2026 年 73 巻 3 号 論文ID: 7303102
Lytic polysaccharide monooxygenases (LPMOs) are activated by electrons supplied from external redox enzymes. Cellobiose dehydrogenase (CDH) and pyrroloquinoline quinone (PQQ)-dependent pyranose dehydrogenase (PDH) are electron donors to LPMOs. CDH and PDH each consist of two domains: a catalytic domain that oxidizes substrates to extract electrons, and an AA8 domain that receives and transfers these electrons to LPMOs. Although non-catalytic AA8 proteins comprising only an AA8 domain and a cellulose-binding module have been reported, their electron donors and physiological roles remain largely unclear. In this study, we characterized a protein from Armillaria tabescens (syn. Desarmillaria tabescens) that lacks a catalytic domain but includes both an AA8 domain and a cellulose-specific binding module, CBM104, which we designated AtAA8-CBM104. Electron-transfer assays demonstrated that the AA8 domain can accept electrons from an external AA12 catalytic domain, strongly suggesting that AtAA8-CBM104 act as an electron mediator transferring electrons from redox partners such as AA12 to adjacent LPMOs. Further, adsorption experiments showed that the CBM104 domain binds to crystalline cellulose but with lower adsorption efficiency than a previously characterized CBM104 appended to LPMO9. This observation suggests that despite AtAA8-CBM104 localization on cellulose being necessary for electron transfer to cellulose-bound LPMO9, excessive retention on the cellulose surface may interfere with LPMO9 catalysis. Furthermore, phylogenetic analysis indicated that AA8 appended to CBM104 lacking catalytic domains share similar electrostatic properties. These findings advance our understanding of non-catalytic AA8 proteins, underscoring their potential involvement in fungal redox networks linked to cellulose decomposition.
AA, auxiliary activities; CAZy, carbohydrate-active enzymes; CBM, carbohydrate-binding module; CDH, cellobiose dehydrogenase; CV, cyclic voltammetry; JGI, Joint Genome Institute; LPMO, lytic polysaccharide monooxygenase; MWCO, molecular weight cut-off; PCR, polymerase chain reaction; PDH, pyrroloquinoline quinone-dependent pyranose dehydrogenase; PQQ, pyrroloquinoline quinone; SAM, self-assembled monolayer.
Wood-rotting fungi, which include specific basidiomycetes and ascomycetes, are the only known organisms capable of fully degrading wood independently. These fungi utilize cellulose in wood as a nutrient source. Wood cellulose comprises both crystalline regions―where cellulose chains are tightly packed via hydrogen bonding and exhibit strong resistance to degradation―and amorphous regions, which are less ordered and more easily degraded. Wood-rotting fungi can degrade not only the amorphous regions but also the crystalline regions of cellulose [1, 2, 3, 4]. The crystalline regions are initially cleaved oxidatively by lytic polysaccharide monooxygenases (LPMOs), which disrupt the ordered structure and enhance cellulose accessibility to cellulases, thereby facilitating hydrolytic depolymerization [3, 4]. The oxidative function of LPMOs depends on an external electron donor to initiate the reaction, and cytochrome proteins, such as cellobiose dehydrogenase (CDH) and pyrroloquinoline quinone (PQQ)-dependent pyranose dehydrogenase (PDH) have been identified as oxidoreductases capable of supplying electrons to LPMOs [5, 6, 7, 8].
CDH and PDH consist of two domains: a catalytic domain belonging to Auxiliary Activities family 3 (AA3) or AA12, which oxidizes specific sugar substrates to generate electrons, and an AA8 domain that transfers these electrons to LPMOs [9, 10, 11, 12, 13]. In some cases, a cellulose-binding module (CBM1) is also attached to the C-terminal end. Particularly, AA8 is a cytochrome domain that folds into an immunoglobulin-like β-sandwich, containing a single b-type heme as a prosthetic group, where the central iron ion of the heme is coordinated by histidine and methionine residues [13]. These AA8 domains mediate electron transfer by accepting electrons generated through catalytic oxidation and relaying them to other redox enzymes [8, 14]. Although AA3 and AA12 domains alone can directly supply electrons to LPMOs, AA8 has only electron-transfer functionality; therefore, it cannot donate electrons to LPMOs in isolation. Although AA8-containing proteins lacking catalytic domains but featuring a CBM1, which binds cellulose, have also been identified [15], the electron donors and biological roles of such non-catalytic cytochrome proteins in cellulose degradation remain unresolved.
