Photo-driven decarboxylation for sustainable biofuel production: a review on harnessing the potential of fatty acid decarboxylases

Abstract

Fatty acid photodecarboxylase (FAP) has emerged as a green biocatalyst to produce biofuel from sustainable feedstocks under the illumination of visible light. However, oxidative damage by photogenerated radicals and structural destabilization of enzymes by organic solvents result in the poor stability of FAP. Meanwhile, the low efficiency of light and mass transfer in conventional photoreactors and poor substrate availability limit the catalytic performance of FAP. This review covers three strategies to improve FAP stability: (1) the removal of the radical effect, (2) the improvement of enzyme tolerance to organic solvents, and (3) the directed evolution of FAP by protein engineering. Furthermore, the design of a photobioreactor and enzyme engineering are elaborately discussed to enhance catalytic performance.

---

Feng Li

Feng Li is currently pursuing his PhD at Chongqing University under Prof. Ao Xia. Prior to this, he received his BS degree in Energy and Environmental Engineering from the Fujian University of Technology in 2019 and his MS degree in Power Engineering and Engineering Thermophysics from Chongqing University in 2022. His research focuses on the stability of photoenzymes and the production of sustainable aviation fuel.

Yaqi Sui

Yaqi Sui received her BS degree in Biomedical Engineering from Shandong Second Medical University in 2021 and her MS degree in Biology and Medicine from Chongqing University in 2024, where she focused on protein post-translational modifications. Her current doctoral research centers on developing advanced optical imaging techniques and exploring their applications in life sciences, with the aim of providing new insights into cellular processes and disease mechanisms.

Kai Lin

Dr Kai Lin is currently a lecturer in the State Key Laboratory of Photocatalysis on Energy and Environment, College of Chemistry, Fuzhou University. He obtained his BS degree in 2018 and his PhD degree in 2024 from Chongqing University, under the supervision of Prof. Qiang Liao and Ao Xia. He conducted research at the Department of Chemical and Biomolecular Engineering, National University of Singapore, as a visiting PhD student from 2022 to 2023. His research primarily focuses on biomass energy, photocatalytic hydrogen production and the design of highly efficient reactors.

Xiaobo Guo

Dr Xiaobo Guo is a research associate at the Biogas Institute of Ministry of Agriculture and Rural Affairs and at Research Center for Rural Energy and Ecology, Chinese Academy of Agricultural Science. He received his master's degree (2016) from the School of Chemistry and Chemical Engineering at Guangxi University and his PhD degree (2022) from the Institute of Engineering Thermophysics, Chongqing University. His research focuses on energy and resource utilization of agricultural waste through biochemical conversion.

Ao Xia

Dr Ao Xia is a professor in the School of Energy and Power Engineering at Chongqing University. He received his bachelor's degree (2008) and PhD degree (2013) from Zhejiang University. He worked as a post-doctoral researcher in the Environmental Research Institute at University College Cork from 2014 to 2016. His research focuses on bioenergy, biofuels, and renewable distributed energy systems.

Vinod Kumar

Dr Vinod Kumar is a senior lecturer in sustainable biomanufacturing in the Magan Centre for Applied Mycology at Cranfield University. Before joining Cranfield University, Dr Vinod Kumar was a Marie Curie International Incoming Fellow working at the EPSRC/BBSRC Synthetic Biology Research Centre (SBRC) at the University of Nottingham, UK. Prior to this, he worked as a Post-Doctoral Fellow in France, South Korea and the United Kingdom. He graduated from the School of Chemistry at Hindu College, University of Delhi, India, with a BS degree. He earned his MS degree (Chemistry) and PhD degree (Biochemical Engineering & Biotechnology) from the Indian Institute of Technology Delhi, India. His expertise in developing cleaner synthetic routes and designing robust cell factories and processes makes the production of a range of chemicals possible in a way that maximises efficiency and minimises waste using a circular economy approach.

Xun Zhu

Dr Xun Zhu is a professor in the School of Energy and Power Engineering at Chongqing University. She obtained her bachelor's degree (1990) from the School of Mechanical and Power Engineering at Shanghai Jiao Tong University. Then, she received her master's (1993) and PhD (2002) degrees from the School of Energy and Power Engineering at Chongqing University. Her current research focuses on microbial energy conversion, waste heat recovery, and hydrogen and fuel cell technologies.

Wenhui Zhao

Dr Wenhui Zhao is a professor in the School of Life Sciences at Chongqing University. He obtained his BS degree (1991) from the Department of Biology at Nankai University. Then, he earned his PhD degree (2000) from the Shanghai Institute of Cell Biology at the Chinese Academy of Sciences. Before join Chongqing University, he worked as a Postdoc at the Medical School, Yale University (2000–2001), Weill Medical College, Cornell University (2001–2003), Institute for Cancer Genetics, Columbia University (2003–2008). Then, he worked as an Associate Scientist at the Institute for Cancer Genetics, Columbia University (2008–2013) and as a researcher at the Health Science Center, Peking University (2004–2020). His research focuses on the role of post-translational protein modification and RNA acetylation in major diseases.

Qiang Liao

Dr Qiang Liao is a professor in the School of Energy and Power Engineering at Chongqing University. He obtained his bachelor's degree (1985) in mechanical power from the Huazhong University of Science and Technology. Then, he earned his PhD degree (1993) in Power Engineering and Engineering Thermophysics from Chongqing University. He worked as a post-doctoral researcher in the Mechanical Engineering Institute at the Hong Kong University of Science and Technology from 1997 to 1999. His current research focuses on biomass energy, heat and mass transfer, and energy conservation and emission reduction technologies.

