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Probing of the plasticity of the active site in pinene synthase elucidates its potential evolutionary mechanism
Phytochemistry 181 (2021) 112573
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Probing of the plasticity of the active site in pinene synthase elucidates its potential evolutionary mechanism Jingwei Xu 1, Guanzu Peng 1, Jinkun Xu, Yi Li, Li Tong, Dong Yang * Gene Engineering and Biotechnology Beijing Key Laboratory, College of Life Sciences, Beijing Normal University, Beijing, 100875, China
A R T I C L E I N F O
A B S T R A C T
Keywords: Paeonia lactiflora Paeoniaceae GC-MS Molecular dynamics Monoterpene Terpenene synthase Pinene Sabinene
Terpenes form a class of highly diverse natural products. The diversity of terpenes is created by terpene syn thases. During the reaction, carbocation intermediates form and their rearrangement could lead to the formation of various products. Terpene synthases determine the final product profile by controlling the conformation of the intermediate or stabilizing the carbocation. Pinene synthase is a monoterpene synthase catalyzing the cyclization of geranyl pyrophosphate (GPP) to form pinene. Our study suggests that by mutating the aromatic residue F482 to Ala, Val, Ile and Thr, the enzyme can be converted to sabinene synthase, with more than 90% of its total terpene products becoming sabinene, which indicates the aromaticity of this residue is essential for stabilizing the pinyl carbocation. We also identified a mutation S491A that could cause an about 29% increase in the overall activity of the enzyme without altering its produce selectivity. Molecular dynamic simulation indicates this mutation could decrease the flexibility of the enzyme when it forms a complex with the pinyl carbocation. Our study suggests the active pocket of pinene synthase has a certain level of plasticity, making it relatively easy to change the product selectivity or overall activity. This property could have an important implication in the evolution of terpene synthases and thereby terpene diversity, as by changing a few residues an enzyme could synthesize a completely different terpene product in a significant amount, which allows selection to take place.
1. Introduction Terpenes form one of the largest groups of natural products (Wise and Croteau, 1999; Tholl, 2006; Christianson, 2006, 2007; Degenhardt et al., 2009; Gao et al., 2012). As of 2012, more than 55,000 different types of terpenes have been reported (Gao et al., 2012). Terpenes play many biological roles. For example, they can serve as signaling mole cules mediating interactions between plants and insects (Pare and Tumlinson, 1999; Umehara et al., 2008). Cholesterol, also belonging to the terpene family, plays important roles in regulating membrane fluidity. Terpenes also have applications in food, cosmetics, medicine and other industries (Ajikumar et al., 2008; Kirby and Keasling, 2009). All terpenes are derived from a five-carbon precursor, isopentenyl pyrophosphate (IPP) (Wise and Croteau, 1999; Tholl, 2006; Christian son, 2006, 2007; Degenhardt et al., 2009; Gao et al., 2012). IPP and its isomer dimethylallyl pyrophosphate (DMAPP) are synthesized via either the mevalonate or the non-mevalonate pathway. The head-to-tail condensation of DMAPP and IPP forms geranyl pyrophosphate (GPP). Further condensation with more IPP units forms farnesyl pyrophosphate
(FPP), geranylgeranyl pyrophosphate (GGPP), etc. Cyclization of these linear products by terpene synthases is the key step to form the terpene diversity (Wise and Croteau, 1999; Tholl, 2006; Christianson, 2006, 2007; Degenhardt et al., 2009; Gao et al., 2012). Cyclized products can be modified by oxidation and other steps, further increasing the di versity of terpenoids. Cyclization is usually initiated by creating a carbocation in the active site of terpene synthases. The rearrangement of carbocations leads to the formation of a variety of skeletons of terpenes. This process is controlled by terpene synthases as they have evolved multiple mechanisms to limit the conformation of the substrate or stabilize a certain carbocation in termediate and prevent its rearrangement (Lesburg et al., 1997; Starks et al., 1997; Christianson, 2006, 2007; Gao et al., 2012). Aromatic or polar side chains are often involved in the latter process (Christianson, 2007; Zhou and Peters, 2011; Gao et al., 2012). Because of differences in these controlling mechanisms, the product selectivity of terpene syn thases could have large variations. Some enzymes are specific, pre dominantly producing one product, while others could be promiscuous and form a wide range of different products (O’Maille et al., 2008;
* Corresponding author. E-mail address: [email protected] (D. Yang). 1 These authors contributed equally to this work. https://doi.org/10.1016/j.phytochem.2020.112573 Received 27 June 2019; Received in revised form 19 October 2020; Accepted 22 October 2020 Available online 1 November 2020 0031-9422/© 2020 Elsevier Ltd. All rights reserved.
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Yoshikuni et al., 2006). The active pockets of terpene synthases often demonstrate a certain level of plasticity, as changing a few residue could completely alter the product profile (Morrone et al., 2008; Wilderman et al., 2007; Xu et al., 2007; Greenhagen et al., 2006; Yoshikuni et al., 2006; Keeling et al., 2008; Zerbe et al., 2012; Katoh et al., 2004). It has been proposed that product specificity of terpene synthases may evolve from their more promiscuous ancestors (Bohlmann et al., 1998; O’Brien and Herschlag, 1999; Copley, 2003; O’Maille et al., 2008). Monoterpenes are terpenes derived from GPP (Fig. 1A). Most of them are volatile scent compounds emitted by plants. The key intermediate for cyclic monoterpenes is the terpinyl cation (Pyun et al., 1993; Wil liams et al., 1998; Schwab et al., 2001; Hyatt et al., 2007; Srividya et al., 2015). If directly deprotonated, it forms limonene (Fig. 1A). The terpinyl cation can also undergo different types of rearrangement, which leads to the formation of other products (Fig. 1A). For example, 2, 7-ring closure forms the pinyl cation that is the precursor of α- and β-pinene. 6, 7-hy dride shift followed by 2, 6-ring closure forms the thujyl cation and then sabinene (Srividya et al., 2015). We previously studied the catalytic mechanism of Mentha spicata Slimonene synthase (Xu et al., 2017). Our data suggests that by intro ducing N345A/L423A/S454A mutations we can convert its activity to that of pinene synthase. Our further studies suggest that polar residues
in the active site of S-limonene synthase may form a “polar pocket” that participates in the stabilization of the terpinyl cation. Disruption of this pocket could lead to the rearrangement of the terpinyl cation and the formation of alternative products. In this study, we probed the key residues of Paeonia lactiflora pinene synthase. Our results suggest that by mutating the aromatic residue F482 to Ala, Val, Ile or Thr, the enzyme could be completely converted to sabinene synthase. We also identified a mutation S491A that can significantly enhance the enzyme activity without changing its product profile. These results show that pinene synthase, like S-limonene syn thase we studied before, has significant plasticity in its active site, which may explain the evolutionary history of monoterpene synthases. 2. Results 2.1. Homology modeling of P. lactiflora pinene synthase The crystal structures of pinene synthases have not been reported. Therefore, we constructed a homolog model of P. lactiflora pinene syn thase using the crystal structure of S-limonene synthase as the template. In order to investigate the catalytic mechanism, we docked three car bocation intermediates (terpinyl, pinyl and thujyl cations) into its active
Fig. 1. a, The proposed reaction mechanism for the synthesis of limonene, pinene and sabinene. b, The structural model of pinene synthase docked with the terpinyl (left), pinyl (middle) and thujyl (right) cations. Carbocations are colored in magenta (terpinyl), wheat (pinyl) and cyan (thujyl). Other atoms are colored according to their types (red: oxygen; yellow: carbon; blue: nitrogen; orange: phosphorus; green: magnesium). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.) 2
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site (Fig. 1B–D). The docking results suggest that carbocations bind to a similar location in the active pocket. Nearby residues include R305, S311, W314, I335, I338, N339, D342, Y417, I443, A444, I448, F482, D486, S491 and H571. Some of these residues could make hydrophobic interactions with the carbon skeleton of carbocations. Certain polar or aromatic residues may stabilize the positive charge and influence the product profile of the enzyme.
investigate this issue, we first docked the pinyl cation into the active pockets of WT and F482Y. It could be seen that in these two structures, the conformation of the docked pinyl cation is quite different (Fig. 4A). We then performed a 2 ns molecular dynamics simulation. The results suggest more conformational changes (Fig. 4B–C). Therefore, the extra hydroxyl group in Tyr could significantly change the conformation of the carbocation, which may explain the decreased activity and altered product profile.
2.2. Alanine scanning mutagenesis identifies mutations that cause an increase in sabinene production
2.4. S491A mutation enhances the overall activity of pinene synthase without changing its product profile
We first identified residues within 5 Å from the docked carbocations and performed alanine-scanning mutagenesis on them. Most of these mutations do not show dramatic changes in their product profiles. However, two mutations show interesting alteration in the product profile. One of them, F482A, essentially converts the enzyme to sabinene synthase. About 90.8% of its products is sabinene, while 97.7% of the products of the WT enzyme is α-pinene (Table 1, Fig. 2A–B). Another mutation, I335A, also causes an increase in sabinene production, though not as dramatic as F482A (Table 1). Importantly, although both mutants show an altered product profile, their overall activities are only moderately decreased (Table 1).
During our Ala scanning experiments, we identified a mutation S491A that enhances overall activities (Fig. 5A, D, Table 1). The overall activity of S491A mutant is about 29% higher than that of the WT enzyme (Table 1). Furthermore, S491A does not change the product selectivity, as more than 90% of their products remains α-pinene (Fig. 5A, Table 1). What is the mechanism of S491A to enhance the enzymatic activity? Since S491A still predominantly produces pinene, we docked the pinyl cation into the active pocket of the mutant enzyme and performed a 2 ns of molecular dynamics simulation. We also performed the same dynamic simulation on the WT enzyme in complex with the pinyl cation. Our data suggests that the active site residues in S491A have a much smaller RMSD than those in the WT enzyme, indicating the mutation S491A could cause the active pocket to be more rigid (Fig. 6A, STable 2). Interestingly, simulation on the WT or S491A enzymes in complex with the terpinyl or thujyl cation does not show the same differences in RMSD values of active site residues (Fig. 6B–C, STable 2). This probably in dicates that the S491A mutation increases the rigidity of the active pocket only when it binds to the pinyl cation. To better quantify the mobility of protein structures, we reanalyzed the simulation data using MDLovoFit (Martínez, 2015). This way we could subdivide the protein into regions with high and low mobility (Fig. 7A–C, STable 3). Interestingly, in S491A, fewer residues are clas sified as of high mobility than residues in WT enzymes (59% vs 63%). Furthermore, for residues classified with the highly mobile subset, those in S491A have lower average RMSD than those in WT proteins (0.253 nm vs. 0.278 nm). Therefore, there is a slight but significant increase in rigidity in S491A mutant protein.
