Article pubs.acs.org/JPCB
On the pH Dependent Behavior of the Firefly Bioluminescence: Protein Dynamics and Water Content in the Active Pocket Hyun Woo Kim and Young Min Rhee* Center for Self-Assembly and Complexity, Institute for Basic Science (IBS), Pohang 790-784, Korea, and Department of Chemistry, Pohang University of Science and Technology (POSTECH), Pohang 790-784, Korea S Supporting Information *
ABSTRACT: Understanding bioluminescence presents fascinating challenges for fundamental sciences and numerous opportunities for practical applications. As a representative example, the firefly bioluminescent system has been intensively studied in both experimental and computational areas. However, there are still remaining questions regarding especially the detailed protein dynamics and the mechanisms of its color modulation. Here, we report on the pH dependent behavior of the firefly bioluminescence primarily based on molecular dynamics simulations. We find that the overall protein structure is generally resilient to pH variations. As the protein does not exhibit any structural distortions that can affect the emission property, we next focus on the dynamics in the active pocket and its effect on color modulation by adopting different protonation states in the pocket. With this, we observe red-shifted emissions at acidic conditions as consistent with previous studies. Most importantly, we find that a water molecule in the active pocket can mediate flexible motions of neighboring groups, which can subsequently modify the emission properties to a substantial degree. Based on the observations, we propose that the active pocket is in a dry condition during the luminescence process. Our results highlight the importance of understanding the role of the dynamics near the active pocket in modulating bioluminescence.
1. INTRODUCTION Bioluminescence has become an important research theme in both theoretical and experimental chemistry,1−3 fueled by its potential applications in various fields.4,5 For practical applications, bioluminescent systems have several advantages with various interesting chemical behaviors. They usually show high quantum yields up to ∼41% as experimentally confirmed in the firefly case.6 It also exhibits a feasible color modulation capability from green to red in response to external stimuli such as pH.6 In fact, several creatures in nature already display luminescence with notably different emission frequencies even with a single luciferin species.1,7,8 Such richness in chemistry indeed necessitates detailed mechanistic studies at various stages of bioluminescence. Understanding fundamentals will be of course helpful in developing next level applications. In regard to figuring out the fundamentals, among many bioluminescent systems, many researchers have focused on the firefly luciferase−luciferin system as a representative example. In the firefly system, with its recently elucidated transition state analog structure,9 the luciferin substrate undergoes adenylation and oxidation processes inside the luciferase to generate the electronically excited oxyluciferin. Subsequently, this excited oxyluciferin emits light and falls down to its ground state. In Scheme 1, we have shown a general mechanism for explaining the generation of the excited oxyluciferin. This mechanism, however, still does not explain the color modulation effect of firefly bio© XXXX American Chemical Society
luminescence. Moreover, there is not even a complete consensus on the exact chemical structure of intermediates and the actual emitting form of the oxyluciferin.1−3,10 For example, there is a recent discussion that the firefly dioxetanone is neutral during the oxyluciferin generation.11 Thus, more intensive studies in both experimental and computational areas are still highly required. To reveal the process of the oxyluciferin production in detail, high-level computations have been continuously found to be fruitful. For example, Morokuma and co-workers studied the decomposition of a firefly dioxetanone to the excited oxyluciferin with the exploration for energy surfaces involved in the oxyluciferin generation.12 In addition, systematic studies on decompositions of several 1,2-dioxetanes and 1,2dioxetanones were performed by Lindh and colleagues13−15 and by Yamaguchi and co-workers.16 With these, a charge transfer induced luminescence (CTIL) mechanism was suggested as a plausible explanation for the oxyluciferin production.1 Recently, Yue et al. computed the reaction pathway of oxyluciferin generation with a similar approach as the one taken by Lindh and co-workers.17 Their results showed that a deprotonated keto form of the oxyluciferin can be produced more plausibly than its neutral form. They also Received: March 11, 2013 Revised: May 23, 2013
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Scheme 1. General Mechanism of Oxyluciferin Generation in Fireflies
these computations have focused on a small number of specific conformations of the oxyluciferin/luciferase complex unavoidably yields certain level of limitations. To overcome these limitations, directly inspecting the dynamical behaviors of the protein−ligand complex will be highly desirable. In this respect, similarly to studying other biological complexes,36−39 molecular dynamics (MD) simulations can complement previous studies in elucidating the relationship between the emission color changes and the dynamic aspects of the luciferase−oxyluciferin system.40 In this article, we report our study on the pH dependence of the firefly bioluminescence, with the purpose of revealing the origin of the color changes and the role of the protein dynamics in various pH conditions. We first investigate structural effects of different protonations on the luciferase by obtaining plausible protonation states in a range of pH conditions and by subsequently performing MD simulations in those states. We also inspect structural effects induced by altering protonation on a histidine residue and AMP in the active pocket to examine the specific role of the species inside the pocket. In all adopted protonation states, we find that the luciferase maintains its structure without any meaningful distortions. As different protonation states affect insignificantly on the overall protein structure, we then scrutinize electronic structure modification effects on oxyluciferin caused by protonations on its neighboring residues. With this, we demonstrate that differently protonated proteins can modulate the emission energy to a large enough extent to explain the color changes at acidic and basic conditions. Most importantly, we find that water content in the active pocket can affect the side chain flexibility inside the pocket and subsequently modify the color modulation pattern. Specifically, we observe that a single water molecule can lead to flexible motions of oppositely charged residues with respect to the oxyluciferin molecule, resulting in reductions in the amounts of red shifts at low pH conditions. On the contrary, when the active pocket is in a dry condition, it forms a rigid environment around the oxyluciferin and sensitively affects the luminescence. These findings emphasize the importance of the relationship between color changes in various pH conditions and the dynamical aspects near the active pocket. Such understanding will surely provide essential insights in elucidating the detailed mechanism of the bioluminescence.
