- Email: [email protected]
In-situ monitoring of the interaction of lactate dehydrogenase with NAD on a gold electrode by FT-SERS
Journal of Electroanalytical Chemistry 465 (1999) 187 – 194
In-situ monitoring of the interaction of lactate dehydrogenase with NAD on a gold electrode by FT-SERS Yi-Jin Xiao a,*, Xiao-Xia Gao a, John Markwell b b
a Department of Chemistry, Peking Uni6ersity, Beijing 100871, PR China Department of Biochemistry, Uni6ersity of Nebraska-Lincoln, Lincoln, NE 68588, USA
Received 7 September 1998; received in revised form 2 February 1999
Abstract The FT-SERS method was used to observe directly and monitor the combination of dehydrogenases with NAD pre-adsorbed onto the surface of a roughened gold electrode. At negative electrode potentials, the SERS spectra of NAD exhibit significant changes upon the addition of lactate dehydrogenase. The enzyme active site may approach the electrode closely enough to produce several strong bands attributable to backbone peptide. The experimental results may be explained by binding of the NAD adenine moiety in a hydrohobic region of the enzyme while the nicotinamide group is left exposed near the surface of the electrode. This competitive binding state appears to be strongly dependent on the applied electrode potentials. Comparative experiments were carried out with lactate dehydrogenase and glutamate dehydrogenase. The SERS of NAD upon binding with lactate dehydrogenase show some features significantly different from those in the case of glutamate dehydrogenase. This difference may result from the different architectures within binding sites of type-A and type-B dehydrogenases. © 1999 Elsevier Science S.A. All rights reserved. Keywords: Gold electrode; Lactate dehydrogenase; FT-SERS; NAD
1. Introduction The requisite step for an enzymatic redox process is the binding of substrates within a specific intraprotein environment which stimulates reactivity, reduces the transition state barrier between substrates and products, and defines the specificity of the enzymatic reactions. As oxidoreductase enzymes important in diverse metabolic pathways, NAD-dependent dehydrogenases have been extensively studied by many methods [1–24]. Whereas much of our structural knowledge of this class of enzymes is derived from X-ray crystallographic measurement, the development of methods to study the behavior and properties of these enzymes in solution would be highly desirable; techniques such as infrared * Corresponding author. E-mail address: yjxiaopku.edu.cn (Y.-J. Xiao)
spectroscopy, capable of making measurements in solution, have been suggested to provides insight into molecular structures of proteins in solution [25–30]. However, the general applicability of infrared spectroscopy is hampered by the practical experimental difficulties in obtaining spectra of proteins in H2O, which absorbs strongly in the same spectral region as the conformation-sensitive amide bands in the polypeptide background. Thus, the conformational changes in tertiary structure induced by interaction with the NAD cofactor would be largely masked by the aqueous solvent. A complicated computational procedure would be needed for deconvolution of the complex amide absorption bands into their constituent components [25]. Recent reports [5,17–23] have employed normal Raman (NR) and resonance Raman (RR) spectroscopy to gain additional insights into the molecular structure of the NAD coenzyme upon binding with enzyme
0022-0728/99/$ - see front matter © 1999 Elsevier Science S.A. All rights reserved. PII: S 0 0 2 2 - 0 7 2 8 ( 9 9 ) 0 0 0 9 3 - 5
188
Y.-J. Xiao et al. / Journal of Electroanalytical Chemistry 465 (1999) 187–194
proteins. However, the absence of suitable chromophores from many enzymes makes it difficult to get RR spectra. In addition, the RR vibrational bands arising from the polypeptide backbone of the enzyme, such as the amide I band, are comparatively weak and do not provide much information about secondary or tertiary structure [5,31]. With normal Raman spectra, careful difference spectra are required to reveal the spectrum of the bound coenzyme. Many researchers have demonstrated the potential of SERS techniques for obtaining structure-function information about enzymes. Strong SERS signals from the surface-exposed aromatic residues of proteins have indeed been obtained [32 – 42]. However, it is difficult to get any SERS signals originating from the polypeptide backbone of the enzyme, which would yield information about protein structure. As has been reviewed by Cotton et al. [32], the screening by aromatic groups on the protein surface interferes with surface enhancement of the amide I bands. It has been proposed that structural changes of NAD induced by active-site binding to a dehydrogenase should theoretically be monitored by the SERS technique [43,44]. Unfortunately, no such enzyme-cofactor SERS signal was obtained [43]. The lack of significant surface enhancement was attributed to NAD being buried deeply in the dehydrogenase protein and implied that use of this technique to track the combination of enzymes, coenzyme and substrate in solution was problematic. In the present work, we have designed an original experimental procedure which provides high quality spectra attributable to NAD when bound to a dehydrogenase enzyme using the near infrared Fourier transform surface enhanced Raman scattering (NIR-FT-SERS) technique. This technique permits monitoring the process of combination between an NAD-dependent dehydrogenase and its coenzyme. The SERS behavior of NAD under perturbation by the enzymes showed that NAD molecules adsorbed on the electrode can be progressively ‘accommodated’ by the enzymes and this process can be governed by adjusting the applied voltage.
180° back scattering geometry was used. Laser power at the sample was ca. 100 mW. All of the spectra were acquired by 200 scans at a resolution of 2 cm − 1; none of the spectra presented have been smoothed. A specifically designed 4 ml spectroelectrochemical cell with a CaF2 window was used in the studies. The cell was equipped with a three-electrode system consisting of a Pt counter electrode, a polycrystalline gold plate (f= 5 mm) embedded in a Teflon holder as a working electrode and a saturated calomel reference electrode. The working electrode was polished carefully with emery paper (No. 1500) and cleaned ultrasonically in distilled water before use. The roughening of the electrode was achieved electrochemically by 30 oxidation-reduction cycles (orc) from + 1.1 to − 0.3 V versus SCE. at 0.5 V s − 1. The solutions for the orc performance consisted of 0.1 M KCl+ 10 mM Na2HPO4 + 1.2 mM NAD solution. The voltages for the roughening and the polarization of the electrode were controlled using an M173 potentiostat and an M175 program generator (EG&G, USA). After the roughening, an aliquot of LDH was injected into the electrolyte solution.
3. Results and discussion
3.1. SERS spectra of NAD on a gold electrode As shown in Fig. 1, a very defined SERS pattern was obtained on a roughened Au electrode with NAD solutions of 1.2 mmol l − 1. All of the SERS bands are assigned as in Table 1, based on the suggestion [45] that the Raman spectrum bands of NAD can be simply attributed to the adenine, nicotinamide, ribose and phosphate constituent molecular moieties in the structure of NAD:
2. Experimental Nicotinamide adenine dinucleotide (NAD + ) (grade 1, 100%) and beef liver glutamate dehydrogenase (GDH) were purchased from Sigma. Porcine heart lactate dehydrogenase (LDH) was purchased from Boehringer Mannheim. All of the other reagents were of analytical grade. Fourier transform surface enhanced Raman scattering measurements were carried out on a Bruker IFS 66/FRA 106FT-Raman spectrometer equipped with a diode-pumped Nd-YAG laser exciting at 1064 nm. A
To explain the SERS behavior of NAD on a gold electrode, we previously proposed an adsorption mech-
Y.-J. Xiao et al. / Journal of Electroanalytical Chemistry 465 (1999) 187–194
anism [46,47]. The SERS bands assignable to adenine and nicotinamide moieties both underwent strengthened enhancement, indicating that both are directly adsorbed onto the electrode surface. The ribose and phosphate moieties may be exposed to the solution since very weak SERS signals were observed from these moieties at a positive electrode potential. At a positive electrode potential, the adenine moiety adopted a vertical configuration in which the amino group and N7 are coordinated to the electrode surface, whereas the nicotinamide moiety is apparently adsorbed in a flat orientation with respect to the electrode surface. The substantial changes of the SERS spectra with increasingly negative potential indicate a potential-induced molecular reorientation. Specifically, the adenine moiety changes from an end-on surface interaction with the positively charged electrode dominated by interaction of the extra-ring amino group to a face-on interaction involving the ring electrons. In contrast, the nicotinamide moiety is in an end-on orientation with respect to the surface. Additionally, the changes observed with SERS during potential scanning imply that the adsorption intensity of NAD on the electrode is quite sensitive to the applied voltage. A tight chemical adsorption should occur between the NAD molecule and positively charged electrode surface. However, un-
189
Table 1 SERS bands of NAD and NAD+LDH on a gold electrode with different potentialsa 0.0 V/cm−1 NAD
−0.4 V/cm−1
NAD+LDH NAD
NAD+LDH Assignmentsb 1650vs
1628m 1600m 1571s
1583s
1554m
Amide I(LDH) N N N A
1501w 1468s
1470w 1456m 1415m
1375w 1338s 1317s
1114w 1025vs 825w 729m
1320s
1113w 1028s 828w 732w 681w 640w
1321s
1180m 1116m 1025vs 818m 731s
1322s 1239m 1194m
1028s 737m 692w
A CH2 sc (LDH) N A A A Amide (LDH) Tyr-85(LDH) R2/N R2/P N, ring R A, ring Tyr(LDH) A+N
tom-border\A+N a Abbreviations: A, adenine; N, nicotinamide; P, phosphate; R, ribose; Tyr, tyrosine; vs, very strong; s, strong; m, medium; w, weak. b Refs. [45,46,48] and the references therein.