---

1. Introduction

For achieving net-zero CO₂ emissions before 2050, the International Energy Agency (IEA) aims to adopt carbon-neutral fuels to replace traditional fossil-based feedstocks, mainly responsible for emissions of greenhouse gases.^1,2 Meanwhile, the development and utilization of sustainable bio-liquid fuel and biogas as alternative fuels contribute to reducing the oil dependence and the achievement of global energy targets along with the transition to a carbon-neutral society.^3–5 The aviation sector accounts for ∼2% of global CO₂ emissions every year due to its substantial reliance on petroleum-derived aviation fuel.^6,7 According to the International Air Transport Association (IATA), 302 million tonnes of commercial aviation fuel were consumed in 2023, which caused over 950 million tonnes of CO₂ emissions.^8,9 To achieve the IATA's declared target of 50% CO₂ emission reduction in the aviation industry by 2050, the application of sustainable aviation fuel (SAF) needs to be prioritized,¹⁰ as the use of SAF can achieve up to 90% reduction in CO₂ emissions across its full well-to-wake life cycle (from raw material extraction to aircraft operation).⁹ However, the global share of SAF consumption reached only 0.3% in 2024, and hence the displacement of conventional aviation fuel is marginal.¹¹

The hydrocarbons with lengths ranging from C₈ to C₁₆, comprising n-alkanes, iso-alkanes, aromatics and cycloalkanes, are fundamental components of SAF.^12,13 The renewable feedstocks such as agricultural residues, forestry wastes, waste cooking oils and municipal solid wastes can be upgraded to produce SAF.¹⁴ In particular, waste cooking oils, as widely available feedstocks, are typically converted to SAF by hydrodeoxygenation.¹⁵ Owing to the high technological maturity, hydroprocessed oil accounts for 89% of the current global SAF production capacity.¹⁶ However, energy-intensive consumption of the hydroprocess and low selectivity of the catalyst lead to its production cost being 3–5 times higher than that of conventional petroleum-derived aviation fuel, thereby limiting the large-scale applications.¹⁷ Other routes for SAF production, including Fischer–Tropsch synthesis and alcohol-to-jet and pyrolysis oil upgrading, are difficult to exploit at an industrial scale due to high equipment capital expenditures and difficulty in removing impurities (oxygen and nitrogen compounds).^18,19 Therefore, the development of an alternative pathway for SAF production is essential for the aviation industry to achieve the target of 50% reduction in CO₂ emissions by 2050.

In 2017, Beisson's group reported a novel photoenzyme derived from the microalga Chlorella variabilis NC64A, which catalyzes the decarboxylation of free fatty acids (C_n) to alkanes (C_n−1) under blue light illumination.²⁰ This is an alternative and promising route for SAF production using a mild process. This photo-driven enzyme, subsequently termed fatty acid photodecarboxylase (FAP), possesses flavin adenine dinucleotide (FAD) as a cofactor for capturing light and a hydrophobic substrate tunnel. The investigation of the substrate scope showed that FAP exhibited higher enzyme activity for long-chain fatty (C₁₆–C₁₈) acids, but lower enzyme activity for medium-chain (C₈–C₁₂) and short-chain fatty acids (<C₈). This is ascribed to the inefficient hydrogen bonding and stabilization for relatively shorter hydrophobic chains in medium and short-chain fatty acids, as the substrate pocket is a long and narrow region.

Hollmann et al. first calculated the turnover numbers (TON) of FAP and then demonstrated that the corresponding triglycerides could be converted to hydrocarbons by lipase hydrolysis and FAP decarboxylation with the highest TON of up to 8280.²¹ Focused on the elucidation of the FAP photocycle, different perspectives were proposed by spectroscopic, crystallographic, and computational studies.^22–27 Meanwhile, a few studies have reviewed the development of FAP, providing a summary of the findings related to FAP in sustainable hydrocarbon fuel production, the catalytic mechanism from structural biology perspectives, and biotechnological applications of FAP.^17,28–31 However, the systematic investigation of FAP inactivation dynamics and the targeted activity-enhancing strategies remains underexplored. Considering the huge potential of FAP in sustainable biofuel production, this paper aims to discuss the various insights into the mechanism of FAP catalysis and inactivation, followed by different strategies for enhancing the stability and catalytic performance in FAP-mediated biofuel production.

2. FAP catalytic cycle and inactivation

2.1 Decarboxylation mechanism of FAP

The FAD cofactor functions as the light absorbing molecule in the decarboxylation reaction. However, the insight into the interaction between FAD and fatty acids was not clear (Fig. 1). According to Beisson's group, FAD was photoexcited to ¹FAD* and then an electron was transferred from the fatty acid to ¹FAD* in 300 ps, which was measured by time-resolved optical spectroscopy.^20,22 Zhang et al. found that the photoexcitation of FAP and fatty acid yields a CH₃COO˙···FAD˙ radical pair and the generated CH₃˙ radical binds to the anionic flavin semiquinone, thereby causing the formation of a covalent dearomatized methylquinone flavin adduct (CH₃–FAD⁻).²⁶ The CH₃–FAD⁻ intermediate is protonated to produce an alkane and regenerate the oxidized FAD for closing the photocycle. This photo-driven decarboxylation process is based on the analysis of density functional theory (DFT) and time-dependent (TD-DFT) density functional theory and is conducted in a representative model system. Hong et al. proposed an alternative interaction mechanism.²³ The formed ¹FADH, rather than ¹FAD*, is more likely to withdraw the electron from the fatty acid anion as determined by DFT analysis. Afterwards, alkyl R˙ is bound to a hydrogen atom from the FADH˙ radical to form the final product, RH. However, the decarboxylation mechanism envisioned by Zhang et al. and Hong et al. is based on a computational study. The existence of CH₃–FAD⁻ and FADH˙ in the decarboxylation process has not been demonstrated experimentally. Moreover, it seems challenging for neighboring residues of the active site to donate a proton to FAD˙⁻ and form FADH˙ species.³² Future work on the investigation of the dynamic mechanism of radical intermediates will further reveal the catalytic photocycle.