suggested a modified mechanism, a gradually reversible charge transfer initiated luminescence (GRCTIL).17 With an analysis at DFT computational level, Esteves da Silva and co-workers proposed interstate crossing-induced chemiexcitation (ICIC) mechanism.18 On the other hand, Branchini et al. demonstrated experimentally that the luciferase may alter its conformation during the oxidation step from luciferin to oxyluciferin due to the conformational restriction from the luciferase,19 with the crystal structure of this alterative conformation.20 These experimental and computational studies have indeed provided important insights in explaining the early stage of the firefly bioluminescence with a viewpoint on both the emitter and the protein. Once the oxyluciferin is formed, it will stay in the excited state with a lifetime of nearly 10 ns.21 In a sense, this is a long period of time, and many dynamical changes can take place during that period. In fact, it is natural to anticipate that one of the elusive problems of firefly bioluminescence, namely the color modulation, will be related to this dynamics of nascently formed oxyluciferin. According to recent reports,9,21−24 the emission frequencies are affected by the environment near the luciferase and/or by the form of the oxyluciferin among several possible isomers and tautomers. In fact, there are several possible mechanisms and debates are still ongoing. For details of discussion on various suggestions and their limitations, readers are referred to comprehensive summaries in recent articles,25,26 and we will only comment on brief aspects which are relevant to our discussion in this article. Experimentally, it is observed that firefly bioluminescence can span the range from green to red depending on structural mutations9 and external stimuli such as pH changes.6 At an acidic condition (pH ∼6.5) luciferase−oxyluciferin system emits in red while yellow-green light is observed at pH ∼8.6 Recently, Milne et al. explained this pH dependence by investigating the detailed contribution of residues and other nearby molecules and suggested that the protonation state of AMP may induce significant changes in emission colors.26 They reached to this conclusion by assuming that important factors in the color modulation are the residues in the active pocket of luciferase at a specifically fixed conformation of oxyluciferin and luciferase. Because important residues were explicitly considered, computations based on this assumption can provide more information than the ones carried out only with oxyluciferin in the gas phase or in approximated protein environments.25,27−33 There are several quantum mechanical/molecular mechanical (QM/MM) computational studies that included all residues of luciferase and nearby solvent molecules for better understanding the color modulation in the firefly bioluminescence.23,24,34,35 Still, the fact that
2. COMPUTATIONAL DETAILS MD Simulations. As a starting conformation, we chose the crystal structure of oxylucyferin − luciferase found in a firefly (PDB ID: 2D1S),9,41 attached hydrogen atoms toward proper B
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valencies, and placed the complex in an octahedron box filled with 23992 water molecules. We described luciferase and AMP(−2) with the AMBER99SB force field, which was shown to give reliable protein structures.42 The force field for modeling AMP(−1) was simply obtained by adopting parameters of similar atoms and by changing atomic charges of the phosphate group as listed in Figure S1 and Table S1 in the Supporting Information. For water molecules, we used the TIP3P model.43 To describe the force field of the oxyluciferin (OLU), we adopted the same parameters that were utilized in MD studies of the firefly bioluminescent system.40,44 In ref 44, the OLU model was developed in such a manner that the parameters are determined independently from the protein model. The only exception is on the Lennard-Jones parameters as they are optimized in reference to both quantum chemical calculation data and a few selected amino acid side chains.44 Even though the OLU model was derived in reference to side chains with OPLS,45 the effect on the excited state dynamics caused by the slight mismatches in Lennard-Jones parameters against the AMBER model is usually only marginal.40,46 For the chemical form of OLU, as we were focusing on the pH dependent structural behavior of the luciferase and its effect on the luminescence, we considered only the case where OLU is in the phenolate-keto form. This is in part to simplify the problem by assuming a single form of the emitter, while diversely varying the form of the protein. Also, this is in line with previous studies which have suggested that the phenolate-keto form of OLU will be the most probable emitter.23,40 After this modeling, we performed MD simulations at various protonation states. We will describe the way of obtaining protonation states in a later part of this section. In any case, for each given protonation state, we carried out an equilibration MD simulation for 1 ns to eliminate any potential clash that the PDB-derived structure might contain. This was followed by two types of production MD simulations. In the first case, relatively long MD trajectories of 50 ns were simulated to investigate the structural change of the luciferase in protonation states from pH 5 to 9. The primary purpose of this rather long simulation was to check the impact on the protein structure from different protonation states at different pH conditions. We limited the number of individual trajectories to five due to the practical consideration of the computational cost. In the second type of simulations, we focused on the dynamics during the luminescence lifetime. We carried out MD simulations for 10 ns by