Cold-adapted carboxylesterases from Alcanivoracaceae active with a wide range of synthetic polyesters

Cold-adapted carboxylesterases from Alcanivoracaceae active with a wide range of synthetic polyesters | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Research Article Cold-adapted carboxylesterases from Alcanivoracaceae active with a wide range of synthetic polyesters Hairong Ma, Anna N. Khusnutdinova, Tatyana N. Chernikova, Manuel Ferrer, and 3 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-7917997/v1 This work is licensed under a CC BY 4.0 License Status: Published Journal Publication published 07 Feb, 2026 Read the published version in Applied Microbiology and Biotechnology → Version 1 posted You are reading this latest preprint version Abstract Members of the family Alcanivoracaceae are widespread in marine environments, where they play central roles in hydrocarbon degradation and populate plastics-associated microbiomes, with notable enzymatic potential toward ester- and olefin-based polymers. To further investigate their enzymatic potential, we selected 21 candidate enzymes from the α/β-fold hydrolase superfamily, specifically carboxylesterase Family V from genome-sequenced representatives of the genera Alcanivorax, Alloalcanivorax , and Isoalcanivorax . Seventeen enzymes were cloned and heterologously expressed in E. coli , of which eleven were purified and subjected to substrate specificity analyses alongside six previously reported and partially characterised carboxylesterases from A. borkumensis SK2, used as benchmarks. All enzymes showed activity against soluble model p- nitrophenyl ester substrates with acyl chain lengths ranging from C2 to C12 and against bis(benzoyloxyethyl) terephthalate (3PET) and polycaprolactone 2 kDa (PCL2). During 3PET hydrolysis, product accumulation followed the order: benzoic acid > > MHET > terephthalic acid. Five enzymes hydrolysed polycaprolactone 14 kDa (PCL14), poly-D,L-lactide (PDLLA), and polybutylene adipate (PBA). All five enzymes displayed temperature optima around or below 50°C and retained high activity at low temperatures (5–20°C), consistently with adaptation to marine environments. Enzymes also exhibited moderate solvent tolerance, neutral-to-alkaline pH optima, and low thermostability, with melting temperatures (Tm) between 31°C and 48°C. Overall, enzymes from Alcanivoracaceae exhibited promising potential for synthetic polyesters biodegradation, especially under low-temperature conditions, suggesting potential application for degrading specific polyester-based plastics with lower molecular weight, and their utility in further enzyme engineering for plastic recycling and upcycling. Carboxylestereases polyesters cold-adapted enzymes marine hydrocarbon-degrading bacteria Alcanivoracaceae Alcanivorax Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Figure 7 Figure 8 Figure 9 Figure 10 Key points Members of Alcanivoracaceae are a rich resource of polyester-degrading enzymes All selected and analysed Family V esterases exhibited high activities and stabilities at low temperatures and solvent tolerance Characterised enzymes were active with a broad range of polyesters Introduction Plastic pollution is an ever-escalating issue with detrimental effects on both the environment and human health. Over the years, plastics have become an indispensable part of daily life, serving critical roles in packaging, transportation and healthcare. However, their widespread use has also led to a significant waste accumulation problem, making plastic pollution one of the greatest environmental challenges of the 21st century. Despite growing consumer awareness, the production of single-use plastics continues to escalate (Geyer et al., 2017 ; Stubbins et al., 2021 ). Among various strategies for addressing plastic waste, biological degradation, particularly enzymatic closed-loop recycling has emerged as a promising solution (Abou-Zeid et al., 2004 ; Danso et al., 2019 ; Viljakainen and Hug, 2021). For example, the first enzymatic polyethylene terephthalate (PET) degradation study was reported with an enzyme derived from Thermobifida fusca (Müller et al., 2005 ). Over the past two decades, significant advancements in protein and process engineering have led to substantial improvements in the initially low PET degradation activity of T. fusca (Barth et al., 2015 ; Müller et al., 2005 ; Roth et al., 2014 ). With the advancement of metagenomic approaches, more active enzymes, such as leaf-branch-compost cutinase LCC, have been identified (Sulaiman et al., 2012 ). Furthermore, protein engineering has enabled the development of mutant LCC variants (e.g., LCC ICCG , LCC WCCG ) with enhanced and accelerated depolymerisation efficiency (Makryniotis et al., 2023 ; Tournier et al., 2020 ). Subsequently, a novel bacterium, Ideonella sakaiensis 201-F6, was isolated from plastic-contaminated sediment samples and found to exhibit both IsPETase and monomer-hydrolysing (MHETase) activities (Kan et al., 2021 ; Yoshida et al., 2016 ). IsPETase initially identified and characterised in that study, has served as the foundation for numerous structural and protein engineering studies aimed at further enhancing its PET depolymerisation activity (Dai et al., 2021 ; Feng et al., 2021 ; Joo et al., 2018 ; Haugwitz et al., 2022 ; Wei et al., 2022 ). Ongoing research efforts are focused on modifying the active sites of the wild-type enzymes to improve their catalytic efficiency, as well as engineering enzymes with enhanced thermostabilities trough rational design and directed evolution (Bell et al., 2022 ; Shi et al., 2023 ; Schmidt et al., 2019 ; Williams et al., 2023 ). These advancements hold great potential for the development of efficient biocatalysts capable of supporting sustainable plastic waste management and recycling. The ester bonds in polyesters like PET are chemically different from the carbon-carbon bonds in polymers like polyethylene (PE) and polystyrene (PS) (Mohanan et al., 2020 ). The latter olefin-based polymers require oxygenation, hydroxylation, or monooxygenation of C-C and C-H bonds to enable their degradation in same pathways as for aliphatic hydrocarbons degradations (Bornscheuer et al., 2024 ). In contrast, hydrocarbon contamination from petroleum oil production, processing and transportation significantly contributes to environmental pollution. Like plastic waste, hydrocarbons persist in ecosystems, exacerbating the long-term environmental burden due to their resistance to biodegradation and accumulation in both terrestrial and marine habitats (Brzeszcz and Kaszycki, 2018 ; Park and Park, 2018 ). The degradation of these compounds using bio-based approaches has gained considerable attention, particularly the application of naturally occurring hydrocarbon-degrading marine microorganisms and their enzymes (Beilen et al., 1994 ; Beilen and Funhoff, 2005 , 2007 ; Haines and Alexander, 1974 ; Wei and Zimmermann, 2017 ). Hydrocarbon degrading microorganisms, which are taxonomically diverse, capable to utilise hydrocarbons as their sole carbon and energy source. These include truly marine bacteria, such as Alcanivorax, Oleiphilus, Oleispira, Thalassolituus, Cycloclasticus, Marinobacter and others (Golyshin et al., 2003 ; Gregson et al., 2018 , 2020 ; Schneiker et al., 2006 ; Yakimov et al., 2004 ;), as well as fungi Aspergillus and Penicillium (Zhou et al., 2014 ; Sowmya et al., 2015 ) that harbour enzymes for full mineralisation of alkanes. Key enzymes involved in alkane degradation include alkane hydroxylases and monooxygenases (e.g. AlmA, AlkB, P450), which oxidise alkanes to alcohols, followed by their conversion to aldehydes, fatty acids, acyl-CoA and their further beta-oxidation (Guo et al., 2023 ; Kube et al., 2013 ; Sabirova et al., 2006 ; Wang and Shao, 2014). These metabolic pathways are essential for the complete mineralisation of hydrocarbons and play a key role in the natural bioremediation of oil spills (De Santi et al., 2014 ; Lai et al., 2011 ). Interestingly, many hydrocarbon-degrading microorganisms identified in the marine oil-contaminated environments are also found in the “plastisphere”, the surfaces of plastics and microplastics providing a biological niche where diverse microbial communities establish biofilms (Emadian et al., 2017 ; Wei and Zimmermann, 2017 ; Zettler et al., 2013 ). Both oil spill and plastisphere-associated microbiomes often possess similar metabolic capabilities, such as alkane hydroxylases and esterases, which enable them to degrade complex hydrophobic compounds (Emadian et al., 2017 ; Wei and Zimmermann, 2017 ). This functional overlap suggests that, despite originating from different sources, these environments create comparable ecological niches that select for organisms with similar biodegradation potential (Wilkes and Aristilde, 2017 ; Sabirova et al., 2006 ). For instance, Pseudomonas species are known for their metabolic versatility, producing cutinases and other esterases that facilitate the degradation of synthetic polyesters like PET (Wilkes and Aristilde, 2017 ; Zhou et al., 2024 ). Similarly, Rhodococcus species produce esterases with broad substrate specificity, allowing them to degrade various polyester-based plastics (De Santi et al., 2014 ). These enzymes exhibit significant potential for bioremediation and recycling of plastic waste, offering an environmentally friendly alternative to conventional chemical processes. Polyesterases, such as carboxylesterases, cutinases, lipases, and specific PETases, are particularly effective in degrading synthetic polyesters. Cutinases, found in both bacteria and fungi, hydrolyse the ester bonds in cutin and synthetic polyesters, leading to the formation of monomers such as terephthalic acid and ethylene glycol. PETases, which have gained considerable attention, are specialized enzymes capable of breaking down PET into its constituent monomers under mild conditions (Tournier et al., 2020 ). These enzymes, originally identified in Ideonella sakaiensis , have been found in various alkane-degrading microorganisms, highlighting the potential for dual functionality in hydrocarbon and polyester degradation (Yoshida et al., 2016 ). Some bacteria from the genera Alcanivorax, Oleiphilus, Oleispira, Marinobacter and Thalassolituus , utilise hydrocarbons as their preferred source of carbon and energy (Staley, 2010 ; Yakimov et al., 2004 , 2007 ; Golyshin et al., 2002 ). Among those organisms, which hold significant potential for both natural attenuation of oil hydrocarbons in situ , and degradation of olefins and polyesters, the genus Alcanivorax , a member of the family Alcanivoracaceae within the class Gammaproteobacteria , has attracted particular attention for over 25 years (Golyshin et al., 2003 ; Yakimov et al., 1998 , 2022 ). The genus Alcanivorax (family Alcanivoracaceae ) was recently divided into three genera, including two new (Rai et al., 2023). Several species within this family exhibit a strong preference for aliphatic hydrocarbons, both linear and branched (Yakimov et al., 2022 ). Some Alcanivoracaceae species have also been reported to degrade polycyclic aromatic hydrocarbons (PAHs) such as naphthalene and pyrene, e.g. Alloalcanivorax xenomutans SRM1 (previously known as Alcanivorax xenomutans ) (Dell’Anno et al., 2023 ) and aromatic hydrocarbons, e.g. xylene ( A. xenomutans JC109) (Rahul et al., 2014 ). Typically, Alcanivoracaceae species occur at low or undetectable levels in unpolluted environments; however, their growth is markedly stimulated by the presence of hydrocarbons (Yakimov et al., 2007 ; 2022 ). Many characterised strains do not assimilate sugars or amino acids, but are capable of metabolising fatty acids, alcohols, and aliphatic hydrocarbons (Lai et al., 2011 ). Given their ecological lifestyle, these organisms, and their enzymes, are expected to exhibit tolerance to organic solvents (Bollinger et al., 2020 ). Alcanivoracaceae species along with other hydrocarbon-degrading taxa, are frequently identified as colonisers of plastic surface in marine environment and are considered as potential plastic degraders (Denaro et al., 2020 ; Delacuvellerie et al., 2019 ; Popovic et al., 2017 ; Tulloch et al., 2024 ; Yakimov et al., 2019 ; Zadjelovic et al., 2022 ;). In terms of enzymatic capability, carboxylesterases from the type strain of the family Alcanivorax borkumensis SK2 T have demonstrated broad substrate profiles. These enzymes are active not only against model substrates (i.e. p NP-esters of fatty acids with aliphatic chain lengths between C2 and C16 ), but also against synthetic polyesters such as PLLA, PDLLA, PCL, 3PET (Hajighasemi et al., 2018 , 2016 ; Tchigvintsev et al., 2015 ), indicating the broader biotechnological potential of this organism beyond its established role in the natural attenuation of oil spill pollution. Notably, such substrate promiscuity is common among carboxylesterases and has been extensively studied (Martínez-Martínez et al., 2018 ; Ma et al., 2025 ). Another species within the family, Alloalcanivorax dieselolei (previously known as Alcanivorax dieselolei) , has been reported to encode a promiscuous, solvent-tolerant esterase (Zhang et al., 2014 ). Recent investigations of a strain isolated from plastic marine debris, Alcanivora x sp. 24 (recently renamed to Alloalcanivorax sp. 24), showed that its hydrolase ALC24_4107 exhibited a strong activity vs aliphatic polyester, such as PHB, PHBV, PES, PBS and PCL (Zadjelovic et al., 2020 ). Of note, this very strain was also demonstrated to degrade the weathered low-density polyethylene (LDPE) by recruiting an array of redox enzymes (alkane monooxygenases, P450, laccases) and reactive oxygen species (Zadjelovic et al., 2022 ). As reported elsewhere, a promiscuous hydrolase from metagenomic fragment of Thalassolituus oleivorans (another renowned marine oil-degrader) showed a hybrid ester hydrolase and haloacid- dehalogenase activity (Beloqui et al., 2010 ). As exemplified in the above and in other studies, mining metagenomes of petroleum-enriched microbial communities using activity-based screens is highly regarded as a productive approach to attribute activities to yet uncharacterised enzymes and new biochemical activities from hydrocarbon-based marine environments (Ferrer et al., 2005 ; Popovic et al., 2015 , 2017 ). This study aimed to identify novel polyesterases from Alcanivoracaceae species with the potential to degrade a wide range of synthetic polyesters, such as 3PET, PLLA, PDLA, PDLLA, PCL, PBA, PBS, and PC. While several esterases from Alcanivoracaceae species have been previously characterised, this research extends the investigation to further predicted polyesterases across a broader selection of strains from this family including Alloalcanivorax sp. 24, Alloalcanivorax gelatiniphagus and Isoalcanivorax pacificus (previously known as Alcanivorax gelatiniphagus and Alcanivorax pacificus , respectively). The genomes of these organisms encode a variety of putative α/β hydrolases and carboxylesterases, many of which remain uncharacterised and present exciting opportunities for further exploration. Notably, Alcanivoracaceae species are naturally adapted to marine environments, making them particularly well-suited for applications in low temperatures conditions and energy-efficient industrial processes. Additionally, polyesterases have demonstrated considerable promise in various sectors, such as waste management, bioremediation, and the development of biodegradable plastics, further emphasising the relevance of this research for sustainable and environmentally industrial practices. Materials and Methods Chemicals All chemicals and substrates used in this study were of analytical grade. Substrates chromogenic p -nitrophenyl ( p NP) esters (C 2 -C 16 ) were purchased from Sigma-Aldrich/Merck (Gillingham, UK) and Tokyo Chemical Industry UK Ltd. (TCI, Oxford, UK). 