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. Author manuscript; available in PMC: 2021 Dec 1.
Published in final edited form as: Carbohydr Polym. 2021 Aug 19;273:118564. doi: 10.1016/j.carbpol.2021.118564

Rational enzyme design for controlled functionalization of acetylated xylan for cell-free polymer biosynthesis

Hsin-Tzu Wang a,b,1, Vivek S Bharadwaj c,1, Jeong-Yeh Yang a,b,2, Thomas M Curry a,b, Kelley W Moremen a,b, Yannick J Bomble d, Breeanna R Urbanowicz a,b,*
PMCID: PMC8506904  NIHMSID: NIHMS1745952  PMID: 34560975

Abstract

Xylan O-acetyltransferase 1 (XOAT1) is involved in O-acetylating the backbone of hemicellulose xylan. Recent structural analysis of XOAT1 showed two unequal lobes forming a cleft that is predicted to accommodate and position xylan acceptors into proximity with the catalytic triad. Here, we used docking and molecular dynamics simulations to investigate the optimal orientation of xylan in the binding cleft of XOAT1 and identify putative key residues (Gln445 and Arg444 on Minor lobe & Asn312, Met311 and Asp403 on Major lobe) involved in substrate interactions. Site-directed mutagenesis coupled with biochemical analyses revealed the major lobe of XOAT1 is important for xylan binding. Mutation of single key residues yielded XOAT1 variants with various enzymatic efficiencies that are applicable to one-pot synthesis of xylan polymers with different degrees of O-acetylation. Taken together, our results demonstrate the effectiveness of computational modeling in guiding enzyme engineering aimed at modulating xylan and redesigning plant cell walls.

Keywords: Xylan O-acetylation, Xylooligosaccharide, Enzyme-substrate interaction, Computational modeling, Enzyme engineering, One-pot synthesis

1. Introduction

Acetylation of biomolecules is an important chemical modification employed in nature observed across all domains of life. These modifications have significant implications to myriad biological functions and self-assembly of molecular architectures in biopolymers, including proteins, polysaccharides and polyphenolics. Plant secondary cell walls, which constitute the majority of the primary biomass resources harvested for valorization into fuels, chemicals, textiles and materials, are principally composed of cellulose, hemicelluloses, and lignin (Carpita & McCann, 2020). Xylan is the most abundant hemicellulose in these walls and on Earth, making up to 20–30% of the dry weight of most secondary cell walls (Scheller & Ulvskov, 2010). Glucuronoxylan present in dicots consists of a backbone that is formed by 1,4-linked β-d-xylopyranosyl (Xyl) residues that is further substituted by 1,2-linked α-d-glucuronic acid (GlcA) and/or its 4-O-methyl derivative (MeGlcA). In addition, more than half of the Xyl residues are O-acetylated at the O-2 or O-3 position, or can also be di-acetylated at both the O-2 and O-3 positions (Teleman, Lundqvist, Tjerneld, Stalbrand, & Dahlman, 2000; Teleman, Tenkanen, Jacobs, & Dahlman, 2002). The level of acetylation has been shown to affect the physical properties of polysaccharides such as hydrophobicity (Biely, 2012), may also interfere with the interaction between xylan and other cell wall polymers (Busse-Wicher et al., 2014), and perturbs various physiological functions, including freezing tolerance and pathogen resistance (Diener & Ausubel, 2005; Manabe et al., 2011; Urbanowicz, Peña, Moniz, Moremen, & York, 2014; Vogel, Raab, Somerville, & Somerville, 2004; Xin, Mandaokar, Chen, Last, & Browse, 2007).

Recent advances using solid-state nuclear magnetic resonance (NMR) spectroscopy have begun to reveal detailed molecular-level information about the organization of the supramolecular networks formed and the roles specific polymer structures play within the 3D architecture of secondary cell walls (Kang et al., 2019; Simmons et al., 2016). These groundbreaking studies indicate that xylan plays a more important role in the wall than previously thought, and the O-acetyl moieties along the xylan backbone form intermolecular interactions with lignin aromatics or cellulose microfibrils in a conformation dependent manner. Using mass spectrometry and NMR, it has been observed that substituents are evenly attached to the xylan backbone in an alternating pattern in Arabidopsis, suggesting that this xylan glycoform adopts a twofold helical screw conformation (a 360° twist every two glycosidic bonds) where the unsubstituted side forms hydrogen bonds with the hydrophilic surface of cellulose microfibrils (Duan et al., 2021; Gao, Lipton, Wittmer, Murray, & Mortimer, 2020; Kang et al., 2019). This hypothetical model was later experimentally substantiated by solid-state magic-angle spinning (MAS) NMR spectroscopy of fresh Arabidopsis stems, and supported by in silico predictions, which reveal an induced assembly of xylan on cellulose fibrils as a rigid twofold screw from a flexible threefold conformation (a 360° twist every three glycosidic bonds) in solution (Simmons et al., 2016). Moreover, a recent study of acetylated xylan rearrangement via unrestrained molecular dynamics simulations further indicates that not only evenly spaced substituents but also the position on xylosyl residues play key roles in tuning xylan-cellulose interactions. It has been shown that 2-O-acetylated xylan is required to trigger the transition from a threefold to twofold screw conformation on cellulose surface through specific O–O contacts (Gupta, Rawal, Dupree, Smith, & Petridis, 2021), while 3-O-acetylated xylan maintains a threefold screw that preferentially binds lignin and only interacts weakly with cellulose (Gupta, Rawal, Dupree, Smith, & Petridis, 2021; Kang et al., 2019). Taken together, these data suggest that O-acetyl substituents along the backbone may be the one of the key chemical determinants of xylan confirmation, and thus lignin-cellulose packing, in muro (Duan et al., 2021; Gao, Lipton, Wittmer, Murray, & Mortimer, 2020; Kang et al., 2019).

