In the Laboratory
What Is the True Color of Fresh Meat? A Biophysical Undergraduate Laboratory Experiment Investigating the Effects of Ligand Binding on Myoglobin Using Optical, EPR, and NMR Spectroscopy Kimberly Linenberger, Stacey Lowery Bretz, Michael W. Crowder, Robert McCarrick, Gary A. Lorigan,* and David L. Tierney* Department of Chemistry and Biochemistry, Miami University, Oxford, Ohio 45056, United States *
[email protected] (G.A.L.) and
[email protected] (D.L.T.).
The new ACS Committee on Professional Training guidelines call for more integrated upper-level undergraduate laboratories (1). However, few such experiments have been described in the literature. A review of approximately 250 modern physical chemistry experiments, covering both laboratory instrumentation and current research topics, found only seven could be considered integrated (2). A comparison of the experimental techniques used in chemistry research and those published for use in undergraduate laboratories, presented in this Journal, found several obvious omissions (3). One underrepresented technique is electron paramagnetic resonance (EPR) spectroscopy. Only four experiments that use EPR spectroscopy have appeared in this Journal since 2000 (4-7). The experiment discussed below combines modern spectroscopic techniques (EPR, NMR, and optical spectroscopy), set in the context of better understanding meat packaging practices in the United States. Controversy has been sparked with the revelation that red meat is often packaged in an oxygen free, carbon monoxide rich environment (8-10). The brown color associated with the aging process is chiefly the result of air oxidation of myoglobin (Mb), upon prolonged exposure to oxygen. Packaging in a CO atmosphere preserves the pink color associated with freshness, making the meat appear fresher than it actually is. In this experiment, students investigate this issue by examining the effect of exchanging the distal ligand of Mb on its optical and magnetic properties. Myoglobin is a single chain, 154 amino acid polypeptide (17 kDa) that folds into eight R-helices (11); it contains a single heme, coordinated proximally by a histidine side chain and distally by a water molecule. In its reduced Fe(II) form, Mb, which is ubiquitous in muscle tissue, binds and stores oxygen in muscle for use under conditions of stress and normal cellular metabolism. Oxidation of the heme to Fe(III) eliminates O2 binding and is responsible for the color change that is observed as red meat ages. The color change depends on both the oxidation state of the iron and the identity of the distal ligand. When beef is first cut, the meat is a purple color due to the prevalence of reduced myoglobin. Upon exposure to air, the Fe(II) binds oxygen, forming oxymyoglobin, which is responsible for the bright pink color often associated with freshness. However, upon prolonged exposure to oxygen, the iron oxidizes to form metmyoglobin (metMb), which is brown (12-14). The addition of
_
CO converts high-spin aquometMb to the low-spin metMbCO complex, which is close in color to the pink observed in oxymyoglobin. Addition of azide to aquometMb results in a similar change in spin state and therefore color. Myoglobin has been the focus of multiple laboratory investigations described in this Journal (15-21), including a detailed optical titration of metMb with azide (16). However, the present experiment is the first to incorporate EPR and paramagnetic NMR spectroscopy, for any system. Guided by UV-visible spectroscopy, paramagnetic resonance is used to examine the effect of replacing the coordinated water molecule in resting metMb with the azide anion, which converts the ferric heme from high to low spin. A similar conversion takes place on formation of the CO adduct, making the connection to the meat packing process. The laboratory consists of (i) UV-visible, (ii) continuous wave (CW) EPR, and (iii) NMR spectroscopy of metMb, with (metMbN3) and without (aquometMb) added azide. Given pre-laboratory instruction introducing EPR and paramagnetic NMR spectroscopy, the experiments can be completed in two 3-4 h laboratory periods with students working in pairs. This biophysical chemistry experiment is appropriate for an upper-level undergraduate laboratory course. Experimental Procedure Samples of Mb were prepared for spectroscopy (22) by dissolving commercially available, lyophilized horse skeletal muscle Mb in 0.2 M phosphate buffer at pH 7. Mb was fully oxidized to metMb by the addition of a 2-fold molar excess of K3Fe(CN)6, and subsequent dialysis, according to published procedures (16). NMR samples were prepared using buffer made from 99.8% D2O and adding K3Fe(CN)6 directly. The metMbN3 complex was subsequently formed by adding a 2-fold molar excess of NaN3. UV-Visible Spectroscopy Optical spectra were collected for both 10 and 80 μM samples of aquometMb and metMbN3 using an Agilent model 8435 diode array spectrophotometer. Typical spectra are shown in Figure 1A. Students were asked to determine transition energies in both reciprocal centimeter (cm-1) and joules, from
_
r 2010 American Chemical Society and Division of Chemical Education, Inc. pubs.acs.org/jchemeduc Vol. 88 No. 2 February 2011 10.1021/ed100585t Published on Web 11/17/2010
_
Journal of Chemical Education
223
In the Laboratory
Figure 1. Spectroscopy of aquometMb and metMbN3. (A) UV-visible spectra of 10 μM samples accentuating the Soret band. (Inset) 80 μM spectra accentuating the R, β, and charge-transfer bands. (B) X-band CW EPR at 77 K. (C) 200 MHz 1H NMR.