6-hydroxycaproic acid (6-HHA), lactic acid (LA), succinic acid (SA), adipic acid (AA), terephthalic acid (TA) and bisphenol-A, mono 2-hydroxyethyl terephthalic acid (MHET), and bis(2-hydroxyethyl) terephthalic acid (BHET), Tween-20, dichloromethane (DCM) and 1,1,1,3,3,3-hexafluoro-2-propanol (HFP) were purchased from Sigma-Aldrich/Merck (Gillingham, UK). Polyester substrates, poly-D, L-lactide (PDLLA, molecular weight M W 10,000–18,000), polycaprolactone (PCL14, average M W ~14,000) (PCL2, M W ~2,000), and amorphous polyethylene terephthalate (aPET, thickness 0.25 mm, coil width 600 mm), polybutylene succinate (PBS, M W 45,500), poly-1,4-butylene adipate) (PBA, M W 12,000), poly(1,4-butylene) terephthalate (PBT, M W 38,000), poly-L-lactic acid (PLLA, M W 15,000–25,000), poly-D-lactic acid (PDLA, M W 10,000–15,000), polycarbonate (PC, 3 mm granules) were purchased from Sigma-Aldrich/Merck (Gillingham, UK). The PET model substrate, bis(benzoyloxyethyl) terephthalate (3PET, M W 462.4), was synthesized by CanSyn (Toronto, Canada). Impranil® DLN was kindly donated by WhitChem Ltd, UK (Azelis, www.whitchem.co.uk ). Isopropyl β-D-1-thiogalactopyranoside (IPTG), ampicillin, 2-(cyclohexylamino)ethanesulfonic acid (CHES), sodium chloride (NaCl), tris(hydroxymethyl)aminomethane hydrochloride (Tris-HCl), methanol, sulfuric acid, orthophosphoric acid, and Luria-Bertani (LB) broth were also purchased from Sigma-Aldrich/Merck (Gillingham, UK). Gene cloning and protein purification The coding sequences of selected hydrolase genes were commercially synthesized (without signal peptides, with addition of N-terminal hexahistidine tags) and cloned into a modified p15TVL (TWIST BIOSCIENCE, South San Francisco, USA). All plasmids were transformed into the E. coli Lobstr BL21(DE3) cells (Kerafast, Boston, USA). E. coli cultures were grown aerobically in 2.5 L baffled Erlenmeyer flasks with 1 L Luria-Bertani medium supplemented with 4 g/L glycerol and 100 µg/ml ampicillin at 37°C, in a shaking incubator at 200 rpm, to the optical density (OD 600 ) 0.6–0.8, then spiked with 0.4 mM IPTG, transferred to the 16 ºC and incubated at that temperature in a shaker for further 16 h. The biomass was harvested by centrifugation at 4,000 rpm, 10 min (Avanti J26 rotor JLA8.1, Beckman Coulter Life Sciences, Indianapolis, USA) and disrupted in ice bath by sonication (Q-sonica, Newtown, USA) for 10 min at 70% intensity, in 4 s pulses with 5 s cooling time. Recombinant proteins were purified to near homogeneity (> 95%) using nickel-chelate affinity chromatography on Ni-NTA Superflow resin (QIAGEN, Hilden, Germany) as described previously (Distaso et al., 2023 ). Purity and protein size of purified enzymes were assessed using denaturing 10% polyacrylamide gel (BioRad Laboratories, Hercules, USA) electrophoresis, whereas protein concentration was measured by Bradford assay (Bio-Rad Laboratories, Hercules, USA). Carboxylesterase assays with soluble chromogenic substrates The chromogenic p -nitrophenyl ( p NP) esters of fatty acids with different acyl chain lengths: p NP-acetate (C2), p NP-butyrate (C4), p NP-hexanoate (C6), p NP-octanoate (C8), p eps-decanoate (C10), p NP-dodecanoate (C12), p NP-myristate (C14), p NP-palmitate (C16) were used to test carboxylesterase activity of purified proteins. The activity was screened in 96 well plates spectrophotometrically using SpectraMax M3 (Molecular Devices, San Jose, USA). All assays were conducted in triplicates at indicated temperatures in 96-well plates with reaction mixtures (200 µl) containing 50 mM CHES (pH 9.0) buffer (or as indicated), 1 mM substrate (and 0.1-1 µg of enzyme. The reaction mixtures were incubated for 20 min at 30 ºC, and the activity was calculated based on the absorbance of p -nitrophenol at 410 nm (ε = 17.8 mM − 1 cm − 1 ). Enzyme reaction conditions and stability determination The effect of pH on carboxylesterase activity of purified proteins (pH profile) was determined using the universal Britton-Robinson buffer system (50 mM acetic acid, 50 mM phosphoric acid, 50 mM boric acid, pH range 4.0 to 10.5). Temperature optimum (temperature profile) for carboxylesterase activity of selected enzymes was measured at temperatures from 30 ºC to 90 ºC using 1 mM p NP-octanoate substrate and 0.1-1 µg of protein in 50 mM CHES buffer (pH 9.0). 200 µl assay reactions were incubated in 96 well plate in the Thermomixer (Eppendorf, Hamburg, Germany) at 500 rpm. For the analysis of their thermal stabilities, selected proteins (1 mg/ml) were incubated for 2 h for thermostability and 5 h for cold stability (or as indicated) at different temperatures (from 5 ºC to 90 ºC) in the PCR thin wall tubes in T100 Thermal Cycler (Bio-Rad Laboratories, Hercules, USA), and the residual carboxylesterase activity was measured with 1 mM p NP-octanoate at 30 ºC as described in the carboxylesterase assay methods section. The effect of NaCl and Tween20 on carboxylesterase activity of selected proteins was analysed using 1.0 mM p NP-octanoate and 0.1-1 µg of enzyme. In the reaction mixture of 200 µl containing NaCl at concentrations between 0.1 and 2.0 M and Tween-20 in the range of 0.1- 3.0% in 50 mM CHES, pH 9.0. Reaction mixtures were incubated for 20 min at 30 ºC, and the activity was measured spectrophotometrically at 410 nm. The effect of organic solvents on carboxylesterase activity was analysed in the conditions described in carboxylesterase assays with soluble chromogenic substrates section, but supplied with various concentration of dichloromethane, dimethylsulfoxide, ethanol, and 1,1,1,3,3,3-hexafluoro-2-propanol in the range between 0 and 50%, in 50 mM CHES, pH 9.0. Analysis of temperature-dependent protein denaturation The melting temperature of selected enzymes was measured by differential scanning fluorimetry (DSF) on a QuantStudio 6 Flex system (Applied Biosystems, Thermo Fisher Scientific, Waltham, USA) using SYPRO Orange (Invitrogen, Carlsbad, USA) as a binding dye. Experiments were conducted in triplicate in 30 µl reaction volumes with 10 µg of protein in 50 mM CHES buffer (pH 9.0) and 25X SYPRO Orange in sealed optically clear 96-well plates (Applied Biosystems, Thermo Fisher Scientific, Waltham, USA) with the system set on the ROX as the reference wavelength using the 450/490 nm excitation and 560/580 nm emission filters. The temperature was increased from 25 to 95 ºC with an increment of 1 ºC s − 1 . T m values for selected proteins were determined by non-linear fitting of Sigmoidal-Boltzmann equation using the GraphPad Prism software (version 5.0). Preparation of polyester substrates and polyesterase assays Polyester substrates used in this study (PDLLA, PLLA, PDLA, PCL14, PCL2, 3PET, PBS, PBA, PBT, aPET) were prepared in 50 mM Tris-HCl (pH 8.0), as described previously (Distaso et al., 2023 ). For agarose-based screens, 0.5% polyester emulsions were diluted with three volumes of CHES buffer (pH 9.0) in 2% (w/vol) molten agarose, poured and solidified in the round 90 mm Petri dishes to make opaque gel with the final concentration of polyesters 0.125% (w/vol). Impranil DLN was used only for agarose gel plates preparation. For this, 1 ml Impranil® DLN dissolved in 100 ml 50 mM CHES (pH 9.0), with the final concentration of Impranil 1% in 2% (w/vol) agarose. 50 µg enzyme aliquots were loaded into cylindrical boreholes cored in the agarose using cut 1 ml micropipette tips. Plates were placed into plastic bags to prevent evaporation, incubated at 30 ºC and monitored for 2–3 days. The formation of a clear zone around the wells was considered as indication for the presence of polyester-degrading activity. For polyesterase assays in solution, the 200 µl reaction mixtures with 50 µg of purified protein and 0.125% of emulsified polyester in 50 mM CHES buffer (pH 9.0) were incubated for 12 h at 30 ºC in a shaker at 500 rpm. The reactions were spun down at 14,000 g for 10 min in a minicentrifuge, and supernatant was filtered through 10 kDa spin filters (14,000 g, 15 min, Eppendorf 5424 centrifuge, Eppendorf AG, Hamburg, Germany). The presence of polyester degradation products in filtrates was analysed using HPLC Prominence-I LC-2030C 3D Plus equipped with UV-VIS detector (Shimadzu, Kyoto, Japan). For aPET, 3PET, PBT the formation of terephthalic acid (TA), mono-2-hydroxyethyl terephthalic acid (MHET), bis-2-hydroxyethyl terephthalic acid (BHET) was analysed. For PC depolymerisation the bisphenol-A (BPA), was analysed. For product analysis the reverse-phase hydrophobic chromatography on a Shimadzu C 18 Shim-pack column (150x4.6 mm, 5 µm particle size) (Shimadzu Corporation, Kyoto, Japan), 40 ºC (injection volume 10 µl, detection at 240 nm) was used. For products separation we used the mobile phase of Solvent A (0.1% (vol/vol) of orthophosphoric acid (H 3 PO 4 ) in the HPLC grade water and Solvent B (100% methanol). At flow rate of 0.7 ml/min, 40 ºC, the gradient was: 0–2 min, 25% Solvent B; 2–18 min, linear gradient to 55% of Solvent B; 18–22 min, linear gradient to 25% of Solvent B. For PDLA, PLLA, PDLLA depolymerisation efficiency, lactic acid was quantified; for PCL14, PCL2, 6-hydroxyhexanoic acid; for PBS depolymerisation, succinic acid; for PBAT, adipic acid were quantified using ion-moderated partition chromatography on a Prominence-I LC-2030C 3D Plus HPLC system equipped with an Aminex HPX-87-H column (300x7.8 mm, 9 µm particle size, conditioned at 50 ºC) (Shimadzu Corporation, Kyoto, Japan) and a UV detector (LC-2030C_3Dplus, Shimadzu Corporation, Kyoto, Japan). For analysis, 0.05 N sulfuric acid (H 2 SO 4 ) in HPLC grade water was used as solvent at flow rate of 0.6 ml/min, detection at 210 nm. Polyester degradation products were quantified based on their calibration curves generated using commercially available standards (TA, MHET, BHET, LA, 6-HHA, AA, SA, BPA). Bioinformatic and structural analyses Multiple sequence alignments of selected carboxylesterases were performed using the MAFFT online service, automated regime (Katoh et al., 2019 ). Alignment was visualised and modified using the online software STRAP (Gille et al., 2014 ). For phylogenetic analysis, 5 Alcanivoracaceae strains genomes were screened for IPR029058 (AB hydrolase) signature containing sequences, and the 297 sequences were retrieved, and aligned in Geneious Prime using global alignment with free end gaps, Cost matrix Blosum 62, and Fast tree was used for phylogeny analysis. Structural analysis was performed in ChimeraX (Meng et al., 2023 ). Results Analysis of Alcanivoracaceae genomes for potential carboxylesterases and novel polyesterases In this study, we used five reference Alcanivoracaceae genomes ( Alcanivorax borkumensis SK2 T , A. hongdengensis A-11-3 T , A. sediminis PA15-N-34 T , A. profundi MTE017 T , and A. nanhaiticus 19-m-6 T ), for screening of IPR029058 (ɑ/β hydrolase) signature containing sequences. 297 sequences were retrieved from GenBank, 5 sequences manually selected from Isoalcanivorax pacificus previously known as Alcanivorax pacificus (APA_2, APA_3, APA_4, APA_5, APA_6), 6 sequences (ABO_0116, ABO_1197, ABO_1251, ABO_1483, ABO_1895, ABO_2249) from A. borkumensis , 5 sequences (ALC24_3989, ALC24_1328, ALC24_1162, ALC24_2069, ALC24_4107) from Alloalcanivorax sp. 24 (SI Table 1) and added for multiple sequence analysis (SI Fig. 1). Selected esterases from Alcanivorax borkumensis (ABO_0116, ABO_1197, ABO_1251, ABO_1483, ABO_1895, and ABO_2249) were partially characterised in previous studies (Hajighasemi et al., 2016 , 2018 ; Tchigvintsev et al., 2015 ), which is highlighted in the phylogenetic tree (Fig. 1). Sequence analysis of these proteins revealed that ABO_1197, ABO_1895, APA_6 and APA_5 have signal peptides (S1 Table 1) suggesting that majority of Alcanivoracaceae esterases are involved in the intracellular metabolism. Selected sequences shared identity from 4.6% to 66.5% with characterised enzymes from Alcanivorax borkumensis SK2 (ABO_0116, ABO_1197, ABO_1251, ABO_1483, ABO_1895, and ABO_2249) (SI Table 2) and according to the Arpigny and Jaeger ( 1999 ) classification of lipolytic enzymes, belong to the carboxylesterase family V with the characteristic catalytic Ser motif GX S XGG (SI Fig. 1). Based on the sequence analysis eight proteins (APA_2, ALC24_4107, AGE_2, APA_8, AGE_8, ABO_1898, ABO_0116, ABO_1483) belong to family V, subfamily I, sharing serine motif with GX S XG sequences. There were 5 sequences referred to family V subfamily III (AGE_12, ALC24_2069, ALC24_1162, ABO1197, ABO2449), sharing serine motif with GX S GG, and the rest 8 sequences belonging to family V subfamily IV (APA_5, ALC24_1328, AGE_3, APA_4, ALC24_3989, APA_3, APA_6, ABO1251), with the serine motif GX S XGX (Fig. 1). Carboxylesterase activity against chromogenic monoester substrates The selected 21 genes encoding \(\:{\alpha\:}/{\beta\:}\) -hydrolases and predicted carboxylesterases (S1 Table 1) were recombinantly expressed in E. coli with an N-terminal 6His-tag and affinity purified (SI Fig. 2). From 17 cloned genes, 11 were found to be expressed as soluble proteins, but only 5 clones produced highly purified proteins useful for detailed analysis (AGE_12, APA_2, APA_3, APA_5 and ALC24_1328) (S1 Table 1). Furthermore, six previously published esterases from A. borkumensis SK2 T (ABO_0116, ABO_1197, ABO_1251, ABO_1483, ABO_1895, and ABO_2249) were also induced and expressed and purified in this study (Tchigvintsev et al., 2015 ; Hajighasemi et al., 2016 , 2018 ). These 11 purified esterases were initially screened at 30 ºC using a range of chromogenic p NP-esters with varying acyl chain lengths from C2 to C16 (Fig. 2). These screens revealed the presence of carboxylesterase activity in all purified enzymes, with most enzymes showing the highest activity with short-chain esters (acetyl and butyl- p NP) and low activity against long acyl chain esters. In agreement with previous studies (Hajighasemi et al., 2016 , 2018 ; Tchigvintsev et al., 2015 ; Urbanek et al., 2020 ) ABO_0116, ABO_1197, ABO_1251, ABO_1895, and ABO_2249 showed the preference to the short chain length substrates. Notably, ABO_2249 displayed the highest activity with p NP-hexanoyl, consistent with previous studies (Urbanek et al., 2018 ) highlighting it as promising candidate for polyester plastic depolymerisation. From five novel Alcanivoracaceae proteins (AGE_12, APA_2, APA_3, APA_5 and ALC24_1328), APA_5 and ALC24_1328 exhibited the highest carboxylesterase activity with the C2 and C4 substrates, whereas the remaining three (AGE_12, APA_2 and APA_3) showed low activity (Fig. 2). Activity of carboxylesterases against different polyesters 11 purified carboxylesterases from Alcanivoracaceae were then screened for depolymerisation activity against various polyesters, including 3PET, aPET, PCL14, PCL2, PDLA, PLLA, PDLLA, PBA, PBS, PBT, PC, and Impranil DLN. Polyesterase assays were conducted using both the agarose plate-based screens and HPLC analyses of soluble reaction products after enzyme incubation with plastic emulsions. Visual inspection of agar plates revealed halos only around ABO_2249, ABO_1251, ALC24_1328, and ABO_1197 for 3PET or PCL2 (SI Fig. 3). However, no polyesterase activity against aPET, PLLA/PDLA, PLA, PBT, or PBS were detected in agarose plate screens. HPLC assays (Fig. 3) appeared more sensitive and revealed even slight depolymerisation not detected by agar plate visual inspection. Thus, polyesterase activity with short-chain polymers like 3PET or PCL2 was observed for all esterases (Fig. 3), indicating enzymes’ promiscuity and their potential for plastic-degrading activity. Furthermore, the HPLC analysis was performed to detect soluble reaction products following incubation of purified enzymes with emulsified polyester substrates aPET, PCL14, PDLA, PLLA, PDLLA, PBA, PBT and PC. These assays followed the formation of various polyester degradation products, such as TA, MHET, BA from 3PET, 6-HHA from PCL14 and PCL2, LA from PDLA, PLLA and PDLLA; AA from BPA, SA from PBS, TA from PBT, and BPA from PC. In these experiments, positive results were observed for PCL14, PCL2, PDLLA, PBA and 3PET as substrates, but no polyesterase activity was detected against aPET and PBS, PDLA, PLLA or PC (SI Table 3). Interestingly, the major product during 3PET depolymerisation was MHET, with minor amounts of TA, and none of them producing BHET indicating that the selected enzymes possess high BHETase activity and low MHETase activity (Fig. 3). ABO_1197, ABO_1251, ALC24_1328 and APA_5 exhibited high polyesterase activity against 3PET, generating significant amount of BA, MHET and TA (from 0.2 to 1.35 mM). In contrast, ABO_0116, ABO_1483, ABO_1895, ALC24_1328 and APA_3 showed low polyesterase activity, producing little to no BA, MHET, or TA as products (< 0.1 mM). Surprisingly, the highest accumulation of MHET (at concentration of 0.78 mM) was observed for ABO_1251. Since this enzyme produced no visible halos on the 3PET-agrose plates, this discrepancy highlights the higher precision of HPLC analysis for quantifying the depolymerisation activity. With shorter-chain substrate PCL2, all tested Alcanivoracaceae enzymes were active, generating 6-hydroxyhexanoic acid (6-HHA) at concentrations ranging from 0.1 to 0.5 mM. However, only five enzymes (ABO_1197, ABO_1251, ABO_2249, ALC24_1328 and APA_5) exhibited activity against the longer-chain PCL14 producing 6-HHA in the range 0.2–1.7 mM) (SI Table 3). A similar trend was observed with PDLLA (poly-D, L-lactide, MW range 10–18 kDa), where ABO_1197, ABO_1251, ABO_2249, APA_5, and ALC24_1328 produced significant levels of lactic acid (from 0.1 to 0.6 mM) (SI Table 3). The PDLLA polymer consists of a racemic mix of L- and D-lactic acid monomers, making it less crystalline than PLLA (L-lactic acid polymer) or PDLA (D-lactic acid polymer) (Lim et al., 2008 ), which may facilitate its enzymatic degradation. For PBA degradation, six enzymes (ABO_1197, ABO_1251, ABO_1895, ABO_2249, AGE_12, APA_5) demonstrated hydrolytic activity, producing adipic acid at concentrations ranging from 0.1 to 0.3 mM (SI Table 3). In conclusion, five esterases, ABO_1197, ABO_1251, ABO_2249, APA_5, and ALC24_1328 were selected for further characterisation based on their promising polyesterase activities. Substrates and reaction products analysed in this study are presented in the supplementary SI Table 3. Analysis of optimal reaction conditions for polyester depolymerisation To identify the potential industrial applications of enzymes, they need to be screened for several key parameters, including substrate specificity, thermal stability to withstand industrial processing temperatures, pH stability across various industrial conditions, catalytic efficiency, and resistance to potential inhibitors present in industrial environments. The selected enzymes ABO_1197, ABO_1251, ABO_2249, APA_5, and ALC24_1328 were screened for their optimal conditions regarding pH, salinity (NaCl concentrations), detergent (Tween20), and temperature. The pH range was screened from pH 4.0 to pH 10.5, revealing that all selected enzymes preferred alkaline conditions (pH 8.0–pH 9.0) (Fig. 4A). Salinity tolerance was assessed with NaCl concentrations varying within 0–2.0 M, demonstrating that ABO_1197, ABO_1251, and ALC24_1328 maintained over 90% of activity at concentrations up to 0.1 M NaCl, while ABO_2249 and APA_5 generally exhibited lower salt tolerance. Notably, ABO_1197 showed the highest residual activity at 2 M NaCl concentration, retaining 23% of its initial activity (Fig. 4B). For detergent tolerance, the non-ionic surfactant Tween20 was used, ranging at concentrations from 0.1% to 2.0%. ABO_1197 and ABO_1251 showed increased activity with Tween20, which is often attributed to enhanced substrate accessibility. However, ABO_2249 and APA_5 experienced a decrease in activity even with small traces of detergent, while ALC24_1328 displayed remarkable detergent resilience, increasing its activity by up to 25% at 2.0% Tween20 (Fig. 5). Temperature profiles were assessed from 5–70 ºC, with all selected enzymes showing mesophilic traits, favouring temperatures between 10 ºC and 50 ºC. Most enzymes lost two-thirds of their activity at 60 ºC. ABO_2249 and APA_5 demonstrated notable cold tolerance, with activities at 5 ºC being just 24% less than their respective optima (Fig. 6). Analysis of thermostability In the literature, protein stability is commonly assessed by measuring the melting temperature (Tm), which is defined as the temperature at which equal amounts of the protein are folded and unfolded under specific conditions (Huynh and Partch, 2015 ; Sorgenfrei et al., 2024 ). All applicable methods share the same principle: a sample is gradually heated, protein stability is monitored by melting temperature, which either measured by fluorescence signal of internal tryptophan residues or by using a fluorescence dye that changes its signal upon interacting with the unfolding protein. In this study, we investigated the thermostability of purified polyesterases by analysing the remaining enzyme activity after preincubation at different temperatures, as well as by identifying protein melting temperatures (T m ) using DSF. As shown in Fig. 7, after 5 h of preincubation at temperatures ranging from 5 to 40 ºC, the polyesterases retained almost 100% of their initial activity at 5 ºC. ABO_1197, ABO_1251, and ABO_2249 exhibited increased activity at 5 ºC and 10 ºC, however, ABO_1197 and ABO_1251 lost 40% and 80% of their activity at 30 ºC and completely lost it at 40 ºC. APA_5 and ALC24_1328 showed only a slight (2–5%) decrease in activity after incubation at 5 ºC. Overall, the selected enzymes maintained good activity within the low-temperature range (5–30 ºC) over prolonged incubation, with a maximum tolerance temperature of 30 ºC. Protein melting temperatures (T m ) were determined based by DSF and are shown in SI Fig. 4. The highest melting temperature was observed for APA_5 (T m 48.1 ºC), correlating with its high residual activity at 30 ºC after 5 h of incubation (Fig. 7). Analysis of solvent resistance Organic solvent tolerance of enzymes was determined by measuring activity in presence of solvents concentrations varying from 0–50%. Four different water-miscible organic solvents were chosen: ethanol, DMSO, dichloromethane (DCM), and 1,1,1,3,3,3-hexafluoro-2-propanol (HFP) based on their relevance for synthetic organic chemistry and for synthetic polymers. DCM and HFP are commonly used to dissolve the polyesters and were used for emulsions preparation in this work. In presence of HFP all enzymes retained less than 5% of their initial activity. Similar negative effect was observed for DCM (Fig. 8), with complete loss of the activity with more than 10% of DCM in reaction mixture. DMSO overall had a positive effect on the screened enzymes. ABO_1197, ABO_1251 and ABO_2249 showed 10–20% increase in the activity in presence of up to 20% DMSO, for APA_5 DMSO dramatically increased the activity, adding up to 50% of solvent in reaction mixture. Ethanol negatively influenced ABO_1197 and ABO_1251 activity, leading to decrease even at 5% of solvent in reaction mixture, but at the same time increased activity for the ABO_2244, APA_5 and ALC24_1328. For APA_5 at presence of 20% of ethanol in reaction mixture, activity was increased by 84%. ABO_2249, APA_5 and ALC24_1328 showed relative tolerance to the polar solvents of low polarity, increasing sensitivity with the increase of solvents polar index (DMSO < ethanol < HFP). 50% of DMSO retained positive effect on these enzymes, though for ethanol, less than 1% of activity was retained at 50% reaction mixture solution. Combined effects of solvents and temperature on the enzymatic polyester depolymerisation In this study, selected enzyme-maintained activity at low temperatures 5–40 ºC (Fig. 9). ABO_2249 and APA_5 exhibited highest cold adaptation, with activities maintained at 70–85% at 5 ºC, compared to those at 30 ºC. We tested our enzymes activity with model substrate p NP-hexanoate, in presence of DMSO and ethanol and observed improvement in the activity up to 25 to 100% with presence of 10–20% (v/v) for APA_5 and AlC24_1328. DCM and HFP showed stronger inhibiting effect on target enzymes activity (Fig. 8). Based on these results we selected 20% ethanol and DMSO to increase enzyme activity and 5% for DCM aiming decrease in crystallinity of substrate in the reaction mixture with actual polymeric substrates: 3PET, PCL14 and PDLLA. The incubations took place at 5 ºC and 30 ºC to make later comparisons. The reaction mixtures were filtered (10 kDa spin filters) and analysed by HPLC, the concentration of end products (mM) was used as standard to define the depolymerisation efficiency (Fig. 9). The results from 5 ºC incubation with 3PET, the presence of DMSO for enzyme APA_5 and ALC24_1328 increased production of BA and MHET compared with ‘no solvent’ conditions. For the rest of enzymes (ABO_1197, ABO_1251 and ABO_2249) any solvent addition at 5 ºC, resulted in decrease of depolymerisation efficiency. Addition of organic solvents DMSO, ethyl alcohol and DCM at 30 ºC slightly improved the depolymerisation efficiency for reactions with APA_5 and ALC24_1328, with both PLA and 3PET (Fig. 9, 10) when compared with no solvent conditions. Interestingly, even accounting for the protein activity inhibition at 5% DCM (Fig. 9), the increase in product concentration for 3PET depolymerisation for APA_5 and ALC24_1328 was observed at 30 ºC, however at 5°C, the DCM addition decreased the yield of monomeric products threefold, as compared with 30°C. At the same time, if accounting only temperature effect on the enzyme activity more than twofold reduction in reaction products accumulation was observed in samples without solvent. The DCM has higher solubility in water at 5 vs 30 ºC (U.S. National Institute of Standards and Technology (NIST), Solubility Database, SRD 106, https://srdata.nist.gov/solubility/ ), therefore at lower temperatures the non-polar interaction with protein structures must increase, potentially causing negative structural changes in proteins. DCM showed improved results for 3PET depolymerisation for APA_5 and ALC24_1328 only at 30 ºC. Discussion In this study, eleven α/β-hydrolases from Alcanivoracaceae were purified and biochemically characterised for their carboxylesterase and polyesterase activities, including six previously reported enzymes (Hajighasemi et al., 2016 , 2018 ; Tchigvintsev et al., 2015 ), which were included as reference proteins (ABO_0116, ABO_1197, ABO_1251, ABO_1483, ABO_1895 and ABO_2249). All purified proteins exhibited carboxylesterase activity toward chromogenic p NP-esters with a broad substrate range and a preference for short-chain acyl esters, which is typical for carboxylesterases (Martínez-Martínez et al., 2018 ) (Hajighasemi et al., 2016 , 2018 ; Tchigvintsev et al., 2015 ). HPLC-based assays with emulsified polyesters revealed detectable or significant polyesterase activity in all enzymes against PCL2 and the model PET-like substrate 3PET (Fig. 3). Six enzymes also hydrolysed PBA, five proteins degraded PCL14 and PDLLA, while no polyesterase activity was observed with other tested polyesters (aPET, PLLA, PDLA, PBS, PC, and PBT). These results confirm and extend our recent observations that polyesterase activity is widespread in carboxylesterases from families IV and V (Ma et al., 2025 ). Moreover, our findings reinforce that HPLC-based assays provide greater sensitivity than agarose clearing methods for detecting polyesterase activity in purified proteins (Ma et al., 2025 ). The Alcanivoracaceae enzymes preferentially degraded aliphatic polyesters with shorter chain lengths and lower molecular weights, while aromatic polyesters were recalcitrant. For example, polyesterase activity was observed on substrates such as PCL2, PCL14, PDLLA, PBA, whereas limited or no activity was detected on PC, PBT and aPET. Generally, the polyester chain length strongly influences chain mobility, crystallinity, and thus susceptibility to enzymatic hydrolysis. Longer and more crystalline polymers exhibit enhanced intermolecular interactions, such as Van der Waals forces, requiring higher activation energy for depolymerisation. From an industrial perspective, cold-adapted enzymes offer distinct advantages, including reduced energy costs and protection of thermolabile products and intermediates (Santiago et al., 2016 )). However, because polyester crystallinity remains high at low temperatures, appropriate decrystallinization pretreatments will likely be required to enable complete depolymerization with cold-adapted enzymes. Marine microorganisms represent an important reservoir of cold-active and cold-tolerant enzymes due to the predominantly low temperatures of seawater habitats (Médigue et al., 2005 ; Kube et al., 2013 ; Santiago et al., 2016 ). For instance, EstB from Alloalcanivorax dieselolei , displayed optimal activity around 20 ºC and retained over 95% of activity between 0 and 10 ºC (Degli-Innocenti et al., 2023 ; Zhang et al., 2014 ). Similarly, RhLip from Rhodococcus sp. AW25M09 retained 50% activity at 10 ºC (Hjerde et al., 2013 ), and a carboxylesterase from Psychrobacter cryohalolentis maintained over 90% of its maximal activity at 0–5 ºC (Novototskaya-Vlasova et al., 2012 ). The carboxylesterase OLEI01171 from Oleispira antarctica RB-8, another marine hydrocarbon degrader, also exhibits high activity between 5 and 30 ºC (Lemak et al., 2012 ). Structural analyses of enzymes from O. antarctica RB-8 and their mesophilic homologues suggested that enhanced flexibility at the active sites underlies the catalytic efficiency of cold-active enzymes, compensating for the reduced thermal energy in cold environments (Kube et al., 2013 ). Consistent with this, all enzymes characterised in this study were highly active and stable at low temperatures (5–20°C), with protein melting temperatures (T m ) ranging from 30 to 48 ºC. Enzymes with lower T m values (ABO_1197 and ABO_1251) exhibited partial reactivation at 5°C, whereas Apa_5 (T m 48.1°C) retained activity over a wider temperature range. These results indicate that lower T m values correlate with increased cold tolerance, while higher T m values correspond to greater enzyme thermostability. The enhanced cold tolerance observed in ABO_1197 and ABO_1251 may result from increased conformational flexibility conferred by residues such as Gly, Ser, and Met, and a lower abundance of Pro and Arg residues, which typically constrain structural mobility (Parvizpour et al., 2021 ). However, comparative sequence analysis of cold-active and thermostable esterases did not reveal a consistent correlation between amino acid composition and cold tolerance. Several Alcanivoracaceae polyesterases (ABO_1197, ABO_1251, and ALC24_1328) exhibited high tolerance or partial activation in the presence of Tween 20 (Fig. 5), a feature frequently observed among cold-adapted α/β-hydrolases. For instance, the three thermophilic metagenomic polyesterases from Ischia showed no activation and low tolerance to detergents (Distaso et al., 2023 ), whereas several cold-adapted carboxylesterases from marine bacteria and a cold-active protease from Psychrobacter sp. 94-6PB were stimulated by the addition of detergents, including Tween 20 (Novototskaya-Vlasova et al., 2012 ; Wu et al., 2013 ; Jiang et al., 2016; Perfumo et al., 2020 ). Organic solvents can enhance enzymatic transformations by increasing substrate solubility and modulating enzyme flexibility via alterations in hydrogen bonding (Osbon and Kumar, 2019; Sorgenfrei et al., 2024 ). However, this simultaneously requires a certain level of enzyme tolerance to organic solvents. Given that Alcanivoracaceae species thrive in oil- and hydrocarbon-rich environments (Sabirova et al., 2006 ; Zhang et al., 2014 ), their enzymes are expected to exhibit tolerance to both low temperatures and organic solvents. Nevertheless, the inherent structural flexibility of cold-adapted enzymes may render them more susceptible to the destabilizing effects of elevated temperatures and organic solvents (Sellek and Chaudhuri, 1999 ). To date, only a limited number of cold-adapted esterases and lipases with solvent tolerance have been described (Lee et al., 2017 ). For instance, high solvent resistance was reported for the EstB esterase from Alloalcanivorax dieselolei B-5 T with the retention of 80% activity in the presence of isopropanol (70%) or acetone (up to 70%) (Zhang et al., 2014 )). In the presence of 50% dimethyl sulfoxide (DMSO) or ethanol, EstB retained 83% and 87% of activity, respectively, but it was strongly inhibited by acetonitrile and methanol (Zhang et al., 2014 ). In the present study, the addition of DMSO up to 20% increased the carboxylesterase activity of all tested enzymes from Alcanivoracaceae (Fig. 8). Ethanol at 10–20 % stimulated the activity of three enzymes but was inhibitory at higher concentrations, while DCM (dichloromethane) and HFP (hexafuoroisopropanol) caused strong inhibition. The addition of DMSO or ethanol enhanced 3PET depolymerisation by APA_5 and ALC24_1328 at 30°C, whereas no stimulation was observed at 5°C (Fig. 9). These findings suggest that, within the optimal range, higher reaction temperatures exert a more pronounced positive effect on enzymatic polyester depolymerisation than the presence of organic solvents. Conclusion This study expands the current understanding of Alcanivoracaceae α/β-hydrolases by demonstrating that polyesterase activity is a common feature among their carboxylesterases, particularly within families IV and V. Eleven enzymes tested exhibited carboxylesterase activity with a preference for short-chain monoesters, as well as polyesterase activity toward aliphatic polyesters such as PCL and PBA, while showing limited activity toward aromatic polyesters. Their notable activity and stability at low temperatures, together with tolerance to detergents and certain organic solvents, reflect their adaptation to hydrocarbon-rich marine environments. These findings provide new insights into the biochemical diversity and ecological roles of Alcanivoracaceae enzymes and highlight their potential for environmentally friendly biotransformations, particularly in low-temperature, energy-efficient plastic biodegradation and recycling. Abbreviations 3PET, bis(benzoyloxyethyl) terephthalate; PET, poly(ethylene terephthalate); aPET, amorphous poly(ethylene terephthalate); PU, polyurethane; PE, polyethylene; PS, polystyrene; PC, polycarbonate; PCL14, polycaprolactone (14kDa); PCL2, polycaprolactone (2 kDa); PBS, polybutylene succinate; PBA, polybutylene adipate; PBT, polybutylene terephthalate; PLLA, poly-L-lactide; PDLA, poly-D-lactide; PDLLA, poly-D,L-lactide; MHET, mono(2-hydroxyethyl) terephthalate; BHET, bis(2-hydroxyethyl) terephthalate; TA, terephthalic acid; BPA, bisphenol-A; DMSO, dimethyl sulfoxide; DCM, dichloromethane; HFP, 1,1,1,3,3,3-hexafluoro-2-propanol Declarations Funding declaration This research was supported by the FuturEnzyme project funded by the European Union Horizon 2020 Research and Innovation Program under grant agreement 101000327, UKRI-funded projects P3EB (UKRI Engineering Biology Mission Hub, grant BB/Y007972/) and EBIC (UKRI Engineering Biology Mission Hub, grant (grant Nr BB/Y008332/1). PNG and AFY also acknowledge support from the “Plastics Vector” project NE/S004548/1 funded by the Natural Environment Research Council (NERC, UK). OVG, PNG, and AFY are also thankful for support from the European Regional Development Fund (ERDF) through the Welsh Government to the Centre for Environmental Biotechnology, project number 81280. Authors’ contributions PNG, AFY, MF, OVG, and ANK designed experiments and interpreted the data. HM wrote the manuscript draft and edited it with HM, ANK, TNC performed experiments. PNG, MF, AFY and OVG provided the funding. References Abou-Zeid DM, Müller RJ, Deckwer WD (2004) Biodegradation of aliphatic homopolyesters and aliphatic–aromatic copolyesters by anaerobic microorganisms. Biomacromolecules 5(5):1687–1697. https://doi.org/10.1021/bm0499334 Arpigny JL, Jaeger KE (1999) Bacterial lipolytic enzymes: classification and properties. 