From an industrial perspective, acetic acid derived from innate acetylation of lignocellulosic materials could have both positive and negative impacts on biomass conversion. During biomass pretreatment processes, it has been shown that acetic acid itself could be effectively used for delignification (Simon et al., 2014; Young, Davis, & Wiesmann, 1986) and fractionation of cell wall components (Parajó, Alonso, & Santos, 1995; Vásquez, Lage, Parajó, & Vázquez, 1992). A recent study further shows that acetic acid that is liberated from acetylated cell wall polysaccharides in Populus trichocarpa is able to catalyze the breakdown of wood and thus increase sugar release during biomass pretreatment (Johnson, Kim, Ralph, & Mansfield, 2017). In addition to that, the downstream fermentative microorganisms were shown to be inhibited by the released acetic acid (Larsson et al., 1999; Olsson & Hahn-Hägerdal, 1996).

Overall, understanding the process of plant polysaccharide acetylation at the molecular level is essential not only to elucidate this crucial modification, but also to promote our ability to develop targeted genomics or genome engineering approaches to design plant cell walls that can be more efficiently valorized into food, materials and products. Nine members of the TBL family in Arabidopsis have been shown to possess xylan O-acetyltransferase activities, and differ in their regiospecificity for the O-2 and/or O-3 positions on xylosyl residues (Yuan et al., 2016; Yuan, Teng, Zhong, & Ye, 2013, 2015; Yuan, Teng, Zhong, & Ye, 2016; Zhong, Cui, & Ye, 2017). Xylan O-acetyltransferase 1 (XOAT1) has been shown to specifically catalyze the 2-O-acetylation of xylan using an NMR spectroscopy technique that allows the reaction to be monitored in real time (Lunin et al., 2020; Urbanowicz, Peña, Moniz, Moremen, & York, 2014), diminishing the ambiguity caused by non-enzymatic acetyl migration observed in end-point analyses (Brecker, Mahut, Schwarz, & Nidetzky, 2009; Lassfolk et al., 2019; Mastihubová & Biely, 2004; Roslund et al., 2008; Yoshimoto & Tsuda, 1983). The esk1 mutant is a null mutant in the XOAT1 gene that has collapsed xylem vessels, an ~60% reduction of xylan O-acetylation (Lefebvre et al., 2011; Xiong, Cheng, & Pauly, 2013), and is more tolerant to freezing, drought and salt stress (Bouchabke-Coussa et al., 2008; Xin & Browse, 1998). Further, it has been shown that the even-pattern of acetyl and Me(GlcA) substituents on alternate xylosyl residues along the backbone is disrupted in esk1 mutants, suggesting a role of XOAT1 in patterning xylan decorations (Grantham et al., 2017). However, it is unclear how polysaccharide O-acetyltransferases interact with specific acceptor substrates to potentially control acetyl distribution.

A recent crystallographic study of XOAT1 revealed a heart shaped enzyme formed by two unequal lobes, referred to as major and minor, with a catalytic triad formed by Ser216-His465-Asp462 localized at the bottom of the cleft (Lunin et al., 2020). One of the key challenges in gaining insight into the enzymatic mechanisms of plant biopolymer synthesis and modification in general, and in XOAT1 in particular, is the paucity of experimentally characterized enzyme-substrate co-crystal structures. Molecular modeling techniques involving docking and molecular dynamics (MD) simulations have been effective in successfully bridging this crucial gap (Lunin et al., 2020; Urbanowicz et al., 2017). In our previous study, a combination of biochemical analyses and molecular simulations indicated that XOAT1 utilizes a double displacement/Ping Pong Bi Bi mechanism (Lunin et al., 2020). In addition to revealing the root mean square fluctuations of the various domains of the protein, the molecular simulations also enabled the identification of donor and substrate binding sites on XOAT1. While the substrate bound simulations revealed that the active site and substrate binding groove were able to stabilize the bound states of XOAT1 over timescales relevant for the catalytic mechanism, identification of the specific residues involved in substrate binding remained to be explored. In this study, we extend our analyses to investigate key residues involved in enzyme-acceptor substrate interactions through a combination of computational modeling followed by site-directed mutagenesis (SDM), kinetic analysis, and in vitro synthesis of O-acetylated xylan. Taken together, our results show that the major lobe of XOAT1 interacts more with the substrate, and two of the major lobe residues, M311 and D403, are crucial for hydrolysis of the acetyl donors to form the acyl-enzyme intermediate.

The use of computational techniques greatly speeds up the exploration of protein design space, facilitating the identification of key residues for protein engineering. We envisage that our study lays the foundations to provide insight into the relationships between the conformation of TBL proteins and their corresponding substrate specificities, allowing catalytic parameters to be precisely modulated to synthesize xylan with different degrees of O-acetylation and/or substitution patterns. Understanding the molecular levers that can be used to control a key modification such as polysaccharide O-acetylation opens the door towards new synthetic biology strategies aimed at fine tuning 3-D cell wall architecture. Post-synthetic modification of plant cell walls through in muro expression of acetyl xylan esterases (AXE) has been shown to reduce biomass recalcitrance without perturbing plant growth, the obtained transgenic lines exhibited reduced xylan acetylation and are more susceptible to β−1,4-endoxylanase during saccharification, which further results in increased sugar yields (Pawar et al., 2016; Pawar et al., 2017; Wang et al., 2020). In addition to modification of cell walls via post-synthesis, our research provides candidates for direct replacement of synthetic genes involved in pathways of xylan acetylation to improve lignocellulosic materials for benefits in industrial applications including valorization into food, fuels, and chemicals.

