nature CHeMICaL BIOLOGY | VOL 13 | SEPTEMBER 2017 | www.nature.com/naturechemicalbiology 1009

articlepuBLIsHed OnLIne: 24 juLY 2017 | dOI: 10.1038/nCHeMBIO.2435

Nonribosomally synthesized peptides represent a class of natural products, including therapeutically established com-pounds such as cyclosporine and vancomycin, with a plethora

of chemical structures and bioactivities1. Their biosynthesis occurs on large protein templates—the NRPSs2–4. The multimodular NRPSs facilitate an ATP-driven stepwise biosynthesis of the peptide chain in a mechanism reminiscent of an assembly line.

In a typical NRPS the number and arrangement of modules cor-relate in a colinear fashion with the amino acid sequence in the product5. Modules can be further subdivided into domains for the individual catalytic steps and the central peptidyl-carrier-protein (PCP) domain, which shuttles the reactive intermediates between the catalytic centers and to the next module. The PCP domain covalently binds a single aminoacyl or peptidyl intermediate as a thioester on the terminal sulfhydryl group of its prosthetic group 4′-phosphopantetheine (HS-Ppant). The A domain activates its cognate amino acid with ATP to the aminoacyl adenylate (amino acid–AMP) and charges it onto the PCP domain to give the amino acid–S-Ppant-PCP thioester. The PCP domain can then interact with a condensation (C) domain from the upstream and/or downstream module to effect peptide bond formation, with other domains for chemical diversification or with a thioesterase (TE) domain, which is typically involved in product release from the protein template.

What is poorly understood about this assembly-line arrangement is how interactions between catalytic and carrier domains are con-trolled and timed6,7. The length of the HS-Ppant moiety (maximum ~20 Å) is not sufficient to reach all catalytic sites. Crystal structures of NRPS modules have revealed that only one of the catalytic sites present is in reach of the Ppant’s action radius in a single confor-mation8–11. These findings indicate that substantial conformational changes, including PCP translocations, must occur during each elongation cycle. Similar domain movements have been proposed for acyl carrier proteins in modular polyketide synthases and fatty acid synthases12,13. However, experimental verification of PCP trans-locations in solution has not yet been reported, and how these are orchestrated is currently unknown.

Crystal structures of A domains have also revealed at least three different functional conformations8,14,15. A domains are members of the acyl-CoA synthetase, NRPS adenylation domain, luciferase enzyme (ANL) superfamily of adenylating proteins2,16. They com-prise a large N-terminal subdomain (AN) and a small C-terminal subdomain (AC) linked through a short hinge region, shown for the first time for the phenylalanine-activating A domain (PheA) from gramicidin S synthetase I (ref. 14). In the absence of sub-strates the two subdomains show no or only weak interaction, and the A domain can be observed in an ensemble of open (O) confor-mations. In the presence of substrates, two distinct closed confor-mations can be adopted to form different active sites in the cleft between the subdomains. According to the domain-alternation model15,16, the enzyme in the adenylation conformation (A confor-mation) converts the amino acid and ATP into amino acid–AMP and PPi. Subsequently, the AC subdomain rotates by 140° relative to the AN subdomain to adopt the thiolation conformation (T confor-mation) for the second half-reaction of thioester formation15,17–21. In the A conformation, a strictly conserved lysine from the A10 sequence motif2 coordinates the amino acid and ATP (K517 in PheA)14, whereas in the T conformation, a conserved lysine of the A8 sequence motif from the opposite side of AC coordinates the aminoacyl adenylate from a nearly identical position relative to AN (K434 in PheA)15 (Supplementary Results, Supplementary Fig. 1). Mutation of one of these residues abolishes or impairs the respective half-reaction but leaves the protein catalytically competent for the other half-reaction22–25. The displacement of pyrophosphate (PPi) generated in the adenylation reaction is proposed to occur with the rotation into the T conformation. In the latter conformation, a conserved arginine from the A8 motif (R439 in PheA) forms a salt bridge with a glutamate from the A5 motif (E327 in PheA). It occupies the space taken by the β- and γ-phosphates of ATP in the A conformation, effectively creating a control switch15,26 (Supplementary Fig. 1f). In the T conformation, formation of a new composite interaction surface helps recruit the PCP domain and leads to the transfer conformation of the A-PCP

1Institute of Biochemistry, Department of Chemistry and Pharmacy, University of Muenster, Münster, Germany. 2Department of Chemistry, Princeton University, Princeton, New Jersey, USA. 3Molecule and Life Nonlinear Sciences Laboratory, Research Center of Mathematics for Social Creativity, Research Institute for Electronic Science (RIES), Hokkaido University, Sapporo, Japan. *e-mail: henning.mootz@uni-muenster.de

Fret monitoring of a nonribosomal peptide synthetasejonas alfermann1, Xun sun2   , Florian Mayerthaler1, thomas e Morrell2   , eva dehling1, Gerrit Volkmann1, tamiki Komatsuzaki3, Haw Yang2 & Henning d Mootz1*   

Nonribosomal peptide synthetases (NRPSs) are multidomain enzyme templates for the synthesis of bioactive peptides. Large-scale conformational changes during peptide assembly are obvious from crystal structures, yet their dynamics and coupling to catalysis are poorly understood. We have designed an NRPS FRET sensor to monitor, in solution and in real time, the adoption of the productive transfer conformation between phenylalanine-binding adenylation (A) and peptidyl-carrier-protein domains of gramicidin synthetase I from Aneurinibacillus migulanus. The presence of ligands, substrates or intermediates induced a distinct fluorescence resonance energy transfer (FRET) readout, which was pinpointed to the population of specific confor-mations or, in two cases, mixtures of conformations. A pyrophosphate switch and lysine charge sensors control the domain alternation of the A domain. The phenylalanine–thioester and phenylalanine–AMP products constitute a mechanism of product inhibition and release that is involved in ordered assembly-line peptide biosynthesis. Our results represent insights from solution measurements into the conformational dynamics of the catalytic cycle of NRPSs.

© 2

017

Nat

ure

Am

eric

a, In

c., p

art

of

Sp

rin

ger

Nat

ure

. All

rig

hts

res

erve

d.

1010 nature CHeMICaL BIOLOGY | VOL 13 | SEPTEMBER 2017 | www.nature.com/naturechemicalbiology

article NATuRE chEMIcAL bIOLOgy dOI: 10.1038/nCHeMBIO.2435

ensemble. Crystal structures using vinylsulfonamide analogs of the aminoacyl-AMP capture this state with a covalently trapped Ppant thiol group9,20,21.

Structural investigation of NRPS by NMR in solution is com-plicated by the size of the domains (the A domain is ~60 kDa) and has been restricted so far to PCP, TE and PCP-TE constructs27–29. A previous study proposed that internal conformational changes of the PCP domain contribute to specific interactions with the domain partners27, but these have not been observed in structures of multidomain NRPS constructs6,28. Conformational changes coupled to A domain activity were also investigated by limited proteolysis30,31, native PAGE, gel filtration and accessibility of the Ppant thiol group31.

Owing to the lack of solution studies on catalytically competent NRPS constructs, particularly those containing an A domain, the structural dynamics of NRPS under conditions of catalysis remain largely unknown. Key questions about the directionality of the NRPS assembly-line process include how domain interactions are coordinated and how such interactions and conformational changes are coupled to the progression of the reaction (i.e., to the chemi-cal nature of substrates and intermediates bound to the enzyme). Ultimately, these relationships must drive the multistep peptide synthesis in a directional manner.

Here we report a FRET approach to studying conformational dynamics of NRPS in an A-PCP di-domain system. We observed changes in the FRET ratio that could be assigned to conformational changes within the A domain and between the A and the PCP domains in response to ligands and substrates. We followed for-ward and reverse progression of the domain alternation mechanism and revealed a conformational state of product inhibition. Our data suggest the presence of mixtures of conformations in two cases of substrate and ligand binding.

RESuLTSAn intramolecular FRET sensor of a di-domain model NRPSTo develop a FRET sensor of conformational changes between neigh-boring A and PCP domains, we chose the A-PCP di-domain derived from GrsA (wild-type (WT) A-PCP)31–33 (Fig. 1a). By removing four native cysteines31 we obtained A-PCP(Δ4Cys) (Supplementary Table 1). We fused EGFP C-terminally to the PCP domain and introduced a single cysteine (N152C). The resulting construct, here referred to as A-PCP-EGFP, was chemically conjugated with Alexa Fluor 546 (AF546) at N152C to give a doubly labeled protein, referred to as the apo sensor (Fig. 1 and Supplementary Figs. 1a and 2). N152C is in the AN subunit close to the predicted binding interface of the PCP domain14,20,21. With this design we expected closer proximity between the fluorophores in the transfer confor-mation of the A-PCP ensemble than in the A domain in the A or O conformations, which do not support productive interaction of the A and PCP domains. Computational modeling supported this design on the qualitative level by sampling possible dye and EGFP positions and provided a molecular picture for possible domain and fluorophore localizations (Supplementary Table 2, Supplementary Figs. 3–5 and Online Methods).

We converted the apo sensor into a holo sensor by incubation with the phosphopantetheine transferase Sfp and coenzyme A34 (Supplementary Fig. 6). Compared to WT A-PCP control constructs, the sensors showed ~19% and ~28% lower observed k (kobs) for the ATP-PPi exchange reaction (Table 1). A more pronounced effect, of ~17-fold rate reduction, was observed in the thioester formation reac-tion with L-[3H]Phe and ATP (Table 1). However, similar plateaus were reached in all cases (Supplementary Fig. 7). MS analysis showed complete formation of the phenylalanyl-thioester on the holo sensor with rates that were unaffected by the presence of the AF546 label (Supplementary Fig. 8). Furthermore, preincubation of the unlabeled apo and holo A-PCP-EGFP constructs with the high affinity inhibitor

5′-O-[N-(L-phenyl)-sulfamoyl] adenosine (Phe-AMS), a nonhydrolys-able analog of Phe-AMP (Supplementary Fig. 9), resulted in increased electrophoretic mobility in native PAGE, indicative of a closed confor-mation of the A domain31 (Fig. 2a). Together, these assays indicated that the sensor proteins remained enzymatically functional, as expected.