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1","display":"","copyAsset":false,"role":"figure","size":1474447,"visible":true,"origin":"","legend":"\u003cp\u003ePhylogenetic analysis of 297 α/β hydrolase sequences from the genomes of Alcanivoracaceae species, including Alcanivorax borkumensis SK2\u003csup\u003eT\u003c/sup\u003e, Alcanivorax hongdengensis A-11-3\u003csup\u003eT\u003c/sup\u003e, Alcanivorax sediminis PA15-N-34\u003csup\u003eT\u003c/sup\u003e, Alcanivorax profundi MTEO17\u003csup\u003eT\u003c/sup\u003e, and Alcanivorax nanhaiticus 19-m-6\u003csup\u003eT\u003c/sup\u003e, all containing the IPR029058 (AB hydrolase) signature. Red stars indicate previously published proteins (ABO_0116, ABO_1483, ABO_1895, ABO_1251, ABO_1197, and ABO_2249), while blue stars denote proteins identified in this study (AGE_2, AGE_3, AGE_8, AGE_12, APA_2, APA_3, APA_4, APA_5, APA_6, APA_8, ALC24_1162, ALC24_3989, ALC24_1328, ALC24_2069, and ALC24_4107). The scale bar represents one substitution per position.\u003c/p\u003e","description":"","filename":"1.png","url":"https://assets-eu.researchsquare.com/files/rs-7917997/v1/dce83e9f851093d41036b037.png"},{"id":96244234,"identity":"7c478e0d-12a3-45d1-b840-b51f6d201b2f","added_by":"auto","created_at":"2025-11-19 07:17:59","extension":"png","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":910090,"visible":true,"origin":"","legend":"\u003cp\u003eAssays of α/β-hydrolases for carboxylesterase activity using chromogenic \u003cem\u003ep\u003c/em\u003eNP-esters with varying acyl chain lengths (C2-C16). Activities were measured in a reaction mixture with 1 µg of enzyme in 50 mM CHES (pH 9.0) and the specified substrates (1 mM) at 30 ºC for 20 minutes. All assays were conducted in triplicates; results represent the means derived from at least two independent determinations. \"n.d.\", not detectable.\u003c/p\u003e","description":"","filename":"2.png","url":"https://assets-eu.researchsquare.com/files/rs-7917997/v1/7e40cfd54d54d851cc058529.png"},{"id":96243567,"identity":"efe80c0a-2d26-4948-ae32-b1089f443391","added_by":"auto","created_at":"2025-11-19 07:16:40","extension":"png","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":369309,"visible":true,"origin":"","legend":"\u003cp\u003ePolyesterase activities of selected carboxylesterases as revealed by reactions products analyses using HPLC. 50 µg of enzymes were incubated with emulsified polyesters at 30 ºC for 16 h, and the indicated reaction products were analysed by HPLC, as described in Materials and Methods.\u003c/p\u003e","description":"","filename":"3.png","url":"https://assets-eu.researchsquare.com/files/rs-7917997/v1/7c1254a5e49337d30778504c.png"},{"id":96242629,"identity":"ce5d5a13-c92e-4d75-bc87-f3790215a2c1","added_by":"auto","created_at":"2025-11-19 07:13:50","extension":"png","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":557022,"visible":true,"origin":"","legend":"\u003cp\u003eEffect of pH \u003cstrong\u003e(A)\u003c/strong\u003e and\u003cstrong\u003e \u003c/strong\u003eNaCl concentrations \u003cstrong\u003e(B) \u003c/strong\u003eon carboxylesterase activity of selected polyesterases with soluble chromogenic substrates. \u003cstrong\u003e(A),\u003c/strong\u003e Reactions were carried out in Britton–Robinson buffer over a pH range of 4.0–10.5 using 1 mM \u003cem\u003ep\u003c/em\u003eNP-octanoate as substrate and 1 µg of enzyme per reaction, incubated for 20 min at 30 ºC. \u003cstrong\u003e(B), \u003c/strong\u003eThe effect of NaCl was tested in the concentration range of 0–2 M in 50 mM CHES buffer (pH 9.0) using 1 mM \u003cem\u003ep\u003c/em\u003eNP-octanoate and 1 µg of enzyme per reaction (incubation for 20 min at 30 ºC).\u003c/p\u003e","description":"","filename":"4.png","url":"https://assets-eu.researchsquare.com/files/rs-7917997/v1/abb326a0db48218f36a6cc77.png"},{"id":95905593,"identity":"5523487e-3929-4b2b-afb7-3889a6f1c4de","added_by":"auto","created_at":"2025-11-14 09:23:11","extension":"png","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":264837,"visible":true,"origin":"","legend":"\u003cp\u003eEffect of Tween20 on carboxylesterase activity of selected polyesterases at concentrations of Tween20 between 0% and 3% (v/v). Assays were done with 1 mM \u003cem\u003ep\u003c/em\u003eNP-octanoate as substrate and 1 µg of enzyme per reaction at 30 ºC for 20 min in 50 mM CHES buffer (pH 9.0).\u003c/p\u003e","description":"","filename":"5.png","url":"https://assets-eu.researchsquare.com/files/rs-7917997/v1/d7dfcde5cc76a32c80821d15.png"},{"id":95905609,"identity":"c2899035-aff7-4ae0-b2a7-aa7369f114d8","added_by":"auto","created_at":"2025-11-14 09:23:11","extension":"png","order_by":6,"title":"Figure 6","display":"","copyAsset":false,"role":"figure","size":399247,"visible":true,"origin":"","legend":"\u003cp\u003eEffect of reaction temperatures on carboxylesterase activity of selected polyesterases. Reaction mixtures contained 1 µg of enzyme, 1 mM \u003cem\u003ep\u003c/em\u003eNP-octanoate as substrate and were incubated at indicated temperatures from 5 ºC to 70 ºC for 20 min in 50 mM CHES buffer (pH 9.0).\u003c/p\u003e","description":"","filename":"6.png","url":"https://assets-eu.researchsquare.com/files/rs-7917997/v1/0becb6e3b2d7696b2c6b6b50.png"},{"id":95905606,"identity":"da2278c4-892c-4038-9bd2-460399639907","added_by":"auto","created_at":"2025-11-14 09:23:11","extension":"png","order_by":7,"title":"Figure 7","display":"","copyAsset":false,"role":"figure","size":721484,"visible":true,"origin":"","legend":"\u003cp\u003eCold tolerance of purified polyesterases: activity-based analysis. Following 5 h pre-incubation at different temperatures from 5 ºC to 40 ºC, residual carboxylesterase activities were measured at 20 ºC with 1 mM \u003cem\u003ep\u003c/em\u003eNP-octanoate as substrate, 1 µg of enzyme per reaction in 50 mM CHES buffer at pH 9.0. “n.d.”, not detectable.\u003c/p\u003e","description":"","filename":"7.png","url":"https://assets-eu.researchsquare.com/files/rs-7917997/v1/694fa8b45d2752a0dd12bb61.png"},{"id":95905599,"identity":"3210e229-946b-4df0-a417-a3fe9ee58743","added_by":"auto","created_at":"2025-11-14 09:23:11","extension":"png","order_by":8,"title":"Figure 8","display":"","copyAsset":false,"role":"figure","size":607329,"visible":true,"origin":"","legend":"\u003cp\u003eEffects of various organic solvents at concentrations from 0 % to 50 % (v/v) on carboxylesterase activity of selected polyesterases. Reaction mixtures contained 1 µg of purified enzyme in 50 mM CHES buffer (pH 9.0) and were incubated for 20 min at 30 ºC . Solvents (0 % to 50 % (v/v): DCM, dichloromethane; DMSO, dimethyl sulfoxide; HPF, 1,1,1,3,3,3-hexafluoro-2-propanol.\u003cstrong\u003eNote:\u003c/strong\u003e assays of ABO_2249 vs DMSO and ALC24_1328 vs ethanol were conducted with enzyme samples derived from separate purification batches\u003c/p\u003e","description":"","filename":"8.png","url":"https://assets-eu.researchsquare.com/files/rs-7917997/v1/1783b3055d223a1363de6c91.png"},{"id":95905616,"identity":"958822f8-f47b-47a6-bdd7-9beef152a615","added_by":"auto","created_at":"2025-11-14 09:23:11","extension":"png","order_by":9,"title":"Figure 9","display":"","copyAsset":false,"role":"figure","size":648271,"visible":true,"origin":"","legend":"\u003cp\u003eEffects of solvents and incubation temperature on 3PET degradation:\u003cstrong\u003e \u003c/strong\u003eHPLC analysis of reaction products. Reaction mixtures contained 50 µg of purified enzyme in 50 mM CHES buffer (pH 9.0) with the indicated solvents and were incubated overnight at 30 ºC (\u003cstrong\u003eA\u003c/strong\u003e) and 5 ºC (\u003cstrong\u003eB\u003c/strong\u003e). Reaction products (BA, MHET, and TA) were analysed by HPLC as described in Materials and Methods.\u003c/p\u003e","description":"","filename":"9.png","url":"https://assets-eu.researchsquare.com/files/rs-7917997/v1/dfef5311cf9de1379b38f48d.png"},{"id":95905613,"identity":"341e316e-42e8-4e8e-b6dd-9498ffc4922a","added_by":"auto","created_at":"2025-11-14 09:23:11","extension":"png","order_by":10,"title":"Figure 10","display":"","copyAsset":false,"role":"figure","size":881430,"visible":true,"origin":"","legend":"\u003cp\u003eEffects of solvents and incubation temperature on the degradation of PCL14 and PDLLA: HPLC analysis of hydrolysis products. PCL14 and PDLLA were incubated with 50 µg of enzyme per reaction in three different organic solvents (DCM, DMSO and ethanol (EtOH)) overnight at 30 ºC (A) and 5 ºC (B), and the formation of reaction products (6-HHA and LA) was analysed using HPLC as described in Materials and Methods.\u003c/p\u003e","description":"","filename":"10.png","url":"https://assets-eu.researchsquare.com/files/rs-7917997/v1/b8d2686b2a497dafb7e4132b.png"},{"id":102234174,"identity":"9cfbd712-bf37-4921-9f4d-10173c48bacd","added_by":"auto","created_at":"2026-02-09 16:07:11","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":9024744,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-7917997/v1/ae93ddfb-54aa-4168-9394-6c4fa4ba62b4.pdf"},{"id":95905585,"identity":"57da3bc7-74da-404a-abc5-480d559d2760","added_by":"auto","created_at":"2025-11-14 09:23:11","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"supplement","size":1718149,"visible":true,"origin":"","legend":"","description":"","filename":"SupplementaryInformation.pdf","url":"https://assets-eu.researchsquare.com/files/rs-7917997/v1/1ab1a5711e41335efac8a20a.pdf"}],"financialInterests":"No competing interests reported.","formattedTitle":"Cold-adapted carboxylesterases from Alcanivoracaceae active with a wide range of synthetic polyesters","fulltext":[{"header":"Key points","content":"\u003cp\u003e\u003cul\u003e\u003cli\u003e\u003cp\u003eMembers of \u003cem\u003eAlcanivoracaceae\u003c/em\u003e are a rich resource of polyester-degrading enzymes\u003c/p\u003e\u003c/li\u003e\u003cli\u003e\u003cp\u003eAll selected and analysed Family V esterases exhibited high activities and stabilities at low temperatures and solvent tolerance\u003c/p\u003e\u003c/li\u003e\u003cli\u003e\u003cp\u003eCharacterised enzymes were active with a broad range of polyesters\u003c/p\u003e\u003c/li\u003e\u003c/ul\u003e\u003c/p\u003e"},{"header":"Introduction","content":"\u003cp\u003ePlastic pollution is an ever-escalating issue with detrimental effects on both the environment and human health. Over the years, plastics have become an indispensable part of daily life, serving critical roles in packaging, transportation and healthcare. However, their widespread use has also led to a significant waste accumulation problem, making plastic pollution one of the greatest environmental challenges of the 21st century. Despite growing consumer awareness, the production of single-use plastics continues to escalate (Geyer et al., \u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e2017\u003c/span\u003e; Stubbins et al., \u003cspan citationid=\"CR70\" class=\"CitationRef\"\u003e2021\u003c/span\u003e). Among various strategies for addressing plastic waste, biological degradation, particularly enzymatic closed-loop recycling has emerged as a promising solution (Abou-Zeid et al., \u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e2004\u003c/span\u003e; Danso et al., \u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e2019\u003c/span\u003e; Viljakainen and Hug, 2021). For example, the first enzymatic polyethylene terephthalate (PET) degradation study was reported with an enzyme derived from \u003cem\u003eThermobifida fusca\u003c/em\u003e (Müller et al., \u003cspan citationid=\"CR52\" class=\"CitationRef\"\u003e2005\u003c/span\u003e). Over the past two decades, significant advancements in protein and process engineering have led to substantial improvements in the initially low PET degradation activity of \u003cem\u003eT. fusca\u003c/em\u003e (Barth et al., \u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e2015\u003c/span\u003e; Müller et al., \u003cspan citationid=\"CR52\" class=\"CitationRef\"\u003e2005\u003c/span\u003e; Roth et al., \u003cspan citationid=\"CR60\" class=\"CitationRef\"\u003e2014\u003c/span\u003e). With the advancement of metagenomic approaches, more active enzymes, such as leaf-branch-compost cutinase LCC, have been identified (Sulaiman et al., \u003cspan citationid=\"CR71\" class=\"CitationRef\"\u003e2012\u003c/span\u003e). Furthermore, protein engineering has enabled the development of mutant LCC variants (e.g., LCC\u003csub\u003eICCG\u003c/sub\u003e, LCC\u003csub\u003eWCCG\u003c/sub\u003e) with enhanced and accelerated depolymerisation efficiency (Makryniotis et al., \u003cspan citationid=\"CR47\" class=\"CitationRef\"\u003e2023\u003c/span\u003e; Tournier et al., \u003cspan citationid=\"CR73\" class=\"CitationRef\"\u003e2020\u003c/span\u003e). Subsequently, a novel bacterium, \u003cem\u003eIdeonella sakaiensis\u003c/em\u003e 201-F6, was isolated from plastic-contaminated sediment samples and found to exhibit both IsPETase and monomer-hydrolysing (MHETase) activities (Kan et al., \u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e2021\u003c/span\u003e; Yoshida et al., \u003cspan citationid=\"CR88\" class=\"CitationRef\"\u003e2016\u003c/span\u003e). IsPETase initially identified and characterised in that study, has served as the foundation for numerous structural and protein engineering studies aimed at further enhancing its PET depolymerisation activity (Dai et al., \u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e2021\u003c/span\u003e; Feng et al., \u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e2021\u003c/span\u003e; Joo et al., \u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e2018\u003c/span\u003e; Haugwitz et al., \u003cspan citationid=\"CR33\" class=\"CitationRef\"\u003e2022\u003c/span\u003e; Wei et al., \u003cspan citationid=\"CR78\" class=\"CitationRef\"\u003e2022\u003c/span\u003e).\u003c/p\u003e\u003cp\u003eOngoing research efforts are focused on modifying the active sites of the wild-type enzymes to improve their catalytic efficiency, as well as engineering enzymes with enhanced thermostabilities trough rational design and directed evolution (Bell et al., \u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e2022\u003c/span\u003e; Shi et al., \u003cspan citationid=\"CR66\" class=\"CitationRef\"\u003e2023\u003c/span\u003e; Schmidt et al., \u003cspan citationid=\"CR63\" class=\"CitationRef\"\u003e2019\u003c/span\u003e; Williams et al., \u003cspan citationid=\"CR81\" class=\"CitationRef\"\u003e2023\u003c/span\u003e). These advancements hold great potential for the development of efficient biocatalysts capable of supporting sustainable plastic waste management and recycling.