a plot of absorbance versus wavelength. The two concentrations allow for calculation of molar absorptivities for the charge transfer region.
respectively) are a vibronically coupled, mostly metal-based pair of π f π* transitions that are enhanced in the low-spin state.
EPR Spectroscopy
The EPR spectra offer a nice demonstration of the effect of orbital angular momentum on the magnetic symmetry of the complex. Deviations of g from the free electron value of 2.0023 arise due to interactions between the electron spin, S, and orbital angular momentum, L. Typically, S = 1/2 complexes display minimal spin-orbit coupling, with g values close to 2. In contrast, S > 1/2 complexes often exhibit much larger anisotropy in g (24). Here, the observed g values collapse from an axial signal (g = [5.76, 5.76, 2.00]) for high-spin aquometMb to a rhombic signal (g = [2.78, 2.21, 1.73]) for the low-spin azide complex. Students are instructed to calculate the observed g values and describe the EPR spectra as isotropic (gx = gy = gz), axial (gx = gy 6¼ gz), or rhombic (gx 6¼ gy 6¼ gz).
X-band CW EPR spectra were collected on 5 mM samples of aquometMb and metMbN3 (30% v/v glycerol was added as a glassing agent) using a Bruker EMX EPR spectrometer operating at 77 K, with temperature maintained by an Oxford Instruments ER 4112HV continuous flow cryostat. Results similar to those shown in Figure 1B can be easily obtained using a liquid nitrogen finger dewar. NMR Spectroscopy NMR spectra were collected on 5 mM samples of aquometMb and metMbN3 with a Bruker ASX 200 NMR spectrometer (νH = 200 MHz). Using the parameters included in the supporting information, spectra such as those shown in Figure 1C can be obtained in 10 min or less. Concentrations as low as 2 mM can be used to conserve material, without significantly affecting data acquisition. Students were asked to determine how many paramagnetically shifted 1H resonances were observed (10 for aquometMb and 7 for metMbN3), and to report the shifts in both parts per million (ppm) and hertz (Hz). Chemical shifts were referenced to the water resonance at 4.7 ppm. Interpretation of Data Central to this experiment is the concept of ligand field strength. The hydroxide anion, as a moderately weak-field ligand, produces a high-spin complex in aquometMb. In contrast, the azide anion is a fairly strong field ligand, leading to a low-spin complex in metMbN3. All of the spectroscopic methods employed show a clear effect of the change in spin state. UV-Visible Spectroscopy Using the spectra in Figure 1A, students are asked to explain the changes in the electronic spectra that accompany complex formation. The Soret band, which is a π f π* transition originating in the porphyrin macrocycle, red shifts from 409 to 420 nm on formation of the low-spin azide complex due to decreased interaction with the Fe(III) ion's frontier orbitals (23). The high-spin aquometMb complex favors the two ligand-tometal charge-transfer (LMCT) transitions at 503 and 633 nm. The LMCT at 503 nm vanishes on conversion to the low-spin complex, and the 633 nm band is diminished. In contrast, the R and β bands in the low-spin complex (λ = 575 and 545 nm, 224
Journal of Chemical Education
_
Vol. 88 No. 2 February 2011
_
EPR Spectroscopy
NMR Spectroscopy Most students' prior exposure to NMR spectroscopy will have been in the organic and possibly analytical laboratories. Their studies will have been focused on diamagnetic molecules, where structural information can be derived from a combination of chemical shift and spin-spin, J, couplings. The presence of a paramagnetic center dramatically alters the NMR properties of surrounding nuclei, leading to an expanded chemical shift range, unresolved J couplings, and enhanced relaxation rates (25). What should be immediately apparent in comparison of the spectra in Figure 1C is the collapse of the pattern, on the highspin to low-spin conversion. The observed chemical shift, δobs H (eq 1), is the sum of the diamagnetic contribution, δdia H , and two paramagnetic components, the through-space dipolar (pseudoC contact) shift, δD H (eq 2), and the contact shift, δH (eq 3) that is the result of through-bond polarization of electron spins (25). Both depend directly on the spin state of the complex, which changes 5-fold (from 5/2 in aquometMb to 1/2 in metMbN3), and on the average g-value, which is reduced 2-fold (from 4.51 to 2.24). dia D C δobs H ¼ δH þ δH þ δH
δD H
¼
! C D gav SðS þ 1Þ r3 kT
C D ¼ ð10 - 7 Þ
ð1Þ μ2B ð3 cos2 θ - 1Þ 9 ð2Þ
!