In this study, we identified a novel non-catalytic AA8 protein in the genome of the wood-rotting fungus Armillaria tabescens (syn. Desarmillaria tabescens). This protein consists of an AA8 domain and a recently identified cellulose-binding domain, CBM104 (designated AtAA8-CBM104). CBM1 binds to cellulose without discriminating among different crystalline forms, whereas CBM104 exhibits a specific binding affinity for native crystalline cellulose (cellulose I) [16].
Here, we first performed functional analyses of the individual AA8 and CBM104 domains of AtAA8-CBM104 and investigated their physiological roles. Additionally, to infer the potential functions of other non-catalytic AA8 proteins aside from AtAA8-CBM104, we performed a molecular phylogenetic analysis based on amino acid sequences. To our knowledge, this is the first description of an AA8 protein lacking a catalytic domain, harboring a CBM104 at its C-terminus. The present findings provide significant insights into the potential function of non-catalytic cytochrome proteins involved in electron-transfer processes during cellulose degradation.
Strains. The A. tabescens strain NBRC31620 was used as the source of the target genes. Escherichia coli strain TOP10 (Thermo Fisher Scientific Inc., Waltham, MA, USA) and Komagataella phaffii (formerly Pichia pastoris) strain KM71H (Invitrogen Corp., Waltham, MA, USA) were employed as hosts for subcloning and heterologous expression of recombinant proteins, respectively.
Sequence analysis. A search for cytochrome protein exhibiting novel domain architectures was performed using BLASTp against fungal genome sequences available in the Fungal Genome Database hosted by the Joint Genome Institute (JGI) Genome Portal (https://genome.jgi.doe.gov/programs/fungi/index.jsf). The AA8 domain sequence from CBCyt.b562―a protein consisting of AA8 and CBM1 domains from Phanerodontia chrysosporium (formerly Phanerochaete chrysosporium; Protein ID: 6112339)―served as the query, using the BLOSUM62 substitution matrix and an E-value cutoff of 1 × 10−3 [15]. This search led to the identification of a cytochrome with a previously unreported domain arrangement in the genome of A. tabescens (Protein ID: 1636565). The sequence was subsequently designated as AtAA8-CBM104.
To determine the AA8 domain sequence of this protein, the amino acid sequence predicted to adopt an AA8-like structure was identified using AlphaFold 3 and PrDOS [17, 18]. The resulting AA8 domain sequence was subsequently aligned with four previously characterized AA8 domains (GenBank: EAU83856.1, ABS45567.2, EAA27355.1, AAC49277.1) using MAFFT Multiple Sequence Alignment Software version 7 [19]. To investigate the domain appended at the C-terminal end of the AA8 domain, a homology search was performed via BLASTp (BLOSUM62, E-value < 1 × 10−3), indicating that the domain corresponded to CBM104. This sequence was then aligned with the CBM104 amino acid sequence from Gloeophyllum trabeum using MAFFT [19].
The three-dimensional structures were predicted using AlphaFold 3 [17].