2.3. Phenylalanine at the 482 position is required for pinene production We hypothesized that the aromatic ring of F482 might play a role in stabilizing the pinyl cation. To investigate this possibility, we mutated F482 to Val and Ile, two nonpolar residues with different side chain sizes. We also mutated it to Thr, Arg and Trp, representing polar, charged or non-Phe aromatic residues. The results suggest that any mutations that changes the phenylalanine at this position leads to an increase in sabinene production and almost abolishes the α-pinene production (Table 1). Replacing phenylalanine with nonpolar residues (F482I, F482V) all effectively convert the enzyme to sabinene synthase (Fig. 2C–D). More than 90% of products made by these mutants are sabinene (Table 1). Interestingly, F482T also predominantly produces sabinene, which seems to suggest that a polar residue here is not suffi cient to stabilize the pinyl cation (Table 1, Fig. 2E). Other mutations (F482Y, F482R and F482W) retain very little activities (Table 1). In order to verify that the mutant enzymes indeed produce sabinene, we analyzed the reaction product of F482L using the high resolution hybrid quadrupole-orbitrap GC-MS/MS system (Fig. 3A–C). By comparing with the sabinene standard, the peak eluted at 9.29 min was identified as sabinene. Its mass spectrum was measured and could be readily matched with that of sabinene standard (Fig. 3B–C). It is interesting to note that replacement of F482 to other aromatic residues (Tyr or Trp) causes a drastic decrease in enzyme activities. To
2.5. Combination of S491A with F482 mutants has mixed results in the overall activity Since all four sabinene synthase mutants show some decreases in the catalytic activity, we wondered whether we may restore their activities to that of the WT pinene synthase by combing with S491A. We produced
Table 1 Major products (%±SD) of WT α-pinene synthase and enzymes with mutations. Enzymes
PS-WT F482A S491A F482I F482L F482V F482T F482Y F482R F482W I335A F482A/S491A F482I/S491A
Relative Proportion of Products (%) Unknown
α-pinene
sabinene
limonene
Unknown
Unknown
RT = 4.52
RT = 7.05
RT = 8.55
RT = 11.05
RT = 11.58
RT = 12.72
0.31 ± 0.02 0.05 ± 0.00 0.46 ± 0.01 1.27 ± 0.02 0.96 ± 0.06 1.03 ± 0.01 0.00 ± 0.00 0.00 17.62 ± 6.21 16.32 ± 5.33 3.52 ± 0.02 1.34 ± 0.11 0.63 ± 0.02
97.86 ± 0.21 4.43 ± 0.03 97.28 ± 0.09 3.17 ± 0.08 2.03 ± 0.03 2.12 ± 0.03 3.53 ± 0.00 2.71 0.00 ± 0.00 0.00 ± 0.00 78.00 ± 0.03 4.17 ± 0.01 2.72 ± 0.00
0.96 ± 0.00 93.21 ± 0.06 1.14 ± 0.00 92.35 ± 0.21 94.26 ± 0.30 94.51 ± 0.07 93.91 ± 0.00 31.00 18.96 ± 7.19 29.92 ± 17.90 12.94 ± 0.00 88.40 ± 1.43 92.88 ± 0.45
0.26 ± 0.01 1.59 ± 0.05 0.12 ± 0.00 0.66 ± 0.09 0.31 ± 0.01 0.30 ± 0.03 1.64 ± 0.04 63.61 31.04 ± 19.27 21.67 ± 6.84 0.57 ± 0.02 3.38 ± 0.14 2.40 ± 0.09
0.54 ± 0.05 0.72 ± 0.03 0.99 ± 0.06 2.33 ± 0.08 1.91 ± 0.17 1.82 ± 0.06 0.92 ± 0.05 2.68 27.33 ± 14.94 27.35 ± 14.96 4.28 ± 0.03 2.36 ± 0.34 1.19 ± 0.08
0.07 ± 0.00 ± 0.00 ± 0.23 ± 0.29 ± 0.23 ± 0.00 ± 0.00 5.05 ± 4.74 ± 0.67 ± 0.35 ± 0.18 ±
3
0.00 0.00 0.00 0.00 0.01 0.00 0.00 0.51 0.45 0.00 0.01 0.00
Activity(% WT) 100 46.63 ± 4.47 128.90 ± 4.19 18.36 ± 2.51 49.98 ± 39.78 29.50 ± 12.39 52.69 ± 3.17 0.62 1.28 ± 0.01 1.54 ± 0.02 54.36 ± 2.17 18.73 ± 0.13 28.74 ± 5.73
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Fig. 2. Converting pinene synthase to sabinene synthase by mutations on 482 position. Chromatograms in a, b, c, d and e show the GC-MS analysis of terpene products for WT, F482A, F482I, F482V and F482T, respectively. The x-axis is the retention time and the y-axis is the relative abundance of each species. The numbers in each peak correspond to α-pinene (1), sabinene (2) and limonene (3).
double mutants F482A/S491A and F482I/S491A. Both seem to retain the same product selectivity of the original single mutants, though F482A/S491A shows an insignificant decrease in sabinene production (P = 0.39; Fig. 5B–C, Table 1). In overall activities, F482I/S491A shows an insignificant increase in activity (P = 0.27), while F482A/S491A has a significant decrease in activity (P < 0.05; Fig. 5D, Table 1). Therefore, it seems that the effect of combining two mutants is more complicated than we expected. We asked whether the combination of F482A and S491A mutations could affect the rigidity of the protein. After a 2 ns of molecular dy namics simulation and data analysis using MDLovoFit, we found the rigidity of F482A/S491A is similar to that of S491A (STable 3). Appar ently, the decrease of activity by the combination of F482A with S491A cannot be explained by the overall RMSD change. To further investigate this issue, we analyze the RMSF values based on the simulation data (SFig. 1A–C). Compared with S491A and WT, F482A/S491A shows an RMSF spike at a region between residues 219 and 223. In the structural model, this region forms a loop in the protein structural model (SFig. 2). However, this loop is quite far from the active site (about 40 Å). Therefore, its mobility could only affect the enzyme activity indirectly. For example, it could influence the closure of the active pocket. We also superimposed the structural model of WT, S491A and F482A/S491A in complex with the docked pinyl cation (SFig. 3). In the
structural model, the pinyl cation is docked between the residue 491 and 482 (SFig. 3). Interestingly, in WT and S491A, the position of the pinyl cation is similar, while in F482A/S491A, the pinyl cation is shifted to wards the A482. This change in the carbocation position may impair the enzyme activity (SFig. 3). 3. Discussion The plasticity of the active pocket has been investigated in several terpene synthases. Our previous study in S-limonene synthase has shown that mutating three residues could readily convert limonene synthase to pinene synthases (Xu et al., 2017). In this case, about 70% of the total products become pinene or phellandrene in mutant enzymes. Here, we performed a similar study in pinene synthase. By mutating a single residue, we achieved an almost complete conversion of pinene synthase to sabinene synthase, resulting more than 90% of the total products to be sabinene. These findings prove that the active pocket of monoterpene synthases have a certain level of plasticity and dramatic changes in product profile can be achieved by a small number of mutations. In limonene synthase, we showed that polar residues are important to stabilize the terpinyl cation (Xu et al., 2017). We identified a polar pocket and found that mutating key residues in this pocket could lead to the rearrangement of the terpinyl cation and cause the production of alternative products (Xu et al., 2017). In pinene synthase, however, we 4
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Fig. 3. Hybrid quadrupole-orbitrap GC-MS/MS. a shows the chromatogram of terpene products of F482L. The elution peak corresponding to sabinene is labeled. The mass spectrum of the product eluted at 9.29 min from a is shown in b. The mass spectrum of the standard sabinene is shown in c.
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synthase do not change the product profile, as α-pinene still composes of more than 90% of the total products. Dynamic simulation suggests that the mutation S491A could decrease the flexibility of the active pocket only when it binds the pinyl cation. Therefore, it is possible that this mutation makes an enzyme that is already specific for pinene production more rigid. Probably this rigidness stabilizes the pinyl cation intermediate in the active pocket, which enhances the overall activity without changing the product se lection. Interestingly, during the maturation of antibodies, a decrease in protein flexibility is often associated with the increased affinity, prob ably due to the decrease in the entropic cost for antigen binding (Thorpe and Brooks, 2007). Here, in a completely different setting, we found this mechanism may also play a similar role. However, we have to be prudent to apply this mechanism. The enhancement of rigidity of the overall protein may indeed play a role in activity enhancement in S491A. But in F482A/S491A, the RMSD of the overall structure seems to change little. The decrease of activity in F482A/S491A could probably be caused by the positional change of the pinyl cation in the active pocket or the increased flexibility in the loop between 219 and 223. Other causes could also be possible and further investigations are required. One question remains. Why only when S491A binds the pinyl cation does the protein rigidity increase? As our simulation on the F482Y mutant protein indicates, the conformation of the carbocation is highly sensitive to the slight change in the amino acid in the active pocket. Probably, the opposite is also true. Therefore, the difference in the carbocation may also play an important role in the conformation of the enzyme. The basis of terpene diversity is terpene synthases. Since a slight alteration by a few mutations in the active pocket could significantly change the product profile, the expansion of the chemical space of ter penoids could be easily achieved. It is likely that the ancestor of modern terpene synthases are quite promiscuous and produce a multitude of products. New enzymes that produce novel terpene products could evolve by a few mutations in the active pocket. If these new products are beneficial to the organism (e.g. attracting pollinating insects), enzymes
Fig. 4. Conformational differences in the pinyl cation in WT and F482Y as demonstrated by molecular dynamics simulation. The structures of WT and F482Y are superimposed. Results after 0, 1 and 2 ns of simulation are shown in a, d and c, respectively. Carbon in WT is colored in yellow and carbon in F482Y is colored in green. Oxygen is colored in red. Nitrogen is colored in blue. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)
found the aromatic residue F482 is a key player. Mutating this residue to other polar residues could not maintain the pinene synthase activity, as the result of F482T proves. Actually, it seems the residue in this position must be a Phe for pinene synthase, as a similar aromatic residue, Tyr, does not work (Table 1). This phenomenon could probably be explained by the possible geometric constraint in this area so even the addition of a single hydroxyl group in Tyr can significantly change the conformation of the pinyl cation, as our simulation suggests (Fig. 4). This conforma tional change could explain the decrease of the overall enzyme activity as well as the change in the product profile. Therefore, it seems that within monoterpene synthases, distinct mechanisms are used to stabilize the carbocation intermediate and determine the product selectivity. The plasticity of the active pocket is also reflected on the relative ease of changing the overall activity of the enzyme. In both the previous study on limonene synthase and this study on pinene synthase, we showed the alteration of a single amino acid is sufficient to enhance the overall enzymatic activity, which means the mutant enzyme can pro duce more terpenoid products, regardless of its types. For example, the mutation S454G is about 7% more active than the WT limonene syn thase. In pinene synthase, S491A is about 29% more active than the WT enzyme. Different from S454G in limonene synthase, S491A in pinene
Fig. 5. The effect of the S491A mutation. Chromatograms in a, b and c show the GC-MS analysis of terpene production for S491A, F482A/S491A, and F482I/S491A, respectively. The x-axis is the retention time and the y-axis is the relative abundance of each species. The numbers in each peak correspond to α-pinene (1), sabinene (2) and limonene (3). d shows the overall activity and the percentage of sabinene within total products produced by F482A, F482A/S491A, F482I and F482I/S491A. The asterisk indicates P < 0.05. 6
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Fig. 6. S491A mutation increases the rigidity of the active pocket when it binds the pinyl cation. RMSD values of the active site residues between WT and S491A in complex with the pinyl cation (a), terpinyl cation (b) and thujyl cation (c) are compared.
