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Protonation-Dependent Structural Heterogeneity in the Chromophore Binding Site of Cyanobacterial Phytochrome Cph1 Francisco Javier Velazquez Escobar, Christina Lang, Aref Takiden, Constantin Schneider, Jens Balke, Jon Hughes, Ulrike Alexiev, Peter Hildebrandt, and Maria Andrea Mroginski J. Phys. Chem. B, Just Accepted Manuscript • DOI: 10.1021/acs.jpcb.6b09600 • Publication Date (Web): 14 Dec 2016 Downloaded from http://pubs.acs.org on December 16, 2016

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The Journal of Physical Chemistry

Protonation-dependent Structural Heterogeneity in the Chromophore Binding Site of Cyanobacterial Phytochrome Cph1

Francisco Velazquez Escobar1, Christina Lang2, Aref Takiden2, Constantin Schneider3, Jens Balke3, Jon Hughes2, Ulrike Alexiev3, Peter Hildebrandt1 and Maria Andrea Mroginski1*

1

Technische Universität Berlin, Institut für Chemie, Sekr. PC 14, Straße des 17. Juni 135, D-10623

Berlin 2

Plant Physiology, Justus-Liebig University Gießen, Senckenbergstr. 3, D-35390 Giessen, Germany

3

Freie Universität Berlin, Institut für Experimentalphysik, Arnimallee 14, 14195 Berlin, Germany

Corresponding author: [email protected] Abstract Phytochromes are biological red/far-red light sensors found in many organisms. Photoisomerization of the linear methine-bridged tetrapyrrole triggers transient proton translocation events in the chromophore binding pocket (CBP) leading to major conformational changes of the protein matrix which are in turn associated with signaling. By combining pH-dependent resonance Raman and UV-visible absorption spectroscopy we analyzed protonation-dependent equilibria in the CBP of Cph1 involving the proposed Pr-I and Pr-II substates that prevail below and above pH 7.5, respectively. The protonation pattern and vibrational properties of these states were further characterized by means of hybrid quantum mechanics / molecular mechanics (QM/MM) calculations. From this combined experimental-theoretical study we were able to identify His260 as the key residue controlling

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pH-dependent equilibria. This residue is not only responsible for the conformational heterogeneity of CBP in the Pr state of prokaryotic phytochromes, discussed extensively in the past, but it constitutes the sink and source of protons in the proton release/uptake mechanism involving the tetrapyrrole chromophore which finally leads to the formation of the Pfr state. Thus, this work provides valuable information which may guide further experiments towards the understanding of the specific role of protons in controlling structure and function of phytochromes in general.


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Introduction Phytochromes are biological red-light photoreceptors found not only in plants but also in cyanobacteria, bacteria and fungi.1-5 Light sensing is based on an open-chain-tetrapyrrole chromophore which is covalently attached to the protein matrix via a thioether linkage.1,5 Upon

light

absorption,

the

chromophore

switches

between

the

red-absorbing

thermodynamically stable state Pr and the far-red absorbing Pfr state, which in prototypical phytochromes can also thermally revert to Pr.1,6 In all phytochromes characterized so far, the chromophore in Pr adopts a periplanar ZZZssa (C5-Z, C10-Z, C15-Z, C5-syn, C10-syn, C15anti) conformation (Figure 1), whereas Pfr is ZZEssa.1,5 The first step of the photoinduced Pr/Pfr transformations is the Z/E photo-isomerization of the chromophore at the C15=C16 double bond of the CD methine-bridge, which is followed by thermal relaxations of the cofactor and the protein including proton transfer steps.7 Despite the large amount of structural and spectroscopic data available for phytochromes,1-18 a complete description of the ground state structural properties of the chromophore binding pocket is still lacking. This particularly refers to the structural heterogeneity of the chromophore in the parent states.10, 12 The structural heterogeneity of Pr was initially observed in plant phytochromes.19,20 The multi-component kinetics of the Pr→Pfr phototransformation indicated the presence of at least two isoforms absorbing at slightly different wavelengths. The same conclusion was derived for Cph1 phytochrome from the cyanobacterium Synechocystis 6803. Temperaturedependent changes of the emission spectra with the excitation wavelength were detected, pointing to at least two different ground state species of Pr in a thermal equilibrium.13 In contrast, Matthies and coworkers claimed a homogeneous structure on the basis of an absorption and stimulated RR spectroscopic analysis.21, 22 However, two sub-states of Pr were identified and structurally characterized by solid-state NMR both in Cph1 and oat phyA3.12, 23 These findings prompted a re-evaluation of ultrafast absorption spectroscopy data by Larsen



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and co-workers.10 These authors provided evidence for the coexistence of at least two subpopulations of Pr in Cph1, which exhibit different excited state properties and therefore different photochemical activities. While one population is associated with a rapid Lumi-R formation, the other subpopulation is characterized by slower excited state decay but higher fluorescent quantum yield. These conclusions are consistent with a recent spectroscopic analysis of Pr of the bacteriophytochrome P2 from Rhodopseudomonas palustris.24 Structural heterogeneity was also shown for Pfr in prototypical prokaryotic phytochromes from Deinococcus radiodurans and Agrobacterium tumefaciens

25

and Cph18,

18

, whereas bathy

phytochromes, in which Pfr represents the thermodynamically stable state, reveal no indication of heterogeneity.26, 27 The conformational sub-states Pr-I and Pr-II of Cph1 were analyzed on the basis of 1H-13C contact maps from MAS (magic-angle spinning) NMR spectroscopy.12 These substates were proposed to differ with respect to the protonation state of His260 and His290 and the orientation of the pyrrole water while the tetrapyrrole chromophore is protonated in both species. As a consequence of the change in the protonation state of the neighboring histidines, a significantly different hydrogen network around the chromophore involving the pyrrole water (W1), as well as the water molecules W4 and W5 was suggested (figure 1). Accordingly, Pr-I is characterized by a cationic His260 and a neutral His290 protonated at Nδ while in Pr-II both histidines are neutral and protonated at Nε. Furthermore, it was postulated that solely the Pr-II form prevails in the crystal phase. In this work we have employed a different combined experimental-theoretical approach to analyze the protonation-dependent structural heterogeneity in the chromophore pocket of Pr in Cph1. On the basis of the crystal structure of the photosensory module of Cph1 (Cph1∆2),28 a hybrid quantum mechanics - molecular mechanics (QM-MM) method was used to calculate the structures of the chromophore pocket.29 In this technique, the protein fragment of interest, ACS Paragon Plus Environment