varying protonations near OLU. In this case, we performed ten individual MD simulations for each given protonation state. In both types of simulations, the time step was 2 fs with constraints on chemical bonds involving hydrogen atoms with the LINCS algorithm.47 Temperature and pressure were maintained at 298.15 K and 1 atm with Berendsen’s weak coupling schemes.48 Long range interactions were tapered from 10 Å and were gradually decreased to zero at 12 Å. We performed all MD simulations with GROMACS software package.49 Protonation State Determination. Since the luciferase contains ∼100 titratable residues (Asp, Glu, Lys, and His), there are numerous possibilities for differently protonating all the residues. Therefore, we generated plausible protonation states in a range of pH by applying a Monte Carlo (MC) method.50 This method is based on intrinsic pKa of acidic and basic residues and their Coulomb interactions. In fact, similar methods have been continuously adopted by other researchers.51−55 Intrinsic pKa and residue−residue Coulomb inter-
actions were computed with the program TAPBS56 which solves the linearized Poisson−Boltzmann equation (LPBE). We utilized the optimized structure at molecular mechanics (MM) computational level. Because the geometry optimization requires an initial starting protonation state as an input, it was assumed that all titratable residues except histidines were in their forms at pH 7. For histidines, we added two protons at nitrogen atoms in the side chain. The geometry and the protonation states were then searched iteratively. In addition, atomic charge lists of differently protonated residues as inputs for solving LPBE were derived from AMBER99SB force field.42 The intrinsic pKa values thus found and the Coulomb interactions are used as inputs of Karlsberg 2.050 for determining possible protonation states within an MC scheme. After performing the MC simulation at the optimized conformation, we determined possible protonation states of the luciferase in five conditions from pH 5−9. Comparing the initial and final protonation states from our iterative processes, we found that the state of His247 was prone to changes at pH 5−7. Indeed, because His247 is inside the active pocket, and because the MC scheme has some level of uncertainty, it will be desirable to carefully investigate the role of its protonation. Thus, for checking potential conformational changes with pH 5−7 conditions, His247 was set to its protonated form. After observing that the conformation does not depend on pH to a meaningful degree, all possible protonated forms of this residue were examined again at pH 7 to inspect the emission changes by protonations. See the next section for more details. In addition, it should be noted that our scheme for determining protonation state may sometimes fail to converge as in many other iterative procedures. In fact, at pH 5, the MC scheme suggested a few protonation states in an oscillatory manner. In this case, we chose the most heavily protonated one among those states. This was based on reasoning that structural distortion at lower pH will likely be more severe with more protonations on the protein. In the end, we will show that the protein structure is robust against the pH changes even with such multiple protonations. More detailed analysis on the protein structure and the protonation states will of course be given in the Results and Discussion. Emission Energy Calculations. We adopted a QM/MM approach in computing emission energies of OLU with the protein and solvent. For this, we considered various conformations that were sampled at 100 ps intervals along each 10 ns long MD trajectory. For the QM part of calculations, RI-CIS(D) method was applied with the 6-31+G(d) basis set. As noted previously,40 this method can explain the dynamical electron correlation at a reasonable computing cost. However, RI-CIS(D) systematically underestimates the emission energy. Therefore, we corrected the RI-CIS(D)/6-31+G(d) results so that they can have similar reliability as obtained with the EOMCCSD method57 with the same basis set. Our correction scheme for obtaining the emission energy (E) can be written as E = EQM/MM + ΔEgas
(1)
where EQM/MM represents the energy obtained at the RICIS(D)/6-31+G(d) level in QM/MM. The correction factor, ΔEgas, adjusts the amount of underestimated emission energy of a given conformation of OLU. This correction factor can be found based on the near-linear relationship between EOMCCSD and RI-CIS(D) emission energies of isolated OLU in the gas phase. (See Figure S2 for more details.) For quantum C
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significantly narrowed into a plausible subset. After reducing the number of candidates in this manner, a small set of titratable sites with more importance can be additionally considered in a full-scan manner. To investigate the effect of explicit protonations on the structure of the luciferase, we have simulated 50 ns MD trajectories with these MC-determined states. This consideration is crucial as changes on luciferase structure may alter emission patterns to a significant degree. To be complete, instantaneous deprotonation and reprotonation from the dynamic aspect of equilibrium will need to be included. For simplicity, however, we assumed that protonation states remain the same during the simulation time scale. We examined the overall structural changes by following root-mean-square deviation (RMSD) of the luciferase backbone along MD trajectories. The reference conformation for computing RMSD was the optimized structure at MM level which was adopted in obtaining the representative conformation during the MC scheme application. Average RMSDs obtained from analyses of five MD trajectories in each pH condition are shown in Figure 2a.
