Structural and dynamic studies of the peptidase domain from Clostridium thermocellum PCAT1

Abstract

The export of antimicrobial peptides is mediated by diverse mechanisms in bacterial quorum sensing pathways. One such binary system employed by gram‐positive bacteria is the PCAT1 ABC transporter coupled to a cysteine protease. The focus of this study is the N‐terminal C39 peptidase (PEP) domain from Clostridium thermocellum PCAT1 that processes its natural substrate CtA by cleaving a conserved ‐GG‐ motif to separate the cargo from the leader peptide prior to secretion. In this study, we are primarily interested in elucidating the dynamic and structural determinants of CtA binding and how it is coupled to cleavage efficiency in the PCAT1 PEP domain. To this end, we have characterized CtA interactions with PEP domain and PCAT1 transporter in detergent micelles using solution nuclear magnetic resonance spectroscopy. The bound CtA structure revealed the disordered C‐terminal cargo peptide is linked by a sterically hindered cleavage site to a helix docked within a hydrophobic cavity in the PEP domain. The wide range of internal motions detected by amide nitrogen (N¹⁵) relaxation measurements in the free enzyme and substrate‐bound complex suggests the binding site is relatively floppy. This flexibility plays a key role in the structural rearrangement necessary to relax steric inhibition in the bound substrate. In conjunction with previously reported PCAT1 structures, we offer fresh insight into the ATP‐mediated association between PEP and transmembrane domains as a putative mechanism to optimize peptide cleavage by regulating the width and flexibility of the enzyme active site.

Short abstract

PDB Code(s): 7N87 and 7S5J;

Abbreviations

ABC

ATP‐binding cassette transporters

CSP

chemical shift perturbation

HMQC

heteronuclear multiple‐quantum coherence

HSQC

heteronuclear single‐quantum coherence

ILV

isoleucine–leucine–valine

EMF

Extended ModelFree

NB

nucleotide binding

NOE

nuclear Overhauser effect

NMR

nuclear magnetic resonance

OD

optical density

PCAT

peptidase‐containing ATP‐binding cassette transporters

PEP

peptidase

TM

transmembrane

TROSY

transverse relaxation optimized spectroscopy

1. INTRODUCTION

A diverse array of quorum sensing circuits is responsible for bacterial cell‐to‐cell communication by regulating gene expression in response to fluctuations in cell‐population density. ¹ The ability to self‐monitor bacterial population is vital for coordinating a myriad of group activities such as symbiosis, virulence, competence, and biofilm formation. In general, gram‐negative bacteria rely upon the acylated homoserine lactones as autoinducers for LuxR‐LuxI family of quorum sensing transcriptional regulators. Gram‐positive bacteria secrete cleaved peptides by peptidase‐containing ABC transporters (PCATs) that harness the energy of ATP hydrolysis to translocate substrates across the bilayer. Typically, these dual‐functional ATP‐binding cassette transporters (ABC) transporters share similar multidomain architecture with a cytoplasmic protease domain (PEP), α‐helical transmembrane domain (TMD), and a nucleotide‐binding domain (NBD).

The X‐ray structure of dimeric PCAT1 from Clostridium thermocellum revealed the peptide maturation and translocation through the α‐helical barrel utilized the alternate‐access mechanism common to many ABC transporters. ² In this mechanism, the transporter alternates between “inward‐ and outward”‐facing states regulated by ATP‐dependent conformational changes in the inter‐subunit interactions. ³ The N‐terminal peptidase (PEP) domain from PCAT1 recruits and processes a specific substrate CtA, which consists of a 24‐residue leader sequence with a highly conserved double‐glycine motif followed by a 66‐residue cargo peptide. ⁴ The PEP active site configuration is homologous to the C39 family of cysteine proteases, with a nucleophilic cysteine thiol (Cys21) forming a catalytic triad with His99 and Asp115. ⁵ As summarized in Scheme 1, the reaction cycle is initiated by the conformational switch between the “inward‐ and outward”‐facing configurations of the transporter triggered by ATP binding and hydrolysis coupled to proteolytic cleavage of the substrate. ² ATP binding releases the PEP domain and simultaneously reorients the TMDs to stabilize the “outward‐facing” configuration that facilitates the release of the sequestered cargo peptide. This reaction cycle is reset by ATP hydrolysis that causes the transporter to revert to an inward‐facing conformation. The PCAT1 transporter structure in proteoliposomes has been solved in three different states to confirm the key steps of the proposed mechanism: (i) wild‐type protein in the “inward”‐facing configuration free of ATP and substrate (X‐ray, 4RY2), ² (ii) an intermediate configuration (E648Q mutant) captured by ATPγS binding in NBD, with closed translocation channel and flexibly linked PEP domains (X‐ray, 4S0F), ² (iii) mutant of PCAT1 (C21A) with inactive PEP domain bound to substrate in the “inward”‐facing configuration (CryoEM, 6V9Z). ⁶

SCHEME 1.

(a) Displayed a schematic representation of the alternate access mechanism of PCAT1 activity triggered by ATP binding. The reaction cycle starts with “inward”‐facing configuration of the TMDs in the ATP‐free state, with the substrate CtA bound to the N‐terminal PEP domain. ATP binding reorients the TMDs to stabilize the “outward” configuration that triggers the release of the cargo peptide. (b) Domain structure of PCAT1 consisting of N‐terminal peptidase (PEP) domain, transmembrane domain (TMD), and C‐terminal nucleotide binding domain (NBD); full‐length CtA with N‐terminal leader sequence before the double‐glycine cleavage site. The mutation C21A in the catalytic triad abrogates proteolytic activity in both PCAT1 and the isolated PEP domain. The peptide CtA‐37 used for the NMR studies corresponds to the optimal PEP binding site spanning residues 5–37

The central question in the transport mechanism of various PCATs is what factors determine the specificity of the leader sequence toward the PEP domain and the nature of interactions between the cargo peptide and the TM domain. The substrate recognition mechanism of isolated and active PEP domains has been widely studied in various organisms. ⁷ , ⁸ , ⁹ In all these structures, the leader sequence forms a short amphipathic helix that docks within a hydrophobic groove adjacent to the active site of the PEP domain. The lack of proteolytic activity of excised C39 domains in the absence of TMDs in PCAT1 ² and NukT ¹⁰ is poorly understood.

Until recently, the structural determinants of the cargo peptide enclosed by the TM conduit had proven to be elusive owing to the dynamic association, a natural prerequisite for rapid passage through the TM channel. This lack of atomic‐level insight has been remedied by recent studies that have focused on both the structure and binding affinity of the C‐terminal cargo peptide. The very first Cryo‐EM images of dimeric PCAT1^C21A complexed with nonproteolyzed CtA protein ⁶ revealed important details such as an asymmetric binding mode. The two subunits of PCAT1, instead of exporting two peptides simultaneously, primed only one for export while the second molecule remained tethered to the PEP domain in the cytosol. An independent study using electron paramagnetic resonance (EPR), and fluorescence spectroscopy ¹¹ concluded that the cargo peptide contributes as much to the substrate binding affinity as the leader sequence despite the diffuse structure in the alpha‐helical TM barrel.