2. Materials and methods

2.1. Molecular dynamics simulations

The recently published protein structure of XOAT1 (PDB ID: 6CCI) in its apo state was used to develop the donor (p-nitrophenyl acetate, pNP-Ac) and acceptor (xylodecaose) bound complexes of XOAT1. A combined docking and molecular dynamics protocol was employed to predict binding poses for donor-XOAT1 and acceptor-XOAT1 complexes. The detailed set-up for docking and molecular dynamics simulations of this complex are outlined in our previous publication (Lunin et al., 2020). For this publication, only simulations of the acceptor-bound XOAT1 complex were considered. Briefly, the production run consisted of an unbiased 50 ns trajectory of a fully solvated acceptor-bound (xylodecaose) XOAT1 complex. The MD trajectory was analyzed to quantify interaction energies between XOAT1 amino acid residues and the xylodecaose substrate using CHARMM (Brooks et al., 2009; Hynninen & Crowley, 2014) and VMD (Humphrey, Dalke, & Schulten, 1996). These per-residue interaction energies were used as the metric to rank all the amino-acid interactions stabilizing acceptor binding to XOAT1 and identify targets for experimental mutagenesis.

2.2. Fusion protein expression and purification

Briefly, the truncated XOAT1 coding-region was cloned into a mammalian expression vector, pGEn2-DEST, as previously described (Moremen et al., 2018). A truncated form of XOAT1 (amino acids 133–487) was used in all experiments that encodes the catalytic domain and lacks the N-terminal variable region. Specifically, the fusion proteins contain an NH2-terminal signal sequence followed by an 8xHis tag, an AviTag, ‘superfolder’ GFP, and the Tobacco Etch Virus (TEV) protease recognition site followed by amino acids 133–487 of Arabidopsis XOAT1 (Lunin et al., 2020). XOAT1 variants mutated at the predicted substrate-binding residue(s) were generated using the Q5 Site-Directed Mutagenesis Kit (New England Biolabs) and the aforementioned plasmid construct, pGEn2-DEST-XOAT1, as a template. Oligonucleotide primers (Eurofins, USA) used to generate the base changes were designed using NEBaseChanger (http://nebasechanger.neb.com/) and are listed in Supplemental Table A.1. The introduction of mutations into pGEn2-DEST-XOAT1 was confirmed by DNA sequencing (Eurofins, USA). Fusion proteins were expressed in HEK cells (FreeStyle™ 293-F cell line, Life Technologies) and purified from the culture media using HisTrap HP columns (Cytiva, USA) using an ÄKTA Pure 25 L protein purification system (GE Healthcare Life Sciences) as described previously (Prabhakar et al., 2020). Purification of XOAT1 and each variant was carried out on individual 1-mL HisTrap HP columns (Cytiva, USA) to exclude the risk of protein cross-contamination.

The purity of proteins was confirmed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) followed by Coo-massie Brilliant Blue R-250 (Bio-Rad) staining. The obtained fusion proteins were concentrated by using Amicon Ultra centrifugal filter devices (30-kDa molecular weight cutoff, EMD Millipore), then buffer exchanged into 75 mM HEPES sodium salt-HCl (pH 7.0) via dialysis. The dialysis buffer was supplemented with Chelex-100 resin (0.5 g/L, BioRad) to remove any potential contamination by divalent metal ions for activity assays. The concentrations of the dialyzed proteins were measured by Pierce™ BCA Protein Assay Kit (Thermo Fisher Scientific) using bovine serum albumin (BSA) as a standard. Protein expression levels of wild-type XOAT1 and the mutant variants, together with the primers used for generating mutations, are listed in Supplemental Table A.1.

2.3. In vitro activity assay

To monitor the ability of each enzyme to catalyze the transfer of O-acetyl moieties to xylan, each variant was evaluated using the standard in vitro acetyltransferase assay that was described previously (Lunin et al., 2020). Briefly, reactions (20 μL) consisted of 4 μM purified enzyme, 1 mM acetyl donor substrate and 0.1 mM 2-aminobenzamide-labeled xylohexaose (Xyl6-2AB) as an acceptor substrate, unless otherwise indicated, in 75 mM HEPES sodium salt-HCl (pH 6.8). All reactions were carried out at room temperature. Acetyl donor substrates used in this study include acetylsalicylic acid and 4-methylumbelliferyl acetate (4MU-Ac). After overnight incubation, a 5 μL aliquot of the reaction was mixed with 1 μL Dowex-50 (Bio-Rad, USA) cation exchange resin (suspended in water), and incubated at room temperature for 30 min. The mixture was then centrifuged at 2000 ×g for a minute to pellet the resin, and 1 μL of the supernatant was mixed with 1 μL of matrix solution (2% (w/v) 2,5-dihydroxybenzoic acid (DHB) in 50% (v/v) methanol) directly on a stainless steel MALDI target plate. The mixture was dried under a stream of warm air by a hair dryer (Conair) and analyzed by matrix-assisted laser desorption/ionization-time of flight mass spectrometry (MALDI-TOF MS) using a Microflex LT spectrometer (Bruker). Positive ion spectra of the reaction products were generated by summation of 200 laser shots at a frequency of 10 shots/s.