Fig. 1 Interaction between FAD and fatty acids and FAP inactivation. (a) Bession's trimolecular mechanism.²² Reproduced from ref. 22 with permission from American Association for the Advancement of Science, copyright 2021. (b) Hong's mechanism with FADH biomolecular species.²³ Reproduced from ref. 23 with permission from Elsevier, copyright 2021. (c) Zhang's mechanism with the CH₃–FAD⁻ methylquinone flavin adduct.²⁶ Reproduced from ref. 26 with permission from American Chemical Society, copyright 2019. (d) The generation pathway for protein- and oxygen-based radicals.

After the decarboxylation of the fatty acid, the formation process or pathway of alkanes remains controversial (Fig. 2). Heyes et al. proposed that cysteine 432 (Cys 432) mediated H atom transfer (HAT) is the formation pathway of alkanes during FAP photocatalysis (Fig. 2a).²⁵ Specifically, the deprotonated state of Cys 432 coupled with the H atom transferred to the alkyl radical (R˙) further yields a Cys thiyl radical and an alkane product, which is ascribed to the Cys 432 thiol group being sufficiently close to the alkyl radical. Subsequently, the electron transferred from the FAD semiquinone to Cys 432 quenches the Cys thiyl radical to form the thiolate species. The thiolate is reprotonated to restart a new photocycle. However, Beisson's group has indicated that HAT is unlikely to occur based on experimental findings. Firstly, Cys 432 was demonstrated to have no effect on alkane formation since the Cys 432 mutant also showed a significant enzymatic activity. The decrease in enzyme activity due to the replacement of the Cys 432 mutant, reported by Heyes et al., may result from structural changes in the protein, rather than a point mutation. Meanwhile, no cysteine deprotonation was observed in the red-shifted state of FAD, but only thiol S–H vibration shift occurred according to Fourier transform infrared spectroscopy (FTIR) experiments. In another alkane formation mechanism reported by Beisson's group, the PCET pathway enabled the formation of alkanes (Fig. 2b).²² In the PCET pathway, a proton sourced from an arginine residue (Arg 451) or a water molecule close to FAD is transferred to R˙, accompanied by an electron transfer from anionic FAD semiquinone. Arg 451 is considered to be a proton donor in the FAP photocycle, which is proven by cryotrapping FTIR experiments.

Fig. 2 Alkane formation mechanism in the FAP photocycle. Alkane formation by (a) H transfer.²⁵ Reproduced from ref. 25 with permission from American Chemical Society, copyright 2020. (b) Proton-coupled electron transfer (PCET).²² Reproduced from ref. 22 with permission from American Association for the Advancement of Science, copyright 2021.

From the perspective of enzyme engineering, different substrate chain lengths may trigger different pathways of alkane formation. The catalysis of long-chain fatty acids (e.g., C16 and C18) by the wild type FAP may prefer Arg451-mediated PCET, which is ascribed to the contribution of the Arg451 residue to electrostatic adsorption onto the hydrophobic carbon chain. Meanwhile, Arg451 helps the spatial localization of long-chain fatty acids through salt bridging.^22,32 Therefore, we assume that Arg451 not only provides a proton for alkane formation, but also is a recognition site for long-chain fatty acids. However, this phenomenon did not exist in the engineered mutant for decarboxylation of short-chain fatty acids (e.g., C3 and C4). The Arg451 residue exhibited electrostatic repulsion to propionic acid in our recent study. This caused the short-chain fatty acids to not be able to localize, thereby limiting the proton transfer between Arg451 and the substrate. We suspected that short-chain acids may shift to the Cys432 HAT pathway, as Cys432 significantly contributes to the enhancement of enzyme activity for short-chain fatty acids.²⁵

2.2 Inactivation of FAP

During the reaction, both prolonged photoexcitation and organic solvent exposure contribute to FAP inactivation through distinct mechanisms. Current research attributes FAP inactivation to two primary pathways.

The first is the photodamage of protein under light illumination. Lakavath et al. first reported a catalytically inactive enzyme produced under light conditions.³³ By pre-illumination of FAP at 1000 μmol m⁻² s⁻¹ light intensity, the catalytic activation of FAP was significantly decreased to 40% in 30 min with a FAP half-life of 5 min. Such a phenomenon was considered to be the photoinactivation of FAP and an intrinsic property, which is not dependent on assay reagents or oxygen. To verify the mechanism of photoinactivation, an electron paramagnetic resonance (EPR) test was adopted to detect the radical signal. Interestingly, large radical signals were observed when FAP was exposed to blue light. Such radical signals were attributed to the formation of a FAD semiquinone and a protein-based organic radical. For example, in a typical generation process of the radical (Fig. 1d), FAD is firstly transferred to the triple excited state (³FAD*) by intersystem crossing under 450 nm light irradiation. For oxygen-based radicals, with the involvement of the oxygen molecule, O²˙⁻ radicals are produced along with an electron transfer from ³FAD* to O₂; ¹O₂ is produced after transferring the excitation energy of ³FAD* to O₂. For protein-based radicals, in the presence of tryptophan (Typ) and tyrosine (Tyr), the indole cationic radical and the phenoxyl radical are produced by the energy transfer from ³FAD* along with electron transfer, as aromatic rings in Typ and Tyr could not directly absorb the energy from 450 nm light.³⁴ Finally, these radicals destroy the structure of FAP by crosslinking and oxidation, which further causes the enzyme instability.³⁵ For example, two phenoxyl radicals combine to form dityrosine, generating stable covalent crosslinking by dimerization, which inevitably causes the conformational change of FAP.

