Multimodular flavobacterial enzymes specialised in coordinated decomposition of cellulose and alginate in brown algal cell walls.

Multimodular flavobacterial enzymes specialized in coordinated decomposition of cellulose and alginate in brown algal cell walls - PMC Skip to main content An official website of the United States government Here's how you know Here's how you know Search Log in Primary site navigation Search Log in Search PMC Full-Text Archive Search in PMC Journal List User Guide As a library, NLM provides access to scientific literature. Inclusion in an NLM database does not imply endorsement of, or agreement with, the contents by NLM or the National Institutes of Health. Learn more: PMC Disclaimer | PMC Copyright Notice ISME J . 2026 May 8;20(1):wrag112. doi: 10.1093/ismejo/wrag112 Multimodular flavobacterial enzymes specialized in coordinated decomposition of cellulose and alginate in brown algal cell walls Fei Xu 1, # , Xiao-Hui Sun 2, # , Xiao-Dong Zhang 3, # , Xiao-Fei Wang 4 , Yan Wang 5 , Hai-Yan Cao 6 , Peng Wang 7, 8 , Jian-Xun Li 9 , Xi-Ying Zhang 10 , Qi-Long Qin 11 , Xiu-Lan Chen 12 , Yu-Zhong Zhang 13, 14, 15, ✉ , Yin Chen 16, 17, 18, ✉ , Yu-Qiang Zhang 19, ✉ Author information Article notes Copyright and License information 1 State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China 2 State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China 3 State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China 4 State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China 5 State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China 6 MOE Key Laboratory of Evolution and Marine Biodiversity, Frontiers Science Center for Deep Ocean Multispheres and Earth System & College of Marine Life Sciences, Ocean University of China, Qingdao 266203, Shandong Province, China 7 MOE Key Laboratory of Evolution and Marine Biodiversity, Frontiers Science Center for Deep Ocean Multispheres and Earth System & College of Marine Life Sciences, Ocean University of China, Qingdao 266203, Shandong Province, China 8 Laboratory for Marine Biology and Biotechnology, Qingdao Marine Science and Technology Center & Laoshan Laboratory, Qingdao 266237, Shandong Province, China 9 State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China 10 State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China 11 State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China 12 State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China 13 MOE Key Laboratory of Evolution and Marine Biodiversity, Frontiers Science Center for Deep Ocean Multispheres and Earth System & College of Marine Life Sciences, Ocean University of China, Qingdao 266203, Shandong Province, China 14 Laboratory for Marine Biology and Biotechnology, Qingdao Marine Science and Technology Center & Laoshan Laboratory, Qingdao 266237, Shandong Province, China 15 Marine Biotechnology Research Center, State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China 16 MOE Key Laboratory of Evolution and Marine Biodiversity, Frontiers Science Center for Deep Ocean Multispheres and Earth System & College of Marine Life Sciences, Ocean University of China, Qingdao 266203, Shandong Province, China 17 School of Life Sciences, University of Warwick, Coventry CV4 7AL, West Midlands, United Kingdom 18 School of Biosciences, University of Birmingham, Edgbaston B15 2TT, Birmingham, United Kingdom 19 State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China ✉ Corresponding authors. Yu-Zhong Zhang, MOE Key Laboratory of Evolution and Marine Biodiversity, Frontiers Science Center for Deep Ocean Multispheres and Earth System & College of Marine Life Sciences, Ocean University of China, 238 Songling Road, Qingdao 266003, Shandong Province, China. E-mail: zhangyz@sdu.edu.cn ; Yin Chen, MOE Key Laboratory of Evolution and Marine Biodiversity, Frontiers Science Center for Deep Ocean Multispheres and Earth System & College of Marine Life Sciences, Ocean University of China, 238 Songling Road, Qingdao 266003, Shandong Province, China. E-mail: y.chen.22@bham.ac.uk ; Yu-Qiang Zhang, State Key Laboratory of Microbial Technology, Shandong University, 72 Binhai Road, Qingdao 266237, Shandong Province, China. E-mail: yqzh2024@sdu.edu.cn # Fei Xu, Xiao-Hui Sun and Xiao-Dong Zhang contributed equally to this work. Received 2025 Dec 19; Revised 2026 Apr 12; Accepted 2026 May 5; Collection date 2026 Jan. © The Author(s) 2026. Published by Oxford University Press on behalf of the International Society for Microbial Ecology. This is an Open Access article distributed under the terms of the Creative Commons Attribution License ( https://creativecommons.org/licenses/by/4.0/ ), which permits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited. PMC Copyright notice PMCID: PMC13235733  PMID: 42102175 Abstract Brown algal cell walls are complex matrices composed primarily of alginate, cellulose, and fucoidan. Their depolymerization is important in marine carbon cycling. Although numerous algal polysaccharide-degrading enzymes have been characterized, most studies focus on breaking down single, purified polysaccharides, leaving the degradation mechanisms of native cell walls containing mixed polysaccharides poorly understood. Here, we report the integrated modular enzymes involved in brown algal cell wall polysaccharide (BACWP) degradation. Using the marine flavobacterium Aquimarina sp. 2-A2 as a model, we isolated a bifunctional enzyme, CelAly, which integrates a glycoside hydrolase family 5 cellulase domain and a polysaccharide lyase family 31 alginate lyase domain within a single polypeptide, enabling the degradation of cellulose and alginate in brown algal cell walls. In vivo relevance of CelAly was confirmed by upregulation of its gene during growth on algal biomass. CelAly also contains three distinctive substrate-binding modules (B1, B2, UKD) that support its multimodular functionality; among these, UKD is notable for its dual substrate-binding capability. CelAly’s modular architecture and interdomain flexibility may facilitate coordinated degradation of BACWPs. Bioinformatic analyses and biochemical validation revealed three additional types of such modular enzymes from marine microbes. CelAly and related modular enzymes are strongly associated with marine environments and exhibit conserved modular strategy for substrate recognition and catabolism. Thus, these enzyme architectures represent a previously unrecognized strategy specialized for BACWP decomposition. This study elucidates the unique structural and functional adaptations of the integrated multimodular enzymes and highlights their ecological prevalence among marine bacteria, providing insights into natural biomass decomposition. Keywords: marine algae, cell wall polysaccharides, Flavobacteriaceae, polysaccharide-degrading enzyme, modular enzyme Introduction Polysaccharides constitute the primary structural scaffold of plant and algal cell walls, serving as both mechanical barriers and carbon reservoirs for microbial communities [ 1 , 2 ]. These polysaccharides play essential roles in the global carbon cycle and serve as attractive substrates for biotechnological applications [ 3 , 4 ]. However, their structural complexity poses a significant challenge for efficient utilisation by microbes. To overcome this structural complexity, microorganisms have evolved sophisticated enzymatic systems mainly relying on diverse carbohydrate-active enzymes (CAZymes) [ 5 , 6 ]. A prominent example is the efficient multimodular cellulase CelA from Caldicellulosiruptor bescii , which combines two catalytic domains with endoglucanase and exoglucanase activities, along with multiple carbohydrate-binding modules (CBMs) [ 7 ]. Another case is CbXyn10C/Cel48B, which features a glycoside hydrolase family 10 (GH10) domain (a typical xylanase domain) and a glycoside hydrolase family 48 (GH48) domain (a canonical cellulase domain), both exhibiting bifunctional activity against xylan and cellulose [ 8 ]. These examples demonstrate that multimodular CAZymes act as powerful tools for complex terrestrial plant cell wall degradation by combining domains with closely related functional activities. In marine ecosystems, macroalgae represent major contributors to primary production, releasing vast amounts of organic matter through seasonal blooms and detrital export [ 9 , 10 ]. These inputs trigger successions of heterotrophic bacteria, with members of Bacteroidota (particularly flavobacteria) acting as dominant degraders [ 11 , 12 ]. Their ecological success relies on CAZyme-rich genomes, extensive polysaccharide utilization loci (PULs) coupled to SusCD-like uptake systems, and the presence of the type IX secretion system (T9SS) [ 12 , 13 ]. These organisms therefore play a central ecological role in determining not only the rate of macroalgal carbon turnover but also the chemical form and size distribution of released carbohydrates (e.g. oligosaccharides versus larger particulate fragments), with implications for cross-feeding interactions, microbial loop efficiency, and carbon cycling [ 14 , 15 ]. Model flavobacteria, such as Zobellia galactanivorans , exemplify how these mechanisms support the breakdown of fresh macroalgae, releasing metabolites that serve as public goods for the broader microbial community [ 12 ]. However, although we understand how these bacteria process individual polysaccharides [ 16–19 ], it remains unclear how they process the intricate physical architecture of the intact cell wall. The bacterial decomposition of brown algae ( Phaeophyceae ) may serve as a model for studying the bacteria–alga interactions, owing to the high biomass and complex cell wall matrix of brown algae [ 6 , 9 ]. This matrix consists of alginate [a gel-forming heteropolymer of β-D-mannuronic acid (M) and α-L-guluronic acid (G)] embedding cellulose microfibrils (β-1,4-linked glucan) and other