We then evaluated the holo sensor as a FRET reporter of confor-mational changes. Control experiments using fluorescence anisot-ropy measurements confirmed that the fluorophores were not limited in their movement (Supplementary Fig. 10), and chemical denatur-ation with urea confirmed proper protein folding was required for the FRET effect (Supplementary Fig. 11). We used the FRET ratio (Ia/Id) as a measure of relative energy transfer, normalized to the enzyme in the absence of ligands as a reference (ratio = 1; with Id measured at λ = 510 nm and Ia at λ = 570 nm) (Fig. 2b,c). To test our expected assignment of FRET readout to conformational state(s), we added Phe-AMS with the assumption that it would induce the pre-transfer conformation. The FRET ratio increased to 1.10 ± 0.02 (P < 0.005), indicative of a shorter inter-dye distance than in the protein without ligands and consistent with the expected recruitment of PCP to the A domain31. However, we were surprised by an even higher FRET ratio (1.29 ± 0.03; P < 0.005) when the substrates phenylalanine and ATP were added (Fig. 2b,c). In this case, we had expected a lower FRET ratio than that observed with Phe-AMS, because with the formed Phe-S-Ppant-PCP thioester, the PCP is primed for downstream reac-tions and should detach from the A domain. Thus, one of our ideas was incorrect: the biochemical model of aminoacylated PCP release from the active site or the assumption that the holo sensor reported a high FRET ratio on the transfer conformation.

1 431 524536

618 1,098 aa

C60F1 N152C C331A

C376SC473A C666S 866 aa

K434 K517 His6R439 S573

N152C N152

kDa150120100

706050

C UV C UV

2.0

1.6

1.2

0.8

0.4

0350 450 550 650

Wavelength (nm)

Apo A-PCP-EGFPApo sensor

* Abs

orpt

ion

(a.u

.)

AN AC PCP EGFP

a

AN AC PCP E

b c

Figure 1 | Design of an intramolecular FRET sensor of an A-PcP di-domain. (a) Domain structures of GrsA (top) and the A-PCP-EGFP constructs used as FRET sensors (bottom). Residues mutated in this study are indicated. (b) Representative maleimide labeling of the N152C mutant with AF546 to give the apo sensor and the N152 control construct. Shown are SDS–PAGE gels under ultraviolet illumination (right) and after Coomassie staining (left). Full gels are provided in Supplementary Figure 14. (c) Representative absorption spectra of the apo sensor before and after AF546 modification. Results represent at least three independent repeats.

© 2

017

Nat

ure

Am

eric

a, In

c., p

art

of

Sp

rin

ger

Nat

ure

. All

rig

hts

res

erve

d.

nature CHeMICaL BIOLOGY | VOL 13 | SEPTEMBER 2017 | www.nature.com/naturechemicalbiology 1011

articleNATuRE chEMIcAL bIOLOgy dOI: 10.1038/nCHeMBIO.2435

correlation of FRET changes with catalysisTo investigate the high FRET ratio induced by ATP and phenylala-nine, we first asked whether it correlated with enzymatic thioester formation. Upon addition of the substrates, the time courses for the FRET ratios of the holo sensor and of Phe-S-Ppant formation of the corresponding holo A-PCP(Δ4Cys), as measured by MS35, were virtually superimposable (Fig. 3a) and very similar to that of [3H]Phe binding in a radioactive assay (Supplementary Fig. 7). ATP and phenylalanine added individually only had a marginal or little effect on the FRET ratio (Fig. 3b). Thus, the conformational changes monitored by FRET correlated with the catalytic conversion of ATP and phenylalanine to Phe-S-Ppant. In a control experiment, we prepared a defined apo form of the FRET sensor by mutating the serine residue of the Ppant attachment site (referred to below as apo sensor(S573A)). This protein failed to produce high FRET ratio in the presence of ATP and phenylalanine, underlining the importance of the HS-Ppant group (Fig. 3b).

We also considered whether an elevated FRET ratio of the holo sensor reports only on the transfer conformation or if the closed A conformation of the A domain also contributes to the FRET effect. The formation of Phe-AMP can be reversed by adding excess PPi; this is the basis for the ATP–PPi exchange reaction to assay the adenylation reaction of A domains. We added PPi to the holo sen-sor in various ways (Fig. 3c); when added as the only ligand, PPi (10 mM) reduced the FRET ratio to 0.95 ± 0.01 (P < 0.005). When PPi was added at saturating concentrations (2 or 10 mM) as a mixture with ATP and phenylalanine, we observed no change in the FRET ratio (Fig. 3c). Under these conditions, no substantial amount of Phe-AMP should be present; thus the enzyme is exposed to ATP and phenylalanine, which trigger the A conformation. This finding supported our postulated assignment that only the transfer conformation with the A domain in T conformation contributes to a high FRET ratio. Notably, addition of PPi 20 min after incubation with ATP and phenylalanine, when the plateau of Phe-S-Ppant for-mation had already occurred, led to a FRET ratio modestly above that of the buffer control (1.06 ± 0.01; P < 0.005). We observed a rapid decrease from this plateau (Fig. 3d). By ESI-MS we could rule out that excess PPi also reversed the Phe-S-Ppant thioester to the uncharged HS-Ppant in the presence of saturating concentrations of ATP and phenylalanine (Supplementary Fig. 12). Therefore, the difference between FRET ratios observed at 0 min and 20 min after PPi addition was due to the presence of Phe-S-Ppant thioester in the latter case. Further, these results show that excess PPi not only shifts the reaction equilibrium of the adenylation reaction but also is cou-pled to conformational changes involving the interaction of A and

PCP domains. A concentration-dependent analysis revealed the PPi has a half-maximal effective concentration (EC50) of 320 ± 50 μM in the presence of ATP and phenylalanine (2 mM each; Fig. 3e).

These results suggested that the aminoacylated holo A-PCP di-domain accounted for the high FRET ratio. One possible expla-nation is that some or all of the enzyme population adopts a post-transfer conformation, corresponding to a product-inhibited state. To further test this hypothesis, we sought to measure the pre-transfer conformation, assuming that pre- and post-transfer conformations are very similar. We prepared a catalytically impaired sensor protein, with a Ppant analog lacking the terminal thiol group, here referred to as desulfo sensor. This enzyme is blocked for the conversion of Phe-AMP to Phe-S-Ppant-PCP and thus cannot progress beyond a pre-transfer state. In support of our hypothesis, the desulfo sen-sor’s FRET responses to ligands and substrates were similar to that of the holo sensor (Fig. 3b). We attributed the similarly high FRET ratios with ATP and phenylalanine (FRET ratio = 1.32 ± 0.04; P < 0.005 compared to buffer control), to the pre-transfer state of the A-PCP ensemble, with the A domain in T conformation and the PCP bound. A similar time course was also observed after addi-tion of ATP and phenylalanine and after the later addition of excess PPi (Fig. 3d), yet the plateau reached after PPi addition showed a lower FRET ratio than the buffer control, in marked contrast to the experiment with the holo sensor (P < 0.005). This finding is consis-tent with our conclusion that the plateau difference observed for the holo sensor (see above) stemmed from the stabilization effect of the Phe-S-Ppant intermediate on the post-transfer conformation. The idea that a fraction of the protein population is forced into the A

Table 1 | Enzymatic activities determined by ATP–PPi exchange and labeling of proteins with radioactive amino acids

Constructa

kobs (adenylation) min−1

kobs (thioester) min−1

WT A-PCP (apo) 140 ± 10WT A-PCP (holo) 96 ± 2 >4.90 ± 1.78A-PCP(Δ4Cys) (apo) 59 ± 1A-PCP(Δ4Cys) (holo) 16 ± 1 0.24 ± 0.02Unlabeled apo sensor 114 ± 2Unlabeled holo sensor 69 ± 2 0.29 ± 0.02Unlabeled apo sensor(S573A) 102 ± 3 NAUnlabeled holo sensor(K517A) 0.05 ± 0.02 NAUnlabeled holo sensor(K434A) 82 ± 4 0.11 ± 0.03Results are mean ± s.d. of 2 biological repeats, each consisting of 3 technical repeats. NA, no increase of radioactive signal detectable over 20 min.aAll assays were performed with proteins not conjugated with AF546.

Holo A-PCP-EGFP

Phe-AMS

1.4

1.3

1.2

1.1

1.0

0.9– +– +– +–

––ATP

PhePhe-AMS

500

400

300

200

100

0550 600500 650

Wavelength (nm)

Fluo

resc

ence

(rfu

)

Nor

m. F

RET

ratio

a

b

c

+ 2 mM ATP+Phe+ 10 µM Phe-AMS

+ Bu�er

Apo Holo– + – +

Figure 2 | The A-PcP FRET sensor undergoes ligand-dependent conformational changes. (a) Proteins were preincubated with or without Phe-AMS and then analyzed by native PAGE. Shown is a gel under UV illumination to detect EGFP fluorescence representing at least three independent experiments (full gel shown in Supplementary Fig. 14). (b) Fluorescence emission spectra of the holo sensor preincubated with the indicated ligands. (c) FRET ratios of experiments in b, calculated from the emission intensities (donor intensity at 510 nm/acceptor intensity at 570 nm) using excitation at 470 nm (cutoff 495 nm) and by normalization to buffer reference (ratio = 1). Results in b and c are mean ± s.d. of 3 independent measurements.