However, an oxygen-involved environment demonstrated significant inhibition of fatty acid decarboxylation and caused the inactivation of FAP.³⁶ By the EPR test, the superoxide radical (O₂˙⁻) and singlet oxygen radical (¹O₂), as reactive oxygen species (ROS), were probed in an aerobic environment under light illumination. Such radicals damaged the FAP protein by oxidation, thereby decreasing the catalytic activation of FAP. The detection of amino acid-based radicals was not performed in this study. Therefore, the formation mechanism of protein-based radicals is unclear. Nevertheless, radicals inevitably result in photoinactivation of FAP and urgently need to be eliminated to enhance the stability of FAP for biofuel production.

Organic solvent exposure represents another critical inactivation pathway. According to the previous studies reported, dimethyl sulfoxide (DMSO), the most commonly used solvent for fatty acid decarboxylation, can strip water molecules from the vicinity of FAP, thereby breaking the hydrogen bond and altering the tertiary structure of FAP.³⁷ At 60% (v/v) DMSO, fatty acid conversion plummeted to 3.3%, while reducing solvent concentration to 30% restored conversion to 94.1%, establishing this threshold as optimal for FAP catalysis.¹⁷ Focused on the various inactivation mechanisms of FAP, the corresponding strategies to enhance its catalysis from stability and performance perspectives were proposed and discussed.

3. Strategies to enhance the stability of FAP

3.1 The removal of radical inhibition

3.1.1 Inhibition of ROS radical formation. The limitation of the involvement of oxygen is a valid method to inhibit radical formation. The establishment of an anaerobic environment was documented as an efficient approach to improve the photostability of FAP.³⁶ The pentadecane yield was reported to be increased by 44.7% in an anaerobic environment for palmitic acid decarboxylation by FAP within 3 h, as compared to that under aerobic conditions (Fig. 3a and b).

Fig. 3 The inhibition mechanism of oxygen in fatty acid decarboxylation by FAP.³⁶ Reproduced from ref. 36 with permission from Elsevier, copyright 2023. (a) Schematic diagram of FAP catalysis during aerobic and anaerobic conditions. (b) Inhibition of alkane biofuel production by oxygen through photoenzymatic decarboxylation. (c) EPR spectra of DMPO/O₂˙⁻. (d) EPR spectra of TEMP/¹O₂. (e) Schematic diagram of FAP inhibition by O₂.

Moreover, upon preilluminating FAP with blue light in an aerobic environment, FAP showed a short activity retention time of 30 min, as compared to a FAP activity retention time of 180 min under anaerobic preillumination. This is ascribed to the anaerobic environment with reduced production of O₂˙⁻ and ¹O₂ (Fig. 3c and d), thereby reducing the oxidative damage caused by O₂˙⁻ and ¹O₂ to FAP protein and improving the photostability of FAP (Fig. 3e). However, an anaerobic environment can just inhibit the formation of radicals arising from the presence of oxygen. For radicals generated by the photosensitization of FAD, the elimination needs to be further explored. Moreover, the inhibition of enzyme activity through the destruction of amino acid structures in the active site by ROS remains merely a hypothesis, and this assumption requires further investigation.

Shifting illumination wavelengths provides a complementary stabilization strategy. Green light irradiation (vs. native blue light) extended FAP's operational lifetime by 12-fold during palmitic acid decarboxylation, enabling pentadecane yields to increase from 7.2 mM (1 h blue light plateau) to 14 mM over 36 h (Fig. 4a).³⁸ Meanwhile, for 240 min preillumination, the residual activity of FAP was 47.4% in green light, which was 5.8-fold higher as compared to blue light (Fig. 4b). The EPR test showed that green light reduced the formation of ROS, O₂˙⁻ and ¹O₂, thereby reducing the damage caused by ROS to amino acid residues of FAP (Fig. 4c and d). However, FAP catalytic activity under green light is relatively low, and enhancing enzyme activity through methods such as spectral combinations remains to be studied.

Fig. 4 Influence of blue and green light illumination on the photostability of FAP.³⁸ Reproduced from ref. 38 with permission from the Royal Society of Chemistry, copyright 2023. (a) Yield of alkane biofuel under green and blue light illumination. (b) Yield of alkane biofuel with the preillumination of green and blue light. (c) EPR spectrum of DMPO/O₂˙⁻ under green and blue light illumination. (d) EPR spectrum of TEMP/¹O₂ under green and blue light illumination.

Despite these interventions, optimizing green-light photocatalysis while maintaining strict anaerobic environments imposes rigorous bioprocessing constraints, and the enhancement efficacy toward FAP stability proves inherently limited.

3.1.2 Elimination of the produced radical. In the absence of intervention into the FAP's catalytic environment, radical formation becomes inherently uncontrollable, necessitating the adoption of targeted strategies to eliminate the generated radicals. Considering the amino acid-based radical, L-cysteine was reported to quench the ³FAD* radical, which is a photosensitization intermediate for amino acid-based radical production.³⁹ In an experimental study, FAP activity increased by 34.6% by adding L-cysteine in the photodecarboxylation process as compared to the control.³⁶ For elimination of ROS, researchers designed various natural enzyme-like nanomaterials for radical scavenging, which were inspired by natural antioxidant enzymes (i.e., catalase [CAT] and superoxide dismutase [SOD]).⁴⁰ Among them, O₂˙⁻ can be converted to H₂O₂ and O₂ molecules by the action of SOD (Fig. 5a). Then, H₂O₂ can be detoxified by decomposition of CAT and converted to H₂O and O₂ molecules.⁴¹ Thus, such enzyme-like nanomaterials, further called nanozymes, mimic natural enzymes and their mechanisms to convert ROS into non-toxic products. As compared to natural enzymes, nanozymes exhibit higher stability and robustness over a wide range of temperatures and pH values, lower cost, and higher activity in ROS scavenging. These features make nanozymes more suitable for the radical scavenging applications.⁴²

Fig. 5 Strategies for enhancing the stability of FAP. (a) The illustration of CAT-like and SOD-like nanozymes for radical scavenging.⁴⁴ Reproduced from ref. 44 with permission from Elsevier, copyright 2024. (b) The immobilization of enzyme by NiFe₂O₄ to improve the tolerance to organic solvents.⁴⁶ Reproduced from ref. 46 with permission from Elsevier, copyright 2023. (c) Interfacial enzymatic catalysis of fatty acid in a Pickering droplet.⁴⁷ Reproduced from ref. 47 with permission from American Chemical Society, copyright 2016.