der negative potentials, the NAD molecule mainly undertakes a physical adsorption and contacts the electrode surface loosely. This difference in the interaction of NAD with the electrode is the basis of our experiments below to attempt observation and monitoring of the interactions of NAD with dehydrogenase enzymes on the electrode by the SERS method.
3.2. LDH-induced changes in NAD SERS
Fig. 1. SERS spectra of NAD adsorbed on a gold electrode. 1.2 mmol l − 1 NAD, 10 mmol l − 1 H2PO4 and 100 mmol l − 1 KCl (pH 7.0). Applied voltages (V vs. SCE): (a), + 0.4; (b), 0.0; (c), − 0.2; (d), − 0.4; (e), − 0.6; (f), −0.8; (g), −1.0. A waiting time of ca. 5 min at each voltage was used before recording the spectra.
As mentioned above, NAD molecules are adsorbed onto the electrode surface and give strong SERS under either positive or negative potentials. However, as shown in Fig. 2, significant changes in the SERS of NAD are produced by the addition of LDH. It is known that dehydrogenase enzymes have binding sites for the NAD cofactor with convergent structural motifs. This nucleotide-binding domain is made up of four a-helices and a b-sheet of six parallel strands. The adenine moiety is contained in a hydrophobic crevice, whereas the nicotinamide group is bound in an amphipathic environment. The NAD molecule is bound within the dehydrogenase active site in an extended conformation. The experimental results show that the effect of LDH on the SERS of NAD depend upon the polarization
190
Y.-J. Xiao et al. / Journal of Electroanalytical Chemistry 465 (1999) 187–194
state of the electrode surface. With significantly positive potentials (\ 0.2 V vs. SCE), the addition of LDH produced minimal changes in the NAD SERS signals. This implies a tight chemical adsorption of NAD on the positively charged electrode. In contrast, at electrode potentials of 0 V or less, the addition of LDH results in significant changes in the SERS of NAD. Time-dependent SERS spectra under open circuit after the addition of LDH are shown in Fig. 3. Substantial decreases in the intensities of the SERS bands of the adenine moiety upon the addition of LDH are found in the 1554, 1470, 1339 and 730 cm − 1 bands. In addition, the spectra display a slight decrease in intensity for the 1028 cm − 1 band, which is assignable to the breathing vibration of the nicotinamide moiety and an upward shift of several wave numbers for the 1317 cm − 1 band. Obviously, two factors should contribute to the changes in the SERS spectrum of NAD. First is that the interaction of NAD with the enzyme should cause some separation of the cofactor from the electrode surface and consequently lead to general losses of the Raman enhancements. This phenomenon is probably the reason for the slight decrease of the 1028 cm − 1 band and the larger decrease of the 730 cm − 1 band, which have been assigned to the ring breathing vibrations of the nicotinamide and the adenine moieties respectively. Second, the partial disappearance of the
Fig. 3. Time-dependent SERS spectra of NAD pre-adsorbed on a gold electrode after addition of LDH into the electrolyte solution under open circuit conditions. (a) Before addition of LDH. (b) Ten minutes after the addition of LDH. (c) Thirty minutes after the addition of LDH. The other conditions are the same as those in Fig. 1.
Fig. 2. SERS spectra of NAD adsorbed on a gold electrode in the solution containing 10 mmol l − 1 LDH. Applied voltages/V: (a), 0.0; (b), −0.2; (c),−0.4; (d), − 0.6; (e),− 0.8; (f), −1.0. The other conditions are the same as those in Fig. 1.
adenine bands could represent a characteristic interaction between NAD and LDH. This SERS behavior, especially the decrease of the 1338 and 1470 cm − 1 bands, is in agreement with the changes of NR and RR spectra of NAD + or NADH bound to dehydrogenases in aqueous solutions [5,17,19,23]. It has been observed that the 1335 and 1483 cm − 1 bands of NAD were greatly lowered in intensity or completely vanished, and the 1308 cm − 1 band was shifted up to 1314 cm − 1, upon binding to dehydrogenases in solution [5,17,23,48]. These data were tentatively interpreted as an indication of protonation at N7 or N1 of the adenine ring [49]. As can be seen in Fig. 3(b), only a slight reduction for the SERS band near 1320 cm − 1 occurs with the addition of the enzymes. This band has been assigned to the C6 –NH2 stretching vibration of the amino group of the adenine moiety [46]. The strong band at 1320 cm − 1 is interpreted as showing the direct contact of the NH2 group to the electrode surface. This phenomenon is consistent with the structural features of NAD in all enzymatic environments [5]. The edge of the adenine ring is generally oriented toward the enzyme whereas the C6-NH2 is oriented toward the solution. This binding feature is the structural basis for the general application of affinity chromatography utilizing immobilized
Y.-J. Xiao et al. / Journal of Electroanalytical Chemistry 465 (1999) 187–194
NAD linked to a solid support through the N6 atom [5]. The potential dependence of the spectra on the presence of the enzyme reveals an extremely interesting and surprising effect. As shown in Fig. 2, upon scanning the potential in a negative direction, several new bands appear and increase in intensity up to the limit of − 0.4 V. It is worth noting that some of these bands cannot be attributed to any known vibration mode of the NAD molecule. A comparison of the SERS bands in Fig. 4(B) with that in Fig. 4(A) illustrates significant LDH-induced changes in the SERS spectra of NAD on the negatively charged electrode. The bands at 1628, 1501, 1180, 818 and 640 cm − 1 disappear completely and the 1116 cm − 1 band decreases significantly and several new bands near 1650, 1456, 1415, 1239, 1194 and 692 cm − 1 appear. In addition, the 1571 cm − 1 band has been shifted to 1583 cm − 1 and the relative intensity of the 1028 cm − 1 band is decreased significantly. It is necessary to explain the changes in the NAD SERS spectra upon addition of the enzyme. Upon addition of LDH, the band which has been assigned to a mix of CC and CO vibrations of the nicotinamide moiety shifts from 1571 to 1583 cm − 1 in the SERS spectrum compared with a shift from 1570 to 1580 cm − 1 in the NR spectrum. This wave number shift of the SERS band in Fig. 4(B) is evidence for formation of the NAD-LDH complex near the electrode surface. The 1194 cm − 1 signal is attributed to residue Tyr 84. Based on both theoretical calculations and empirical observation of bands in the spectra of a-helical polypeptides [17,50–52], the bands centered at ca. 1650 cm − 1 are
Fig. 4. Comparison of the SERS spectrum of NAD with that of NAD+ LDH on negatively charged electrodes. (a) In the absence of LDH (data from Fig. 1(d)); (b) in the presence of 10 mmol l − 1 LDH in solution (data from Fig. 2(c)). The other conditions are the same as those in Fig. 1.