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i.e. the chromophore and important surrounding amino acids, was treated quantummechanically (QM partition) whereas for the remainder of the protein and the solvent a classical force field was applied (MM partition). Thus, the impact of the protein environment on the chromophore structure is considered explicitly, allowing for a more accurate calculation of the spectral properties compared to a pure QM method.18,

30

Here we have

focused on the calculation of Raman spectra of the PCB cofactor which were compared with the experimental RR spectra measured as a function of the pH. The potential of QM-MM techniques

was

photoreceptors.18,

already 30-32

demonstrated

previously

for

phytochromes

and

related

Now we have applied this approach to determine the nature of the

conformational sub-states Pr-I and Pr-II of Cph1∆2, focusing on the influence of the protonation states of His260 and His290 and the chromophore structure.

Materials and Methods: Sample Preparation. Cph1∆2 wild-type and H260Q mutant holoprotein expression in E. coli, production of

13

C5-labelled PCB, in vitro autoassembly with the apoprotein and subsequent

purification were performed according to established protocols.28, 33- 35 Resonance Raman Spectroscopy. Measurements were performed with 1064-nm excitation using a Fourier-transform Raman spectrometer, following the same procedure as in previous works.6,14,18,25-27 Protein solutions were prepared in Tris buffer (100 mM Tris-HCl, 300 mM NaCl, 5 mM EDTA). The pH(D) was adjusted with H(D)Cl or NaOH(D) solution (2 M). RR spectra of Pr following irradiation with far-red light (incandescent source with 730 nm cutoff filter) were recorded in the dark, whereas partial conversion to Pfr was achieved after 2 min red light irradiation (646 or 670 nm LED). In addition, RR spectra of Cph1∆2 were recorded between pH = 6 and 9.0 in steps of 0.5 units. RR spectra of Cph1∆2 crystals 28 were obtained



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as described previously.18 All spectra were measured at 80 K. Reversibility of the pHdependent processes was checked on the basis of RR spectra measured prior (pH 7.8) and after pH titration by re-exchanging the buffer back to pH 7.8. Global analysis. After background subtraction, the RR spectra were subjected to a global component analysis that allows extracting the spectra of individual components from a series of experimental spectra.36 A detailed description of the component analysis and determination of the apparent (pKA,app) and true pKA has been provided elsewhere.17,

37

The spectral

contribution of the protein was negligible for the native Cph1∆2 species. The H260Q-mutant displays a lower chromophore binding affinity than the wild-type (WT) protein,33 such that the apoprotein contribution had to be considered in the global analysis as described in detail in the Supporting Information (figure S1). UV-Vis Spectroscopy and pH titration. Absorption spectra were measured with a Shimadzu UV2450 spectrometer. Cph1∆2 was prepared in Tris-Citrate buffer (10 mM Tris-Citrate, 15mM NaCl). Titration and measurements were carried out in a 1x1 cm quartz cuvette at 20°C. The pH was adjusted with small aliquots of 1 M NaOH and HCl in the range from 6.5 to 10.3. Pr was generated by saturating irradiation with a LED at 735 nm. Photoequilibrium corresponding to 70% Pfr occupancy was achieved by irradiation with a LED at 640 nm for 2 minutes. The spectra of pure Pfr were calculated by subtracting the spectra of the corresponding Pr state normalized to 30%. The spectra were scaled to the extinction coefficient ε658 of Pr (85 mM-1cm-1 at λmax for pH 7.8).38 Only low intensity light was used for the measurements to avoid photoconversion of Pr.17 The pH titration curves were generated from the respective absorbance values at 9 distinct wavelengths in the range between 638 and 682 nm for Pr and between 682 and 726 nm for Pfr. A global fit of the HendersonHasselbalch equation was performed.


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Electrostatic Calculations. The protonation probabilities of all titrable amino acids in the crystal structure of the photosensory domain of Cph1 as Pr (PDB entry: 2VEA)28 were determined by solving the linearized Poisson-Boltzmann equation of the electrostatic potential.39 A modified version of the Karlsberg+ software package,40 which also accounts for the presence of a tetrapyrrole-prosthetic group, was employed for this purpose. The computation of the electrostatic energy was performed using the CHARMM22 force field.41 Quantum mechanics - molecular mechanics calculations. Structural models were constructed from the crystal structure of the photosensory module of Cph1 as Pr (PDB entry: 2VEA) 28. The remaining missing hydrogen atoms were added to the protein matrix with the Karlsberg+ software package.40 In addition, based on spectroscopic evidence,18 the PCB chromophore was nitrogen-protonated whereas the propionic side chains were deprotonated yielding a total charge of -1e. Altogether nine models were considered that differ with respect to the protonation states of His260 and His290. In the following we will term these models as H260X-H290Y, where X and Y specify the respective protonation state. In this nomenclature, X (Y) = E and X (Y) = D refer to a protonation at the Nε and Nδ nitrogens, respectively. X (Y) = P denotes the case of double protonation. Subsequently, the protein was solvated in a thermally-equilibrated box of TIP3P water molecules and its charge was neutralized by adding chloride and sodium ions to the solution. The geometry of the PCB chromophore and all side chains within 20 Å of the N_A nitrogen in PCB (active region) were optimized within a hybrid QM/MM approach with the modular program package ChemShell.42,

