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    Topic review

    Infrared and Resonance Raman Spectroscopies

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    Definition

    Vibrational spectroscopy and in particular, resonance Raman (RR) spectroscopy can provide molecular details on metalloproteins, including those containing multiple cofactors, which are often challenging for other spectroscopies. Due to distinct spectroscopic fingerprints, RR spectroscopy has a unique capacity to monitor simultaneously and independently different metal cofactors that can have particular roles in metalloproteins. These include e.g., (i) different types of hemes, as well as their spin and redox states, (ii) different types of Fe-S clusters, and (iii) bi-metallic center and electron transfer (ET) Fe-S cluster in hydrogenases. IR spectroscopy can provide un-matched molecular details on specific enzymes like hydrogenases that possess catalytic centers coordinated by CO and CN- ligands, which exhibit spectrally well separated IR bands.

    1. Introduction

    Vibrational spectroscopy has, over the past couple of decades, provided valuable information on structure – function relationship in metalloproteins, contributing to the elucidation of molecular mechanisms, dynamics and interactions on the level that goes beyond high resolution crystallographic structures. It does not depend on size or paramagnetic properties of the protein like nuclear magnetic resonance (NMR) and electron paramagnetic resonance (EPR) spectroscopies, and furthermore, it can be coupled with electrochemical methods and employed in time-resolved mode, offering the possibility to probe redox and transient molecular events down to femto-second time scale.[1]

    1.1 Resonance Raman and surface-enhanced (resonance) Raman spectroscopy

    In the case of metalloproteins, the intrinsically low sensitivity and the selectivity of Raman spectroscopy are strongly increased when the energy of the incident laser light is in resonance with an electronic transition of the chromophore. The resultant resonance Raman (RR) spectra display several orders of magnitude higher intensity for the vibrational modes of the cofactor, regardless of the size of the protein matrix.[1] RR spectroscopic studies have mainly focused onto metalloproteins containing iron and copper ions, but they have also been reported for other transition metals like nickel, cobalt and molybdenum.[2][3][4][5] In particular, the studies of heme and non-heme iron, iron-sulfur cluster and copper center containing metalloproteins have provided a wealth of details on their active sites, hence contributing to understanding of their mechanistic properties.

    RR spectra of heme proteins obtained upon Soret-band excitation are, in the high frequency (1300–1700 cm-1) region, dominated by the core-size marker bands νi (ν4, ν3, ν2 and ν10) originating from complex porphyrin vibrations that are sensitive to the redox, spin, and coordination state of the heme iron (Figure 1 and Table 1). The spectral distinction between LS and HS heme populations is therefore straightforward in RR spectra, which has helped addressing e.g., processes at catalytic HS vs. electron transfer (ET) LS hemes in oxygen and nitrite reductases and the activation mechanism of cytochrome c peroxidases (CcPs).[6][7][8][9] Similarly the capacity of RR spectroscopy to distinguish between ferric and ferrous heme populations contributed to disentangling of the ET processes in numerous metalloproteins, as well as those that involve semi-reduced forms of Complex III and dihemic bacterial CcPs.[9][10]

    Figure 1. Simulated Resonance Raman (RR) spectra of different oxidation, spin and coordination states of cytochrome c, with depicted marker bands, left and corresponding heme configurations, right. (a) Ferrous (Fe2+) heme in low spin (LS) state, coordinated by distal methionine and proximal histidine (6cLS). (b) Ferric (Fe3+) heme in 6cLS state, coordinated by distal methionine and proximal histidine. (c) Ferric heme in a high spin (HS) state, coordinated by proximal histidine and vacant 6th axial position (5cHS). (d) Ferric heme in 6cHS state, coordinated by a distal water molecule and proximal histidine.

    Table 1. Frequencies of characteristic (resonance) Raman and IR active vibrational modes of metalloprotein cofactors. Heme proteins - typical frequency range of the marker bands (νi) of ferric and ferrous His/Met and His/- coordinated c-type heme in RR spectra. Iron-sulfur proteins - typical frequency range of predominantly bridging (b) and terminal (t) Fe-S RR modes for different cluster types. Hydrogenases - typical frequency regions of IR and RR bands for metal bound CO and CN ligand vibrations.