The conformational switch in ABC transporters observed by different structural biology techniques ³ , ⁷ , ¹² validates the recurrent paradigm of modular allostery in multidomain proteins. ¹³ There is a growing wealth of experimental evidence from nuclear magnetic resonance (NMR), ¹⁴ neutron spin‐echo, ¹⁵ and fluorescence resonance energy transfer to show long‐range dynamic allostery is coupled to local dynamics in super‐tertiary structures. ¹⁶ , ¹⁷ The intrinsic flexibility of proteins is an indisputable fact with the three‐dimensional structures best viewed as a dynamic ensemble of interconverting states instead of static entity. ¹³ , ¹⁸ Structural malleability plays a fundamental role in the manifold of biological functions executed by proteins. Among the various techniques available for studying protein motions, site‐specific nuclear spin relaxation measurements by NMR provide a powerful technique to map dynamic hot spots over a broad range of timescales and amplitudes.

In this study, we are primarily interested in elucidating the dynamic and structural determinants of substrate recognition coupled with cleavage efficiency in the PEP domain from PCAT1. A highly relevant question in the reaction cycle of PCAT1 (Scheme 1) is the mechanism by which the substrate is prevented from proteolysis in the pre‐translocation state. Previous studies have demonstrated the cleavage efficiency of PEP domain is optimal when associated with TMD in the ATP‐free state but significantly inhibited (~80%) when isolated. ² The significance of the interdomain interactions on PEP activity was confirmed by trans‐complementation assay in which isolated PEP was added in trans to a construct containing only the TMD and NBD resulting in enhanced activity. However, the root cause of the drastic reduction in protease activity is unclear from the existing structures of intact PCAT1 where PEP is always associated with the TM domain. The lack of insight on the nature of interactions between the substrate and isolated enzyme provides an impetus to investigate the complex in solution by NMR.

To advance this goal, the 90‐residue substrate (9.9 kDa, 90 AA) has been successfully reconstituted with the isolated PEP^C21A domain (16.4 kDa, 148 AA) and separately with dimeric PCAT1^C21A in detergent micelles. The ability to generate ²H/¹³C/¹⁵N‐labeled samples with protonated isoleucine–leucine–valine (ILV) methyl groups of the substrate by recombinant methods facilitated transverse relaxation optimized spectroscopy (TROSY)‐based NMR studies of the detergent‐solubilized PCAT1 (160 kDa) complex in solution. Using static and time‐resolved ultrafast NMR experiments, we were able to probe the conformational properties of CtA with and without cleavage in the PCAT1 complex. Having verified the precise length of the PEP‐binding motif from CtA, the peptide sequence (residues 5–37) complexed with catalytically inactive PEP^C21A mutant was used for subsequent structural and dynamic studies by solution NMR techniques. Magnetic field–dependent amide nitrogen (N¹⁵) spin–lattice (T₁), spin–spin (T₂), and heteronuclear ¹⁵N‐{¹H} nuclear Overhauser effect (NOE) data were acquired and analyzed using ModelFree ¹⁹ , ²⁰ to characterize multiple timescale motions in free PEP^C21A and its complex with the amino‐terminal leader peptide sequence. The results of the analysis provide novel insight into the role of internal dynamics in molecular recognition and PEP domain enzyme activity.

2. RESULTS

2.1. Resonance assignments of CtA and PEP^C21A domain

The backbone chemical shifts of the substrate CtA and PEP^C21A in the free and complexed states, respectively, were assigned using TROSY‐based triple‐resonance experiments. ²¹ The extensive perturbation of amide chemical shifts in the ¹H‐¹⁵N TROSY–heteronuclear single‐quantum coherence (HSQC) of PEP^C21A upon binding CtA (Figure 1a) and the complementary complex with labeled CtA (Figure 1b) is consistent with 1:1 stoichiometry and low‐micromolar binding affinity. ² The secondary structure predicted by the backbone (N, H^N, H^α, C^α, C^β, C′) chemical shifts in TALOS+ ²² (Figure 1c) confirms the isolated PEP^C21A domain adopts the canonical C39 PEP fold. A notable difference from the X‐ray structure of PCAT1 ² is the presence of well‐defined helix α4 connecting β1‐β2 strands in solution. In Figure 1d, the largest binding induced chemical shift changes (>0.2 ppm) in PEP^C21A domain map to residues in α2‐α3 helices, L2 loop, and β1‐β2‐β3 strands enclosing the binding site for the leader sequence. ⁶ , ⁹ , ²³ The secondary structure analysis of full‐length CtA encompasses a stable N‐terminal amphipathic helix (Residues 15–20) that corresponds to the canonical double‐glycine binding motif followed by an extended disordered region that is eventually excised as cargo for export (Figure 1e). Given the typical orientation of the helical peptide, ⁶ , ⁷ , ⁸ the conspicuously large chemical shift perturbation (CSP > 0.2 ppm) observed along the N‐terminal CtA sequence in Figure 1f (residues 7–11) suggests the binding motif extends beyond the amino‐terminus of the helical region (residues 15–20) than purported based on similarities to other C39 structures. A truncated version of the CtA peptide exhibited nearly identical CSP profile compared to that of the full‐length protein (Figure S1). Thus, a shorter peptide that could be expressed by recombinant methods was used for subsequent structural and dynamic studies.

FIGURE 1.

Overlay of N¹⁵‐HSQC, (a) ²H^/13C/¹⁵N‐labeled PEP^C21A in the presence (1:1 ratio, red contour) and absence (black contour) of unlabeled substrate CtA acquired at 15°C. (b) ¹⁵N/¹³C‐labeled CtA substrate in the presence (1:1 ratio, red contour) and absence (black contour) of unlabeled peptidase domain acquired at 25°C. The secondary structure probability from TALOS+ analysis of the backbone chemical shifts (C′, C^α, C ^β , N, H^N); (c) Isolated PEP^C21A domain and (e) CtA. Secondary structure propensity is color coded with α‐helices indicated in black and β‐strands by blue bars. (d) Bar plot of chemical shift perturbation (CSP) calculated from the relation √(0.5 × (δ _H ²+(0.2 × δ _N)²)) using the amide nitrogen (δ _Ν) and proton (δ _Η) chemical shift differences. (f) CSP analysis of ¹³C/¹⁵N‐labeled CtA in the presence and absence of unlabeled peptidase domain