© 2

017

Nat

ure

Am

eric

a, In

c., p

art

of

Sp

rin

ger

Nat

ure

. All

rig

hts

res

erve

d.

1012 nature CHeMICaL BIOLOGY | VOL 13 | SEPTEMBER 2017 | www.nature.com/naturechemicalbiology

article NATuRE chEMIcAL bIOLOgy dOI: 10.1038/nCHeMBIO.2435

conformation of the A domain by excess PPi while another fraction is in the post-transfer conformation indicates a dynamic equilib-rium between these conformational states.

Importance of the nucleotide phosphate groupsAMP is the other product of the aminoacyl-transfer reaction. It is a competitive inhibitor of the adenylation reaction30. We found that AMP at saturating concentrations (2 mM) induced a FRET ratio of 1.40 ± 0.06 (P < 0.005) (Fig. 3a,b). We observed this strong effect only for the holo sensor and not for apo sensor(S573A) (1.08 ± 0.02;

P < 0.005), consistent with the inability of the latter to adopt a stable transfer conformation. AMP had an EC50 = 300 ± 50 μM for this conformational change (Fig. 3f). These results suggest that the sta-bilization of the post-transfer conformation has a role in the action of AMP as a competitive inhibitor.

ADP (2 mM) induced a FRET ratio of 1.23 ± 0.03 (P < 0.005) with the holo sensor (Fig. 3b). Although this value was lower than that seen for AMP, it indicates that the γ-phosphate of ATP is the main factor preventing adoption of the closed T conformation of the A domain. The predicted steric clash of the β-phosphate with R439 of the A8 motif (see above) appears to be malleable to a sub-stantial extent.

Phe-AMS induces a mixture of conformationsGiven our findings so far, the modest FRET ratio changes in the presence of Phe-AMS indicated that this synthetic ligand induced the transfer conformation to a substantially lower extent than Phe-AMP (pre-transfer conformation with the desulfo sensor) and Phe-AMP or Phe-S-Ppant (partial post-transfer conformation with the holo sensor). This finding was surprising, because Phe-AMS is an isostere of Phe-AMP, binds with high affinity36 and effectively induces a closed conformation31 (Fig. 2a).

We hypothesized that a partial adoption of the A conforma-tion as the alternative closed conformation might account for this effect. The imperfect mimic of the α-phosphate group by the sulfa-mate group in Phe-AMS, which lacks a negative charge, may have caused a less-defined control of the conformational switch by the A domain. To further test this hypothesis, we needed to monitor the population of the A conformation. This was not possible with our initial overall FRET sensor design. Therefore, we aimed for an indirect approach that explored destabilization of the A confor-mation by building on the previous proposal of similar energies of the electrostatically controlled A and T conformations and the similar positioning of the charges of the involved K517 and K434

1.0

0.8

0.6

0.4

0.2

0

0 2 4 6 8 10 12 14

1.5

1.4

1.3

1.2

1.1

1.0

0.91.3

1.2

1.1

1.0

0.91.4

1.3

1.2

1.1

1.0

0.9ATPPhePhe-AMSAMPCPPPPiAMPADP

Holo sensor

Apo sensor(S573A)

ndnd nd nd

ndndnd ndnd

Desulfo sensor

1.4

1.3

1.2

1.1

1.0

0.9ATPPhePPi

– + – +– + – +

– +– + +++

0 20 min

Holo sensorDesulfo sensor

ATP + Phe PPi1.4

1.3

1.2

1.1

1.0

0.9

1.3

1.2

1.1

1.010 100 1,000 10,000

t (min)0 10 20 30

1.4

1.3

1.2

1.1

1.01 10 100 1,000 10,000

AMP (µM)

PPi (µM)

Nor

m. F

RET

ratio

Nor

m. F

RET

ratio

Nor

m. F

RET

ratio

Nor

m. F

RET

ratio

Nor

m. F

RET

ratio

Nor

m. F

RET

ratio

Nor

m. F

RET

ratio

FRET (ATP+Phe)FRET (AMP)Thioester (ESI-TOF)

t (min)

****

**

*

* **

********

*

******

Rela

tive

sign

al

– + +++

++ +

+ + ++ +

++ +

++

+

++–

––––– –

––

––– – – –

–––– – – –

–– –

– –– –

–– – –

––––

–––––

–– –––

–––

––––––– –

––––

–– ––

––––

– – – –

–– –

a

b

c

d

e

f

–

** **

Figure 3 | correlation of FRET ratio with catalysis and ligand binding. (a) Overlay of normalized FRET ratio time courses of the holo sensor with the indicated substrates or ligands with a normalized time course of thioester formation using holo A-PCP(Δ4Cys) incubated with ATP and phenylalanine and assayed by ESI-TOF MS35 (data are mean ± s.d. from 3 independent repeats). (b) FRET ratios of the indicated proteins, determined after incubation with ligands for 20 min. The ratios are normalized to the buffer control for each protein. nd, not determined. (c) Effect of ligand addition on the holo sensor at the indicated time points. PPi was added at 10 mM. (d) Time-course experiments using the holo sensor and the desulfo sensor. ATP and phenylalanine (2 mM each) and PPi (10 mM) were added at the indicated time points. Data shown are from 3 independent measurements. (e) Dose-response curve of the holo sensor for PPi, added to the enzyme preincubated with ATP and phenylalanine (2 mM each) for 20 min. (f) Dose-response curve of the holo sensor for AMP, added to the enzyme and incubated for 20 min before determination of the FRET ratio. In b–f results show the mean of at least 3 biological repeats, each with 3 technical repeats. Error bars represent the combined s.d. using error propagation. *P < 0.05, **P < 0.005 (Student’s t-test).

1.3

1.2

1.1

1.0

0 2 4 6 8 10 12 14 16 18 20t (min)

Phe-AMS

1.4

1.3

1.2

1.1

1.0

0.9

1.4

1.3

1.2

1.1

1.0

0.9

Holo sensor(K517A)

Nor

m. F

RET

ratio

Nor

m. F

RET

ratio

ATPPhePhe-AMSAMPCPPAMP

Holo sensor(K434A)

a b

c

Nor

m. F

RET

ratio

Bu�erATP + PhePhe-AMS

–– –

–

– – – ––– –

– –– –

– –– – –

– –––––

––– – – – –

–+

+ ++

+ ++ +

+ ++

****

***

****

A-PCP(∆4Cys, K517A)

A-PCP(∆4Cys, K434A)

Apo Holo– + – +

Apo Holo– + – +

Figure 4 | A-PcP FRET sensors with selectively destabilized A or T conformations. (a) Coomassie-stained native PAGE analysis of A-PCP(Δ4Cys) constructs with the K517A and K434A mutations were preincubated with Phe-AMS (gels represent at least three independent experiments; full gels shown in Supplementary Fig. 14). (b) Ligand-dependent FRET ratios of the indicated proteins and ligands after 20 min incubation. Ratios are normalized to buffer controls. (c) Time-dependent FRET ratios of the holo sensor(K517A) after addition of ligands. Results in b and c are mean of at least 3 biological repeats, each with 3 technical repeats. Error bars represent the combined s.d. using error propagation. *P < 0.05, **P < 0.005 (Student’s t-test).

© 2

017

Nat

ure

Am

eric

a, In

c., p

art

of

Sp

rin

ger

Nat

ure

. All

rig

hts

res

erve

d.

nature CHeMICaL BIOLOGY | VOL 13 | SEPTEMBER 2017 | www.nature.com/naturechemicalbiology 1013

articleNATuRE chEMIcAL bIOLOgy dOI: 10.1038/nCHeMBIO.2435

side chains in the active site15 (Supplementary Fig. 1). We reasoned that a ligand directing the WT enzyme (measured with the holo sensor) into A conformation, by engaging an interaction with K517, would be impaired in this mechanism in context of a K517A mutant and instead would gain electrostatic binding energy by stabilizing the T conformation through interaction with K434. The FRET of the holo sensor(K517A) would thus report on the original T con-formation and the additional T conformation resulting from the disfavored A conformation. The latter contribution in response to a certain ligand should be represented by the relative increase of the FRET ratio with the holo sensor(K517A) within the total ligand profile of this reporter in comparison to the profile measured for the holo sensor.

A gel-shift assay of a separately prepared A(K517A)-PCP mutant incubated with Phe-AMS indicated that the mutant enzyme could still properly fold into a closed conformation (Fig. 4a). As expected, the ligand profile of the holo sensor(K517A) was markedly different from that of the holo sensor (Fig. 4b). Phe-AMS was one of the two cases with a relatively increased FRET ratio (1.30 ± 0.03 for the holo sensor(K517A); P < 0.005). On the basis of our design concept for the holo sensor(K517A), these findings supported our hypothesis that Phe-AMS binding at least partially induces the A conformation in WT A-PCP, in addition to the population of T conformation with the overall pre-transfer domain orientation. This case represented a second example of a population of mixtures of conformations in response to a defined ligand.