    RR spectra of Fe-S cluster containing proteins, obtained with a laser wavelength that matches the energy of S → Fe charge transfer transitions, show selectively enhanced modes involving the metal–ligand stretching coordinates in the low-frequency (200–450 cm−1) region (Table 1). The spectra are sensitive to the cluster type, structure and symmetry and moreover allow for distinguishing between bridging vs. terminal Fe-S vibrational modes that involve inorganic and cysteine bound S atom in each cluster, respectively.[11] Unlike in the case of heme proteins, in Fe-S proteins only one redox state is usually RR active; [2Fe–2S]2+, [3Fe–4S]1+ and [4Fe–4S]2+ (and [4Fe–4S]3+ in HiPIPs). RR spectroscopy can simultaneously probe different types of clusters present in the same protein, alongside with process that account for their interconversion.[11]

    Large enhancements of Raman signals are obtained by measuring the spectra of molecules in close proximity to plasmonic metals (e.g. Ag and Au), due to the surface enhanced Raman (SER) scattering effect.[1] Accordingly, SER spectroscopy can provide sensitive (bio)chemical information about the studied system, and is therefore viewed as a promising analytical tool for e.g. medical diagnostics.[12][13][14] The combination of the SER and RR effects (SERR spectroscopy) enables further enhancement of the signal by selectively probing the chromophoric part of molecule attached to SER active metal surfaces.[1] SERR spectroscopy can provide specific molecular information about proteins that perform their physiological function in the immobilized state (e.g. membrane proteins). The nanostructured metal that amplifies the signals can serve as a working electrode, allowing for spectro-electrochemical SERR studies capable of monitoring the changes of the heme group as a consequence of variations of the electrode potential.[1][15]

    1.2 Infrared (IR) and surface-enhanced IR absorption spectroscopy

    Infrared (IR) spectroscopy most commonly provides information on the secondary structure of proteins based on the analysis of the amide I (1600–1700 cm-1) and amide II (1480–1580 cm-1) bands.[1] However, when measured in the difference mode, IR spectra can probe structural changes of individual cofactors and sensitively detect redox-linked structural changes, protonation events or amino acids and specific groups upon isotopic labeling (e.g. 13C and 15N labeled heme).[16] Analogous to SERR, surface enhanced infrared absorption (SEIRA) spectroscopy probes protein molecules that are found in the vicinity of nanostructured metal surfaces deposited on a transmission window or an inert ATR (attenuated total reflection) crystal, experiencing an enhanced absorption of the incident IR radiation. SEIRA can be used to control the protein attachment and orientation at the biocompatible surface upon immobilization, but more importantly it can also provide fine details about redox-linked changes of the secondary structural elements when employed in spectro-electrochemical mode (vide supra).[1][15] For certain metalloenzymes, where the active site harbors ligands that possess spectrally isolated IR absorption bands, such as hydrogenases that contain bimetallic catalytic centers with unusual CO and CN- ligands, IR (transmission) spectroscopy and its surface sensitive variants offer unique insights in the redox behavior of the catalytic center.[17]

    2. Heme Proteins

    2.1. Nitrite reductases (NiR)

    NrfHA menaquinol: nitrite oxidoreductase complex catalyzes the six-electron reduction of nitrite to ammonia in a reaction that involves eight protons. It houses a total of 28 heme groups in the biological unit and, as such, represents a challenge for every experimental approach. 22 of heme groups have 6cLS and 6 have 5cHS configuration; two of the HS hemes are membrane-integrated and the other four represent catalytic sites. The HS and LS hemes can be easily distinguished in the RR spectra of NrfHA, with redox sensitive ν4 and redox/spin state sensitive ν3 modes of the HS species at 1366 and 1493 cm-1, and of the LS at 1373 and 1501 cm-1, allowing for independent monitoring of the processes that involve these two populations. Binding of nitrite to the catalytic HS hemes and the resulting spin configuration of the initial enzyme / substrate complex have profound consequences for the reaction mechanism of nitrite reduction, indicating whether the N-O bond cleavage follows homolytic or heterolytic route, and the subsequent steps of the catalytic cycle. RR data have provided the first experimental evidence that nitrite binding to NrfHA active site HS hemes causes a spin conversion from HS to LS configuration, which implies that the heterolytic cleavage of the N-O bond is favored in the first step of the catalytic reaction.[18]