191
believed to reflect the amide I vibration of a-helical segments while the 1239 cm − 1 band is attributed to amide vibration of LDH. Generally, the amide I band represents the CO stretching vibrations of the amide groups [25]. The exact frequency of the band depends on the nature of the hydrogen bonding involving the amide CO and NH moieties. The bands in this frequency region usually reflect the particular secondary structure adopted by the polypeptide chains. However, because an enzyme generally contains a variety of tertiary structure domains, a conventional vibration spectrum usually gives complex, overlapping amide bond contours representing a-helices, b-sheets, turns and non-ordered structures. Due to the inherently large widths of these overlapping component bands and the poor signal-to-noise ratio in aqueous solutions, individual features cannot normally be resolved, making any interpretation questionable, even at a qualitative level [25]. However, the appearance of well resolved SERS bands assignable to the vibration modes of the polypeptide backbone suggests a potential use of the SERS method in the study of the secondary structures of enzymes. In the light of the suggestion [41] that SERS signals of macromolecules are detected only from components in direct contact with the electrode surface, we propose that specific signals can be ascribed to the formation of the enzyme-NAD binary complex upon the electrode. The strong, sharp band at 1650 cm − 1 strongly implies a-helical segments close to the electrode surface. Because the enzyme itself does not give a SERS band in this region, it may reflect the NAD-dependent approach of the enzyme to the electrode surface. It has been suggested by X-ray analysis that an enzyme may change its conformation when the coenzyme or substrate enters the active site. Specifically, in the case of LDH, a loop (a-E and a-D involving residues 98–120) closes over the active site. The increase of the amide I signal at 1650 cm − 1 with more negative electrode potentials may implicate a surface potential-induced approach of the enzyme to the electrode and interaction with the NAD cofactor to produce the Raman enhancement. The dipole moments of the helices within the active site contribute to a positive environment [5,53] and could contribute to an electrostatic attraction of this face of the protein to the electrode surface containing the SERS-responsive NAD. When the electrode potential is made more negative than − 0.4 V, the SERS bands arising from LDH decrease and then disappear, producing a spectrum with bands characteristic of the system in the absence of enzyme, while some of the bands become even more intense than in the case of NAD alone, as in the SERS spectra of Figs. 1(h) and 2(f). This extra enhancement of the SERS bands remains to be elucidated. The
192
Y.-J. Xiao et al. / Journal of Electroanalytical Chemistry 465 (1999) 187–194
accompanying progressive disappearance of the SERS bands of LDH may indicate the desorption of the enzyme from the electrode surface; then the SERS bands completely disappear when the electrode potential is made more negative than −1.0 V, suggesting the complete desorption of the adsorbents.
3.3. A possible model for the interaction between LDH and NAD on the electrode Generally, NAD is regarded as a loosely-bound substrate rather than a prosthetic group in dehydrogenase reactions [5]. However, LDH has an ordered kinetic mechanism in which NAD must bind to the active site first, and follows with the binding of other substrates. In the first step of the enzymatic reaction, the enzyme binds the coenzyme and then closes off the active site from solvent by conformational changes. In the case of LDH, the major conformational change is a large movement of the loop region of 98 – 120 (E – D) towards the coenzyme [5,7]. The potential dependent SERS behavior of NAD in the presence of LDH demonstrates that, under certain potentials, at least a portion of the NAD molecules on the electrode can interact with the enzyme. When this occurs, the adenine moiety function appears to partition into a hydrophobic pocket of enzyme whereas the nicotinamide moiety is left partly exposed near the surface of the electrode. The reversible NAD binding to LDH is sensitive to the applied potential. Potentials near 0 V seem to favor cofactor binding. With increasingly negative potentials, the electrostatic attraction between the electrode and the positively charged nicotinamide moiety increases. The coincident strong SERS band at 1650 cm − 1 indicates a close contact of a-helical segments of the active site to the surface, likely the loop region 98 – 120 consisting of helix E–D. Gradual escape of the coenzyme from binding sites of the enzyme is reflected by the SERS behavior of NAD when the potential is made more negative than − 0.4 V. In the potential region of − 0.4 to − 1.0 V, the SERS spectra show typical adsorption features of NAD on an electrode in the absence of enzyme. After this, when the potential is further moved more negative than − 1.0 V, the irreversible disappearance of all the SERS bands indicates the desorption of the coenzyme from the electrode surface. To determine whether the observed SERS behavior observed with NAD and LDH can be generalized to other NAD-dependent dehydrogenases, a second set of experiments was performed on glutamate dehydrogenase (GDH). It is known [54] that GDH is a type-B dehydrogenase, i.e. when the NAD molecule is reduced catalyzed by glutamate dehydrogenase, the enzyme transfers the hydride ion from its substrate to the B side of the nicotinamide ring. In contrast, LDH is a type A
dehydrogenase, i.e. the hydride ion is transferred to the A side of the nicotinamide ring. The SERS of NAD perturbed by GDH is also potential dependent and similar to the experiments with LDH. No influence of the enzyme on the SERS of NAD was observed with positive electrode potentials. However, with negative shifts of the electrode potential, two bands due to the enzyme appeared at 1665 and 1230 cm − 1. Meanwhile, the ring vibration band of the nicotinamide moiety of NAD moved from 1571 to 1584 cm − 1, consistent with binding with GDH. In addition, the SERS of NAD upon binding with GDH show some features significantly different from those induced by LDH under negative potentials. Particularly, the amide I and amide bands of GDH appear at 1665 and 1230 cm − 1, instead of 1650 and 1240 cm − 1. This is most probably due to the different structures of active sites in the LDH and GDH enzymes and raises the possibility that the FT-SERS technique may eventually be used to recognize different structural elements within an enzyme. From an understanding of the general spectroscopic properties of proteins [25,50,51], the frequency of an amide I mode, arising from polypeptide backbone amide CO stretching motions, is dependent on its hydrogen bonding environment and hence its secondary structure. Specifically, a frequency near 1650 cm − 1 is suggested to be related to an a-helical structure, whereas frequencies near 1665 cm − 1 are typically found in turn motifs [25,55–60]. The amide regions, due to polypeptide backbone N–H in-plane bending and C–N stretching motions, is also sensitive to secondary structures. Therefore, differences between the SERS spectra of NAD-LDH and those of NADGDH are reasonably attributed to the different secondary structures of binding sites in the two enzymes. The prominent structures of the LDH active site interacting with the coenzyme appear to be a-helical regions, whereas turns appear to be the main elements of the GDH active site interacting with NAD coenzyme on a negatively charged electrode. Because turns are often found near the surface of a protein [54], the SERS bands may indicate a greater interaction of these structural elements with the surface of the enzyme. However, we cannot explain the development of a strong band at 1415 cm − 1 with increasing negative potential. This band has not been reported previously in either normal Raman or SERS of NAD in solution or bound to a enzyme. It is interesting to note that the reduced coenzyme, NADH, shows a strong band near 1422 cm − 1 in solution and 1416 cm − 1 when bound to dehydrogenases [5,17,19,23]. It therefore seems possible that the NAD molecules are reduced by the enzyme to NADH on the negatively charged electrode. Unfortunately, this proposition was not supported by the electrochemical data. In the potential range 0.0 to −1.0 V, no reduction peaks in the cyclic voltammograms of
Y.-J. Xiao et al. / Journal of Electroanalytical Chemistry 465 (1999) 187–194
either NAD or NAD-LDH systems were observed. So, the exact explanation of the mechanism of the reactions and the assignment of the 1415 cm − 1 band still remain to be elucidated by further experimental and theoretical studies. In addition, the relative intensity of the 1028 cm − 1 band was reduced significantly by interaction with the dehydrogenase under conditions of negative potential. This suggests a partial reduction of the nicotinamide ring. Only the oxidized nicotinamide moity of NAD shows an aromatic nature and gives a ring breathing motion near 1030 cm − 1. In contrast, reduced nicotinamide is not aromatic and produces no band near 1030 cm − 1. This band is quite insensitive to its molecular environment and all Raman spectra of NAD, whether in solution [45], binding with a dehydrogenase [17], or adsorbed on a metal surface [43,44,46,47], give the ring breathing vibration mode near 1030 cm − 1. Thus, it is reasonable for the position of the 1028 cm − 1 band to be unchanged in the potential scanning. Obviously, in this case, an unfavorable orientation of the nicotinamide moiety may also cause the decrease of the relative intensity of the 1028 cm − 1 band. The absence of the 1680 cm − 1 band, which is normally apparent in spectra of NADH in solution, is probably due to the unfavorable orientation on the electrode surface of the reduced nicotinamide carboxamide moiety. Obviously, further systematic studies will be required to validate the ability of SERS spectra to provide structural information about the structure and reaction mechanisms of NAD-dependent dehydrogenases and other enzymes, which will complement the existing methods currently employed in this active area of research.