43

Energy minimization was performed using the limited

memory quasi Newton L-BFGS algorithm working with delocalized internal coordinates.44 The QM fragment was described with DFT at the B3LYP/6-31G* level of theory, whereas the MM part was treated by an empirical CHARMM22 force field. The coupling between the QM and MM regions was described through the electrostatic embedding model using the charge-



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shift scheme.45 The QM partition comprised the PCB cofactor, the Cys258 side chain, the pyrrole water, Asp207 and Iso208, the side chains of Ser272, Thr274, Y263, His260, and His290, as well as six water molecules located in the tetrapyrrole cavity, summing up to a total of 167 to 169 atoms depending on the protonation state of the H260 and H290. The Raman spectrum of the protein-bound PCB chromophore was computed by the QM/MM method as described previously.18,

26, 30

For a better description of the experimental RR

intensities, the calculated Raman intensities of the PCB chromophore were scaled by a factor that depended on the contribution of internal coordinates to the corresponding normal mode.46 Results pH-dependence of UV-vis absorption. The UV-Vis absorption spectra of the WT 514-residue sensory module of Cph1 (Cph1∆2) and its H260Q mutant as studied here were essentially identical to those reported previously

33

with λmax for Pr and Pfr at 659 nm and 700 nm for

Cph1∆2 WT. Under more alkaline conditions absorbance of the WT weakened and was largely abolished in H260Q, as described earlier.33 Absorption spectra were measured at different pH values (figure 2). Titration curves from the absorbance changes at nine distinct wavelengths were used to calculate the pKa values. In the Pr and Pfr states the titration isosbestic points at 575 nm and 615 nm, respectively, are blueshifted by more than 10 nm compared to those reported by van Thor et al.17 For the Pr state, the analysis based on a global fit of the Henderson-Hasselbalch equation yielded two pKa values of 7.55±0.11 and 9.04±0.02. Whereas the lower value is in line with published data and agrees well with the value of 7.4 for the Pr-I/Pr-II equilibrium derived from the RR experiments described below, the higher value of 9.04 determined here differs significantly from the pKa reported previously by van Thor et al.17 On the other hand, our pKa value for chromophore deprotonation in the Pr state is similar to the pKa value of 9.13±0.02 we determined for Pfr deprotonation. From the titration curves we could determine the residual ACS Paragon Plus Environment

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absorbance of the deprotonated Pr and Pfr species at alkaline pH. The values are 26% and 2%, respectively, and were estimated as fractions of the maximum absorbance of the protonated chromophore. Residual absorbance of Pr agrees with spectrophotometric titrations of model compounds indicating that the absorbance of the deprotonated form is about a third of that of the protonated state.47, 48 Resonance Raman spectroscopic analysis of the pH-dependence of the Pr state of WT Cph1∆2. RR spectra were measured between pH 6.0 and 9.0 in steps of 0.5 units (figure 3). In general, spectral contributions of the protein and the buffer were very small except for very alkaline solutions (pH 8.5 and 9.0). Here we note a broadening and intensity reduction of the chromophore bands such that spectral contributions of the protein and the buffer gain weight. These spectral changes also include a decrease of the marker band for the protonated chromophore at ca. 1570 cm−1 that originates from the in-plane bending mode of the N-H groups of rings B and C (N-H ip).31 Whereas below pH 8.5 all four pyrrole nitrogens are fully protonated, under alkaline conditions the chromophore becomes deprotonated. However, also below pH 8.5 the spectra vary with pH, indicating a pH-dependent equilibrium between species in which the chromophore is protonated but the protonation state of a nearby amino acid changes. The latter observation fits nicely into the previously proposed scheme of a protonation-dependent equilibrium between Pr-I and Pr-II.12 Consequently, the quantitative analysis of the RR spectra is based on the assumption of two pH-dependent equilibria, i.e. (a)

Pr-I + AH+ Pr-II + A + H+

(b)

Pr-II Prdep + H+

where A denotes an amino acid residue and Prdep refers to a state including a deprotonated chromophore. Initial spectra of the three species required for a component analysis were obtained by mutual subtraction of the experimental spectra (Supporting Information). Indeed,



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subsequent refinement of the spectra within a global analysis yielded a good description of all experimental RR spectra on the basis of three species (figure 3). In this way, the three refined component spectra and their relative spectral contributions at each pH were obtained. The latter values were converted into relative concentrations and analyzed on the basis of the Henderson-Hasselbalch equation (figure 4). For the deprotonation of the chromophore (red data points), the values below pH 8.5 are associated with relatively large errors, since due to the low Raman scattering cross section of the deprotonated form, its spectral weight in the component spectra is rather low (figure 3). Nevertheless, the data in figure 4 indicate a relatively sharp pH-dependent transition, and a fit of the Henderson-Hasselbalch equation to the data between 8.0 and 9.5 (solid circles, dotted line) and in the entire pH range (open and solid circles) yields essentially the same pKa of 8.7 – 8.8, although in the latter case the slope deviates more strongly from the theoretical value of 1.0. Large errors associated with the spectra analysis can hardly account for the non-ideal behavior of the Pr-I/Pr-II equilibrium which appears to approach a constant value at pH < 7.0. We note, however, that in this case the Pr-II/Pr-I only indirectly mirrors the acid/base transition which takes place at a nearby amino acid. Thus, it may well be that the limiting value for lg[Pr-II]/[Pr-I] of ca. 0.4 simply reflects the stable conformational distribution for the fully-protonated amino acid residue. We therefore restricted the fit of the Henderson-Hasselbalch equation to the data above pH 6.5, leading to a pKa of 7.4. The RR spectroscopic experiments and the spectra analysis were repeated for samples in D2O and Cph1∆2 adducts with PCB 13C-labelled at the C5 position to assist the vibrational assignment. The component spectra of Pr-I and Pr-II are shown in figure 5 and compared with the RR spectrum measured from Pr crystals of Cph1∆2. We note an excellent agreement of the spectra of Pr-I, which is the prevailing state in solution at pH 6.0 (figure 5), and the Pr crystal. This finding is in line with the fact that crystals were grown under mildly acidic conditions.28