    SERR spectro-electrochemistry helped disentangle the ET pathway in NrfHA. Potentiometric titrations of NrfHA immobilized on biocompatible Ag electrode, followed by the analysis of the ν4 band of the HS hemes provided the redox potentials of the catalytic and the non-catalytic HS centers, revealing the first evidence for the downhill biological electron flow in the integral NrfHA.[6]

    2.2. Heme containing respiratory chain and analogous complexes

    Complex III (ubiquinol : cyt c oxidoreductase or bc1 complex) catalyzes the transfer of two electrons from ubiquinol to two cyt c molecules. bc1 complexes are formed by a minimum of three subunits; one contains a c1-type heme (or structurally and functionally analogous cytochrome f in plants, cyanobacteria and green algae that house cytochrome b6f complex), one holds a Rieske type [2Fe–2S] cluster, and the third one contains two LS b-type hemes, designated bL and bH.[10] RR spectroscopy has been employed in the initial characterization of heme groups of the complex at different stages of reduction. Namely, due to the differences in midpoint redox potentials of the heme c1, low potential heme b (bL) and high potential heme b (bH), it is possible to selectively reduce the higher potential sites (heme c1 and bH). A selective resonance enhancement using multiple excitation lines and sequential stoichiometric reduction of the complex allowed for spectral distinction between the c and b-type hemes, providing insights into the differences in peripheral heme-protein interactions, and in particular to the conformation of vinyl substituents of the pyrrole rings.[10]

    RR spectra of cyt c1 and cyt f containing subunits of bc1 and b6f complexes employing Q-band (550 nm) excitation effectively probe the respective local heme environments of these sites, indicating remarkably similar macrocycle geometry of the two hemes.[19] Cyt b6f participates in the oxygenic photosynthesis as a redox link between the two reaction center complexes. RR spectroscopy initially helped identify the chromophoric groups in b6f complexes isolated from different species, revealing a presence of chlorophyll a, β-carotene and an additional 5cHS c1-type heme.[20][21] A functional analogue to bc1 complex carrying two b- and one a-type heme was found in an archaeon. RR spectroscopy helped characterize the three heme groups in this enzyme that shows bc1 complex activity.[22]

    IR difference spectroscopic studies of bc1 complex and more recently, ATR-IR spectroscopy of thin layers of bovine bc1 complex that have been deposited on the surface of a silicon microprism revealed redox-induced structural changes in these enzymes. Redox-sensitive IR absorption bands in the potential-induced difference spectra of c1, bH and bL hemes, ubiquinone and surrounding protein were identified upon separation by selective reduction. A similar approach was used to probe the [2Fe–2S] cluster of the complex.[23][24][25]

    Oxygen reductase - Complex IV (cyt c : oxygen oxidoreductase, CcO, or heme copper oxygen reductase, or heme copper oxidase, HCO) catalyzes the reduction of molecular oxygen to water by utilizing four electrons and four protons. This is one of the most fundamental reactions in living organisms. HCOs pump protons from the N to the P side of the membrane, contributing to the generation of transmembrane electrochemical potential, which drives the ATP synthesis.[26] The catalytic reaction occurs at a binuclear center formed by a HS heme (e.g. heme a3 in mitochondrial enzyme, here designated as CcO) and a copper atom (CuB). The ET to the binuclear center is mediated by a LS heme group in the catalytic subunit (heme a in CcO) and a copper center in the non-catalytic subunit, which is composed of two copper atoms (CuA) that hold one redox equivalent. The number of non-catalytic subunits and their cofactors vary in HCOs. Bacteria and archaea have complexes that are simpler than the mammalian CcO, and if required, the expression of the appropriate HCO (e.g. aa3-, cbb3- or ba3- type) can be fine-tuned as a function of oxygen levels in the environment. RR spectroscopy played a fundamental role in identification and description of the cofactors in these enzymes, such as the type and spin, oxidation and coordination state of the heme groups in structurally diverse HCOs of different origin [27][28][29][30][31] as well as interactions, conformations and the dynamics of small ligand (CO, CN-, N3, NO) binding.[32][33][34][35][36][37][38] The low frequency region of RR spectra has provided unprecedented details on (i) mechanistic properties of HCOs, including detection and identification of the short living catalytic intermediates formed upon oxygen binding to the catalytic HS heme (i.e. a3, b3, o3) and (ii) molecular environment and conformation of the catalytic site, based on correlation of Fe-CO and Fe-C-O stretching mode frequencies of HS heme-CO adduct.[39][40]

