Crystal Structures of a Thermophilic Cutinase from Chaetomium thermophilum Reveal Conformational Dynamics of the Catalytic Lid Loop

Nojima, Ryohei; Chen, Lirong; Kurokawa, Minami; Ito, Sho; Nishino, Tatsuya

doi:10.3390/cryst16040217

Open AccessArticle

Crystal Structures of a Thermophilic Cutinase from Chaetomium thermophilum Reveal Conformational Dynamics of the Catalytic Lid Loop

by

Ryohei Nojima

^1,†,

Lirong Chen

^1,†,

Minami Kurokawa

²,

Sho Ito

¹

and

Tatsuya Nishino

^1,*

¹

Faculty of Advanced Engineering, Tokyo University of Science, 6-3-1 Niijyuku, Katsushika, Tokyo 125-8585, Japan

²

Institute for Integrated Radiation and Nuclear Science, Kyoto University, 2-1010 Asashironishi, Kumatori, Osaka 590-0494, Japan

^*

Author to whom correspondence should be addressed.

^†

These authors contributed equally to this work.

Crystals 2026, 16(4), 217; https://doi.org/10.3390/cryst16040217

Submission received: 9 March 2026 / Revised: 20 March 2026 / Accepted: 21 March 2026 / Published: 24 March 2026

(This article belongs to the Special Issue Crystallography of Enzymes (2nd Edition))

Download

Browse Figures

Versions Notes

Abstract

Microbial cutinases are promising biocatalysts for polymer recycling. Here, we investigated the structural basis of catalytic activation in a thermophilic cutinase from Chaetomium thermophilum (CtCut). Differential scanning calorimetry revealed a three-state thermal unfolding pathway (Tm = 66.4 °C and 69.5 °C), indicating hierarchical stability. To capture distinct conformational states while avoiding affinity-tag artifacts, we employed both tag-free and tagged constructs. We determined apo-structures of wild-type and S136A mutant CtCut at 1.7 Å resolution and a complementary inhibitor complex at 2.65 Å. In the apo-state, a chloride ion coordinated the electrostatically pre-organized active site, while the catalytic H204 adopted a solvent-exposed, inactive loop conformation. In the inhibitor complex, p-nitrophenol displaced the chloride, establishing a characteristic oxyanion hole network. Concomitantly, the “lid” loop transitioned to an open state, with H204 exhibiting pronounced conformational heterogeneity across eight independent molecules. These complementary structures provide structural evidence for conformational dynamics of the catalytic lid loop, consistent with the conformational cycling model previously proposed for a mesophilic homolog.

Keywords:

cutinase; thermophilic enzyme; crystal structure; conformational dynamics; Chaetomium thermophilum; oxyanion hole; lid loop dynamics; thermostability

1. Introduction

Cutinases (E.C. 3.1.1.74) are versatile extracellular enzymes originally evolved by phytopathogenic fungi and bacteria to hydrolyze cutin, a complex, insoluble polyester that forms the structural framework of plant cuticles [1]. Belonging to the α/β-hydrolase superfamily, cutinases possess the unique ability to bridge the functional gap between true esterases and lipases; they efficiently hydrolyze a wide range of substrates without requiring the substantial interfacial activation characteristic of classical lipases.

In recent years, microbial cutinases and their homologs have attracted immense attention as robust industrial biocatalysts for depolymerizing synthetic plastics, particularly poly(ethylene terephthalate) (PET). Following the discovery of the leaf–branch compost cutinase (LCC) [2] and the highly active IsPETase from Ideonella sakaiensis [3], significant progress has been made in elucidating their catalytic mechanisms [4,5] and engineering highly efficient variants, such as LCC-ICCG [6], FAST-PETase [7], and HotPETase [8]. More recently, comprehensive structural and biochemical analyses have further expanded the repertoire of PET-hydrolyzing enzymes and provided a framework for their industrial application [9,10]. For practical industrial applications, enzymatic depolymerization is most efficient near the glass transition temperature of PET (~70 °C), making thermophilic enzymes highly desirable [11]. However, the rational design of such enzymes is often hindered by a fundamental thermodynamic trade-off: global structural rigidity is required to endure high temperatures, whereas local active-site flexibility is necessary to accommodate bulky polymer substrates which is a feature identified as a critical “hallmark” for efficient polyester degradation [12].

Pioneering structural studies on the mesophilic cutinase from Fusarium solani (FsCut) established two key principles of cutinase catalysis: the catalytic triad (Ser–Asp–His) operates within an accessible active site, and the oxyanion hole is “preformed” in the apo-state, ready to stabilize the tetrahedral intermediate [13,14,15]. A contrasting observation was reported for Glomerella cingulata cutinase (GcCut), in which the catalytic histidine (His204) was found to adopt a solvent-exposed, inactive conformation in the crystal structure. Nyon et al. demonstrated through H204N mutagenesis and activity assays that this residue is essential for catalysis and proposed a “conformational cycling” mechanism in which the histidine must reposition from an inactive to an active state during the catalytic cycle [16]. However, whether such conformational cycling occurs in thermophilic cutinases and how it is reconciled with the preformed oxyanion hole architecture remain open questions. Addressing this question crystallographically is further complicated by the well-documented effects of affinity tags on crystal packing and active-site conformation [17].

To investigate these structural questions, we chose the cutinase from Chaetomium thermophilum (CtCut), a thermophilic fungus widely utilized as a model organism for structural biology [18]. We employed a complementary structural approach: a tagged construct was used to capture a ligand-bound state at 2.65 Å resolution, while a fully tag-free strategy yielded high-resolution (1.7 Å) apo-structures of both the wild-type and S136A mutant enzymes. Combined with differential scanning calorimetry measurement, these structures provide structural evidence for conformational dynamics of the catalytic lid loop region and offer new insights into the balance between thermal stability and active-site flexibility in thermophilic cutinases.