Phylogenetic analysis. To compile amino acid sequences of the AA8 domain, the National Center for Biotechnology Information protein database was queried using the BLASTp algorithm [20], with AA8 domain sequences from AtAA8-CBM104 and 20 characterized cytochrome proteins as queries. The 21 query sequences used for the BLASTp search are listed in Table S1 (see J. Appl. Glycosci. Web site). All searches were conducted with standard settings using the BLOSUM62 substitution matrix. For each query, 30 amino acid sequences were retrieved, resulting in a total of 651 sequences, including the original queries. After removal of duplicate full-length sequences, multiple sequence alignment was performed using Clustal Omega (ver. 1.2.4) [21, 22]. Non-AA8 domain regions were manually excised using SeaView (ver. 4) [22]. Redundant sequences within the AA8 domain region were then eliminated, followed by a second round of alignment using Clustal Omega. To identify the optimal amino acid substitution model, the resulting alignment was saved as a text file using SeaView and analyzed with ModelTest-NG (ver. 0.1.7) [23]. Phylogenetic analysis was conducted using RAxML-NG (Randomized Axelerated Maximum Likelihood-Next Generation, version 1.2.2) under the selected model (WAG+I+G4+F) [24]. Bootstrap analysis was carried out with 1,000 replicates. The resulting phylogenetic tree was rooted using the midpoint rooting method and visualized using Interactive Tree of Life (iTOL, ver. 6) [25].
Sequence logos depicting the conservation of amino acid residues were generated for each of the six clades in the resulting phylogenetic tree using WebLogo [26].
Cloning of genes encoding AtAA8-CBM104. Armillaria tabescens was statically cultured at 25 °C on YMC agar medium (yeast extract 0.4 %, malt extract 1.0 %, Avicel 0.4 %) until visible mycelial growth was observed. The resulting mycelium was excised and transferred into P-YPG liquid medium (yeast extract 0.2 %, malt extract 0.2 %, D(+)-glucose 1.0 %) and incubated at 25 °C with agitation at 180 rpm. After 9 days of cultivation, total RNA was extracted from fungal cells using the RNeasy Plant Mini Kit (QIAGEN N.V., Venlo, Netherlands). The mRNA was then purified from total RNA using the Oligotex-dT30 Super mRNA Purification Kit (Takara Bio Inc., Shiga, Japan). First-strand cDNA was synthesized from the isolated mRNA via reverse transcription using SuperScript IV Reverse Transcriptase (Thermo Fisher Scientific Inc.). Polymerase chain reaction (PCR) was conducted with KOD-Plus-Neo polymerase (Toyobo Co., Ltd., Osaka, Japan), employing gene-specific primers targeting AtAA8-CBM104 and the synthesized cDNA as the template. Primers were designed based on the nucleotide sequence available in the JGI database to amplify the coding region excluding the signal peptide: forward primer, 5′-AGGGGTATCTCTCGAGAAAAGACAGTCCGCCTCTGCCTACAC-3′; reverse primer, 5′-AGAAAGCTGGCGGCCGCCTACAACCGACGTCCCAGGTTG-3′. The PCR product was purified using the Wizard SV Gel and PCR Clean-Up System (Promega Corp., Madison, WI, USA). The purified DNA fragment was subsequently subcloned into the pPICZα vector (Invitrogen Corp.) that had been digested with NotI (Invitrogen Corp.) and XhoI (Invitrogen Corp.) using the In-Fusion HD Cloning Kit (Takara Bio Inc.). The resulting construct was used for protein expression.
Heterologous expression and purification of the recombinant proteins. Approximately 10 µg of pPICZα expression plasmid DNA encoding AtAA8-CBM104 and the AA8 domain alone of AtAA8-CBM104 (AtAA8) were linearized with BlpI (New England Biolabs, Ipswich, MA, USA) and subsequently transformed into K. phaffii. Selection of transformants and production of recombinant proteins were conducted according to previously described methods [16]. Culture supernatants were concentrated using 20 mM sodium acetate buffer (pH 5.2) containing 1 M ammonium sulfate. The concentrates were applied to a HiTrap Phenyl HP column (Cytiva, Marlborough, MA, USA) pre-equilibrated with the same buffer, and proteins were eluted using the identical buffer. Fractions containing recombinant proteins were collected, pooled, and buffer-exchanged into 20 mM sodium phosphate buffer (pH 6.5) using Vivaspin 20 centrifugal concentrator tubes with a 10,000 Da molecular weight cut-off (MWCO) membrane (Sartorius AG, Göttingen, Germany). The pooled fractions were then applied to a HiTrap DEAE FF column (Cytiva) equilibrated with the same buffer. Elution was carried out using a linear gradient of NaCl (0 to 1 M) in the same buffer. Protein-containing fractions were collected, pooled, and buffer-exchanged into 10 mM sodium phosphate buffer (pH 6.5) using Vivaspin 20 centrifugal concentrator tubes with a 10,000 MWCO membrane (Sartorius AG). The purity of the recombinant proteins was verified by SDS-PAGE on 10 % polyacrylamide gels. Protein concentration was estimated by absorbance at 420 nm (e420 = 130 mM−1 cm−1). Deglycosylation of the recombinant proteins was performed using Endoglycosidase H (Endo H) (New England Biolabs) as described previously [11].