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It further implies that the crystal structure refers to a homogeneous state, i.e. Pr-I, which hence constitutes the reference point for the QM/MM calculations (vide infra). The vibrational spectra of Pr has been analyzed in detail before

27, 49

. The strongest band

between 1500 and 1800 cm−1 originates from a mode dominated by the C=C stretching of the CD methine bridge.18, 31 It is observed at similar frequencies in Pr-I and Pr-II (1629 and 1631 cm−1) and undergoes a 7-cm−1 downshift upon H/D exchange at the pyrrole nitrogens, due to the admixture of the N-H ip coordinates of the adjacent pyrrole rings. A second mode of lower RR intensity close to this position is due to the ring D C=C stretching which is invariant to 13C5 labelling and H/D exchange and can only be resolved upon 13C15 labelling.49 The A-B stretching may couple with the B-C stretching coordinate giving rise to two bands above and below the C-D stretching. The Raman-active high and IR-active low-frequency component are dominated by the A-B stretching and B-C stretching, respectively. These two modes reflect the main differences between Pr-I and Pr-II. In Pr-I, the high frequency component is found at 1654 cm−1, 8 cm−1 higher than in Pr-II. This high frequency of the mode in Pr-I is associated with a distinctly larger downshift (−17 cm−1) upon 13C-labelling at C5 than in Pr-II. Although for the latter spectrum the 13C5 downshift cannot be determined precisely, an upper limit of −4 cm−1 indicates a lower contribution of the A-B stretching coordinate and thus a stronger coupling of the A-B and B-C stretching coordinates in Pr-II. Accordingly, the lowerfrequency mode at 1608 cm−1 shows an appreciable downshift of −4 cm−1 in Pr-II, corresponding to a substantial contribution of the A-B stretching, in contrast to Pr-I where this mode seems to remain invariant upon

13

C5-labelling. Both modes show similar downshifts

upon H/D exchange in Pr-I and Pr-II, reflecting the admixture of the adjacent N-H ip coordinates. The main contributions of the ring B and C N-H ip coordinates are at 1566 (Pr-I) and 1570 cm−1 (Pr-II). This frequency difference is most likely due to different hydrogen bonding interactions. Both Pr-I and Pr-II show a weak band at ca.1712 cm−1. According to



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FT-IR measurements, this band can be assigned to the C=O stretching mode of ring D.50, 51 The corresponding mode of ring A, observed at higher frequencies in the IR spectra, cannot be unambiguously identified in the Pr-I and Pr-II substates. Resonance Raman spectra of the WT Pfr state. In contrast to Pr, the RR spectra of Pfr remained unchanged between pH 6.0 and 9.0 (figure S2). However, the spectra do not represent a structurally homogeneous chromophore. Although fully protonated and in a periplanar ZZEssa conformation, the cofactor in Pfr is structurally heterogeneous, as in all prototypical phytochromes studied so far.6,26,27 Specifically, two sub-states are involved which differ with respect to the AB methine bridge bond angle and the CD methine bridge dihedral angle. However, this conformational heterogeneity is not governed by (de)protonation processes of the surrounding amino acids. Only above pH 9.0 is the decrease of the N-H ip of rings B and C at 1553 cm−1 accompanied by a broadening of the C=C stretching modes, indicating the onset of the chromophore deprotonation. We estimate the corresponding pKa to be ca. 9.2 (Supporting Information, figure S2), in good agreement with the value from absorbance data (see above). The component spectrum of the deprotonated Pfr chromophore is shown in the Supporting Information (figure S4). Resonance Raman spectra of H260Q. The substitution of His260 by Gln has a pronounced effect on the pH dependence of the RR spectra of Pr. Below pH 8.5, the RR spectra do not respond to changes of the pH, indicating a single conformational state with a protonated cofactor as reflected by the N-H ip at 1575 cm−1 (figure S5). The spectrum, however, differs from both, the Pr-I and Pr-II, sub-states of the WT protein. The C-D stretching is shifted by ca. 15 cm−1 to higher frequencies such that it overlaps with the A-B stretching. This finding points to a slightly distorted geometry compared to the WT chromophore structure. The stable photoconversion product of the mutant Pr following irradiation at room temperature was clearly deprotonated even between pH 7.8 and 8.5. Only at pH 7.0 was a ACS Paragon Plus Environment

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protonated species apparent, indicating a single pH-dependent transition as in WT Pfr, albeit at a distinctly lower pKa. Back conversion of the mutant to Pr by far-red irradiation was ineffective under alkaline conditions as expected from the low extinction coefficient of the deprotonated species but was readily achieved in acidic buffers, as observed earlier33 The RR spectrum of the protonated photoconversion product of H260Q displays characteristic features of a Pfr spectrum such as the intense C-H out-of-plane mode of the CD methine bridge at ca. 800 cm−1, however, the overall comparison with the spectrum of the WT Pfr implies that the ZZE geometry is distorted (figure S3). QM/MM geometry optimization. The geometry of nine structural models of Cph1 as Pr, differing with respect to the protonation pattern of the His260 and His290, were optimized using the hybrid QM/MM approach described in the experimental section. The deviation from the X-ray structure (PDB entry 2VEA) was quantified by computing root-mean-square deviations (rmsd) of heavy atoms belonging to the chromophore, His260, and His290, after alignment of the chromophore’s heavy atoms. The resulting rmsd values for each model are listed in table 1, while relevant structural parameters involving the chromophore are listed in table 2, restricted to those models which did not change their original protonation pattern upon energy minimization. Structure of the phycocyanobilin cofactor. The structure of the PCB cofactor seems to remain unaffected by the protonation pattern of the neighboring His residues as reflected by the rmsd values for the heavy atoms and the structural parameters of the bilin backbone which are essentially the same for all models. The rmsd values for the bilin heavy atoms vary between 0.42 and 0.66 Å within the nine models. These relatively large rmsd values result from the reorientation of the propionic side chain of ring C. Here, the carboxyl group rotates and moves towards H260 to strengthen the O⋅⋅⋅Nε hydrogen bond interaction. In average, an increase of the C-C-C=O torsional angle of 23° is predicted for the QM/MM models