Lysine charge sensors control Ac domain alternationNotably, the addition of ATP and phenylalanine to the holo sensor(K517A) also produced an increase in FRET ratio (1.23 ± 0.05; P < 0.005; Fig. 4b), although the K517A mutation prevented the enzyme from forming Phe-AMP and the Phe-S-Ppant-PCP inter-mediates, as expected and verified by biochemical assays (Table 1 and Supplementary Fig. 13). We propose that the adoption of transfer conformation implied by the high FRET ratio can again be explained by the K517A mutation in this sensor. The A confor-mation would be induced in the unmutated enzyme by the sub-strates ATP and phenylalanine, but in the mutant the alternative gain in binding energy with the K434 residue in T conformation can at least partially override the steric clash between the β- and γ-phosphates and R439 of the A8 motif. Consistent with this idea, we observed that the time-dependent increase of the FRET ratio with ATP and phenylalanine was much slower than with Phe-AMS, indicative of a different mechanism to adopt the transfer conformation (Fig. 4c). In contrast, no overriding of the control switch was required with Phe-AMS (and the kinetics were very similar to that of the WT reporter holo sensor). The slower rate to adopt the conformation in the presence of ATP and phenylalanine can be interpreted as a consequence of internal friction from the two usually mutually exclusive mechanisms. This interpretation of a certain structural malleability is consistent with the partial over-riding of the R439–PPi control switch observed in the presence of ADP (Fig. 3b).

LowFRET

HSPCP

AC

LowFRET

HS

No ligand

O conformation A conformation

Phe + ATPPhe-AMP+ Ppant

T conformation(pre-transfer)

Phe-S-Ppantdischarge atC domain

HighFRET

HighFRET

–AMP

LowFRET

Phe

Repetitivereaction

cycle

O conformation

ATPPhe

(excess)LowFRETLow

FRETPhePhe

s

T conformation A conformation

Phe + ATPPhe-AMP–PPi

First reaction cycle

–PPi

AN

SH

AMP+ Phe-Ppant

T conformation(post-transfer)

No ligand

ExcessPPi

(added to in vitro system)

1 2

3

4

5

67

S573

HS

K517K434

AC

AN

Active site

AF546

EGFP

Ppant

ATP

Phe-AMP

AMP

Phe

HSPCP

s

in vitrosystem)

(addedto

ExcessPPi

s

ATPPhe

(excess)

Figure 5 | Model of the catalytic A-PcP reaction cycle and its coupling to conformational states. Reaction model deduced from experimental conditions with ATP and phenylalanine at saturating concentrations (2 mM each). In state (1), the enzyme is in the O conformation in the absence of substrates. Binding of ATP and phenylalanine leads to adoption of the A conformation (state (2)). After Phe-AMP formation and PPi release, rotation of the AC domain brings about the T conformation of the A domain and allows binding of the PCP to give the pre-transfer conformation (state (3)). State (4) shows the enzyme in the post-transfer conformation after thioester formation, which represents a product-inhibited conformation. The A domain is still in T conformation. To start the next adenylation reaction, the enzyme is assumed to go through an open state again (state (5)). As in the first reaction cycle, binding of the substrates induces the A conformation; however, the enzyme now carries a thioester bound phenylalanine (state (6)). Formation of Phe-AMP triggers the T conformation, but in contrast to state (3), the prosthetic group is charged and therefore in competition with the Phe-AMP intermediate for the active site (state (7)). We propose that under these conditions an equilibrium between states (4) and (7) is adopted. Gray boxes indicate conformational states and ligands present in the A domain active site.

© 2

017

Nat

ure

Am

eric

a, In

c., p

art

of

Sp

rin

ger

Nat

ure

. All

rig

hts

res

erve

d.

1014 nature CHeMICaL BIOLOGY | VOL 13 | SEPTEMBER 2017 | www.nature.com/naturechemicalbiology

article NATuRE chEMIcAL bIOLOgy dOI: 10.1038/nCHeMBIO.2435

A similar argument can be made for the experiment with AMP and phenylalanine as ligands, the second case in which the holo sensor(K517A) showed a higher relative FRET ratio (1.22 ± 0.01; P < 0.005) than the original WT-like holo sensor (1.14 ± 0.02; P < 0.005). This differential response underlined the additive con-tributions of the AMP 5′-phosphate group and the carboxylate moi-ety of phenylalanine. Whereas in the unmutated holo sensor the high FRET transfer conformation induced by AMP was suppressed by AMP and phenylalanine, this was not the case for the holo sensor(K517A). Therefore, we concluded that the presence of both ligands directed the unmutated enzyme into the A conformation. K517 appears to act as the preferred binding partner for this con-stellation of negative charges. By extension, a similar charge sensing in presence of ATP and phenylalanine would direct the unmutated A domain into the A conformation. This mechanism would ensure a directional sequence of conformational changes in the catalytic cycle of the A domain from an O to an A conformation and then, after catalysis, to a T conformation.

We also mutated K434 to study its importance for the confor-mational changes. The mutant enzyme could still adopt a compact conformation (Fig. 4a) and remained catalytically active at reduced rates (Table 1). The holo sensor(K434A) did not produce a high FRET ratio with any of the ligands tested (Fig. 4b). These observa-tions were consistent with the importance of K434 to progress the enzyme to the T and transfer conformations and our assignment of the high FRET states. Our model is summarized in Figure 5.

DIScuSSIONWe report the first study on conformational changes of an NRPS under catalytic conditions in solution with a novel FRET sensor concept consisting of a pair of A and PCP domains. By using vari-ous substrate and ligand combinations, computational modeling, as well as mutated and chemically modified enzyme variants as FRET sensors, we were able to assign the conformational state that yielded a high FRET signal to the transfer conformation, here observed as either pre-transfer or post-transfer conformation. In these con-formations the A domain is in T conformation and binds the PCP domain in a productive fashion (states (3) and (4) in Fig. 5).

Our data provides further and independent support for the domain alternation mechanism15,16 of the A domain to switch between the conformations required for the adenylation and thioes-terification half reactions (Fig. 5, from state (2) to (3) and from (6) to (7)). We could monitor in real time the reversal of the T confor-mation to the A conformation by excess PPi. The shift of the con-formational equilibrium closely mirrors the kinetics of the shifted chemical equilibrium of the aminoacylation reaction37 (Fig. 5, from state (4) to (6) and from (7) to (6)). These findings experimentally validate that PPi release is an essential step in the reaction pathway to switch from an A to a T conformation. It was previously pro-posed that the rotation of the AC subunit between the A and T con-formations is facilitated through electrostatic interactions that are almost equivalent energetically19. In agreement with this model, we observe selective destabilization of one conformation by mutat-ing the key lysine residue in the A8 or A10 motif (K434 or K517 in GrsA, respectively).

The reaction cycle of an A-PCP di-domain unit begins upon binding of the substrate amino acid and ATP. We propose that the positively charged side chain of the A10-K517 first interacts with the negatively charged substrate’s α-phosphate and the carboxylate of phenylalanine to clamp down AC from the O conformation to the A conformation (from state (1) to (2) in Fig. 5). Once Phe-AMP has formed and PPi has left the active site, the salt-bridge between R439 of the A8 motif and E327 favors the rotation into the T conformation (Fig. 5, state (3)). K434 contributes to the stabilization of this con-formation by electrostatic interaction with the intermediate product. Additional residues, such as R428 and D430, in the hinge between

the subdomains could be involved in AC rotation19. We propose that K517 and K434 are involved in the fine tuning of AC rotation in response to the ligand or substrate occupying the active site. Solution measurements with our FRET approach will be very useful for more detailed mechanistic investigations on this issue in the future. In the presence of Phe-AMS, both charge sensors K517 and K434 were confronted with altered electrostatic characteristics of the sulfamate group and therefore failed to adopt the same proportion of A and T conformations as with Phe-AMP. For this reason, AMS analogs appear to be imperfect tools to arrest NRPS enzymes in a defined conformation—for example, to facilitate protein crystallography.

Once in the T conformation, PCP can bind to give the pre- transfer conformation (Fig. 5, state (3)) and the Phe-S-Ppant thioester is formed (Fig. 5, state (4)). Notably, our data show that the covalently bound intermediate largely remains bound to the A domain, in competition with Phe-AMP formed in the active site in the next cycle of the A domain, giving rise to a dynamic equilibrium between conformational states (Fig. 5, states (4) and (7)). The post-transfer conformation corresponds to a state of product inhibition (Fig. 5, state (4)), which is further supported by the holo enzyme’s lower rate compared to the apo enzyme in the ATP–PPi exchange reaction (Table 1), as also reported earlier for a stably aminoacylated EntF NRPS38.

The phenomenon of product inhibition is relevant to under-standing the control of directionality in the overall assembly-line process, in which the covalently bound intermediates are required to be translocated to the next catalytic center(s) by the PCP domain. On the basis of our results, we propose a mechanism of product inhibition and release in which the aminoacyl-AMP formation in the next cycle39 contributes to the disengagement of aminoacyl-S-Ppant-PCP from the A domain through dynamic competition in the active site, thereby helping release it for downstream interac-tions (Fig. 5, from state (4) to (7)). The partial sequestering of the thioester might also be important to protect this reactive inter-mediate from undesired hydrolysis when exposed to bulk solvent. Another factor previously proposed40–42 to drive the assembly-line multistep synthesis is the affinity of the thioester intermediate to the next catalytic domain—for example, a C domain. It appears likely that both mechanisms are important for the directionality of NRPS-mediated biosynthesis. Neither a C domain to complete the catalytic cycle nor another downstream domain was present in our experimental setup (Fig. 5, from state (7) to (3)).

Our data show that the presence of bound substrates or inter-mediates correlates with the conformation of the enzyme. Notably, more than one conformation can be populated at a certain stage on the reaction coordinate of the enzyme, as shown for the com-petition between Phe-AMP and Phe-Ppant-PCP in the active site and the two closed conformations of the A domain in the presence of Phe-AMS. Given these observations and considering the general flexibility of the multidomain NRPSs, it is conceivable that even more catalytic states of the enzyme will be represented by a mixture of conformations that will undergo a population shift as the enzy-matic reaction progresses. Such mixtures of conformations will be difficult to delineate in detail by bulk studies like those done in this work and would call for single-molecule techniques in subsequent studies. The FRET methodology holds great promise to uncover further details of the dynamic assembly-line mechanism of NRPS in the future.

received 22 august 2016; accepted 14 June 2017; published online 24 July 2017

METhODSMethods, including statements of data availability and any associated accession codes and references, are available in the online version of the paper.