producing them is likely to be selected and further evolution may nar row the range of products, producing a new enzyme that specifically produces this compound. One important difficulty that a new terpene synthase faces is the fact that normally a mutation decreases the overall activity of the enzyme. Therefore, although the new enzyme may produce a novel compound, because its activity is lower, it may not be able to produce a sufficient amount of new compounds that could benefit the organism. However, our study has demonstrated that the overall activity of the enzyme could be enhanced by a single mutation. The cause of the activity increase could be through the increase of the rigidity of the overall protein or the
alteration of fluctuation of some structural elements. Other mechanisms may also be involved. Obviously, more research should still be per formed to delineate the possible mechanism involved in this process. Another important issue is about the chirality of products in our enzyme. Unfortunately, due to the difficulty of obtaining chiral stan dards, we could not perform a chiral GC-MS. Furthermore, because of the low yield of the enzymatic reaction, we were not able to perform NMR to determine the absolute configuration of terpene products. However, by analyzing protein sequences of pinene and sabinene syn thases with known chirality of their products, we could still make certain conclusions. 7
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Fig. 7. The RMSD profiles of the region with high (RMSD H) and low (RMSD L) mobility from the simulation of WT (a), S491A (b) and F482/S491A (c) in complex with the pinyl cation.
Several studies on pinene synthases reported the chirality of its products (Hall et al., 2013; McKay et al., 2003; Phillips et al., 2003; Williams et al., 1998). By making a multiple sequence alignment, we could identify possible key residues that may determine the chirality of the product (SFig. 4). The most interesting residue is the one corre sponding to I443 in our enzyme. In reported (+)-α-pinene synthases, this residue is always a Phe, while in reported (− )-α-pinene synthases, it is always a Ser. By looking at the homology model of our enzyme, we found I443 is close to the docked pinyl cation (Fig. 1C). So the size of this residue is likely to influence the chirality of the product. For example, a bulky side chain such as Phe may cause steric clash and prevent the formation of (− )-α-pinene. Therefore, as in our enzyme this residue is a bulky Ile, the wild type form of our enzyme is likely to produce (+)-α-pinene. As a further proof, phylogenic analysis also shows our enzyme is more closely related to (+)-α-pinene synthases than (− )-α-pinene synthases (SFig. 5). Unfortunately, we could not find any reports on (− )-sabinene syn thases, which makes it difficult to make sequence analysis directly. However, it is known that both (+)-pinene and (+)-sabinene require the carbocation intermediate (3R)-linalyl diphosphate (Wise et al., 1998). Therefore, if the wild type form of our enzyme indeed produces (+)-pinene, it is most likely the sabinene produced in our mutant enzyme is (+)-sabinene, since they share the same carbocation intermediate. Certainly, our interpretation on the rigidity of the enzyme as well as the chirality of products is still relatively speculative. In the future, more experiments should be performed to validate our predictions on both cases.
sequence, we only synthesized the sequence encoding the protein sequence from R51 to the C-terminus. The codon usage was optimized for the expression in E. coli. It was cloned into pGEX-4T-1 vector. Point mutations were introduced by the site-directed mutagenesis kit (Trans gen and Takara). Primers used were listed in STable 1. The amplification products were digested by DpnI for 5 h at 37 ◦ C and transformed into E. coli DH5α cells. Correct mutations were identified by sequencing. 5.2. Protein expression and purification Plasmids were transformed into E. coli BL21 (DE3) cells and were then cultured in LB medium with 50 mM ampicillin. Expression was induced by 0.5 mM IPTG at 20 for 19 h. Cells were harvested by centrifugation at 3000 g for 10 min, resuspended by buffer A (25 mM Tris, pH 8.0, 150 mM NaCl, 1 mM DTT, 0.2 mM PMSF) and lysed with sonication. The lysate was applied to glutathione sepharose resin and proteins were then cleaved on-column by thrombin that was diluted in 25 mM Tris, pH 8.0, 150 mM NaCl and 1.25 mM CaCl2 to 1 KU. Finally, proteins were eluted with buffer E (25 mM Tris, pH 8.0, 150 mM NaCl, 1 mM DTT, and 0.5 mM EDTA). 5.3. Enzymatic reaction and GC-MS assay The reaction was performed in a 400 μl reaction system that contains 25 mM Mops, pH 7.0, 15 mM MgCl2, 1.0 mM DTT, 10% (v/v) glycerol, 68.5 μM GPP and 1 μg pinene synthase. The reaction was performed at 37 ◦ C for 40 min, and then 400 μl of hexane was added to extract terpene. The mixture was incubated at 37 ◦ C for 20 min and vortexed for 30 min. Then it was centrifuged at 8000 g for 5 min to separate the organic layer that contains terpene products. 1 μl of extraction was analyzed by GC-MS (Thermo Trace GC Ultra & ISQ). The extraction was injected into a HP-88 column (100 mm × 250 μM x 0.20 μM thickness; Agilent), with ultra-purity helium as the carrier gas at the flow rate 1.0 ml/min. The oven temperature was set to 3 phases; the temperature was first increased to 50 ◦ C and kept for 1 min, then increased to 130 ◦ C at the increment of 3 ◦ C/min, and increased to 220 ◦ C at the increment of 50 ◦ C/min and held for 5 min. The injector and transfer line temperature were set at 250 ◦ C. Terpenes were identified by GC retention times by comparing with available authentic standards. The proportion of each product was based on the ratio of the peak area of the particular product and the peak area of total terpene products. The relative activity of an enzyme is calculated by dividing the total peak area for all terpene products of one mutant enzyme by that of the wild type (WT) pinene synthase.
4. Conclusions In conclusion, we have demonstrated the plasticity of the active pocket of pinene synthase in our study. Our data suggested mutating a single residue could dramatically change the product selectivity of the enzyme, converting it to sabinene synthase. We also demonstrated a single mutation could significantly enhance the overall activity of the enzyme without changing its product profile, probably caused by the increase in rigidity of the protein. Our findings could provide important insights on the evolution of new monoterpene synthases and the di versity of monoterpenes. 5. Methods 5.1. Gene synthesis and site-directed mutagenesis The P. lactiflora α-pinene synthase (PS) cDNA was synthesized by Sangon Biotech (China). According to the findings of Williams and his colleagues (Williams et al., 1998), we identified the potential plastidial targeting sequence by aligning its protein sequence with that of M. spicata S-limonene synthase. To eliminate this plastidial targeting
5.4. Hybrid quadrupole-orbitrap GC-MS/MS system analysis The reaction product was analyzed by the Hybrid QuadrupoleOrbitrap GC-MS/MS System (ThermoFisher Q Exactive GC). 1 μl of extraction was injected with ultra purity helium as the carrier gas at the 8
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flow rate 1.0 ml/min, with split in 20 ml/min. The oven temperature was set to 3 phases, the temperature was first increased to 50 ◦ C and kept for 1 min, then increased to 100 ◦ C at the increment of 3 ◦ C/min, and increased to 220 ◦ C at the increment of 50 ◦ C/min and held for 5 min. The injector and transfer line temperature were set at 250 ◦ C. High resolution EI fragment spectra were acquired using 60,000 FWHM res olution, with full scan mode mass range of 50–550 Da. To identify the reaction products, their gas chromatography retention time and mass spectra were compared with those of the sabinene standard.
Declaration of competing interest The authors declare no competing financial interest. Acknowledgements This work is supported by grants from the Fok Ying Tung Education Foundation (#132025), the National Science Foundation of China (#31070682 and #31640004), State Education Ministry and the Pro gram of the Co-Construction with Beijing Municipal Commission of Education of China. We thank the Center of Biomedical Analysis, Tsinghua University, for the technical assistance in GC-MS.
5.5. Construction of structural models for protein-ligand complexes For proteins, the homology model of the α-pinene synthase from P. lactiflora (GenBank: ANA91931.1) was constructed by Swiss-model (Waterhouse et al., 2018) using the structure of 4S-limonene synthase from M. spicata (2ONH) as the template. Based on the structural model of the WT α-pinene synthase, a series of mutants structural models were constructed by PyMoL version 2.1.1. For ligands, the structure of carbocation intermediates (the terpinyl, pinyl and thujyl cations) were generated by MOE v2008.1 (Chemical Computing Group Inc.). They were then docked into the active site of pinene synthase models in Autodock 4.2.5.1 using default settings (Morris et al., 2009). The structures of protein-carbocation complexes were then optimized in ATB (Malde et al., 2011). The coordinate of Mg2+ was adapted from the coordinate of Mn2+ in the limonene syn thase structure by superimposing the limonene synthase structure with the pinene synthase structural model. In the same way, the pyrophos phate group was adapted from the pyrophosphate tail in the limonene synthase structure. The topology and force field files for carbocations and the pyrophosphate tail were generated and optimized by ATB.