chemical calculations, we have utilized the developers’ version of Q-Chem 4.0.58
3. RESULTS AND DISCUSSION Structural Effect of Protonations on Luciferase. As explained in the previous section, we determined possible protonation states of the luciferase−oxyluciferin system in its optimized conformation at the MM computational level. We observed that a few titratable residues are in states that are different from their conventional protonation states at pH 7. These residues are listed in Table 1 and pictorially illustrated Table 1. Protonation States of Titratable Residues in Different pH Conditions, Obtained by the Monte Carlo Approach pH titratable residues
5
6
7
8
9
Glu89 Glu457 His118 His214 His223 His246 His247 His421 His433 His463 Lys432
GluH GluH HisP δ-His ε-His ε-His HisP HisP HisP HisP LysH
GluH Glu ε-His δ-His ε-His ε-His HisP HisP HisP ε-His LysH
Glu Glu ε-His δ-His ε-His ε-His HisP HisP HisP ε-His LysH
Glu Glu ε-His δ-His ε-His ε-His ε-His HisP HisP ε-His LysH
Glu Glu ε-His δ-His ε-His ε-His ε-His ε-His HisP ε-His Lys
together with OLU in Figure S3 of the Supporting Information. In Figure 1, we show the nominal abbreviations that we will
Figure 2. Backbone RMSDs of the luciferase with different protonation states on titratable residues (a) at pH 5−9 conditions and (b) at pH 7 condition plus all possible protonation states of His247 and AMP.
The result shows that RMSDs do not show significant deviations and their values remain at around 1.5 Å. This implies that the luciferase−oxyluciferin system does not exhibit any meaningful structural distortions in this pH range. This is likely because the pH change affects at most 11 residues out of more than 500 residues in the luciferase. In addition, RMSD values have already reached almost the plateauing value at the beginning (time zero in Figure 2a). As the time zero configuration has been actually obtained with 1 ns equilibration simulation, we can infer that the small structural deviation of ∼1.5 Å is caused by relatively fast local structure fluctuations, not by slow and long-ranged motions. Among titratable residues listed in Table 1, His247 is actually close to OLU while other residues are relatively far from OLU. In addition, it can be inferred from the crystal structure that His247 may form a hydrogen bond with the neighboring
Figure 1. Titratable residues and their abbreviations adopted in this study. Structures of the two additional conjugate forms, anionic Glu and cationic LysH, are not shown.
adopt for discussion in this section. There are a number of aspartate residues in the luciferase, but all of them were suggested to exist in deprotonated forms according to the MC procedure. From Table 1, one can notice that protonations of only a few residues are affected in the pH range from 5 to 9. We note again that our purpose of adopting MC scheme is not to find the exact protonation state of the luciferase in the given pH. Ideally, testing structural effects of every different possible protonation would be more desirable and less prone to computational errors. However, this will lead to an impractically large set of simulations with ∼2100 ≈ 1030 candidate states. By adopting the MC scheme, the range of candidates can be D
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AMP.9 Due to these proximities to OLU and AMP, His247 may exert a significant effect on the active pocket structure when its protonation state is altered. AMP may also affect the protein structure through pH dependent protonation change, from AMP(−2) to AMP(−1). To inspect these possibilities, we have additionally simulated 50 ns MD trajectories by varying the protonation states of these two groups while fixing the protonations on other titratable residues to the pH 7 condition. Figure 2b compares RMSD variations of the luciferase backbone thus obtained with respect to the optimized structure of the luciferase. For conformations with ε-His and HisP, RMSDs stay almost constant around at 1 − 2 Å regardless of the protonation on AMP. On the contrary, it is interesting to see a noticeable RMSD difference with δ-His in response to the change from AMP(−2) to AMP(−1). When the deviating trajectory was closely inspected, we found that this different behavior in RMSD was related to a domain movement19,20 of the luciferase (Figure S4). RMSDs also do not vary much within ∼10 ns, additionally suggesting that the deviation is likely related to relatively slow and large-scale motion. This peculiarity can be understood from the lack of the hydrogen bond between AMP and δ-His247/OLU. Because neither of δHis and AMP(−2) possesses a donatable hydrogen atom toward neighboring H-bond acceptors, structural tightness will be different compared to the cases with differently protonated His247 forms or AMP(−1) in terms of H-bonding capabilities. However, this RMSD deviation was observed only in one of the five attempted independent trajectories and care must be taken when this aspect is to be considered in a statistically meaning way. In addition, the lack of the H-bond will be an energetic disadvantage for the δ-form of His247 with AMP(−2), and this protonation state will not likely be an important participant in the actual process.59 Even still, it is interesting to see that δ-His and AMP(−2) in the pocket may induce a large enough structural distortion in the active pocket which may subsequently trigger a domain motion in the overall protein structure. Overall, we have observed that protonations on titratable residues