\u003c/p\u003e\u003cp\u003eThe ester bonds in polyesters like PET are chemically different from the carbon-carbon bonds in polymers like polyethylene (PE) and polystyrene (PS) (Mohanan et al., \u003cspan citationid=\"CR51\" class=\"CitationRef\"\u003e2020\u003c/span\u003e). The latter olefin-based polymers require oxygenation, hydroxylation, or monooxygenation of C-C and C-H bonds to enable their degradation in same pathways as for aliphatic hydrocarbons degradations (Bornscheuer et al., \u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e2024\u003c/span\u003e). In contrast, hydrocarbon contamination from petroleum oil production, processing and transportation significantly contributes to environmental pollution. Like plastic waste, hydrocarbons persist in ecosystems, exacerbating the long-term environmental burden due to their resistance to biodegradation and accumulation in both terrestrial and marine habitats (Brzeszcz and Kaszycki, \u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e2018\u003c/span\u003e; Park and Park, \u003cspan citationid=\"CR55\" class=\"CitationRef\"\u003e2018\u003c/span\u003e). The degradation of these compounds using bio-based approaches has gained considerable attention, particularly the application of naturally occurring hydrocarbon-degrading marine microorganisms and their enzymes (Beilen et al., \u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e1994\u003c/span\u003e; Beilen and Funhoff, \u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e2005\u003c/span\u003e, \u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e2007\u003c/span\u003e; Haines and Alexander, \u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e1974\u003c/span\u003e; Wei and Zimmermann, \u003cspan citationid=\"CR79\" class=\"CitationRef\"\u003e2017\u003c/span\u003e). Hydrocarbon degrading microorganisms, which are taxonomically diverse, capable to utilise hydrocarbons as their sole carbon and energy source. These include truly marine bacteria, such as \u003cem\u003eAlcanivorax, Oleiphilus, Oleispira, Thalassolituus, Cycloclasticus, Marinobacter\u003c/em\u003e and others (Golyshin et al., \u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e2003\u003c/span\u003e; Gregson et al., \u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e2018\u003c/span\u003e, \u003cspan citationid=\"CR28\" class=\"CitationRef\"\u003e2020\u003c/span\u003e; Schneiker et al., \u003cspan citationid=\"CR64\" class=\"CitationRef\"\u003e2006\u003c/span\u003e; Yakimov et al., \u003cspan citationid=\"CR84\" class=\"CitationRef\"\u003e2004\u003c/span\u003e;), as well as fungi \u003cem\u003eAspergillus\u003c/em\u003e and \u003cem\u003ePenicillium\u003c/em\u003e(Zhou et al., \u003cspan citationid=\"CR93\" class=\"CitationRef\"\u003e2014\u003c/span\u003e; Sowmya et al., \u003cspan citationid=\"CR68\" class=\"CitationRef\"\u003e2015\u003c/span\u003e) that harbour enzymes for full mineralisation of alkanes. Key enzymes involved in alkane degradation include alkane hydroxylases and monooxygenases (e.g. AlmA, AlkB, P450), which oxidise alkanes to alcohols, followed by their conversion to aldehydes, fatty acids, acyl-CoA and their further beta-oxidation (Guo et al., \u003cspan citationid=\"CR29\" class=\"CitationRef\"\u003e2023\u003c/span\u003e; Kube et al., \u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e2013\u003c/span\u003e; Sabirova et al., \u003cspan citationid=\"CR61\" class=\"CitationRef\"\u003e2006\u003c/span\u003e; Wang and Shao, 2014). These metabolic pathways are essential for the complete mineralisation of hydrocarbons and play a key role in the natural bioremediation of oil spills (De Santi et al., \u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e2014\u003c/span\u003e; Lai et al., \u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e2011\u003c/span\u003e).\u003c/p\u003e\u003cp\u003eInterestingly, many hydrocarbon-degrading microorganisms identified in the marine oil-contaminated environments are also found in the “plastisphere”, the surfaces of plastics and microplastics providing a biological niche where diverse microbial communities establish biofilms (Emadian et al., \u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e2017\u003c/span\u003e; Wei and Zimmermann, \u003cspan citationid=\"CR79\" class=\"CitationRef\"\u003e2017\u003c/span\u003e; Zettler et al., \u003cspan citationid=\"CR91\" class=\"CitationRef\"\u003e2013\u003c/span\u003e). Both oil spill and plastisphere-associated microbiomes often possess similar metabolic capabilities, such as alkane hydroxylases and esterases, which enable them to degrade complex hydrophobic compounds (Emadian et al., \u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e2017\u003c/span\u003e; Wei and Zimmermann, \u003cspan citationid=\"CR79\" class=\"CitationRef\"\u003e2017\u003c/span\u003e). This functional overlap suggests that, despite originating from different sources, these environments create comparable ecological niches that select for organisms with similar biodegradation potential (Wilkes and Aristilde, \u003cspan citationid=\"CR80\" class=\"CitationRef\"\u003e2017\u003c/span\u003e; Sabirova et al., \u003cspan citationid=\"CR61\" class=\"CitationRef\"\u003e2006\u003c/span\u003e). For instance, \u003cem\u003ePseudomonas\u003c/em\u003e species are known for their metabolic versatility, producing cutinases and other esterases that facilitate the degradation of synthetic polyesters like PET (Wilkes and Aristilde, \u003cspan citationid=\"CR80\" class=\"CitationRef\"\u003e2017\u003c/span\u003e; Zhou et al., \u003cspan citationid=\"CR94\" class=\"CitationRef\"\u003e2024\u003c/span\u003e). Similarly, \u003cem\u003eRhodococcus\u003c/em\u003e species produce esterases with broad substrate specificity, allowing them to degrade various polyester-based plastics (De Santi et al., \u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e2014\u003c/span\u003e). These enzymes exhibit significant potential for bioremediation and recycling of plastic waste, offering an environmentally friendly alternative to conventional chemical processes.\u003c/p\u003e\u003cp\u003ePolyesterases, such as carboxylesterases, cutinases, lipases, and specific PETases, are particularly effective in degrading synthetic polyesters. Cutinases, found in both bacteria and fungi, hydrolyse the ester bonds in cutin and synthetic polyesters, leading to the formation of monomers such as terephthalic acid and ethylene glycol. PETases, which have gained considerable attention, are specialized enzymes capable of breaking down PET into its constituent monomers under mild conditions (Tournier et al., \u003cspan citationid=\"CR73\" class=\"CitationRef\"\u003e2020\u003c/span\u003e). These enzymes, originally identified in \u003cem\u003eIdeonella sakaiensis\u003c/em\u003e, have been found in various alkane-degrading microorganisms, highlighting the potential for dual functionality in hydrocarbon and polyester degradation (Yoshida et al., \u003cspan citationid=\"CR88\" class=\"CitationRef\"\u003e2016\u003c/span\u003e).\u003c/p\u003e\u003cp\u003eSome bacteria from the genera \u003cem\u003eAlcanivorax, Oleiphilus, Oleispira, Marinobacter\u003c/em\u003e and \u003cem\u003eThalassolituus\u003c/em\u003e, utilise hydrocarbons as their preferred source of carbon and energy (Staley, \u003cspan citationid=\"CR69\" class=\"CitationRef\"\u003e2010\u003c/span\u003e; Yakimov et al., \u003cspan citationid=\"CR84\" class=\"CitationRef\"\u003e2004\u003c/span\u003e, \u003cspan citationid=\"CR87\" class=\"CitationRef\"\u003e2007\u003c/span\u003e; Golyshin et al., \u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e2002\u003c/span\u003e). Among those organisms, which hold significant potential for both natural attenuation of oil hydrocarbons \u003cem\u003ein situ\u003c/em\u003e, and degradation of olefins and polyesters, the genus \u003cem\u003eAlcanivorax\u003c/em\u003e, a member of the family \u003cem\u003eAlcanivoracaceae\u003c/em\u003e within the class \u003cem\u003eGammaproteobacteria\u003c/em\u003e, has attracted particular attention for over 25 years (Golyshin et al., \u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e2003\u003c/span\u003e; Yakimov et al., \u003cspan citationid=\"CR86\" class=\"CitationRef\"\u003e1998\u003c/span\u003e, \u003cspan citationid=\"CR83\" class=\"CitationRef\"\u003e2022\u003c/span\u003e). The genus \u003cem\u003eAlcanivorax\u003c/em\u003e (family \u003cem\u003eAlcanivoracaceae\u003c/em\u003e) was recently divided into three genera, including two new (Rai et al., 2023). Several species within this family exhibit a strong preference for aliphatic hydrocarbons, both linear and branched (Yakimov et al., \u003cspan citationid=\"CR83\" class=\"CitationRef\"\u003e2022\u003c/span\u003e). Some \u003cem\u003eAlcanivoracaceae\u003c/em\u003e species have also been reported to degrade polycyclic aromatic hydrocarbons (PAHs) such as naphthalene and pyrene, e.g. \u003cem\u003eAlloalcanivorax xenomutans\u003c/em\u003e SRM1 (previously known as \u003cem\u003eAlcanivorax xenomutans\u003c/em\u003e) (Dell’Anno et al., \u003cspan citationid=\"CR17\" class=\"CitationRef\"\u003e2023\u003c/span\u003e) and aromatic hydrocarbons, e.g. xylene (\u003cem\u003eA. xenomutans\u003c/em\u003e JC109) (Rahul et al., \u003cspan citationid=\"CR37\" class=\"CitationRef\"\u003e2014\u003c/span\u003e).\u003c/p\u003e\u003cp\u003eTypically, \u003cem\u003eAlcanivoracaceae\u003c/em\u003e species occur at low or undetectable levels in unpolluted environments; however, their growth is markedly stimulated by the presence of hydrocarbons (Yakimov et al., \u003cspan citationid=\"CR87\" class=\"CitationRef\"\u003e2007\u003c/span\u003e; \u003cspan citationid=\"CR83\" class=\"CitationRef\"\u003e2022\u003c/span\u003e). Many characterised strains do not assimilate sugars or amino acids, but are capable of metabolising fatty acids, alcohols, and aliphatic hydrocarbons (Lai et al., \u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e2011\u003c/span\u003e). Given their ecological lifestyle, these organisms, and their enzymes, are expected to exhibit tolerance to organic solvents (Bollinger et al., \u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e2020\u003c/span\u003e). \u003cem\u003eAlcanivoracaceae\u003c/em\u003e species along with other hydrocarbon-degrading taxa, are frequently identified as colonisers of plastic surface in marine environment and are considered as potential plastic degraders (Denaro et al., \u003cspan citationid=\"CR18\" class=\"CitationRef\"\u003e2020\u003c/span\u003e; Delacuvellerie et al., \u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e2019\u003c/span\u003e; Popovic et al., \u003cspan citationid=\"CR58\" class=\"CitationRef\"\u003e2017\u003c/span\u003e; Tulloch et al., \u003cspan citationid=\"CR74\" class=\"CitationRef\"\u003e2024\u003c/span\u003e; Yakimov et al., \u003cspan citationid=\"CR85\" class=\"CitationRef\"\u003e2019\u003c/span\u003e; Zadjelovic et al., \u003cspan citationid=\"CR90\" class=\"CitationRef\"\u003e2022\u003c/span\u003e;).\u003c/p\u003e\u003cp\u003eIn terms of enzymatic capability, carboxylesterases from the type strain of the family \u003cem\u003eAlcanivorax borkumensis\u003c/em\u003e SK2\u003csup\u003eT\u003c/sup\u003e have demonstrated broad substrate profiles. These enzymes are active not only against model substrates (i.e. \u003cem\u003ep\u003c/em\u003eNP-esters of fatty acids with aliphatic chain lengths between C2 and C16 ), but also against synthetic polyesters such as PLLA, PDLLA, PCL, 3PET (Hajighasemi et al., \u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e2018\u003c/span\u003e, \u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e2016\u003c/span\u003e; Tchigvintsev et al., \u003cspan citationid=\"CR72\" class=\"CitationRef\"\u003e2015\u003c/span\u003e), indicating the broader biotechnological potential of this organism beyond its established role in the natural attenuation of oil spill pollution. Notably, such substrate promiscuity is common among carboxylesterases and has been extensively studied (Martínez-Martínez et al., \u003cspan citationid=\"CR48\" class=\"CitationRef\"\u003e2018\u003c/span\u003e; Ma et al., \u003cspan citationid=\"CR46\" class=\"CitationRef\"\u003e2025\u003c/span\u003e). Another species within the family, \u003cem\u003eAlloalcanivorax dieselolei\u003c/em\u003e (previously known as \u003cem\u003eAlcanivorax dieselolei)\u003c/em\u003e, has been reported to encode a promiscuous, solvent-tolerant esterase (Zhang et al., \u003cspan citationid=\"CR92\" class=\"CitationRef\"\u003e2014\u003c/span\u003e). Recent investigations of a strain isolated from plastic marine debris, \u003cem\u003eAlcanivora\u003c/em\u003ex sp. 24 (recently renamed to \u003cem\u003eAlloalcanivorax\u003c/em\u003e sp. 24), showed that its hydrolase ALC24_4107 exhibited a strong activity \u003cem\u003evs\u003c/em\u003e aliphatic polyester, such as PHB, PHBV, PES, PBS and PCL (Zadjelovic et al., \u003cspan citationid=\"CR89\" class=\"CitationRef\"\u003e2020\u003c/span\u003e). Of note, this very strain was also demonstrated to degrade the weathered low-density polyethylene (LDPE) by recruiting an array of redox enzymes (alkane monooxygenases, P450, laccases) and reactive oxygen species (Zadjelovic et al., \u003cspan citationid=\"CR90\" class=\"CitationRef\"\u003e2022\u003c/span\u003e). As reported elsewhere, a promiscuous hydrolase from metagenomic fragment of \u003cem\u003eThalassolituus oleivorans\u003c/em\u003e (another renowned marine oil-degrader) showed a hybrid ester hydrolase and haloacid- dehalogenase activity (Beloqui et al., \u003cspan citationid=\"CR8\" class=\"CitationRef\"\u003e2010\u003c/span\u003e). As exemplified in the above and in other studies, mining metagenomes of petroleum-enriched microbial communities using activity-based screens is highly regarded as a productive approach to attribute activities to yet uncharacterised enzymes and new biochemical activities from hydrocarbon-based marine environments (Ferrer et al., \u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e2005\u003c/span\u003e; Popovic et al., \u003cspan citationid=\"CR59\" class=\"CitationRef\"\u003e2015\u003c/span\u003e, \u003cspan citationid=\"CR58\" class=\"CitationRef\"\u003e2017\u003c/span\u003e).\u003c/p\u003e\u003cp\u003eThis study aimed to identify novel polyesterases from \u003cem\u003eAlcanivoracaceae\u003c/em\u003e species with the potential to degrade a wide range of synthetic polyesters, such as 3PET, PLLA, PDLA, PDLLA, PCL, PBA, PBS, and PC. While several esterases from \u003cem\u003eAlcanivoracaceae\u003c/em\u003e species have been previously characterised, this research extends the investigation to further predicted polyesterases across a broader selection of strains from this family including \u003cem\u003eAlloalcanivorax\u003c/em\u003e sp. 24, \u003cem\u003eAlloalcanivorax gelatiniphagus\u003c/em\u003e and \u003cem\u003eIsoalcanivorax pacificus\u003c/em\u003e (previously known as \u003cem\u003eAlcanivorax gelatiniphagus\u003c/em\u003e and \u003cem\u003eAlcanivorax pacificus\u003c/em\u003e, respectively). The genomes of these organisms encode a variety of putative α/β hydrolases and carboxylesterases, many of which remain uncharacterised and present exciting opportunities for further exploration. Notably, \u003cem\u003eAlcanivoracaceae\u003c/em\u003e species are naturally adapted to marine environments, making them particularly well-suited for applications in low temperatures conditions and energy-efficient industrial processes. Additionally, polyesterases have demonstrated considerable promise in various sectors, such as waste management, bioremediation, and the development of biodegradable plastics, further emphasising the relevance of this research for sustainable and environmentally industrial practices.\u003c/p\u003e"},{"header":"Materials and Methods","content":"\u003cp\u003e\u003cstrong\u003eChemicals\u003c/strong\u003e\u003c/p\u003e\u003cp\u003eAll chemicals and substrates used in this study were of analytical grade. Substrates chromogenic \u003cem\u003ep\u003c/em\u003e-nitrophenyl (\u003cem\u003ep\u003c/em\u003eNP) esters (C\u003csub\u003e2\u003c/sub\u003e-C\u003csub\u003e16\u003c/sub\u003e) were purchased from Sigma-Aldrich/Merck (Gillingham, UK) and Tokyo Chemical Industry UK Ltd. (TCI, Oxford, UK). 