Nanozymes can be constructed using metal compounds, metal nanoparticles, nanoclusters, organic networks, and amorphous coordination polymers. For example, recent studies have demonstrated that cerium oxide-based nanomaterials possess SOD activity.⁴³ With the help of mixed valence states of Ce³⁺ and Ce⁴⁺, O₂˙⁻ can be converted to H₂O₂ and O₂ molecules in the Ce³⁺→ Ce⁴⁺→ Ce³⁺ regeneration process (eqn (1)–(4)).⁴⁴ The elimination of O₂˙⁻ by nanoceria was proven by EPR measurements. Moreover, nanoceria with a size of 3–5 nm exhibits a higher catalytic rate constant of 3.6 × 10⁹ M⁻¹ s⁻¹, as compared to native SOD with a rate constant value of 1.3–2.8 × 10⁹ M⁻¹ s⁻¹. According to Self and coworkers' report, nanoceria exhibits a CAT-like property at a low Ce³⁺/Ce⁴⁺ ratio, while exhibiting an SOD-like property at a high Ce³⁺/Ce⁴⁺ ratio.⁴⁵ Therefore, by controlling the ratio of Ce³⁺/Ce⁴⁺, the addition of nanoceria would contribute to the ROS elimination in FAP catalysis.

2Ce³⁺ + H₂O₂ + 2H⁺ → 2H₂O + 2Ce⁴⁺ (1)

Ce⁴⁺ + H₂O₂ → 2Ce³⁺ + 2H⁺ + HOO˙ (2)

Ce⁴⁺ + HOO˙ → Ce³⁺ + O₂ + H⁺ (3)

2Ce⁴⁺ + H₂O₂ → 2Ce³⁺ + O₂ + 2H⁺ (4)

Inspired by the metal catalytic center of SOD, other nanozymes, such as Mn₃O₄ nanoparticles (NPs), iron oxide NPs, CuZn carbon dots, also showed the ability to quench O₂˙⁻.⁴³ Additionally, Pt, Au, Co, Mo, and V and their oxides and alloys show SOD-like activity.⁴⁸ Interestingly, carbon-based nanomaterials, such as fullerene, carbon nanotubes (CNTs) and graphene, show SOD mimetic activity. Choi and co-workers found that polyhydroxylated fullerenes exhibit SOD mimetic activity by forming non-harmful adducts.⁴⁹ The number and distribution of carboxyl groups on the fullerene ball significantly determine the activity of ROS elimination. The alloy nanostructures of Au, Ag, Pt, Pd, Co, Cu, Mn, Fe, etc. were recently found to possess multiple enzyme-mimetic activities.⁵⁰ For example, Au@Pt (an alloy material) exhibits SOD and CAT mimetic activity. Nevertheless, the application of nanozymes in FAP photo-driven catalysis for ROS elimination remains at an early stage and has not yet been implemented. The biocompatibility of nanozymes, the effect on light transfer in the catalytic reactor, and involvement in the FAP photocycle should be investigated for further applications.

3.2 The improvement of enzyme tolerance to organic solvents

3.2.1 Immobilization of FAP. Under natural conditions, the conformational structure of enzyme molecules is prone to disruption by organic solvents in the reaction system, leading to a reduction in enzymatic catalytic activity.⁵¹ Additionally, enzymes in their free state are difficult to separate from reaction products, which limits their recovery and reuse, thereby increasing industrial production costs.⁵² By immobilizing enzymes onto carriers through adsorption, entrapment, covalent binding, or cross-linking, enzyme molecules can be stabilized within the reaction system.⁵³ This enhances the stability of the enzyme and enables continuous catalytic processes.

Simić et al. selected beads of Eupergit C250 L for immobilizing FAP onto the surface via covalent linkage as the beads allow selective binding of His-tagged FAP.⁵⁴ A FAP protein content of 7 μM was confirmed to immobilize onto beads, thereby achieving a reuse time of up to 8. Similarly, Kara et al. adopted EziG Amber beads to immobilize FAP.⁵⁵ The protein content of 3 mg mL⁻¹ was adsorbed onto beads from cell extract containing FAP. Moreover, in reusability research, immobilized FAP exhibited 52% of enzyme activity retention after continuous catalysis for 3780 min within 3 cycles. In comparison, the free state FAP enzyme inactivation just happened after 240 min. This result revealed that the photostability of FAP was significantly enhanced by immobilization. Li et al. reported that FAP was immobilized in magnetic nickel ferrite (NiFe₂O₄) nanoparticles for facile recovery by a simple procedure (Fig. 5b).⁴⁶ The residual enzyme activities of 79.7% and 68% at pH 9.0 and an organic solvent ratio of 60% were observed, respectively, which proved that the enhanced stability of FAP was achieved by this immobilization method, especially using a high organic solvent ratio.