may provide only minor structural effects on the luciferase. Even with changes on His247 which is in the active pocket, the structure is still relatively well-maintained as long as the energetically stabilizing H-bonding in the active pocket is preserved. This suggests that the experimentally observed pH dependent luminescence change will not likely come from the protein structure modification. Then, what characteristics are the sources of the observed change in the emission color? Indeed, the luciferase possesses multiple sites in the active pocket that can vary their protonation states in a number of different ways. Even though such differences are not inducing large structural modifications, they may still induce color modulations at least from the already known electrostatic effects.24,26,34,40,60 Therefore, we will next focus on color modulations induced by the charge state changes on the titratable residues in the active pocket. Emission Color Changes upon Protonations in the Active Pocket. To reveal the effect of protonations in the active pocket, we chose a small set of important groups in that region (Figure 3). Among these, we first focused on His247, Lys531, and AMP. In addition, we considered a water molecule in the first solvation shell of OLU, which is denoted as wat1 in Figure 3. Wat1 can be important in the dynamics of the active pocket because water may not freely move near the thiazole ring.40 To reduce the number of possible combinations of
Figure 3. Structure of the active pocket in the luciferase−oxyluciferin system. The residues that are explicitly discussed in the text are shown with their abbreviated names. For visual clarity, hydrogen atoms are not displayed. The figure was generated with the VMD software.61
protonation states, we ignored potential deprotonation of arginines because an isolated arginine has a relatively high pKa value (∼12) compared to other less basic residues such as Lys and His. Glu313 is set to its deprotonated form. However, the issue of the protonation state of Glu313 deserves some attention, and we will discuss it in a later part of this section. We prepared 16 different states as listed in Table 2 by varying protonations on AMP, Lys531 and His247, and by Table 2. Adopted Combinations of Protonation States and the Absence/Presence of a Water Molecule in the Active Pocket, Together with Their Average Emission Energies entry His247 no water molecule in the active pocket
with a water molecule in the active pocket
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
ε-His ε-His ε-His ε-His δ-His δ-His HisP HisP ε-His ε-His ε-His ε-His δ-His δ-His HisP HisP
Lys531
AMP
average emission energy (eV)
LysH LysH Lys Lys LysH LysH LysH LysH LysH LysH Lys Lys LysH LysH LysH LysH
AMP(−1) AMP(−2) AMP(−1) AMP(−2) AMP(−1) AMP(−2) AMP(−1) AMP(−2) AMP(−1) AMP(−2) AMP(−1) AMP(−2) AMP(−1) AMP(−2) AMP(−1) AMP(−2)
1.93 2.01 2.02 2.01 1.95 2.00 1.87 1.89 1.95 1.97 1.99 1.99 1.94 1.97 1.92 1.90
considering the absence or presence of wat1. Of course, we do not mean that all these states may appear in the physiological condition. The intention is to check whether there is a key residue that may affect the emission more strongly than others. In any case, when we tried these different forms with MD simulations, we found notable changes in the emission energy. Because all the distributions of the emission energies can be reasonably well represented with single Gaussian functions (Figures S5 and S6), we will only use the average values in our discussion in the following. Table 2 shows these computed average energies, and one can see that the values range from 1.87 to 2.02 eV. Although this range is somewhat in the redder side than what has been observed with experiments, the E
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His247, it is easily observed that HisP induces red side emissions while both ε-His and δ-His similarly exhibit emissions in the bluer side. This trend with His is not noticeably affected by the presence of wat1 molecule in the active pocket. Overall, with Table 2, one can observe that the emission shifts to the red side when the number of protonation sites increases, and that the luciferse−oxyluciferin system with AMP(−1), LysH, and HisP exhibit redder emissions than systems with their deprotonated counterparts. This is consistent with the trend reported in previous experimental6 and computational studies,26 which demonstrated that the firefly bioluminescence became red at acidic conditions while it is yellow-green at the physiological condition. This is also consistent with physical reasoning that protonations near the thiazole oxygen of OLU can stabilize the excited state more than the ground state.40,44 It is also worth mentioning that we have not exhausted all possible protonation states in the active pocket. For example, we did not include hydronium ion explicitly in our simulation. However, considering the trend observed in the above, other protonations near the oxygen at the thiazole moiety of OLU will likely cause increased red shifts in the emission. So far, we have focused on variations near the thiazole side of the luminophore. The benzothiazole side is surrounded by water with higher mobility40 and the emission will be affected by the average equilibrium nature of the surrounding groups. This calls for additional investigations on the effects of protonation in that region. For this purpose, we have prepared an imaginary system with protonated Glu313, which is