2. Materials and Methods

2.1. Gene Cloning and Construct Design

Several expression constructs were designed to optimize the expression and solubility of Chaetomium thermophilum cutinase (CTHT_0064000; CtCut) for structural analysis. These included constructs encoding the full-length protein without a fusion tag (CtCutΔM1), a truncated version lacking the predicted N-terminal signal peptide (CtCutΔM1–V19), and a version fused with an Escherichia coli alkaline phosphatase signal peptide (PhoA–CtCutΔM1–V19). To achieve high yields of soluble protein, two types of thioredoxin (TrxA) fusion constructs were ultimately engineered using the In-Fusion HD Cloning Kit (Takara Bio, Shiga, Japan).

One construct, cloned into a pRSFDuet-1 vector (kanamycin resistance), consisted of an N-terminal TrxA tag, a thrombin cleavage site (LVPR|GS), the CtCutΔM1–V19 sequence, and a C-terminal 8 × His-tag. After thrombin cleavage, the resulting protein retained the C-terminal His-tag (approx. 23.1 kDa).

To observe the enzyme’s conformation without tag-mediated packing constraints, another construct was cloned into a pCold-I vector (ampicillin resistance). This construct placed a 6 × His-tag and the TrxA fusion tag upstream of the thrombin cleavage site (6 × His–TrxA–LVPR|GS–CtCutΔM1–V19). Thrombin cleavage of this construct allowed for the complete removal of all affinity tags, yielding the pristine tag-free WT and S136A proteins (approx. 22.0 kDa). The inactive S136A mutation was introduced into both TrxA-fusion constructs via site-directed mutagenesis.

2.2. Protein Expression and Purification

The expression plasmids were transformed into E. coli Rosetta (DE3) cells (Novagen, Madison, WI, USA). Transformants were selected on LB agar plates containing the appropriate antibiotic (0.1 mg/mL ampicillin for the pCold construct or kanamycin for the pRSF construct) at 37 °C overnight. A single colony was inoculated into 1 L of LB medium supplemented with the same antibiotic and cultured at 37 °C with shaking at 120 rpm until the OD₆₀₀ reached 0.4. Protein expression was then induced by the addition of isopropyl β-D-1-thiogalactopyranoside (IPTG) at a final concentration of 0.1 mM, and cells were cultured at 18 °C with shaking at 90 rpm for 18 h. The cells were harvested by centrifugation (4500 rpm, 4 °C, 45 min), resuspended in A0 buffer (10 mM Tris-HCl, pH 8.0), and stored at −80 °C.

Frozen cell pellets were resuspended in 5 volumes of A0 buffer containing 1× BugBuster Protein Extraction Reagent (Merck, Darmstadt, Germany) and house-made recombinant Serratia marcescens nuclease (1 μg/mL final) and incubated at room temperature for 30 min to achieve cell lysis. The lysate was clarified by centrifugation (12,000 rpm, 4 °C, 15 min) and filtered through a 0.22 μm membrane. For the tag-free constructs, the supernatant was applied to a 10 mL HisTrap FF crude column (Cytiva, Marlborough, MA, USA) and eluted with B0 buffer (10 mM Tris-HCl pH 8.0, 500 mM imidazole). After desalting (HiPrep Desalting column, Cytiva), the eluate was treated with thrombin (Sigma-Aldrich, St. Louis, MO, USA, 400 μL of 1 U/μL) at room temperature for 24 h to cleave the N-terminal TrxA–His fusion. Thrombin was removed by passage through a Benzamidine Sepharose column (Cytiva), and cleaved tags were separated from CtCut by a second HisTrap column. The tag-free CtCut, collected in the flow-through fraction, was concentrated to 5 mL using a 10,000 MWCO centrifugal filter (Amicon Ultra, Millipore, Billerica, MA, USA) and further purified by size-exclusion chromatography on a Superdex 200 column (Cytiva) pre-equilibrated with A1 buffer (10 mM Tris-HCl pH 8.0, 500 mM NaCl). The single-peak fraction was collected, buffer-exchanged into A0 buffer, and concentrated to approximately 50 mg/mL for crystallization. Protein purity was confirmed by SDS-PAGE.

2.3. Differential Scanning Calorimetry

The thermal stability of the purified WT enzyme was evaluated using a differential scanning calorimeter (NanoDSC, TA Instruments, New Castle, DE, USA). The protein sample (0.9 mg/mL in 10 mM Tris-HCl, pH 8.0; 700 μL) was degassed under vacuum for 10 min prior to measurement. Temperature scans were performed from 30 to 100 °C at a heating rate of 1 °C/min. Baseline correction was applied by subtracting the buffer-only scan, and the resulting thermogram was analyzed in the temperature range of 50–90 °C [19,20,21].

2.4. Crystallization

All crystallization experiments were conducted using the sitting-drop vapor diffusion method on 96-well plates.

Apo CtCut^WT (Tag-free): Purified tag-free CtCut^WT (25 mg/mL in 10 mM Tris-HCl, pH 8.0) was mixed in a 1:1 ratio with a reservoir solution containing 16% (w/v) PEG 3350, 0.3 M CaCl₂, and 0.2 M NaCl, and incubated at 20 °C. This condition yielded multi-layered, sea-urchin-shaped crystal clusters. To obtain single crystals suitable for high-resolution diffraction, individual layers were physically exfoliated from the clustered aggregates using a microneedle under a microscope. The isolated crystals were briefly soaked in a cryoprotectant solution consisting of the reservoir solution supplemented with ethylene glycol (in an 8:2 ratio) before flash-cooling in liquid nitrogen.