2.2. Substrate interactions with PCAT1 in micelles

The secondary structure predictions from the chemical shifts of the isolated substrate support a predominantly disordered polypeptide chain with a single N‐terminal helix (Figure 1e). To map the binding‐induced conformational changes in CtA, we assigned the complete backbone and side‐chain resonances of the free substrate and transferred the same to a 1:1 complex with the inactive mutant of PCAT1^C21A in micelles. The ability to observe the resonances of CtA bound to the 160 kDa PCAT1^C21A transporter in micelles by ¹⁵N/¹³C‐edited TROSY‐NMR is a methodological benchmark. The substrate bound to the inactive transporter shows significant line broadening of the amide resonances accompanied by complete loss of signal intensity at the N‐terminus (residues 3–37; Figure 2a). Despite the prohibitively large size of the complex, the favorable relaxation properties of the fast‐rotating methyl groups enabled the shifted resonances from Leu13 and Leu18 to be detected from the binding sequence in the methyl‐TROSY spectrum (Figure 2b). The sharpness of the unperturbed resonances originating from the disordered C‐terminus of CtA is consistent with some fraction of the bound substrate remaining occluded from the TM channel in agreement with the previously reported CryoEM studies. ⁶ Owing to practical challenges of preparing stable samples with the transporter at sufficiently high concentration required for NMR, we were unable to detect the TM‐domain bound conformation of CtA in solution.

FIGURE 2.

Overlay of (a) ¹H‐¹⁵N TROSY, and (b) ¹H‐¹³C HMQC of specifically ¹³CH₃‐ILV, ²H/¹⁵N‐labeled CtA in the absence (black contour) and presence of inactive mutant PCAT1^C21A in UDM detergent micelles at 25°C (red contour), (c) ¹H‐¹⁵N HSQC, and (d) ¹H‐¹³C HMQC of ²H/¹⁵N, ¹³C‐labeled ILV‐methyl protonated intact CtA (black contour) and CtA cleaved by wild‐type PCAT1 at 5°C (1:0.1 ratio, red contour). The methyl resonances of CtA were assigned in a uniformly C¹³/N¹⁵‐labeled sample using standard triple resonance experiments (vide infra—Methods)

To measure the kinetic parameters of the catalytically active PEP domain attached to the PCAT1 transporter in the micelles, we acquired a series of ¹H‐¹⁵N SOFAST‐HMQC spectra as a function of time to follow the chemical shift changes in CtA during cleavage in solution. During the reaction, we follow the differential intensity of two peaks corresponding to the original and cleaved substrate (Figure 2c,d) change in response to the relative concentrations of the reactant and product. Based on the CSP, cleavage after the ‐GG‐ motif results in two fragments representing the N‐terminal leader sequence (residues 1–24) and the C‐terminal cargo peptide (Residues 25–90) dissociating from the enzyme. Further analysis of the progression curve (Figure S2) by applying the integrated Michaelis–Menten curve ²⁴ , ²⁵ yields K _m (0.21 ± 0.06 mM at 5°C) and V _max (7.2 ± 0.2 μM/min) values within the expected range of similar C39 enzymes. ⁹ The enzyme turnover with k _cat ~ 0.7/min at 5°C (~0.1/min at 37°C) ² is significantly slower than the rate of ATP hydrolysis in the NBD (10/min at 37°C). ² This implies the cleavage of the bound substrate in the PEP domain is intrinsically slow and thus prevents the accumulation of the cargo peptide in the cytoplasm before transport.

2.3. Structure determination of PEP^C21A complex with the leader peptide

To investigate the extent of structural coupling between the TM and PEP domains and its impact on the association with the substrate, we determined the ab initio NMR structure of the isolated PEP domain, PEP^C21A (Figure S3a), and its complex with CtA‐37 peptide in solution (Figure 3a). The intra‐ and intermolecular distance restraints derived from NOE data, dihedral angle restraints from TALOS+, and the final statistics for the ensemble of structures are summarized in Table 1. The αβ‐fold in the isolated PEP domain confirms the prototype C39 or papain‐like domain, with the active site enclosed by an open‐twist β‐sheet (right) and α‐helical (left) lobe, respectively (Figure 3b). The buried hinge region between the two lobes is shored up by extensive hydrophobic contacts between α1, β2‐β3, and α3, β4‐β5 strands, while the flexible loop L2 (α2‐α3; Figure S3b) modulates the accessibility of the binding cavity.

FIGURE 3.

(a) Stereoview of the ensemble of 20 PEP^C21A peptidase domain NMR structures (Molmol 2K.2) using line representation of the backbone (black) with the bound CtA‐37 peptide backbone colored red. (b) Ribbon representation of a single model structure of PEP^C21A domain (olive green) complexed with the CtA‐37 peptide (red) from the NMR ensemble shown in Panel A. The peptidase domain consists of an α‐helical (L, olive) and β‐sheet (R, sandy brown) subdomain forming a bilobe structure. The bound CtA‐37 peptide forms a single amphipathic helix between Residues 15–20. (c) Annotated display of the intermolecular interactions at the peptidase domain binding site between side chains from the protein (yellow) and CtA‐37 peptide (red) in stick representation. The zoomed inset (left) shows intermolecular H‐bonds (dashed lines) between main‐chain atoms from Table 2. (d) The peptidase domain surface color coded based on Kyte–Doolittle hydrophobicity in Chimera 1.15 ⁵² with a variable color range from blue (charged) to orange (hydrophobic) to indicate the type of side chain. The principal hydrophobic side‐chain anchors from the bound peptide are exhibited in wire‐and‐stick representation. (e) Displayed the charged side‐chain groups from the peptide complemented by the coulombic surface of the protein with a color scale ranging from positive (blue) to negative (red) charge. (f) Amino‐acid sequence alignment of PCAT1 substrates from different sources. The top row has the sequence number listed with reference to the cleaved peptide bond (‐GS‐)

TABLE 1.

NMR ensemble statistics

Constraints	C39	Complex
		C39	Peptide	C39 + peptide
Intraresidue	815	532	77
Sequential (|i − j| = 1)	830	498	48
Medium range(|i − j| < 4)	571	287	28
Long range (|i − j| > 5)	1,205	482	—
Intermolecular	−	77	—
Total	3,421	1,854	153
Phi	87	111	15
Psi	88	113	13
Total	175	224	28
Precision a
Backbone (Å)	0.55 ± 0.08	0.66 ± 0.10	0.16 ± 0.05	0.68 ± 0.09
Heavy atoms (Å)	0.98 ± 0.10	0.98 ± 0.09	0.91 ± 0.10	1.00 ± 0.09
PDB	7N87	7S5J
BMRB	30,926	>30,950
Ramachandran
Favored regions	89.3 ± 2%	92.2 ± 1.8%
Additional allowed regions	9.6 ± 1.9%	7.1 ± 1.7%
Clashscore	5.6 ± 1.3	8.3 ± 4.7
MolProbity	2.0 ± 0.2	2.1 ± 0.3
Violations	C39	Complex
Distance (d > 0.5Å)	0.0	0.1
Distance (d > 0.3Å)	0.9	0.2
Distance (d > 0.1Å)	4.3	12.1
Angle (θ > 5°)	0.3	1.1
E Total(kcal mol−1)	−5,453.2 ± 78.7	−6,493.9 ± 187.4
E noe	13.7 ± 2.2	25.3 ± 14.0
E vdw	−1,343.2 ± 10.4	−1,565.4 ± 15.7

^a The backbone RMSD and Ramachandran plot calculated for Residues 21–137 (PEP^C21A) and residues 15–20 (peptide).