Acknowledgements The research was supported by the National Science Foundation of China (No. 29675001).
References [1] H. Eklund, C. Branden, in: D. Dolphin, O. Avramovic, R. Poulson Jr. (Eds.), Pyridine Nucleotides Coenzymes: Part A, Chapter 4, Wiley, New York, 1987, pp. 51–98. [2] H. Sund, H. Theorell, Enzymes 7 (1963) 25. [3] A.M. Gronenborn, G.M. Clore, J. Mol. Biol. 157 (1982) 155. [4] A.G.W. Leslie, A.J. Wonacott, J. Mol. Biol. 178 (1984) 743. [5] J.C. Austin, C.W. Wharton, R.E. Hester, Biochemistry 28 (1989) 1533. [6] U.M. Grau, W.E. Trommer, M.G. Rossmann, J. Mol. Biol. 151 (1981) 289. [7] J.L. White, M.L. Hacker, M. Buehner, M.J. Adams, G.C. Ford, P.J. Lentz Jr., I.E. Smiley, S.J. Steindel, M.G. Rossmann, J. Mol. Biol. 102 (1976) 759.
193
[8] H. Eklund, J.-P. Samama, L. Wallen, C.-I. Branden, A. Akeson, T.A. Jones, J. Mol. Biol. 146 (1981) 561. [9] H. Eklund, J.-P. Samama, T.A. Jones, Biochemistry 23 (1984) 5982. [10] H. Eklund, C.-I. Branden, Pyridine Nucleotides, Wiley, New York, 1986. [11] A. Bardea, E. Katz, A.J. Buckmann, I. Willner, J. Am. Chem. Soc. 119 (1997) 9114. [12] P. Gros, C. Zaborosch, H.G. Schlegel, A. Bergel, J. Electroanal. Chem. 405 (1996) 189. [13] J.J. Holbrook, A. Lijas, S. J. Steindel, M.G. Rossmann, in: P.D. Boyer (Ed.), The Enzymes, 3rd edition, Academic Press, New York, 1975, p. 191. [14] A.R. Clarke, D.B. Wigley, W.N. Chia, J.J. Holbrook, Nature 324 (1986) 699. [15] D.M. Parker, J.J. Holbrook, in: H. Sund (Ed.), Pyridine Nucleotide-Dependent Dehydrogenases, de Gruyter, Berlin, 1977, pp. 485 – 502. [16] J.J. Birktoft, R.T. Fernley, R.A. Bradshaw, L.J. Banaszak, Proc. Natl. Acad. Sci. USA 79 (1982) 6166. [17] H. Deng, J. Zheng, J. Burgner, D. Sloan, R. Callender, Biochemistry 28 (1989) 1525. [18] H. Deng, J. Zheng, J. Burgner, R. Callender, J. Phys. Chem. 93 (1989) 4710. [19] H. Deng, K.T. Yue, C. Martin, K.W. Rhee, D. Sloan, R. Callender, Biochemistry 26 (1987) 4776. [20] H. Deng, J. Burgner, R. Callender, J. Am. Chem. Soc. 114 (1992) 7997. [21] H. Deng, J. Zheng, J. Burgner, R. Callender, Proc. Natl. Acad. Sci. USA 86 (1989) 4484. [22] R. Callender, H. Deng, Annu. Rev. Biophys. Biomol. Struct. 23 (1994) 215. [23] H. Deng, J. Burgner, R. Callender, Biochemistry 30 (1991) 8804. [24] L. Young, C.B. Post, Biochemistry 35 (1996) 15129. [25] W.K. Surewicz, H.H. Mantsch, Biochim. Biophys. Acta 952 (1988) 115. [26] A. Elliot, E.J. Ambrose, Nature 165 (1950) 921. [27] H. Susi, in: S.N. Timasheff, G.D. Fasman (Eds.), Structure and Stability of Biological Macromolecules, Marcel Dekker, New York, 1969, pp. 575 – 663. [28] H. Susi, Methods Enzymol. 26 (1972) 445. [29] F.S. Parker, Applications of Infrared, Raman and Resonance Raman Spectroscopy in Biochemistry, Plenum Press, New York, 1983. [30] W.-Z. He, W.R. Newell, P.I. Haris, D. Chapman, J. Barber, Biochemistry 30 (1991) 4552. [31] R.A. Copeland, T.G. Spiro, Biochemistry 26 (1987) 2134. [32] T.M. Cotton, J.-H. Kim, G.D. Chumanov, J. Raman Spectrosc. 22 (1991) 729. [33] X. Dou, T. Takama, Y. Yamaguchi, H. Yamamoto, Anal. Chem. 69 (1997) 1492. [34] T.M. Cotton, in: R.J.H. Clark, R.E. Hester (Eds.), Spectroscopy of Surfaces, Wiley, New York, 1988, pp. 91 – 153. [35] G.D. Chumanov, R.G. Efremov, I.R. Nabiev, J. Raman Spectrosc. 21 (1990) 43. [36] E. Grabbe, R.P. Buck, J. Am. Chem. Soc. 111 (1989) 8362. [37] A.M. Ahern, R.L. Garrell, Langmuir 7 (1991) 254. [38] R.A. Copeland, S.P.A. Fodor, T.G. Spiro, J. Am. Chem. Soc. 106 (1984) 3872. [39] N.-S. Lee, Y.-Z. Hsieh, M.D. Morris, L.M. Schopfer, J. Am. Chem. Soc. 109 (1987) 1358. [40] R.E. Holt, T.M. Cotton, J. Am. Chem. Soc. 111 (1989) 2815. [41] E. Koglin, J.-M. Sequaris, Top. Curr. Chem. 134 (1986) 39. [42] J. Hu, R.S. Sheng, Z.S. Xu, Y.-E. Zeng, Spectrochim. Acta Part A 51 (1995) 1087. [43] J.C. Austin, R.E. Hester, J. Chem. Soc. Faraday Trans. I 85 (1989) 1159.
194
Y.-J. Xiao et al. / Journal of Electroanalytical Chemistry 465 (1999) 187–194
[44] O. Siiman, R. Rivellini, R. Patel, Inorg. Chem. 27 (1988) 3940. [45] K.T. Yue, C.L. Martin, D. Chen, P. Nelson, D.L. Sloan, R. Callender, Biochemistry 25 (1986) 4941. [46] Y.-J. Xiao, J.P. Markwell, Langmuir 13 (1997) 7068. [47] Y.-J. Xiao, T. Wang, X.-Q. Xang, X.-X. Gao, J. Electroanal. Chem. 433 (1997) 49. [48] D. Chen, K.T. Yue, C. Martin, K.W. Rhee, D. Sloan, R. Callender, Biochemistry 26 (1987) 4776. [49] C. Nadolny, G. Zundel, J. Mol. Struct. 385 (1996) 81. [50] P.R. Carey, Biochemical Applications of Raman and Resonance Raman Spectroscopy, Academic Press, New York, 1982. [51] J. Ryttersgaard, B.D. Larsen, A. Holm, D.H. Christensen, O.F. Nielsen, Spectrochim. Acta Part A 53 (1997) 91. [52] S. Krimm, J. Bandekar, Adv. Protein Chem. 38 (1986) 181.
.
[53] W.G.J. Hol, P.T. van Duijnen, H.J.C. Berendsen, Nature (London) 273 (1978) 443. [54] A. L. Lehninger, D.L. Nelson, M.M. Cox, Principles of Biochemistry, 2nd edition, Worth, New York, 1993, pp. 390–392. [55] H. Susi, M. Byler, Methods Enzymol. 130 (1986) 290. [56] M. Byler, H. Susi, Biopolymers 25 (1986) 469. [57] P.I. Harris, D.C. Lee, D. Chapman, Biochim. Biophys. Acta 874 (1986) 255. [58] P.W. Yang, H.H. Mantsch, J.L.R. Arrondo, I. Saint-Girons, Y. Guillou, G.N. Cohen, O. Barzu, Biochemistry 26 (1987) 2706. [59] W.K. Surewicz, M.A. Moscarello, H.H. Mantsch, Biochemistry 26 (1987) 3881. [60] J.L.R. Arrondo, H.H. Mantsch, N. Mullner, S. Pikula, A. Martonosi, J. Biol. Chem. 262 (1987) 9037.