polysaccharides such as fucoidan (a sulfated fucose-rich polysaccharide) [ 20 ]. This architecture creates a pronounced accessibility barrier, implying that successful decomposition requires more than isolated single-function CAZymes. Though some specialists, like Verrucomicrobia, manage this by encoding hundreds of enzymes for specific polymers like fucoidan [ 21 ], it is unclear whether flavobacteria and even other abundant algal-associated bacteria employ distinct enzymatic strategies to overcome the structural heterogeneity of the entire brown algal cell wall matrix. In this study, we report the identification and ecological significance of the integrated multimodular enzyme CelAly that enables the coordinated breakdown of native brown algal cell wall polysaccharides (BACWPs). CelAly from the marine flavobacterium Aquimarina sp. 2-A2 comprises a cellulase domain (Cel5), an alginate lyase domain (Aly31) and three noncatalytic domains with distinct substrate-binding capacities. In vitro and in vivo experiments demonstrate that CelAly plays an important role in the bacterial degradation of brown algae. Integrated bioinformatic and biochemical analyses reveal that such CAZymes are particularly abundant in marine flavobacteria and represent an adaptive strategy employed by these bacteria to overcome the structural heterogeneity of algal cell walls, thereby shedding light on microbial degradation of native macroalgal biomass. Materials and methods Materials and strains Aquimarina sp. 433 was isolated from seaweed collected from seawater in Qingdao, China (120 o 29′42″E, 36 o 5′21″N), and Aquimarina sp. 2-A2 was isolated from a sediment sample collected from the Southwest Indian Ocean (50 o 37′56″E, 37 o 34′60″S). Both strains were preserved in our lab, and their genome sequences have been deposited to the GenBank database under accession numbers JBSWZK000000000.1 and JBJKFI000000000.1 , respectively. Escherichia coli strains were purchased from Vazyme (China). Phosphoric acid swollen cellulose (PASC) and heteropolymers of M and G (PMG) were prepared as previously described [ 22 , 23 ]. All other carbohydrate substrates were purchased at analytical grade. Detailed information on materials is provided in Supplementary Materials and Methods. Real-time quantitative PCR analysis To verify the role of the multimodular enzyme in brown algae decomposition, the strain Aquimarina sp. 433, which harbors the CELALY’ gene, was cultured in 2216E medium at 30°C to an OD 600 of 0.8. Cells were collected by centrifugation at 4000 × g for 10 min at 4°C, washed three times with a 3% sea salt solution, and then inoculated into the medium containing Laminaria japonica and 3% sea salt (pH 8.0). Bacterial samples were collected at 12, 24, and 36 h after inoculation. Total RNA was extracted by using the RNeasy Mini Kit (Qiagen, Germany), followed by cDNA synthesis with the TransScript All-in-One First-Strand cDNA Synthesis SuperMix (TransGen, China). RT-qPCR assays were subsequently performed on a LightCycler II 480 System (Roche, Switzerland) using SYBR Premix Ex Taq (TaKaRa, Japan), with the 16S rRNA gene used as the internal reference. Gene cloning, mutation, and protein expression, and purification Genes encoding CelAly (GenBank: WP_407141409.1 ), its truncated mutations, and selected BACWPs-degrading enzymes (Enzyme A–C) were amplified or synthesized and cloned into the pET-22b vector with a His-tag. Site-directed mutagenesis was performed using the QuikChange kit (Agilent, USA), and the recombinant plasmids were transformed into E. coli BL21 (DE3) for protein expression. Recombinant proteins were purified via nickel-affinity and gel filtration chromatography (GE Healthcare, Germany), and protein concentrations were measured using the bicinchoninic acid (BCA) protein assay kit (Thermo, USA). The primers for construction of recCelAly and its truncated mutations are listed in Table S1 , and detailed protocols are described in Supplementary Materials and Methods. Enzyme assays The glycoside hydrolase (GH) activities of recCelAly and its mutants, as well as other multimodular enzymes, were determined by the dinitrosalicylic acid (DNS) method [ 24 ]. The polysaccharide lyase (PL) activities were determined by the ultraviolet absorption method [ 25 ]. After confirming that E173A lacked cellulase activity and Y501A lacked alginate lyase activity, the abilities of both mutants to degrade alginate and cellulose-rich fractions of L. japonica , respectively, were assessed using the DNS method [ 24 ]. Detailed protocols are described in Supplementary Materials and Methods. Biochemical characterization The optimum temperature, pH, and NaCl concentration for recCelAly, Cel5, and Aly31 activities on carboxymethyl cellulose (CMC) or sodium alginate were determined by individually varying each reaction condition. Degradation products of recCelAly and its mutants toward CMC or sodium alginate were analyzed by high-performance liquid chromatography (HPLC) after 24-h reactions at 40°C. Kinetic parameters for depolymerization of PASC and sodium alginate were determined by nonlinear curve fitting based on the Michaelis–Menten equation using Origin 8.5 software. Substrate binding analysis Insoluble cellulose substrates (PASC, Avicel, and bacterial cellulose) were incubated with proteins, and unbound proteins in the supernatant were quantified by the BCA assay to calculate binding ratios. The UKD domain’s interaction with soluble cellohexaose was analyzed by isothermal titration calorimetry (ITC). Protein binding to alginate substrates, including sodium alginate, PMG, polyguluronate (PG), and polymanuronate (PM), was evaluated by gel filtration chromatography or by incubation with alginate gel beads, with bovine serum albumin (BSA) used as a control. Binding ratios were calculated as the fraction of protein bound relative to the total protein. Detailed protocols are described in Supplementary Materials and Methods. Crystallization, data collection, structure determination, and refinement B1 (the TM4 mutant, 10 mg/ml) was crystallized at 18°C in the buffer containing 0.1 M Bis-Tris propane (pH 8.5) and 28% (w/v) polyethylene glycol (PEG) 6000. UKD (the TM6 mutant, 8 mg/ml) was crystallized in the buffer containing 3.25 M sodium formate (pH 6.8). X-ray diffraction data were collected on the BL02U1 Beamline at Shanghai Synchrotron Radiation Facility (SSRF). The initial diffraction datasets were processed by XDS. Initial model building was finished by the Phenix program AutoBuild [ 26 ]. Refinement of the B1 and UKD structures was performed by the Phenix program Refine [ 26 ] and Coot [ 27 ] alternately. The statistics for data collection and the final model quality are presented in Table S2 . All the structure figures were processed using the program PyMOL. The structures of B1 and UKD have been deposited in PDB under identifiers 9KVU and 9KVN, respectively. Analysis of alginate assimilation genes The genomes of strains containing CelAly-like modular enzymes were downloaded from the National Center for Biotechnology Information (NCBI) database and then annotated using the rapid annotation using subsystem technology (RAST) server ( http://rast.nmpdr.org/ ). The prevalence of genes involved in alginate assimilation was statistically analyzed, including aly (alginate lyase gene), oal (oligoalginate lyase gene), kdgF (pectin degradation protein gene), sdr (short-chain dehydrogenase/reductase gene), kdgK (2-dehydro-3-deoxygluconokinase gene), and eda (2-dehydro-3-deoxyphosphogluconate aldolase gene). Small-angle X-ray scattering measurement Small-angle X-ray scattering (SAXS) data for CelAly were collected at SSRF BL19U2 and processed using BioXTAS RAW [ 28 ] and ATSAS [ 29 ] ( Table S3 ). Scattering profiles were obtained after buffer subtraction. Structural parameters were evaluated using Guinier approximation and GNOM [ 30 ]; flexibility was assessed by Kratky plots [ 31 ]. Ab initio envelopes were generated with DAMMIF [ 32 ] and DAMMIN [ 33 ]. Rigid-body modeling was performed with CORAL [ 34 ] using individual domain models (crystal structures for B1 and UKD and AlphaFold2 predictions for others), with theoretical curves compared to experimental data via CRYSOL [ 35 ] and models docked into envelopes using SUBCOMB [ 36 ]. Detailed protocols are described in Supplementary Materials and Methods. Bioinformatic analysis The residues encoding a putative signal peptide were predicted by the SignalP 5.0 [ 37 ]. Domain analysis was performed in the NCBI Conserved Domain Database (CDD) and dbCAN meta server [ 38 , 39 ]. Sequence alignments were performed by ClustalW, and alignment figures were processed using WebLogo ( http://weblogo.threeplusone.com ) [ 40 ]. A phylogenetic tree was constructed using the MEGA X software [ 41 ] with sequences of related CBMs. Specifically, the tree was constructed through the neighbor-joining approach with a bootstrap analysis of 1000 replicates. The three-dimensional structures of the Cel5, Aly31, and B2 domains of CelAly were predicted by AlphaFold2 with default settings [ 42 ], and the models with the highest predicted local distance difference test (pLDDT) scores were selected for further analysis. Results and discussion Identification of a multimodular enzyme from a marine flavobacterium for efficient degradation of native brown algal cell wall polysaccharides To investigate the enzymatic strategies adopted by marine bacteria for algal biomass decomposition, Aquimarina sp. 433, belonging to Flavobacteriaceae, was isolated from seaweed collected from seawater in Qingdao, China (120 o 29′42″E, 36 o 5′21″N). When cultured in artificial seawater supplemented with L. japonica (brown alga medium), Aquimarina sp. 433 exhibited rapid growth, concomitant with significant degradation of algal biomass, with nearly complete breakdown of the algal substrate observed within 36 h ( Fig. 1a ), indicating a