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compared to the crystal structure. Furthermore, independent of the protonation state of the histidines, the QM/MM calculations allow for the refinement of the structural parameters of the chromophore backbone. Geometry optimization of all QM/MM models yielded structures with longer AB C=C (0.012 Å) and shorter AB and BC C-C bond lengths (0.06 Å and 0.09 Å, respectively) than the standard restraint values in the crystal structure, implying incomplete electron π-conjugation at the CD methine bridge. Whereas the calculated tetrapyrrole geometries showed little change in A-B tilt, the B-C and C-D tilt angles were >14° greater than the experimental values. These large tilt angles, predicted for all models, result from an out-of-plane displacement of ring C towards the pyrrole water, strengthening the hydrogen bond interactions. Structure of the chromophore binding pocket. Out of the nine structural models of Cph1, the four models H260E-H290D, H260P-H290D, H260D-H290P and H260E-H290P undergo proton translocation, reflecting the unlikeliness of the corresponding starting protonation patterns. Interestingly, all four models converged to geometries in which the two neighboring histidines were protonated only at their Nδ and with a protonated carboxylic side chain of ring C (psC) or/and a deprotonated PCB chromophore backbone. In particular, in the H260PH290D and H260D-H290P models we observed that the Nε-proton of the cationic histidine, which in both cases is oriented towards the carboxylic group of the psC, is transferred either directly (H260P-H290D) or via a water molecule (H260D-H290P) to the psC (figure S7). Although these two models converge to a similar geometry, the QM/MM potential energy difference between them is very large (ca. 50 a.u.) (see Supplementary Information). This energy difference can be rationalized in terms of distinct hydrogen bonding network around the chromophore, in particular in the vicinity of the psC. Furthermore, the position of the H290 side chain in the optimized geometries is significantly shifted with respect to the crystal arrangement as reflected by the high rmsd values for the heavy atoms of 1.41 Å (H260P-


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H290D) and 1.53 Å (H260D-H290P), as also observed for the H260D-H290D and H260PH290P models. These particularly high rmsd values which are computed when the His290 is protonated only at Nδ, imply that it is highly unlikely to find a proton at this site, in contrast to the protonated structural models suggested on the basis of NMR experiments.12 For the H260E-H290D and H260E-H290P model, proton translocation events do not only involve the histidines but also the PCB chromophore. In the case of the H260E-H290D model, the geometry converges to a structure characterized by a neutral PCB backbone and a protonated (neutral) psC. The formation of this state is associated with three proton translocation events, i.e. from Nε of His260 to psC, from the pyrrole water to Nδ of His260, and from the PCB pyrrole nitrogen to the pyrrole water. The same effect is predicted for the H260E-H290P model. In this case, the additional proton contributes to the formation of an Eigen-type water cluster involving the crystallographic waters W4 and W5. These QM/MM geometry optimizations emphasize the bridging role played by the His260 and the pyrrole water for the intramolecular proton transfer within the PCB chromophore. Table 1 shows that the lowest rmsd values for the heavy atoms of the His260 and His290 are estimated for the H260E-H290E, H260D-H290E and H260P-H290E models, clearly indicating that the His290 must be protonated at the Nε, as predicted by the electrostatic calculations. Indeed, protonation at Nδ leads to large displacement of the His290 with respect to its crystallographic position as mentioned above. The position and orientation of the pyrrole water (W1) in the chromophore pocket influence the strength of the hydrogen bond interactions with the pyrroles A, B and C. The position of W1 varies strongly within the different models, as reflected by the N⋅⋅⋅O distances between the pyrrole ring nitrogens (N(A), N(B) or N(C) ) and the water oxygen. In general the N(Ring)⋅⋅⋅O(W1) distances are predicted to be 0.2 Å shorter than the 2VEA values of ca. 3.1



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Å, indicating a relatively strong hydrogen bond interaction between water W1 and the pyrrole ring nitrogens. An exception is model H260P-H290E where W1 is shifted away from N(A) by 3.4 Å. The remaining residues which were treated quantum mechanically in the QM/MM approach, namely the Asp207, Ser272, Thr274 and the Tyr263, were only slightly displaced from their original position during geometry optimization. Rmsd values of less than 1 Å were predicted, except for Thr274 (1.3 Å) and, only in the H260P-H290P model, Tyr263 (1 Å). In summary, four of the nine models considered for QM/MM geometry optimization converged to geometries with protonation patterns in disagreement with experimental evidence and two models converged to structures where the H290 is strongly displaced from its position in the crystal structure. From the three remaining models, the H260E-H290E and H260D-H290E models can be considered as potential candidates for the alkaline Pr-II substate, while only the H260P-H290E model can represent the acidic Pr-I. Analysis of calculated Raman spectra. To validate the potential structural models for the Pr-I and Pr-II substates of Cph1 we compared the respective calculated Raman spectra with the experimental RR spectra (figure 6). The corresponding normal mode frequencies and assignments are given in table S1 (Supplementary Information). The calculated Raman spectrum of the H260P-H290E model, as the only candidate for the PrI substate, provides an excellent description of the experimental frequencies and intensities and, in particular, for the isotopic shifts. This is not questioned by the underestimation of the A-B stretching mode frequency by 11 cm−1, since this deviation is still within the accuracy of the QM/MM calculations.18, 30, 31 In view of this good agreement, it appears to be justified to determine the structural model for Pr-II on the basis of the calculated Raman spectra.