The immobilization of FAP stabilizes the conformation of enzymes and improves the activity retention time (80–3780 min) and reusability of FAP compared to the free state of FAP in the batch system, which exhibited a short activity retention time (10–225 min) and could not be repeatedly used for fatty acid decarboxylation and biofuel production (Table S1).^{21,37,54,56–64} However, the conversion of fatty acid by immobilized FAP was found to be significantly decreased, which may be ascribed to the shading of the immobilization carrier from light. As a result, the energy production rate, which ranged from 2.6 to 136.7 kJ L⁻¹ h⁻¹ with the immobilized system (Table S1), shows no enhancement as compared to the batch system. Moreover, the low efficiency of immobilization necessitates substantial enzyme usage during the immobilization process, significantly impacting the economic feasibility of biofuel production.

3.2.2 Interfacial catalysis of FAP in Pickering emulsion. In previous studies, DMSO was used as the organic phase to dissolve fatty acids, and Tris–HCl buffer was adopted as the water phase to load FAP.²⁸ Due to mutual dissolution in water, DMSO inevitably came into contact with the enzyme in the water phase, which disrupted the structure of the enzyme and decreased the stability of FAP in biofuel production.⁶⁵ Focused on this issue, an interfacial catalysis or a two-phase catalysis system is developed to prevent enzymes in the water phase from the damage by the organic phase.⁶⁶ However, traditional two-phase catalysis is hampered by low efficiency of mass transfer, poor recyclability of the enzyme, and difficulty in product separation.⁶⁷

In recent years, Pickering emulsion has emerged as a platform for enzymatic catalysis, which holds the advantage of a huge interface area for catalysis, enhanced catalysis efficiency (CE), and simple product separation.^68,69 Yang's group developed a series of SiO₂-based colloidosome stabilized Pickering emulsions, which contained molecular catalysts and enzymes for chemical or biofuel production.^70–72 In a typical description, a nonpolar reactant located in the oil phase (continuous phase) flowed to the oil/water interphase, catalyzed by the enzyme in water droplets, and then the product flowed back to the oil phase (Fig. 5c). In such systems, enzymes work in the oil/water interphase, rather than entering into the oil phase for catalysis, which prevent enzymes from toxic effects of organic solvents. In the case of kinetic resolution of racemic esters to chiral alcohol, the ee values of the corresponding chiral alcohol and chiral ester exceeded 98%.⁴⁷ It can be observed that each milligram of CALB can resolve 34 g of ester in a 720 h running time, highlighting high reaction productivity and enzyme stability in the Pickering emulsion system.

Hang et al. reported polyethylene glycol (PEG)-based hydrogel microparticles (HMPs) as a stabilizer to form Pickering emulsion with adsorption of lipase and FAP onto the oil/water interphase for the enzymatic conversion of acylglycerols into biofuel.⁷³ In such a catalytic system, HMPs can stabilize the oil-in-water Pickering emulsion for at least 14 days, which enables light-driven cascade conversion of triacylglycerols to renewable hydrocarbons in a green approach. More importantly, compared with free lipase and FAP catalysis, the interfacial catalysis of enzymes in Pickering emulsion enhanced the relative activity of enzymes by nearly 90%. Such improvement was ascribed to the enzyme protection by Pickering emulsion interfacial interaction from a harsh and detrimental environment in the organic solvent.

3.2.3 Hydrophobic modification of FAP. Environmental conditions, such as pH and temperature, significantly affect the enzyme stability and activity in biocatalytic reactions. Under harsh conditions, especially in an organic solvent, the stability of the enzyme can be enhanced by protein modification.⁷⁴ For example, lipase prefers to be adsorbed onto a hydrophobic surface due to the existence of a lid.⁷⁵ Meanwhile, the hydrophobic pocket of lipase is prone to bind to hydrophobic substrate molecules in an organic solvent. The enhanced hydrophobicity of lipase can also improve the enzyme tolerance to organic solvents. Therefore, the hydrophobic modification of lipase could contribute to its activity and stability enhancement. Mortazavi et al. grafted a hydrophobic surfactant (cetrimonium bromide) onto lipase by ion-exchange.⁷⁵ Their results showed that the hydrophobic modification of lipase increased the thermostability of lipase. The relative activity of hydrophobic modification of lipase at 86.4% increased by 23% compared with free lipase at 50 °C. In the case of glucose oxidase (GOx), Xu et al. investigated the enzymatic bioactivity and stability of GOx after the modification via in situ hydrophobic polymerization.⁷⁶ The results showed that tert-butyl acrylate polymerized GOx retained 73–79% relative activity at 50 °C after 1 h, as compared to free GOx with no activity. Meanwhile, in the case of the enzymatic storage bioactivity, hydrophobic polymerized GOx retained a bioactivity of 37% after 30 days, compared to no bioactivity with native GOx. These results revealed that the stability of GOx was enhanced after hydrophobic modification.

FAP was proven to interact with the interface in phospholipid liposomes and preferred to be adsorbed onto the oil/water interface, which is similar to the characteristics of lipase.⁷⁷ Therefore, we can conclude that the surface hydrophobic modification of FAP would enhance the binding between the enzyme and hydrophobic fatty acids. Moreover, the stability of FAP would also be improved due to the enhancement of the dissolution of FAP in organic solvents. The relevant experiment will be carried out in our laboratory. Focused on the hydrophobic modification of FAP, the chemical and biological modifications are the rational methods. For example, the grafting of hydroxyl groups of polyethylene glycol (PEG), covalently conjugated to the lysine residues on the FAP surface through Michael addition, may increase the stability in the organic phase, similar to the chemical modification of lipase.⁷⁸ Considering the biological modification of FAP, the site-specific mutagenesis of hydrophilic amino acid (e.g., aspartic acid) into hydrophobic amino acid (e.g., phenylalanine) or the fusion of a hydrophobic peptide into the FAP gene for expression would be an effective attempt.⁷⁹