In the PEP complex, the bound CtA‐37 peptide forms a short amphipathic helix (residues 15–20) lodged in the cavity between the two lobes making extensive contacts with α3 helix, β1‐β2 strands, and L2 loop (Figure 3b). The side chains from Leu(‐7) and Met(‐4) form a hub of hydrophobic interactions with the protein (Ala55, Tyr56, Ile59, Phe100, and Val137) that aligns the helix along the backbone of the L2 loop (Figure 3c,d). As shown in Figure 3e, the intermolecular contacts are augmented by favorable electrostatic forces between positively charged protein surface and the negatively charged peptide side chains at positions (‐8, ‐10, and ‐13).

The docked helix is supported by additional interactions spanning the unstructured N‐ and C‐terminal backbone. The N‐terminal end of the peptide (residues 5–14) is tethered to a distal hydrophobic patch on the R‐lobe (Figure 3c) by interactions with Ile(‐16) and Leu(‐18) side chains along with a putative H‐bond between Leu(‐12)^N‐Gly74^O main‐chain atoms. The H‐bond pair is also detected in a structurally similar position in other complexes (Table 2).

TABLE 2.

Summary of hydrogen‐bonding interactions at the substrate binding site

PEPC21A‐CtA a	Distance b Å (%)	6v9z c	6mpz d	Distance (Å)
Thr53N‐Gly(‐2) O	—	3.3 e	Ser59N‐Gly(‐2) O	2.9
Ala55N‐Met(‐4) O	2.9±0.1 (100)	3.45±0.05	Ala61N‐Val(‐4) O	2.8
Tyr56N‐Leu(‐7) O	3.3±0.2 (70)	3.05±0.15	Leu62N‐Leu(‐7) O	3.4
Ala98N‐Ser(+1) O	2.8±0.2 (100)	2.8 e	—	—
Leu(‐12) N ‐Gly71O	2.9±0.3 (85)	3.55±0.05	Leu(‐12) N ‐Gly77O	3.6
Gly(‐2) N ‐Thr53O	2.9±0.2 (100)	3.05±0.05	Gly(‐2) N ‐Ser59O	2.9
Ser(+1) N ‐Ala98O	3.3±0.2 (80)	—	—	—

^a The peptide atom in the H‐bond pair is indicated in bold.

^b The H‐bonded pairs are predicted based on published statistics ²⁶ for distances (<3.4 Å) commonly observed between donor–acceptor atom pairs in high‐resolution structures. The number in the bracket denotes the percentage fraction of the NMR ensemble where the distance is less than the cutoff.

^c Average H‐bond distance from the two asymmetric subunits (PDB 6v9z) determined by CryoEM.

^d Crystal structure of a double‐glycine motif protease from AMS/PCAT transporter—Lachnospiraceae bacterium C6A11—in complex with the leader peptide.

^e The H‐bond is only observed in a single subunit from the dimer structure solved by CryoEM (PDB 6v9z).

The C‐terminal cleavage sequence (residues 21–25) with the double‐glycine motif (‐MTGGS‐) is anchored by a network of H‐bond interactions with residues along the exposed backbone of the L2 loop (inset in Figure 3c). The minimum requirements for donor–acceptor distance pairs ²⁶ are satisfied by a significant fraction (>80%) of structures from the NMR ensemble (Table 2). Based on the known high‐resolution structures, the pairwise interactions involving the backbone atoms of Thr53, Ala55, and Tyr56 are remarkably well preserved in the complexes (Table 2) which underscores the importance of the L2 loop from C39 PEPs in target recognition.

In the transition state of the enzyme, the acyl intermediate with Cys21 typically occupies a narrow cleft between α1 and β3 which requires the ‐TGGS‐ motif to adopt an extended backbone to minimize steric clashes (Figure 3). However, CtA‐37 bound to the isolated PEP domain forms an atypical Type I turn ²⁷ at the cleavage sequence which sterically hinders access to the main‐chain atoms of Gly24 (‐1) and Ser25 (+1) (Figure 3c) by the catalytic triad (Cys21, His99, and Asp115). Unlike a typical Type 1 turn, the i to i+3 (N‐H..O=C) distance is not favorable for H‐bond formation between the main‐chain atoms even if the dihedral angles and Ca ⁱ ‐Ca ⁱ⁺³ distances (<7 Å) are in the recommended range. ²⁸ The unusual backbone fold is stabilized mainly by close hydrophobic contacts between the aromatic side chains of Phe(+3), Ile92, and Ala98 located in β2‐β3 strands. Based on the distance, the main‐chain atoms from Ala98^N‐Ser(+1)^O are also favorably poised for H‐bond formation. The archetype cysteine proteases are known to cleave longer substrates with the order of efficiency dictated by the bulk of the group at positions following the ‐GG‐ motif. ²⁹ , ³⁰ We can extend this analogy to the interaction hub anchored by Phe(+3) which essentially restricts access to the double‐glycine motif and thus offers a plausible mechanism for inhibition. The observed steric hindrance leads to a pertinent question on the mechanism that might enable structural rearrangement in the substrate or enzyme necessary for the formation of the transition state complex. In this context, given the flexible nature of the L2 loop and its importance in binding the target peptide provided a mechanistic rationale to investigate further the backbone dynamics of the enzyme in the free and substrate‐bound states, respectively.

