitt, M.; Gerstein, M.; Huang, E.; Subbiah, S.; Tsai, J. Annu. Rev. Biochem. 1997, 66, 549–579. 4. Thomas, P. J.; Qu, B. H.; Pederson, P. L. Trends Biochem. Sci. 1995, 20, 456–459. 5. Sifers, R. N. Struct. Biol. 1995, 2, 355–357. 6. Chiti, F.; Webster, P.; Taddei, N.; Clark, A.; Stefani, M.; Ramponi, G.; Dobson, C. M. Proc. Natl. Acad. Sci. USA 1999, 96, 3590–3594. 7. Cammers-Goodwin, A.; Allen, T. J.; Oslick, S. L.; McClure, K. F.; Lee, J. F.; Kemp, D. S. J. Am. Chem. Soc. 1996, 118, 3082. 8. Pelton, J. T.; McLean, L. R. Anal. Biochem. 2000, 277, 167–176. 9. Greenfield, N. J. Trends Anal. Chem. 2000, 18, 236–244. 10. Kelly, S. M.; Price, N. C. Biochim. Biophys. Acta 1997, 1338, 161–185. 11. Kuroki, R.; Yamada, H.; Moriyama, T.; Imoto, T. J. Biol. Chem. 1986, 261, 13571–13574. 12. Christie, D. J.; Alter, G. M.; Magnuson, J. A. Biochemistry 1978, 17, 4425–4430. 13. Margoliash, E.; Frohwirt, N. Biochem. J. 1959, 71, 570–572. 14. Puett, D. J. Biol. Chem. 1973, 248, 4623–4634. 15. Greenfield, N.; Fasman, G. D. Biochemistry 1969, 8, 4108– 4116. 16. Marmorino, J. L.; Pielak, G. Biochem. 1995, 34, 3140–3143. 17. Baldwin, R. L. Chemtracts—Biochem. Mol. Biol. 1991, 2, 379. 18. Cantor, C. R.; Schimmel, P. R. Biophysical Chemistry; Freeman: San Francisco, 1980; pp 1088–1090. 19. Ikai, A.; Tanford, C. Nature 1971, 230, 100–102.
JChemEd.chem.wisc.edu • Vol. 78 No. 9 September 2001 • Journal of Chemical Education
1247