Apo CtCut^S136A (Tag-free): Purified tag-free CtCut^S136A (27 mg/mL in 10 mM Tris-HCl, pH 8.0, 0.2 M NaCl) was mixed in a 1:1 ratio with a reservoir solution containing 0.05 M MgCl₂, 0.1 M Tris-HCl (pH 8.5), and 40% (v/v) ethanol, and incubated at 20 °C. Similar to the WT, this yielded clustered crystals that required physical exfoliation. The isolated single crystals were cryoprotected in a solution containing the reservoir solution and ethylene glycol (in a 7:3 ratio) prior to flash-cooling.

CtCut^S136A mutant/inhibitor complex (C-terminal His-tag): The C-terminally His-tagged CtCut^S136A (40 mg/mL in 10 mM Tris-HCl, pH 8.0) was pre-incubated with 10 mM of the esterase inhibitor ME-600. The mixture was crystallized by mixing with a reservoir solution containing 0.1 M imidazole (pH 8.0) and 10% (v/v) isopropanol at 17 °C. Plate-like single crystals were obtained directly. Crystals were cryoprotected using a solution containing the reservoir solution, 4.0 M trimethylamine N-oxide dihydrate, and ethylene glycol in an 8:1:1 ratio, and subsequently flash-cooled in liquid nitrogen.

2.5. X-Ray Data Collection and Structure Determination

X-ray diffraction data were collected at beamline PF-BL1A (Photon Factory, Tsukuba, Japan) under a nitrogen cryostream at 100 K. Diffraction data were indexed, integrated, and scaled using XDS [22]. The data completeness for the wild-type apo structures was slightly lower in the outermost shell, likely due to the presence of minor crystal anisotropy. The structures were determined by molecular replacement using the PHENIX suite [23], with AlphaFold3-predicted structures [24] of CtCut^WT and CtCut^S136A serving as initial search models for the tag-free constructs, and an AlphaFold2-predicted structure [25] for the C-terminal His-tag construct. Iterative model building and refinement were performed using Coot [26] and Phenix.refine. Both tag-free crystals contained 2 molecules per asymmetric unit, and strict non-crystallographic symmetry (NCS) restraints were applied during early stages of the refinement, but they were released at the later stages. For the CtCut^S136A/pNP complex, strict NCS restraints were similarly applied and subsequently released during refinement to accommodate the 8 molecules per asymmetric unit. Structure figures were prepared using PyMOL (version 3.1.5.1; Schrödinger, LLC, New York, NY, USA) and Discovery Studio 2025 (version 25.1.0.24284; Dassault Systèmes BIOVIA, San Diego, CA, USA). The atomic coordinates and structure factors have been deposited in the Protein Data Bank under the accession codes 24KD (WT apo), 24KG (S136A apo), and 24KH (S136A/pNP complex). Detailed data collection and refinement statistics are summarized in Table 1.

3. Results

3.1. Construct Optimization, Expression, and Crystallization

CtCut proved to be a difficult-to-express protein in E. coli. Constructs encoding the full-length protein (CtCut^ΔM1) or the version lacking the signal peptide (CtCut^ΔM1–V19) without fusion tags yielded no detectable target protein upon IPTG induction. While the addition of a PhoA signal peptide (PhoA–CtCut^ΔM1–V19) enabled protein expression, a large proportion of the protein remained insoluble. In contrast, N-terminal fusion with Thioredoxin (TrxA) dramatically improved both the expression and soluble recovery of the protein (Figure 1A–C). This is consistent with TrxA’s known ability to act as a covalently linked “molecular chaperone,” effectively circumventing inclusion body formation in the bacterial cytoplasm [27]. Furthermore, extracting the cells with BugBuster reagent significantly increased the recovery of the target protein compared to conventional sonication (Figure 1D,E).

Co-crystallization of the His-tagged CtCut^S136A with ME-600 yielded P2₁ crystals with eight molecules per asymmetric unit, in which the His-tag participated in intermolecular crystal contacts (Table 1). To complement this, tag-free CtCut^WT and CtCut^S136A were crystallized in space group C2 with two molecules per asymmetric unit (Table 1). Data collection and refinement statistics for all three structures are summarized in Table 1.

3.2. Thermostability of CtCut^WT

Preliminary assays confirmed that CtCut exhibits esterase activity, and that substitution of Ser136 abolishes catalytic activity, consistent with its role as the catalytic nucleophile. The thermal stability of CtCut^WT was evaluated by differential scanning calorimetry (DSC) between 30 and 100 °C (Figure 2). The enzyme exhibited a gradual endothermic transition starting at approximately 60 °C, followed by a rapid transition from 65 °C onward. The thermogram was best fit by an equilibrium three-state model (N ⇌ I ⇌ D), as judged by the residuals and χ² values, consistent with the reversible unfolding framework established by Tamura and Sturtevant [19,20,21]. The first transition from the native (N) to an intermediate (I) state occurred with a Tm of 66.4 ± 0.08 °C, closely followed by a second transition to the fully denatured (D) state at 69.5 ± 0.01 °C. These two closely spaced but distinct Tm values indicate the sequential destabilization of structurally independent subdomains within the enzyme.

3.3. Overall Structure and Electrostatic Pre-Organization of the Apo-State

In the tag-free CtCut^WT and CtCut^S136A apo-structures, the enzyme adopts a highly stable monomeric α/β-hydrolase fold with a closed “lid” loop (Figure 3A). Superposition of Chain A and Chain B in the WT structure yielded an RMSD of 0.109 Å over 2184 atoms, confirming near-identical conformations (Figure 3B). Notably, the catalytic triad residue H204 resides 13.2 Å away from S136, fully exposed to the solvent in an inactive loop conformation (Figure 3B, inset). During refinement, a highly coordinated spherical electron density was observed near the active site in both the WT and S136A apo-structures. Despite the absence of a substrate, distance analyses revealed this density to be a chloride ion (Cl⁻) robustly coordinated by A57, S58, N100, Q137, F199, and P202 (distances 2.3–3.7 Å) with an additional contribution from the side chain of S136 in the WT structure (replaced by A136 in the S136A mutant) (Figure 3C). Notably, the chloride ion was observed in both structures, indicating that the electrostatic pre-organization of the oxyanion hole is maintained independently of the catalytic serine. The ability to strongly capture this solvent-derived anion confirms that the apo-enzyme maintains a pre-organized, positively charged electrostatic microenvironment. Comparison of the CtCut^WT and CtCut^S136A structures (RMSD = 0.256 Å over 2547 atoms) reveals conformational heterogeneity of the H204-containing lid loop (Figure 3D).