Appendix A. Supplementary data Supplementary data to this article can be found online at https://doi. org/10.1016/j.phytochem.2020.112573. References Ajikumar, P.K., Tyo, K., Carlsen, S., Mucha, O., Phon, T.H., Stephanopoulos, G., 2008. Terpenoids: opportunities for biosynthesis of natural product drugs using engineered microorganisms. Mol. Pharm. 5, 167–190. Berendsen, H.J.C., Spoel, D.V.D., Drunen, R.V., 1995. GROMACS: a message-passing parallel molecular dynamics implementation. Comput. Phys. Commun. 91, 43–56. Bohlmann, J., Meyer-Gauen, G., Croteau, R., 1998. Plant terpenoid synthases: molecular biology and phylogenetic analysis. Proc. Natl. Acad. Sci. U.S.A. 95, 4126–4133. Christianson, D.W., 2006. Structure biology and chemistry of the terpenoid cyclases. Chem. Rev. 106, 3412–3442. Christianson, D.W., 2007. Roots of biosynthetic diversity. Science 316, 60–61. Copley, S.D., 2003. Enzymes with extra talents: moonlighting functions and catalytic promiscuity. Curr. Opin. Chem. Biol. 7, 265–272. Degenhardt, J., K¨ ollner, T.G., Gershenzon, J., 2009. Monoterpene and sesquiterpene synthases and the origin of terpene skeletal diversity in plants. Phytochemistry 70, 1621–1637. Gao, Y., Honzatko, R.B., Peters, R.J., 2012. Terpenoid synthase structures: a so far incomplete view of complex catalysis. Nat. Prod. Rep. 29, 1153–1175. Greenhagen, B.T., O’Maille, P.E., Noel, J.P., Chappell, J., 2006. Identifying and manipulating structural determinates linking catalytic specificities in terpene synthases. Proc. Natl. Acad. Sci. U.S.A. 103, 9826–9831. Hall, D.E., Yuen, M.M., Jancsik, S., Quesada, A.L., Dullat, H.K., Li, M., Henderson, H., Arango-Velez, A., Liao, N.Y., Docking, R.T., Chan, S.K., Cooke, J.E., Breuil, C., Jones, S.J., Keeling, C.I., Bohlmann, J., 2013. Transcriptome resources and functional characterization of monoterpene synthases for two host species of the mountain pine beetle, lodgepole pine (Pinus contorta) and jack pine (Pinus banksiana). BMC Plant Biol. 13, 80. Hyatt, D.C., Youn, B., Zhao, Y., Santhamma, B., Coates, R.M., Croteau, R.B., Kang, C., 2007. Structure of limonene synthase, a simple model for terpenoid cyclase catalysis. Proc. Natl. Acad. Sci. U.S.A. 104, 5360–5365. Katoh, S., Hyatt, D., Croteau, R., 2004. Altering product outcome in Abies grandis (-)-limonene synthase and (-)-limonene/(-)-alpha-pinene synthase by domain swapping and directed mutagenesis. Arch. Biochem. Biophys. 425, 65–76. Keeling, C.I., Weisshaar, S., Lin, R.P.C., Bohlmann, J., 2008. Functional plasticity of paralogous diterpene synthases involved in conifer defense. Proc. Natl. Acad. Sci. U. S.A. 105, 1085–1090. Kirby, J., Keasling, J.D., 2009. Biosynthesis of plant isoprenoids: perspectives for microbial engineering. Annu. Rev. Plant Biol. 60, 335–355. Lesburg, C.A., Zhai, G., Cane, D.E., Christianson, D.W., 1997. Crystal structure of pentalenene synthase: mechanistic insights on terpenoid cyclization reactions in biology. Science 277, 1820–1824. Malde, A.K., Zuo, L., Breeze, M., et al., 2011. An automated force field topology builder (ATB) and repository: version 1.0. J. Chem. Theor. Comput. 7, 4026–4037. Martínez, L., 2015. Automatic identification of mobile and rigid substructures in molecular dynamics simulations and fractional structural fluctuation analysis. PloS One 10, e0119264. McKay, S.A., Hunter, W.L., Godard, K.A., Wang, S.X., Martin, D.M., Bohlmann, J., Plant, A.L., 2003. Insect attack and wounding induce traumatic resin duct development and gene expression of (-)-pinene synthase in Sitka spruce. Plant Physiol. 133, 368–378. Morris, G.M., Huey, R., Lindstrom, W., Sanner, M.F., Belew, R.K., Goodsell, D.S., Olson, A.J., 2009. Autodock4 and AutoDockTools4: automated docking with selective receptor flexibility. J. Comput. Chem. 16, 2785–2791. Morrone, D., Xu, M., Fulton, D.B., Determan, M.K., Peters, R.J., 2008. Increasing complexity of a diterpene synthase reaction with a single residue switch. J. Am. Chem. Soc. 130, 5400–5401. O’Brien, P.J., Herschlag, D., 1999. Catalytic promiscuity and the evolution of new enzymatic activities. Chem. Biol. 6, R91–R105. O’Maille, P.E., Malone, A., Dellas, N., Hess Jr., B.A., Smentek, L., Sheehan, I., Greenhagen, B.T., Chappell, J., Manning, G., Noel, J.P., 2008. Quantitative
5.6. Molecular simulation The protein-ligand complex was then solvated in a 0.707 × 6 × 6 × 6 nm3 dodecahedron water box (spc216 water model) with Na+ and CL− ions in GROMACS version 5.1.4 (Berendsen et al., 1995; Oostenbrink et al., 2010). For energy minimization, 200ps NVT ensemble (constant Number of particles, Volume, and Temperature) and 200ps NPT ensemble (constant Number of particles, Pressure, and Temperature) were performed in GROMACS version 5.1.4 using GROMOS96 54a7 force field (Berendsen et al., 1995; Oostenbrink et al., 2010). The equilibrated systems were then used for molecular dynamics simulation in GROMACS using GROMOS96 54a7 force field for 2ns in NPT ensemble (300K, 1 atm). To investigate the RMSD of the active pocket residues, we first defined the active pocket as the residues within 5 Å from the docked carbocations. 5.7. Analysis of the simulation data RMSD values of these residues were calculate based on MD trajec tories produced by the molecular dynamic simulation mentioned above. The simulation data was further analyzed with MDLovoFit-16.264 (Martínez, 2015). Each protein was divided into two subsets according to the structural fluctuation. Then RMSD L (low fluctuation fraction), RMSD H (high fluctuation fraction) and RMSF were obtained. Author contributions J.X. and G.P. carried out GC-MS, molecular dynamic simulation, wrote the paper and made the figures; Y.L. carried out molecular dy namic simulation; J.X., G.P. and J.X. carried out cloning, mutagenesis and protein production; L.T. interpreted the data; D.Y. designed research, interpreted the data and wrote the paper. 9
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Phytochemistry 181 (2021) 112573 Waterhouse, A., Bertoni, M., Bienert, S., Studer, G., Tauriello, G., Gumienny, R., Heer, F. T., de Beer, T.A.P., Rempfer, C., Bordoli, L., Lepore, R., Schwede, T., 2018. SWISSMODEL: homology modelling of protein structures and complexes. Nucleic Acids Res. 46 (W1), W296–W303. Wilderman, P.R., Peters, R.J., 2007. A single residue switch converts abietadiene synthase into a pimaradienen specific cyclase. J. Am. Chem. Soc. 129, 15736–15737. Williams, D.C., McGarvey, D.J., Katahira, E.J., Croteau, R., 1998. Truncation of limonene synthase preprotein provides a fully active ‘pseudomature’ form of this monoterpene cyclase and reveals the function of the amino-terminal arginine pair. Biochemistry 37, 12213–12220. Wise, M.L., Savage, T.J., Katahira, E., Croteau, R., 1998. Monoterpene synthase from common sage (Salvia officinalis). cDNA isolation, characterization, and functional expression of (+)-sabinene synthase, 1,8-cineole synthase and (+)-bornyl diphosphate synthase. J. Biol. Chem. 273, 14891–14899. Wise, M.L., Croteau, R., 1999. In: Cane, D.E. (Ed.), Comprehensive Natural Products Chemistry: Isoprenoids Including Carotenoids and Steroids, vol. 2. Elsevier Science, Oxford, UK, pp. 97–153. Xu, M., Wilderman, P.R., Peters, R.J., 2007. Following evolution’s lead to a single residue switch for diterpene synthase product outcome. Proc. Natl. Acad. Sci. U.S.A. 104, 7397–7401. Xu, J., Ai, Y., Wang, J., Xu, J., Zhang, Y., Yang, D., 2017. Converting S-limonene synthase to pinene and phellandrene synthases reveals the plasticity of the active site. Phytochemistry 137, 34–41. Yoshikuni, Y., Ferrin, T.E., Keasling, J.D., 2006. Designed divergent evolution of enzyme function. Nature 440, 1078–1082. Zerbe, P., Chiang, A., Bohlmann, J., 2012. Mutational analysis of white spruce (Picea glauca) ent-kaurene synthase (PgKS) reveals common and distinct mechanisms of conifer diterpene synthases of general and specialized metabolism. Phytochemistry 74, 30–39. Zhou, K., Peters, R.J., 2011. Electrostatic effects on (di) terpene synthase product outcome. Chem. Commun. 47, 4074–4080.
exploration of the catalytic landscape separating divergent plant sesquiterpene synthases. Nat. Chem. Biol. 4, 617–623. Oostenbrink, C., Villa, A., Mark, A.E., et al., 2010. A biomolecular force field based on the free enthalpy of hydration and solvation: the GROMOS force-field parameter sets 53A5 and 53A6. J. Comput. Chem. 25, 1656–1676. Pare, P.W., Tumlinson, J.H., 1999. Plant volatiles as a defense against insect herbivores. Plant Physiol. 121, 325–331. Phillips, M.A., Wildung, M.R., Williams, D.C., Hyatt, D.C., Croteau, R., 2003. cDNA isolation, functional expression, and characterization of (+)-alpha-pinene synthase and (-)-alpha-pinene synthase from loblolly pine (Pinus taeda): stereocontrol in pinene biosynthesis. Arch. Biochem. Biophys. 411, 267–276. Pyun, H.-J., Coates, R.M., Wagschal, K.C., McGeady, P., Croteau, R.B., 1993. Regiospecificity and isotope effects associated with the methy-methylene eliminations in the enzyme-catalyzed biosynthesis of (R) - and (S)-limonene. J. Org. Chem. 58, 3998–4009. Schwab, W., Williams, D.C., Davis, E.M., Croteau, R., 2001. Mechanism of monoterpene cyclization: stereochemical aspects of the transformation of noncyclizable substrates analog by recombinant (-)-limonene synthase, (+)-bornyl diphosphate synthase, and (-)-pinene synthase. Arch. Biochem. Biophys. 392, 123–136. Srividya, N., Davis, E.M., Croteau, R.B., Lange, B.M., 2015. Functional analysis of (4S)limonene synthase mutants reveals determinants of catalytic outcome in a model monoterpene synthase. Proc. Natl. Acad. Sci. U.S.A. 112, 3332–3337. Starks, C.M., Back, K., Chappell, J., Noel, J., 1997. Structural basis for cyclic terpene biosynthesis by tobacco 5-epi-aristolochene synthase. Science 277, 1815–1820. Tholl, D., 2006. Terpene synthases and the regulation, diversity and biological roles of terpene metabolism. Curr. Opin. Plant Biol. 9, 297–304. Thorpe, I.F., Brooks III, C.L., 2007. Molecular evolution of affinity and flexibility in the immune system. Proc. Natl. Acad. Sci. U.S.A. 104, 8821–8826. Umehara, M., Hanada, A., Yoshida, S., Akiyama, K., Arite, T., Takeda-Kamiya, N., Magome, H., Kamiya, Y., Shirasu, K., Yoneyama, K., Kyozuka, J., Yamaguchi, S., 2008. Inhibition of shoot branching by new terpenoid plant hormones. Nature 455, 195–200.