Adsorption properties for crystalline cellulose. All experiments were conducted according to procedures described in previous studies [16]. Cellulose Iβ from Halocynthia was prepared using the following protocol. Halocynthia roretzi specimens were repeatedly treated with 5 % KOH and 0.3 % NaClO2 solutions for purification. Subsequently, hydrolysis was performed using 4 M hydrochloric acid, followed by centrifugation, water washing, and collection of the supernatant [27]. A reaction mixture containing 0.1 % (w/v) Halocynthia cellulose Iβ, 100 mM sodium acetate buffer (pH 5.2), and various concentrations of recombinant proteins (0, 0.2, 0.25, 0.5, 0.75, 1.0, 1.25, 1.5, 1.75, 2.0, 3.5 µM) in a total volume of 100 µL was incubated at 4 °C for 2 h. After incubation, the reaction mixture was filtered through a 96-well filter plate (MilliporeSigma, Burlington, MA, USA) to eliminate recombinant proteins bound to the substrate. The protein level in the filtrate was quantified by recording absorbance at 420 nm with a microplate reader (Fluoroskan Ascent, Thermo Fisher Scientific Inc.). All measurements were carried out in triplicate, and the mean values were plotted. The adsorption parameters were obtained by fitting the plots to the Hill equation using KaleidaGraph version 5.0 (Synergy Software, Reading, PA, USA).
UV-visible absorption spectrum analysis. All spectral analyses were carried out in 20 mM sodium phosphate buffer (pH 7.0) at room temperature. Ultraviolet-visible absorption spectroscopy of the purified recombinant protein was performed using a U-3900 spectrophotometer (Hitachi High-Tech Corporation, Tokyo, Japan). The reduced form of the protein was generated by adding 1 mM L(+)-ascorbic acid (Fujifilm Wako Pure Chemical Corp., Osaka, Japan) to achieve the desired working concentration.
Spectral variations in AtAA8 upon interaction with the AA12 domain of PDH from Coprinopsis cinerea (CcAA12) were analyzed using the following methodology. Hololization of CcAA12 was accomplished by incubating a solution comprising 5 μM CcAA12, 50 μM PQQ disodium salt, 1 mM CaCl2, and 50 mM sodium acetate buffer (pH 6.0) at 4 °C for 2 h. The formation of the holoenzyme was subsequently verified through UV-visible absorption spectroscopy. Unbound PQQ was removed using a Vivaspin 500 centrifugal concentrator equipped with a polyethersulfone membrane and a 5,000 MWCO (Sartorius AG), followed by buffer exchange with 50 mM sodium acetate (pH 6.0) containing 1 mM CaCl2. To assess electron transfer activity from CcAA12 to AtAA8, enzymatic assays were conducted according to previously established protocols [12]. A reaction mixture containing 0.05 μM holo-CcAA12, 2.04 μM AtAA8, 100 mM L(-)-fucose (Fujifilm Wako Pure Chemical Corp.), 1 mM CaCl2, and 50 mM sodium phosphate buffer (pH 6.0) was assembled, and spectral shifts were continuously recorded using a UV-visible spectrophotometer U-3900 (Hitachi High-Tech Corp.).