3.4. Structural Dynamics upon Ligand Accommodation

To understand how the enzyme responds to substrate analogs, we compared the apo-structure with the 2.65 Å structure of the S136A mutant/inhibitor complex (8 molecules/ASU) (Figure 4A). Here, an unambiguous extra electron density corresponding to p-nitrophenol (p-NP) which is thought to be incorporated after spontaneous degradation of the ME-600 inhibitor was observed (Figure 4B). Notably, the p-NP molecule directly displaced the aforementioned Cl⁻ ion, establishing robust hydrogen bonds with S58, N100, and Q137 to form the characteristic “oxyanion hole” network (Figure 4C). Triggered by this specific local binding, the lid loop transitioned to an open conformation. Superposition of the eight molecules revealed pronounced conformational heterogeneity in the loop region containing H204 (Leu195–Phe205), with the histidine side chain adopting distinctly different orientations in each chain.

3.5. Structural Comparison with Homologous Cutinases

To contextualize these conformational states, we compared CtCut with characterized fungal cutinases (Figure 5A). CtCut shares moderate sequence identity (55–69%) with Fusarium solani cutinase (FsCut) and other homologous enzymes, including Humicola insolens cutinase (HiCut), Glomerella cingulata cutinase (GcCut), and Aspergillus oryzae cutinase (AoCut). While the overall folds are highly conserved, critical differences emerge in the conformation of the lid loop (Figure 5B). In catalytically active cutinases such as FsCut, AoCut and HiCut, this region forms a short α-helix, directing the imidazole side chain toward the active serine (~4 Å distance) to complete the catalytic triad [28,29]. Conversely, in the apo-structures of CtCut and Glomerella cingulata cutinase [16,30], this region adopts an extended loop conformation, orienting the histidine away from the active site (~13 Å distance) (Figure 5C). This structural dichotomy is consistent with the conformational cycling model proposed for GcCut [16,30], in which the lid loop must undergo a loop-to-helix transition to position the catalytic histidine for triad formation.

4. Discussion

4.1. Complementary Structural Insights from Tagged and Tag-Free Constructs

Affinity tags can influence crystal packing and potentially obscure intrinsic conformational features [17]. In the present study, the C-terminal His-tag in the S136A/inhibitor complex participates in intermolecular contacts, while the tag-free constructs yielded a simpler crystal form. The inactive conformation of H204 was consistently observed in both tagged and tag-free structures, confirming that this configuration is an inherent property of the CtCut lid loop rather than an artifact of crystal packing. AI-based structure prediction tools such as AlphaFold3 [24] were useful for molecular replacement; however, the conformational heterogeneity of H204 and the coordinated chloride ion represent local structural features that extend beyond a single predicted conformation, underscoring the continued importance of experimental structural biology.

4.2. Active-Site Architecture and Conformational Cycling

The apo-structures reveal a preformed oxyanion hole occupied by a chloride ion, consistent with the electrostatic pre-organization described for FsCut [14]. In the S136A/inhibitor complex, the Cl⁻ is displaced by p-NP, accompanied by lid loop opening and increased H204 heterogeneity, consistent with the conformational cycling model proposed by Nyon et al. for GcCut [16]. Crucially, they demonstrated through H204N mutagenesis that this residue is essential for catalysis despite its large distance from the active-site serine in the resting state [16]. Analogous mutagenesis of H204 in CtCut would be a valuable future experiment to confirm this requirement in a thermophilic context. The dynamic behavior of the lid loop may facilitate accommodation of bulky polymer substrates, as proposed for other polyester-degrading enzymes [12].

4.3. Thermodynamic Decoupling of Global Stability and Local Flexibility

The three-state unfolding profile (Section 3.2) suggests the presence of structurally distinct regions with different thermal stabilities [19,20,21]. We hypothesize that the first transition reflects destabilization of the lid loop region, which exhibits the highest B-factors and greatest conformational variability, while the second transition corresponds to global unfolding of the α/β-hydrolase core. This thermodynamic decoupling, in which local flexibility is maintained within an overall thermostable framework, may be functionally relevant, although direct verification by hydrogen–deuterium exchange mass spectrometry would be required to map regional unfolding.

4.4. Functional Role of Local Flexibility: Comparison with PET Hydrolases

The structural heterogeneity observed in the H204-containing lid loop provides insight into the functional role of local flexibility in CtCut. In Ideonella sakaiensis PETase (IsPETase), flexibility of the active-site W-loop, particularly involving Trp185, has been implicated in substrate accommodation [12,32]. While the specific residues differ, both systems exhibit conformational variability near the active site. Our results suggest a possible functional analogy, in which local structural flexibility adjacent to a preformed oxyanion hole contributes to substrate recognition and positioning. Although the precise mechanistic roles of these flexible elements may differ between enzymes, the presence of such dynamic regions appears to be a recurring feature among polyester hydrolases.

4.5. Implications for Understanding Cutinase Diversity

Recent PET hydrolase engineering has highlighted the importance of balancing global stability with local flexibility [6,7,8,9,10]. CtCut exemplifies this balance structurally: a rigid α/β-hydrolase core coupled with a dynamic lid loop. For future engineering, stabilizing mutations may be most effectively introduced into the global scaffold while preserving flexibility in the active-site region.