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Phytochemistry journal homepage: www.elsevier.com/locate/phytochem
Probing of the plasticity of the active site in pinene synthase elucidates its potential evolutionary mechanism Jingwei Xu 1, Guanzu Peng 1, Jinkun Xu, Yi Li, Li Tong, Dong Yang * Gene Engineering and Biotechnology Beijing Key Laboratory, College of Life Sciences, Beijing Normal University, Beijing, 100875, China
A R T I C L E I N F O
A B S T R A C T
Keywords: Paeonia lactiflora Paeoniaceae GC-MS Molecular dynamics Monoterpene Terpenene synthase Pinene Sabinene
Terpenes form a class of highly diverse natural products. The diversity of terpenes is created by terpene syn thases. During the reaction, carbocation intermediates form and their rearrangement could lead to the formation of various products. Terpene synthases determine the final product profile by controlling the conformation of the intermediate or stabilizing the carbocation. Pinene synthase is a monoterpene synthase catalyzing the cyclization of geranyl pyrophosphate (GPP) to form pinene. Our study suggests that by mutating the aromatic residue F482 to Ala, Val, Ile and Thr, the enzyme can be converted to sabinene synthase, with more than 90% of its total terpene products becoming sabinene, which indicates the aromaticity of this residue is essential for stabilizing the pinyl carbocation. We also identified a mutation S491A that could cause an about 29% increase in the overall activity of the enzyme without altering its produce selectivity. Molecular dynamic simulation indicates this mutation could decrease the flexibility of the enzyme when it forms a complex with the pinyl carbocation. Our study suggests the active pocket of pinene synthase has a certain level of plasticity, making it relatively easy to change the product selectivity or overall activity. This property could have an important implication in the evolution of terpene synthases and thereby terpene diversity, as by changing a few residues an enzyme could synthesize a completely different terpene product in a significant amount, which allows selection to take place.
1. Introduction Terpenes form one of the largest groups of natural products (Wise and Croteau, 1999; Tholl, 2006; Christianson, 2006, 2007; Degenhardt et al., 2009; Gao et al., 2012). As of 2012, more than 55,000 different types of terpenes have been reported (Gao et al., 2012). Terpenes play many biological roles. For example, they can serve as signaling mole cules mediating interactions between plants and insects (Pare and Tumlinson, 1999; Umehara et al., 2008). Cholesterol, also belonging to the terpene family, plays important roles in regulating membrane fluidity. Terpenes also have applications in food, cosmetics, medicine and other industries (Ajikumar et al., 2008; Kirby and Keasling, 2009). All terpenes are derived from a five-carbon precursor, isopentenyl pyrophosphate (IPP) (Wise and Croteau, 1999; Tholl, 2006; Christian son, 2006, 2007; Degenhardt et al., 2009; Gao et al., 2012). IPP and its isomer dimethylallyl pyrophosphate (DMAPP) are synthesized via either the mevalonate or the non-mevalonate pathway. The head-to-tail condensation of DMAPP and IPP forms geranyl pyrophosphate (GPP). Further condensation with more IPP units forms farnesyl pyrophosphate
(FPP), geranylgeranyl pyrophosphate (GGPP), etc. Cyclization of these linear products by terpene synthases is the key step to form the terpene diversity (Wise and Croteau, 1999; Tholl, 2006; Christianson, 2006, 2007; Degenhardt et al., 2009; Gao et al., 2012). Cyclized products can be modified by oxidation and other steps, further increasing the di versity of terpenoids. Cyclization is usually initiated by creating a carbocation in the active site of terpene synthases. The rearrangement of carbocations leads to the formation of a variety of skeletons of terpenes. This process is controlled by terpene synthases as they have evolved multiple mechanisms to limit the conformation of the substrate or stabilize a certain carbocation in termediate and prevent its rearrangement (Lesburg et al., 1997; Starks et al., 1997; Christianson, 2006, 2007; Gao et al., 2012). Aromatic or polar side chains are often involved in the latter process (Christianson, 2007; Zhou and Peters, 2011; Gao et al., 2012). Because of differences in these controlling mechanisms, the product selectivity of terpene syn thases could have large variations. Some enzymes are specific, pre dominantly producing one product, while others could be promiscuous and form a wide range of different products (O’Maille et al., 2008;
* Corresponding author. E-mail address: [email protected] (D. Yang). 1 These authors contributed equally to this work. https://doi.org/10.1016/j.phytochem.2020.112573 Received 27 June 2019; Received in revised form 19 October 2020; Accepted 22 October 2020 Available online 1 November 2020 0031-9422/© 2020 Elsevier Ltd. All rights reserved.
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Yoshikuni et al., 2006). The active pockets of terpene synthases often demonstrate a certain level of plasticity, as changing a few residue could completely alter the product profile (Morrone et al., 2008; Wilderman et al., 2007; Xu et al., 2007; Greenhagen et al., 2006; Yoshikuni et al., 2006; Keeling et al., 2008; Zerbe et al., 2012; Katoh et al., 2004). It has been proposed that product specificity of terpene synthases may evolve from their more promiscuous ancestors (Bohlmann et al., 1998; O’Brien and Herschlag, 1999; Copley, 2003; O’Maille et al., 2008). Monoterpenes are terpenes derived from GPP (Fig. 1A). Most of them are volatile scent compounds emitted by plants. The key intermediate for cyclic monoterpenes is the terpinyl cation (Pyun et al., 1993; Wil liams et al., 1998; Schwab et al., 2001; Hyatt et al., 2007; Srividya et al., 2015). If directly deprotonated, it forms limonene (Fig. 1A). The terpinyl cation can also undergo different types of rearrangement, which leads to the formation of other products (Fig. 1A). For example, 2, 7-ring closure forms the pinyl cation that is the precursor of α- and β-pinene. 6, 7-hy dride shift followed by 2, 6-ring closure forms the thujyl cation and then sabinene (Srividya et al., 2015). We previously studied the catalytic mechanism of Mentha spicata Slimonene synthase (Xu et al., 2017). Our data suggests that by intro ducing N345A/L423A/S454A mutations we can convert its activity to that of pinene synthase. Our further studies suggest that polar residues
in the active site of S-limonene synthase may form a “polar pocket” that participates in the stabilization of the terpinyl cation. Disruption of this pocket could lead to the rearrangement of the terpinyl cation and the formation of alternative products. In this study, we probed the key residues of Paeonia lactiflora pinene synthase. Our results suggest that by mutating the aromatic residue F482 to Ala, Val, Ile or Thr, the enzyme could be completely converted to sabinene synthase. We also identified a mutation S491A that can significantly enhance the enzyme activity without changing its product profile. These results show that pinene synthase, like S-limonene syn thase we studied before, has significant plasticity in its active site, which may explain the evolutionary history of monoterpene synthases. 2. Results 2.1. Homology modeling of P. lactiflora pinene synthase The crystal structures of pinene synthases have not been reported. Therefore, we constructed a homolog model of P. lactiflora pinene syn thase using the crystal structure of S-limonene synthase as the template. In order to investigate the catalytic mechanism, we docked three car bocation intermediates (terpinyl, pinyl and thujyl cations) into its active
Fig. 1. a, The proposed reaction mechanism for the synthesis of limonene, pinene and sabinene. b, The structural model of pinene synthase docked with the terpinyl (left), pinyl (middle) and thujyl (right) cations. Carbocations are colored in magenta (terpinyl), wheat (pinyl) and cyan (thujyl). Other atoms are colored according to their types (red: oxygen; yellow: carbon; blue: nitrogen; orange: phosphorus; green: magnesium). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.) 2
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site (Fig. 1B–D). The docking results suggest that carbocations bind to a similar location in the active pocket. Nearby residues include R305, S311, W314, I335, I338, N339, D342, Y417, I443, A444, I448, F482, D486, S491 and H571. Some of these residues could make hydrophobic interactions with the carbon skeleton of carbocations. Certain polar or aromatic residues may stabilize the positive charge and influence the product profile of the enzyme.
investigate this issue, we first docked the pinyl cation into the active pockets of WT and F482Y. It could be seen that in these two structures, the conformation of the docked pinyl cation is quite different (Fig. 4A). We then performed a 2 ns molecular dynamics simulation. The results suggest more conformational changes (Fig. 4B–C). Therefore, the extra hydroxyl group in Tyr could significantly change the conformation of the carbocation, which may explain the decreased activity and altered product profile.
2.2. Alanine scanning mutagenesis identifies mutations that cause an increase in sabinene production
2.4. S491A mutation enhances the overall activity of pinene synthase without changing its product profile
We first identified residues within 5 Å from the docked carbocations and performed alanine-scanning mutagenesis on them. Most of these mutations do not show dramatic changes in their product profiles. However, two mutations show interesting alteration in the product profile. One of them, F482A, essentially converts the enzyme to sabinene synthase. About 90.8% of its products is sabinene, while 97.7% of the products of the WT enzyme is α-pinene (Table 1, Fig. 2A–B). Another mutation, I335A, also causes an increase in sabinene production, though not as dramatic as F482A (Table 1). Importantly, although both mutants show an altered product profile, their overall activities are only moderately decreased (Table 1).