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At first sight, there is a good overall match between the calculated Raman spectra of the H260E-H290E and H260D-H290E models and the pure experimental RR spectrum of Pr II. Both models reproduce most of the spectral features specifically in the region between 1500 and 1750 cm−1, namely the strong peak at 1631 cm−1 (C-D stretching), its two shoulders at 1646 and 1608 cm−1 (A-B and B-C stretching, respectively) and the band at 1570 cm−1 (N-H ip of rings B and C). An additional mode localized in ring D is predicted to be between the CD and A-B stretching but cannot be resolved experimentally. Here

13

C15-labelled PCB

cofactors are required as shown previously.49 The calculations also confirm the assignment of the weak RR band at 1712 cm−1 to the C=O stretching mode of ring D, independent of the structural model, whereas the corresponding mode of ring A is predicted at distinctly higher wavenumbers (table S1). Upon closer inspection, however, significant deviations between the two models become apparent. The C-D stretching is predicted to be at 1635 cm−1 in the H260E-H290E model, in very good agreement with the experimental data, whereas in H260D-H290E the C-D stretching is not only overestimated by 13 cm−1, but it also does not correspond to the mode of highest Raman activity, in contrast to the experimental findings. This is due to the different contributions of the C-D and A-B stretching coordinates to the corresponding normal mode. In the H260E-H290E model, the A-B stretching coordinate couples with the B-C stretching, thereby accounting for the downshift of the 1608 cm−1 band upon 13C5-labelling. Altogether, H260E-H290E reproduces the C=C stretching modes more satisfactorily than H260D-H290E, with respect to the order of the modes, [that is ν(B-C) < ν(C-D) < ν(A-B)], their isotopic shifts and their relative intensities. The N-H ip mode of the rings B and C deserves particular attention. This mode is calculated to be at 1593 and at 1559 cm−1 for the H260E-H290E (Pr-II) and H260P-H290E (Pr-I) models, respectively. The former mode is significantly upshifted with respect to the ACS Paragon Plus Environment


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experimental values at 1570 and 1566 cm-1 for the Pr-II and Pr-I substates. Since the position of this mode strongly depends on the orientation of the pyrrole water in the cavity, a more precise vibrational frequency can only be determined by averaging over different possible arrangements and orientations of the water molecule as shown previously for phycoerythrocyanin.52 The 4 cm−1 frequency difference of the N-H ip bending mode observed for Pr-I and Pr-II cannot be reproduced since the errors associated with the static QM/MM calculations are clearly larger than 10 cm−1.18

Discussion Protonation-dependent equilibria in the chromophore pocket. The present RR spectroscopic results demonstrate that in Cph1∆2 Pr forms two coupled pH-dependent equilibria between pH 6 and 9, involving (de)protonation of the chromophore and an amino acid residue adjacent to the chromophore, identified here to be H260. Accordingly, the low-pH transition between Pr-I and Pr-II refers to the (de)protonation of His260 whereas deprotonation of the chromophore (ring B or C) occurs at higher pH. Titration experiments based on RR and UVvis absorption spectroscopic monitoring afforded very similar pKa values for the respective transitions (that is ca. 7.5 and 8.9 for the Pr-II/Pr-I, and Prdep/Pr-II transitions, respectively). Also for the Pfr/Pfrdep transition nearly the same pKa of ca. 9.1 was obtained by both methods. The pKa values determined here not only for the deprotonation of the chromophore in Pfr but also for the Pr-I/Pr-II transition are in good agreement with published data. However, there are notable discrepancies for the chromophore deprotonation in Pr for which a distinctly higher pKa (+0.7 units) was previously reported.17 A possible origin for this disagreement may root in the protocol for Cph1∆2 assembly as the latter work used in-vitro assembled holoprotein. In fact, a similar observation was reported for a cyanobacteriochrome and attributed to improper folding of the protein upon in-vitro assembly.53 ACS Paragon Plus Environment

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Structural differences between the Pr substates. QM/MM calculations attribute Pr-I to a state with a cationic (protonated) His260, carrying a proton at both Nδ and Nε whereas in Pr-II the proton at Nδ is removed. In both Pr substates His290 remains neutral with the proton bound to Nε. While the description of the Pr-II substate is in agreement with the NMR data and electrostatic calculations, Pr-I differs from Song’s model, which suggested protonation of Nδ in His290.12 The present calculations showed that such a protonation pattern would lead to significant structural rearrangements of the side chains and water molecules in the chromophore binding pocket, arguing against this protonation pattern. This conclusion is in line with the fact that substitution of His260 by a glutamine abolishes the Pr-I/Pr-II transition. Moreover, the results obtained for this mutant also demonstrate the crucial role of His260 for the chromophore structure, most likely via controlling the electrostatics in the chromophore binding pocket. The structural impact of this substitution is reflected particularly by the substantial changes of the C-D stretching mode not only in Pr but also in Pfr, which in the latter state accompanied by an extraordinarily lowered pKa for chromophore deprotonation (Figures S3 – S5). Conversely, for the WT protein, the structural consequences of His260 protonation on the PCB chromophore are relatively small [by comparing H260E-H290E (Pr-II) and H260PH290E (Pr-I)]. Only a slight twist of the AB methine bridge is observed together with a reorientation of the psC pyrrole water (figure 7). This distortion leads to a downshift of the AB stretching mode by 4 cm-1, reproducing the tendency in the experimental RR spectra (∆νΑ−Β = 8 cm-1). The experimental RR spectra show that the CD methine bridge seems to be robust to pH changes, in line with the calculated structural parameters of the CD methine bridge which are essentially the same in both models (table 2). Thus, it is surprising that the C-D stretching mode is downshifted by 17 cm−1 upon protonation of the His260. The reason for this large ACS Paragon Plus Environment