5. Conclusions

These crystal structures of a thermophilic cutinase from C. thermophilum provide three key structural insights. First, the apo-structures reveal a preformed oxyanion hole occupied by a chloride ion, demonstrating electrostatic pre-organization independent of the catalytic serine. Second, the ligand-bound complex captures pronounced conformational heterogeneity of the catalytic H204, consistent with the conformational cycling model established for G. cingulata cutinase [16]. Third, the DSC data suggest thermodynamic decoupling between the rigid core and flexible lid loop. Together, these findings expand the structural diversity of characterized cutinase architectures and may inform future engineering of thermostable polyester hydrolases.

Author Contributions

Conceptualization, T.N. and S.I.; Methodology, R.N., L.C., M.K., S.I. and T.N.; Validation, R.N., L.C., M.K., S.I. and T.N.; Formal Analysis, R.N., L.C., M.K., S.I. and T.N.; Investigation, R.N., L.C., M.K., S.I. and T.N.; Resources, T.N. and S.I.; Data Curation, T.N. and S.I.; Writing—Original Draft, R.N., L.C. and T.N.; Writing—Review and Editing, T.N. and S.I.; Supervision, T.N. and S.I.; Funding Acquisition, T.N. All authors have read and agreed to the published version of the manuscript.

Funding

This work was partly supported by the Japan Society for the Promotion of Science (JSPS) KAKENHI under Grant Numbers 20K06512 and 23K05671 (to T.N.).

Data Availability Statement

The atomic coordinates have been deposited in the PDB with accession numbers 24KD, 24KG, and 24KH.

Acknowledgments

We thank Ed Hurt for the generous gift of Chaetomium thermophilum cDNA and Minako Maeda for her kind assistance in DSC measurement. The Synchrotron experiments were performed under the approval of the Photon factory Program Advisory Committee (Proposal Nos 2020G099, 2022G118, 2024G111) and diffraction data were collected at BL1A. This work was carried out in part using the Joint Usage/Research Program of Institute for Protein Research, Osaka University (Project No.CR-21-35, CR-22-37, CR-23-25, CR24-7). During the preparation of this manuscript, the authors used ChatGPT (OpenAI), Gemini (Google), and Claude (Anthropic) for language refinement and suggestions. The authors have reviewed and edited all outputs and take full responsibility of the content.

Conflicts of Interest

The authors declare no conflicts of interest.

Abbreviations

The following abbreviations are used in this manuscript:

PET	poly(ethylene terephthalate)
LCC	leaf–branch compost cutinase
DSC	differential scanning calorimetry
LB	Luria–Bertani
pNP	p-nitrophenol
WT	Wild-type