Measurement of midpoint potential. Following a previously reported procedure [28], AtAA8-immobilized electrodes were fabricated as working electrodes. A polycrystalline gold electrode (AuE) with a 1.6 mm diameter was modified using gold nanoparticles (AuNPs). The AuNPs, with particle sizes of 14-15 nm, were synthesized by citrate reduction of H4AuCl4, following the method of Frens [29].The resulting AuNP solution was concentrated by centrifugation at 10,000 × G for 30 min at 4 °C, and the supernatant was discarded to obtain a 50-fold concentrated AuNP dispersion. One microliter of the concentrated AuNP solution was drop-cast onto the AuE surface and then air-dried. Three AuNP-coated electrodes (AuNPs/AuEs) were prepared by repeating this process three times. This procedure generates three-dimensionally structured electrode surfaces with a high surface area-to-volume ratio, thereby increasing protein loading. AuNPs/AuEs were cleaned in 500 mM sulfuric acid by cyclic voltammetry between −0.2 and +1.5 V vs. Ag|AgCl (3 M NaCl) for 20 cycles at a scan rate of 0.1 V s-1 prior to modification with a self-assembled monolayer (SAM). The electrode surface was rinsed and then immersed in 20 mM 6-mercapto-1-hexanol for 3 h at room temperature to form the SAM. The SAM-modified electrode was rinsed with Milli-Q (Merck KGaA, Darmstadt, Germany) water to remove excess thiol molecules. The AtAA8 domain was adsorbed onto SAM-modified AuNPs/AuEs by applying 2 μL of purified AtAA8 solution and incubating overnight at 4 °C. Cyclic voltammetry (CV) measurements were conducted using an ALS Model 612Dx electrochemical analyzer (BAS Inc., Tokyo, Japan). A conventional three-electrode cell was employed, with a platinum wire as the counter electrode and Ag|AgCl (3 M NaCl) as the reference electrode. All potentials reported in this study are referenced to the Ag|AgCl (3 M NaCl) electrode, which corresponds to +209 mV vs. the normal hydrogen electrode. CV measurements were carried out in 50 mM buffer solutions used as electrolytes: citrate buffer (pH 3), sodium acetate buffer (pH 4-6), sodium phosphate buffer (pH 7), and Tris-HCl buffer (pH 8-9) [12].
We identified a putative gene encoding an AA8 domain appended with a CBM104 in the A. tabescens genome using BLASTp. The transcript was subsequently cloned and sequenced, confirming that it represents a single open reading frame lacking other catalytic domains such as AA3 or AA12. A multiple sequence alignment was performed using this sequence alongside several characterized AA8 domains (Figs. 1A and B). The analysis revealed that the A. tabescens sequence retains key conserved residues typical of known AA8 domains, including methionine and histidine residues that coordinate the fifth and sixth ligands of the heme iron, as well as cysteine residues involved in disulfide bond formation, contributing to structural stability. Additionally, the sequence preserves a tyrosine residue (Y93), proposed to play a critical role in electron transfer from the AA3 domain to the AA8 domain and subsequently to LPMOs in CDHs [30]. These findings suggest that the A. tabescens sequence constitutes an AA8 domain that coordinates a heme group and functions as an electron transfer domain. Subsequently, the amino acid sequence appended to the C-terminus of the AA8 domain was compared with the CBM104 domain previously characterized in LPMO9 from G. trabeum (GtCBM104), revealing a sequence identity of 72.7 %. Notably, four cysteine residues responsible for forming two disulfide bonds reported to be essential for specific binding to cellulose I were conserved [31]. Furthermore, among the residues previously identified as conserved in CBM104 homologs, three aromatic, two acidic, and four basic residues were also conserved in the C-terminal region of this protein (Fig. 1C) [16]. From here onward, this protein is called AtAA8-CBM104.