During our Ala scanning experiments, we identified a mutation S491A that enhances overall activities (Fig. 5A, D, Table 1). The overall activity of S491A mutant is about 29% higher than that of the WT enzyme (Table 1). Furthermore, S491A does not change the product selectivity, as more than 90% of their products remains α-pinene (Fig. 5A, Table 1). What is the mechanism of S491A to enhance the enzymatic activity? Since S491A still predominantly produces pinene, we docked the pinyl cation into the active pocket of the mutant enzyme and performed a 2 ns of molecular dynamics simulation. We also performed the same dynamic simulation on the WT enzyme in complex with the pinyl cation. Our data suggests that the active site residues in S491A have a much smaller RMSD than those in the WT enzyme, indicating the mutation S491A could cause the active pocket to be more rigid (Fig. 6A, STable 2). Interestingly, simulation on the WT or S491A enzymes in complex with the terpinyl or thujyl cation does not show the same differences in RMSD values of active site residues (Fig. 6B–C, STable 2). This probably in dicates that the S491A mutation increases the rigidity of the active pocket only when it binds to the pinyl cation. To better quantify the mobility of protein structures, we reanalyzed the simulation data using MDLovoFit (Martínez, 2015). This way we could subdivide the protein into regions with high and low mobility (Fig. 7A–C, STable 3). Interestingly, in S491A, fewer residues are clas sified as of high mobility than residues in WT enzymes (59% vs 63%). Furthermore, for residues classified with the highly mobile subset, those in S491A have lower average RMSD than those in WT proteins (0.253 nm vs. 0.278 nm). Therefore, there is a slight but significant increase in rigidity in S491A mutant protein.
2.3. Phenylalanine at the 482 position is required for pinene production We hypothesized that the aromatic ring of F482 might play a role in stabilizing the pinyl cation. To investigate this possibility, we mutated F482 to Val and Ile, two nonpolar residues with different side chain sizes. We also mutated it to Thr, Arg and Trp, representing polar, charged or non-Phe aromatic residues. The results suggest that any mutations that changes the phenylalanine at this position leads to an increase in sabinene production and almost abolishes the α-pinene production (Table 1). Replacing phenylalanine with nonpolar residues (F482I, F482V) all effectively convert the enzyme to sabinene synthase (Fig. 2C–D). More than 90% of products made by these mutants are sabinene (Table 1). Interestingly, F482T also predominantly produces sabinene, which seems to suggest that a polar residue here is not suffi cient to stabilize the pinyl cation (Table 1, Fig. 2E). Other mutations (F482Y, F482R and F482W) retain very little activities (Table 1). In order to verify that the mutant enzymes indeed produce sabinene, we analyzed the reaction product of F482L using the high resolution hybrid quadrupole-orbitrap GC-MS/MS system (Fig. 3A–C). By comparing with the sabinene standard, the peak eluted at 9.29 min was identified as sabinene. Its mass spectrum was measured and could be readily matched with that of sabinene standard (Fig. 3B–C). It is interesting to note that replacement of F482 to other aromatic residues (Tyr or Trp) causes a drastic decrease in enzyme activities. To
2.5. Combination of S491A with F482 mutants has mixed results in the overall activity Since all four sabinene synthase mutants show some decreases in the catalytic activity, we wondered whether we may restore their activities to that of the WT pinene synthase by combing with S491A. We produced
Table 1 Major products (%±SD) of WT α-pinene synthase and enzymes with mutations. Enzymes
PS-WT F482A S491A F482I F482L F482V F482T F482Y F482R F482W I335A F482A/S491A F482I/S491A
Relative Proportion of Products (%) Unknown
α-pinene
sabinene
limonene
Unknown
Unknown
RT = 4.52
RT = 7.05
RT = 8.55
RT = 11.05
RT = 11.58
RT = 12.72
0.31 ± 0.02 0.05 ± 0.00 0.46 ± 0.01 1.27 ± 0.02 0.96 ± 0.06 1.03 ± 0.01 0.00 ± 0.00 0.00 17.62 ± 6.21 16.32 ± 5.33 3.52 ± 0.02 1.34 ± 0.11 0.63 ± 0.02
97.86 ± 0.21 4.43 ± 0.03 97.28 ± 0.09 3.17 ± 0.08 2.03 ± 0.03 2.12 ± 0.03 3.53 ± 0.00 2.71 0.00 ± 0.00 0.00 ± 0.00 78.00 ± 0.03 4.17 ± 0.01 2.72 ± 0.00
0.96 ± 0.00 93.21 ± 0.06 1.14 ± 0.00 92.35 ± 0.21 94.26 ± 0.30 94.51 ± 0.07 93.91 ± 0.00 31.00 18.96 ± 7.19 29.92 ± 17.90 12.94 ± 0.00 88.40 ± 1.43 92.88 ± 0.45
0.26 ± 0.01 1.59 ± 0.05 0.12 ± 0.00 0.66 ± 0.09 0.31 ± 0.01 0.30 ± 0.03 1.64 ± 0.04 63.61 31.04 ± 19.27 21.67 ± 6.84 0.57 ± 0.02 3.38 ± 0.14 2.40 ± 0.09
0.54 ± 0.05 0.72 ± 0.03 0.99 ± 0.06 2.33 ± 0.08 1.91 ± 0.17 1.82 ± 0.06 0.92 ± 0.05 2.68 27.33 ± 14.94 27.35 ± 14.96 4.28 ± 0.03 2.36 ± 0.34 1.19 ± 0.08
0.07 ± 0.00 ± 0.00 ± 0.23 ± 0.29 ± 0.23 ± 0.00 ± 0.00 5.05 ± 4.74 ± 0.67 ± 0.35 ± 0.18 ±
3
0.00 0.00 0.00 0.00 0.01 0.00 0.00 0.51 0.45 0.00 0.01 0.00
Activity(% WT) 100 46.63 ± 4.47 128.90 ± 4.19 18.36 ± 2.51 49.98 ± 39.78 29.50 ± 12.39 52.69 ± 3.17 0.62 1.28 ± 0.01 1.54 ± 0.02 54.36 ± 2.17 18.73 ± 0.13 28.74 ± 5.73
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Fig. 2. Converting pinene synthase to sabinene synthase by mutations on 482 position. Chromatograms in a, b, c, d and e show the GC-MS analysis of terpene products for WT, F482A, F482I, F482V and F482T, respectively. The x-axis is the retention time and the y-axis is the relative abundance of each species. The numbers in each peak correspond to α-pinene (1), sabinene (2) and limonene (3).
double mutants F482A/S491A and F482I/S491A. Both seem to retain the same product selectivity of the original single mutants, though F482A/S491A shows an insignificant decrease in sabinene production (P = 0.39; Fig. 5B–C, Table 1). In overall activities, F482I/S491A shows an insignificant increase in activity (P = 0.27), while F482A/S491A has a significant decrease in activity (P < 0.05; Fig. 5D, Table 1). Therefore, it seems that the effect of combining two mutants is more complicated than we expected. We asked whether the combination of F482A and S491A mutations could affect the rigidity of the protein. After a 2 ns of molecular dy namics simulation and data analysis using MDLovoFit, we found the rigidity of F482A/S491A is similar to that of S491A (STable 3). Appar ently, the decrease of activity by the combination of F482A with S491A cannot be explained by the overall RMSD change. To further investigate this issue, we analyze the RMSF values based on the simulation data (SFig. 1A–C). Compared with S491A and WT, F482A/S491A shows an RMSF spike at a region between residues 219 and 223. In the structural model, this region forms a loop in the protein structural model (SFig. 2). However, this loop is quite far from the active site (about 40 Å). Therefore, its mobility could only affect the enzyme activity indirectly. For example, it could influence the closure of the active pocket. We also superimposed the structural model of WT, S491A and F482A/S491A in complex with the docked pinyl cation (SFig. 3). In the
structural model, the pinyl cation is docked between the residue 491 and 482 (SFig. 3). Interestingly, in WT and S491A, the position of the pinyl cation is similar, while in F482A/S491A, the pinyl cation is shifted to wards the A482. This change in the carbocation position may impair the enzyme activity (SFig. 3). 3. Discussion The plasticity of the active pocket has been investigated in several terpene synthases. Our previous study in S-limonene synthase has shown that mutating three residues could readily convert limonene synthase to pinene synthases (Xu et al., 2017). In this case, about 70% of the total products become pinene or phellandrene in mutant enzymes. Here, we performed a similar study in pinene synthase. By mutating a single residue, we achieved an almost complete conversion of pinene synthase to sabinene synthase, resulting more than 90% of the total products to be sabinene. These findings prove that the active pocket of monoterpene synthases have a certain level of plasticity and dramatic changes in product profile can be achieved by a small number of mutations. In limonene synthase, we showed that polar residues are important to stabilize the terpinyl cation (Xu et al., 2017). We identified a polar pocket and found that mutating key residues in this pocket could lead to the rearrangement of the terpinyl cation and cause the production of alternative products (Xu et al., 2017). In pinene synthase, however, we 4
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Fig. 3. Hybrid quadrupole-orbitrap GC-MS/MS. a shows the chromatogram of terpene products of F482L. The elution peak corresponding to sabinene is labeled. The mass spectrum of the product eluted at 9.29 min from a is shown in b. The mass spectrum of the standard sabinene is shown in c.