References

Kolattukudy, P.E. Polyesters in Higher Plants. Adv. Biochem. Eng. Biotechnol. 2001, 71, 1–49. [Google Scholar] [CrossRef] [PubMed]
Sulaiman, S.; Yamato, S.; Kanaya, E.; Kim, J.-J.; Koga, Y.; Takano, K.; Kanaya, S. Isolation of a Novel Cutinase Homolog with Polyethylene Terephthalate-Degrading Activity from Leaf-Branch Compost by Using a Metagenomic Approach. Appl. Environ. Microbiol. 2012, 78, 1556–1562. [Google Scholar] [CrossRef]
Yoshida, S.; Hiraga, K.; Takehana, T.; Taniguchi, I.; Yamaji, H.; Maeda, Y.; Toyohara, K.; Miyamoto, K.; Kimura, Y.; Oda, K. A Bacterium That Degrades and Assimilates Poly (Ethylene Terephthalate). Science 2016, 351, 1196–1199. [Google Scholar] [CrossRef] [PubMed]
Austin, H.P.; Allen, M.D.; Donohoe, B.S.; Rorrer, N.A.; Kearns, F.L.; Silveira, R.L.; Pollard, B.C.; Dominick, G.; Duman, R.; El Omari, K.; et al. Characterization and Engineering of a Plastic-Degrading Aromatic Polyesterase. Proc. Natl. Acad. Sci. USA 2018, 115, E4350–E4357. [Google Scholar] [CrossRef] [PubMed]
Joo, S.; Cho, I.J.; Seo, H.; Son, H.F.; Sagong, H.-Y.; Shin, T.J.; Choi, S.Y.; Lee, S.Y.; Kim, K.-J. Structural Insight into Molecular Mechanism of Poly(Ethylene Terephthalate) Degradation. Nat. Commun. 2018, 9, 382. [Google Scholar] [CrossRef]
Tournier, V.; Topham, C.M.; Gilles, A.; David, B.; Folgoas, C.; Moya-Leclair, E.; Kamionka, E.; Desrousseaux, M.-L.; Texier, H.; Gavalda, S.; et al. An Engineered PET Depolymerase to Break down and Recycle Plastic Bottles. Nature 2020, 580, 216–219. [Google Scholar] [CrossRef]
Lu, H.; Diaz, D.J.; Czarnecki, N.J.; Zhu, C.; Kim, W.; Shroff, R.; Acosta, D.J.; Alexander, B.R.; Cole, H.O.; Zhang, Y.; et al. Machine Learning-Aided Engineering of Hydrolases for PET Depolymerization. Nature 2022, 604, 662–667. [Google Scholar] [CrossRef]
Bell, E.L.; Smithson, R.; Kilbride, S.; Foster, J.; Hardy, F.J.; Ramachandran, S.; Tedstone, A.A.; Haigh, S.J.; Garforth, A.A.; Day, P.J.R.; et al. Directed Evolution of an Efficient and Thermostable PET Depolymerase. Nat. Catal. 2022, 5, 673–681. [Google Scholar] [CrossRef]
Oda, K.; Wlodawer, A. Development of Enzyme-Based Approaches for Recycling PET on an Industrial Scale. Biochemistry 2024, 63, 369–401. [Google Scholar] [CrossRef]
Kawai, F.; Iizuka, R.; Kawabata, T. Engineered Polyethylene Terephthalate Hydrolases: Perspectives and Limits. Appl. Microbiol. Biotechnol. 2024, 108, 404. [Google Scholar] [CrossRef]
Kawai, F.; Kawabata, T.; Oda, M. Current Knowledge on Enzymatic PET Degradation and Its Possible Application to Waste Stream Management and Other Fields. Appl. Microbiol. Biotechnol. 2019, 103, 4253–4268. [Google Scholar] [CrossRef]
Fecker, T.; Galaz-Davison, P.; Engelberger, F.; Narui, Y.; Sotomayor, M.; Parra, L.P.; Ramírez-Sarmiento, C.A. Active Site Flexibility as a Hallmark for Efficient PET Degradation by I. sakaiensis PETase. Biophys. J. 2018, 114, 1302–1312. [Google Scholar] [CrossRef] [PubMed]
Martinez, C.; De Geus, P.; Lauwereys, M.; Matthyssens, G.; Cambillau, C. Fusarium solani Cutinase Is a Lipolytic Enzyme with a Catalytic Serine Accessible to Solvent. Nature 1992, 356, 615–618. [Google Scholar] [CrossRef] [PubMed]
Martinez, C.; Nicolas, A.; van Tilbeurgh, H.; Egloff, M.P.; Cudrey, C.; Verger, R.; Cambillau, C. Cutinase, a Lipolytic Enzyme with a Preformed Oxyanion Hole. Biochemistry 1994, 33, 83–89. [Google Scholar] [CrossRef] [PubMed]
Nicolas, A.; Egmond, M.; Verrips, C.T.; de Vlieg, J.; Longhi, S.; Cambillau, C.; Martinez, C. Contribution of Cutinase Serine 42 Side Chain to the Stabilization of the Oxyanion Transition State. Biochemistry 1996, 35, 398–410. [Google Scholar] [CrossRef]
Nyon, M.P.; Rice, D.W.; Berrisford, J.M.; Hounslow, A.M.; Moir, A.J.G.; Huang, H.; Nathan, S.; Mahadi, N.M.; Bakar, F.D.A.; Craven, C.J. Catalysis by Glomerella cingulata Cutinase Requires Conformational Cycling between the Active and Inactive States of Its Catalytic Triad. J. Mol. Biol. 2009, 385, 226–235. [Google Scholar] [CrossRef]
Majorek, K.A.; Kuhn, M.L.; Chruszcz, M.; Anderson, W.F.; Minor, W. Double Trouble—Buffer Selection and His-Tag Presence May Be Responsible for Nonreproducibility of Biomedical Experiments. Protein Sci. 2014, 23, 1359–1368. [Google Scholar] [CrossRef]
Amlacher, S.; Sarges, P.; Flemming, D.; van Noort, V.; Kunze, R.; Devos, D.P.; Arumugam, M.; Bork, P.; Hurt, E. Insight into Structure and Assembly of the Nuclear Pore Complex by Utilizing the Genome of a Eukaryotic Thermophile. Cell 2011, 146, 277–289. [Google Scholar] [CrossRef]
Tamura, A.; Sturtevant, J.M. A Thermodynamic Study of Mutant Forms of Streptomyces Subtilisin Inhibitor. I. Hydrophobic Replacements at the Position of Met103. J. Mol. Biol. 1995, 249, 625–635. [Google Scholar] [CrossRef]
Tamura, A.; Kojima, S.; Miura, K.; Sturtevant, J.M. A Thermodynamic Study of Mutant Forms of Streptomyces Subtilisin Inhibitor. II. Replacements at the Interface of Dimer Formation, Val13. J. Mol. Biol. 1995, 249, 636–645. [Google Scholar] [CrossRef]
Tamura, A.; Sturtevant, J.M. A Thermodynamic Study of Mutant Forms of Streptomyces Subtilisin Inhibitor. III. Replacements of a Hyper-Exposed Residue, Met73. J. Mol. Biol. 1995, 249, 646–653. [Google Scholar] [CrossRef] [PubMed]
Kabsch, W. XDS. Acta Crystallogr. D Biol. Crystallogr. 2010, 66, 125–132. [Google Scholar] [CrossRef] [PubMed]
Liebschner, D.; Afonine, P.V.; Baker, M.L.; Bunkóczi, G.; Chen, V.B.; Croll, T.I.; Hintze, B.; Hung, L.-W.; Jain, S.; McCoy, A.J.; et al. Macromolecular Structure Determination Using X-Rays, Neutrons and Electrons: Recent Developments in Phenix. Acta Crystallogr. Sect. Struct. Biol. 2019, 75, 861–877. [Google Scholar] [CrossRef] [PubMed]
Abramson, J.; Adler, J.; Dunger, J.; Evans, R.; Green, T.; Pritzel, A.; Ronneberger, O.; Willmore, L.; Ballard, A.J.; Bambrick, J.; et al. Accurate Structure Prediction of Biomolecular Interactions with AlphaFold 3. Nature 2024, 630, 493–500. [Google Scholar] [CrossRef]
Jumper, J.; Evans, R.; Pritzel, A.; Green, T.; Figurnov, M.; Ronneberger, O.; Tunyasuvunakool, K.; Bates, R.; Žídek, A.; Potapenko, A.; et al. Highly Accurate Protein Structure Prediction with AlphaFold. Nature 2021, 596, 583–589. [Google Scholar] [CrossRef]
Emsley, P.; Lohkamp, B.; Scott, W.G.; Cowtan, K. Features and Development of Coot. Acta Crystallogr. D Biol. Crystallogr. 2010, 66, 486–501. [Google Scholar] [CrossRef]
LaVallie, E.; DiBlasio, E.; Kovacic, S.; Grant, K.; Schendel, P.; McCoy, J. A Thioredoxin Gene Fusion Expression System That Circumvents Inclusion Body Formation in the E. coli Cytoplasm. Nat. Biotechnol. 1993, 11, 187–193. [Google Scholar] [CrossRef]
Liu, Z.; Gosser, Y.; Baker, P.J.; Ravee, Y.; Lu, Z.; Alemu, G.; Li, H.; Butterfoss, G.L.; Kong, X.-P.; Gross, R.; et al. Structural and Functional Studies of Aspergillus oryzae Cutinase: Enhanced Thermostability and Hydrolytic Activity of Synthetic Ester and Polyester Degradation. J. Am. Chem. Soc. 2009, 131, 15711–15716. [Google Scholar] [CrossRef]
Kold, D.; Dauter, Z.; Laustsen, A.K.; Brzozowski, A.M.; Turkenburg, J.P.; Nielsen, A.D.; Koldsø, H.; Petersen, E.; Schiøtt, B.; De Maria, L.; et al. Thermodynamic and Structural Investigation of the Specific SDS Binding of Humicola insolens Cutinase. Protein Sci. 2014, 23, 1023–1035. [Google Scholar] [CrossRef]
Nyon, M.P.; Rice, D.W.; Berrisford, J.M.; Huang, H.; Moir, A.J.G.; Craven, C.J.; Nathan, S.; Mahadi, N.M.; Farah Diba, A.B. Crystallization and Preliminary X-Ray Analysis of Recombinant Glomerella cingulata Cutinase. Acta Crystallograph. Sect. F Struct. Biol. Cryst. Commun. 2008, 64, 504–508. [Google Scholar] [CrossRef]
Madeira, F.; Madhusoodanan, N.; Lee, J.; Eusebi, A.; Niewielska, A.; Tivey, A.R.N.; Lopez, R.; Butcher, S. The EMBL-EBI Job Dispatcher Sequence Analysis Tools Framework in 2024. Nucleic Acids Res. 2024, 52, W521–W525. [Google Scholar] [CrossRef]
Han, X.; Liu, W.; Huang, J.-W.; Ma, J.; Zheng, Y.; Ko, T.-P.; Xu, L.; Cheng, Y.-S.; Chen, C.-C.; Guo, R.-T. Structural Insight into Catalytic Mechanism of PET Hydrolase. Nat. Commun. 2017, 8, 2106. [Google Scholar] [CrossRef]