2.4. Backbone dynamics from N ¹⁵ ‐relaxation data analysis

Amide nitrogen (N¹⁵) relaxation rates exhibit a variable response to backbone dynamics on distinct timescales. ³¹ Typically, the ¹⁵N‐{¹H} NOE values are most sensitive to picosecond bond fluctuations, ¹⁵N‐R₁ to pico‐nanosecond motions, and ¹⁵N‐R₂ to nanosecond rotational tumbling and micro‐to‐milliseconds conformational exchange processes. ³² To evaluate the role of dynamics in PEP domain function, we have acquired a suite of amide nitrogen (N¹⁵) relaxation rates for PEP^C21A and the substrate in the free and complexed states, respectively. The magnetic field–dependent N¹⁵ relaxation rates measured at 11.73T and 21.12T were analyzed by applying the Extended ModelFree (EMF) formalism ³³ with internal motions on two different timescales represented by Equation 1. Within the EMF framework, J(ω), the spectral density function is described by five parameters, global rotational correlation time (τ _m), fast picosecond (τ _f), slow nanosecond motion (τ _f << τ _s < τ _m), and square of the order parameters to represent the spatial restriction of the amide bond on fast (S _f ²) or slow timescales (S _s ²), respectively. The magnitude of the order parameters associated with each timescale (S ², S _i ² where i = f, s) ranges from 0 for free motion to 1 for static bond vector.

$$J\left(\omega \right)=\frac{2}{5}\left[\frac{{\tau }_{\mathrm{m}}{S}^2}{1+{\left(\omega {\tau }_{\mathrm{m}}\right)}^2}+\frac{\left(1-S_{\mathrm{f}}^2\right){\tau }_{\mathrm{f}}^{\prime }}{1+{\left(\omega {\tau }_{\mathrm{f}}^{\prime }\right)}^2}+\frac{S_{\mathrm{f}}^2\left(1-S_{\mathrm{s}}^2\right){\tau }_{\mathrm{s}}^{\prime }}{1+{\left(\omega {\tau }_{\mathrm{s}}^{\prime }\right)}^2}\right],{S^2=S_{\mathrm{f}}^2{S}_{\mathrm{s}}^2,\tau }_{\mathrm{i}}^{\prime }=\frac{{\tau }_{\mathrm{i}}{\tau }_{\mathrm{m}}}{{\tau }_{\mathrm{i}}+{\tau }_{\mathrm{m}}}\left(\mathrm{i}=\mathrm{f},\mathrm{s}\right).$$ (1)

Based on the above analytical form of J(ω), protein backbone motions fall under three broad categories: small amplitude fluctuations (0 < S _f ²  < 1, 10⁻¹² < τ _f (s) < 10⁻¹¹), correlated torsional movement in loops or terminal residues (0 < S _s ² < 1, 10⁻¹¹ < τ _s (s) < 10⁻⁹), and micro‐to‐millisecond timescale conformational exchange manifested as line broadening (R _ex ≠ 0). ¹⁸ , ³² , ³³

Applying the above description to EMF analysis of the N¹⁵‐relaxation rates measured on the free enzyme revealed a diverse range of motions (Figure S4) in the two‐domain structure represented as a worm plot in Figure 4a. In the L‐lobe, the terminal residues and loops connecting the three helices exhibit notable dips in the S ² values (Figure 4c) a direct consequence of both fast (S _f ²) and slow (S _s ²) timescale order parameters deviating from a static bond vector (Figure S4a, S4b) due to backbone motions on the pico‐to‐nanosecond time regime (Figure S4c, S4d). In addition to correlated motion along the unstructured backbone, the L2 loop also exchanges between different conformational states on a much slower timescale (micro‐milliseconds) resulting in R _ex terms for residues G52–G57 (Figure S4e). Together, the L1 loop (K36 and M37), α2 helix (I42, E44, and M45), and L2 loop (G47‐N54) constitute the dynamic core of the L‐Lobe that plays a crucial role in bridging the substrate binding site with TM domain in PCAT1 (Figure 5a).

FIGURE 4.

The radius‐of‐worm representation of the backbone of PEP^C21A domain scaled by the magnitude of internal motions on the picosecond–nanosecond timescale regime: (a) Free protein and (b) PEP^C21A complexed with CtA‐37 peptide. The presence of internal motions is represented by increasing tube radius and blue color gradient ranging from light (τ _f > 40ps) to dark shade (τ _s < 3ns). Residues with field‐dependent exchange terms (R _ex > 5 Hz at 21.12T) painted in orange. (c) Overlay of generalized order (S ² = S _f ² S _s ²) parameters of the free protein (black triangles) and the peptide complex (red circles) with flexible parts of the backbone in orange color bars and the more rigid secondary structure in blue. The dotted line indicates the separation between the L‐ and R‐lobe. Panels (d)–(f) displayed amide ¹⁵N‐{¹H} NOE, N¹⁵‐R₁ and N¹⁵‐R₂ relaxation measurements at 18.77T and 15°C for free CtA and Panels (g)–(i) CtA‐37 peptide bound to the PEP^C21A domain

FIGURE 5.

Effect of interdomain contacts on conformational changes triggered in the PEP domain coupled to substrate binding in intact PCAT1. The membrane‐bound PCAT1 structure has been solved in the free (X‐ray, PDB 4RY2) and substrate‐bound states (CryoEM, PDB 6V9Z). (a) Displayed single subunit from dimeric PCAT1 (4RY2), with the PEP domain in ribbon representation (green), space filling model of TM (cyan), and NBD (sandy brown). The average separation between the L‐ and R‐lobes from the peptidase domain is represented by the dashed line connecting the centroids calculated in Chimera. (b) Backbone superposition of PEP domain (residues 18–137) from solution NMR (blue ribbon) and PCAT1 X‐ray structure (4RY2, green ribbon) in the absence of substrate (backbone RMSD 1.8 Å). The bilobe structure is wider in PCAT1 (16.6 Å, Panel A) compared to the isolated domain (15.8 Å). (c) In the substrate‐bound state, superposition of the solution PEP domain (blue ribbon) and PCAT1 complex structure (6V9Z, green; backbone RMSD 1.4 Å) reveals the interlobe distance of 16.8 Å in the isolated protein is comparable with PCAT1 (17.1 Å)

Relative to the floppiness of the L‐lobe, the β‐sheet in the R‐Lobe undergoes only fast timescale picosecond breathing motions (τ _f < 25ps) and is practically devoid of nanosecond dynamics (Figure S4d). The exceptions include the inherently unstable 3₁₀‐helix (α4) and scattered residues at the peptide binding cavity flanked by β1‐β2‐β3 strands (Figure 4a).

Substrate binding triggers a notable rigidification of the enzyme backbone (Figure 4b) as reflected by the flatter profile of order parameters with elevated values in the flexible regions highlighted in Figure 4c. However, the motions in the L2 loop and β2‐β3 strands are only partially quenched, and some flexibility persists at the protein–peptide interaction surface in the complex. These conclusions are well supported by the relaxation data fitted by simpler motional models for a large fraction of residues without the need to invoke multiple timescales or exchange terms except for selected residues (Figure S4f–j).