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frequency shift is most likely the mixing with the C=O(C) mode predicted to be at 1621 cm−1 for the H260P-H290E model structure. On the other hand, recent polarization- and timeresolved infrared spectroscopic studies on Cph1 by Heyne and coworkers revealed the existence of two Pr ground state species, which were suggested to differ with respect to the ring D orientation.54,55 However, this conclusion cannot be confirmed by the present study. Role of His260 (de)protonation. His290 and His260 are highly conserved residues in phytochromes and are both essential for proton transfer processes associated with the last steps of the photoinduced reaction pathways. In bathy phytochromes, the decay of the Meta-F intermediate to Pr as the last step of the Pfr→Pr photoconversion includes three important events, namely the deprotonation of the ring C propionic side chain of the biliverdin cofactor, the functional secondary structure change in the tongue region, and the protonation of His290 which eventually promotes the thermal isomerization of chromophore.6 In prototypical phytochromes, however, protein structural changes in the opposite Pr→Pfr pathway follow chromophore deprotonation in the Meta-Rc intermediate. Earlier studies of Cph1∆2 showed that about two protons per monomer were released to the medium upon irradiation of Pr.17,33 These suggestions were substantiated by studies of Agp1 from A. tumefaciens.7,56,57 Only one of them was uptaken again upon Pfr formation, which leads to the reprotonation of the chromophore. This process is perturbed in the H260A mutant. It is therefore concluded that His260 is essential for proton re-uptake from the solution phase. In the WT protein, the chromophore is protonated in the pH range up to 9.0 in both Pr and Pfr. Thus, upon Pr→Pfr photoconversion at physiological pH, deprotonation of the chromophore occurs only in the Meta-Rc intermediate (see figure S6). Meta-Rc is associated with the release of two protons to the medium, however, only one of which is likely to originate from the chromophore. Subsequent decay of Meta-Rc to Pfr is associated with re-uptake of approximately one proton per monomer, so that Pfr formation is accompanied by the net


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release of ca. one proton to the solution phase.17 Thus, an alteration of the protonation pattern in the chromophore binding pocket is required to ensure the fully protonated chromophore in Pfr. It is very likely that this pattern involves a deprotonated His260, in line with the fact that re-protonation of the chromophore in Pfr is perturbed in the H260Q variant of Cph1∆2 and even blocked in the H260A variant of Agp1.33, 56 Furthermore, unlike to the conformational equilibrium in Pr that is controlled by the protonation state of His260, the chromophore structure in Pfr does not respond to variations of the pH, pointing to a pKa of His260 below 6.0. This observation is in agreement with previous NMR studies suggesting a unique neutral form of His260 in Pfr.12 Thus, this interpretation suggests that the lowering of the pKa of His260 during the Pr→Pfr phototransformation is the driving force for the proton release to the solution phase. The original pKa is then recovered upon back conversion to Pr, thereby completing proton uptake from the solution to satisfy the proton balance for the entire photocycle. Altogether this hypothesis may guide further experiments, also directed to putative role of proton translocations for functionally relevant structural changes during photoconversion in prototypical phytochromes. Acknowledgements The work was supported by the Deutsche Forschungsgemeinschaft (SFB1078 A2, C2, C3, B6 to MAM, UA and PH, Hu702/8 to JH, Mr81/3-1 to MAM). The authors thank Wolfgang Gärtner for providing Cph1 adducts with

13

C5 labeled PCB and Ernst Walter Knapp for

fruitful discussions. The computations were performed with resources provided by the NorthGerman Supercomputing Alliance HLRN. Supporting Information Available Supporting information includes a detailed description of the spectra analysis, as well as figures with pH-dependent RR spectra of Pfr, the complete component spectra of the



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protonated chromophore species (Pr, Pfr) of the WT Cph1∆2 and the H260Q variant, the pH dependent RR spectra of Pr state of H260Q- mutant and WT-Cph1∆2, RR spectra of the Meta-Rc intermediate of WT-Cph1∆2 in H2O and D2O and a figure of the chromophore binding site of QM/MM optimized H260P-H290D and H260D-H290P structural models with the corresponding energy values. The results of normal mode analyses for the Pr-I and Pr-II substates are listed in table S1. This information is available free of charge via the

Internet at http://pubs.acs.org

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Figure captions Figure 1: (A) Chromophore binding pocket of Cph1∆2 Pr crystal structure (pdb entry: 2VEA). Chromophore as well as the Cys259 binding site are shown in grey, whereas remaining conserved or relevant residues are displayed in yellow. Crystal waters (W1 to W7) in close vicinity to chromophore are plotted as spheres in cyan. The figure was generated with VMD1.86.58 (B) Structure, pyrrole ring and carbon-atom notation of the protonated ZZZssa PCB chromophore. Figure 2: Top panels, pH-dependent absorption spectra of Pr (A) and Pfr (B). The direction of the pH-titration is indicated by the black arrow. Bottom left panel (C), the nine wavelengths for Pr ( 638(),643.5(), 649(), 654.5(), 660(), 665.5(⊳), 671(), 676.5() and 682