6-hydroxycaproic acid (6-HHA), lactic acid (LA), succinic acid (SA), adipic acid (AA), terephthalic acid (TA) and bisphenol-A, mono 2-hydroxyethyl terephthalic acid (MHET), and bis(2-hydroxyethyl) terephthalic acid (BHET), Tween-20, dichloromethane (DCM) and 1,1,1,3,3,3-hexafluoro-2-propanol (HFP) were purchased from Sigma-Aldrich/Merck (Gillingham, UK). Polyester substrates, poly-D, L-lactide (PDLLA, molecular weight M\u003csub\u003eW\u003c/sub\u003e 10,000–18,000), polycaprolactone (PCL14, average M\u003csub\u003eW\u003c/sub\u003e ~14,000) (PCL2, M\u003csub\u003eW\u003c/sub\u003e ~2,000), and amorphous polyethylene terephthalate (aPET, thickness 0.25 mm, coil width 600 mm), polybutylene succinate (PBS, M\u003csub\u003eW\u003c/sub\u003e 45,500), poly-1,4-butylene adipate) (PBA, M\u003csub\u003eW\u003c/sub\u003e 12,000), poly(1,4-butylene) terephthalate (PBT, M\u003csub\u003eW\u003c/sub\u003e 38,000), poly-L-lactic acid (PLLA, M\u003csub\u003eW\u003c/sub\u003e 15,000–25,000), poly-D-lactic acid (PDLA, M\u003csub\u003eW\u003c/sub\u003e 10,000–15,000), polycarbonate (PC, 3 mm granules) were purchased from Sigma-Aldrich/Merck (Gillingham, UK). The PET model substrate, bis(benzoyloxyethyl) terephthalate (3PET, M\u003csub\u003eW\u003c/sub\u003e 462.4), was synthesized by CanSyn (Toronto, Canada). Impranil® DLN was kindly donated by WhitChem Ltd, UK (Azelis, \u003cspan class=\"ExternalRef\"\u003e\u003cspan class=\"RefSource\"\u003ewww.whitchem.co.uk\u003c/span\u003e\u003c/span\u003e). Isopropyl β-D-1-thiogalactopyranoside (IPTG), ampicillin, 2-(cyclohexylamino)ethanesulfonic acid (CHES), sodium chloride (NaCl), tris(hydroxymethyl)aminomethane hydrochloride (Tris-HCl), methanol, sulfuric acid, orthophosphoric acid, and Luria-Bertani (LB) broth were also purchased from Sigma-Aldrich/Merck (Gillingham, UK).\u003c/p\u003e\u003cp\u003e\u003cstrong\u003eGene cloning and protein purification\u003c/strong\u003e\u003c/p\u003e\u003cp\u003eThe coding sequences of selected hydrolase genes were commercially synthesized (without signal peptides, with addition of N-terminal hexahistidine tags) and cloned into a modified p15TVL (TWIST BIOSCIENCE, South San Francisco, USA). All plasmids were transformed into the \u003cem\u003eE. coli\u003c/em\u003e Lobstr BL21(DE3) cells (Kerafast, Boston, USA). \u003cem\u003eE. coli\u003c/em\u003e cultures were grown aerobically in 2.5 L baffled Erlenmeyer flasks with 1 L Luria-Bertani medium supplemented with 4 g/L glycerol and 100 µg/ml ampicillin at 37°C, in a shaking incubator at 200 rpm, to the optical density (OD\u003csub\u003e600\u003c/sub\u003e) 0.6–0.8, then spiked with 0.4 mM IPTG, transferred to the 16 ºC and incubated at that temperature in a shaker for further 16 h. The biomass was harvested by centrifugation at 4,000 rpm, 10 min (Avanti J26 rotor JLA8.1, Beckman Coulter Life Sciences, Indianapolis, USA) and disrupted in ice bath by sonication (Q-sonica, Newtown, USA) for 10 min at 70% intensity, in 4 s pulses with 5 s cooling time. Recombinant proteins were purified to near homogeneity (\u0026gt; 95%) using nickel-chelate affinity chromatography on Ni-NTA Superflow resin (QIAGEN, Hilden, Germany) as described previously (Distaso et al., \u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e2023\u003c/span\u003e). Purity and protein size of purified enzymes were assessed using denaturing 10% polyacrylamide gel (BioRad Laboratories, Hercules, USA) electrophoresis, whereas protein concentration was measured by Bradford assay (Bio-Rad Laboratories, Hercules, USA).\u003c/p\u003e\u003cp\u003e\u003cstrong\u003eCarboxylesterase assays with soluble chromogenic substrates\u003c/strong\u003e\u003c/p\u003e\u003cp\u003eThe chromogenic \u003cem\u003ep\u003c/em\u003e-nitrophenyl (\u003cem\u003ep\u003c/em\u003eNP) esters of fatty acids with different acyl chain lengths: \u003cem\u003ep\u003c/em\u003eNP-acetate (C2), \u003cem\u003ep\u003c/em\u003eNP-butyrate (C4), \u003cem\u003ep\u003c/em\u003eNP-hexanoate (C6), \u003cem\u003ep\u003c/em\u003eNP-octanoate (C8), \u003cem\u003ep\u003c/em\u003eeps-decanoate (C10), \u003cem\u003ep\u003c/em\u003eNP-dodecanoate (C12), \u003cem\u003ep\u003c/em\u003eNP-myristate (C14), \u003cem\u003ep\u003c/em\u003eNP-palmitate (C16) were used to test carboxylesterase activity of purified proteins. The activity was screened in 96 well plates spectrophotometrically using SpectraMax M3 (Molecular Devices, San Jose, USA). All assays were conducted in triplicates at indicated temperatures in 96-well plates with reaction mixtures (200 µl) containing 50 mM CHES (pH 9.0) buffer (or as indicated), 1 mM substrate (and 0.1-1 µg of enzyme. The reaction mixtures were incubated for 20 min at 30 ºC, and the activity was calculated based on the absorbance of \u003cem\u003ep\u003c/em\u003e-nitrophenol at 410 nm (ε = 17.8 mM\u003csup\u003e− 1\u003c/sup\u003e cm\u003csup\u003e− 1\u003c/sup\u003e).\u003c/p\u003e\u003cp\u003e\u003cstrong\u003eEnzyme reaction conditions and stability determination\u003c/strong\u003e\u003c/p\u003e\u003cp\u003eThe effect of pH on carboxylesterase activity of purified proteins (pH profile) was determined using the universal Britton-Robinson buffer system (50 mM acetic acid, 50 mM phosphoric acid, 50 mM boric acid, pH range 4.0 to 10.5). Temperature optimum (temperature profile) for carboxylesterase activity of selected enzymes was measured at temperatures from 30 ºC to 90 ºC using 1 mM \u003cem\u003ep\u003c/em\u003eNP-octanoate substrate and 0.1-1 µg of protein in 50 mM CHES buffer (pH 9.0). 200 µl assay reactions were incubated in 96 well plate in the Thermomixer (Eppendorf, Hamburg, Germany) at 500 rpm. For the analysis of their thermal stabilities, selected proteins (1 mg/ml) were incubated for 2 h for thermostability and 5 h for cold stability (or as indicated) at different temperatures (from 5 ºC to 90 ºC) in the PCR thin wall tubes in T100 Thermal Cycler (Bio-Rad Laboratories, Hercules, USA), and the residual carboxylesterase activity was measured with 1 mM \u003cem\u003ep\u003c/em\u003eNP-octanoate at 30 ºC as described in the carboxylesterase assay methods section.\u003c/p\u003e\u003cp\u003eThe effect of NaCl and Tween20 on carboxylesterase activity of selected proteins was analysed using 1.0 mM \u003cem\u003ep\u003c/em\u003eNP-octanoate and 0.1-1 µg of enzyme. In the reaction mixture of 200 µl containing NaCl at concentrations between 0.1 and 2.0 M and Tween-20 in the range of 0.1- 3.0% in 50 mM CHES, pH 9.0. Reaction mixtures were incubated for 20 min at 30 ºC, and the activity was measured spectrophotometrically at 410 nm.\u003c/p\u003e\u003cp\u003eThe effect of organic solvents on carboxylesterase activity was analysed in the conditions described in carboxylesterase assays with soluble chromogenic substrates section, but supplied with various concentration of dichloromethane, dimethylsulfoxide, ethanol, and 1,1,1,3,3,3-hexafluoro-2-propanol in the range between 0 and 50%, in 50 mM CHES, pH 9.0.\u003c/p\u003e\u003cp\u003e\u003cstrong\u003eAnalysis of temperature-dependent protein denaturation\u003c/strong\u003e\u003c/p\u003e\u003cp\u003eThe melting temperature of selected enzymes was measured by differential scanning fluorimetry (DSF) on a QuantStudio 6 Flex system (Applied Biosystems, Thermo Fisher Scientific, Waltham, USA) using SYPRO Orange (Invitrogen, Carlsbad, USA) as a binding dye. Experiments were conducted in triplicate in 30 µl reaction volumes with 10 µg of protein in 50 mM CHES buffer (pH 9.0) and 25X SYPRO Orange in sealed optically clear 96-well plates (Applied Biosystems, Thermo Fisher Scientific, Waltham, USA) with the system set on the ROX as the reference wavelength using the 450/490 nm excitation and 560/580 nm emission filters. The temperature was increased from 25 to 95 ºC with an increment of 1 ºC s\u003csup\u003e− 1\u003c/sup\u003e. \u003cem\u003eT\u003c/em\u003e\u003csub\u003em\u003c/sub\u003e values for selected proteins were determined by non-linear fitting of Sigmoidal-Boltzmann equation using the GraphPad Prism software (version 5.0).\u003c/p\u003e\u003cp\u003e\u003c/p\u003e\u003cp\u003e\u003cstrong\u003ePreparation of polyester substrates and polyesterase assays\u003c/strong\u003e\u003c/p\u003e\u003cp\u003ePolyester substrates used in this study (PDLLA, PLLA, PDLA, PCL14, PCL2, 3PET, PBS, PBA, PBT, aPET) were prepared in 50 mM Tris-HCl (pH 8.0), as described previously (Distaso et al., \u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e2023\u003c/span\u003e). For agarose-based screens, 0.5% polyester emulsions were diluted with three volumes of CHES buffer (pH 9.0) in 2% (w/vol) molten agarose, poured and solidified in the round 90 mm Petri dishes to make opaque gel with the final concentration of polyesters 0.125% (w/vol). Impranil DLN was used only for agarose gel plates preparation. For this, 1 ml Impranil® DLN dissolved in 100 ml 50 mM CHES (pH 9.0), with the final concentration of Impranil 1% in 2% (w/vol) agarose. 50 µg enzyme aliquots were loaded into cylindrical boreholes cored in the agarose using cut 1 ml micropipette tips. Plates were placed into plastic bags to prevent evaporation, incubated at 30 ºC and monitored for 2–3 days. The formation of a clear zone around the wells was considered as indication for the presence of polyester-degrading activity.\u003c/p\u003e\u003cp\u003eFor polyesterase assays in solution, the 200 µl reaction mixtures with 50 µg of purified protein and 0.125% of emulsified polyester in 50 mM CHES buffer (pH 9.0) were incubated for 12 h at 30 ºC in a shaker at 500 rpm. The reactions were spun down at 14,000 g for 10 min in a minicentrifuge, and supernatant was filtered through 10 kDa spin filters (14,000 g, 15 min, Eppendorf 5424 centrifuge, Eppendorf AG, Hamburg, Germany). The presence of polyester degradation products in filtrates was analysed using HPLC Prominence-I LC-2030C 3D Plus equipped with UV-VIS detector (Shimadzu, Kyoto, Japan). For aPET, 3PET, PBT the formation of terephthalic acid (TA), mono-2-hydroxyethyl terephthalic acid (MHET), bis-2-hydroxyethyl terephthalic acid (BHET) was analysed. For PC depolymerisation the bisphenol-A (BPA), was analysed. For product analysis the reverse-phase hydrophobic chromatography on a Shimadzu C\u003csub\u003e18\u003c/sub\u003e Shim-pack column (150x4.6 mm, 5 µm particle size) (Shimadzu Corporation, Kyoto, Japan), 40 ºC (injection volume 10 µl, detection at 240 nm) was used. For products separation we used the mobile phase of Solvent A (0.1% (vol/vol) of orthophosphoric acid (H\u003csub\u003e3\u003c/sub\u003ePO\u003csub\u003e4\u003c/sub\u003e) in the HPLC grade water and Solvent B (100% methanol). At flow rate of 0.7 ml/min, 40 ºC, the gradient was: 0–2 min, 25% Solvent B; 2–18 min, linear gradient to 55% of Solvent B; 18–22 min, linear gradient to 25% of Solvent B. For PDLA, PLLA, PDLLA depolymerisation efficiency, lactic acid was quantified; for PCL14, PCL2, 6-hydroxyhexanoic acid; for PBS depolymerisation, succinic acid; for PBAT, adipic acid were quantified using ion-moderated partition chromatography on a Prominence-I LC-2030C 3D Plus HPLC system equipped with an Aminex HPX-87-H column (300x7.8 mm, 9 µm particle size, conditioned at 50 ºC) (Shimadzu Corporation, Kyoto, Japan) and a UV detector (LC-2030C_3Dplus, Shimadzu Corporation, Kyoto, Japan). For analysis, 0.05 N sulfuric acid (H\u003csub\u003e2\u003c/sub\u003eSO\u003csub\u003e4\u003c/sub\u003e) in HPLC grade water was used as solvent at flow rate of 0.6 ml/min, detection at 210 nm. Polyester degradation products were quantified based on their calibration curves generated using commercially available standards (TA, MHET, BHET, LA, 6-HHA, AA, SA, BPA).\u003c/p\u003e\u003cp\u003e\u003cstrong\u003eBioinformatic and structural analyses\u003c/strong\u003e\u003c/p\u003e\u003cp\u003eMultiple sequence alignments of selected carboxylesterases were performed using the MAFFT online service, automated regime (Katoh et al., \u003cspan citationid=\"CR40\" class=\"CitationRef\"\u003e2019\u003c/span\u003e). Alignment was visualised and modified using the online software STRAP (Gille et al., \u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e2014\u003c/span\u003e). For phylogenetic analysis, 5 \u003cem\u003eAlcanivoracaceae\u003c/em\u003e strains genomes were screened for IPR029058 (AB hydrolase) signature containing sequences, and the 297 sequences were retrieved, and aligned in Geneious Prime using global alignment with free end gaps, Cost matrix Blosum 62, and Fast tree was used for phylogeny analysis. Structural analysis was performed in ChimeraX (Meng et al., \u003cspan citationid=\"CR50\" class=\"CitationRef\"\u003e2023\u003c/span\u003e).\u003c/p\u003e"},{"header":"Results","content":"\u003cp\u003e\u003cstrong\u003eAnalysis of Alcanivoracaceae genomes for potential carboxylesterases and novel polyesterases\u003c/strong\u003e\u003c/p\u003e\u003cp\u003eIn this study, we used five reference \u003cem\u003eAlcanivoracaceae\u003c/em\u003e genomes (\u003cem\u003eAlcanivorax borkumensis\u003c/em\u003e SK2\u003csup\u003eT\u003c/sup\u003e, \u003cem\u003eA. hongdengensis\u003c/em\u003e A-11-3\u003csup\u003eT\u003c/sup\u003e, \u003cem\u003eA. sediminis\u003c/em\u003e PA15-N-34\u003csup\u003eT\u003c/sup\u003e, \u003cem\u003eA. profundi\u003c/em\u003e MTE017\u003csup\u003eT\u003c/sup\u003e, and \u003cem\u003eA. nanhaiticus\u003c/em\u003e 19-m-6\u003csup\u003eT\u003c/sup\u003e), for screening of IPR029058 (ɑ/β hydrolase) signature containing sequences. 297 sequences were retrieved from GenBank, 5 sequences manually selected from \u003cem\u003eIsoalcanivorax pacificus\u003c/em\u003e previously known as \u003cem\u003eAlcanivorax pacificus\u003c/em\u003e (APA_2, APA_3, APA_4, APA_5, APA_6), 6 sequences (ABO_0116, ABO_1197, ABO_1251, ABO_1483, ABO_1895, ABO_2249) from \u003cem\u003eA. borkumensis\u003c/em\u003e, 5 sequences (ALC24_3989, ALC24_1328, ALC24_1162, ALC24_2069, ALC24_4107) from \u003cem\u003eAlloalcanivorax\u003c/em\u003e sp. 24 (SI Table\u0026nbsp;1) and added for multiple sequence analysis (SI Fig.\u0026nbsp;1). Selected esterases from \u003cem\u003eAlcanivorax borkumensis\u003c/em\u003e (ABO_0116, ABO_1197, ABO_1251, ABO_1483, ABO_1895, and ABO_2249) were partially characterised in previous studies (Hajighasemi et al., \u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e2016\u003c/span\u003e, \u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e2018\u003c/span\u003e; Tchigvintsev et al., \u003cspan citationid=\"CR72\" class=\"CitationRef\"\u003e2015\u003c/span\u003e), which is highlighted in the phylogenetic tree (Fig.\u0026nbsp;1).\u003c/p\u003e\u003cp\u003eSequence analysis of these proteins revealed that ABO_1197, ABO_1895, APA_6 and APA_5 have signal peptides (S1 Table\u0026nbsp;1) suggesting that majority of \u003cem\u003eAlcanivoracaceae\u003c/em\u003e esterases are involved in the intracellular metabolism. Selected sequences shared identity from 4.6% to 66.5% with characterised enzymes from \u003cem\u003eAlcanivorax borkumensis\u003c/em\u003e SK2 (ABO_0116, ABO_1197, ABO_1251, ABO_1483, ABO_1895, and ABO_2249) (SI Table\u0026nbsp;2) and according to the Arpigny and Jaeger (\u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e1999\u003c/span\u003e) classification of lipolytic enzymes, belong to the carboxylesterase family V with the characteristic catalytic Ser motif GX\u003cb\u003eS\u003c/b\u003eXGG (SI Fig.\u0026nbsp;1). Based on the sequence analysis eight proteins (APA_2, ALC24_4107, AGE_2, APA_8, AGE_8, ABO_1898, ABO_0116, ABO_1483) belong to family V, subfamily I, sharing serine motif with GX\u003cb\u003eS\u003c/b\u003eXG sequences. There were 5 sequences referred to family V subfamily III (AGE_12, ALC24_2069, ALC24_1162, ABO1197, ABO2449), sharing serine motif with GX\u003cb\u003eS\u003c/b\u003eGG, and the rest 8 sequences belonging to family V subfamily IV (APA_5, ALC24_1328, AGE_3, APA_4, ALC24_3989, APA_3, APA_6, ABO1251), with the serine motif GX\u003cb\u003eS\u003c/b\u003eXGX (Fig.\u0026nbsp;1).\u003c/p\u003e\u003cp\u003e\u003cstrong\u003eCarboxylesterase activity against chromogenic monoester substrates\u003c/strong\u003e\u003c/p\u003e\u003cp\u003eThe selected 21 genes encoding \u003cspan class=\"InlineEquation\"\u003e\u003cspan class=\"mathinline\"\u003e\\(\\:{\\alpha\\:}/{\\beta\\:}\\)\u003c/span\u003e\u003c/span\u003e-hydrolases and predicted carboxylesterases (S1 Table\u0026nbsp;1) were recombinantly expressed in \u003cem\u003eE. coli\u003c/em\u003e with an N-terminal 6His-tag and affinity purified (SI Fig.\u0026nbsp;2).\u003c/p\u003e\u003cp\u003eFrom 17 cloned genes, 11 were found to be expressed as soluble proteins, but only 5 clones produced highly purified proteins useful for detailed analysis (AGE_12, APA_2, APA_3, APA_5 and ALC24_1328) (S1 Table\u0026nbsp;1). Furthermore, six previously published esterases from \u003cem\u003eA. borkumensis\u003c/em\u003e SK2\u003csup\u003eT\u003c/sup\u003e (ABO_0116, ABO_1197, ABO_1251, ABO_1483, ABO_1895, and ABO_2249) were also induced and expressed and purified in this study (Tchigvintsev et al., \u003cspan citationid=\"CR72\" class=\"CitationRef\"\u003e2015\u003c/span\u003e; Hajighasemi et al., \u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e2016\u003c/span\u003e, \u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e2018\u003c/span\u003e). These 11 purified esterases were initially screened at 30 ºC using a range of chromogenic \u003cem\u003ep\u003c/em\u003eNP-esters with varying acyl chain lengths from C2 to C16 (Fig.