To probe the loss of mobility in the bound substrate, relaxation rates for free CtA and CtA‐37 complex with PEP^C21A were measured at a single field. In free CtA, the low average ¹⁵N‐{¹H} NOE values (~0.38 ± 0.12) along with slower N¹⁵‐R₂ relaxation rates (~5.2 ± 0.9 s⁻¹) is consistent with random segmental motion in disordered proteins ³⁴ (Figure 4d,f). The restricted mobility in the single α‐helix (Figure 1e) is reflected by elevated ¹⁵N‐{¹H} NOE (~0.60 ± 0.01) and N¹⁵‐R₂ values (6.4 ± 0.3 s⁻¹) relative to the remaining unstructured polypeptide chain. This distinction between the ordered and disordered regions of the backbone is amplified further in the bound peptide. The ¹⁵N‐{¹H} NOE values increase (~0.76 ± 0.09) for residues 11–21 immobilized in the PEP domain binding site (blue bars; Figure 4g). The dramatic drop in N¹⁵‐R₁ (0.6 ± 0.1 s⁻¹) with increasing R₂ (32.6 ± 2.1 s⁻¹) values (Figure 4h,i) is primarily a function of slower tumbling time of a larger complex. The mobility observed beyond the cleavage site at the double‐glycine motif (residues > 25) corroborates the structural flexibility noted previously in the C‐terminal region of bound CtA (Figure 1e). To summarize, our data presents direct evidence of a dynamic association between the PEP domain and substrate which allows structural rearrangement at the binding site to optimize cleavage and translocation.

3. DISCUSSION

High‐resolution NMR in concert with nuclear spin relaxation measurements offers a powerful tool to probe the inherent flexibility of enzymes and illustrates how these motions impact catalytic activity and substrate binding. ³² A comparison of the free and substrate‐bound PCAT1 PEP domain NMR structures reveals a flexible binding surface that molds itself to the leader sequence derived from its cognate binding partner. The unbound substrate is mostly disordered, except for the leader sequence forming a stable two‐turn helix in solution. In the bound state, the helix is embedded in the hydrophobic groove enclosed by two lobes of the structure (Figure 3b).

The structural determinants of the C39 enzyme‐substrate selectivity and binding affinity are encoded by a combination of hydrophobic, electrostatic, and sequence‐specific H‐bond interactions. A sequence alignment of PEP domains from different sources shows residues critical for orienting the cleavage site (‐MTGGS‐) along the L2 loop from S1 (Cys21, Gly22), S2 (Gly52, Thr53), and S5 (His99, Phe100) subsites are highly conserved (Figure S5) which results in a generic affinity for cognate substrates. Notably, the peptide‐to‐protein H‐bonded pairs Gly(‐2)‐Thr53 and Met(‐4)‐Ala55 (Table 2) are preserved in all the known complexes which highlights the importance of these interactions in determining the proteolytic selectivity of the bacteriocin‐based PEP enzymes. ⁷ , ⁸ , ⁹ , ³⁵

In contrast to the conserved scaffold at the cleavage site, the remaining N‐terminal leader sequence of the peptide including the helix interacts with a variable but predominantly hydrophobic surface formed by the S2‐S4 subsites from the PEP domain. Despite the sequence diversity in the S2‐S4 subsites, structural studies of CtA, ⁶ LahT147, ⁷ and ComC ⁹ complexes have revealed a common mode of interaction involving hydrophobic side chains at ‐4, ‐7, and ‐12 positions from the substrate adapting to the topology of the protein binding surface. The remarkably similar structures of the complexes underscore the significance of mutational studies of substrates of ABC transporter ComA from Streptococcus that concluded the PEPs discriminate between substrates by enzyme efficiency rather than binding affinities. ⁸ , ⁹

For the first time in the NMR structure of the longer peptide, we observe distal contacts that extend beyond the nominal binding site at both ends of the helix. The release of the leader peptide after cleavage suggests these peripheral contacts have marginal effect on the substrate binding affinity but instead regulate catalytic efficiency by steric inhibition. ³⁰ Normally, the double‐glycine motif adopts an extended backbone configuration to minimize steric clashes during the formation of the acyl intermediate in the transition state. ³⁶ However, the interactions anchored by Ser(+1) and Phe(+3) from CtA stabilize a kinked and catalytically incompetent substrate conformation that explains the drastic loss of activity in the detached PEP complex in the ATP bound state of PCAT1. ² Clearly, the kinetic bottleneck prevents buildup of nonsecreted cargo peptide during ATP hydrolysis and leads to the obvious question if this mode of substrate binding is also relevant in the pre‐translocation state of PCAT1 where the domains are coupled (Scheme 1).

While the asymmetric CryoEM structure showed at least one of the PCAT1 subunits bound to the substrate with a similar fold, ⁶ the ability to observe the resonances of CtA bound to the intact PCAT1^C21A transporter in detergent micelles by TROSY‐NMR offers further proof of the presence of dynamics in the pre‐translocation complex in solution. The loss of amide signals originating from the immobilized leader peptide in the PEP domain compared to the sharpness of the signal arising from the C‐terminal residues of CtA (Figure 2a,b) confirms the latter is excluded from the TM channel. This result can be rationalized if the leader sequence from PCAT1 bound CtA adopts a fold similar to CtA‐37 which leaves the disordered C‐terminus to flop in solution. Nonetheless, the increased efficiency of cleavage when the PEP and TM domains are coupled together ² suggests the bound substrate has some flexibility to reorganize and in process release the steric hindrance that inhibits cleavage in the isolated complex.

The dynamic reorganization of the binding site interactions is supported by a key finding of this study: The accessibility of the active site opening is regulated by a flexible L2 loop and correlated motions in the β2‐β3 strands that function like jaws of a clamp to enclose the substrate. The dynamics exhibited in the PEP domain subsites S2 (L2 loop), S3 (α4 helix), and S5 (β2‐β3 turn; Figures 4b and S5) are reminiscent of a selected fit model of enzyme action where the bound substrate has wiggle room to achieve the active state configuration prior to cleavage. Similar conformational variability has been reported in the active site of papain homologue cathepsin K and implicated in the allosteric inhibition by small effector molecules. ³⁷ The remarkable plasticity of the L‐lobe from the PEP domain also sheds new insight into a plausible mechanism to mediate enzyme activity by modifying the dynamics at the binding site through association with the TM domain (Figure 5a).

A notable difference between the isolated PEP domain and the intact PCAT1 structure is in the juxtaposition of interdomain contacts with the substrate binding L2 loop. In the ATP‐free state of PCAT1, these bridging contacts between the PEP domain, and the TM or NBD helices are mediated by residues located at the N‐terminus, α2, and L2 loop (Figure 5a). The major deviations in the backbone superposition of the isolated enzyme with the X‐ray structure of the PEP domain attached to PCAT1 (PDB 4ry2, RMSD 1.8 Å) can be traced to the flexible loop L2, β2‐β3 turn, and the altered position of secondary structure (α2, β4) at the surface buried by the TM domain (Figure 5b). The mechanistic role of the domain coupling on the enzyme activity is evident from the enlarged binding cavity enclosed by the L‐ and R‐lobes of PEP in the isolated protein (15.8 Å) compared to PCAT1 (16.6 Å, Figure 5b). In contrast, the backbone of the substrate‐bound enzyme structures is superimposable (1.4 Å) with similar lobe positions (~16.8 Å) irrespective of the association with the TM domain. Thus, even if the width of the active site has nominal effect on the mode of substrate binding in PEP domain, it is conceivable the floppiness in the L‐lobe is partially restrained by the interdomain contacts which reduces the entropic cost of any structural rearrangement triggered in the substrate (Figure 5a).