© 2

017

Nat

ure

Am

eric

a, In

c., p

art

of

Sp

rin

ger

Nat

ure

. All

rig

hts

res

erve

d.

nature CHeMICaL BIOLOGY | VOL 13 | SEPTEMBER 2017 | www.nature.com/naturechemicalbiology 1015

articleNATuRE chEMIcAL bIOLOgy dOI: 10.1038/nCHeMBIO.2435

references1. Arnison, P.G. et al. Ribosomally synthesized and post-translationally modified

peptide natural products: overview and recommendations for a universal nomenclature. Nat. Prod. Rep. 30, 108–160 (2013).

2. Marahiel, M.A., Stachelhaus, T. & Mootz, H.D. Modular peptide synthetases involved in nonribosomal peptide synthesis. Chem. Rev. 97, 2651–2674 (1997).

3. Hur, G.H., Vickery, C.R. & Burkart, M.D. Explorations of catalytic domains in non-ribosomal peptide synthetase enzymology. Nat. Prod. Rep. 29, 1074–1098 (2012).

4. Koglin, A. & Walsh, C.T. Structural insights into nonribosomal peptide enzymatic assembly lines. Nat. Prod. Rep. 26, 987–1000 (2009).

5. Mootz, H.D., Schwarzer, D. & Marahiel, M.A. Ways of assembling complex natural products on modular nonribosomal peptide synthetases. ChemBioChem 3, 490–504 (2002).

6. Weissman, K.J. The structural biology of biosynthetic megaenzymes. Nat. Chem. Biol. 11, 660–670 (2015).

7. Marahiel, M.A. A structural model for multimodular NRPS assembly lines. Nat. Prod. Rep. 33, 136–140 (2016).

8. Tanovic, A., Samel, S.A., Essen, L.O. & Marahiel, M.A. Crystal structure of the termination module of a nonribosomal peptide synthetase. Science 321, 659–663 (2008).

9. Drake, E.J. et al. Structures of two distinct conformations of holo-non-ribosomal peptide synthetases. Nature 529, 235–238 (2016).

10. Reimer, J.M., Aloise, M.N., Harrison, P.M. & Schmeing, T.M. Synthetic cycle of the initiation module of a formylating nonribosomal peptide synthetase. Nature 529, 239–242 (2016).

11. Kittilä, T., Mollo, A., Charkoudian, L.K. & Cryle, M.J. New structural data reveal the motion of carrier proteins in nonribosomal peptide synthesis. Angew. Chem. Int. Edn Engl. 55, 9834–9840 (2016).

12. Maier, T., Jenni, S. & Ban, N. Architecture of mammalian fatty acid synthase at 4.5 Å resolution. Science 311, 1258–1262 (2006).

13. Dutta, S. et al. Structure of a modular polyketide synthase. Nature 510, 512–517 (2014).

14. Conti, E., Stachelhaus, T., Marahiel, M.A. & Brick, P. Structural basis for the activation of phenylalanine in the non-ribosomal biosynthesis of gramicidin S. EMBO J. 16, 4174–4183 (1997).

15. Yonus, H. et al. Crystal structure of DltA. Implications for the reaction mechanism of non-ribosomal peptide synthetase adenylation domains. J. Biol. Chem. 283, 32484–32491 (2008).

16. Gulick, A.M. Conformational dynamics in the Acyl-CoA synthetases, adenylation domains of non-ribosomal peptide synthetases, and firefly luciferase. ACS Chem. Biol. 4, 811–827 (2009).

17. Reger, A.S., Wu, R., Dunaway-Mariano, D. & Gulick, A.M. Structural characterization of a 140 degrees domain movement in the two-step reaction catalyzed by 4-chlorobenzoate:CoA ligase. Biochemistry 47, 8016–8025 (2008).

18. Wu, R., Reger, A.S., Lu, X., Gulick, A.M. & Dunaway-Mariano, D. The mechanism of domain alternation in the acyl-adenylate forming ligase superfamily member 4-chlorobenzoate: coenzyme A ligase. Biochemistry 48, 4115–4125 (2009).

19. Branchini, B.R. et al. Bioluminescence is produced from a trapped firefly luciferase conformation predicted by the domain alternation mechanism. J. Am. Chem. Soc. 133, 11088–11091 (2011).

20. Mitchell, C.A., Shi, C., Aldrich, C.C. & Gulick, A.M. Structure of PA1221, a nonribosomal peptide synthetase containing adenylation and peptidyl carrier protein domains. Biochemistry 51, 3252–3263 (2012).

21. Sundlov, J.A., Shi, C., Wilson, D.J., Aldrich, C.C. & Gulick, A.M. Structural and functional investigation of the intermolecular interaction between NRPS adenylation and carrier protein domains. Chem. Biol. 19, 188–198 (2012).

22. Branchini, B.R., Murtiashaw, M.H., Magyar, R.A. & Anderson, S.M. The role of lysine 529, a conserved residue of the acyl-adenylate-forming enzyme superfamily, in firefly luciferase. Biochemistry 39, 5433–5440 (2000).

23. Horswill, A.R. & Escalante-Semerena, J.C. Characterization of the propionyl-CoA synthetase (PrpE) enzyme of Salmonella enterica: residue Lys592 is required for propionyl-AMP synthesis. Biochemistry 41, 2379–2387 (2002).

24. Branchini, B.R. et al. Mutagenesis evidence that the partial reactions of firefly bioluminescence are catalyzed by different conformations of the luciferase C-terminal domain. Biochemistry 44, 1385–1393 (2005).

25. Wu, R. et al. Mechanism of 4-chlorobenzoate:coenzyme a ligase catalysis. Biochemistry 47, 8026–8039 (2008).

26. Kochan, G., Pilka, E.S., von Delft, F., Oppermann, U. & Yue, W.W. Structural snapshots for the conformation-dependent catalysis by human medium-chain acyl-coenzyme A synthetase ACSM2A. J. Mol. Biol. 388, 997–1008 (2009).

27. Koglin, A. et al. Conformational switches modulate protein interactions in peptide antibiotic synthetases. Science 312, 273–276 (2006).

28. Frueh, D.P. et al. Dynamic thiolation-thioesterase structure of a non-ribosomal peptide synthetase. Nature 454, 903–906 (2008).

29. Goodrich, A.C., Harden, B.J. & Frueh, D.P. Solution structure of a nonribosomal peptide synthetase carrier protein loaded with its substrate reveals transient, well-defined contacts. J. Am. Chem. Soc. 137, 12100–12109 (2015).

30. Dieckmann, R., Pavela-Vrancic, M., von Döhren, H. & Kleinkauf, H. Probing the domain structure and ligand-induced conformational changes by limited proteolysis of tyrocidine synthetase 1. J. Mol. Biol. 288, 129–140 (1999).

31. Zettler, J. & Mootz, H.D. Biochemical evidence for conformational changes in the cross-talk between adenylation and peptidyl-carrier protein domains of nonribosomal peptide synthetases. FEBS J. 277, 1159–1171 (2010).

32. Hoyer, K.M., Mahlert, C. & Marahiel, M.A. The iterative gramicidin s thioesterase catalyzes peptide ligation and cyclization. Chem. Biol. 14, 13–22 (2007).

33. Stachelhaus, T. & Marahiel, M.A. Modular structure of peptide synthetases revealed by dissection of the multifunctional enzyme GrsA. J. Biol. Chem. 270, 6163–6169 (1995).

34. Lambalot, R.H. et al. A new enzyme superfamily—the phosphopantetheinyl transferases. Chem. Biol. 3, 923–936 (1996).

35. Sun, X., Li, H., Alfermann, J., Mootz, H.D. & Yang, H. Kinetics profiling of gramicidin S synthetase A, a member of nonribosomal peptide synthetases. Biochemistry 53, 7983–7989 (2014).

36. Finking, R. et al. Aminoacyl adenylate substrate analogues for the inhibition of adenylation domains of nonribosomal peptide synthetases. ChemBioChem 4, 903–906 (2003).

37. Luo, L. & Walsh, C.T. Kinetic analysis of three activated phenylalanyl intermediates generated by the initiation module PheATE of gramicidin S synthetase. Biochemistry 40, 5329–5337 (2001).

38. Liu, Y. & Bruner, S.D. Rational manipulation of carrier-domain geometry in nonribosomal peptide synthetases. ChemBioChem 8, 617–621 (2007).

39. Kittilä, T., Schoppet, M. & Cryle, M.J. Online pyrophosphate assay for analyzing adenylation domains of nonribosomal peptide synthetases. ChemBioChem 17, 576–584 (2016).

40. Linne, U. & Marahiel, M.A. Control of directionality in nonribosomal peptide synthesis: role of the condensation domain in preventing misinitiation and timing of epimerization. Biochemistry 39, 10439–10447 (2000).

41. Mootz, H.D. & Marahiel, M.A. Biosynthetic systems for nonribosomal peptide antibiotic assembly. Curr. Opin. Chem. Biol. 1, 543–551 (1997).