(A) Domain architecture of AtAA8-CBM104 and five known cytochrome proteins with different domain architectures. CBCyt.b562 (AA8-CBM1) from the basidiomycete Phanerodontia chrysosporium; PcCDH, a CDH from the basidiomycete P. chrysosporium; CcPDH, a PDH from Coprinopsis cinerea; MtCDH, a CDH from the ascomycete Myriococcum thermophilum; NcCDH, a CDH from Neurospora crassa. (B) The amino acid sequence of AtAA8-CBM104 from Armillaria tabescens (residues 1-188) was aligned with the representative five AA8 domains. The methionine residue coordinating the fifth position of the heme iron is highlighted in magenta, the histidine residue coordinating the sixth position in blue, cysteine residues forming a disulfide bond in orange, and the tyrosine residue important for electron transfer in green. (C) Alignment of AtAA8-CBM104 (residues 274-328) with a known CBM104. The CBM104 sequence from an LPMO9 of the basidiomycete Gloeophyllum trabeum (GtCBM104; residues 296-351) was used as a reference. Thirteen amino acid residues highly conserved among CBM104 homologs are highlighted in red.
To investigate their respective functions, both the full-length AtAA8-CBM104 and the AtAA8 domain alone were heterologously expressed as recombinant proteins in the yeast K. phaffii. The proteins were purified via column chromatography, treated with EndoH, and subsequently analyzed by SDS-PAGE. Although Endo H treatment reduced the apparent molecular weights of both the AtAA8 domain and AtAA8-CBM104, they still migrated at higher molecular weights than the theoretical values (19.6 and 33.2 kDa, respectively), indicating that both proteins are likely subject to extensive O-linked glycosylation (Fig. S1; see J. Appl. Glycosci. Web site).
The GtCBM104 appended to LPMO9 exhibits high, specific binding affinity for native crystalline cellulose [16]. To investigate CBM104 function in this context, the binding ability of AtAA8-CBM104 to native crystalline cellulose, specifically Halocynthia cellulose Iβ, was examined. AtAA8 alone showed little or no binding whereas AtAA8-CBM104 did the opposite (Fig. 2). The adsorption parameters obtained by fitting the adsorption plots of AtAA8-CBM104 to the Hill equation were compared with previously reported values for GtCBM104 and the CBM1 domain of cellobiohydrolase from Trichoderma reesei (TrCBM1) (Table 1) [16]. The dissociation constant (Kd) of AtAA8-CBM104 was comparable to that of GtCBM104 and approximately one order of magnitude lower than that of TrCBM1. Conversely, despite the higher adsorption efficiency of AtAA8-CBM104 compared to TrCBM1, it was approximately one order of magnitude lower than that of GtCBM104. Compared with the CBM104 appended to AA9, the CBM104 appended to AtAA8 contains a C-terminal flexible region enriched in basic amino acids such as arginine (Fig. S2; see J. Appl. Glycosci. Web site). Such structural features have been associated with reduced adsorption efficiency, suggesting that the lower adsorption efficiency of AtAA8-CBM104 is likely attributable to this C-terminal region [31]. The lower adsorption efficiency of AtAA8-CBM104 relative to GtCBM104 appended to LPMO9 may reflect functional differences. Since LPMO9 directly catalyzes cellulose degradation, the presence of a CBM104 with high adsorption efficiency is advantageous for maintaining frequent contact with the cellulose surface. In contrast, although AA8 must localize to cellulose to transfer electrons to cellulose-bound LPMO9, its excessive retention on the cellulose surface may interfere with LPMO9 catalysis. Therefore, a CBM104 with lower adsorption efficiency than the one appended to LPMO9 may be better suited to AA8’s functional role.
The adsorption plots are shown as follows: closed red circles, AtAA8-CBM104; closed blue squares, AtAA8. Each measurement was performed in triplicate, and the plotted values represent the mean of the three measurements. The data for AtAA8-CBM104 were fitted to the Hill equation. The data for AtAA8 were not fitted, as it exhibited negligible adsorption.