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synthase do not change the product profile, as α-pinene still composes of more than 90% of the total products. Dynamic simulation suggests that the mutation S491A could decrease the flexibility of the active pocket only when it binds the pinyl cation. Therefore, it is possible that this mutation makes an enzyme that is already specific for pinene production more rigid. Probably this rigidness stabilizes the pinyl cation intermediate in the active pocket, which enhances the overall activity without changing the product se lection. Interestingly, during the maturation of antibodies, a decrease in protein flexibility is often associated with the increased affinity, prob ably due to the decrease in the entropic cost for antigen binding (Thorpe and Brooks, 2007). Here, in a completely different setting, we found this mechanism may also play a similar role. However, we have to be prudent to apply this mechanism. The enhancement of rigidity of the overall protein may indeed play a role in activity enhancement in S491A. But in F482A/S491A, the RMSD of the overall structure seems to change little. The decrease of activity in F482A/S491A could probably be caused by the positional change of the pinyl cation in the active pocket or the increased flexibility in the loop between 219 and 223. Other causes could also be possible and further investigations are required. One question remains. Why only when S491A binds the pinyl cation does the protein rigidity increase? As our simulation on the F482Y mutant protein indicates, the conformation of the carbocation is highly sensitive to the slight change in the amino acid in the active pocket. Probably, the opposite is also true. Therefore, the difference in the carbocation may also play an important role in the conformation of the enzyme. The basis of terpene diversity is terpene synthases. Since a slight alteration by a few mutations in the active pocket could significantly change the product profile, the expansion of the chemical space of ter penoids could be easily achieved. It is likely that the ancestor of modern terpene synthases are quite promiscuous and produce a multitude of products. New enzymes that produce novel terpene products could evolve by a few mutations in the active pocket. If these new products are beneficial to the organism (e.g. attracting pollinating insects), enzymes
Fig. 4. Conformational differences in the pinyl cation in WT and F482Y as demonstrated by molecular dynamics simulation. The structures of WT and F482Y are superimposed. Results after 0, 1 and 2 ns of simulation are shown in a, d and c, respectively. Carbon in WT is colored in yellow and carbon in F482Y is colored in green. Oxygen is colored in red. Nitrogen is colored in blue. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)
found the aromatic residue F482 is a key player. Mutating this residue to other polar residues could not maintain the pinene synthase activity, as the result of F482T proves. Actually, it seems the residue in this position must be a Phe for pinene synthase, as a similar aromatic residue, Tyr, does not work (Table 1). This phenomenon could probably be explained by the possible geometric constraint in this area so even the addition of a single hydroxyl group in Tyr can significantly change the conformation of the pinyl cation, as our simulation suggests (Fig. 4). This conforma tional change could explain the decrease of the overall enzyme activity as well as the change in the product profile. Therefore, it seems that within monoterpene synthases, distinct mechanisms are used to stabilize the carbocation intermediate and determine the product selectivity. The plasticity of the active pocket is also reflected on the relative ease of changing the overall activity of the enzyme. In both the previous study on limonene synthase and this study on pinene synthase, we showed the alteration of a single amino acid is sufficient to enhance the overall enzymatic activity, which means the mutant enzyme can pro duce more terpenoid products, regardless of its types. For example, the mutation S454G is about 7% more active than the WT limonene syn thase. In pinene synthase, S491A is about 29% more active than the WT enzyme. Different from S454G in limonene synthase, S491A in pinene
Fig. 5. The effect of the S491A mutation. Chromatograms in a, b and c show the GC-MS analysis of terpene production for S491A, F482A/S491A, and F482I/S491A, respectively. The x-axis is the retention time and the y-axis is the relative abundance of each species. The numbers in each peak correspond to α-pinene (1), sabinene (2) and limonene (3). d shows the overall activity and the percentage of sabinene within total products produced by F482A, F482A/S491A, F482I and F482I/S491A. The asterisk indicates P < 0.05. 6
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Fig. 6. S491A mutation increases the rigidity of the active pocket when it binds the pinyl cation. RMSD values of the active site residues between WT and S491A in complex with the pinyl cation (a), terpinyl cation (b) and thujyl cation (c) are compared.
producing them is likely to be selected and further evolution may nar row the range of products, producing a new enzyme that specifically produces this compound. One important difficulty that a new terpene synthase faces is the fact that normally a mutation decreases the overall activity of the enzyme. Therefore, although the new enzyme may produce a novel compound, because its activity is lower, it may not be able to produce a sufficient amount of new compounds that could benefit the organism. However, our study has demonstrated that the overall activity of the enzyme could be enhanced by a single mutation. The cause of the activity increase could be through the increase of the rigidity of the overall protein or the
alteration of fluctuation of some structural elements. Other mechanisms may also be involved. Obviously, more research should still be per formed to delineate the possible mechanism involved in this process. Another important issue is about the chirality of products in our enzyme. Unfortunately, due to the difficulty of obtaining chiral stan dards, we could not perform a chiral GC-MS. Furthermore, because of the low yield of the enzymatic reaction, we were not able to perform NMR to determine the absolute configuration of terpene products. However, by analyzing protein sequences of pinene and sabinene syn thases with known chirality of their products, we could still make certain conclusions. 7
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Fig. 7. The RMSD profiles of the region with high (RMSD H) and low (RMSD L) mobility from the simulation of WT (a), S491A (b) and F482/S491A (c) in complex with the pinyl cation.
Several studies on pinene synthases reported the chirality of its products (Hall et al., 2013; McKay et al., 2003; Phillips et al., 2003; Williams et al., 1998). By making a multiple sequence alignment, we could identify possible key residues that may determine the chirality of the product (SFig. 4). The most interesting residue is the one corre sponding to I443 in our enzyme. In reported (+)-α-pinene synthases, this residue is always a Phe, while in reported (− )-α-pinene synthases, it is always a Ser. By looking at the homology model of our enzyme, we found I443 is close to the docked pinyl cation (Fig. 1C). So the size of this residue is likely to influence the chirality of the product. For example, a bulky side chain such as Phe may cause steric clash and prevent the formation of (− )-α-pinene. Therefore, as in our enzyme this residue is a bulky Ile, the wild type form of our enzyme is likely to produce (+)-α-pinene. As a further proof, phylogenic analysis also shows our enzyme is more closely related to (+)-α-pinene synthases than (− )-α-pinene synthases (SFig. 5). Unfortunately, we could not find any reports on (− )-sabinene syn thases, which makes it difficult to make sequence analysis directly. However, it is known that both (+)-pinene and (+)-sabinene require the carbocation intermediate (3R)-linalyl diphosphate (Wise et al., 1998). Therefore, if the wild type form of our enzyme indeed produces (+)-pinene, it is most likely the sabinene produced in our mutant enzyme is (+)-sabinene, since they share the same carbocation intermediate. Certainly, our interpretation on the rigidity of the enzyme as well as the chirality of products is still relatively speculative. In the future, more experiments should be performed to validate our predictions on both cases.
sequence, we only synthesized the sequence encoding the protein sequence from R51 to the C-terminus. The codon usage was optimized for the expression in E. coli. It was cloned into pGEX-4T-1 vector. Point mutations were introduced by the site-directed mutagenesis kit (Trans gen and Takara). Primers used were listed in STable 1. The amplification products were digested by DpnI for 5 h at 37 ◦ C and transformed into E. coli DH5α cells. Correct mutations were identified by sequencing. 5.2. Protein expression and purification Plasmids were transformed into E. coli BL21 (DE3) cells and were then cultured in LB medium with 50 mM ampicillin. Expression was induced by 0.5 mM IPTG at 20 for 19 h. Cells were harvested by centrifugation at 3000 g for 10 min, resuspended by buffer A (25 mM Tris, pH 8.0, 150 mM NaCl, 1 mM DTT, 0.2 mM PMSF) and lysed with sonication. The lysate was applied to glutathione sepharose resin and proteins were then cleaved on-column by thrombin that was diluted in 25 mM Tris, pH 8.0, 150 mM NaCl and 1.25 mM CaCl2 to 1 KU. Finally, proteins were eluted with buffer E (25 mM Tris, pH 8.0, 150 mM NaCl, 1 mM DTT, and 0.5 mM EDTA). 5.3. Enzymatic reaction and GC-MS assay The reaction was performed in a 400 μl reaction system that contains 25 mM Mops, pH 7.0, 15 mM MgCl2, 1.0 mM DTT, 10% (v/v) glycerol, 68.5 μM GPP and 1 μg pinene synthase. The reaction was performed at 37 ◦ C for 40 min, and then 400 μl of hexane was added to extract terpene. The mixture was incubated at 37 ◦ C for 20 min and vortexed for 30 min. Then it was centrifuged at 8000 g for 5 min to separate the organic layer that contains terpene products. 1 μl of extraction was analyzed by GC-MS (Thermo Trace GC Ultra & ISQ). The extraction was injected into a HP-88 column (100 mm × 250 μM x 0.20 μM thickness; Agilent), with ultra-purity helium as the carrier gas at the flow rate 1.0 ml/min. The oven temperature was set to 3 phases; the temperature was first increased to 50 ◦ C and kept for 1 min, then increased to 130 ◦ C at the increment of 3 ◦ C/min, and increased to 220 ◦ C at the increment of 50 ◦ C/min and held for 5 min. The injector and transfer line temperature were set at 250 ◦ C. Terpenes were identified by GC retention times by comparing with available authentic standards. The proportion of each product was based on the ratio of the peak area of the particular product and the peak area of total terpene products. The relative activity of an enzyme is calculated by dividing the total peak area for all terpene products of one mutant enzyme by that of the wild type (WT) pinene synthase.
4. Conclusions In conclusion, we have demonstrated the plasticity of the active pocket of pinene synthase in our study. Our data suggested mutating a single residue could dramatically change the product selectivity of the enzyme, converting it to sabinene synthase. We also demonstrated a single mutation could significantly enhance the overall activity of the enzyme without changing its product profile, probably caused by the increase in rigidity of the protein. Our findings could provide important insights on the evolution of new monoterpene synthases and the di versity of monoterpenes. 5. Methods 5.1. Gene synthesis and site-directed mutagenesis The P. lactiflora α-pinene synthase (PS) cDNA was synthesized by Sangon Biotech (China). According to the findings of Williams and his colleagues (Williams et al., 1998), we identified the potential plastidial targeting sequence by aligning its protein sequence with that of M. spicata S-limonene synthase. To eliminate this plastidial targeting
5.4. Hybrid quadrupole-orbitrap GC-MS/MS system analysis The reaction product was analyzed by the Hybrid QuadrupoleOrbitrap GC-MS/MS System (ThermoFisher Q Exactive GC). 1 μl of extraction was injected with ultra purity helium as the carrier gas at the 8
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flow rate 1.0 ml/min, with split in 20 ml/min. The oven temperature was set to 3 phases, the temperature was first increased to 50 ◦ C and kept for 1 min, then increased to 100 ◦ C at the increment of 3 ◦ C/min, and increased to 220 ◦ C at the increment of 50 ◦ C/min and held for 5 min. The injector and transfer line temperature were set at 250 ◦ C. High resolution EI fragment spectra were acquired using 60,000 FWHM res olution, with full scan mode mass range of 50–550 Da. To identify the reaction products, their gas chromatography retention time and mass spectra were compared with those of the sabinene standard.
Declaration of competing interest The authors declare no competing financial interest. Acknowledgements This work is supported by grants from the Fok Ying Tung Education Foundation (#132025), the National Science Foundation of China (#31070682 and #31640004), State Education Ministry and the Pro gram of the Co-Construction with Beijing Municipal Commission of Education of China. We thank the Center of Biomedical Analysis, Tsinghua University, for the technical assistance in GC-MS.