42. Belshaw, P.J., Walsh, C.T. & Stachelhaus, T. Aminoacyl-CoAs as probes of condensation domain selectivity in nonribosomal peptide synthesis. Science 284, 486–489 (1999).

acknowledgmentsWe are grateful to C.-B. Li and S. Kawai for fruitful discussions. We thank J. Diecker for technical support with radioactive assays and FRET experiments and W. Dörner for support with MS measurements. This work was funded by the Human Frontier Science Program (RGP0031/2010 to H.D.M., H.Y. and T.K.) and the National Science Foundation (graduate research fellowship DGE-0646086 to T.E.M.).

author contributionsJ.A., X.S., T.K., H.Y. and H.D.M. conceived the study. J.A., X.S., F.M., H.Y. and H.D.M. planned the experiments. J.A. developed the FRET sensor and carried out FRET, gel-shift and enzyme assays. X.S. performed MS experiments and photophysical controls of the FRET dye pair. F.M. performed FRET measurements with the desulfo sensor and the AF546 control experiments. E.D. and G.V. established and carried out enzyme assays. T.E.M. performed the computational modeling. J.A., X.S., F.M., H.Y. and H.D.M. interpreted the data. J.A., X.S. and H.D.M. wrote the manuscript.

Competing financial interestsThe authors declare no competing financial interests.

additional informationAny supplementary information, chemical compound information and source data are available in the online version of the paper. Reprints and permissions information is available online at http://www.nature.com/reprints/index.html. Publisher’s note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Correspondence and requests for materials should be addressed to H.D.M.

© 2

017

Nat

ure

Am

eric

a, In

c., p

art

of

Sp

rin

ger

Nat

ure

. All

rig

hts

res

erve

d.

nature CHeMICaL BIOLOGY doi:10.1038/nchembio.2435

ONLINE METhODSGeneral. β-D-1-thiogalactopyranoside (IPTG), ADP, ATP, HEPES and MgCl2 were purchased from AppliChem. 5′-O-[N-(L-phenyl)-sulfamoyl] adenosine (Phe-AMS) was purchased from Integrated DNA Technologies. TCEP was purchased from Fluka. Acetyl-coenzyme A, desulfo-coenzyme A, AMP and AMPCPP were purchased from Jena Bioscience. AF546 C5 maleimide was purchased from Life Technologies. PerfectPro Ni-NTA agarose was purchased from 5Prime GmbH. Oligonucleotides were purchased from Biolegio. L-[3,4,5-3H]phenylalanine was purchased from Hartmann Analytic. 32P-labeled PPi was purchased from PerkinElmer. Standard chemicals were purchased from AppliChem, Roth or Thermo Scientific unless otherwise stated. Experimental data were analyzed using Origin Pro 9.1 or MatLab.

Cloning of expression plasmids. To introduce the N152C substitution, plasmid pJZ13, encoding A-PCP(Δ4Cys) in a pET16b vector31, was mutated using prim-ers oJZ22 (5′-GTTCATTTAATTCATtgTATaCAATTTAATGGGCAAG-3′) and oJZ23 (5′-CTTGCCCATTAAATTGtATAcaATGAATTAAATGAAC-3′) to give plasmid pJZ23. The insert was amplified with primers oJA40 (5′-CCCTCTAGAAATAATTTTGTTTAACTTTAAGAAGGAGATATACCCGAGATGGTAAACAG-3′) and oJA41 (5′-GCTTAGTGATGGTGATGGTGATGccATggcCTCACTTC-3′), replacing a 5′ NcoI site with a BamHI site and a 3′ BglII site with a NcoI site. Plasmid pAB40 (pET28a backbone) encoding EGFP-His6 was digested with XbaI and NcoI and ligated with the PCR-amplified frag-ment digested with the same enzymes to give plasmid pJA50. The surface- exposed C666 of EGFP was removed by mutation to serine using primers oJA48 (5′-CTGAAGTTCATCTcCACCACCGGCAAGC-3′) and oJA49 (5′-GCT TGCCGGTGGTGgAGATGAACTTCAG-3′). The resulting pJA55 encodes pro-tein A-PCP-EGFP. pJA55 was used as a template to introduce mutation S573A using primers oJA59 (5′-GCATTAGGTGGAGATgcgATTAAAGCAATACAG-3′) and oJA60 (5′-CTGTATTGCTTTAATcgcATCTCCACCTAATGC-3′) to yield pJA82 encoding A-PCP(S573)-EGFP. pJA55 was used as the template to introduce mutation K517A using primers oJA01 (5′-CTTACATCAAATGGGgcgATcGATCGAAAGCAGTTG-3′) and oJA02 (5′-CAACTGCTTTCGATCgATcgcCCCATTTGATGTAAG-3′) to give pJA83 encoding A(K517A)-PCP-EGFP. pJA55 was used as the template to introduce mutation K434A in the encoded gene using primers oJA03(5′-GAATAGATAACCAGGTAgcGATTAGAGGTCACCGAG-3′) and oJA04 (5′-CTCGGTGACCTCTAATCgcTACCTGGTTATCTATTC-3′) to give pJA70 encoding A(K434A)-PCP-EGFP. Plasmid pJA81 encoding N152 control construct was prepared by inserting the EcoRI-NheI fragment from pJZ13 into pJA55.

Protein expression and purification. Proteins were expressed as described before31, with the exception that Escherichia coli Rosetta (DE3) was used as expression strain. Cells were grown to an OD600 of 0.6 in LB medium with additional 10 mM MgCl2 at 37 °C. After induction of expression with IPTG (0.4 mM) cells were grown at 16 °C for 16 h. Cells were harvested by centrifu-gation and lysed by two passages through an Avestin Emulsiflex C5 emulsifier. Proteins were purified via their C-terminal His6 tag as described31 and subse-quently dialyzed twice against assay buffer (50 mM HEPES, pH 7.0, 100 mM NaCl, 1 mM EDTA, 10 mM MgCl2) and once against assay buffer with 10% glycerol (v/v) before shock freezing in liquid nitrogen for storage.

Protein concentrations were determined by absorption measure-ments at 490 nm for EGFP fusion proteins using extinction coefficient ε = 55,000 M−1cm−1 and at 280 nm for proteins without EGFP. In these cases, the extinction coefficient at 280 nm was calculated using DNASTAR Lasergene Protean. Supplementary Table 1 shows the entire amino acid sequence of construct A-PCP-EGFP.

Chemical modification with AF546. C152 is the only surface-exposed cysteine side chain in all constructs based on A-PCP-EGFP and was modified by AF546. Proteins (4 μM) in assay buffer were incubated with 2 mM TCEP for 15 min at 18 °C. The AF546 stock solution in DMSO (10 mM) was diluted with 100 μL assay buffer and added to the protein solution to a final concentration of 40 μM. The labeling reaction was conducted at 18 °C for 30 min and quenched by the addi-tion of 2 mM DTT for 5 min. Proteins were applied to a Ni-NTA agarose column (200 μL bed volume), washed with 20 mL Ni-NTA buffer (Tris-HCl 100 mM,

NaCl 300 mM, pH 8.0), then 20 mL Ni-NTA buffer including 40 mM imidazole and eluted with Ni-NTA buffer including 250 mM imidazole. Fractions con-taining the protein of interest were identified by SDS–PAGE and pooled.

Post-translational conversion of apo into holo proteins. Apo constructs, typically at 1 μM, were incubated in assay buffer with 2 mM TCEP, 0.05 equivalents phos-phopantetheine transferase Sfp from Bacillus subtilis and 100 equivalents coen-zyme A or delsulfo coenzyme A. The reaction mixture was incubated at 4 °C for 7 h or overnight, followed by removal of excess CoA by dialysis against assay buffer.

ATP–PPi exchange and thioester formation assays. The ATP–PPi exchange assay to monitor the adenylation reaction was conducted as described35. Thioester formation was monitored with a tritium-labeled substrate amino acid. The respective enzyme (300 nM in assay buffer) was incubated with 2 mM ATP, a 1:175 mixture of L-[3,4,5-3H]Phe and phenylalanine (4 μM total concentration, ~0.23 μCi) in assay buffer, 5 mM TCEP and 10 mM MgCl2. L-phenylalanine was used throughout this study and is referred to as phenyla-lanine here. Reaction cups were coated with BSA overnight beforehand. At the respective time points, 100 μL of the reaction mixture was quenched by addi-tion of 0.8 mL TCA (10%) and 15 μL BSA solution (25 mg/mL). Samples were incubated on ice for 30 min, and the precipitate was collected by centrifugation in a microfuge (30 min, 13,000 r.p.m., 1 °C). The pellet washed twice with ice cold TCA (10%, 0.8 mL) and then solubilized in 150 μL formic acid (98%). For counting, 3.5 mL scintillation fluid (Aquasafe 500 Plus, Zinsser Analytic GmbH) was added and disintegrations per min (dpm) were counted over 10 min using a Beckman LS6500 Multi-Purpose Scintillation Counter.

Native PAGE assay. Native PAGE assays were conducted as described31. In brief, proteins were incubated with Phe-AMS (100 μM) for 15 min at 18 °C. Then 4× loading buffer (500 mM Tris-HCl, glycerol 40% (v/v), β-mercaptoethanol 20% (v/v), 5 mg/L bromophenol blue, pH 6.8) was added to the solution, and samples were subjected to native PAGE (running buffer: 25 mM Tris, 250 mM glycine). The gel apparatus was cooled on ice during electrophoresis at 80 V.