The hinge movement of domains coupled to gated access to active sites is a common structural and dynamic paradigm in enzymes and a critical component of the mechanism. ³⁸ , ³⁹ In PCAT1, the dynamics in the PEP domain are fine‐tuned for substrate recognition and activity by enabling the latter to switch between different bound conformations with the L2 loop acting as a flexible gate to the active site. Comparison of the isolated enzyme and TMD‐coupled PCAT1 structures is consistent with domain movement in the PEP structure that provides facile entry of the substrate to the active site followed by reorganization in the complex prior to cleavage. The proposed mechanism is expedited by dynamic PEP interactions with the TM domain which widens the active site and stabilizes substrate conformations that are favorable for cleavage compared to the isolated protein. To conclude, PCAT1 functionality is a sum of moving parts with the coupled domains operating in a concerted fashion to transport the cargo peptide.

4. MATERIALS AND METHODS

The mutant PEP domain (PEPC21A, residues 1–148), wild‐type, and mutant (C21A) PCAT1 genes (1‐722) were cloned into the pMSCG20 plasmid with N‐terminal GST‐fusion tag and a TEV cleavage site. For structural characterization of the enzyme–substrate complexes, the single C21A mutation in the PEP domain (Scheme 1) known to abrogate all protease activity was used. The substrate, CtA, and the N‐terminal peptide CtA‐37 (Residues 5–37) were cloned into vector pMCSG7 with N‐terminal 6xHis tag. Details of the various protein expression and purification strategies are described in Supplementary Methods. Summarized below are only the modified expression protocols for uniform and selective isotope enrichment of proteins used to prepare the NMR samples.

5. LABELED PROTEIN EXPRESSION

5.1. Uniformly ¹⁵ N‐ and ¹³ C/ ¹⁵ N‐labeled protein expression

The cells from the 50 ml starter culture of LB grown overnight 37°C were harvested by centrifugation at 4°C (4,000 rpm for 15 min), washed and resuspended in 10–20 ml of M9 media. One fifth of the total culture volume, after washing, was used to inoculate 1 L of M9 media supplemented with trace metal ions. ⁴⁰ Depending on the labeling pattern, 1 g/L ¹⁵N/¹³C‐labeled ISOGRO (Sigma), 1 g/L ¹⁵NH₄Cl, or 2 g/L U‐¹³C D‐glucose (CIL) was substituted as sole nitrogen and carbon source for the unlabeled reagents. Freshly inoculated cultures were incubated at 37°C/250 rpm until optical density (OD)₆₀₀ = 0.4 and then cooled to 20°C. At OD₆₀₀ = 0.5–0.6, the cultures were induced with 1 mM IPTG and incubated for 24 hr at 20°C/250 rpm; 1 L cultures were harvested by centrifugation at 4°C/4,000 rpm for 15 min and stored at −80°C.

5.2. Uniformly ² H/ ¹³ C/ ¹⁵ N‐labeled and ¹ H/ ¹³ C methyl ILV, ² H/ ¹⁵ N‐labeled protein expression

The cells harvested from an overnight 20 ml LB culture was washed and resuspended in 50 ml M9 media in 50% D₂O incubated at 37°C/250 rpm for 8 hr before harvesting, washing, and resuspending the cells in 50 ml M9 media in 100% D₂O incubated at 37°C/250 rpm overnight. The cells adapted to D₂O were used to inoculate 1 L of M9 Media, supplemented with ampicillin, and modified for respective labeling schemes. For ²H/¹³C/¹⁵N‐labeled expression, ¹⁵NH₄Cl, U‐¹³C, D₇‐glucose and ¹³C/¹⁵N, D‐Isogro were substituted for unlabeled reagents. For ILV‐methyl protonated samples, ¹⁵NH₄Cl and D₇‐glucose were substituted for unlabeled reagents and Isogro was omitted. After inoculation, cultures were incubated at 37°C/250 rpm. At OD₆₀₀ = 0.4, the cultures were cooled to 20°C. To enable ILV‐methyl specific protonation, 120 mg/L α‐ketoisovaleric acid sodium salt (3‐methyl‐¹³C, 3, 4, 4, 4‐D₄) and 70 mg/L α‐ketobutyric acid sodium salt (methyl ¹³C, 3, 3‐D₂) were added to each culture at OD₆₀₀ = 0.4. At OD₆₀₀ = 0.5–0.6, cultures were induced with 1 mM IPTG and incubated for 24 hr at 20°C/250 rpm. Cultures were then harvested by centrifugation at 4°C/4,000 rpm for 15 min, and the pellets were frozen and stored in −80°C.

6. NMR SAMPLES

The following samples were used for chemical shift assignments, map binding site, and structural characterization: (a) U‐¹³C/¹⁵N‐labeled PEP^C21A (675 μM) in buffer A, (b) U‐¹³C/¹⁵N‐labeled CtA (150 μM) in buffer B, (c) U‐¹³C/¹⁵N‐labeled PEP^C21A (170 μl, 165 μM) and unlabeled CtA‐37 (30 μl, 1.7 mM) at protein:peptide ratio 1:1.8 in buffer A, (d) U‐¹³C/¹⁵N‐labeled CtA‐37 (180 μl, 222 μM) and unlabeled PEP^C21A (37 μl, 1.75 mM) at peptide:protein ratio 1:1.6 in buffer A, (e) U‐²H/¹³C/¹⁵N‐labeled PEP^C21A (100 μM) and unlabeled CtA in buffer A at pH 8.0, and (f) U‐²H/¹³C/¹⁵N‐labeled CtA (100 μM) and unlabeled PEP^C21A in buffer A at pH 8.0, (g) ¹³CH₃‐ILV, ²H/¹⁵N‐labeled CtA (100 μM) complexed with proteolytically inactive PCAT1^C21A mutant in buffer C. All three binary complexes at 1:1 ratio in samples (e), (f), and (g) were copurified by size exclusion chromatography to wash away unbound substrate. The buffer constituents are as follows: (A) 50 mM sodium phosphate, 150 mM NaCl, pH 7.0, 5 mM DTT; (B) 20 mM Tris buffer, 150 mM NaCl, pH 7.0, 5 mM DTT; and (C) 50 mM sodium phosphate buffer, 150 mM NaCl, pH 7.0, 5 mM DTT, 2mM n‐Undecyl β‐D‐Maltopyranoside detergent.