\u0026nbsp;2). These screens revealed the presence of carboxylesterase activity in all purified enzymes, with most enzymes showing the highest activity with short-chain esters (acetyl and butyl-\u003cem\u003ep\u003c/em\u003eNP) and low activity against long acyl chain esters. In agreement with previous studies (Hajighasemi et al., \u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e2016\u003c/span\u003e, \u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e2018\u003c/span\u003e; Tchigvintsev et al., \u003cspan citationid=\"CR72\" class=\"CitationRef\"\u003e2015\u003c/span\u003e; Urbanek et al., \u003cspan citationid=\"CR75\" class=\"CitationRef\"\u003e2020\u003c/span\u003e) ABO_0116, ABO_1197, ABO_1251, ABO_1895, and ABO_2249 showed the preference to the short chain length substrates. Notably, ABO_2249 displayed the highest activity with \u003cem\u003ep\u003c/em\u003eNP-hexanoyl, consistent with previous studies (Urbanek et al., \u003cspan citationid=\"CR76\" class=\"CitationRef\"\u003e2018\u003c/span\u003e) highlighting it as promising candidate for polyester plastic depolymerisation. From five novel \u003cem\u003eAlcanivoracaceae\u003c/em\u003e proteins (AGE_12, APA_2, APA_3, APA_5 and ALC24_1328), APA_5 and ALC24_1328 exhibited the highest carboxylesterase activity with the C2 and C4 substrates, whereas the remaining three (AGE_12, APA_2 and APA_3) showed low activity (Fig.\u0026nbsp;2).\u003c/p\u003e\u003cp\u003e\u003cstrong\u003eActivity of carboxylesterases against different polyesters\u003c/strong\u003e\u003c/p\u003e\u003cp\u003e11 purified carboxylesterases from \u003cem\u003eAlcanivoracaceae\u003c/em\u003e were then screened for depolymerisation activity against various polyesters, including 3PET, aPET, PCL14, PCL2, PDLA, PLLA, PDLLA, PBA, PBS, PBT, PC, and Impranil DLN. Polyesterase assays were conducted using both the agarose plate-based screens and HPLC analyses of soluble reaction products after enzyme incubation with plastic emulsions. Visual inspection of agar plates revealed halos only around ABO_2249, ABO_1251, ALC24_1328, and ABO_1197 for 3PET or PCL2 (SI Fig.\u0026nbsp;3). However, no polyesterase activity against aPET, PLLA/PDLA, PLA, PBT, or PBS were detected in agarose plate screens. HPLC assays (Fig.\u0026nbsp;3) appeared more sensitive and revealed even slight depolymerisation not detected by agar plate visual inspection. Thus, polyesterase activity with short-chain polymers like 3PET or PCL2 was observed for all esterases (Fig.\u0026nbsp;3), indicating enzymes’ promiscuity and their potential for plastic-degrading activity. Furthermore, the HPLC analysis was performed to detect soluble reaction products following incubation of purified enzymes with emulsified polyester substrates aPET, PCL14, PDLA, PLLA, PDLLA, PBA, PBT and PC. These assays followed the formation of various polyester degradation products, such as TA, MHET, BA from 3PET, 6-HHA from PCL14 and PCL2, LA from PDLA, PLLA and PDLLA; AA from BPA, SA from PBS, TA from PBT, and BPA from PC. In these experiments, positive results were observed for PCL14, PCL2, PDLLA, PBA and 3PET as substrates, but no polyesterase activity was detected against aPET and PBS, PDLA, PLLA or PC (SI Table\u0026nbsp;3).\u003c/p\u003e\u003cp\u003eInterestingly, the major product during 3PET depolymerisation was MHET, with minor amounts of TA, and none of them producing BHET indicating that the selected enzymes possess high BHETase activity and low MHETase activity (Fig.\u0026nbsp;3). ABO_1197, ABO_1251, ALC24_1328 and APA_5 exhibited high polyesterase activity against 3PET, generating significant amount of BA, MHET and TA (from 0.2 to 1.35 mM). In contrast, ABO_0116, ABO_1483, ABO_1895, ALC24_1328 and APA_3 showed low polyesterase activity, producing little to no BA, MHET, or TA as products (\u0026lt; 0.1 mM). Surprisingly, the highest accumulation of MHET (at concentration of 0.78 mM) was observed for ABO_1251. Since this enzyme produced no visible halos on the 3PET-agrose plates, this discrepancy highlights the higher precision of HPLC analysis for quantifying the depolymerisation activity.\u003c/p\u003e\u003cp\u003eWith shorter-chain substrate PCL2, all tested \u003cem\u003eAlcanivoracaceae\u003c/em\u003e enzymes were active, generating 6-hydroxyhexanoic acid (6-HHA) at concentrations ranging from 0.1 to 0.5 mM. However, only five enzymes (ABO_1197, ABO_1251, ABO_2249, ALC24_1328 and APA_5) exhibited activity against the longer-chain PCL14 producing 6-HHA in the range 0.2–1.7 mM) (SI Table\u0026nbsp;3). A similar trend was observed with PDLLA (poly-D, L-lactide, MW range 10–18 kDa), where ABO_1197, ABO_1251, ABO_2249, APA_5, and ALC24_1328 produced significant levels of lactic acid (from 0.1 to 0.6 mM) (SI Table\u0026nbsp;3). The PDLLA polymer consists of a racemic mix of L- and D-lactic acid monomers, making it less crystalline than PLLA (L-lactic acid polymer) or PDLA (D-lactic acid polymer) (Lim et al., \u003cspan citationid=\"CR45\" class=\"CitationRef\"\u003e2008\u003c/span\u003e), which may facilitate its enzymatic degradation. For PBA degradation, six enzymes (ABO_1197, ABO_1251, ABO_1895, ABO_2249, AGE_12, APA_5) demonstrated hydrolytic activity, producing adipic acid at concentrations ranging from 0.1 to 0.3 mM (SI Table\u0026nbsp;3).\u003c/p\u003e\u003cp\u003eIn conclusion, five esterases, ABO_1197, ABO_1251, ABO_2249, APA_5, and ALC24_1328 were selected for further characterisation based on their promising polyesterase activities. Substrates and reaction products analysed in this study are presented in the supplementary SI Table\u0026nbsp;3.\u003c/p\u003e\u003cp\u003e\u003cstrong\u003eAnalysis of optimal reaction conditions for polyester depolymerisation\u003c/strong\u003e\u003c/p\u003e\u003cp\u003eTo identify the potential industrial applications of enzymes, they need to be screened for several key parameters, including substrate specificity, thermal stability to withstand industrial processing temperatures, pH stability across various industrial conditions, catalytic efficiency, and resistance to potential inhibitors present in industrial environments.\u003c/p\u003e\u003cp\u003eThe selected enzymes ABO_1197, ABO_1251, ABO_2249, APA_5, and ALC24_1328 were screened for their optimal conditions regarding pH, salinity (NaCl concentrations), detergent (Tween20), and temperature. The pH range was screened from pH 4.0 to pH 10.5, revealing that all selected enzymes preferred alkaline conditions (pH 8.0–pH 9.0) (Fig.\u0026nbsp;4A). Salinity tolerance was assessed with NaCl concentrations varying within 0–2.0 M, demonstrating that ABO_1197, ABO_1251, and ALC24_1328 maintained over 90% of activity at concentrations up to 0.1 M NaCl, while ABO_2249 and APA_5 generally exhibited lower salt tolerance. Notably, ABO_1197 showed the highest residual activity at 2 M NaCl concentration, retaining 23% of its initial activity (Fig.\u0026nbsp;4B).\u003c/p\u003e\u003cp\u003eFor detergent tolerance, the non-ionic surfactant Tween20 was used, ranging at concentrations from 0.1% to 2.0%. ABO_1197 and ABO_1251 showed increased activity with Tween20, which is often attributed to enhanced substrate accessibility. However, ABO_2249 and APA_5 experienced a decrease in activity even with small traces of detergent, while ALC24_1328 displayed remarkable detergent resilience, increasing its activity by up to 25% at 2.0% Tween20 (Fig.\u0026nbsp;5).\u003c/p\u003e\u003cp\u003eTemperature profiles were assessed from 5–70 ºC, with all selected enzymes showing mesophilic traits, favouring temperatures between 10 ºC and 50 ºC. Most enzymes lost two-thirds of their activity at 60 ºC. ABO_2249 and APA_5 demonstrated notable cold tolerance, with activities at 5 ºC being just 24% less than their respective optima (Fig.\u0026nbsp;6).\u003c/p\u003e\u003cp\u003e\u003cstrong\u003eAnalysis of thermostability\u003c/strong\u003e\u003c/p\u003e\u003cp\u003eIn the literature, protein stability is commonly assessed by measuring the melting temperature (Tm), which is defined as the temperature at which equal amounts of the protein are folded and unfolded under specific conditions (Huynh and Partch, \u003cspan citationid=\"CR36\" class=\"CitationRef\"\u003e2015\u003c/span\u003e; Sorgenfrei et al., \u003cspan citationid=\"CR67\" class=\"CitationRef\"\u003e2024\u003c/span\u003e). All applicable methods share the same principle: a sample is gradually heated, protein stability is monitored by melting temperature, which either measured by fluorescence signal of internal tryptophan residues or by using a fluorescence dye that changes its signal upon interacting with the unfolding protein. In this study, we investigated the thermostability of purified polyesterases by analysing the remaining enzyme activity after preincubation at different temperatures, as well as by identifying protein melting temperatures (T\u003csub\u003em\u003c/sub\u003e) using DSF.\u003c/p\u003e\u003cp\u003eAs shown in Fig.\u0026nbsp;7, after 5 h of preincubation at temperatures ranging from 5 to 40 ºC, the polyesterases retained almost 100% of their initial activity at 5 ºC. ABO_1197, ABO_1251, and ABO_2249 exhibited increased activity at 5 ºC and 10 ºC, however, ABO_1197 and ABO_1251 lost 40% and 80% of their activity at 30 ºC and completely lost it at 40 ºC. APA_5 and ALC24_1328 showed only a slight (2–5%) decrease in activity after incubation at 5 ºC. Overall, the selected enzymes maintained good activity within the low-temperature range (5–30 ºC) over prolonged incubation, with a maximum tolerance temperature of 30 ºC.\u003c/p\u003e\u003cp\u003eProtein melting temperatures (T\u003csub\u003em\u003c/sub\u003e) were determined based by DSF and are shown in SI Fig.\u0026nbsp;4. The highest melting temperature was observed for APA_5 (T\u003csub\u003em\u003c/sub\u003e 48.1 ºC), correlating with its high residual activity at 30 ºC after 5 h of incubation (Fig.\u0026nbsp;7).\u003c/p\u003e\u003cp\u003e\u003cstrong\u003eAnalysis of solvent resistance\u003c/strong\u003e\u003c/p\u003e\u003cp\u003eOrganic solvent tolerance of enzymes was determined by measuring activity in presence of solvents concentrations varying from 0–50%. Four different water-miscible organic solvents were chosen: ethanol, DMSO, dichloromethane (DCM), and 1,1,1,3,3,3-hexafluoro-2-propanol (HFP) based on their relevance for synthetic organic chemistry and for synthetic polymers. DCM and HFP are commonly used to dissolve the polyesters and were used for emulsions preparation in this work.\u003c/p\u003e\u003cp\u003eIn presence of HFP all enzymes retained less than 5% of their initial activity. Similar negative effect was observed for DCM (Fig.\u0026nbsp;8), with complete loss of the activity with more than 10% of DCM in reaction mixture. DMSO overall had a positive effect on the screened enzymes. ABO_1197, ABO_1251 and ABO_2249 showed 10–20% increase in the activity in presence of up to 20% DMSO, for APA_5 DMSO dramatically increased the activity, adding up to 50% of solvent in reaction mixture. Ethanol negatively influenced ABO_1197 and ABO_1251 activity, leading to decrease even at 5% of solvent in reaction mixture, but at the same time increased activity for the ABO_2244, APA_5 and ALC24_1328. For APA_5 at presence of 20% of ethanol in reaction mixture, activity was increased by 84%.\u003c/p\u003e\u003cp\u003eABO_2249, APA_5 and ALC24_1328 showed relative tolerance to the polar solvents of low polarity, increasing sensitivity with the increase of solvents polar index (DMSO \u0026lt; ethanol \u0026lt; HFP). 50% of DMSO retained positive effect on these enzymes, though for ethanol, less than 1% of activity was retained at 50% reaction mixture solution.\u003c/p\u003e\u003cp\u003e\u003cstrong\u003eCombined effects of solvents and temperature on the enzymatic polyester depolymerisation\u003c/strong\u003e\u003c/p\u003e\u003cp\u003eIn this study, selected enzyme-maintained activity at low temperatures 5–40 ºC (Fig.\u0026nbsp;9). ABO_2249 and APA_5 exhibited highest cold adaptation, with activities maintained at 70–85% at 5 ºC, compared to those at 30 ºC. We tested our enzymes activity with model substrate \u003cem\u003ep\u003c/em\u003eNP-hexanoate, in presence of DMSO and ethanol and observed improvement in the activity up to 25 to 100% with presence of 10–20% (v/v) for APA_5 and AlC24_1328. DCM and HFP showed stronger inhibiting effect on target enzymes activity (Fig.\u0026nbsp;8). Based on these results we selected 20% ethanol and DMSO to increase enzyme activity and 5% for DCM aiming decrease in crystallinity of substrate in the reaction mixture with actual polymeric substrates: 3PET, PCL14 and PDLLA. The incubations took place at 5 ºC and 30 ºC to make later comparisons. The reaction mixtures were filtered (10 kDa spin filters) and analysed by HPLC, the concentration of end products (mM) was used as standard to define the depolymerisation efficiency (Fig.\u0026nbsp;9). The results from 5 ºC incubation with 3PET, the presence of DMSO for enzyme APA_5 and ALC24_1328 increased production of BA and MHET compared with ‘no solvent’ conditions. For the rest of enzymes (ABO_1197, ABO_1251 and ABO_2249) any solvent addition at 5 ºC, resulted in decrease of depolymerisation efficiency.\u003c/p\u003e\u003cp\u003eAddition of organic solvents DMSO, ethyl alcohol and DCM at 30 ºC slightly improved the depolymerisation efficiency for reactions with APA_5 and ALC24_1328, with both PLA and 3PET (Fig.\u0026nbsp;9, 10) when compared with no solvent conditions. Interestingly, even accounting for the protein activity inhibition at 5% DCM (Fig.\u0026nbsp;9), the increase in product concentration for 3PET depolymerisation for APA_5 and ALC24_1328 was observed at 30 ºC, however at 5°C, the DCM addition decreased the yield of monomeric products threefold, as compared with 30°C. At the same time, if accounting only temperature effect on the enzyme activity more than twofold reduction in reaction products accumulation was observed in samples without solvent. The DCM has higher solubility in water at 5 vs 30 ºC (U.S. National Institute of Standards and Technology (NIST), Solubility Database, SRD 106, \u003cspan class=\"ExternalRef\"\u003e\u003cspan class=\"RefSource\"\u003ehttps://srdata.nist.gov/solubility/\u003c/span\u003e\u003cspan address=\"https://srdata.nist.gov/solubility/\" targettype=\"URL\" class=\"RefTarget\"\u003e\u003c/span\u003e\u003c/span\u003e), therefore at lower temperatures the non-polar interaction with protein structures must increase, potentially causing negative structural changes in proteins. DCM showed improved results for 3PET depolymerisation for APA_5 and ALC24_1328 only at 30 ºC.\u003c/p\u003e"},{"header":"Discussion","content":"\u003cp\u003eIn this study, eleven \u0026alpha;/\u0026beta;-hydrolases from \u003cem\u003eAlcanivoracaceae\u003c/em\u003e were purified and biochemically characterised for their carboxylesterase and polyesterase activities, including six previously reported enzymes (Hajighasemi et al., \u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e2016\u003c/span\u003e, \u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e2018\u003c/span\u003e; Tchigvintsev et al., \u003cspan citationid=\"CR72\" class=\"CitationRef\"\u003e2015\u003c/span\u003e), which were included as reference proteins (ABO_0116, ABO_1197, ABO_1251, ABO_1483, ABO_1895 and ABO_2249). All purified proteins exhibited carboxylesterase activity toward chromogenic \u003cem\u003ep\u003c/em\u003eNP-esters with a broad substrate range and a preference for short-chain acyl esters, which is typical for carboxylesterases (Martínez-Martínez et al., \u003cspan citationid=\"CR48\" class=\"CitationRef\"\u003e2018\u003c/span\u003e) (Hajighasemi et al., \u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e2016\u003c/span\u003e, \u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e2018\u003c/span\u003e; Tchigvintsev et al., \u003cspan citationid=\"CR72\" class=\"CitationRef\"\u003e2015\u003c/span\u003e).\u003c/p\u003e\u003cp\u003eHPLC-based assays with emulsified polyesters revealed detectable or significant polyesterase activity in all enzymes against PCL2 and the model PET-like substrate 3PET (Fig.