3.3 Directed evolution technology to enhance the stability of FAP

FAP stability can be enhanced through targeted mutagenesis, ancestral sequence reconstruction (ASR), and computational design. Chanquia et al. demonstrated that the Y466F mutation stabilizes productive conformations and improves selectivity for C18:0,⁸⁰ while the Y466T/P460A/G462I mutant, designed via the ‘Focused Rational Iterative Site-specific Mutagenesis’ (FRISM) strategy, increases productivity 6.3-fold.^81,82 ASR further reinforces stability, with the ANC1 mutant showing a 15 °C higher melting temperature than wild-type FAP.^83–85

Computational tools like FRESCO and Rosetta design accelerate the identification of thermostable variants, such as ANC1.^83,86 Comparative studies of FAP homologs, including McFAP from Micractinium conductrix, highlight industrial potential due to solvent tolerance and pH stability.^61,87 Zhong et al. developed enzyme immobilization strategies that improve substrate transport efficiency by 50%, while radical-based photoinactivation studies show that substrate binding prevents enzyme degradation under blue light.³³ Stability improvements via ASR and solvent engineering are further validated in phenolic acid decarboxylases.⁸⁸

4. Strategies to enhance the catalytic performance of FAP

4.1 Enzyme engineering to enhance the catalytic performance of FAP

Globally, over 2 billion tons of agricultural residues (such as crop straw, bagasse, corn cobs, etc.) are produced annually.⁹¹ These biogenic residues can yield approximately 300–600 million tons of short-chain fatty acids via anaerobic fermentation, with butyric acid accounting for 20%.^92,93 Therefore, the catalytic conversion of short-chain fatty acids into alkane biofuels via FAP holds significant potential and promising prospects. However, FAP exhibited lower enzyme activity for short- or medium-chain fatty acids (C4–C12 < 11%) compared to long-chain fatty acids (C16–C20 > 96%).^17,21 This is attributed to the less efficient stabilization of short hydrophobic chains in a narrow hydrophobic tunnel, observed by crystal form diffraction.²⁰ Zhang et al. first proposed that the use of decoy molecules in the FAP enzyme increased the conversion of short-chain fatty acids.²⁶ For example, in the presence of tridecane as a decoy molecule, the decarboxylation rate of CH₃–COOH was increased by 2-fold and the yield of CH₄ was increased by 3-fold. This is attributed to tridecane entering into substrate channeling of FAP and thereby improving the stabilization of CH₃–COOH. However, such a method showed modest enhancement in short- or medium-chain fatty acid decarboxylation, with the TON (C2–C8) ranging from 5 to 2034, which is far from industrial applications.

Enzyme engineering is considered a more efficient platform for enhancing FAP catalytic activity for short- or medium-chain fatty acids. Amer et al. documented the first butyric acid decarboxylation by the FAP mutant G462V.⁶² The key amino acid residue G462 was mutated to reduce the channel size and the distance between the fatty acid and FAD. As a result, compared to wild FAP with no activity on butyric acid, the G462V mutant could decarboxylate butyric acid into propane at a concentration of 3.41 mg L⁻¹. Xu et al. designed two mutants (I398L and G462A) based on the strategy of ‘Focused Rational Iterative Site-specific Mutagenesis’ (FRISM) (Fig. 6a), which exhibited 29- to 552-fold enhancement in catalytic efficiency (CE) for C3–C14 fatty acid decarboxylation, as compared to wild FAP.⁸² Ma et al. identified the putative FAP from Micractinium conductrix (McFAP), based on publicly available sequence databases, which was capable of decarboxylating short- and medium-chain fatty acids.⁶¹ Focused on the decarboxylation of C8 fatty acid by McFAP, the conversion of over 90% was achieved at a 50 mM substrate concentration.

Fig. 6 Strategies for enhancing the catalytic performance of FAP. (a) The application of the FRISM strategy for enhancing the decarboxylation of various fatty acids by variants.⁸⁹ Reproduced from ref. 89 with permission from Elsevier, copyright 2024. (b) Continuous fatty acid decarboxylation by immobilized photodecarboxylase in a membrane reactor.⁹⁰ Reproduced from ref. 90 with permission from Wiley, copyright 2024.

4.2 The design of a photoreactor to enhance the catalytic performance of FAP

The typical batch reactor was first used to investigate the photo-driven decarboxylation of fatty acids for biofuel production. Guo et al. optimized the reaction conditions, including light intensity and temperature, for the decarboxylation of a long-chain fatty acid (C16) in a batch reactor.⁵⁷ Ma et al. designed FAP mutant decarboxylate medium- and short-chain fatty acids based on the FRISM strategy for gasoline biofuel production.⁶¹ However, in the case of a batch reactor for biofuel production, a low energy production rate at 10.8–183.1 kJ L⁻¹ h⁻¹ (Table S1) was achieved with a high catalyst concentration added.^{21,56,57,59–61} Moreover, the FAP-based catalyst for biofuel production in the batch reactor could not be reused. Therefore, the low energy production rate and failure of continuous production in a batch reactor limited the industrial applications of FAP for biofuel production. Such an issue encourages researchers to design a high-efficiency photoreactor.

Duong et al. designed an internal intensified illumination reactor by replacing a conventional external illumination setup, significantly accelerating fatty acid decarboxylation.⁶³ In such a proposed internal illumination system, palmitic acid was fully converted to pentadecane biofuel in less than 20 min, corresponding to a 22-fold energy production rate compared with that of a traditional reactor. Meanwhile, the turnover frequency at 6500 h⁻¹ and the energy production rate at 389.3 kJ L⁻¹ h⁻¹ (Table S1) were achieved, respectively, which were also significantly higher than those obtained in the previously reported batch reactor. The light transfer enhancement is responsible for accelerated product formation. More light was scattered back into the reaction suspension rather than being lost by reflection.