Table 1. Adsorption parameters obtained via fitting to the Hill equation.
| CBMs | Amax [μmol/g cellulose] | n | [μM−1] | Kd [μM] | Adsorption efficiency [L/g-cellulose] |
| AtAA8-CBM104 | (3.60 ± 0.50) × 10−1 | (1.70 ± 0.56) × 100 | (7.06 ± 7.01) × 100 | 3.17 × 10−1 | 1.14 × 100 |
| GtCBM104 | (3.22 ± 0.31) × 100 | (8.60 ± 1.30) × 10−1 | (4.39 ± 1.96) × 100 | 1.79 × 10−1 | 1.80 × 101 |
| TrCBM1 | (3.54 ± 1.36) × 100 | (8.30 ± 1.20) × 10−1 | (3.30 ± 1.80) × 10−1 | 3.80 × 100 | 9.31 × 10−1 |
The values for AtAA8-CBM104 were obtained in the present study. For GtCBM104 and TrCBM1, which were fused to a red fluorescent protein, the values were taken from the following publication [26]. The adsorption parameters obtained by fitting the data to the Hill equation were rounded to three significant figures.
The absorption spectrum of purified AtAA8 is presented in Fig. 3A. A Soret band peak was observed at 421 nm, and upon reduction with ascorbic acid, the α- and β-bands emerged at 564 nm and 534 nm, respectively, accompanied by a red-shifted Soret band at 430 nm. A comparable absorption profile was recorded for purified AtAA8-CBM104 (data not shown). These findings indicate that purified AtAA8 is in an oxidized state and contains a b-type cytochrome, consistent with AA8 domains found in characterized CDHs and PDHs. The redox potential of the AA8 domain was determined via protein film voltammetry. As illustrated in Fig. 3B, redox peak currents attributed to the heme b of AtAA8 were detected. At pH 7.0, the midpoint potential was calculated as −105 ± 3 mV. The peak widths at half-height for oxidation and reduction were 85 mV and 81 mV, respectively (from Fig. 3B), indicating that these values are consistent with a 1e− process (theoretical value: 90 mV under these conditions). The potential declined linearly from pH 3.0 to 6.0, with a slope of −23 mV/pH, and remained stable between pH 6.0 and 8.0 (Fig. 3C). These measurements align with those reported for AA8 domains of CDHs from the basidiomycete P. chrysosporium and the ascomycete Myriococcum thermophilum, as well as for the AA8 domain of PDH from the basidiomycete C. cinerea, suggesting that AtAA8 can accept electrons from the AA3 or AA12 and donate them to LPMO9 [32, 33, 34]. To experimentally test this possibility, we conducted an electron transfer assay by incubating AtAA8 with the AA12 domain. Since A. tabescens lacks a gene encoding an AA12 domain, CcAA12 was utilized. The spectral changes of AtAA8 upon incubation with CcAA12 and L-fucose (the substrate of AA12) are shown in Fig. 4. Compared to the spectrum obtained prior to mixing AtAA8 with the reaction solution, the absorbances of the α- and β-bands―characteristic of reduced cytochrome b―increased approximately 5 min after mixing. These absorbances further increased around 40 min post-incubation. These results demonstrate that AtAA8 was progressively reduced, suggesting its ability to accept electrons generated by the external AA12 protein during L-fucose oxidation. Thus, AtAA8-CBM104 likely receives electrons from electron-donating proteins such as AA12 and transfers them to LPMO9 on cellulose, thereby facilitating its catalytic activity.
(A) UV-visible absorption spectra of AtAA8. Solid line: oxidized form; dotted line: reduced form upon addition of ascorbic acid. All spectra were recorded in 20 mM sodium phosphate buffer (pH 7.0) at room temperature. (B) Cyclic voltammograms of AtAA8 immobilized on 6-mercapto-1-hexanol SAM-modified AuNPs/AuEs. The measurements were performed in 50 mM HEPES buffer, pH 7.0, at a scan rate of 100 mV/s. Raw data are shown as solid lines, and baseline-subtracted data as dashed lines. (C) Plots of the midpoint potentials of AtAA8 immobilized on 6-mercapto-1-hexanol SAM-modified AuNPs/AuEs against pH. The buffers used for these measurements were citrate buffer (pH 3), sodium acetate buffer (pH 4-6), sodium phosphate buffer (pH 7), and Tris-HCl buffer (pH 8-9).