    Reduced HCOs bind molecular oxygen via formation of short living A, P, F, and H intermediates,[41] which were initially only tentatively described by electronic absorption spectroscopy. RR experiments performed independently by the groups of Kitagawa, Rousseau and Babcock led to a consistent mechanistic model of HCOs, employing isotopic labeling, mutagenesis studies, a number of structurally different HCOs and individually developed TR RR experimental approaches.[26][41][42][43][44][45] In particular, iron-oxygen stretching frequencies obtained by TR RR 16O2 -18O2 difference spectra, measured in H2O and D2O media, helped identifying proton coupled ET reactions and established structural fingerprints of each catalytic intermediate. A presence of a-type hemes in some HCOs offers further advantages as the effect of mutations and other molecular perturbations can be monitored via porphyrin formyl stretching C=O modes, which are well resolved in RR spectra of both ferrous HS and LS hemes at 1661 and 1628 cm-1, respectively.[46]

    IR difference spectroscopy recorded under steady state conditions and in the TR mode identified protonation/deprotonation and re-protonation events during the catalytic cycle of aa3-type HCO. The IR data in particular point out the role of a proton shuttle of glutamic acid found at ~ 11 Å from the active site, which changes its protonation state a number of times during the catalytic cycle. The role of the Tyr, located in the proximity of the binuclear center, in the splitting of the O-O bond during the catalysis was also highlighted by IR spectroscopy.[41]

    More recently, SERR spectroscopy has provided a novel platform for investigations of HCOs under conditions that can mimic some basic features of their natural environment. Since these enzymes exert their function integrated into a phospholipid bilayer under i) restricted mobility, ii) directionalized ET from electron donor to LS heme and binuclear site and iii) influence of strong interfacial electric fields, the immobilization onto biocompatible electrodes mimics far better these conditions than the solution studies.[47] Furthermore, SERR potentiometric titrations represent a powerful alternative to common methods for the determination of the E0 value, which is a prerequisite for understanding the ET pathway in these enzymes. The determination of E0 of the individual heme groups in HCOs by conventional electronic absorption titrations is hampered by strong overlapping of the spectra of individual hemes and by complex cooperative effects that modulate the electroprotonic energy transduction.[47] Deconvolution of the (SE)RR spectra of aa3-type quinol oxidase (aa3 QO) by component analysis allowed separation of the contributions from the LS and the HS hemes based on their respective ν3 and νC=O modes, among over 40 vibrational modes originating from the two hemes.[47] The potential dependence of these spin and redox-state sensitive marker bands allowed for determined midpoint potentials of hemes a (E0 = 320 mV vs. NHE) and a3 (E0 = 390 mV vs. NHE), which reveal a reversed order of reduction compared to mitochondrial-like HCOs. This suggests a distinct mechanism of electroprotonic energy transduction in aa3 QO in comparison with e.g. CcO. A downhill ET is already guaranteed by the order of the midpoint potentials at the onset of enzyme reduction, indicating that this enzyme does not require a complex network of cooperativities to ensure exergonicity. SERR studies of redox properties of aa3-type and a more complex, pentahemic cbb3-type HCOs employed enzymes anchored to nanostructured Ni-NTA coated Ag electrodes and further embedded into lipid bilayer. This strategy allows for even better mimicking of the natural membrane together with the control of the enzyme orientation that can be probed via SEIRA spectroscopy.[48][49][50][51] SERR analysis focusing onto the protonation dependent CH2 propionate bending modes, detected by H2O - D2O (SE)RR difference spectroscopy, supported the hypothesis that heme a3 propionates act as possible proton loading sites in HCOs.[52]

    IR difference spectroscopy was employed to specifically probe the heme propionate protonation events using a CcO mutant with 13C labeled propionates to distinguish its signal from possible concomitant changes of CO stretching modes of other carboxylic acids side chains.[16] Differential IR spectra, employing isotopically labeled ligands or different redox states of the protein (e.g. oxidized minus reduced), provided further structural details on active site conformations in HCOs and specific protonatable sites.[53][54] Step-scan IR spectroscopy, using CuB-CO adduct as a probe, showed that the protonation events in the binuclear site of ba3-type HCO involve Tyr residues.[55] Moreover, electrochemical redox titrations of aa3- and caa3-type HCOs, performed using IR spectroscopy in the transmission and the ATR mode, helped identify the redox sensitive bands, originating from heme groups, their ligands, amino acid residues and/or protein backbone.[23][53][56]