2.4. Kinetic analysis

To determine the kinetic parameters, assays were carried out in 75 mM HEPES sodium salt-HCl (pH 6.8) with enzyme concentrations at 1 μM (WT, N312A, R444A, Q445A, N312A/R444A, N312A/Q445A and R444A/Q445A), 2 μM (D403A), 3 μM (M311A and D403A/Q445A) or 6 μM (M311A/R444A, M311A/Q445A and D403A/R444A), depending on the catalytic performance of each variant. The concentration of the acceptor substrate, xylohexaose (Xyl6, Megazyme, Ireland), used in the assays ranged from 0 to 400 μM (WT, R444A, Q445A and R444A/Q445A), 0 to 2 mM (N312A, D403A, N312A/R444A, N312A/Q445A, D403A/R444A and D403A/Q445A) or 0 to 3 mM (M311A, M311A/R444A and M311A/Q445A) according to each variant’s catalytic velocity with a constant concentration of 1.5 mM 4MU-Ac as the donor substrate. Reactions were initiated with addition of 4MU-Ac, and the hydrolysis of 4MU-Ac by each variant was determined by measuring the emission of blue fluorescence from the released 4MU at 460 nm (excitation at 360 nm) using Synergy LX Multi-Mode Microplate Spectrometer (BioTek, USA) every minute during the 20-min reaction period at room temperature. Reaction rates with different Xyl6 concentrations were calculated from the linear portion of each variant’s hydrolysis curve. Steady-state parameters (kcat, KM and Vmax) were calculated by fitting the initial reaction velocities to the Michaelis–Menten equation with nonlinear curve fitting using GraphPad Prism version 7.0d Macintosh Version (GraphPad Software Inc., USA; www.graphpad.com).

2.5. Hydrolytic activity determination

Previously, we confirmed an acetyl-enzyme intermediate is formed at Ser-216 of XOAT1 using 4MU-Ac as an acetyl donor (Lunin et al., 2020). To monitor the ability of each enzyme to form the acyl enzyme intermediate, the esterase activity of acetyltransferases can be measured by quantification of the released deacylated donor, which indicates their ability to form the acyl-enzyme intermediate in the absence of acceptor. The ability of each variant to hydrolyze 4MU-Ac in the absence of acceptor substrate was determined through the aforementioned method described by Lunin et al., without adding Xyl6 or any other acceptor substrates in the reaction (Lunin et al., 2020). The autohydrolytic background of 4MU-Ac was determined by measuring the released 4MU from the incubation of 1.5 mM 4MU-Ac in 75 mM HEPES sodium salt-HCl (pH 6.8), and was subtracted from the value obtained from each enzyme-containing hydrolytic assay.

2.6. Protein thermal shift assays

Thermal shift analysis of the protein stability of XOAT1 and its variants was compared using SYPRO Orange Fluorescent Dye (Huynh & Partch, 2015). Protein thermal shift assays were carried out using a CFX96™ Real-Time System (Bio-Rad, USA) to detect fluorescent dye-bound denatured proteins according to the protocol from the manufacturer (https://www.biorad.com/webroot/web/pdf/lsr/literature/Bulletin_7180.pdf). In brief, 45 μL of protein (2 μM) in 75 mM HEPES-sodium salt buffer (pH 6.8) was mixed with 5 μL of 100× SYPRO Orange Protein Gel Stain (diluted from 5000× stock supplied by Sigma-Aldrich, USA; λex 470 nm/λem 570 nm) in 96-well Hard-Shell® PCR plates (Bio-Rad, USA) and sealed with PCR Sealers™ Microseal® ‘B’ Film (Bio-Rad, USA). Controls contain an equal volume of 75 mM HEPES-sodium salt buffer (pH 6.8) in lieu of protein. The program included an initial temperature hold of 10 °C for 31 s, followed by a temperature ramp from 10 °C to 95 °C at 0.5 °C increments with a 30-s hold at each temperature. Fluorescence reads using the “FRET” channel to measure the SYPRO Orange fluorescence signal were taken at the end of each hold, and the obtained data were processed using CFX Maestro™ Software for calculation of the melting temperature (Tm) of each XOAT1 variant.

2.7. In vitro synthesis of acetylated xylan

For cell-free, in vitro synthesis of acetylated xylo-oligosaccharides with adjusted acetylation levels, we carried out a one-pot reaction by co-incubating heterologously expressed xylan synthase from Klebsormidium flaccidum (KfXYS1)(Jensen et al., 2018) and XOAT1 or its variants at an enzyme concentration of 3 μM each in the presence of 60 mM uridine 5′-diphosphate (UDP)-xylose as an activate nucleotide sugar donor, 2 mM xylobiose (Xyl2)as an acceptor for the xylan synthase, 3.5 mM acetylsalicylic acid as an acetyl donor in 75 mM HEPES-sodium salt (pH 6.8). After overnight incubation, the products were analyzed by MALDI-TOF MS according to the method described above.

3. Results and discussion

In our previous study, we provided several lines of evidence that suggest XOAT1 uses a double displacement Bi–Bi reaction to catalyze 2-O-acetylation of xylan that occurs in two stages. First, Ser-216 of the catalytic triad is acetylated, forming an acyl-enzyme intermediate that has been experimentally confirmed by LC-MS/MS (Lunin et al., 2020). The transfer of the proton from Ser-216 to the phenolic oxygen of the donor results in release of deacylated donor, which can be measured spectrophotometrically when pNP-Ac or 4MU-Ac are used. After formation of the acyl-enzyme intermediate, the xylan substrate binds to the substrate binding groove to get into alignment with the active site. Next, the acetyl group is transferred from Ser216 onto the acceptor xylan substrate (Lunin et al., 2020). The main objective of this study is to understand mechanisms of acceptor substrate recognition and the residues dictating the acetylation propensities of XOAT1. Thus, we started by identifying the key residues involved in acceptor binding followed by detailed biochemical characterization of the impact of mutations on the overall acetylation mechanism, kinetic parameters, and finally the ability of putative acceptor binding mutants to synthesize xylan with different degrees of O-acetyl substitution that could be potentially applied in planta to generate xylan with different physicochemical properties to modulate lignin-carbohydrate interactions in muro.