δCH ¼
pubs.acs.org/jchemeduc
_
CC Aiso gav SðS þ 1Þ kT
CC ¼
μB 2π3γI
ð3Þ
r 2010 American Chemical Society and Division of Chemical Education, Inc.
In the Laboratory
The NMR aspect of this experiment opens a number of avenues for more advanced students. For example, the paramagnetic shift is inversely dependent on temperature and Mb is quite thermally stable. Further, the electronic relaxation rate of highspin Fe(III) is as many as 3 orders of magnitude faster than for low-spin Fe(III), and this directly affects the observed relaxation rates (which could be explored using T1 measurements) and line widths (dependent on T2, accessible through a suitable line-shape analysis). The dipolar shift also depends on the angle (θ) between the metal-proton vector and the magnetic z axis of the complex (eq 2). For the current systems, the magnetic z axis is perpendicular to the plane of the porphyrin ring, and consequently, this term is the same (3 cos2 θ - 1 = -1) for all observable protons. However, addition of a proton-containing axial ligand, such as imidazole, would afford the opportunity to explore the angular dependence further. Hazards Standard laboratory safety precautions should be followed, including the use of goggles and gloves. While weighing lyophilized myoglobin, students should be careful not to inhale the protein powder and avoid contact with the skin and eyes. It is important to note that both ferricyanide and azide are toxic. These compounds both liberate a very toxic gas should they come in contact with acid. Typical Student Results The spectra collected by students, shown in Figure 1, are similar to those found in the literature (22, 23, 26-30). Students typically struggled with EPR conceptually, especially g values. The comparison between NMR, where field is constant and frequency is “swept”, and EPR, where the frequency is fixed and it is the magnetic field that is swept, also proved conceptually difficult for the students and should therefore be addressed in any pre-laboratory discussion. In addition, students found it difficult to interpret the paramagnetic NMR spectra. After multiple laboratories using NMR to determine the structure of diamagnetic molecules, it was hard for students to embrace the use of fundamental properties of the spectra (i.e., chemical shift and line width) to extract structural information. We encourage the use of pre-lab discussions to solidify these concepts prior to the collection of data. The spectroscopic investigations clearly demonstrate that the azide anion is a strong-field ligand, producing a low-spin complex. Students were asked to explain how packaging meat in a carbon monoxide environment keeps meat a pink color and state their opinion as to the soundness of this practice. Students commonly cited the “opportunity to explore various instruments (especially EPR)” as a strength of the experiment. Finally, several students indicated an appreciation for solving a “real-life problem with chemistry”. Conclusion This experiment provides an everyday context for the integration of multiple spectroscopic techniques, through examination of the electronic environment of the heme center of metMb. Analysis of multiple spectra provides students with exposure to ligand field theory, EPR spectroscopic methods and theory, and paramagnetic NMR theory. Universities without access to an
r 2010 American Chemical Society and Division of Chemical Education, Inc.