potent algolytic capability. Figure 1. Open in a new tab Role of CELALY’ from Aquimarina sp. 433 in brown algae decomposition. (a) Decomposition of brown alga by Aquimarina sp. 433. Strain 433 was cultured at 30°C, 180 rpm in the brown alga medium, which contained 4 pieces (1 cm × 1 cm) of L. japonica blade and 3% sea salts. Samples were observed at 12, 24, and 36 h post-inoculation. The brown alga medium without strain 433 incubated was taken as the control. (b) Alginate lyase genes and alginate metabolic gene cluster in Aquimarina sp. 433. kdgF , pectin degradation protein gene, sdr , short-chain dehydrogenase/reductase gene. kdgK , 2-dehydro-3-deoxygluconokinase gene. eda , 2-dehydro-3-deoxyphosphogluconate aldolase gene. (c) RT-qPCR analysis of the transcriptional levels of CELALY’ and alginate metabolic genes of Aquimarina sp. 433 in response to brown alga. The 16S rRNA gene was used as an internal reference. The relative expression levels were normalized to the corresponding gene expression in 2216E medium at the same time points. The graphs show data from triplicate experiments (mean ± SD). Genome analysis revealed that Aquimarina sp. 433 encodes 13 putative alginate lyases, likely important for brown algal degradation. Among these, seven are scattered throughout the genome, whereas the remaining six are organized into two distinct gene clusters, designated Cluster 1 and Cluster 2 ( Fig. 1b ). Cluster 2 resembles a classical alginate utilization locus, containing genes encoding transporters, a regulator, and enzymes involved in monomer assimilation [ 43 , 44 ] ( Fig. 1b ). In contrast, Cluster 1 primarily consists of CAZyme-encoding genes ( Fig. 1b ). Within Cluster 1, we noticed a gene (designated CELALY’ ) that encodes a hexa-modular protein containing a GH family 5 subfamily 2 (GH5_2) domain and a PL family 31 (PL31) domain, as well as 4 other domains. Given that GH5_2 enzymes typically act on cellulose and PL31 enzymes on alginate [ 45 ], two important components of brown algal cell walls, this architecture suggested a potential role in concerted cell wall deconstruction. To examine this, we monitor its expression along with that of several other functionally relevant genes during algal decomposition. All tested genes, including CELALY’ , were upregulated during the early growth phase in brown alga medium, supporting the involvement of CELALY’ in the degradation process ( Fig. 1c ). Due to difficulties in purifying recombinant enzyme encoded by CELALY’ , we turned to its homolog CelAly from Aquimarina sp. 2-A2, which shares 69.0% amino acid sequence identity and retains all conserved domains with CELALY’. Aquimarina sp. 2-A2 was isolated from a sediment sample collected from the Southwest Indian Ocean (50 o 37′56″E, 37 o 34′60″S). In contrast with strain 433, strain 2-A2 cannot utilize brown algae for growth. We hypothesize that strain 2-A2 may have lost the metabolic genes during evolution due to the absence of brown algae in its deep-sea habitat. Alternatively, strain 2-A2 may produce CelAly to degrade algal polysaccharides without utilizing the released products, which may instead be assimilated by other bacteria acting as exploiters, thereby facilitating the assembly of polysaccharide-degrading communities [ 46 ]. Consistent with CELALY’, CelAly contains a GH5_2 domain (Cel5) and a PL31 domain (Aly31) ( Fig. 2a ). In addition, CelAly also contains three conserved domains for which no enzymatic activity has been reported including two bacterial immunoglobulin-like (Big) domains (B1 and B2) and one UKD domain, as well as a Por secretion system C-terminal sorting domain (CTD) domain likely involved in T9SS-dependent secretion ( Fig. 2a ) [ 47 ]. To confirm the function of CelAly, we expressed and purified the recombinant CelAly (recCelAly) lacking the signal peptide and the CTD domain ( Fig. S1 ). A range of polysaccharides known to serve as substrates for GHs and PLs ( Table S4 ) were tested to assess the substrate selectivity of CelAly. recCelAly showed significant GH activity toward barley β-glucan, hydroxyethyl cellulose (HEC), CMC, and PASC but showed much lower activity against xylans ( Fig. 2b ), indicating that recCelAly functions as a cellulase. For PL-related substrates, recCelAly was exclusively active against alginate substrates, including PM, PMG, and sodium alginate, but not PG ( Fig. 2c ), indicating that CelAly cleaves the glycosidic bonds between M and G, and M-M linkages but not G-G linkages in alginate. Collectively, these results demonstrate that CelAly is a bifunctional enzyme with both cellulase and alginate lyase activities. Figure 2. Open in a new tab Activities of CelAly toward brown algal cell wall polysaccharides. (a) Schematic diagram of CelAly and its truncated mutants. The signal peptide was predicted by SignalP 5.0 Server. The domain composition was analyzed by the NCBI’s Conserved Domain Database and dbCAN meta server. Cel5, the domain from GH5 family; Aly31, the domain from PL31 family; B1 and B2, two bacterial Ig-like domains of group 7 (Big_7 domains); UKD, an unknown functional domain; CTD, a Por secretion system C-terminal sorting domain. (b) Substrate specificity of recCelAly toward cello- and xylo-configured polysaccharides. CMC, carboxymethyl cellulose. HEC, hydroxyethyl cellulose. PASC, phosphoric acid swollen cellulose. (c) Substrate specificity of recCelAly toward various polysaccharides. PMG, heteropolymers consisting of M and G alternately. PM, polymannuronate. PG, polyguluronate. (d) Cellulase activities of recCelAly and its truncated mutants toward PASC. (e) Alginate lyase activities of recCelAly and its truncated mutants toward sodium alginate. (f) The degradation of L. japonica by CelAly mutants. Pictures are representative of three independent experiments. The graphs in (b)–(e) show data from triplicate experiments (mean ± SD). Further domain analysis using a series of truncated mutants (TM1–TM6) ( Fig. S1 ) showed that TM2, containing the Cel5 domain, retained cellulase activity comparable to recCelAly, whereas TM1, which lacked the Cel5 domain, showed no cellulase activity ( Fig. 2a and d ), confirming that Cel5 is responsible for cellulose degradation. Similarly, TM3 demonstrated alginate lyase activity, whereas the other mutants did not, indicating that Aly31 domain is responsible for alginate degradation ( Fig. 2a and e ). These results demonstrated that CelAly’s bifunctional activity is driven by its Cel5 and Aly31 domains and that the B1, B2, and UKD domains are not catalytic domains because their removal had minimal impact on substrate specificity ( Figs S2a and b ), degradation products ( Fig. S2c and d ), or optimal reaction conditions ( Fig. S2e–g ). We further verified CelAly’s activity against native BACWPs in vitro using the representative brown alga L. japonica as an example. Because L. japonica cell walls are rich in alginate but contain low levels of cellulose [ 48 ], we performed a sequential degradation assay to distinguish between the two activities. To facilitate this approach, the active-site mutants for both catalytic modules (Cel5: E173A and E259A; Aly31: Y501A and K538A) were constructed based on structural and sequence alignments with their homologs. Activity determination confirmed that E173A and Y501A completely lost the cellulase and alginate lyase activities of recCelAly, respectively ( Table S5 ), thereby enabling selective inactivation of one catalytic module while retaining the activity of the other within the intact multimodular enzyme. In the sequential assay, L. japonica was first treated with the mutant E173A to selectively degrade alginate. The generation of reducing sugars was detected by colour development via the DNS method ( Fig. 2f ). Next, a cellulose-enriched fraction was prepared by removing alginate via hot water extraction, followed by incubation with the Y501A mutant. This treatment also produced significant DNS-reactive products ( Fig. 2f ). These results demonstrate that CelAly is a BACWPs-degrading enzyme capable of breaking down both alginate and cellulose in brown algal cell walls during algal decomposition. Diversity and distribution of CelAly homologs CelAly shares 61% sequence identity with ClGP from Cellulophaga lytica and CelA from Cellulophaga sp. RHA 52, both are multimodular enzymes from the ocean ( Fig. S3 ) [ 49 , 50 ]. ClGP shows cellulase, alginate lyase, and glucuronan lyase activities, whereas CelA exhibits activities toward cellulose, laminarin, and alginate. However, their ability toward native BACWPs has not been reported. Given their sequence similarity to CelAly, it is plausible that they may also function as BACWPs-degrading enzymes. Therefore, the multimodular architecture of CelAly-like enzymes likely represents an adaptive strategy for efficient degradation of BACWPs in natural marine environments. To further analyze the ecological significance of such enzymes, we searched the homologs of CelAly against the NCBI nonredundant protein sequences (nr) database (coverage ≥75% and identity ≥40%). A total of 58 homologs were identified, and their conserved domain architectures were further analyzed ( Fig. 3 ). Due to the similarity between domains B1 and B2 that hinders their distinction, both were categorized as Big domains, while the number of Big domains was counted. The result showed that all CelAly homologs contain the catalytic GH5 and PL31 domains, as well as the noncatalytic UKD domain and a T9SS-related CTD domain, with most also carrying one or two Big domains. All the homologs are exclusively within Bacteroidota . The majority of these homologs (55/58) are from the Flavobacteriaceae family, particularly the genus Aquimarina . (36/58), others are from Marinigracilibium of Flammeovirgaceae (1/58) and Neolewinella of Lewinellaceae (2/58). These homologs are predominantly marine in origin and enriched in seaweed-associated habitats ( Fig. 