In-situ monitoring of the interaction of lactate dehydrogenase with NAD on a gold electrode by FT-SERS Yi-Jin Xiao a,*, Xiao-Xia Gao a, John Markwell b b
a Department of Chemistry, Peking Uni6ersity, Beijing 100871, PR China Department of Biochemistry, Uni6ersity of Nebraska-Lincoln, Lincoln, NE 68588, USA
Received 7 September 1998; received in revised form 2 February 1999
Abstract The FT-SERS method was used to observe directly and monitor the combination of dehydrogenases with NAD pre-adsorbed onto the surface of a roughened gold electrode. At negative electrode potentials, the SERS spectra of NAD exhibit significant changes upon the addition of lactate dehydrogenase. The enzyme active site may approach the electrode closely enough to produce several strong bands attributable to backbone peptide. The experimental results may be explained by binding of the NAD adenine moiety in a hydrohobic region of the enzyme while the nicotinamide group is left exposed near the surface of the electrode. This competitive binding state appears to be strongly dependent on the applied electrode potentials. Comparative experiments were carried out with lactate dehydrogenase and glutamate dehydrogenase. The SERS of NAD upon binding with lactate dehydrogenase show some features significantly different from those in the case of glutamate dehydrogenase. This difference may result from the different architectures within binding sites of type-A and type-B dehydrogenases. © 1999 Elsevier Science S.A. All rights reserved. Keywords: Gold electrode; Lactate dehydrogenase; FT-SERS; NAD
1. Introduction The requisite step for an enzymatic redox process is the binding of substrates within a specific intraprotein environment which stimulates reactivity, reduces the transition state barrier between substrates and products, and defines the specificity of the enzymatic reactions. As oxidoreductase enzymes important in diverse metabolic pathways, NAD-dependent dehydrogenases have been extensively studied by many methods [1–24]. Whereas much of our structural knowledge of this class of enzymes is derived from X-ray crystallographic measurement, the development of methods to study the behavior and properties of these enzymes in solution would be highly desirable; techniques such as infrared * Corresponding author. E-mail address: yjxiaopku.edu.cn (Y.-J. Xiao)
spectroscopy, capable of making measurements in solution, have been suggested to provides insight into molecular structures of proteins in solution [25–30]. However, the general applicability of infrared spectroscopy is hampered by the practical experimental difficulties in obtaining spectra of proteins in H2O, which absorbs strongly in the same spectral region as the conformation-sensitive amide bands in the polypeptide background. Thus, the conformational changes in tertiary structure induced by interaction with the NAD cofactor would be largely masked by the aqueous solvent. A complicated computational procedure would be needed for deconvolution of the complex amide absorption bands into their constituent components [25]. Recent reports [5,17–23] have employed normal Raman (NR) and resonance Raman (RR) spectroscopy to gain additional insights into the molecular structure of the NAD coenzyme upon binding with enzyme
0022-0728/99/$ - see front matter © 1999 Elsevier Science S.A. All rights reserved. PII: S 0 0 2 2 - 0 7 2 8 ( 9 9 ) 0 0 0 9 3 - 5
188
Y.-J. Xiao et al. / Journal of Electroanalytical Chemistry 465 (1999) 187–194
proteins. However, the absence of suitable chromophores from many enzymes makes it difficult to get RR spectra. In addition, the RR vibrational bands arising from the polypeptide backbone of the enzyme, such as the amide I band, are comparatively weak and do not provide much information about secondary or tertiary structure [5,31]. With normal Raman spectra, careful difference spectra are required to reveal the spectrum of the bound coenzyme. Many researchers have demonstrated the potential of SERS techniques for obtaining structure-function information about enzymes. Strong SERS signals from the surface-exposed aromatic residues of proteins have indeed been obtained [32 – 42]. However, it is difficult to get any SERS signals originating from the polypeptide backbone of the enzyme, which would yield information about protein structure. As has been reviewed by Cotton et al. [32], the screening by aromatic groups on the protein surface interferes with surface enhancement of the amide I bands. It has been proposed that structural changes of NAD induced by active-site binding to a dehydrogenase should theoretically be monitored by the SERS technique [43,44]. Unfortunately, no such enzyme-cofactor SERS signal was obtained [43]. The lack of significant surface enhancement was attributed to NAD being buried deeply in the dehydrogenase protein and implied that use of this technique to track the combination of enzymes, coenzyme and substrate in solution was problematic. In the present work, we have designed an original experimental procedure which provides high quality spectra attributable to NAD when bound to a dehydrogenase enzyme using the near infrared Fourier transform surface enhanced Raman scattering (NIR-FT-SERS) technique. This technique permits monitoring the process of combination between an NAD-dependent dehydrogenase and its coenzyme. The SERS behavior of NAD under perturbation by the enzymes showed that NAD molecules adsorbed on the electrode can be progressively ‘accommodated’ by the enzymes and this process can be governed by adjusting the applied voltage.
180° back scattering geometry was used. Laser power at the sample was ca. 100 mW. All of the spectra were acquired by 200 scans at a resolution of 2 cm − 1; none of the spectra presented have been smoothed. A specifically designed 4 ml spectroelectrochemical cell with a CaF2 window was used in the studies. The cell was equipped with a three-electrode system consisting of a Pt counter electrode, a polycrystalline gold plate (f= 5 mm) embedded in a Teflon holder as a working electrode and a saturated calomel reference electrode. The working electrode was polished carefully with emery paper (No. 1500) and cleaned ultrasonically in distilled water before use. The roughening of the electrode was achieved electrochemically by 30 oxidation-reduction cycles (orc) from + 1.1 to − 0.3 V versus SCE. at 0.5 V s − 1. The solutions for the orc performance consisted of 0.1 M KCl+ 10 mM Na2HPO4 + 1.2 mM NAD solution. The voltages for the roughening and the polarization of the electrode were controlled using an M173 potentiostat and an M175 program generator (EG&G, USA). After the roughening, an aliquot of LDH was injected into the electrolyte solution.
3. Results and discussion
3.1. SERS spectra of NAD on a gold electrode As shown in Fig. 1, a very defined SERS pattern was obtained on a roughened Au electrode with NAD solutions of 1.2 mmol l − 1. All of the SERS bands are assigned as in Table 1, based on the suggestion [45] that the Raman spectrum bands of NAD can be simply attributed to the adenine, nicotinamide, ribose and phosphate constituent molecular moieties in the structure of NAD:
2. Experimental Nicotinamide adenine dinucleotide (NAD + ) (grade 1, 100%) and beef liver glutamate dehydrogenase (GDH) were purchased from Sigma. Porcine heart lactate dehydrogenase (LDH) was purchased from Boehringer Mannheim. All of the other reagents were of analytical grade. Fourier transform surface enhanced Raman scattering measurements were carried out on a Bruker IFS 66/FRA 106FT-Raman spectrometer equipped with a diode-pumped Nd-YAG laser exciting at 1064 nm. A
To explain the SERS behavior of NAD on a gold electrode, we previously proposed an adsorption mech-
Y.-J. Xiao et al. / Journal of Electroanalytical Chemistry 465 (1999) 187–194
anism [46,47]. The SERS bands assignable to adenine and nicotinamide moieties both underwent strengthened enhancement, indicating that both are directly adsorbed onto the electrode surface. The ribose and phosphate moieties may be exposed to the solution since very weak SERS signals were observed from these moieties at a positive electrode potential. At a positive electrode potential, the adenine moiety adopted a vertical configuration in which the amino group and N7 are coordinated to the electrode surface, whereas the nicotinamide moiety is apparently adsorbed in a flat orientation with respect to the electrode surface. The substantial changes of the SERS spectra with increasingly negative potential indicate a potential-induced molecular reorientation. Specifically, the adenine moiety changes from an end-on surface interaction with the positively charged electrode dominated by interaction of the extra-ring amino group to a face-on interaction involving the ring electrons. In contrast, the nicotinamide moiety is in an end-on orientation with respect to the surface. Additionally, the changes observed with SERS during potential scanning imply that the adsorption intensity of NAD on the electrode is quite sensitive to the applied voltage. A tight chemical adsorption should occur between the NAD molecule and positively charged electrode surface. However, un-
189
Table 1 SERS bands of NAD and NAD+LDH on a gold electrode with different potentialsa 0.0 V/cm−1 NAD
−0.4 V/cm−1
NAD+LDH NAD
NAD+LDH Assignmentsb 1650vs
1628m 1600m 1571s
1583s
1554m
Amide I(LDH) N N N A
1501w 1468s
1470w 1456m 1415m
1375w 1338s 1317s
1114w 1025vs 825w 729m
1320s
1113w 1028s 828w 732w 681w 640w
1321s
1180m 1116m 1025vs 818m 731s
1322s 1239m 1194m
1028s 737m 692w
A CH2 sc (LDH) N A A A Amide (LDH) Tyr-85(LDH) R2/N R2/P N, ring R A, ring Tyr(LDH) A+N
tom-border\A+N a Abbreviations: A, adenine; N, nicotinamide; P, phosphate; R, ribose; Tyr, tyrosine; vs, very strong; s, strong; m, medium; w, weak. b Refs. [45,46,48] and the references therein.