    Nitric oxide reductase (NOR) catalyzes the two electron reduction of NO to N2O at the di-nuclear heme b3-FeB center. Similarly to the active site of a HCO, the heme b3 is in the HS state, while FeB is a non-hemic iron atom. NORs in addition house c-type and b-type LS hemes. RR spectroscopy provided evidence about NO reduction by monitoring the events that occur upon NO binding to catalytic b3-type heme. Specifically, formation of the N-N bond and the fate of proximal His-heme b3 bond along this process, as well as the recovery of the b3-O-FeB state, have been disentangled by RR spectroscopy.[57]

    3. Fe-S proteins

    3.1. Hydrogenases

    Hydrogenases catalyze the reversible splitting of molecular dihydrogen (H2) into protons and electrons (or hydride species), often with high catalytic rates, and can be classified into [NiFe], [FeFe], and [Fe]-hydrogenases depending on the particular metal composition of their catalytic centers.[58] These metalloenzymes have been extensively studied in recent years revealing new insights on the mechanistic details of the H2 splitting, which are regarded as benchmarks for the development of biomimetic and bioinspired catalysts for H2 production and/or uptake.[59] In this context, vibrational spectroscopy has greatly contributed to the understanding of hydrogenases, targeting inorganic cofactors, such as the particular catalytic site and Fe-S cluster relays, as well as organic protein cofactors (e.g. FAD and FMN) and certain adjacent protein residues (e.g. cysteine, glutamate) all involved in the biological H2 conversion.[60] Thereby, IR spectroscopy has played a major role in the characterization of these enzymes, which is related to the presence of the unusual inorganic carbon monoxide (CO) and cyanide (CN) ligands coordinated to the particular active sites (Figure 2a, Table 1) that exhibit characteristic IR bands in a spectral region free of other protein absorptions. Noteworthy, the specific band position of the CO (1780 – 2030 cm–1) and CN (2030 – 2150 cm–1) stretching modes (Figure 2b) are particularly sensitive to changes of the local protein environment, oxidation states of the metal centers, as well as external perturbations induced by changes of temperature and pH.[61][62] Solution IR spectro-electrochemical studies[63] were thus extremely helpful for the structural and electronic characterization of the different redox states, providing key insights in the catalytic mechanism. Briefly, the [NiFe]-hydrogenase catalytic cycle comprises four intermediates, i.e. Nia-S, Nia-L, Nia-C and Nia-SR differing in the oxidation state of nickel (Ni1+/2+/3+) while iron retains a Fe2+ LS configuration throughout the entire catalytic cycle. In the [FeFe]-hydrogenase catalytic cycle, the di-iron center, 2FeH, has been isolated in a variety of intermediates, some of which are well understood (Hox, Hred/HredH+, Hhyd) while other presumed intermediates (Hred´H, Hsred, HhydH+, HoxH+) are still matter of intense discussion.[64][65] For understanding the assignment of the catalytic competency of hydrogenase intermediates, a number of studies employing ATR-FTIR[66] and SEIRA,[67] both under the control of gas atmosphere or/and potential, photogating with time-resolved IR spectroscopy,[68] and protein film IR electrochemistry (PFIRE)[69] have been performed.

    Figure 2. (a) Schematic representation of the active sites of [NiFe]-, [FeFe]- and [Fe]-hydrogenase. In [NiFe]-hydrogenases, the nickel ion is coordinated by four cysteine residues, two of which are shared with the iron ion. One CO and CN– ligands complete the coordination sphere of the iron. In [FeFe]-hydrogenases, the catalytic center is the H-cluster comprising a di-iron (2FeH) and a [4Fe–4S]H site sharing a cysteine coordination. Each iron of the 2FeH is coordinated by one CN and one CO ligand. A bridging CO and an azadithiolate ligand complete the [2Fe]H coordination. In [Fe]-hydrogenases, the single iron is coordinated by two CO ligands, a cysteine thiolate and a guanylyl pyridinol cofactor bound to the iron via its acyl-carbon and pyridinol-nitrogen. Substrates and inhibitors bind at the coordination site marked by X. (b)  IR spectra of as-isolated (top trace, Nia-S intermediate) and H2-reduced (bottom, Nia-C and Nia-SR intermediates) RH (HoxBC) at 283 K displaying the ligand stretching vibrations of the CO and CN ligands of the active site, located between 1890−2000 cm−1 and 2030−2120 cm−1, respectively. (c) Low temperature RR spectra at 80 K of the regulatory [NiFe] hydrogenase (RH) recorded with excitation at 458 (blue trace) and 514 (green trace) nm in its as-isolated form. Color code of various metal ligand vibrations: Fe-CN/CO (red); Ni/Fe-S of bridging cysteines (black); Fe-S of [4Fe-4S] clusters (dark yellow).