3.1. In-silico studies enable identification of mutagenesis targets

Molecular dynamics simulations of the XOAT1-xylodecaose complex revealed important residues involved in stabilizing the binding of the acceptor substrate. Table A.2 in the supplemental information lists these amino acid residues ranked according to the strength of their interaction energies. The amino acids identified to be interacting with the substrate span the whole binding groove and involve residues on both the major and the minor lobes. The residues that were observed to have the highest binding energies were situated closest to the catalytic triad. Amongst the top-five ranked residues involved in interacting with xylodecaose, two are part of the minor lobe (R444 and Q445), and the other three residues are part of the more structured major lobe (M311, N312 and D403) (Fig. 1). Interestingly, the two distinct lobes interact with different sections of xylodecaose, with residues localized on the major lobe (M311, N312 and D403) binding to the terminal (non-reducing) region, while the residues on the minor lobe (R444 and Q445) interacting with the reducing end of xylodecaose (Fig. 1).

Fig. 1.

Fig. 1.

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Surface topology of XOAT1 in the substrate bound state. (a, b) The major and minor lobes colored in cyan and green respectively with the substrate show in Licorice representation with carbons colored yellow. The substrate residues have been labeled with increasing numbers from the non-reducing end to the reducing end with the residue at the active site being labeled 0. (c) A close-up view of the bound substrate, the catalytic triad (Ser216-His465-Asp462 depicted in VDW representation) and the residues identified to be having the highest binding energies from MD represented in licorice representation with the carbons colored gray.

3.2. Biochemical analysis of XOAT1 variants

Although molecular simulations enable the identification of putative residues in substrate binding, the roles played by these residues in governing the acetylation mechanism still needs to be established. Thus, we generated a series of XOAT1 variants using site-directed mutagenesis, determined kinetic parameters and analyzed the reaction products to investigate their potential role in xylan acetylation. This was executed by substituting each of the five hypothesized binding residues to alanine in various combinations resulting in five single mutants (M311A, N312A, D403A, R444A and Q445A) and nine double mutants (M311A/D403A, M311A/R444A, M311A/Q445A, N312A/D403A, N312A/R444A, N312A/Q445A, D403A/R444A, D403A/Q445A and R444A/Q445A) (Lunin et al., 2020). Kinetic parameters were determined by measuring the initial reaction rates using the fluorogenic substrate 4MU-Ac in the presence and absence of acceptor (Lunin et al., 2020). The ability of the enzymes to transfer an acetyl group onto a saccharide acceptor was measured using and an in vitro acetyltransferase assay that is based on the same principles as β-(1,4)-xylosyltransferase assays, comprised of acetylsalicylic acid as an acetyl donor and 2-aminobenzamide β-(1,4)-xylohexaose (Xyl6-2AB) as an acceptor (Urbanowicz, Peña, Moniz, Moremen, & York, 2014).

The wild-type XOAT1 enzyme and the R444A, Q445A and R444A/Q445A variants show similar acetyltransferase activity and are able to transfer up to four acetyl groups to Xyl6-2AB (Fig. 2a, b). On the other hand, mutants of the major lobe residues showed decreased acetyltransferase activities, with the single mutants M311A, N312A and D403A generating xylooligosaccharides with up to 2 (M311A) or 3 (N312A and D403A) O-acetyl substituents (Fig. 2a). It is worth noting that substrate-binding residues on the two distinct lobes are in close proximity to the opposite ends of the xylosyl chain as observed in simulations, where major lobe residues are near the terminal non-reducing end while interactions with the minor lobe occur closer to the reducing end. Double substitutions of the major lobe residues also resulted in drastic reductions of enzyme activity, with N312A/D403A only producing mono-acetylated xylohexaose after overnight incubation, and M311A/D403A demonstrating no acetyltransferase activity (Fig. 2b). The thermal stability of the mutants was compared with that of the wild-type XOAT1 protein by measuring temperature-induced changes in the fluorescence SYPRO® Orange. Wild-type XOAT1 exhibited a melting temperature (Tm) of 53 ± 0.5 °C. Analysis of the mutant variants showed that M311A and N312A showed decreases (−1.7 and −2.0, respectively) compared to the wild type (Fig. 2a), indicating mutation of residues causes loss of acetyltransferase activities without significantly affecting protein stability. D403A, R444A, and Q445A did not show significant changes in Tm in comparison with wild type. Taken together, mutation of different residues on the major lobe reduced XOAT1’s acetyltransferase activity to a greater or lesser extent, while substituting minor lobe residues, in contrast, allowed the enzyme to retain an activity that is comparable to the wild type, suggesting a more important role for the major lobe in substrate binding and the acetylation mechanism.

Fig. 2.

Fig. 2.

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Analysis of acetyltransferase activity of XOAT1 variants. (a) Single mutants of XOAT1 are sorted by increasing enzymatic efficiency (kcat/KM) from left to right. Protein thermal stability (Tm) and kcat/KM of each variant are shown in the top panel. For reference, the Tm and kcat/KM of wild-type XOAT1 are 53 ± 0.5 °C and 68.51 M−1⋅S−1, respectively. In the second panel, sequence logos showing conservation of amino acids are depicted based on an alignment of nine xylan-specific TBLs, and the location of each mutated residues is marked by an asterisk. In the third panel, local areas inside the binding pocket of XOAT1 with point mutations and the docked xylodecaose are shown. Xylosyl residues are numbered according to the proximity to the catalytic site, the closest residue is assigned as 0, and the number increases towards reducing end. The major lobe is colored in cyan, while the minor lobe is in green. In the bottom panel, MALDI-TOF MS spectra of acetylated Xyl6-2AB products generated by XOAT1 variants after overnight reactions are shown. (b) MALDI-TOF MS spectra of acetylated products generated by the double mutants of XOAT1. M311A/D403A variant generated no acetylated products, and the spectrum is shown in Fig. A.1. An addition of one acetyl group increases the mass of Xyl6-2AB by 42 Da as indicated by [M + H]+ ions.