3 ), suggesting a key role in BACWPs degradation. Figure 3. Open in a new tab Distribution of CelAly and its homologous proteins in bacteria. The homologs of CelAly were searched against the NCBI nr database using a threshold of ≥75% coverage and ≥40% identity. Domain composition was analyzed using the NCBI Conserved Domain Database (CDD). Filled boxes indicate the presence of a domain, whereas empty boxes indicate its absence. GH5, glycoside hydrolase family 5; PL31, polysaccharide lyase family 31. Big, bacterial Ig-like domain; UKD, domain of unknown function. CTD, a Por secretion system C-terminal sorting domain. The number of Big domains is indicated by the numbers. The restricted phylogenetic distribution of CelAly homologs implied that the enzymes may not be a general feature of marine polysaccharide degraders but rather a specialized enzymatic strategy for marine Flavobacteriaceae. Members of Aquimarina and related Flavobacteriaceae are frequently enriched on macroalgal surfaces and in decaying seaweed biomass [ 51 , 52 ], environments with high concentrations of structurally intact BACWPs. At such particle-associated niches, efficient access to insoluble, composite substrates is critical for resource acquisition. The conserved fusion of GH5 and PL31 domains may contribute to coordinated depolymerization of cellulose and alginate directly on the algal surface. The universal presence of the CTD domains among CelAly homologs further supports this interpretation because T9SS-mediated secretion and surface anchoring are hallmarks of Bacteroidota specialized in degrading insoluble biopolymers [ 53 ]. The restricted taxonomic distribution, conserved multimodular architecture, and functional activity against native BACWPs suggest that CelAly-like enzymes may represent a niche-adapted solution for BACWPs degradation, likely contributing to localized carbon turnover during macroalgal decomposition in marine ecosystems. Noncatalytic accessory domains of CelAly show novel polysaccharide-binding mechanisms and enhance CelAly’s affinities for BACWPs Given that the B1, B2, and UKD domains of CelAly are noncatalytic, we next investigated whether they contribute to the binding of BACWPs. Both domains B1 and B2 specifically interacted with alginate and alginate gel beads but not linear alginate derivatives (PMG, PM, or PG) or the cellulose substrate PASC ( Fig. S4a–c ), indicating a requirement for a 3D alginate gel network; notably, B1 showed stronger binding ( Fig. S4c ). Structure analysis revealed that B1 adopts a typical Ig-like β-sandwich fold with a positively charged, flat surface ( Fig. S5a and b , Table S2 ), unlike the weaker-binding B2 ( Fig. S5b ) and the cleft- or pocket-type alginate-binding CBMs ( Fig. S5c ) [ 54–56 ]. Key residues Lys40 B1 , Arg49 B1 , and Arg75 B1 (residue numbering refers to positions within the indicated domain) were confirmed to be essential for binding via this binding interface ( Fig. S5d–f). The UKD domain, located immediately downstream of B2 ( Fig. 1a ), has no previously known function. We analyzed its binding capacity to various polysaccharides and found that it exhibited a distinct substrate binding pattern compared to B1. In addition to selectively binding to sodium alginate and alginate gel beads ( Fig. 4a and b ), UKD also bound to insoluble cellulose substrates, such as PASC and Avicel ( Fig. 4c ), but showed no binding to soluble cellohexaose ( Fig. 4d ). These results suggest that UKD is a unique domain capable of binding both insoluble cellulose and native alginate. Typically, CBMs can only recognize one substrate type (e.g. polysaccharides with identical or structurally similar units) [ 57 ]. Thus, UKD represents the first reported instance of a single domain capable of recognizing two chemically distinct polysaccharides. This dual substrate-binding capability, likely an evolutionary adaptation, allows CelAly to efficiently degrade BACWPs by targeting multiple substrates within the complex algal cell walls. Figure 4. Open in a new tab The substrate-binding ability and binding mechanism analyses of the UKD domain. (a) Gel filtration chromatography analysis of the binding ability of the UKD domain toward the alginate-related substrates. SA, sodium alginate. PMG, heteropolymers consisting of M and G alternately. PM, polymannuronate. PG, polyguluronate. (b) Binding ability of the UKD domain toward alginate gel beads. (c) Binding ability of the UKD domain toward phosphoric acid swollen cellulose (PASC) and Avicel. (d) ITC analysis of the binding ability of UKD domain toward cellohexose. Data are representative of three independent experiments. (e) The overall structure of the UKD domain (Cys884-Cys1010) (PDB ID: 9KVN). (f) The key residues involved in cellulose binding in the UKD domain. (g) The cellulose-binding ability of UKD and its mutants toward bacterial cellulose (BC), Avicel, and PASC. (h) The residues involved in alginate binding in the UKD domain. (i) The binding ability of the mutants of the UKD domain toward alginate gel beads. The graphs in (b), (c), (g), and (i) show data from triplicate experiments (mean ± SD), with bovine serum albumin (BSA) serving as the control. To elucidate the binding mechanisms of the UKD domain to cellulose and alginate, we solved its structure ( Table S2 ), revealing a β-barrel fold comprising nine β-strands and one 3 10 helix (η) ( Fig. 4e ), a folding architecture that has never been reported among the structure-solved CBMs [ 45 , 58 ]. Structural analysis identified a potential planar binding region, including Tyr50 UKD , Tyr63 UKD, and Tyr64 UKD , which are conserved across UKD homologs ( Figs 4f and S6a ). Site-directed mutagenesis followed by binding assays with various insoluble cellulose substrates, including bacterial cellulose (BC), Avicel, and PASC, confirmed the importance of these residues ( Fig. 4g ). Furthermore, the conserved residues across the UKD surface, Arg36 UKD , Lys100 UKD, and Lys110 UKD , may be involved in alginate binding based on mutagenesis analysis ( Figs 4h and i and S6b ). The significance of these binding domains was further illustrated through kinetic assays ( Table 1 ). Compared with recCelAly, the Cel5 and Aly31 variants lacking the three accessory domains showed higher K m values on PASC and alginate, respectively, suggesting that the incorporation of these binding modules improves CelAly’s affinity for BACWPs. Table 1. K m values of recCelAly, Cel5, and Aly31 domains toward different BACWP substrates. Substrate Enzyme K m (mg/ml) PASC recCelAly 50.91 ± 4.6 Cel5 54.68 ± 4.6 Sodium alginate recCelAly 0.31 ± 0.06 Aly31 0.71 ± 0.08 Open in a new tab Noncatalytic accessory domains are prevalent in marine flavobacteria Given the unique binding characteristics of UKD compared to previously reported alginate-binding CBMs [ 54–56 , 59–61 ], we performed comprehensive bioinformatic analysis to determine the relationship of the UKD domain to known CBM families and its distribution across microorganisms. UKD and its homologs formed an independent clade, supporting its classification as the founding member of a new CBM family, CBM113 ( Fig. 5a ). They share a close phylogenetic relationship with families CBM96 and CBM80, which were reported to bind alginate and to exhibit broad specificity for β-glycans, respectively [ 55 , 62 ]. A total of 553 UKD homologs were found. More than three-quarters of UKD-containing proteins are predicted to be CAZymes, with GHs (222/432) being the most abundant group. Based on their CAZymes family classification, most of these GHs are glucanase, especially cellulase. The second most common type is PLs (96/432), the majority of which are putative alginate lyases. These are followed by auxiliary activity (AA) enzymes (55/432) and then modular enzymes combining GH and PL domains (51/432). The AA enzymes are exclusively from the AA10 family and are predicted to be lytic cellulose monooxygenases, a class of enzymes crucial for initiating cellulose degradation. The modular enzymes combining GH and PL domains may simultaneously act on cellulose and alginate, akin to CelAly. These findings indicate that the UKD domain frequently co-occurs with enzymes targeting cellulose and alginate (including bifunctional modular enzymes), reinforcing its functional association in the degradation of these polysaccharides ( Fig. 5b ). Figure 5. Open in a new tab Bioinformatic analyses of the UKD and B1 domains. (a) Phylogenetic analysis of the UKD domain. The phylogenetic tree was constructed using the MEGA X software via the neighbor-joining method. Bootstrap analysis of 1000 replicates was conducted. (b) Distribution of the UKD domain and its homologous proteins in carbohydrate-active enzymes. AA, auxiliary activity. GH, glycoside hydrolase. PL, polysaccharide lyase. GH + PL, enzymes containing GH and PL modules. (c) Taxonomic distribution of UKD homologs in bacteria. (d) Taxonomic distribution of B1 homologs in bacteria. The homologous proteins of UKD and B1 were searched against the NCBI nr database using a threshold of E-value ≤3e-9 and 1e-25, respectively. The majority of the UKD homologs belong to the Bacteroidota phylum, with a very small number originating from Fibrobacterota ( Fig. 5c ). Within Bacteroidota , UKD homologs are predominantly within the Flavobacteriales order (83.7%), particularly in the genus Aquimarina . The second and third most abundant orders are Cytophagales (11.9%) and Bacteroidales (2.0%), respectively. We also observed that the distribution of B1 homologs is highly similar to that of UKD. B1 homologs were predominantly in Bacteroidota (82.4%), mainly within Flavobacteriales (54.1%) and Cytophagales (23.0%), with minor occurrences in Myxococcota, Planctomycetota , and Bdellovibrionota ( Fig. 5d ). Although the continuous discovery of novel GHs and PLs from Flavobacteria underscores their quantitative advantage in enzymatic repertoires