der negative potentials, the NAD molecule mainly undertakes a physical adsorption and contacts the electrode surface loosely. This difference in the interaction of NAD with the electrode is the basis of our experiments below to attempt observation and monitoring of the interactions of NAD with dehydrogenase enzymes on the electrode by the SERS method.
3.2. LDH-induced changes in NAD SERS
Fig. 1. SERS spectra of NAD adsorbed on a gold electrode. 1.2 mmol l − 1 NAD, 10 mmol l − 1 H2PO4 and 100 mmol l − 1 KCl (pH 7.0). Applied voltages (V vs. SCE): (a), + 0.4; (b), 0.0; (c), − 0.2; (d), − 0.4; (e), − 0.6; (f), −0.8; (g), −1.0. A waiting time of ca. 5 min at each voltage was used before recording the spectra.
As mentioned above, NAD molecules are adsorbed onto the electrode surface and give strong SERS under either positive or negative potentials. However, as shown in Fig. 2, significant changes in the SERS of NAD are produced by the addition of LDH. It is known that dehydrogenase enzymes have binding sites for the NAD cofactor with convergent structural motifs. This nucleotide-binding domain is made up of four a-helices and a b-sheet of six parallel strands. The adenine moiety is contained in a hydrophobic crevice, whereas the nicotinamide group is bound in an amphipathic environment. The NAD molecule is bound within the dehydrogenase active site in an extended conformation. The experimental results show that the effect of LDH on the SERS of NAD depend upon the polarization
190
Y.-J. Xiao et al. / Journal of Electroanalytical Chemistry 465 (1999) 187–194
state of the electrode surface. With significantly positive potentials (\ 0.2 V vs. SCE), the addition of LDH produced minimal changes in the NAD SERS signals. This implies a tight chemical adsorption of NAD on the positively charged electrode. In contrast, at electrode potentials of 0 V or less, the addition of LDH results in significant changes in the SERS of NAD. Time-dependent SERS spectra under open circuit after the addition of LDH are shown in Fig. 3. Substantial decreases in the intensities of the SERS bands of the adenine moiety upon the addition of LDH are found in the 1554, 1470, 1339 and 730 cm − 1 bands. In addition, the spectra display a slight decrease in intensity for the 1028 cm − 1 band, which is assignable to the breathing vibration of the nicotinamide moiety and an upward shift of several wave numbers for the 1317 cm − 1 band. Obviously, two factors should contribute to the changes in the SERS spectrum of NAD. First is that the interaction of NAD with the enzyme should cause some separation of the cofactor from the electrode surface and consequently lead to general losses of the Raman enhancements. This phenomenon is probably the reason for the slight decrease of the 1028 cm − 1 band and the larger decrease of the 730 cm − 1 band, which have been assigned to the ring breathing vibrations of the nicotinamide and the adenine moieties respectively. Second, the partial disappearance of the
Fig. 3. Time-dependent SERS spectra of NAD pre-adsorbed on a gold electrode after addition of LDH into the electrolyte solution under open circuit conditions. (a) Before addition of LDH. (b) Ten minutes after the addition of LDH. (c) Thirty minutes after the addition of LDH. The other conditions are the same as those in Fig. 1.
Fig. 2. SERS spectra of NAD adsorbed on a gold electrode in the solution containing 10 mmol l − 1 LDH. Applied voltages/V: (a), 0.0; (b), −0.2; (c),−0.4; (d), − 0.6; (e),− 0.8; (f), −1.0. The other conditions are the same as those in Fig. 1.
adenine bands could represent a characteristic interaction between NAD and LDH. This SERS behavior, especially the decrease of the 1338 and 1470 cm − 1 bands, is in agreement with the changes of NR and RR spectra of NAD + or NADH bound to dehydrogenases in aqueous solutions [5,17,19,23]. It has been observed that the 1335 and 1483 cm − 1 bands of NAD were greatly lowered in intensity or completely vanished, and the 1308 cm − 1 band was shifted up to 1314 cm − 1, upon binding to dehydrogenases in solution [5,17,23,48]. These data were tentatively interpreted as an indication of protonation at N7 or N1 of the adenine ring [49]. As can be seen in Fig. 3(b), only a slight reduction for the SERS band near 1320 cm − 1 occurs with the addition of the enzymes. This band has been assigned to the C6 –NH2 stretching vibration of the amino group of the adenine moiety [46]. The strong band at 1320 cm − 1 is interpreted as showing the direct contact of the NH2 group to the electrode surface. This phenomenon is consistent with the structural features of NAD in all enzymatic environments [5]. The edge of the adenine ring is generally oriented toward the enzyme whereas the C6-NH2 is oriented toward the solution. This binding feature is the structural basis for the general application of affinity chromatography utilizing immobilized
Y.-J. Xiao et al. / Journal of Electroanalytical Chemistry 465 (1999) 187–194
NAD linked to a solid support through the N6 atom [5]. The potential dependence of the spectra on the presence of the enzyme reveals an extremely interesting and surprising effect. As shown in Fig. 2, upon scanning the potential in a negative direction, several new bands appear and increase in intensity up to the limit of − 0.4 V. It is worth noting that some of these bands cannot be attributed to any known vibration mode of the NAD molecule. A comparison of the SERS bands in Fig. 4(B) with that in Fig. 4(A) illustrates significant LDH-induced changes in the SERS spectra of NAD on the negatively charged electrode. The bands at 1628, 1501, 1180, 818 and 640 cm − 1 disappear completely and the 1116 cm − 1 band decreases significantly and several new bands near 1650, 1456, 1415, 1239, 1194 and 692 cm − 1 appear. In addition, the 1571 cm − 1 band has been shifted to 1583 cm − 1 and the relative intensity of the 1028 cm − 1 band is decreased significantly. It is necessary to explain the changes in the NAD SERS spectra upon addition of the enzyme. Upon addition of LDH, the band which has been assigned to a mix of CC and CO vibrations of the nicotinamide moiety shifts from 1571 to 1583 cm − 1 in the SERS spectrum compared with a shift from 1570 to 1580 cm − 1 in the NR spectrum. This wave number shift of the SERS band in Fig. 4(B) is evidence for formation of the NAD-LDH complex near the electrode surface. The 1194 cm − 1 signal is attributed to residue Tyr 84. Based on both theoretical calculations and empirical observation of bands in the spectra of a-helical polypeptides [17,50–52], the bands centered at ca. 1650 cm − 1 are
Fig. 4. Comparison of the SERS spectrum of NAD with that of NAD+ LDH on negatively charged electrodes. (a) In the absence of LDH (data from Fig. 1(d)); (b) in the presence of 10 mmol l − 1 LDH in solution (data from Fig. 2(c)). The other conditions are the same as those in Fig. 1.