_
EPR or NMR spectrometer can use the spectra in the supporting information and the students will still be able to gain a solid understanding and appreciation of this unique biophysical chemistry experiment. Literature Cited 1. ACS Committee on Professional Training, American Chemical Society: Washington, DC, 2008. 2. Abrash, S. A. In Advances in Teaching Physical Chemistry, ACS Symposium Series Vol. 973; Ellison, M.D., Schoolcraft, T.A., Eds.; American Chemical Society: Washington, DC, 2008; pp 115-151. 3. Sojka, Z.; Che, M. J. J. Chem. Educ. 2008, 85, 934–940. 4. Basu, P. J. Chem. Educ. 2001, 78, 666–669. 5. Berry, D. E.; Hicks, R. G.; Gilroy, J. B. J. Chem. Educ. 2009, 86, 76–79. 6. Garribba, E.; Micera, G. J. J. Chem. Educ. 2006, 83, 1229–1232. 7. Vincent, J. B.; Woski, S. A. J. Chem. Educ. 2005, 82, 1211–1214. 8. Diaz, K. Juicy Debate: What is Fresh Meat's True Color? Star Tribune of Minneapolis, MN, Nov. 14, 2007. 9. Schmit, J. E-Mails Surface about Safety of Meat Packaging, in USA Today, Nov. 13, 2007. 10. Weiss, R.; . Meat Treatment Got Approval Despite Safety Concerns. Washington Post, Nov. 14, 2007. 11. Voet, D.; Voet, J. G.; , Pratt, C. W. Fundamentals of Biochemistry: Life at the Molecular LevelI, 3rd ed.; John Wiley & Sons, Inc.: Hoboken, NJ, 2008. 12. Luno, M.; Roncales, P.; Djenane, D.; Beltran, J. A. Meat Sci. 2000, 55, 413–419. 13. Seyfert, M.; Mancini, R. A.; Hunt, M. C.; Tang, J.; Faustman, C. Meat Sci. 2007, 75, 432–442. 14. Jeong, J. Y.; Claus, J. R. Meat Sci. 2010, 85, 525–530. 15. Bylkas, S. A.; Andersson, L. A. J. Chem. Educ. 1997, 74, 426–430. 16. Marcoline, A. T.; Elgren, T. E. J. Chem. Educ. 1998, 75, 1622–1623. 17. Olchowicz, J.; Coles, D. R.; Kain, L. E.; MacDonald, G. J. J. Chem. Educ. 2002, 79, 369–371. 18. Stynes, H. C.; Layo, A.; Smith, R. W. J. Chem. Educ. 2004, 81, 266–269. 19. Nilsson, M. R. J. Chem. Educ. 2007, 84, 112–114. 20. Mehl, A. F.; Crawford, M. A.; Zhang, L. J. Chem. Educ. 2009, 86, 600–602. 21. Miller, S.; Indivero, V.; Burkhard, C. J. Chem. Educ. 2010, 87, 303–305. 22. Bizzarri, A. R.; Cannistraro, S. B. Bull. Mag. Res. 1992, 14, 234–239. 23. Lin, J.; Merryweather, J.; Vitello, L. B.; Erman, J. E. Arch. Biochem. Biophys. 1999, 362, 148–158. 24. Jones, R. In An Introduction to Spectroscopy for Biochemists; Brown., S. B., Ed.; Academic Press, Inc.: New York, 1980; pp 279-319. 25. F. A. Walker. In Spectroscopic Methods in Bioinorganic Chemistry, ACS Symposium Series, Vol. 692; Solomon, E. I., Hodgson, K. O., Eds.; American Chemical Society: Washington, DC, 1998; pp 30-61. 26. Brill, A. S.; Williams, R. J. P. Biochem. J. 1961, 78, 246–253. 27. Morishima, I.; Iizuka, T. J. Am. Chem. Soc. 1974, 96, 5279–5283. 28. Morishima, I.; Ogawa, S.; Inubushi, T.; Yonezawa, T.; Iizuka., T. Biochemistry 1977, 16, 5109–5115. 29. LaMar, G. N.; Budd, D. L.; Smith, K. M. Biochim. Biophys. Acta 1980, 622, 210–218. 30. Lukat, G. S.; Goff, H. M. J. Biol. Chem. 1986, 261, 16528–16534.
Supporting Information Available Notes for the instructor, including a description of chemicals and hazards and detailed procedures; answers to the assigned questions; the student laboratory handout. This material is available via the Internet at http://pubs.acs.org.
pubs.acs.org/jchemeduc
_
Vol. 88 No. 2 February 2011
_
Journal of Chemical Education
225