The in vitro analysis reveals the overall effect of the mutations on XOAT1 activity which could be manifested via the impact on binding or catalysis or a combination of the two. In order to probe this further, we determined the kinetic parameters of XOAT and its variants for the acceptor by generating a substrate saturation curve for each variant. The impact of mutations on xylohexaose binding affinity to XOAT1 variants was quantified by determining the Michaelis constant (KM) for each XOAT1 variant. The minor lobe mutants (R444A, Q445A and R444A/Q445A) showed a similar or slightly higher KM compared to the wild type (Table 1), indicating that they maintained a comparable xylohexaose-binding affinity to that of the wild type. Contrarily, all the major lobe mutants demonstrated highly impaired xylohexaose-binding abilities as shown by the significantly higher KM, and which is consistent with the results of the acetyltransferase assays that showed the reduced acetylation levels of the products generated by these variants (Fig. 2a and b). Taken together, these data suggest that the major lobe plays an important role in interacting with the xylan acceptor.

Table 1.

Kinetic parameters for XOAT1 variants.

XOAT1 variant KM (μM) Vmax (pmol·min−1·nmol−1) kcat (S−1) kcat/KM (M−1·S−1)
WT 51.5 ± 6.5 211.5 ± 8.6 0.0035 ± 0.00014 68.5
M311A 414.0 ± 144.5 29.6 ± 3.2 0.00049 ± 0.000053 1.2
N312A 493.3 ± 49.6 389.9 ± 13.9 0.0065 ± 0.00023 13.2
D403A 250.1 ± 34.6 110.9 ± 4.9 0.0018 ± 0.000081 7.4
R444A 51.5 ± 5.8 304.3 ± 11.1 0.0051 ± 0.00018 98.5
Q445A 56.9 ± 5.8 261.9 ± 8.8 0.0044 ± 0.00015 76.8
M311A/D403A n.d.a n.d.a n.d.a n.d.a
M311A/R444A 497.8 ± 266.3 26.1 ± 4.8 0.00043 ± 0.000080 0.9
M311A/Q445A 575.4 ± 260.3 24.6 ± 4.00 0.00041 ± 0.000066 0.7
N312A/D403A n.d.a n.d.a n.d.a n.d.a
N312A/R444A 447.5 ± 23.7 279.3 ± 5.5 0.0047 ± 0.000092 10.4
N312A/Q445A 445.2 ± 25.5 251.7 ± 5.4 0.0042 ± 0.000090 9.4
D403A/R444A 890.9 ± 264.3 56.6 ± 7.8 0.00094 ± 0.00013 1.1
D403A/Q445A 420.5 ± 125.8 87.2 ± 9.6 0.0015 ± 0.00016 3.5
R444A/Q445A 75.8 ± 9.3 279.9 ± 12.2 0.0047 ± 0.00020 61.6
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a

n.d., not detected.

The turnover number (kcat) of each variant varies, and does not correspond to their localizations on the major or minor lobe of XOAT1. For example, point mutation of N312 residue on the major lobe resulted in a low affinity (high KM) for xylohexaose, while the kcat values of the N312-related mutants, on the contrary, were dramatically increased, especially for the single mutant N312A (Table 1). The increased kcat of N312-related mutants satisfied a catalytic efficiency (kcat/KM) that allows those variants, although with low substrate binding affinities, to still be able to generate multi-acetylated xylohexaoses as shown in the in vitro assays (Fig. 2b. MALDI — major). Besides N312, mutating other two putative substrate-binding residues localized on the major lobe, M311 and D403, significantly decreased kcat, especially for M311-related single and double mutants. The reduced turnover numbers (low kcat) and decreased substrate-binding affinities (high KM) resulted in low catalytic efficiencies of the M311 and D403-related mutants. On the other hand, substituting the two residues R444 and Q445 on the minor lobe to alanine caused the kcat to increase. R444A, Q445A and R444A/Q445A all presented higher kcat, demonstrating that switching from bulky to small and non-polar side chains on the residues at these positions promotes the number of reactions that the enzyme catalyzes per unit time. Without losing the substrate-binding affinity, R444A, Q445A and R444A/Q445A variants revealed much improved catalytic efficiencies compared to the wild type.

It has been shown that targeting substrate-binding residues along the active site cleft, based on computational modeling, is an efficient strategy to evaluate the protein design space to mediate enzymatic activities of TBL proteins. An example of such rationally designed enzymes has been reported in a previous study of Candida antarctica lipase A (CAL-A), which generates fatty acid esters that are useful in the biodiesel, cosmetic, and food industries (Müller, Sowa, Fredrich, Brundiek, & Bornscheuer, 2015). The in silico designed CAL-A mutant D122L, showed lower hydrolysis activity that reduces the production of unwanted fatty acid by-products, while retaining the process-relevant acyltransferase activity and thermostability (Müller, Sowa, Fredrich, Brundiek, & Bornscheuer, 2015). Another example of targeting binding pocket residues based on sequence alignment and 3D modeling to improve enzymatic activities has been reported in lipases/acyltransferases in Candida species (Jan, Subileau, Deyrieux, Perrier, & Dubreucq, 2016). In addition to enhancing enzyme activities, switching substrate specificity through mutating key residues in binding channels is an effective enzyme engineering strategy. Substituting small amino acids with the ones that contain bulky side chains inside the tunnel of Candida rugose lipase resulted in shifting the substrate preference from long chain fatty acids towards those with shorter acyl groups (Schmitt, Brocca, Schmid, & Pleiss, 2002).