7. NMR SPECTROSCOPY

7.1. PEP^C21A and CtA‐37 peptide complex

The NMR data were acquired on Bruker AVANCE spectrometers equipped with TCI CryoProbes at 18.77T or 21.12T. A standard suite of triple‐resonance backbone (HNCO, HNCA, HN(CO)CA, HNCACB, and CBCA(CO)NH) and side‐chain experiments (CCH‐TOCSY, HBHA(CO)NH, CC(CO)NH, and HC(CCO)NH) were employed for chemical shift assignments of PEPC21A in the free (Sample A) and CtA‐37 peptide bound states (Sample C), respectively. The resonances of ¹³C/¹⁵N‐labeled CtA‐37 peptide bound to unlabeled PEP^C21A (Sample D) were assigned using 3D HNCA and HCCH‐COSY/TOCSY experiments. Distance restraints required for structure calculations were obtained from 100 ms mixing time ¹⁵N‐edited and ¹³C‐edited 3D‐NOESY‐HSQC (aliphatic and aromatic regions separately). Intermolecular NOEs were obtained from 2D‐¹⁵N,¹³C f ₂‐filtered NOESY recorded with 150 ms mixing time.

7.2. PEP^C21A and full‐length CtA complex

The complete resonance assignments of isolated CtA (Sample B) were obtained from standard backbone and side‐chain experiments described above for nondeuterated proteins at 21.12T B₀ field and 25°C temperature. For the larger complexes involving deuterated PEP^C21A domain (Samples E and F), TROSY based three‐dimensional backbone (TR‐HNCO, TR‐HNCA, TR‐HN(CO)CA, TR‐HNCACB, and TR‐HN(CO)CACB) experiments were employed for backbone chemical shift assignment at 21.12T B₀ field and 15°C temperature.

7.3. Wild‐type and mutant PCAT1‐C21A in detergent micelles complexed with substrate

To map the binding‐induced CSPs in the substrate, we acquired N¹⁵‐edited TROSY‐HSQC (backbone) and C¹³‐edited HMQC spectrum (methyl region) of CtA in the free state (Sample B) and PCAT1^C21A complex (Sample G) in the absence of proteolytic activity. The backbone and methyl resonance assignments for PCAT1^C21A–CtA complex were transferred from PEP^C21A–CtA complex.

To monitor the kinetics of proteolysis in real time, a sample consisting of 100 μM uniformly N¹⁵‐labeled CtA and 10 μM wild‐type PCAT1 in Buffer D was used to record a series of ¹⁵N‐edited SOFAST‐HMQC ⁴¹ spectra (~3 min) at 16.44T and 5°C temperature (to slow the reaction) until the reaction is complete (~1 hr). The data were processed, and amide cross‐peaks were integrated in Topspin 3.5 to obtain the reaction curves of the substrate. Residues 3–37 all show two sets of peaks corresponding to the reactant and product during cleavage. The curves were fitted using the closed form of the Michaelis–Menten equation using an in‐house python script. ²⁴ , ²⁵ Figure S2 summarizes the data for selected residues from the peptide and the final fitted parameters.

7.4. Backbone amide nitrogen relaxation experiments

The laboratory frame R₁, R₂, and ¹⁵N‐{¹H} NOE relaxation spectra on free C¹³/N¹⁵‐labeled PEP^C21A domain and CtA‐37 bound state were recorded at two static fields (11.73T and 21.12T) and 15°C sample temperature according to established methods. ⁴² , ⁴³ The relaxation data sets on free CtA and C¹³/N¹⁵‐labeled CtA‐37 peptide complex were acquired at 15°C and 18.77T. The pseudo 3D experiments were acquired with variable relaxation delays for R₁ (16, 80 × 3, 160, 240, 320, 400, 480, 560, 640, 720, 800, 880, 960, 1,040 ms) and R₂ (0.0, 16, 32, 48 × 3, 64, 80, 96, 112 ms) measurements. The heteronuclear ¹⁵N‐{¹H} NOE experiments were acquired with 3 s saturation plus 2 s recycle delay, and the reference experiment with 7 s recycle delay. The two‐dimensional data sets were processed in NMRPipe, ⁴⁴ and the relaxation rates extracted from the mono‐exponential decay of the intensity using rate analysis tools in NmrViewJ 8.0.3. ⁴⁵ The field‐dependent relaxation rates for free and peptide‐complexed PEPC21A domain were analyzed in the RELAX 4.0.3 program ⁴⁶ using the EMF formalism ³³ with local correlation time to account for the diffusion anisotropy. ⁴⁷ The 3 × 2 amide nitrogen relaxation rates measured for each residue are fit up to 10 different motional models derived from Equation 1, with and without slow exchange (R _ex) contribution to ¹⁵N‐R₂ on the micro‐millisecond timescales. The dynamic model that best fits the experimental data is selected based on Akaike information criterion (AIC) implemented in RELAX 4.0.3. ⁴⁶ The results of the best‐fitted model for each residue are summarized in Figure 4 and Figure S4.

8. STRUCTURE CALCULATIONS

The NMR data were processed in Topspin 2.1 from Bruker Biospin and the spectra analyzed in CARA 1.5. ⁴⁸ Two structures were calculated of free and CtA‐37 peptide bound PEP^C21A domain, respectively. The ¹⁵N‐edited and ¹³C‐edited aliphatic/aromatic 3D‐NOESY‐HSQC spectra of free and peptide bound U‐¹³C/¹⁵N‐PEP^C21A domain and CtA‐37 peptide were assigned initially with the CANDID‐based automation routine embedded in CYANA 2.1 ⁴⁹ followed by manual inspection of lone/lower quality peak assignments and consistent NOE violations. The intermolecular NOEs were manually assigned with the aid of the 2D‐¹⁵N,¹³C f ₂‐filtered NOESY spectrum. The unambiguous distance restraints output from the automation run was recalibrated by increasing all the upper distance limits by ~10% and the dihedral angle restraints derived from the analysis of the backbone chemical shifts in the TALOS program. ²² The optimum set of distance and angle restraints were imported in CNS 1.1/ARIA 2.3 to generate an ensemble of 500 structures with explicit water refinement. ⁵⁰ The structure calculation and refinement were guided by MolProbity ⁵¹ quality checks at each iteration, and the statistics of the final 20 lowest energy structures are reported in Table 1. The coordinates and chemical shifts have been deposited in RCSB and BMRB databases (Table 1).

AUTHOR CONTRIBUTIONS

Shibani Bhattacharya: Conceptualization (equal); formal analysis (lead); methodology (equal); writing – original draft (lead); writing – review and editing (lead). Anthony Palillo: Conceptualization (equal); formal analysis (supporting); methodology (equal); writing – original draft (supporting); writing – review and editing (supporting).

Supporting information

Appendix S1: Supporting Information

Bhattacharya S, Palillo A. Structural and dynamic studies of the peptidase domain from Clostridium thermocellum PCAT1 . Protein Science. 2022;31:498–512. 10.1002/pro.4248