for algal polysaccharide degradation [ 63 ], UKD represents another aspect of adaptation: the evolution of unusual accessory modules that expand polysaccharide recognition. Namely, unlike typical CBMs, which target a single polysaccharide class, UKD exhibits dual-substrate binding. Its frequent co-occurrence with cellulases, alginate lyases, and GH + PL modular enzymes suggest that UKD is a broadly selected functional module for polysaccharide binding. UKD likely enhances enzyme localization on complex algal cell wall matrices, thereby increasing effective substrate concentration and catalytic efficiency in natural environments. In comparison, B1 represents a distinct strategy: rather than inventing a new CBM fold, a ubiquitous Big domain, typically involved in protein stability/adhesion [ 64 , 65 ], has been repurposed to specifically recognize native alginate via a positively charged planar surface. These findings indicate that marine flavobacteria gain competitiveness not only by expanding CAZyme repertoires but also by diversifying accessory modules, through both the emergence of novel fold and functional rewiring of existing scaffold, supporting efficient degradation of complex BACWPs. Diversity and distribution of CelAly-like modular polysaccharide-degrading enzymes Given the potential of modular architectures combining glucan hydrolase and alginate lyase domains to represent a model for enzymes degrading BACWPs, we analysed sequences in the CAZy database from all 13 PL families (PL5, −6, −7, −14, −15, −17, −18, −31, −34, −36, −39, −41, and −44), which are predominantly composed of alginate lyases, and specifically selected those sequences containing additional GH domains. As a result, we systematically identified four distinct enzyme types: GH5 + PL31 (containing both GH5 domain and PL31 domains; including CelAly), PL14 + GH9, PL7 + GH16 + PL6, and PL7 + GH16. To experimentally validate activities beyond that of CelAly (a GH5 + PL31 enzyme), one representative marine-derived sequence from each of the other three types (Enzyme A, B, and C) was selected for enzymatic characterization ( Table S6 ). Enzymes A–C exhibited alginate lyase activity, along with activity against glucan substrates, including β-1,3/1,4-glucan and the cellulose substrate CMC ( Fig. 6a ). β-1,3/1,4-Glucan has recently been identified as a common component of brown algal cell walls, where it is encapsulated within the alginate matrix [ 66 ]. These results confirm their potential as genuine BACWP-degrading enzymes. Figure 6. Open in a new tab Diversity and distribution of putative BACWP-degrading enzymes. (a) Activities of putative BACWP-degrading enzymes with different domain architectures toward different BACWP substrates. The presence (solid box) or absence (hollow box) of detectable activity is shown. CMC, carboxymethyl cellulose. The putative BACWP-degrading enzymes were searched in the CAZy database and annotated using the dbCAN meta server. (b) The abundance of marine-derived BACWP-degrading enzymes across various marine environments. (c) Schematic diagram of the modular architectures of marine-derived BACWP-degrading enzymes. (d) The prevalence of genes related to alginate assimilation in marine bacterial isolates harboring CelAly-like BACWPs-degrading enzymes. aly , alginate lyase gene. oal , oligoalginate lyase gene. kdgF , pectin degradation protein gene, sdr , short-chain dehydrogenase/reductase gene. kdgK , 2-dehydro-3-deoxygluconokinase gene. eda , 2-dehydro-3-deoxyphosphogluconate aldolase gene. Genes encoding PL15 or PL17 enzymes were classified as oal . Expanding our search using CelAly and Enzymes A–C as query sequences against the NCBI nr database (coverage ≥75%, identity ≥55%) yielded 85 marine-derived homologs, with GH5 + PL31 enzymes showing remarkable predominance ( Fig. 6b and Table S6 ). The limited number is potentially due to the metagenomic assembly biases for multimodular enzymes. Further analysis of their origins revealed that these enzymes are from diverse marine environments, including seaweed, seawater, sediment, animal, and beach ( Fig. 6b ). Approximately 31.8% of them were isolated from seaweed-associated habitats, suggesting their specialized adaptation to algal polysaccharide degradation in marine ecosystems ( Fig. 6b ). Detailed domain architecture analysis revealed distinct, yet convergent evolutionary strategies among the BACWP-degrading enzymes ( Fig. 6c ). PL7 + GH16 + PL6 enzymes displayed complex CBM32-PL7-GH16-CBM32-PL6 arrangements, whereas PL7 + GH16 enzymes exhibit simpler CBM32-PL7-GH16 architecture, both utilizing the CBM32 domain for alginate recognition [ 61 ]. PL14 + GH9 enzymes combine PL14 alginate lyase and GH9 glucanase catalytic domains with one or two CBM2 family domains for cellulose-binding [ 58 ]. GH5 + PL31 enzymes showed greater diversity in their auxiliary domains while maintaining a conserved GH5-PL31 catalytic core. The predominant pattern is exemplified by CelAly (31/58), and variants included single Big domains (16/58), Big + CBM6 combinations (8/58), or CBM6 replacements (2/58), with CBM6 known as the cellulose-binding module [ 58 ]. The atypical utilization of Big and UKD domains may confer enhanced substrate accessibility to these enzymes during BACWP degradation, thereby explaining their evolutionary advantage and numerical predominance over other types of BACWP-degrading enzymes. These findings reveal that all four types of BACWP-degrading enzyme share a conserved principle for architecture arrangement: catalytically essential domains are consistently paired with specialized binding modules targeting BACWP components. This convergence suggests that diverse microorganisms may evolve similar enzymatic strategies to efficiently degrade BACWPs, highlighting the evolutionary importance of this degradation strategy. Such substrate-driven convergence is not limited to extracellular depolymerases and was also observed in polysaccharide uptake systems, exemplified by the SusCD complexes of Bacteroidota, where SusC (outer-membrane transporter) and SusD (surface glycan-binding protein) homologs cluster by polysaccharide type (e.g. alginate, laminarin) rather than taxonomy [ 63 ]. This suggests that structurally complex marine algal polysaccharides impose similar selective pressures across multiple functional layers of utilization systems, from extracellular depolymerization to substrate-specific uptake, thereby facilitating efficient processing of algal polysaccharides in marine environments. Considering that all the modular enzymes demonstrate alginate-degrading activity, we also analyzed the abundance of genes related to alginate assimilation in the corresponding strains, including alginate lyase genes ( aly ) , oligoalginate lyase genes ( oal ), and genes involved in monosaccharide assimilation ( Fig. 6d ) [ 43 ]. In our annotation, alginate lyases from the PL15 and PL17 families were classified as oligoalginate lyases (Oals) because most characterized enzymes in these families exhibit Oal activity [ 43 ]. However, Oal activity has also been reported in other PL families. Due to the ambiguity of functional prediction based solely on sequence or family membership, all putative alginate lyase genes from other PL families were conservatively assigned as aly . As a result, the number of genes annotated as oal is likely an underestimate. Among strains carrying GH5 + PL31-type enzymes, nearly all the alginate assimilation-related genes were detected in the majority of isolates. Among strains harbouring other types of modular enzymes, only a subset of these metabolic genes was identified. However, the reaction catalysed by KdgF can also occur spontaneously [ 67 ], and most of these strains still possess multiple alginate lyase genes ( Fig. 6d ). The absence of certain functional genes may be due to substantial sequence divergence from known references, because these strains (primarily Actinomycetes ) are phylogenetically distant from the typical sources of reference genes (mostly Bacteroides ). Collectively, these results suggest that these strains likely have the capacity to assimilate BACWPs, as demonstrated here with alginate, and that CelAly-like modular enzymes in these strains may play an important role in brown algae degradation, akin to the function verified in Aquimarina sp. 433. BACWP degradation model by CelAly-like modular enzymes Due to difficulties in determining the complete structure of recCelAly, we employed SAXS to investigate its overall conformation and domain organization ( Table S3 and Fig. S7 ). The ab initio beads model revealed an elongated molecular shape with approximate dimensions of 163 × 52 × 60 Å ( Fig. S8 ). Rigid body modelling incorporating the crystal structures of B1 and UKD domains along with predicted models of Cel5, Aly31, and B2 domains supported a side-by-side domain arrangement ( Figs S8 and S9a ). The Kratky plot displayed an upward trend at high q values, indicating the presence of partial disorder and flexible linkers between domains ( Fig. S9b ) [ 68 ]. Based on the structural and biochemical results, we propose the putative mechanism for BACWP degradation by CelAly. The process initiates with binding of the B1 and UKD domains to the alginate matrix through distinct binding mechanisms, enhancing substrate affinity and positioning Aly31 for efficient alginate cleavage. As alginate degradation progresses, exposed cellulose microfibrils become accessible for UKD binding, which subsequently positions Cel5 for cellulose hydrolysis. The proximity of B1 and Aly31 mirrors the canonical CBM-catalytic domain arrangement observed in other highly efficient CAZymes [ 69 ]. In contrast, the spatial separation of the UKD and two catalytic domains (Cel5 and Aly31) likely allow dynamic interactions. When UKD binds alginate, it may lead to conformational changes that optimize Aly31’s access to alginate