Overall, our research on the substrate-binding mode of XOAT1 revealed the important residues involved in binding the xylooligosaccharides that were initially suggested by computational modeling and then confirmed by SDM and activity assays. The kinetic parameters strongly revealed that mutating the hypothesized binding residues on the major lobe, M311, N312 and D403, critically interferes with XOAT1’s ability to interact with xylohexaose, suggesting that the major lobe is critical for effective substrate binding and can be used as a future target for generation of modified lignocellulosic materials with reduced acetylation levels through transgenic biotechnology. It has been recently shown that heterologously expressed fungal acetyl xylan esterase from Hypocrea jecorina (HjAXE) in hybrid aspen using wood-specific promoter helped reduce recalcitrance for enzymatic saccharification and thus improved sugar yield by 27% in transgenic lines (Wang et al., 2020). A similar strategy of expressing a fugal AXE in muro has also been applied in Arabidopsis and aspen wood, and the resulting transgenic lines showed increased susceptibility to hydrolytic enzymes that led to improved yields of reducing sugars (Pawar et al., 2016; Pawar et al., 2017), suggesting that it’s a promising approach to optimize lignocellulosic biomass for biofuel production through changing cell wall architecture but without interfering with plant growth. The major lobe of XOAT1 has been shown to be determinant in binding the substrate in our research, and could be a potential transgenic target to reduce recalcitrant acetyl content in biomass through up-stream modification of xylan structures in the cell wall biosynthetic pathway.

Although the residues chosen for mutagenesis were identified to be involved in acceptor binding and are most likely to be involved in the second stage (acetyl transfer) of the XOAT1 catalytic mechanism, it is important to recognize that they could plausibly also be involved or impact the first stage i.e. the esterase activity of XOAT1. To study if mutations of the hypothesized binding residues affect XOAT1’s hydrolytic activity, the variants were examined using 4MU-Ac as the donor substrate and measuring the release of 4MU in the absence of any acceptor substrates. All mutant variants were capable of releasing 4MU from 4MU-Ac in vitro. However, M311A and D403A single mutants presented significantly reduced hydrolytic activities compared to other single mutants, and all the M311 and D403-related double mutants, except N312A/D403A, showed decreased hydrolytic rates (Fig. A.2). The obtained esterase activities of the XOAT1 variants demonstrated that the two residues, M311 and D403, on the major lobe are not only critical to binding the acceptor substrate, xylohexaose, but decrease the rate that the enzyme forms an acyl-enzyme intermediate using the donor substrate, 4MU-Ac. The third major lobe residue, N312 and its related double mutants displayed hydrolytic activities at a similar level as the wild type, despite its close proximity to the catalytic triad. It is worth noticing that the double mutant N312A/D403A maintains a hydrolysis rate comparable to the wild type while the single mutant D403A does not, suggesting that the loss of activity by mutating D403 can somehow be compensated by mutation of N312. Finally, mutating the two residues on the minor lobe R444 and Q445 showed no difference in hydrolysis activity relative to wild-type.

3.3. XOAT1 variants are able to synthesize xylo-oligosaccharides with different degrees of O-acetylation

Previously, we have identified a highly active xylan synthase from Klebsormidium flaccidum, designated K. flaccidum XYLAN SYNTHASE-1 (KfXYS1), which is able to synthesize xylan polymers with a degree of polymerization of up to 29 Xyl residues (Jensen et al., 2018). We have taken advantage of the highly active nature of KfXYS1 to develop a model in vitro xylan synthesis platform to evaluate XOAT1 variants. Mutants with different enzymatic efficiencies were reacted together with KfXYS1 to evaluate production of xylo-oligosaccharides (XOS) with adjusted degrees of acetylation in vitro, to more closely mimic the long chain acceptors that the enzymes would encounter in vivo. One-pot reactions were carried out by co-incubating KfXYS1 and XOAT1 or its variants in reactions comprised of uridine 5′-diphosphate-xylose (UDP-Xyl) and xylobiose (Xyl2) as the donor and acceptor for xylan synthesis, respectively, together with and acetylsalicylic acid as an acetyl donor (Fig. 3a). MALDI-TOF MS analysis of the products from the overnight reaction shows that KfXYS1 was able to generate XOS with degree of polymerization (DP) ranging from 8 to around 20, and up to 2 to 4 acetyl groups were attached to the XOS depending on their DP (Fig. 3b). Consistent with our hypothesis, the acetylation levels of the xylo-oligo products produced by different XOAT1 variants differ according to their enzymatic efficiencies, suggesting a potential strategy to synthesize xylan polymers and XOS in vitro or going further and replacing XOAT1 in planta with mutant variants to produce plant secondary cell walls with tuned properties through adjusted acetylation levels.

Fig. 3.

Fig. 3.

Open in a new tab

Acetylated xylo-oligosaccharides (XOS) synthesized through one-pot reaction catalyzed by xylan synthase and XOAT1 or its variants. (a) Scheme of the experimental procedure. (b) MALDI-TOF spectra of acetylated products generated by five variants (M311A, N312A, D403A, R444A, and Q445A) are shown and sorted from top to bottom by decreasing enzymatic efficiency. Control contains no XOAT1 or its variants. Degree of polymerization (DP) of XOS are labeled on top of the signals, and an asterisk represents one acetyl group attached to the xylosyl species. An addition of one acetyl group increases the mass of xylo-oligomers by 42 Da as indicated by [M + H]+ ions.