also titratable and relatively close to the benzothiazole ring as mentioned before. The distance between OLU and Glu313 is ∼7 Å when measured with the distance between neighboring oxygen atoms in the two groups at the optimal conformation obtained at the MM computational level. The result shows that the protonation at Glu313 can blue-shift the emission by about 0.06 eV (Figure S7). This shift can be well explained by the electrostatic interaction: the emission becomes bluer when the electrostatic potential on the oxygen atom of the benzothiazole ring becomes higher.40 At a first glance, this blue shift may appear to be contradictory to the trend that has been observed with experiments. However, we should note that the nominal pKa of Glu is quite lower than the pH values explored in experiments.6 In addition, there are neighboring two arginine residues (Arg220 and Arg339) around oxygen on the benzothiazole ring of OLU. Because these arginines will be in protonated forms at low pH conditions, their positive charges will hinder the entrance of additional protons near OLU. Thus, the protonation on Glu313 is even less likely. Of course, the positive charges on arginine residues should already have the same blue-shifting influence on the emission energy, and this is in good accord with the fact that OLU emission is on the bluer side with the protein in the physiological condition than without the protein. In any case, we can conclude that the pH changes will not likely affect the emission through modulation on the benzothiazole side unless extremely low pH condition is adopted. At such extreme conditions, the luciferase will likely denature first. Dynamics of Charged Species and Their Effects on Emission. From the analysis of snapshots from MD trajectories, we showed that the low pH condition can induce considerable red shift (∼0.15 eV) when the active site is in a dry condition. However, protonation state changes with water in the pocket do not contribute significantly to the color
modulation range (0.15 eV) is quite close to the experimentally observed emission energy change of ∼0.2 eV.6 The systematic underestimation stems from the error of the adopted level of theory. However, even with the systematic error, the adopted method is still sensitive to the change of the emission environment,62 and the spectral shifts can be reliably adopted for further analyses. In Table 2, the combination of HisP/ LysH/AMP(−1) without wat1 shows the red-most emission at 1.87 eV while the blue side extreme at 2.02 eV is observed when ε-His/Lys/AMP(−1) are in the luciferase active pocket. Combinations of ε-His/Lys/AMP(−2) and ε-His/LysH/ AMP(−2) also exhibit emissions very close to the blue side extreme (2.01 eV). When we consider only the cases without wat1, the appropriate protein form in the high pH limit will contain ε-His/Lys/AMP(−2),63 while the form in the low pH limit will be with HisP/LysH/AMP(−1). These two cases, which correspond to entries 4 and 7 respectively in Table 2, are essentially at blue side and red side extremes (Figure 4). Thus, we can infer that the experimentally observed color modulation occurred from the combined effects from multiple residues.
Figure 4. Emission energy distributions obtained with low and high pH limiting conditions.
One may wonder that this idea of multiple protonations with a pH change by one or two units may be too extreme as additional protons at neighboring residues need to overcome severe electrostatic repulsions. However, a case with multiple protonations in neighboring residues is not difficult to find out. For example, in human galectin-1, there are two histidine residues which are sequentially protonated at pH 6.3 and 5.7.64 This can be achieved by residues with similar pKa values when the electrostatic destabilization is compensated by other favorable interactions such as cation−π interaction.64 Thus, in the pH range inspected in our study, luciferase may well undergo multiple protonations on His (pKa ∼6.0) and AMP (pKa ∼6.2), and the active pocket of the luciferase can be multiply protonated with the pH changes. In general, when only AMP(−2) is switched to AMP(−1) as in entry 2 → entry 1 in the table, the emission exhibits red shifts as consistent with a previous computational study.26 However, AMP is not the single “key” residue that affects the emission in a dominant manner. Rather, the change is only marginal in many situations and there are even two instances of slight blue shifts (entries 4 → 3 and 16 → 15). What is also intriguing with the AMP effect is its correlation with wat1 in the pocket: one can see from Table 2 that the red shift effect from the protonation on AMP(−2) is much reduced when wat1 is present in the pocket. It indicates that this water molecule can mediate the relationship between the residues in the active pocket and subsequently affect the emission energy. For F
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Figure 5. (a) Distributions of distance differences between OLU-AMP and OLU-LysH pairs, with and without wat1 in the active pocket. The distributions are compared for cases with AMP(−1) and AMP(−2). The distances are measured with atoms highlighted in red in the structure shown at the right. When the distance difference is small, OLU effectively moves toward AMP as pictorially depicted with arrows and residue names shown at the bottom. (b) The corresponding emission energy changes by AMP(−1)/AMP(−2) switches in relation to the presence and absence of wat1.