Solid line: spectrum immediately after the addition of the reaction mixture; long dashed line: spectrum approximately 5 min after the addition of reaction mixture; dotted line: spectrum approximately 40 min after the addition of reaction mixture.
Finally, a phylogenetic analysis of fungal AA8 sequences was performed to determine whether the functional characteristics of AtAA8-CBM104 also apply to other fungal cytochrome proteins containing both an AA8 domain and CBM104. Using the amino acid sequences of AtAA8 and 20 representative AA8 domains as queries, a BLASTp search yielded 630 sequences. Non-AA8 domain sequences were removed, and the remaining sequences were subjected to molecular phylogenetic analysis. The resulting phylogenetic tree was broadly divided into six clades (A-F) (Fig. 5A and Fig. S3; see J. Appl. Glycosci. Web site). Clades A-C consisted of AA8 domains from basidiomycetes, clades D and F consisted of those from ascomycetes, and clade E consisted of AA8 domains from both phyla. All AA8 domains lacking catalytic domains and appended with CBM104 belong to clade E, and the AA8 sequence of AtAA8-CBM104 is also classified within this clade. A notable feature characterizing the six clades is variation at the residue immediately preceding the axial histidine, a position known to locally affect the heme’s surface charge (Fig. 5B). Clade E, to which AtAA8 belongs, exhibited high conservation of neutral aromatic residues, such as tyrosine or phenylalanine, at this position. Since the surface charge surrounding the heme significantly influences electron transfer to donors such as LPMOs [13], AA8 domains within the same clade are likely to interact with electron donors possessing similar electrostatic characteristics.
(A) Midpoint rooted tree based on fungal AA8 sequences. Sequences corresponding to the catalytic domain, linker region, and binding domain were excluded from the analysis. AtAA8-CBM104 is indicated by an arrow. The phylogenetic tree was broadly divided into six clades (A-F). The taxonomic distribution of the organisms harboring the sequences is indicated by the background color, and the domains constituting the sequences are denoted by the color of the terminal labels. Bootstrap support values are shown as pink circles on the branches (see legend in the figure). (B) Sequence logos generated using WebLogo for each clade corresponding to residues 250-270 of AtAA8. Aromatic residues are shown in green, histidine in red, acidic residues in blue, basic residues in magenta, and neutral non-aromatic residues in black.
In this study, we conducted a functional analysis of AtAA8-CBM104, which lacks a catalytic domain. The results demonstrated that AtAA8 coordinates a b-type cytochrome in a manner similar to known AA8 domains and can accept electrons from AA12. The CBM104 domain appended to AtAA8 was shown to bind to cellulose I. In addition, phylogenetic analysis suggested that all non-catalytic AA8 proteins appended with CBM104, including AtAA8-CBM104, share similar electrochemical properties. These findings indicate that AA8-CBM104s likely function by receiving electrons from external electron-donating proteins and transferring them to cellulose-bound LPMO9, thereby enhancing its catalytic activity. Future studies will be required to experimentally validate this hypothesis.
The authors declare no conflicts of interest.
This study was supported by Grant-in-Aid for Challenging Research (Exploratory) (Grant Number 23K18046), Grant-in-Aid for Scientific Research (B) (Grant Number 25K02069), and Grant-in-Aid for Scientific Research (A) (Grant Number 23H00341) from the Japan Society for the Promotion of Science (JSPS). Additionally, we thank the 1000 Fungal Genomes consortia for access to unpublished genome data. The genome sequence data were produced by the US Department of Energy Joint Genome Institute in collaboration with the user community.