The human genome does not encode genes to produce enzymes to deconstruct dietary xylan. However, xylan has been shown to exhibit prebiotic effects of stimulating a limited number of beneficial bacteria in gastrointestinal microbiota such as Bifidobacterium, which can beneficially affect the host’s health (Mussatto & Mancilha, 2007; Samanta et al., 2015). Besides exhibiting prebiotic effects, XOS has also been shown to help prevent diabetes and colon cancer (Aachary, Gobinath, Srinivasan, & Prapulla, 2015; Yang et al., 2015), and other properties such as antioxidant, anti-inflammatory, and thermal (up to 100 °C) and pH (pH 2.5–8) stability have made XOS suitable for applications in various marketable areas such as cosmetic, pharmaceutical, and food industries (Amorim, Silvério, Prather, & Rodrigues, 2019; Vázquez, Alonso, Domínguez, & Parajó, 2000). Moreover, XOS has been applied in microencapsulation techniques used in the field of dietary and medical supplements for microbiota balancing through improvement of digestion resistance and viability of the encapsulated probiotic organisms under gastrointestinal conditions (Liao et al., 2019; Martins, Silva, Ávila, Sato, & Goldbeck, 2021). Different from traditional methods using lignocellulose as raw materials for XOS production (Amorim, Silvério, Prather, & Rodrigues, 2019; Carvalho, Neto, da Silva, & Pastore, 2013; Samanta et al., 2015), the results of one-pot XOS synthesis shown in this study provide a different strategy that allows us to finely adjust the acetylation level, which could have significant impacts on the physiochemical property of xylan (Biely, 2012), of XOS during synthesis, and could thus generate modified features of XOS to meet the requirement for different purposes in various areas mentioned above.

4. Conclusions

The ability to modulate the acetylation activity of XOAT1 presents unique opportunities for the engineering of plant biomass with highly controlled degrees of acetylation without compromising on plant growth and stature. This will yield unique insights into the role of acetylated biopolymers in enabling healthy plant cell wall architectures that could also be easily deconstructed. Additionally, this capability to exploit natural plant enzymatic pathways would also open doors for the development of biosynthetic technologies directed towards the tailored biosynthesis of glycopolymers. In this study, we used computational molecular simulations to bridge gaps in the structural biology of XOAT1 enzyme-substrate complexes. The simulations identified key residues involved in acceptor substrate binding and provided targets for directed mutagenesis of XOAT1. The detailed biochemical characterizations conducted here enabled insights into the role of these residues in substrate binding, esterase and transferase activities.

Amongst the five amino-acid residues identified to play an important role in substrate binding from simulations, two of them are present on the minor lobe (R444, Q445) and three on the major lobe (M311, N312 and D403). The single and double mutant variants of XOAT1 are observed to have differing impacts on substrate binding, esterase and transferase activities. XOAT1 variants of the minor lobe residues (R444A, Q445A, R444A/Q445A) are observed to have the least detrimental impact on acceptor binding affinities, esterase activity and turnover rates for transferase activities. In fact, the R444A variant demonstrates improved catalytic efficiency for substrate acetylation. On the other hand, XOAT1 variants of the major lobe had a far more detrimental impact on acceptor substrate binding and kinetic parameters. All single and double mutants involving any of the major lobe residues, M311A, D403A and N312A showed highly diminished binding affinities for the acceptor xylan substrate. XOAT1 variants involving M311A and D403A also demonstrate highly reduced esterase activities indicating that the transfer of the acetyl group may also be hindered in these cases. Interestingly, all XOAT1 variants involving the N312A mutation seemed to demonstrate restored esterase activity. However, all single and double mutants involving the major lobe residues show impaired xylan acetylation activities. Finally, by combining the xylan synthase enzyme with specific XOAT1 variants described above, we demonstrate a biochemical pathway for the production of xylo-oligomers with highly controlled degrees of acetylation.

This study forms the basis for the quest to gain a detailed and complete understanding of the impact of acetylation on the assembly of biopolymers in the plant cell wall as well as the development of engineered acetylated biopolymers for a circular economy.

Supplementary Material

Supplementary Material

Funding

Funding was provided by the Center for Bioenergy Innovation (CBI), a U.S. Department of Energy Bioenergy Research Center supported by the Office of Biological, Environmental Research in the DOE Office of Science and National Institutes of Health (P41GM103390 and R01GM130915). This work was authored in part by Alliance for Sustainable Energy, LLC, the manager and operator of the National Renewable Energy Laboratory for the U.S. Department of Energy (DOE) under Contract No. DE-AC36-08GO28308. The views expressed in the article do not necessarily represent the views of the DOE or the U.S. Government. The U.S. Government retains and the publisher, by accepting the article for publication, acknowledges that the U.S. Government retains a nonexclusive, paid-up, irrevocable, worldwide license to publish or reproduce the published form of this work, or allow others to do so, for U.S. Government purposes.

Footnotes

CRediT authorship contribution statement

Hsin-Tzu Wang: Conceptualization, Investigation, Methodology, Formal analysis, Visualization, Validation, Data curation, Writing – original draft. Vivek S. Bharadwaj: Conceptualization, Software, Investigation, Methodology, Formal analysis, Visualization, Validation, Data curation, Writing – original draft. Jeong Yeh Yang: Investigation, Methodology, Writing – review & editing. Thomas M. Curry: Investigation, Methodology, Writing – review & editing. Kelley W. Moremen: Funding acquisition, Resources, Methodology, Formal analysis, Project administration, Visualization, Supervision, Writing – review & editing. Yannick J. Bomble: Conceptualization, Funding acquisition, Resources, Methodology, Project administration, Supervision, Writing – review & editing. Breeanna R. Urbanowicz: Conceptualization, Funding acquisition, Resources, Methodology, Formal analysis, Visualization, Validation, Project administration, Supervision, Data curation, Writing – original draft.

Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.carbpol.2021.118564.

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Supplementary Material
NIHMS1745952-supplement-Supplementary_Material.pdf (714.6KB, pdf)

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