modulation. At first, this appears natural as water usually performs a screening effect toward charge−charge interactions. However, it is not clear whether merely a single water molecule may be capable of performing such effective screening. Because all three residues (Lys531, His247, and AMP) are rather directly interacting in close proximity with OLU, the significant reduction in the color modulation by a single water molecule is actually a surprising aspect. One plausible explanation for this peculiarity will be the possibility that the effect of protonation(s) may be somehow compensated by other effects in the active pocket when it is occupied by a water molecule. We have investigated this possibility by inspecting the relationship between emission energies and dynamical motions of these three residues in relation to the position of OLU. After close inspections, we indeed found flexible motions of the anionic AMP and the cationic Lys531 residues, which exhibit striking differences induced by the existence of wat1. Specifically, when the active pocket was occupied by ε-His (His247) and wat1, Lys531 moved in response to the switch between AMP(−2) and AMP(−1). In order to visually illustrate this behavior, we have displayed the histograms of distance relationships of these groups in Figure 5a. In this figure, the horizontal axis represents the difference between the OLU−AMP distance (rOLU‑AMP) and the OLU−LysH distance (rOLU‑LysH). Because these two distances, rOLU‑AMP and rOLU‑LysH, will strongly reflect how OLU will be influenced electrostatically by the two neighboring positively and negatively charged species, the difference between the two (rOLU‑AMP − rOLU‑LysH) can be adopted as a composite indicator for considering the positional relationship between AMP/LysH and OLU. When rOLU‑AMP − rOLU‑LysH becomes smaller, we can interpret that LysH moves farther from OLU and/or AMP migrates relatively closer to OLU.
Indeed, this is the case when wat1 is present in the active pocket. The left panel of Figure 5a shows that the distribution of the distance difference is shifted toward smaller values when AMP(−2) is protonated into AMP(−1), suggesting that AMP moves closer toward OLU with the protonation. In fact, Lys531 also moves away from OLU with this protonation switch (Figure S8). Equivalently, when the active pocket is occupied by AMP(−2), OLU moves relatively closer to Lys531 than AMP. This can be easily understood based on a simple electrostatic reasoning: the cationic Lys531 will compensate the increased electrostatic repulsion energy between the negative end of OLU and AMP(−2). However, when wat1 is not in the active pocket, the positional relationship between Lys531 and AMP does not vary much upon the alternation of the AMP protonation state as displayed with the right panel in Figure 5a. In addition, the distributions of rOLU‑AMP − rOLU‑LysH become narrower in this case. In contrast, the distributions with wat1 are dispersed with two noticeable peaks. From these aspects, we can see that the existence of a water molecule gives much more flexibility in positioning LysH and AMP toward their interaction. This can be understood from the dipolar nature of a water molecule. Because it can perform stabilization of neighboring ionic groups by adjusting its dipole moment direction, and because the dipole direction will have some amount of thermal distribution, the neighboring ionic groups may position in the active pocket with increased degree of flexibility. In a sense, we can conjecture that the existence of even a single water molecule can perform the lubricating role in the active pocket. Without such water in the pocket, the three groups of LysH, OLU, and AMP interact rather stiffly and a change in the protonation state does not lead to significant relocations of the groups. G
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our simulation time of 50 ns. Because the active pocket may have more pronounced effect on the protein and because the simple Monte Carlo approach that we have adopted might not be capable of catching the appropriate protonation state of histidine and AMP in the pocket, we also inspected the structural effect from their protonations by additionally considering all possible states of these groups. In all cases, the structure was not affected to any significant degree with an exception of δ-His and AMP(−2) in a portion of the simulated trajectories. However, δ-His and AMP(−2) cannot readily form an energetically stabilizing hydrogen bond network, and its significance will be very limited. As the structural variations upon pH changes were only marginal, we next moved on to test the effects of the electrostatic modulations from protonations. Of course, only the groups that are proximate to the light emitting ligand will have meaningful effects. Thus, we prepared a series of protonation states by adopting the acidic and basic forms of titratable residues in the active pocket. As previously reported,24,26,34,40,60 we found that the electrostatic interaction is an important factor for governing the emission energy. It should be noted that our approach naturally includes various types of intermolecular interactions31 besides electrostatic effects as the adopted MD and QM/MM calculations can capture such aspects in principle. In general, our results were consistent with many previous reports, where the importance of the environment near oxyluciferin was emphasized.21,23,24,26,34 Specifically, our simulations showed that the emission becomes more red as the number of protons in the active pocket increases as observed in previous computational26 and experimental6 studies. Most importantly, the red shift from the protonation was strongly affected by the water content in the active pocket. When a water molecule that can bridge Lys531−OLU or Lys531−AMP ion pairing was present in the pocket, these ionic groups could fluctuate with enhanced flexibility, leading to compensating motions of cationic and anionic species. Such compensations of course decreased the spectral shift caused by the added proton. On the contrary, when the active pocket was in a dry condition, the ionic groups around OLU were rigidly placed in the pocket, and the change in the electrostatic interaction from (de)protonation directly affected the emission energy without any compensating or screening effect. This aspect clearly indicates that the dynamics near the active pocket can act as a key factor in tuning the firefly bioluminescence color. It also suggests that both dynamical aspect and polarization of the active pocket should be considered in a careful manner especially for further modulating the bioluminescence toward its applications in practical fields.