while prepositioning Cel5 near emerging cellulose microfibrils. The flexible interdomain architecture might allow Cel5 to hydrolyze cellulose while Aly31 continues processing adjacent alginate regions. This coordinated action is mediated by UKD, which functions as a “hook” that captures substrates and orchestrates catalytic domain positioning. Extending from CelAly to the various BACWP-degrading enzymes analysed in the earlier section, we propose a generalized mechanistic paradigm for their action on BACWPs ( Fig. 7 ). This mechanism involves three key elements: (i) specialized binding modules for cello-configured polysaccharides and/or alginate substrate recognition, including both monospecific modules (e.g. B1 and CBMs) and dual-binding specific modules (e.g. UKD); (ii) polysaccharide degradation through coordinated action of at least two catalytic domains targeting on different types of substrates; and (iii) dynamic structural coordination mediated by flexible linkers. In this system, the enzymes may first target the protective alginate matrix, thereby exposing the underlying cello-configured polysaccharides, after which both alginate and cello-configured substrates can be degraded in a coordinated manner. The flexible interdomain organization might allow adaptive repositioning of catalytic modules during this degradation process primarily mediated by the binding modules. Figure 7. Open in a new tab The mechanistic paradigm for modular enzymes on BACWPs. The modular enzyme degrades BACWPs through a three-stage process. (1) Alginate recognition and cleavage: Alginate-specific binding modules and modules with dual binding capability recognize and anchor onto the alginate matrix, enabling the alginate lyase domain to initiate depolymerization. (2) Cellulose microfibril exposure: Degradation of the alginate matrix exposes embedded cello-configured polysaccharides., which allows adaptive repositioning of binding and catalytic modules to the changing substrate landscapes. (3) Coordinated dual degradation: Facilitated by the substrate binding from both alginate/cello-specific modules and modules with dual substrate-binding capability, the alginate lyase and cellulase domains degrade both exposed substrates to generate alginate and glucan oligosaccharides. Flexible linkers ensure optimal domain organization throughout the dynamic degradation process. Though some multimodular CAZymes have been reported for the degradation of terrestrial plant polysaccharides, most of the enzymes combine domains that target a single polysaccharide type (e.g. cellulose or hemicellulose) [ 7 , 70 ]. Distinct from these enzymes, CelAly-like enzymes display a unique catalytic domain architecture and distinct auxiliary modules. Through the coordinated action of these functionally diverse catalytic domains and auxiliary modules, the degradation of native BACWPs is achieved, underscoring its more sophisticated enzymatic digestion strategy specialized for algal metabolism. This mechanistic framework provides insights into the bacterial evolution of multifunctional enzymes for algal biomass degradation. Conclusion Brown algal biomass represents one of the largest pools of marine primary production, yet the enzymatic strategies enabling its efficient microbial decomposition remain incompletely understood, particularly given the composite nature of BACWPs. Here, we identify and characterize a previously underappreciated type of multimodular enzyme specialized in the coordinated degradation of BACWPs, exemplified by CelAly-like enzymes from marine Flavobacteriaceae. CelAly integrates a cellulase and an alginate lyase within a single protein, each independently contributing to the depolymerization of cellulose and alginate. Degradation assays using L. japonica confirm its effectiveness against native brown algal tissues. Noncatalytic accessory domains of CelAly are important for efficient degradation of native BACWPs. The B1 domain selectively recognizes alginate gels through a positively charged planar interface, and the UKD domain possessing a novel fold that binds both insoluble cellulose and native alginate. These modules increase substrate affinity and suggest a mechanistic basis for efficient enzyme localization and coordinated catalysis on complex cell wall assemblies. Genomic surveys reveal that CelAly homologs and associated accessory domains are largely restricted to marine Flavobacteriaceae, and that such GH + PL modular architectures are enriched in strains from seaweed-associated habitats. Together, the findings provide insights into how marine bacteria access and degrade structurally complex macroalgal cell walls. Rather than relying solely on collections of single-function CAZymes, CelAly-like enzymes represent an integrated strategy that targets multiple key BACWP components within the same physical matrix. The presence of specialized binding modules further suggests that efficient degradation depends not only on catalytic capacity but also on the ability of extracellular enzymes to attach to heterogeneous polysaccharide assemblies. Such features likely affect the efficiency of algal tissue breakdown and the forms of released carbohydrates available to other microbes. The strong phylogenetic restriction of these enzymes to marine Bacteroidota indicates that these lineages are adapted for macroalgal polysaccharide degradation. By generating accessible substrates, such degraders may facilitate downstream trophic interactions and carbon flow through microbial consortia. Overall, this study links multimodular enzyme architecture to ecological strategy, providing mechanistic insight into how specific bacterial groups contribute to macroalgal biomass processing. By converting particulate matter into soluble carbohydrates, these enzymatic systems may contribute to carbon cycling in marine ecosystems. Supplementary Material supplementary_materials_for_revision-2_wrag112 supplementary_materials_for_revision-2_wrag112.docx (10.5MB, docx) Acknowledgements We would like to thank Jingyao Qu, Zhifeng Li, and Jing Zhu from the State Key Laboratory of Microbial Technology of Shandong University for help and guidance in ITC and DLS. We thank the staffs from BL02U1&BL19U2 beamlines of National Facility for Protein Sciences Shanghai (NFPS) and Shanghai Synchrotron Radiation Facility, for assistance during data collection. Contributor Information Fei Xu, State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China. Xiao-Hui Sun, State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China. Xiao-Dong Zhang, State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China. Xiao-Fei Wang, State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China. Yan Wang, State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China. Hai-Yan Cao, MOE Key Laboratory of Evolution and Marine Biodiversity, Frontiers Science Center for Deep Ocean Multispheres and Earth System & College of Marine Life Sciences, Ocean University of China, Qingdao 266203, Shandong Province, China. Peng Wang, MOE Key Laboratory of Evolution and Marine Biodiversity, Frontiers Science Center for Deep Ocean Multispheres and Earth System & College of Marine Life Sciences, Ocean University of China, Qingdao 266203, Shandong Province, China; Laboratory for Marine Biology and Biotechnology, Qingdao Marine Science and Technology Center & Laoshan Laboratory, Qingdao 266237, Shandong Province, China. Jian-Xun Li, State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China. Xi-Ying Zhang, State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China. Qi-Long Qin, State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China. Xiu-Lan Chen, State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China. Yu-Zhong Zhang, MOE Key Laboratory of Evolution and Marine Biodiversity, Frontiers Science Center for Deep Ocean Multispheres and Earth System & College of Marine Life Sciences, Ocean University of China, Qingdao 266203, Shandong Province, China; Laboratory for Marine Biology and Biotechnology, Qingdao Marine Science and Technology Center & Laoshan Laboratory, Qingdao 266237, Shandong Province, China; Marine Biotechnology Research Center, State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China. Yin Chen, MOE Key Laboratory of Evolution and Marine Biodiversity, Frontiers Science Center for Deep Ocean Multispheres and Earth System & College of Marine Life Sciences, Ocean University of China, Qingdao 266203, Shandong Province, China; School of Life Sciences, University of Warwick, Coventry CV4 7AL, West Midlands, United Kingdom; School of Biosciences, University of Birmingham, Edgbaston B15 2TT, Birmingham, United Kingdom. Yu-Qiang Zhang, State Key Laboratory of Microbial Technology, Shandong University, Qingdao 266237, Shandong Province, China. Author contributions Y.-Z.Z., X.-L.C., and F.X. conceived the project. F.X., X.-L.C., and X.-H.S. designed and performed most of the experiments and data interpretation. X.-D.Z. and X.-F.W. performed the experiments and helped in data analysis. F.X., X.-H.S, Y.-Z.Z., and X.-L.C. analyzed the data and wrote the paper. Y.W. helped in SAXS analysis. H.-Y.C. P.W. and J.-X.L. helped in structure determination. Q.-L.Q., Y.-Q.Z., and X.-Y.Z. helped to analyze the data. F.X., X.-L.C., Y.-Z.Z., Y.