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nm() with corresponding fit) used for a simultaneous fitted of the Henderson-Hasselbalch equation to calculate the two pKa values (pK1 = 7.55 ; pK2 = 9.04 ). Three traces at selected wavelengths (638 nm, 660 nm, and 676.5 nm) are marked in bold to highlight that the titration spectra are complex and strongly depend on the wavelength. The fit, which assumes two titratable groups, accounts for this complexity. Bottom right panel (D), Thick traces mark the three diagnostic wavelengths, which show the shift of the pK1. Bottom right panel, the nine wavelengths for Pfr (682(),687.5(), 693(), 698.5(), 704(), 709.5(⊳), 715(), 720.5() and 726 nm() with corresponding fit) used for a simultaneous fit of the Henderson-Hasselbalch equation to calculate the pKa (pKa = 9.13). Figure 3: RR spectra of Pr Cph1∆2 measured at different pH in H2O. The experimental spectra (black curves) are shown together with the fitted component spectra of Pr-I (dotted red), Pr-II (dotted blue), and Prdep (dotted green). Figure 4: Plot of the concentration ratios of Pr-I/Pr-II (blue) and Pr-II/Prdeprot (red) of Cph1∆2 versus the pH according to the Henderson-Hasselbalch relation. The dotted blue and red lines represent linear fits. The data displayed by open circles were not considered, as described in the text. The dotted grey lines indicate the respective pKa values. Figure 5: Pure RR spectra of the Pr-I (left) and Pr-II (right) as determined by the component analysis (see text and Supporting Information for further details). The middle black traces show the spectra Cph1∆2 obtained in H2O (natural abundance – n.a.), the blue bottom traces the spectra from samples in D2O (N-D), and the red top traces the spectra measured from 13

C5-labelled PCB adducts of Cph1∆2 in H2O (13C5). The grey line in the left pattern displays

the spectrum measured of Cph1∆2 (Pr) crystals. Figure 6: Experimental RR spectra of Pr-I and Pr-II compared with the calculated Raman spectra of the H260P-H290E model (Pr-I), and the H260E-H290E and H260D-H290E models ACS Paragon Plus Environment


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(Pr-II). The spectra of Cph1∆2 adducts with non-labeled PCB, N(D)-labeled PCB,

13

C5-

labeled PCB are shown by black, blue, and red traces, respectively. Figure 7: Structure of the chromophore binding site in the H260E-H290E (pink) and H260PH290E (cyan). The oxygen atom of the pyrrole-water in the H260E-H290E (H260P-H290E) model is depicted as a pink sphere. The two structures are aligned with respect to the PCB heavy atoms.

Tables Table 1: Root mean square deviations (Å) of the position of heavy atoms from PCB, His260, and His290 relative to the crystal structure geometry. Structural models and crystals structure were aligned at PCB heavy atoms using VMD software.58 Italic letters highlight structural models where protonation pattern was altered in the course of the geometry optimization. H260D- H260E- H260D- H260E- H260P- H260D- H260E- H260P- H260PH290E H290E H290D H290D H290D H290P H290P H290E H290P PCB

0.49

0.57

0.63

0.61

0.62

0.42

0.66

0.64

0.55

H260 1.11

0.67

1.15

0.63

0.72

0.78

1.03

1.06

0.49

H290 1.04

0.80

1.50

1.83

1.41

1.53

1.10

0.82

1.88


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Table 2: Relevant structural parameters of the PCB cofactor and its environment. Bond lengths and distances are given in Å and angles in degree (°). Deviation of average values with respect to the crystal structure (∆) are listed in the last column. H260D- H260E- H260D- H260P- H260P- Average Exp H290E H290E H290D H290E H290P

∆

(2VEA)

AB tilt

8.8

16.6

12.4

6.9

12.0

11.4

11.4

0

BC tilt

24.6

22.6

15.9

13.2

25.6

20.4

1.7

18.7

CD tilt

38.3

37.4

44.8

38.8

40.3

39.9

25.9

14.0

C=O(A) tilt

84.9

94.3

91.4

92.2

90.7

90.7

87.1

3.6

C=O(D) tilt

94.2

94.1

85.3

92.7

90.7

91.4

89.3

2.1

N(A)⋅⋅⋅O(W1) 2.879

2.886

2.900

3.410

2.980

3.011

3.047

-0.036

N(B) ⋅⋅⋅O(W1) 2.960

2.824

2.930

3.074

3.074

2.973

3.170

−0.197

N(C) ⋅⋅⋅O(W1) 2.932

2.887

2.916

2.981

2.981

2.939

3.140

−0.201

C=C (AB)

1.361

1.362

1.354

1.357

1.357

1.358

1.346

0.012

C-C (AB)

1.437

1.435

1.436

1.423

1.424

1.431

1.494

−0.063

C-C-C (AB)

136.1

132.9

134.4

130.8

128.73

132.6

127.89

4.7

C-C (BC)

1.400

1.399

1.402

1.385

1.389

1.395

1.484

−0.089

C-C (BC)

1.392

1.392

1.399

1.396

1.404

1.397

1.469

−0.072

C-C-C (BC)

134.0

134.6

133.3

132.8

134.53

133.9

136.02

-2.2

C=C (CD)

1.359

1.360

1.356

1.368

1.363

1.361

1.479

−0.118

C-C (CD)

1.437

1.439

1.440

1.443

1.445

1.441

1.485

−0.044

C-C-C (CD)

128.6

131.8

127.7

131.3

130.12

129.9

129.59

0.3

C=O(A)

1.215

1.220

1.222

1.212

1.215

1.217

1.329

−0.112



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C=O(pscB)

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1.254

1.251

1.259

1.249

1.250

1.249

1.249

0.004

1.269

1.274

1.270

1.277

1.269

1.253

1.253

0.019

1.281

1.284

1.269

1.278

1.286

1.280

1.248

0.032

1.247

1.246

1.267

1.252

1.246

1.252

1.248

0.004

C=O(D)

1.231

1.232

1.226

1.231

1.245

1.233

1.479

−0.246

CC-CO2

140.6

119.9

120.4

101.9

135.9

123.7

177.5

−53.8

138.0

150.0

141.5

174.7

125.2

145.9

122.7

23.2

H-Nd(H260)

1.011

-

1.008

1.033

1.026

H-Ne(H260)

-

1.032

-

1.030

1.056

H-Nd(H290)

-

-

1.021

-

1.027

H-Ne(H290)

1.046

1.036

-

1.033

1.059

C=O(pscC)

(pscB) CC-CO2 (pscC)


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Figures FIGURE 1:



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Figure 2


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Figure 3



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Figure 4


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Figure 5



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Figure 6

Figure 7


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