    Recently, ultrafast pump-probe and two-dimensional (2D) IR provided novel insights into the dynamic interactions between the hydrogenase CO/CN- ligands and the complex interplay between the [NiFe] site and the protein scaffold.[70] Apart from the distinct CO and CN- markers bands, IR spectroscopy has also unveiled the participation of protein residues in the stabilization of certain intermediates. Combined IR difference spectroscopy and hydrogen/deuterium (H/D) isotopic labeling, allowed for direct monitoring of the changes in the protonation state of conserved glutamate and arginine residues in both [NiFe] and [FeFe] hydrogenases.[71] Noteworthy, the IR detection of S–H/S–D stretching bands at 2505 cm–1 and 1822 cm–1, respectively, confirmed the protonation of a Ni-bound cysteine in the Nia-L intermediate.

    Additional mechanistic understanding of hydrogenases has been derived from RR spectroscopy that enables the detection of multiple protein cofactors. RR spectra of the F-420 [NiFe]-hydrogenase revealed the spectral contributions of FAD, Fe-S clusters, and the [NiFe] active site.[72] Notably, 458 nm laser excitation preferentially enhanced the metal-ligand modes of the Fe-S clusters (Figure 2c, blue trace, Table 1), while the excitation lines distant from the S→Fe charge transfer (CT) transition resulted in a preferential enhancement of the active site vibrational modes (Figure 2c, green trace, Table 1).[73][74] Supported by theoretical methods, RR spectroscopy has shown to be highly versatile in unraveling key electronic and structural details of the [NiFe] and [FeFe] bimetallic centers. Additionally, RR and IR spectroscopic studies were successfully performed on the same single hydrogenase crystals, complementing the crystallographic data and facilitating structural assignment of Additional mechanistic ambiguous inhibited and active redox states, such as the oxygen stable Hinact and the hydrogen binding intermediate Nia-S state in [FeFe] and [NiFe] hydrogenases, respectively.[72][75] Notably, well-defined redox states in hydrogenase single crystals for IR imaging have been generated by electrochemical and gas control.[76][77][78][79] Furthermore, Fe-OH ligation at the unprecedented [4Fe–3S] cluster of the O2-tolerant membrane-bound [NiFe] hydrogenase from R. eutropha has been identified by RR spectroscopy and confirmed by theoretical quantum and molecular mechanics methods.[80][81] The corresponding FeS region of the three different clusters in the enzyme has been disentangled recently by RR spectroscopy in combination with protein engineering and X-ray crystallography,[81] while its coupling with the physiological electron acceptor, cytochrome b3, was investigated by SERR spectro-electrochemistry.[69][82]

    3.2. Multi-cluster containing ferredoxins

    Ferredoxin, Fd, is a small ET Fe-S containing protein in a number of metabolic reactions. Multiple wavelength excitation of di-cluster (e.g., [3Fe–4S] and [4Fe–4S] in Fd) containing Fe-S proteins can assure selective RR enhancement of individual clusters and provide their independent monitoring. IR and RR spectroscopies of a di-cluster Fd have addressed the effect of thermal perturbation on the level of secondary structural elements and the individual clusters, respectively. RR spectroscopy provided fine details about the disassembly of the centers in Fd by using 413, 458 and 514 nm laser excitation lines,[83] which enabled a differential enhancement of the RR bands associated with the [3Fe–4S] and [4Fe–4S] centers. It was shown that the temperature-induced simultaneous degradation of the two metal centers follows the initial loss of the alfa helical structure and triggers major structural changes of the protein matrix, including the loss of tertiary contacts and a secondary structure reorganization.[83]

    This entry is adapted from 10.3390/molecules26164852

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