Steady state anisotropy measurements. Steady state anisotropy measurements were performed using a Fluorolog 3 with L-format detection (Horiba/Jobin Yvon). Excitation wavelength was 470 nm and emission was scanned from 500 nm to 600 nm with a 2-nm interval. Slit widths were set to 5 nm for both exci-tation and emission. The integration time was 0.1 s. The G-factor was determined during the measurements. ~2 nM apo or 6 nM holo A-PCP-EGFP labeled with AF546 was used in measurements (assay buffer at pH 7). For experiments with 1 mM ATP and 1 mM phenylalanine, apo or holo protein was mixed with both substrates at a final concentration of 1 mM each at room temperature (RT) for 15 min before the measurements. For acceptor anisotropy determination of the free AF546 dye (4 nM), the excitation was set to 540 nm and emission was scanned over 560 nm to 600 nm. Error bars represent 1 s.d. from mean for 6 measurements for the holo sensor with both substrates, and 3 measurements for the remaining conditions (Supplementary Fig. 10). The large anisotropy of donor when excited at the donor is consistent with the slow tumbling time of EGFP-tagged construct, consistent with data reported for a similar EGFP-fused construct43.

Fluorescence lifetime measurement. The EGFP fluorophore in apo A-PCP-EGFP was excited using a 450 nm Nano LED with a repetition rate of 500 kHz and sync delay of 50 ns in Fluorolog 3 equipped with a Time-Correlated Single Photon Counting (TCSPC) module. The emission was set to 510 nm. The coaxial decay was set to 85 ns, TAC range was 100 ns, RT preset was 60 s and the peak pre-set was 10,000 counts. The instrument response function (IRF) was determined using 92-nm diameter polymer microspheres (Duke Scientific Corporation). The fluorescence lifetime of apo A-PCP-EGFP was determined to be 2.8 ± 0.1 ns using a single exponential model. This is similar to the reported value44 (~3 ns). Taken together, these results indicated that the fused EGFP in our construct retained its photophysical properties. Similarly, we also determined the fluores-cence lifetime of AF546 attached to C152 to be 3.8 ± 0.1 ns, which is close to the manufacturer’s reported value of free dye (4.1 ns) and by our own measurement (4.0 ± 0.1 ns). Three repeats were performed at 25 °C in the assay buffer at pH 7.0, and the fluorescence lifetime data were reported as mean ± s.d.

© 2

017

Nat

ure

Am

eric

a, In

c., p

art

of

Sp

rin

ger

Nat

ure

. All

rig

hts

res

erve

d.

nature CHeMICaL BIOLOGYdoi:10.1038/nchembio.2435

Förster radius (R0) measurement. The following equation was used to calculate R0 (ref. 45):

R k n Q J02 4 1 60 211= −. [ ( )] /

D l

where k2 is the orientation factor, n is the refractive index of the solution (1.333), QD is the quantum yield of EGFP in the context of our A-PCP-EGFP sensor and J(λ) is the overlap integral, defined as

J F dA( ) ( ) ( )l l e l l l=∞

∫ D0

4

where FD(λ) is the normalized EGFP emission spectrum, εA(λ) is the extinction coefficient of AF546 at λ in the unit of M−1cm−1 and λ is measured in nanom-eters. Measurements were performed for unlabeled A-PCP-EGFP in the assay buffer, and the quantum yield of EGFP in A-PCP-EGFP was measured as 0.69 ± 0.13, using rhodamine 6G as the standard46. To measure FD(λ), A-PCP-EGFP was excited at 470 nm, and emission from 500 nm to 700 nm was scanned. Invitrogen’s coefficient of free AF546 (112,000 M−1cm−1 at pH 7 at 554 nm) was used to calibrate its extinction coefficient in the assay buffer. The R0 was determined as 57 ± 2 Å assuming the mean k2 as 2/3. Replacing the free AF546 with AF546–labeled A-PCP did not change the R0 result.

FRET measurements. Fluorescence measurements were conducted in a SpectraMax M5 Multi-Mode MicroplateReader (Molecular Devices) in either a quartz cuvette (Hellma) or black 96-well half-area microplates (medium binding, Greiner Bio-One). The reader was set to an excitation wavelength of 470 nm (495-nm cutoff) for the EGFP donor, and the detection wavelength was set to 510 nm for EGFP and 570 nm for AF546. Direct excitation of AF546 was done at 540 nm, and detection at 570 nm. Proteins were used at a concentration of 300 nM in assay buffer (pH 7.0) and a total volume of 50 μL. Endpoint meas-urements under equilibrium conditions were performed in the 96-well plates. Proteins were mixed with ligands or substrates and preincubated at 25 °C for 20 min in the dark to ensure complete binding. For time-dependent measure-ments, the protein solution was preincubated with buffer in a cuvette for 3 min before ligands or substrates were added. Fluorescence readings were taken every minute (for measurements with AMP as well as with ATP and phenylalanine and PPi added at different time points) or every 11 s (for measurements with ATP/phenylalanine and with Phe-AMS). Fluorescence data were corrected for background and direct excitation of the acceptor fluorophore using a reference emission spectrum of AF546 (300 nM) after excitation at 540 nm47. A sample of unlabeled A-PCP-EGFP was used to determine the donor bleedthrough as a small fraction of Id (8.16%), according to the published protocol48. The effect of donor bleedthrough on our qualitative explanation of the FRET ratio (Ia/Id) was found to be negligible and was not corrected for. Ia/Id of each measurement in an experiment was calculated47,49 and normalized using the FRET ratio of the buffer control. FRET experiments were conducted as triplicates (technical repeats) in at least 3 independent experiments (biological repeats). The mean and s.d. of each triplicate was calculated. Error propagation was performed as

snn

snn

x xj

j

kj

j

j

k2

1

2

1

2= + −= =∑ ∑ ( )j

where x j is the mean and s j2 is the variance of k sets of data consisting of nj

values for x and s2 is the variance of the new mean x of all n values. A two-tailed Student’s t-test was used to evaluate results, considering results to be significantly different when P < 0.05. Significance is indicated as *P < 0.05 and **P < 0.005

Thioester reversal measurement. 2 μM WT holo A-PCP construct was incu-bated with 1 mM ATP and phenylalanine at RT for 30 s in assay buffer at pH 7.0. PPi was then added the reaction mixture at a final concentration of 10 mM and incubated for another 10 min before the desalting step and subsequent ESI-MS measurements as previously described35. The thioester formation at Ppant was complete at 30 s and was stable under the reaction condition for at least 30 min2.

MS assay for thioester formation assay of FRET sensor constructs. The for-mation of the phenylalanine thioester by the unlabeled (holo A-PCP-EGFP) and labeled enzyme (holo sensor) was followed by ESI-TOF-MS. For the reac-tion the protein was incubated with 2 mM ATP, 2 mM phenylalanine and 10 mM MgCl2 at 25 °C. At different time points 18 μL of the reaction mixture was quenched by adding 2 μL of 10% formic acid. For the 0-min sample, the protein was mixed directly with 10% formic acid solution and afterwards ATP, phenylalanine and MgCl2 were added. All the samples were then centrifuged (14,500 r.p.m., 2 min, room temperature) and directly measured to prohibit precipitation. To quantify the rate of thioester formation, we calculated the relative loading of the substrate as a fraction of the total amount of protein in the sample. According to Mann et al.50, the deconvoluted height of multi-ply charged ions is directly related to their abundance. We therefore used the area under the curve of the respective protein signals in the deconvoluted mass spectrum (using the software Data Analysis 4.4 by Bruker and the Maximum Entropy algorithm)51, assuming that the ionization efficiencies of the loaded and unloaded species are comparable. The relative abundance of the loaded species was plotted against time, and a single exponential function was fitted to the data under the assumption of a pseudo-first-order reaction to yield the rate constant kobs.

Simulation of protein conformations GrsA protein models. Computational modeling was used to determine what types of distance changes could occur when the A-PCP protein transitioned from conformations related to adenyla-tion and thioester formation. The modeling also provided a structural under-pinning for the various conformations discussed in this work. The modeling must take into account the changing adenylation domain conformations, inte-grate the PCP domain and append and sample possible conformations for both the dye and EGFP. The only known GrsA adenylation domain crystal struc-ture, which is in the adenylation conformation, was used for the first model (PDB 1AMU)14. All atoms not resolved in the crystal structure, excluding terminal residues 1–16 and 530–563, were built using the CHARMM c36a2 internal coordinate facility and CHARMM 36 parameters52. The dye, AF546, was built in Avogadro and minimized using the UFF force field53. Initial dye parameters were determined by CGenFF 0.9.7 using the CGenFF force field 2b8 (refs. 54–56). Appropriate mutations were made to generate the cysteine-free construct and the missing loop (residues 192–196) was reconstructed via minimization in CHARMM. The minimization approach employed a combi-nation of 500 steps of steepest decent (SD) minimization and 5,000 steps of adopted-basis Newton–Raphson (ABNR) minimization with a tolerance of 0.01. Minimizations were conducted first with all atoms excluding the missing loop fixed, then with all backbone heavy atoms fixed, then with all atoms free to move. The GBMV2 implicit solvent model was used with the recommended electrostatics57. The protonation state of titratable residues was set to match the experimental pH based on pKa predictions from PROPKA 3.0 (ref. 58). The PCP domain structure was determined by homology modeling with the Phyre2 server and the full A-PCP sequence59. This selected model was based on the complete structure of surfactin A synthetase C (SrfA-C) (PDB 2VSQ)8. The PCP domain was added to the A domain by aligning the C-terminal regions of the two models (residues 431–530) followed by grafting the PCP domain to the A domain structures.