5.5. Construction of structural models for protein-ligand complexes For proteins, the homology model of the α-pinene synthase from P. lactiflora (GenBank: ANA91931.1) was constructed by Swiss-model (Waterhouse et al., 2018) using the structure of 4S-limonene synthase from M. spicata (2ONH) as the template. Based on the structural model of the WT α-pinene synthase, a series of mutants structural models were constructed by PyMoL version 2.1.1. For ligands, the structure of carbocation intermediates (the terpinyl, pinyl and thujyl cations) were generated by MOE v2008.1 (Chemical Computing Group Inc.). They were then docked into the active site of pinene synthase models in Autodock 4.2.5.1 using default settings (Morris et al., 2009). The structures of protein-carbocation complexes were then optimized in ATB (Malde et al., 2011). The coordinate of Mg2+ was adapted from the coordinate of Mn2+ in the limonene syn thase structure by superimposing the limonene synthase structure with the pinene synthase structural model. In the same way, the pyrophos phate group was adapted from the pyrophosphate tail in the limonene synthase structure. The topology and force field files for carbocations and the pyrophosphate tail were generated and optimized by ATB.
Appendix A. Supplementary data Supplementary data to this article can be found online at https://doi. org/10.1016/j.phytochem.2020.112573. References Ajikumar, P.K., Tyo, K., Carlsen, S., Mucha, O., Phon, T.H., Stephanopoulos, G., 2008. Terpenoids: opportunities for biosynthesis of natural product drugs using engineered microorganisms. Mol. Pharm. 5, 167–190. Berendsen, H.J.C., Spoel, D.V.D., Drunen, R.V., 1995. GROMACS: a message-passing parallel molecular dynamics implementation. Comput. Phys. Commun. 91, 43–56. Bohlmann, J., Meyer-Gauen, G., Croteau, R., 1998. Plant terpenoid synthases: molecular biology and phylogenetic analysis. Proc. Natl. Acad. Sci. U.S.A. 95, 4126–4133. Christianson, D.W., 2006. Structure biology and chemistry of the terpenoid cyclases. Chem. Rev. 106, 3412–3442. Christianson, D.W., 2007. Roots of biosynthetic diversity. Science 316, 60–61. Copley, S.D., 2003. Enzymes with extra talents: moonlighting functions and catalytic promiscuity. Curr. Opin. Chem. Biol. 7, 265–272. Degenhardt, J., K¨ ollner, T.G., Gershenzon, J., 2009. Monoterpene and sesquiterpene synthases and the origin of terpene skeletal diversity in plants. Phytochemistry 70, 1621–1637. Gao, Y., Honzatko, R.B., Peters, R.J., 2012. Terpenoid synthase structures: a so far incomplete view of complex catalysis. Nat. Prod. Rep. 29, 1153–1175. Greenhagen, B.T., O’Maille, P.E., Noel, J.P., Chappell, J., 2006. Identifying and manipulating structural determinates linking catalytic specificities in terpene synthases. Proc. Natl. Acad. Sci. U.S.A. 103, 9826–9831. Hall, D.E., Yuen, M.M., Jancsik, S., Quesada, A.L., Dullat, H.K., Li, M., Henderson, H., Arango-Velez, A., Liao, N.Y., Docking, R.T., Chan, S.K., Cooke, J.E., Breuil, C., Jones, S.J., Keeling, C.I., Bohlmann, J., 2013. Transcriptome resources and functional characterization of monoterpene synthases for two host species of the mountain pine beetle, lodgepole pine (Pinus contorta) and jack pine (Pinus banksiana). BMC Plant Biol. 13, 80. Hyatt, D.C., Youn, B., Zhao, Y., Santhamma, B., Coates, R.M., Croteau, R.B., Kang, C., 2007. Structure of limonene synthase, a simple model for terpenoid cyclase catalysis. Proc. Natl. Acad. Sci. U.S.A. 104, 5360–5365. Katoh, S., Hyatt, D., Croteau, R., 2004. Altering product outcome in Abies grandis (-)-limonene synthase and (-)-limonene/(-)-alpha-pinene synthase by domain swapping and directed mutagenesis. Arch. Biochem. Biophys. 425, 65–76. Keeling, C.I., Weisshaar, S., Lin, R.P.C., Bohlmann, J., 2008. Functional plasticity of paralogous diterpene synthases involved in conifer defense. Proc. Natl. Acad. Sci. U. S.A. 105, 1085–1090. Kirby, J., Keasling, J.D., 2009. Biosynthesis of plant isoprenoids: perspectives for microbial engineering. Annu. Rev. Plant Biol. 60, 335–355. Lesburg, C.A., Zhai, G., Cane, D.E., Christianson, D.W., 1997. Crystal structure of pentalenene synthase: mechanistic insights on terpenoid cyclization reactions in biology. Science 277, 1820–1824. Malde, A.K., Zuo, L., Breeze, M., et al., 2011. An automated force field topology builder (ATB) and repository: version 1.0. J. Chem. Theor. Comput. 7, 4026–4037. Martínez, L., 2015. Automatic identification of mobile and rigid substructures in molecular dynamics simulations and fractional structural fluctuation analysis. PloS One 10, e0119264. McKay, S.A., Hunter, W.L., Godard, K.A., Wang, S.X., Martin, D.M., Bohlmann, J., Plant, A.L., 2003. Insect attack and wounding induce traumatic resin duct development and gene expression of (-)-pinene synthase in Sitka spruce. Plant Physiol. 133, 368–378. Morris, G.M., Huey, R., Lindstrom, W., Sanner, M.F., Belew, R.K., Goodsell, D.S., Olson, A.J., 2009. Autodock4 and AutoDockTools4: automated docking with selective receptor flexibility. J. Comput. Chem. 16, 2785–2791. Morrone, D., Xu, M., Fulton, D.B., Determan, M.K., Peters, R.J., 2008. Increasing complexity of a diterpene synthase reaction with a single residue switch. J. Am. Chem. Soc. 130, 5400–5401. O’Brien, P.J., Herschlag, D., 1999. Catalytic promiscuity and the evolution of new enzymatic activities. Chem. Biol. 6, R91–R105. O’Maille, P.E., Malone, A., Dellas, N., Hess Jr., B.A., Smentek, L., Sheehan, I., Greenhagen, B.T., Chappell, J., Manning, G., Noel, J.P., 2008. Quantitative
5.6. Molecular simulation The protein-ligand complex was then solvated in a 0.707 × 6 × 6 × 6 nm3 dodecahedron water box (spc216 water model) with Na+ and CL− ions in GROMACS version 5.1.4 (Berendsen et al., 1995; Oostenbrink et al., 2010). For energy minimization, 200ps NVT ensemble (constant Number of particles, Volume, and Temperature) and 200ps NPT ensemble (constant Number of particles, Pressure, and Temperature) were performed in GROMACS version 5.1.4 using GROMOS96 54a7 force field (Berendsen et al., 1995; Oostenbrink et al., 2010). The equilibrated systems were then used for molecular dynamics simulation in GROMACS using GROMOS96 54a7 force field for 2ns in NPT ensemble (300K, 1 atm). To investigate the RMSD of the active pocket residues, we first defined the active pocket as the residues within 5 Å from the docked carbocations. 5.7. Analysis of the simulation data RMSD values of these residues were calculate based on MD trajec tories produced by the molecular dynamic simulation mentioned above. The simulation data was further analyzed with MDLovoFit-16.264 (Martínez, 2015). Each protein was divided into two subsets according to the structural fluctuation. Then RMSD L (low fluctuation fraction), RMSD H (high fluctuation fraction) and RMSF were obtained. Author contributions J.X. and G.P. carried out GC-MS, molecular dynamic simulation, wrote the paper and made the figures; Y.L. carried out molecular dy namic simulation; J.X., G.P. and J.X. carried out cloning, mutagenesis and protein production; L.T. interpreted the data; D.Y. designed research, interpreted the data and wrote the paper. 9
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exploration of the catalytic landscape separating divergent plant sesquiterpene synthases. Nat. Chem. Biol. 4, 617–623. Oostenbrink, C., Villa, A., Mark, A.E., et al., 2010. A biomolecular force field based on the free enthalpy of hydration and solvation: the GROMOS force-field parameter sets 53A5 and 53A6. J. Comput. Chem. 25, 1656–1676. Pare, P.W., Tumlinson, J.H., 1999. Plant volatiles as a defense against insect herbivores. Plant Physiol. 121, 325–331. Phillips, M.A., Wildung, M.R., Williams, D.C., Hyatt, D.C., Croteau, R., 2003. cDNA isolation, functional expression, and characterization of (+)-alpha-pinene synthase and (-)-alpha-pinene synthase from loblolly pine (Pinus taeda): stereocontrol in pinene biosynthesis. Arch. Biochem. Biophys. 411, 267–276. Pyun, H.-J., Coates, R.M., Wagschal, K.C., McGeady, P., Croteau, R.B., 1993. Regiospecificity and isotope effects associated with the methy-methylene eliminations in the enzyme-catalyzed biosynthesis of (R) - and (S)-limonene. J. Org. Chem. 58, 3998–4009. Schwab, W., Williams, D.C., Davis, E.M., Croteau, R., 2001. Mechanism of monoterpene cyclization: stereochemical aspects of the transformation of noncyclizable substrates analog by recombinant (-)-limonene synthase, (+)-bornyl diphosphate synthase, and (-)-pinene synthase. Arch. Biochem. Biophys. 392, 123–136. Srividya, N., Davis, E.M., Croteau, R.B., Lange, B.M., 2015. Functional analysis of (4S)limonene synthase mutants reveals determinants of catalytic outcome in a model monoterpene synthase. Proc. Natl. Acad. Sci. U.S.A. 112, 3332–3337. Starks, C.M., Back, K., Chappell, J., Noel, J., 1997. Structural basis for cyclic terpene biosynthesis by tobacco 5-epi-aristolochene synthase. Science 277, 1815–1820. Tholl, D., 2006. Terpene synthases and the regulation, diversity and biological roles of terpene metabolism. Curr. Opin. Plant Biol. 9, 297–304. Thorpe, I.F., Brooks III, C.L., 2007. Molecular evolution of affinity and flexibility in the immune system. Proc. Natl. Acad. Sci. U.S.A. 104, 8821–8826. Umehara, M., Hanada, A., Yoshida, S., Akiyama, K., Arite, T., Takeda-Kamiya, N., Magome, H., Kamiya, Y., Shirasu, K., Yoneyama, K., Kyozuka, J., Yamaguchi, S., 2008. Inhibition of shoot branching by new terpenoid plant hormones. Nature 455, 195–200.
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