These contrasting aspects in dynamical behaviors of the charged species are reflected on emission energies as shown in Figure 5b. When wat1 is in the active pocket, the emission energy is only slightly affected by the protonation and deprotonation on AMP (left panel). However, it is significantly affected without wat1 in the pocket (right panel), with ∼0.08 eV shift toward the red side with the protonation. This shift can be easily explained based on the electrostatic interactions and the flexibility inside the active pocket as discussed in the above. When wat1 is in the active pocket, the larger electrostatic effect from AMP(−2) than from AMP(−1) is compensated by the increased canceling effect from positively charged LysH at a closer distance. In other words, AMP(−2) is repelled more in distance by OLU while LysH is attracted more toward it in comparison with the case with AMP(−1). These motions of LysH and AMP effectively cancel their electrostatic effects on the excited state OLU. Namely, these flexible motions with wat1 will reduce the amount of red or blue shifts of the firefly bioluminescence by the pH change. The situation changes drastically when wat1 is missing inside the pocket. In this case, all three moieties (LysH, OLU, and AMP) form a more tightly tied network whose structure does not change very much upon the protonation state change on AMP. Thus, the associated electrostatic modulation with the AMP(−2)/AMP(−1) conversion directly influence the electronic state energies of OLU with little screening or compensation. Because the thiazole oxygen is more negative in the excited state, the decreased repulsive interaction with AMP(−1) at lower pH will stabilize the excited state more, leading to a red shift in the emission. Without wat1 in the active pocket, average values of emission energies are 2.01 eV with AMP(−2) and 1.93 eV with AMP(−1). On the contrary, the difference in average emission energies is only 0.02 eV when wat1 enhances the flexibility of Lys531, AMP, and OLU. The protonation on His247 induces a similarly contrasting behavior with and without wat1: in Table 2, entries 1 and 7 without wat1 exhibit 0.06 eV shift while entries 9 and 15 with wat1 show only 0.03 eV difference. Because experiments find a rather large shift (∼0.2 eV), this finding strongly suggests that the active pocket will likely be in a dry condition without any mediating water molecules. We believe this is also plausible as adenylated and oxidized luciferin before the final step of the biochemical reactions in the enzyme is much bigger than the luminescent oxyluciferin. Namely, the big ligand will not leave much space for water in the pocket. After the final decarboxylation step, luminescence will commence within the relatively short excited state lifetime of oxyluciferin. Because the water migration into and out of the pocket is quite hindered around the thiazole side,40 the dry condition will likely last during this excited state lifetime.
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4. SUMMARY AND CONCLUSION In this article, we have presented our simulation results on the pH dependent behavior of the firefly bioluminescence with a focus on the color modulation effects. We first investigated the possibility of pH dependent structural changes of the luciferase that may cause emission color modulations, by adopting five different pH conditions ranging from 5 to 9 and by testing plausible protonation states of various acidic and basic side chains of the protein at those pH conditions. Through this, we found that different forms of the protein exhibited similar structural fluctuations around a single structure regardless of the adopted protonation states at least during the duration of
ASSOCIATED CONTENT
S Supporting Information *
Complete citations of refs 5 and 58; atomic charges of AMP(−1); correcting emission energies obtained with the RICIS(D) method; structure of the luciferase−oxyluciferin system; luciferase domain motion induced by δ-His and AMP(−2); emission energy distributions with protonation states listed in Table 2; effects of protonation near the benzothiazole end of oxyluciferin; distributions of selected residue−residue distances. This material is available free of charge via the Internet at http://pubs.acs.org. H
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AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This work was supported by the Institute for Basic Science (IBS) in Korea. The supercomputer time from Korea Institute of Science and Technology Information (KISTI) is also gratefully acknowledged.
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