Figure 1. Optimization of CtCut expression constructs and solubility. (A–C) SDS-PAGE analysis of various CtCut expression constructs: (A) CtCutΔM1 and CtCutΔM1-V19 (without fusion tag), (B) PhoA-CtCutΔM1-V19, and (C) TrxA-CtCutΔM1-V19, demonstrating markedly improved soluble expression. (D,E) Comparison of protein extraction methods for TrxA-CtCutΔM1-V19 using (D) sonication and (E) BugBuster extraction reagent. All SDS-PAGE analyses were performed on 14% Tris-Glycine gels (20 mA, 70 min). Lanes are designated as follows: M, molecular weight marker; U, uninduced whole-cell lysate; I, IPTG-induced whole-cell lysate; S, soluble fraction; P, insoluble (pellet) fraction. Black arrowheads indicate the target protein bands. Molecular weight markers (kDa) are shown on the left of each gel, with selected marker positions highlighted in red and green.

Figure 2. Differential scanning calorimetry of CtCut^WT. Baseline-corrected excess specific heat capacity of CtCut^WT (red line) with three-state model fit (blue circles). The vertical dashed lines indicate the transition temperatures: Tm(N ⇌ I) = 66.4 ± 0.08 °C and Tm(I ⇌ D) = 69.5 ± 0.01 °C. The N (native), I (intermediate), and D (denatured) phases are labeled.

Figure 3. Crystal structure of WT CtCut and comparison with the S136A mutant. (A) Ribbon representation of the two WT CtCut molecules in the asymmetric unit (Chain A, orange; Chain B, cyan). (B) Superposition of Chain A (orange) and Chain B (cyan), with an inset showing the active site region where the catalytic triad residues S136, D191, and H204 are displayed as sticks. Note that H204 adopts a solvent-exposed, inactive loop conformation 13.2 Å from S136. (C) Close-up view of the chloride ion (Cl−) coordination near the active site. The Cl− (yellow sphere) is coordinated by backbone and side-chain atoms of A57, S58, N100, S136, Q137, F199 and P202. The protein is shown in cyan, and the interacting residues are represented as sticks. (D) Superposition of CtCut^WT (cyan) and CtCut^S136A (magenta) apo-structures, with an inset showing the active site where H204 electron density is less well-defined in S136A, indicating increased local mobility.

Figure 4. Crystal structure of the CtCut^S136A/p-NP complex at 2.65 Å resolution (P2₁, 8 molecules/ASU). (A) Ribbon representation of the eight CtCut^S136A molecules in the asymmetric unit (P2₁, 2.65 Å). Chains are individually colored: A (green), B (cyan), C (magenta), D (yellow), E (salmon), F (gray), G (purple), H (orange). (B) Superposition of the eight molecules highlighting the conformational heterogeneity of the lid loop (Leu195–Phe205) and the bound p-NP at the oxyanion hole. Each structure is shown in a different color. (C) Active site detail showing the catalytic residues (A136, Asp191, His204) and the oxyanion hole residues (Ser58, Asn100, Gln137) accommodating p-NP. Note the pronounced conformational heterogeneity of His204 among the eight molecules, while the positions of A136 and Asp191 remain conserved. The eight molecules are represented in different colors.