191
believed to reflect the amide I vibration of a-helical segments while the 1239 cm − 1 band is attributed to amide vibration of LDH. Generally, the amide I band represents the CO stretching vibrations of the amide groups [25]. The exact frequency of the band depends on the nature of the hydrogen bonding involving the amide CO and NH moieties. The bands in this frequency region usually reflect the particular secondary structure adopted by the polypeptide chains. However, because an enzyme generally contains a variety of tertiary structure domains, a conventional vibration spectrum usually gives complex, overlapping amide bond contours representing a-helices, b-sheets, turns and non-ordered structures. Due to the inherently large widths of these overlapping component bands and the poor signal-to-noise ratio in aqueous solutions, individual features cannot normally be resolved, making any interpretation questionable, even at a qualitative level [25]. However, the appearance of well resolved SERS bands assignable to the vibration modes of the polypeptide backbone suggests a potential use of the SERS method in the study of the secondary structures of enzymes. In the light of the suggestion [41] that SERS signals of macromolecules are detected only from components in direct contact with the electrode surface, we propose that specific signals can be ascribed to the formation of the enzyme-NAD binary complex upon the electrode. The strong, sharp band at 1650 cm − 1 strongly implies a-helical segments close to the electrode surface. Because the enzyme itself does not give a SERS band in this region, it may reflect the NAD-dependent approach of the enzyme to the electrode surface. It has been suggested by X-ray analysis that an enzyme may change its conformation when the coenzyme or substrate enters the active site. Specifically, in the case of LDH, a loop (a-E and a-D involving residues 98–120) closes over the active site. The increase of the amide I signal at 1650 cm − 1 with more negative electrode potentials may implicate a surface potential-induced approach of the enzyme to the electrode and interaction with the NAD cofactor to produce the Raman enhancement. The dipole moments of the helices within the active site contribute to a positive environment [5,53] and could contribute to an electrostatic attraction of this face of the protein to the electrode surface containing the SERS-responsive NAD. When the electrode potential is made more negative than − 0.4 V, the SERS bands arising from LDH decrease and then disappear, producing a spectrum with bands characteristic of the system in the absence of enzyme, while some of the bands become even more intense than in the case of NAD alone, as in the SERS spectra of Figs. 1(h) and 2(f). This extra enhancement of the SERS bands remains to be elucidated. The
192
Y.-J. Xiao et al. / Journal of Electroanalytical Chemistry 465 (1999) 187–194
accompanying progressive disappearance of the SERS bands of LDH may indicate the desorption of the enzyme from the electrode surface; then the SERS bands completely disappear when the electrode potential is made more negative than −1.0 V, suggesting the complete desorption of the adsorbents.
3.3. A possible model for the interaction between LDH and NAD on the electrode Generally, NAD is regarded as a loosely-bound substrate rather than a prosthetic group in dehydrogenase reactions [5]. However, LDH has an ordered kinetic mechanism in which NAD must bind to the active site first, and follows with the binding of other substrates. In the first step of the enzymatic reaction, the enzyme binds the coenzyme and then closes off the active site from solvent by conformational changes. In the case of LDH, the major conformational change is a large movement of the loop region of 98 – 120 (E – D) towards the coenzyme [5,7]. The potential dependent SERS behavior of NAD in the presence of LDH demonstrates that, under certain potentials, at least a portion of the NAD molecules on the electrode can interact with the enzyme. When this occurs, the adenine moiety function appears to partition into a hydrophobic pocket of enzyme whereas the nicotinamide moiety is left partly exposed near the surface of the electrode. The reversible NAD binding to LDH is sensitive to the applied potential. Potentials near 0 V seem to favor cofactor binding. With increasingly negative potentials, the electrostatic attraction between the electrode and the positively charged nicotinamide moiety increases. The coincident strong SERS band at 1650 cm − 1 indicates a close contact of a-helical segments of the active site to the surface, likely the loop region 98 – 120 consisting of helix E–D. Gradual escape of the coenzyme from binding sites of the enzyme is reflected by the SERS behavior of NAD when the potential is made more negative than − 0.4 V. In the potential region of − 0.4 to − 1.0 V, the SERS spectra show typical adsorption features of NAD on an electrode in the absence of enzyme. After this, when the potential is further moved more negative than − 1.0 V, the irreversible disappearance of all the SERS bands indicates the desorption of the coenzyme from the electrode surface. To determine whether the observed SERS behavior observed with NAD and LDH can be generalized to other NAD-dependent dehydrogenases, a second set of experiments was performed on glutamate dehydrogenase (GDH). It is known [54] that GDH is a type-B dehydrogenase, i.e. when the NAD molecule is reduced catalyzed by glutamate dehydrogenase, the enzyme transfers the hydride ion from its substrate to the B side of the nicotinamide ring. In contrast, LDH is a type A
dehydrogenase, i.e. the hydride ion is transferred to the A side of the nicotinamide ring. The SERS of NAD perturbed by GDH is also potential dependent and similar to the experiments with LDH. No influence of the enzyme on the SERS of NAD was observed with positive electrode potentials. However, with negative shifts of the electrode potential, two bands due to the enzyme appeared at 1665 and 1230 cm − 1. Meanwhile, the ring vibration band of the nicotinamide moiety of NAD moved from 1571 to 1584 cm − 1, consistent with binding with GDH. In addition, the SERS of NAD upon binding with GDH show some features significantly different from those induced by LDH under negative potentials. Particularly, the amide I and amide bands of GDH appear at 1665 and 1230 cm − 1, instead of 1650 and 1240 cm − 1. This is most probably due to the different structures of active sites in the LDH and GDH enzymes and raises the possibility that the FT-SERS technique may eventually be used to recognize different structural elements within an enzyme. From an understanding of the general spectroscopic properties of proteins [25,50,51], the frequency of an amide I mode, arising from polypeptide backbone amide CO stretching motions, is dependent on its hydrogen bonding environment and hence its secondary structure. Specifically, a frequency near 1650 cm − 1 is suggested to be related to an a-helical structure, whereas frequencies near 1665 cm − 1 are typically found in turn motifs [25,55–60]. The amide regions, due to polypeptide backbone N–H in-plane bending and C–N stretching motions, is also sensitive to secondary structures. Therefore, differences between the SERS spectra of NAD-LDH and those of NADGDH are reasonably attributed to the different secondary structures of binding sites in the two enzymes. The prominent structures of the LDH active site interacting with the coenzyme appear to be a-helical regions, whereas turns appear to be the main elements of the GDH active site interacting with NAD coenzyme on a negatively charged electrode. Because turns are often found near the surface of a protein [54], the SERS bands may indicate a greater interaction of these structural elements with the surface of the enzyme. However, we cannot explain the development of a strong band at 1415 cm − 1 with increasing negative potential. This band has not been reported previously in either normal Raman or SERS of NAD in solution or bound to a enzyme. It is interesting to note that the reduced coenzyme, NADH, shows a strong band near 1422 cm − 1 in solution and 1416 cm − 1 when bound to dehydrogenases [5,17,19,23]. It therefore seems possible that the NAD molecules are reduced by the enzyme to NADH on the negatively charged electrode. Unfortunately, this proposition was not supported by the electrochemical data. In the potential range 0.0 to −1.0 V, no reduction peaks in the cyclic voltammograms of
Y.-J. Xiao et al. / Journal of Electroanalytical Chemistry 465 (1999) 187–194
either NAD or NAD-LDH systems were observed. So, the exact explanation of the mechanism of the reactions and the assignment of the 1415 cm − 1 band still remain to be elucidated by further experimental and theoretical studies. In addition, the relative intensity of the 1028 cm − 1 band was reduced significantly by interaction with the dehydrogenase under conditions of negative potential. This suggests a partial reduction of the nicotinamide ring. Only the oxidized nicotinamide moity of NAD shows an aromatic nature and gives a ring breathing motion near 1030 cm − 1. In contrast, reduced nicotinamide is not aromatic and produces no band near 1030 cm − 1. This band is quite insensitive to its molecular environment and all Raman spectra of NAD, whether in solution [45], binding with a dehydrogenase [17], or adsorbed on a metal surface [43,44,46,47], give the ring breathing vibration mode near 1030 cm − 1. Thus, it is reasonable for the position of the 1028 cm − 1 band to be unchanged in the potential scanning. Obviously, in this case, an unfavorable orientation of the nicotinamide moiety may also cause the decrease of the relative intensity of the 1028 cm − 1 band. The absence of the 1680 cm − 1 band, which is normally apparent in spectra of NADH in solution, is probably due to the unfavorable orientation on the electrode surface of the reduced nicotinamide carboxamide moiety. Obviously, further systematic studies will be required to validate the ability of SERS spectra to provide structural information about the structure and reaction mechanisms of NAD-dependent dehydrogenases and other enzymes, which will complement the existing methods currently employed in this active area of research.