Simulation of nonadenylation conformation structures. Homology modeling with the Phyre2 server was used to determine the structure of GrsA in alterna-tive conformations. The complete SrfA-C model provides a good match for the O conformation. For the T conformation, the recent LgrA structure pro-vides a highly homologous structure (42% identity) that includes an attached PCP domain (PDB 5ES8). Alternative less homologous structures include PA1221 (21% identity) (PDB 4DG9)20, a natural PCP-A domain construct from Pseudomonas aeruginosa, as well as EntE-B (26% identity) (PDB 3RG2)21, which joins an adenylation and PCP domain that are part of E. coli enterobactin synthesis system. Additional models can be based on isolated T conformation adenylation domains, such as D-alanine-D-alanyl carrier protein ligase (36% identity) (DltA - PDB 3E7W)15. In order to generate the most faithful model of GrsA, the AN domain of the DltA template was aligned to the GrsA model, and the T conformation AC domain was grafted to the GrsA AN domain starting

© 2

017

Nat

ure

Am

eric

a, In

c., p

art

of

Sp

rin

ger

Nat

ure

. All

rig

hts

res

erve

d.

nature CHeMICaL BIOLOGY doi:10.1038/nchembio.2435

is still substantially different from the T conformation model. A-PCP linker sampling for the O conformation resulted in a distance distribution that is very similar to the distribution observed for the A conformation A-PCP linker sam-pling. While the nonsampled model is more faithful to the crystallographically observed PCP domain orientation, both models support the conclusion that the A and O conformations cannot be distinguished by this experimental con-struct. In the main text we present the most reasonable models, including the A-PCP linker sampling for both the A (PDB 1AMU) and the O conformations (PDB 2VSQ), and T conformation based on LgrA (5ES8).

Statistical methods. All of the data are shown as mean values with error bars representing ± s.d. When stated, P values were calculated using two-tailed Student’s t-test. Data were evaluated to be significantly different with P < 0.05. Significance is indicated as *P < 0.05 and **P < 0.005.

Code availability. The code for the simulation of protein conformations and the sampling of the fluorophores will be made available through GitHub (https://github.com/PrincetonUniversity/GrsA_bulk_FRET).

Data availability. Data generated or analyzed during this study are included in this published article (and its supplementary information files) or are available from the corresponding author upon reasonable request.

at residue 429. Next, the linker (residues 527 to 531) was energy minimized, followed by non-backbone atoms in the C-terminal region and the overall C-terminal region. The PCP domain structure used in the A conformation model was grafted following alignment of the C-terminal regions of the two models (residues 431 to 530) and a ~20-degree swivel around the peptide bond between residues 524 and 525 needed to avoid steric clashes. To generate a rea-sonable structure of the A-PCP linker, the junction between the two segments (residues 524 to 527) was minimized, followed by the entire PCP domain.

Conformation sampling and distance determination. EGFP was appended to the end of the PCP domain by overlapping residue 613 from the minimized crystal structure (PDB 4EUL)60. EGFP was appended at residue 611 in the T conformation PA1221 and EntE-B structures because of the presence of flex-ible terminal residues. Possible configurations of the AF546 were determined by geometric sampling of six TIP3P carbons that connect the aromatic portion of the dye to the protein, as defined by either the α− and β-carbon bond, the β-carbon and sulfur bond, or the central carbon–carbon bonds. Three rotam-ers were sampled for each bond, for a total of 729 possible conformations. To determine the feasibility of a given dye conformation, the dye was energy minimized in vacuum for 50 s.d. steps and for 200 s.d. steps with a tolerance of 0.05 and the GBMV2 implicit solvent model. This energy was compared to the initial energy of the dye, and conformations with equal or lower energy were saved. Possible configurations of the PCP-EGFP linker (residues 613–621 or 611–621 for PA1221 or EntE-B, respectively) were determined by a 5-ns Langevin dynamics simulation with the GBMV2 implicit solvent model, and linker structures every 5 ps were grafted to the PCP domain. EGFP was then added to the end of the linker, and structures within 1,000 kcal/mol of the initial conformation were saved after 100 s.d. steps of energy minimization in vacuum. The EGFP structure that had the center of mass of the fluorophore residues (683–685) closest to the average of all structures was selected. The PCP domain location for the A conformation is not defined by crystal struc-ture, so sampling was also conducted for possible locations of the PCP domain. The linker was defined as residues 527–541, and a 15-ns Langevin dynamics simulation was used to generate conformations of the linker. Each conforma-tion was then grafted to the end of the A domain and the PCP-EGFP average structure. As the PCP domain orientation in the O conformation is not func-tionally restricted, the A-PCP linker sampling procedure was also applied to this structure. The mean distance and s.d. was determined by calculating all distances between the accepted conformations of dye and EGFP.

Comparison of sampled structures. The mean distances of all sampled structures are presented in Supplementary Table 2, and their distributions are presented in Supplementary Figure 4. A visual comparison of the mean structures is shown in Supplementary Figure 5. The A and the O conforma-tion structures have large mean distances that are not distinguishable with the current experimental design. However, the highest homology T conformation structure from LgrA (PDB 5ES8) exhibits a smaller mean distance. The T con-formation model based on the DltA adenylation domain (PDB 3E7W) uses the same PCP domain structure as in the A conformation. In this model the PCP domain does not fully bind to the A domain, and this orientation results in a shorter dye–EGFP distance distribution. The T conformation model based on PA1221 (PDB 4DG9) has a PCP domain that is slightly less bound to the A domain, and the distance distribution for this structure is slightly larger. The T conformation model based on EntE-B (PDB 3RG2) has a highly extended PCP domain because this structure is a dimer with PCP bound to the neighboring chain. While this structure produces a large dye–EGFP distance distribution, it is probably not representative of the monomeric GrsA construct. The LgrA model has the highest homology for GrsA and shows clear binding between the PCP and adenylation domains, so this model is the most reasonable repre-sentation for GrsA in the T conformation. The adenylation conformation crys-tal structure is limited because it does not include the PCP domain. Because the PCP domain is not functionally required in the adenylation conformation, sampling of the A-PCP linker (residues 527–541) was used to model the worst-case scenario where the PCP domain conformation is not restricted. While this results in a shorter dye–EGFP distance distribution, the mean distance

43. Albertazzi, L., Arosio, D., Marchetti, L., Ricci, F. & Beltram, F. Quantitative FRET analysis with the EGFP-mCherry fluorescent protein pair. Photochem. Photobiol. 85, 287–297 (2009).

44. Volkmer, A., Subramaniam, V., Birch, D.J. & Jovin, T.M. One- and two-photon excited fluorescence lifetimes and anisotropy decays of green fluorescent proteins. Biophys. J. 78, 1589–1598 (2000).

45. Lakowicz, J.R. Principles of Fluorescence Spectroscopy (Kluwer Academic/Plenum Publishers, 1999).

46. Magde, D., Wong, R. & Seybold, P.G. Fluorescence quantum yields and their relation to lifetimes of rhodamine 6G and fluorescein in nine solvents: improved absolute standards for quantum yields. Photochem. Photobiol. 75, 327–334 (2002).

47. Kraynov, V.S. et al. Localized Rac activation dynamics visualized in living cells. Science 290, 333–337 (2000).

48. Clegg, R.M. Fluorescence resonance energy transfer and nucleic acids. Methods Enzymol. 211, 353–388 (1992).

49. Miyawaki, A. & Tsien, R.Y. Monitoring protein conformations and interactions by fluorescence resonance energy transfer between mutants of green fluorescent protein. Methods Enzymol. 327, 472–500 (2000).

50. Mann, M., Meng, C.K. & Fenn, J.B. Interpreting mass-spectra of multiply charged ions. Anal. Chem. 61, 1702–1708 (1989).

51. Ferrige, A.G., Seddon, M.J., Jarvis, S., Skilling, J. & Aplin, R. Maximum entropy deconvolution in electrospray mass spectrometry. Rapid Commun. Mass Spectrom. 5, 374–377 (1991).

52. Brooks, B.R. et al. CHARMM: the biomolecular simulation program. J. Comput. Chem. 30, 1545–1614 (2009).

53. Hanwell, M.D. et al. Avogadro: an advanced semantic chemical editor, visualization, and analysis platform. J. Cheminform. 4, 17 (2012).

54. Vanommeslaeghe, K. & MacKerell, A.D. Jr. Automation of the CHARMM General Force Field (CGenFF) I: bond perception and atom typing. J. Chem. Inf. Model. 52, 3144–3154 (2012).

55. Vanommeslaeghe, K., Raman, E.P. & MacKerell, A.D. Jr. Automation of the CHARMM General Force Field (CGenFF) II: assignment of bonded parameters and partial atomic charges. J. Chem. Inf. Model. 52, 3155–3168 (2012).

56. Vanommeslaeghe, K. et al. CHARMM general force field: A force field for drug-like molecules compatible with the CHARMM all-atom additive biological force fields. J. Comput. Chem. 31, 671–690 (2010).

57. Lee, M.S., Feig, M., Salsbury, F.R. Jr. & Brooks, C.L. III. New analytic approximation to the standard molecular volume definition and its application to generalized Born calculations. J. Comput. Chem. 24, 1348–1356 (2003).

58. Olsson, M.H., Søndergaard, C.R., Rostkowski, M. & Jensen, J.H. PROPKA3: consistent treatment of internal and surface residues in empirical pKa predictions. J. Chem. Theory Comput. 7, 525–537 (2011).

59. Kelley, L.A., Mezulis, S., Yates, C.M., Wass, M.N. & Sternberg, M.J. The Phyre2 web portal for protein modeling, prediction and analysis. Nat. Protoc. 10, 845–858 (2015).

60. Arpino, J.A., Rizkallah, P.J. & Jones, D.D. Crystal structure of enhanced green fluorescent protein to 1.35 Å resolution reveals alternative conformations for Glu222. PLoS One 7, e47132 (2012).

© 2

017

Nat

ure

Am

eric

a, In

c., p

art

of

Sp

rin

ger

Nat

ure

. All

rig

hts

res

erve

d.