Figure 5. Structural comparison of CtCut with homologous fungal cutinases. (A) Overall structural superposition of CtCut^S136A (light blue) with Fusarium solani cutinase (FsCut, PDB: 1CUS; blue), Humicola insolens cutinase (HiCut, PDB: 4OYY; orange), Glomerella cingulata cutinase (GcCut, PDB: 3DCN; purple), and Aspergillus oryzae cutinase (AoCut, PDB: 3GBS; yellow-green). (B) Enlarged view of the lid loop region (dashed box in (A)), showing the divergent conformations among the five cutinases. (C) Active site comparison showing the catalytic triad residues as sticks with backbone as wireframe. Note the distinct positioning of the catalytic histidine: the “active/helix” conformation in FsCut and HiCut (~3–4 Å from catalytic Ser) versus the “inactive/loop” conformation in CtCut and GcCut (~13 Å from Ser). p-NP was omitted for clarity. (D) Sequence alignment of five fungal cutinases. Sequences were aligned using the Clustal Omega server (EMBL-EBI, https://www.ebi.ac.uk/Tools/msa/clustalo/, accessed on 16 March 2026) [31]. The blue shading reflects the degree of sequence identity, with darker blue representing higher conservation. The catalytic triad residues (Ser, Asp, and His) are highlighted in red and marked with red dots below the sequence. Residues involved in the formation of the oxyanion hole are indicated by blue diamonds. The lid loop is indicated by dotted box. The numbers above each alignment block correspond to the amino acid positions of CtCut(33-225). Secondary structure elements are indicated as follows: red cylinders represent α-helices, and green arrows represent β-strands. Numbers on the left and right indicate the residue numbers of each protein.

Table 1. Data collection and refinement statistics.

Name and PDBID	Apo CtCut^WT 24KD	Apo CtCut^S136A 24KG	CtCut^S136A-pNP Complex 24KH
Resolution range (Å)	44.29–1.7 (1.73–1.7)	41.34–1.7 (1.73–1.7)	47.84–2.65 (2.7–2.65)
Space group	C2	C2	P2₁
Unit cell (Å, °)	74.033 88.576 56.825 90 94.197 90	75.13 88.248 56.685 90 94.165 90	80.921 104.516 95.709 90 91.239 90
Total reflections	138,869 (5476)	138,945 (5353)	160,614 (8355)
Unique reflections	40,068 (1963)	40,275 (1889)	46,250 (2346)
Multiplicity	3.5 (2.8)	3.4 (2.8)	3.5 (3.6)
Completeness (%)	96.11 (88.89)	95.54 (91.99)	95.10 (94.47)
Mean I/sigma (I)	6.26 (1.16)	6.59 (1.91)	6.78 (1.07)
Wilson B-factor (Å²)	17.23	16.77	51.99
R-merge	0.1366 (0.7305)	0.1147 (0.4424)	0.1435 (1.11)
R-meas	0.1617 (0.9053)	0.1358 (0.54)	0.17 (1.309)
R-pim	0.08559 (0.5265)	0.07188 (0.3056)	0.09028 (0.6883)
CC1/2	0.991 (0.579)	0.989 (0.844)	0.992 (0.517)
CC*	0.998 (0.856)	0.997 (0.957)	0.998 (0.825)
Reflections used in refinement	40,055 (1963)	40,267 (1889)	46,238 (2345)
Reflections used for R-free	1549 (81)	1557 (73)	1991 (99)
R-work	0.1573 (0.3305)	0.1875 (0.3445)	0.2020 (0.3470)
R-free	0.1992 (0.3705)	0.2223 (0.3893)	0.2111 (0.3530)
Number of non-hydrogen atoms	3527	3261	11,870
macromolecules	2872	2870	11,644
ligands	2	2	80
solvent	653	389	146
Protein residues	386	386	1560
RMS(bonds) (Å)	0.006	0.005	0.009
RMS(angles) (°)	0.83	0.78	1.01
Ramachandran favored (%)	96.86	96.34	94.75
Ramachandran allowed (%)	3.14	3.66	5.18
Ramachandran outliers (%)	0.00	0.00	0.06
Rotamer outliers (%)	0.00	0.66	2.76
Clashscore	2.98	1.75	8.71
Average B-factor (Å²)	22.20	23.12	64.03
macromolecules	19.66	22.21	64.19
ligands	22.93	28.16	65.13
solvent	33.35	29.78	50.70

Statistics for the highest-resolution shell are shown in parentheses.

Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Nojima, R.; Chen, L.; Kurokawa, M.; Ito, S.; Nishino, T. Crystal Structures of a Thermophilic Cutinase from Chaetomium thermophilum Reveal Conformational Dynamics of the Catalytic Lid Loop. Crystals 2026, 16, 217. https://doi.org/10.3390/cryst16040217

AMA Style

Nojima R, Chen L, Kurokawa M, Ito S, Nishino T. Crystal Structures of a Thermophilic Cutinase from Chaetomium thermophilum Reveal Conformational Dynamics of the Catalytic Lid Loop. Crystals. 2026; 16(4):217. https://doi.org/10.3390/cryst16040217

Chicago/Turabian Style

Nojima, Ryohei, Lirong Chen, Minami Kurokawa, Sho Ito, and Tatsuya Nishino. 2026. "Crystal Structures of a Thermophilic Cutinase from Chaetomium thermophilum Reveal Conformational Dynamics of the Catalytic Lid Loop" Crystals 16, no. 4: 217. https://doi.org/10.3390/cryst16040217

APA Style

Nojima, R., Chen, L., Kurokawa, M., Ito, S., & Nishino, T. (2026). Crystal Structures of a Thermophilic Cutinase from Chaetomium thermophilum Reveal Conformational Dynamics of the Catalytic Lid Loop. Crystals, 16(4), 217. https://doi.org/10.3390/cryst16040217

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Menu

Crystal Structures of a Thermophilic Cutinase from Chaetomium thermophilum Reveal Conformational Dynamics of the Catalytic Lid Loop

Abstract

1. Introduction