-Q.Z., and Y.C. revised the paper. All authors discussed and commented on the results and the paper. Conflicts of interest None declared. Funding This work was supported by the National Key R&D Program of China (2024YFC2816005), the National Natural Science Foundation of China (3227004, 42206156, 32401351, W2441012, 32370136, and 32330001), Shandong Provincial Natural Science Foundation (ZR2024QC025, ZR2021QC102), Qingdao Natural Science Foundation (24-4-4-zrjj-25-jch), the Marine S&T Fund of Shandong Province for Qingdao Marine Science and Technology Center (grant 2022QNLM030004-3), the Taishan Scholars Program of Shandong Province, China (tspd20240806, tsqn202408064), Shandong Province Science Fund for Excellent Young Scholars (ZR2025QB18), and the SKLMT Frontiers and Challenges Project (grant SKLMTFCP-2023-06). Data availability The genomic data of Aquimarina sp. 433 and Aquimarina sp. 2-A2 are available in NCBI under the accession numbers JBSWZK000000000.1 and JBJKFI000000000.1 , respectively. The structures of B1 and UKD have been deposited in PDB under identifiers 9KVU and 9KVN, respectively. References 1. Shtein  I, Bar-On  B, Popper  ZA. Plant and algal structure: from cell walls to biomechanical function. Physiol Plant 2018;164:56–66. 10.1111/ppl.12727 [ DOI ] [ PubMed ] [ Google Scholar ] 2. Khamassi  A, Dumon  C. Enzyme synergy for plant cell wall polysaccharide degradation. Essays Biochem 2023;67:521–31. 10.1042/EBC20220166 [ DOI ] [ PubMed ] [ Google Scholar ] 3. Sarkar  P, Bandyopadhyay  TK, Gopikrishna  K  et al.  Algal carbohydrates: sources, biosynthetic pathway, production, and applications. Bioresour Technol 2024;413:131489. 10.1016/j.biortech.2024.131489 [ DOI ] [ PubMed ] [ Google Scholar ] 4. Le  Y, Zhang  M, Wu  P  et al.  Biofuel production from lignocellulose via thermophile-based consolidated bioprocessing. Eng Microbiol 2024;4:100174. 10.1016/j.engmic.2024.100174 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 5. López-Mondéjar  R, Zühlke  D, Becher  D  et al.  Cellulose and hemicellulose decomposition by forest soil bacteria proceeds by the action of structurally variable enzymatic systems. Sci Rep 2016;6:25279. 10.1038/srep25279 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 6. Arnosti  C, Wietz  M, Brinkhoff  T  et al.  The biogeochemistry of marine polysaccharides: sources, inventories, and bacterial drivers of the carbohydrate cycle. Annu Rev Mar Sci 2021;13:81–108. 10.1146/annurev-marine-032020-012810 [ DOI ] [ PubMed ] [ Google Scholar ] 7. Brunecky  R, Alahuhta  M, Xu  Q  et al.  Revealing nature’s cellulase diversity: the digestion mechanism of Caldicellulosiruptor bescii CelA. Science 2013;342:1513–6. 10.1126/science.1244273 [ DOI ] [ PubMed ] [ Google Scholar ] 8. Chu  Y, Hao  Z, Wang  K  et al.  The GH10 and GH48 dual-functional catalytic domains from a multimodular glycoside hydrolase synergize in hydrolyzing both cellulose and xylan. Biotechnol Biofuels 2019;12:279. 10.1186/s13068-019-1617-2 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 9. Bligh  M, Nguyen  N, Buck-Wiese  H  et al.  Structures and functions of algal glycans shape their capacity to sequester carbon in the ocean. Curr Opin Chem Biol 2022;71:102204. 10.1016/j.cbpa.2022.102204 [ DOI ] [ PubMed ] [ Google Scholar ] 10. Smith  SV. Marine macrophytes as a global carbon sink. Science 1981;211:838–40. 10.1126/science.211.4484.838 [ DOI ] [ PubMed ] [ Google Scholar ] 11. Fernández-Gómez  B, Richter  M, Schüler  M  et al.  Ecology of marine Bacteroidetes: a comparative genomics approach. ISME J 2013;7:1026–37. 10.1038/ismej.2012.169 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 12. Brunet  M, Le Duff  N, Barbeyron  T  et al.  Consuming fresh macroalgae induces specific catabolic pathways, stress reactions and type IX secretion in marine flavobacterial pioneer degraders. ISME J 2022;16:2027–39. 10.1038/s41396-022-01251-6 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 13. Bauer  M, Kube  M, Teeling  H  et al.  Whole genome analysis of the marine Bacteroidetes ‘ Gramella forsetii ’ reveals adaptations to degradation of polymeric organic matter. Environ Microbiol 2006;8:2201–13. 10.1111/j.1462-2920.2006.01152.x [ DOI ] [ PubMed ] [ Google Scholar ] 14. Thomas  F, Le Duff  N, Wu  T  et al.  Isotopic tracing reveals single-cell assimilation of a macroalgal polysaccharide by a few marine Flavobacteria and Gammaproteobacteria. ISME J 2021;15:3062–75. 10.1038/s41396-021-00987-x [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 15. Li  H, Zhang  Z, Chen  J  et al.  Fate and carbon sequestration potential of sunken macroalgae in coastal oceans from long-term microbial degradation perspective. Natl Sci Rev 2025;12:nwaf273. 10.1093/nsr/nwaf273 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 16. Li  Q, Zheng  L, Guo  Z  et al.  Alginate degrading enzymes: an updated comprehensive review of the structure, catalytic mechanism, modification method and applications of alginate lyases. Crit Rev Biotechnol 2021;41:953–68. 10.1080/07388551.2021.1898330 [ DOI ] [ PubMed ] [ Google Scholar ] 17. Chen  C, Li  X, Lu  C  et al.  Advances in alginate lyases and the potential application of enzymatic prepared alginate oligosaccharides: a mini review. Int J Biol Macromol 2024;260:129506. 10.1016/j.ijbiomac.2024.129506 [ DOI ] [ PubMed ] [ Google Scholar ] 18. Kusaykin  MI, Silchenko  AS, Zakharenko  AM  et al.  Fucoidanases. Glycobiology 2016;26:3–12. 10.1093/glycob/cwv072 [ DOI ] [ PubMed ] [ Google Scholar ] 19. Zueva  AO, Silchenko  AS, Rasin  AB  et al.  Expression and biochemical characterization of two recombinant fucoidanases from the marine bacterium Wenyingzhuangia fucanilytica CZ1127T. Int J Biol Macromol 2020;164:3025–37. 10.1016/j.ijbiomac.2020.08.131 [ DOI ] [ PubMed ] [ Google Scholar ] 20. Beuder  S, Braybrook  SA. Brown algal cell walls and development. Semin Cell Dev Biol 2023;134:103–11. 10.1016/j.semcdb.2022.03.003 [ DOI ] [ PubMed ] [ Google Scholar ] 21. Sichert  A, Corzett  CH, Schechter  MS  et al.  Verrucomicrobia use hundreds of enzymes to digest the algal polysaccharide fucoidan. Nat Microbiol 2020;5:1026–39. 10.1038/s41564-020-0720-2 [ DOI ] [ PubMed ] [ Google Scholar ] 22. Zhang  YH, Cui  J, Lynd  LR  et al.  A transition from cellulose swelling to cellulose dissolution by o-phosphoric acid: evidence from enzymatic hydrolysis and supramolecular structure. Biomacromolecules 2006;7:644–8. 10.1021/bm050799c [ DOI ] [ PubMed ] [ Google Scholar ] 23. Haug  A, Larsen  B, Smidsrød  O  et al.  Studies on the sequence of uronic acid residues in alginic acid. Acta Chem Scand 1967;21:691–704. 10.3891/acta.chem.scand.21-0691 [ DOI ] [ Google Scholar ] 24. Miller  GL. Use of dinitrosalicylic acid reagent for determination of reducing sugar. Anal Chem 1959;31:426–8. 10.1021/ac60147a030 [ DOI ] [ Google Scholar ] 25. Xu  F, Chen  XL, Sun  XH  et al.  Structural and molecular basis for the substrate positioning mechanism of a new PL7 subfamily alginate lyase from the arctic. J Biol Chem 2020;295:16380–92. 10.1074/jbc.RA120.015106 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 26. Adams  PD, Grosse-Kunstleve  RW, Hung  LW  et al. PHENIX : building new software for automated crystallographic structure determination. Acta Crystallogr D Biol Crystallogr 2002;58:1948–54. 10.1107/S0907444902016657 [ DOI ] [ PubMed ] [ Google Scholar ] 27. Emsley  P, Cowtan  K. Coot : model-building tools for molecular graphics. Acta Crystallogr D Biol Crystallogr 2004;60:2126–32. 10.1107/S0907444904019158 [ DOI ] [ PubMed ] [ Google Scholar ] 28. Hopkins  JB, Gillilan  RE, Skou  S. BioXTAS RAW : improvements to a free open-source program for small-angle X-ray scattering data reduction and analysis. J Appl Crystallogr 2017;50:1545–53. 10.1107/S1600576717011438 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 29. Manalastas-Cantos  K, Konarev  PV, Hajizadeh  NR  et al. ATSAS 3.0 : expanded functionality and new tools for small-angle scattering data analysis. J Appl Crystallogr 2021;54:343–55. 10.1107/S1600576720013412 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 30. Svergun  DI. Determination of the regularization parameter in indirect-transform methods using perceptual criteria. J Appl Crystallogr 1992;25:495–503. 10.1107/S0021889892001663 [ DOI ] [ Google Scholar ] 31. Kikhney  AG, Svergun  DI. A practical guide to small angle X-ray scattering (SAXS) of flexible and intrinsically disordered proteins. FEBS Lett 2015;589:2570–7. 10.1016/j.febslet.2015.08.027 [ DOI ] [ PubMed ] [ Google Scholar ] 32. Franke  D, Svergun  DI. DAMMIF , a program for rapid ab-initio shape determination in small-angle scattering. J Appl Crystallogr 2009;42:342–6. 10.1107/S0021889809000338 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 33. Svergun  DI. Restoring low resolution structure of biological macromolecules from solution scattering using simulated annealing. Biophys J 1999;76:2879–86. 10.1016/S0006-3495(99)77443-6 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 34. Petoukhov  MV, Franke  D, Shkumatov  AV  et al.  New developments in the ATSAS program package for small-angle scattering data analysis. J Appl Crystallogr 2012;45:342–50. 10.1107/S0021889812007662 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 35. Svergun  D, Barberato  C, Koch  MHJ. CRYSOL - a program to evaluate X-ray solution scattering of biological macromolecules from atomic coordinates. J Appl Crystallogr 1995;28:768–73. 10.1107/S0021889895007047 [ DOI ] [ Google Scholar ] 36. Kozin  M, Svergun  D. Automated matching of high- and low- resolution structural models. J Appl Crystallogr 2001;34:33–41. 10.1107/S0021889800014126 [ DOI ] [ Google Scholar ] 37. Nielsen  H. Predicting secretory proteins with SignalP. Methods Mol Biol 2017;1611:59–73. 10.1007/978-1-4939-7015-5_6 [ DOI ] [ PubMed ] [ Google Scholar ] 38. Marchler-Bauer  A, Bo  Y, Han  L  et al.  CDD/SPARCLE: functional classification of proteins via subfamily domain architectures. Nucleic Acids Res 2017;45:D200–3. 10.1093/nar/gkw1129 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 39. Zhang  H, Yohe  T, Huang  L  et al.  dbCAN2: a meta server for automated carbohydrate-active enzyme annotation. Nucleic Acids Res 2018;46:W95–101. 10.1093/nar/gky418 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 40. Dong  F, Xu  F, Chen  X  et al.  Alginate lyase Aly36B is a new bacterial member of the polysaccharide lyase family 36 and catalyzes by a novel mechanism with lysine as both the catalytic base and catalytic acid. J Mol Biol 2019;431:4897–909. 