Acknowledgements The research was supported by the National Science Foundation of China (No. 29675001).
References [1] H. Eklund, C. Branden, in: D. Dolphin, O. Avramovic, R. Poulson Jr. (Eds.), Pyridine Nucleotides Coenzymes: Part A, Chapter 4, Wiley, New York, 1987, pp. 51–98. [2] H. Sund, H. Theorell, Enzymes 7 (1963) 25. [3] A.M. Gronenborn, G.M. Clore, J. Mol. Biol. 157 (1982) 155. [4] A.G.W. Leslie, A.J. Wonacott, J. Mol. Biol. 178 (1984) 743. [5] J.C. Austin, C.W. Wharton, R.E. Hester, Biochemistry 28 (1989) 1533. [6] U.M. Grau, W.E. Trommer, M.G. Rossmann, J. Mol. Biol. 151 (1981) 289. [7] J.L. White, M.L. Hacker, M. Buehner, M.J. Adams, G.C. Ford, P.J. Lentz Jr., I.E. Smiley, S.J. Steindel, M.G. Rossmann, J. Mol. Biol. 102 (1976) 759.
193
[8] H. Eklund, J.-P. Samama, L. Wallen, C.-I. Branden, A. Akeson, T.A. Jones, J. Mol. Biol. 146 (1981) 561. [9] H. Eklund, J.-P. Samama, T.A. Jones, Biochemistry 23 (1984) 5982. [10] H. Eklund, C.-I. Branden, Pyridine Nucleotides, Wiley, New York, 1986. [11] A. Bardea, E. Katz, A.J. Buckmann, I. Willner, J. Am. Chem. Soc. 119 (1997) 9114. [12] P. Gros, C. Zaborosch, H.G. Schlegel, A. Bergel, J. Electroanal. Chem. 405 (1996) 189. [13] J.J. Holbrook, A. Lijas, S. J. Steindel, M.G. Rossmann, in: P.D. Boyer (Ed.), The Enzymes, 3rd edition, Academic Press, New York, 1975, p. 191. [14] A.R. Clarke, D.B. Wigley, W.N. Chia, J.J. Holbrook, Nature 324 (1986) 699. [15] D.M. Parker, J.J. Holbrook, in: H. Sund (Ed.), Pyridine Nucleotide-Dependent Dehydrogenases, de Gruyter, Berlin, 1977, pp. 485 – 502. [16] J.J. Birktoft, R.T. Fernley, R.A. Bradshaw, L.J. Banaszak, Proc. Natl. Acad. Sci. USA 79 (1982) 6166. [17] H. Deng, J. Zheng, J. Burgner, D. Sloan, R. Callender, Biochemistry 28 (1989) 1525. [18] H. Deng, J. Zheng, J. Burgner, R. Callender, J. Phys. Chem. 93 (1989) 4710. [19] H. Deng, K.T. Yue, C. Martin, K.W. Rhee, D. Sloan, R. Callender, Biochemistry 26 (1987) 4776. [20] H. Deng, J. Burgner, R. Callender, J. Am. Chem. Soc. 114 (1992) 7997. [21] H. Deng, J. Zheng, J. Burgner, R. Callender, Proc. Natl. Acad. Sci. USA 86 (1989) 4484. [22] R. Callender, H. Deng, Annu. Rev. Biophys. Biomol. Struct. 23 (1994) 215. [23] H. Deng, J. Burgner, R. Callender, Biochemistry 30 (1991) 8804. [24] L. Young, C.B. Post, Biochemistry 35 (1996) 15129. [25] W.K. Surewicz, H.H. Mantsch, Biochim. Biophys. Acta 952 (1988) 115. [26] A. Elliot, E.J. Ambrose, Nature 165 (1950) 921. [27] H. Susi, in: S.N. Timasheff, G.D. Fasman (Eds.), Structure and Stability of Biological Macromolecules, Marcel Dekker, New York, 1969, pp. 575 – 663. [28] H. Susi, Methods Enzymol. 26 (1972) 445. [29] F.S. Parker, Applications of Infrared, Raman and Resonance Raman Spectroscopy in Biochemistry, Plenum Press, New York, 1983. [30] W.-Z. He, W.R. Newell, P.I. Haris, D. Chapman, J. Barber, Biochemistry 30 (1991) 4552. [31] R.A. Copeland, T.G. Spiro, Biochemistry 26 (1987) 2134. [32] T.M. Cotton, J.-H. Kim, G.D. Chumanov, J. Raman Spectrosc. 22 (1991) 729. [33] X. Dou, T. Takama, Y. Yamaguchi, H. Yamamoto, Anal. Chem. 69 (1997) 1492. [34] T.M. Cotton, in: R.J.H. Clark, R.E. Hester (Eds.), Spectroscopy of Surfaces, Wiley, New York, 1988, pp. 91 – 153. [35] G.D. Chumanov, R.G. Efremov, I.R. Nabiev, J. Raman Spectrosc. 21 (1990) 43. [36] E. Grabbe, R.P. Buck, J. Am. Chem. Soc. 111 (1989) 8362. [37] A.M. Ahern, R.L. Garrell, Langmuir 7 (1991) 254. [38] R.A. Copeland, S.P.A. Fodor, T.G. Spiro, J. Am. Chem. Soc. 106 (1984) 3872. [39] N.-S. Lee, Y.-Z. Hsieh, M.D. Morris, L.M. Schopfer, J. Am. Chem. Soc. 109 (1987) 1358. [40] R.E. Holt, T.M. Cotton, J. Am. Chem. Soc. 111 (1989) 2815. [41] E. Koglin, J.-M. Sequaris, Top. Curr. Chem. 134 (1986) 39. [42] J. Hu, R.S. Sheng, Z.S. Xu, Y.-E. Zeng, Spectrochim. Acta Part A 51 (1995) 1087. [43] J.C. Austin, R.E. Hester, J. Chem. Soc. Faraday Trans. I 85 (1989) 1159.
194
Y.-J. Xiao et al. / Journal of Electroanalytical Chemistry 465 (1999) 187–194
[44] O. Siiman, R. Rivellini, R. Patel, Inorg. Chem. 27 (1988) 3940. [45] K.T. Yue, C.L. Martin, D. Chen, P. Nelson, D.L. Sloan, R. Callender, Biochemistry 25 (1986) 4941. [46] Y.-J. Xiao, J.P. Markwell, Langmuir 13 (1997) 7068. [47] Y.-J. Xiao, T. Wang, X.-Q. Xang, X.-X. Gao, J. Electroanal. Chem. 433 (1997) 49. [48] D. Chen, K.T. Yue, C. Martin, K.W. Rhee, D. Sloan, R. Callender, Biochemistry 26 (1987) 4776. [49] C. Nadolny, G. Zundel, J. Mol. Struct. 385 (1996) 81. [50] P.R. Carey, Biochemical Applications of Raman and Resonance Raman Spectroscopy, Academic Press, New York, 1982. [51] J. Ryttersgaard, B.D. Larsen, A. Holm, D.H. Christensen, O.F. Nielsen, Spectrochim. Acta Part A 53 (1997) 91. [52] S. Krimm, J. Bandekar, Adv. Protein Chem. 38 (1986) 181.
.
[53] W.G.J. Hol, P.T. van Duijnen, H.J.C. Berendsen, Nature (London) 273 (1978) 443. [54] A. L. Lehninger, D.L. Nelson, M.M. Cox, Principles of Biochemistry, 2nd edition, Worth, New York, 1993, pp. 390–392. [55] H. Susi, M. Byler, Methods Enzymol. 130 (1986) 290. [56] M. Byler, H. Susi, Biopolymers 25 (1986) 469. [57] P.I. Harris, D.C. Lee, D. Chapman, Biochim. Biophys. Acta 874 (1986) 255. [58] P.W. Yang, H.H. Mantsch, J.L.R. Arrondo, I. Saint-Girons, Y. Guillou, G.N. Cohen, O. Barzu, Biochemistry 26 (1987) 2706. [59] W.K. Surewicz, M.A. Moscarello, H.H. Mantsch, Biochemistry 26 (1987) 3881. [60] J.L.R. Arrondo, H.H. Mantsch, N. Mullner, S. Pikula, A. Martonosi, J. Biol. Chem. 262 (1987) 9037.