\u0026nbsp;3). Six enzymes also hydrolysed PBA, five proteins degraded PCL14 and PDLLA, while no polyesterase activity was observed with other tested polyesters (aPET, PLLA, PDLA, PBS, PC, and PBT). These results confirm and extend our recent observations that polyesterase activity is widespread in carboxylesterases from families IV and V (Ma et al., \u003cspan citationid=\"CR46\" class=\"CitationRef\"\u003e2025\u003c/span\u003e). Moreover, our findings reinforce that HPLC-based assays provide greater sensitivity than agarose clearing methods for detecting polyesterase activity in purified proteins (Ma et al., \u003cspan citationid=\"CR46\" class=\"CitationRef\"\u003e2025\u003c/span\u003e). The \u003cem\u003eAlcanivoracaceae\u003c/em\u003e enzymes preferentially degraded aliphatic polyesters with shorter chain lengths and lower molecular weights, while aromatic polyesters were recalcitrant. For example, polyesterase activity was observed on substrates such as PCL2, PCL14, PDLLA, PBA, whereas limited or no activity was detected on PC, PBT and aPET. Generally, the polyester chain length strongly influences chain mobility, crystallinity, and thus susceptibility to enzymatic hydrolysis. Longer and more crystalline polymers exhibit enhanced intermolecular interactions, such as Van der Waals forces, requiring higher activation energy for depolymerisation. From an industrial perspective, cold-adapted enzymes offer distinct advantages, including reduced energy costs and protection of thermolabile products and intermediates (Santiago et al., \u003cspan citationid=\"CR62\" class=\"CitationRef\"\u003e2016\u003c/span\u003e)). However, because polyester crystallinity remains high at low temperatures, appropriate decrystallinization pretreatments will likely be required to enable complete depolymerization with cold-adapted enzymes.\u003c/p\u003e\u003cp\u003eMarine microorganisms represent an important reservoir of cold-active and cold-tolerant enzymes due to the predominantly low temperatures of seawater habitats (Médigue et al., \u003cspan citationid=\"CR49\" class=\"CitationRef\"\u003e2005\u003c/span\u003e; Kube et al., \u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e2013\u003c/span\u003e; Santiago et al., \u003cspan citationid=\"CR62\" class=\"CitationRef\"\u003e2016\u003c/span\u003e). For instance, EstB from \u003cem\u003eAlloalcanivorax dieselolei\u003c/em\u003e, displayed optimal activity around 20 ºC and retained over 95% of activity between 0 and 10 ºC (Degli-Innocenti et al., \u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e2023\u003c/span\u003e; Zhang et al., \u003cspan citationid=\"CR92\" class=\"CitationRef\"\u003e2014\u003c/span\u003e). Similarly, RhLip from \u003cem\u003eRhodococcus\u003c/em\u003e sp. AW25M09 retained 50% activity at 10 ºC (Hjerde et al., \u003cspan citationid=\"CR34\" class=\"CitationRef\"\u003e2013\u003c/span\u003e), and a carboxylesterase from \u003cem\u003ePsychrobacter cryohalolentis\u003c/em\u003e maintained over 90% of its maximal activity at 0–5 ºC (Novototskaya-Vlasova et al., \u003cspan citationid=\"CR53\" class=\"CitationRef\"\u003e2012\u003c/span\u003e). The carboxylesterase OLEI01171 from \u003cem\u003eOleispira antarctica\u003c/em\u003e RB-8, another marine hydrocarbon degrader, also exhibits high activity between 5 and 30 ºC (Lemak et al., \u003cspan citationid=\"CR44\" class=\"CitationRef\"\u003e2012\u003c/span\u003e). Structural analyses of enzymes from \u003cem\u003eO. antarctica\u003c/em\u003e RB-8 and their mesophilic homologues suggested that enhanced flexibility at the active sites underlies the catalytic efficiency of cold-active enzymes, compensating for the reduced thermal energy in cold environments (Kube et al., \u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e2013\u003c/span\u003e). Consistent with this, all enzymes characterised in this study were highly active and stable at low temperatures (5–20°C), with protein melting temperatures (T\u003csub\u003em\u003c/sub\u003e) ranging from 30 to 48 ºC. Enzymes with lower T\u003csub\u003em\u003c/sub\u003e values (ABO_1197 and ABO_1251) exhibited partial reactivation at 5°C, whereas Apa_5 (T\u003csub\u003em\u003c/sub\u003e 48.1°C) retained activity over a wider temperature range. These results indicate that lower T\u003csub\u003em\u003c/sub\u003e values correlate with increased cold tolerance, while higher T\u003csub\u003em\u003c/sub\u003e values correspond to greater enzyme thermostability. The enhanced cold tolerance observed in ABO_1197 and ABO_1251 may result from increased conformational flexibility conferred by residues such as Gly, Ser, and Met, and a lower abundance of Pro and Arg residues, which typically constrain structural mobility (Parvizpour et al., \u003cspan citationid=\"CR56\" class=\"CitationRef\"\u003e2021\u003c/span\u003e). However, comparative sequence analysis of cold-active and thermostable esterases did not reveal a consistent correlation between amino acid composition and cold tolerance.\u003c/p\u003e\u003cp\u003eSeveral \u003cem\u003eAlcanivoracaceae\u003c/em\u003e polyesterases (ABO_1197, ABO_1251, and ALC24_1328) exhibited high tolerance or partial activation in the presence of Tween 20 (Fig.\u0026nbsp;5), a feature frequently observed among cold-adapted \u0026alpha;/\u0026beta;-hydrolases. For instance, the three thermophilic metagenomic polyesterases from \u003cem\u003eIschia\u003c/em\u003e showed no activation and low tolerance to detergents (Distaso et al., \u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e2023\u003c/span\u003e), whereas several cold-adapted carboxylesterases from marine bacteria and a cold-active protease from \u003cem\u003ePsychrobacter\u003c/em\u003e sp. 94-6PB were stimulated by the addition of detergents, including Tween 20 (Novototskaya-Vlasova et al., \u003cspan citationid=\"CR53\" class=\"CitationRef\"\u003e2012\u003c/span\u003e; Wu et al., \u003cspan citationid=\"CR82\" class=\"CitationRef\"\u003e2013\u003c/span\u003e; Jiang et al., 2016; Perfumo et al., \u003cspan citationid=\"CR57\" class=\"CitationRef\"\u003e2020\u003c/span\u003e).\u003c/p\u003e\u003cp\u003eOrganic solvents can enhance enzymatic transformations by increasing substrate solubility and modulating enzyme flexibility via alterations in hydrogen bonding (Osbon and Kumar, 2019; Sorgenfrei et al., \u003cspan citationid=\"CR67\" class=\"CitationRef\"\u003e2024\u003c/span\u003e). However, this simultaneously requires a certain level of enzyme tolerance to organic solvents. Given that \u003cem\u003eAlcanivoracaceae\u003c/em\u003e species thrive in oil- and hydrocarbon-rich environments (Sabirova et al., \u003cspan citationid=\"CR61\" class=\"CitationRef\"\u003e2006\u003c/span\u003e; Zhang et al., \u003cspan citationid=\"CR92\" class=\"CitationRef\"\u003e2014\u003c/span\u003e), their enzymes are expected to exhibit tolerance to both low temperatures and organic solvents. Nevertheless, the inherent structural flexibility of cold-adapted enzymes may render them more susceptible to the destabilizing effects of elevated temperatures and organic solvents (Sellek and Chaudhuri, \u003cspan citationid=\"CR65\" class=\"CitationRef\"\u003e1999\u003c/span\u003e). To date, only a limited number of cold-adapted esterases and lipases with solvent tolerance have been described (Lee et al., \u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e2017\u003c/span\u003e). For instance, high solvent resistance was reported for the EstB esterase from \u003cem\u003eAlloalcanivorax dieselolei\u003c/em\u003e B-5\u003csup\u003eT\u003c/sup\u003e with the retention of 80% activity in the presence of isopropanol (70%) or acetone (up to 70%) (Zhang et al., \u003cspan citationid=\"CR92\" class=\"CitationRef\"\u003e2014\u003c/span\u003e)). In the presence of 50% dimethyl sulfoxide (DMSO) or ethanol, EstB retained 83% and 87% of activity, respectively, but it was strongly inhibited by acetonitrile and methanol (Zhang et al., \u003cspan citationid=\"CR92\" class=\"CitationRef\"\u003e2014\u003c/span\u003e). In the present study, the addition of DMSO up to 20% increased the carboxylesterase activity of all tested enzymes from \u003cem\u003eAlcanivoracaceae\u003c/em\u003e (Fig.\u0026nbsp;8). Ethanol at 10–20 % stimulated the activity of three enzymes but was inhibitory at higher concentrations, while DCM (dichloromethane) and HFP (hexafuoroisopropanol) caused strong inhibition. The addition of DMSO or ethanol enhanced 3PET depolymerisation by APA_5 and ALC24_1328 at 30°C, whereas no stimulation was observed at 5°C (Fig.\u0026nbsp;9). These findings suggest that, within the optimal range, higher reaction temperatures exert a more pronounced positive effect on enzymatic polyester depolymerisation than the presence of organic solvents.\u003c/p\u003e"},{"header":"Conclusion","content":"\u003cp\u003eThis study expands the current understanding of \u003cem\u003eAlcanivoracaceae\u003c/em\u003e α/β-hydrolases by demonstrating that polyesterase activity is a common feature among their carboxylesterases, particularly within families IV and V. Eleven enzymes tested exhibited carboxylesterase activity with a preference for short-chain monoesters, as well as polyesterase activity toward aliphatic polyesters such as PCL and PBA, while showing limited activity toward aromatic polyesters. Their notable activity and stability at low temperatures, together with tolerance to detergents and certain organic solvents, reflect their adaptation to hydrocarbon-rich marine environments. These findings provide new insights into the biochemical diversity and ecological roles of \u003cem\u003eAlcanivoracaceae\u003c/em\u003e enzymes and highlight their potential for environmentally friendly biotransformations, particularly in low-temperature, energy-efficient plastic biodegradation and recycling.\u003c/p\u003e"},{"header":"Abbreviations","content":"\u003cp\u003e3PET, bis(benzoyloxyethyl) terephthalate; PET, poly(ethylene terephthalate); aPET, amorphous poly(ethylene terephthalate); PU, polyurethane; PE, polyethylene; PS, polystyrene; PC, polycarbonate; PCL14, polycaprolactone (14kDa); PCL2, polycaprolactone (2 kDa); PBS, polybutylene succinate; PBA, polybutylene adipate; PBT, polybutylene terephthalate; PLLA, poly-L-lactide; PDLA, poly-D-lactide; PDLLA, poly-D,L-lactide; MHET, mono(2-hydroxyethyl) terephthalate; BHET, bis(2-hydroxyethyl) terephthalate; TA, terephthalic acid; BPA, bisphenol-A; DMSO, dimethyl sulfoxide; DCM, dichloromethane; HFP, 1,1,1,3,3,3-hexafluoro-2-propanol\u0026nbsp;\u003c/p\u003e\n"},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eFunding declaration\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThis research was supported by the FuturEnzyme project funded by the European Union Horizon 2020 Research and Innovation Program under grant agreement 101000327, UKRI-funded projects P3EB (UKRI Engineering Biology Mission Hub, grant BB/Y007972/) and EBIC (UKRI Engineering Biology Mission Hub, grant (grant Nr BB/Y008332/1). PNG and AFY also acknowledge support from the \u0026ldquo;Plastics Vector\u0026rdquo; project NE/S004548/1 funded by the Natural Environment Research Council (NERC, UK). OVG, PNG, and AFY are also thankful for support from the European Regional Development Fund (ERDF) through the Welsh Government to the Centre for Environmental Biotechnology, project number 81280.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAuthors\u0026rsquo; contributions\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003ePNG, AFY, MF, OVG, and ANK designed experiments and interpreted the data. HM wrote the manuscript draft and edited it with HM, ANK, TNC performed experiments. PNG, MF, AFY and OVG provided the funding.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\n\u003cli\u003eAbou-Zeid DM, M\u0026uuml;ller RJ, Deckwer WD (2004) Biodegradation of aliphatic homopolyesters and aliphatic\u0026ndash;aromatic copolyesters by anaerobic microorganisms. 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Front Bioeng Biotechnol 12:1349010. https://doi.org/10.3389/fbioe.2024.1349010\u003c/li\u003e\n\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":true,"highlight":"","institution":"","isAcceptedByJournal":true,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"researchsquare","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":true,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"/submission","title":"Research Square","twitterHandle":"researchsquare","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"","reportingPortfolio":"","inReviewEnabled":false,"inReviewRevisionsEnabled":true},"keywords":"Carboxylestereases, polyesters, cold-adapted enzymes, marine hydrocarbon-degrading bacteria, Alcanivoracaceae, Alcanivorax","lastPublishedDoi":"10.21203/rs.3.rs-7917997/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-7917997/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003cp\u003eMembers of the family \u003cem\u003eAlcanivoracaceae\u003c/em\u003e are widespread in marine environments, where they play central roles in hydrocarbon degradation and populate plastics-associated microbiomes, with notable enzymatic potential toward ester- and olefin-based polymers. To further investigate their enzymatic potential, we selected 21 candidate enzymes from the α/β-fold hydrolase superfamily, specifically carboxylesterase Family V from genome-sequenced representatives of the genera \u003cem\u003eAlcanivorax, Alloalcanivorax\u003c/em\u003e, and \u003cem\u003eIsoalcanivorax\u003c/em\u003e. Seventeen enzymes were cloned and heterologously expressed in \u003cem\u003eE. coli\u003c/em\u003e, of which eleven were purified and subjected to substrate specificity analyses alongside six previously reported and partially characterised carboxylesterases from \u003cem\u003eA. borkumensis\u003c/em\u003e SK2, used as benchmarks. All enzymes showed activity against soluble model \u003cem\u003ep-\u003c/em\u003enitrophenyl ester substrates with acyl chain lengths ranging from C2 to C12 and against bis(benzoyloxyethyl) terephthalate (3PET) and polycaprolactone 2 kDa (PCL2). During 3PET hydrolysis, product accumulation followed the order: benzoic acid\u0026thinsp;\u0026gt;\u0026thinsp;\u0026gt;\u0026thinsp;MHET\u0026thinsp;\u0026gt;\u0026thinsp;terephthalic acid. Five enzymes hydrolysed polycaprolactone 14 kDa (PCL14), poly-D,L-lactide (PDLLA), and polybutylene adipate (PBA). All five enzymes displayed temperature optima around or below 50\u0026deg;C and retained high activity at low temperatures (5\u0026ndash;20\u0026deg;C), consistently with adaptation to marine environments. Enzymes also exhibited moderate solvent tolerance, neutral-to-alkaline pH optima, and low thermostability, with melting temperatures (Tm) between 31\u0026deg;C and 48\u0026deg;C. Overall, enzymes from \u003cem\u003eAlcanivoracaceae\u003c/em\u003e exhibited promising potential for synthetic polyesters biodegradation, especially under low-temperature conditions, suggesting potential application for degrading specific polyester-based plastics with lower molecular weight, and their utility in further enzyme engineering for plastic recycling and upcycling.\u003c/p\u003e","manuscriptTitle":"Cold-adapted carboxylesterases from Alcanivoracaceae active with a wide range of synthetic polyesters","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2025-11-14 09:23:06","doi":"10.21203/rs.3.rs-7917997/v1","editorialEvents":[{"type":"communityComments","content":0}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"researchsquare","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":true,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"/submission","title":"Research Square","twitterHandle":"researchsquare","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"","reportingPortfolio":"","inReviewEnabled":false,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"9405440a-628c-4267-9788-cf2d5ef23323","owner":[],"postedDate":"November 14th, 2025","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"posted","subjectAreas":[],"tags":[],"updatedAt":"2026-02-09T16:03:34+00:00","versionOfRecord":{"articleIdentity":"rs-7917997","link":"https://doi.org/10.1007/s00253-026-13726-z","journal":{"identity":"applied-microbiology-and-biotechnology","isVorOnly":false,"title":"Applied Microbiology and Biotechnology"},"publishedOn":"2026-02-07 15:59:57","publishedOnDateReadable":"February 7th, 2026"},"versionCreatedAt":"2025-11-14 09:23:06","video":"","vorDoi":"10.1007/s00253-026-13726-z","vorDoiUrl":"https://doi.org/10.1007/s00253-026-13726-z","workflowStages":[]},"version":"v1","identity":"rs-7917997","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-7917997","identity":"rs-7917997","version":["v1"]},"buildId":"8U1c8b4HqxoKbykW_rLl7","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}