10.1016/j.jmb.2019.10.023 [ DOI ] [ PubMed ] [ Google Scholar ] 41. Kumar  S, Stecher  G, Li  M  et al.  MEGA X: molecular evolutionary genetics analysis across computing platforms. Mol Biol Evol 2018;35:1547–9. 10.1093/molbev/msy096 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 42. Jumper  J, Evans  R, Pritzel  A  et al.  Highly accurate protein structure prediction with AlphaFold. Nature 2021;596:583–9. 10.1038/s41586-021-03819-2 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 43. Thomas  F, Barbeyron  T, Tonon  T  et al.  Characterization of the first alginolytic operons in a marine bacterium: from their emergence in marine Flavobacteriia to their independent transfers to marine Proteobacteria and human gut Bacteroides . Environ Microbiol 2012;14:2379–94. 10.1111/j.1462-2920.2012.02751.x [ DOI ] [ PubMed ] [ Google Scholar ] 44. Dudek  M, Dieudonné  A, Jouanneau  D  et al.  Regulation of alginate catabolism involves a GntR family repressor in the marine flavobacterium Zobellia galactanivorans DsijT. Nucleic Acids Res 2020;48:7786–800. 10.1093/nar/gkaa533 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 45. Drula  E, Garron  ML, Dogan  S  et al.  The carbohydrate-active enzyme database: functions and literature. Nucleic Acids Res 2022;50:D571–7. 10.1093/nar/gkab1045 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 46. Pontrelli  S, Szabo  R, Pollak  S  et al.  Metabolic cross-feeding structures the assembly of polysaccharide degrading communities. Sci Adv 2022;8:eabk3076. 10.1126/sciadv.abk3076 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 47. Paillat  M, Lunar Silva  I, Cascales  E  et al.  A journey with type IX secretion system effectors: selection, transport, processing and activities. Microbiology 2023;169:1320. 10.1099/mic.0.001320 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 48. Gong  Y, Shang  DD, Sun  CL  et al.  Direct degradation of fresh and dried macroalgae by Agarivorans albus B2Z047. Mar Drugs 2024;22:203. 10.3390/md22050203 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 49. Lisov  AV, Kiselev  SS, Trubitsina  LI  et al.  Multifunctional enzyme with endoglucanase and alginase/glucuronan lyase activities from bacterium Cellulophaga lytica . Biochemistry 2022;87:617–27. 10.1134/S0006297922070045 [ DOI ] [ PubMed ] [ Google Scholar ] 50. Zhu  B, Wang  H, Zhao  Y  et al.  Elucidation of domain function of a novel multifunctional glycoside hydrolase and its use in efficient preparation of oligosaccharides from kelp powder. J Agric Food Chem 2024;72:28028–39. 10.1021/acs.jafc.4c07333 [ DOI ] [ PubMed ] [ Google Scholar ] 51. Zhang  Y, Zhang  Y, Zhao  X  et al.  Metagenomic insights into the dynamic degradation of brown algal polysaccharides by kelp-associated microbiota. Appl Environ Microbiol 2024;90:e0202523. 10.1128/aem.02025-23 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 52. Hudson  J, Kumar  V, Egan  S. Comparative genome analysis provides novel insight into the interaction of Aquimarina sp. AD1, BL5 and AD10 with their macroalgal host. Mar . Genomics 2019;46:8–15. 10.1016/j.margen.2019.02.005 [ DOI ] [ PubMed ] [ Google Scholar ] 53. Gao  L, Guan  Z, Gao  P  et al. Cytophaga hutchinsonii gldN , encoding a core component of the type IX secretion system, is essential for ion assimilation, cellulose degradation, and cell motility. Appl Environ Microbiol 2020;86:e00220–42. 10.1128/AEM.00242-20 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 54. Teh  A, Sim  P, Hisano  T. Structural basis for binding uronic acids by family 32 carbohydrate-binding modules. Biochem Biophys Res Commun 2020;533:257–61. 10.1016/j.bbrc.2020.09.064 [ DOI ] [ PubMed ] [ Google Scholar ] 55. Ji  S, Tian  X, Li  X  et al.  Identification and structural analysis of a carbohydrate-binding module specific to alginate, a representative of a new family, CBM96. J Biol Chem 2023;299:102854. 10.1016/j.jbc.2022.102854 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 56. Mei  X, Tao  W, Sun  H  et al.  Characterization and structural identification of a novel alginate-specific carbohydrate-binding module (CBM): the founding member of a new CBM family. Int J Biol Macromol 2024;277:134221. 10.1016/j.ijbiomac.2024.134221 [ DOI ] [ PubMed ] [ Google Scholar ] 57. Liu  Y, Wang  P, Tian  J  et al.  Carbohydrate-binding modules of potential resources: occurrence in nature, function, and application in fiber recognition and treatment. Polymers 2022;14:1806. 10.3390/polym14091806 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 58. Boraston  AB, Bolam  DN, Gilbert  HJ  et al.  Carbohydrate-binding modules: fine-tuning polysaccharide recognition. Biochem J 2004;382:769–81. 10.1042/BJ20040892 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 59. Li  S, Yang  X, Bao  M  et al.  Family 13 carbohydrate-binding module of alginate lyase from Agarivorans sp. L11 enhances its catalytic efficiency and thermostability, and alters its substrate preference and product distribution. FEMS Microbiol Lett 2015;362:fnv054. 10.1093/femsle/fnv054 [ DOI ] [ PubMed ] [ Google Scholar ] 60. Mei  X, Chang  Y, Shen  J  et al.  Expression and characterization of a novel alginate-binding protein: a promising tool for investigating alginate. Carbohydr Polym 2020;246:116645. 10.1016/j.carbpol.2020.116645 [ DOI ] [ PubMed ] [ Google Scholar ] 61. Tang  L, Guo  E, Zhang  L  et al.  The function of CBM32 in alginate lyase VxAly7B on the activity on both soluble sodium alginate and alginate gel. Front Microbiol 2021;12:798819. 10.3389/fmicb.2021.798819 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 62. Venditto  I, Luis  AS, Rydahl  M  et al.  Complexity of the Ruminococcus flavefaciens cellulosome reflects an expansion in glycan recognition. Proc Natl Acad Sci USA 2016;113:7136–41. 10.1073/pnas.1601558113 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 63. Kappelmann  L, Krüger  K, Hehemann  J  et al.  Polysaccharide utilization loci of North Sea Flavobacteriia as basis for using SusC/D-protein expression for predicting major phytoplankton glycans. ISME J 2019;13:76–91. 10.1038/s41396-018-0242-6 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 64. Brunecky  R, Alahuhta  M, Bomble  YJ  et al.  Structure and function of the Clostridium thermocellum cellobiohydrolase A X1-module repeat: enhancement through stabilization of the CbhA complex. Acta Crystallogr D Biol Crystallogr 2012;68:292–9. 10.1107/S0907444912001680 [ DOI ] [ PubMed ] [ Google Scholar ] 65. Xing  M, Wang  Y, Zhao  Y  et al.  C-terminal bacterial immunoglobulin-like domain of κ-carrageenase serves as a multifunctional module to promote κ-carrageenan hydrolysis. J Agric Food Chem 2022;70:1212–22. 10.1021/acs.jafc.1c07233 [ DOI ] [ PubMed ] [ Google Scholar ] 66. Salmeán DDJH  AA. Insoluble (1 → 3), (1 → 4)-β-D-glucan is a component of cell walls in brown algae (Phaeophyceae) and is masked by alginates in tissues. Sci Rep 2017;7:2880. 10.1038/s41598-017-03081-5 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 67. Hobbs  JK, Lee  SM, Robb  M  et al.  KdgF, the missing link in the microbial metabolism of uronate sugars from pectin and alginate. Proc Natl Acad Sci USA 2016;113:6188–93. 10.1073/pnas.1524214113 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 68. Holcomb  J, Doughan  M, Spellmon  N  et al.  SAXS analysis of a soluble cytosolic NgBR construct including extracellular and transmembrane domains. PLoS One 2018;13:e0191371. 10.1371/journal.pone.0191371 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 69. Zuhlke  MK, Ficko-Blean  E, Bartosik  D  et al.  Unveiling the role of novel carbohydrate-binding modules in laminarin interaction of multimodular proteins from marine Bacteroidota during phytoplankton blooms. Environ Microbiol 2024;26:e16624. 10.1111/1462-2920.16624 [ DOI ] [ PubMed ] [ Google Scholar ] 70. Su  X, Mackie  RI, Cann  IK. Biochemical and mutational analyses of a multidomain cellulase/mannanase from Caldicellulosiruptor bescii . Appl Environ Microbiol 2012;78:2230–40. 10.1128/AEM.06814-11 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Associated Data This section collects any data citations, data availability statements, or supplementary materials included in this article. Supplementary Materials supplementary_materials_for_revision-2_wrag112 supplementary_materials_for_revision-2_wrag112.docx (10.5MB, docx) Data Availability Statement The genomic data of Aquimarina sp. 433 and Aquimarina sp. 2-A2 are available in NCBI under the accession numbers JBSWZK000000000.1 and JBJKFI000000000.1 , respectively. The structures of B1 and UKD have been deposited in PDB under identifiers 9KVU and 9KVN, respectively. Articles from The ISME Journal are provided here courtesy of Oxford University Press ACTIONS View on publisher site PDF (2.9 MB) Cite Collections Permalink RESOURCES Similar articles Cited by other articles Links to NCBI Databases Follow NCBI NCBI on X (formerly known as Twitter) NCBI on Facebook NCBI on LinkedIn NCBI on GitHub NCBI RSS feed Connect with NLM NLM on X (formerly known as Twitter) NLM on Facebook NLM on YouTube National Library of Medicine 8600 Rockville Pike Bethesda, MD 20894 Web Policies FOIA HHS Vulnerability Disclosure Help Accessibility Careers NLM NIH HHS USA.gov Back to Top

Related documents

• Caracterización microscópica y molecular de eventos de floraciones algales en lagos de la región del Biobío.#1148404
• Oxygen-responsive KoBdcA/KoBdcB/KoBpdA system regulates c-di-GMP levels to control bacterial cellulose biosynthesis and motility in Kosakonia oryzendophytica FY-07.#157043
• Enzymes, Proteins, and Peptides as Promising Biosolutions for Crop Production#14425
• Synthetic communities of Yarrowia lipolytica for efficient betanin bioproduction through metabolic partitioning#1121673
• Tailoring the stress response of Komagataella phaffii for recombinant protein production on sustainable carbon sources#355171