Recently, continuous flow catalysis has been demonstrated to have high efficiency in chemical production. Simić et al. first proposed the strategy for transferring the photodecarboxylation of fatty acid to continuous flow. An alkane production rate of 26.9 mM h⁻¹ and an energy production rate of 240 kJ L⁻¹ h⁻¹ (Table S1) were obtained in the continuous flow system, respectively.⁵⁴ Wang et al. designed a membrane reactor by immobilizing FAP for continuous decarboxylation, which significantly increased the space–time yield of octane with 1.2 mmol L⁻¹ h⁻¹ (Fig. 6b).⁹⁰ Li et al. reduced the diameter of the microtube to 500 μm for the continuous decarboxylation of palmitic acid, in which a high turnover frequency of 19 186 and energy production rate of 533.5 kJ L⁻¹ h⁻¹ were achieved for biofuel production.³⁷ The energy production rate in a microfluidic continuous flow system was significantly higher than that in a conventional batch reactor. This is because (1) the microfluidic reactor enables low light attenuation compared with big-size batch reactors, which is ascribed to the light intensity decreasing exponentially with increasing light length according to the Beer–Lambert law; (2) the continuous flow system accelerates the product transfer for next photocycle generation, rather than staying in catalytic activity sites and blocking substrate molecules from entering the enzyme pocket. Therefore, a microfluidic continuous flow system could reduce the reaction time (no more than 35 min) for fatty acid decarboxylation, as compared to the batch reactor with a long reaction time (hundreds to thousands of minutes) (Table S1).

However, the scale-up of biofuel production by photo-driven decarboxylation of fatty acids faces multiple challenges. Firstly, the multiple microfluidic device unit undoubtedly caused the exponential attenuation of light intensity, which increased with the rise of the optical path. The implementation of embedded optical fibers and the series module unit could be considered as effective measures. Then, the microchannel may be clogged up in a continuous running state as long-chain fatty acids are likely to crystallize (<35 °C).⁹⁴ The static mixing unit and thermal management module (local temperature >40 °C) are required, though this increases the device cost. Moreover, the microchannel materials and the precise pump control unit make the cost of the microfluidic reactor 3–5 times that of a batch reactor.⁵⁴ According to the techno-economic analysis (TEA) by Amer et al., non-optimized technology was considered to be economically unviable due to high production cost. In a basic model of photodecarboxylation for propane biofuel production, the cost of 69.86 $ per kg propane far exceeds the acceptable range of market selling price (0.25–1.50 $ per kg of propane).⁶² The analysis of production cost reveals that the electricity cost of the device running and the input cost of raw materials (containing the enzyme preparation) account for up to 90%. Nevertheless, by analyzing the carbon footprint, the global warming potential (GWP) of HEFA and fossil jet fuel was found to be 53.2–65.3 gCO₂e per MJ and 81.1–94.8 gCO₂e per MJ, respectively.^95,96 In comparison, in the case of FAP-derived SAF, the GWP is significantly decreased to 16.8 gCO₂e per MJ. To achieve the goal of carbon-neutrality, the balance of the economic efficiency of FAP-derived SAF and the potential of decarbonization should be given priority consideration.

Table 1 summarizes the key mechanism, performance gain, limitation, and scalability of various strategies for enhancing FAP-derived biofuel production. To maximize the potential of this unique enzyme, cohesive framework combining pathway enhancement and bioprocess intensification must be implemented. FAP expression should be optimized by employing strong, tunable promoters to improve catalytic efficiency and stability under processing conditions. Furthermore, engineered strains must be exploited in advanced photobioreactor systems. The light delivery, which is often a limiting factor, can be enhanced by employing novel reactor designs that ensure optimal photon distribution. Process parameters, including light intensity, feeding strategy, and gas exchange, must be dynamically controlled. Future advancements are anticipated in the development of immobilized enzyme or whole-cell systems for continuous operation and the application of artificial intelligence for the predictive design of both enzymes and bioprocesses. Ultimately, such a synergistic approach will be crucial for transitioning FAP-based biofuel production from a promising concept to an industrially viable, carbon-neutral technology.

Table 1 Comprehensive comparison of enhancement strategies for FAP-derived biofuel production

Strategy	Key mechanism	Performance gain	Limitations	Ref.
Anaerobic operation	• O2 exclusion	• Pentadecane yield increased by 44.7%	• High cost of inert gas	36
	• Suppresses ROS formation		• Reactor sealing complexity
Green light illumination	• Lower-energy photons	• Yield: 14 mM@36 h	• Average reaction rate decreased	38
	• Suppressing ROS	• Residual activity: 47.4%@4 h
Nanozyme ROS scavenging	• SOD/CAT mimicry	• ROS decreased by 90%	• Biocompatibility risks	44
	• Converting O2^−/^1O2 to H2O	• Activity retention: 79%@4 h	• High synthesis cost
Enzyme immobilization	• Covalent binding to magnetic carriers	• Reuse >8 cycles	• Light scattering by carrier	90
	• Physical shielding	• Tolerance to the high volume ratio of cosolvent	• Activity loss
Interfacial catalysis	• Enzyme confinement at the oil/water interface	• Activity increased by 90%	• Limited light penetration	73
		• Organic solvent protection	• Complex phase separation
Protein engineering	• Substrate tunnel engineering	• Broad substrate range (C4–C22)	• Low expression yield of enzyme	83
	• Thermostabilization	• 15 °C higher melting temperature (Tm)	• High production cost
Microfluidic flow systems	• Enhanced mass/light transfer	• Energy production rate: 533.5 kJ L^−1 h^−1	• Clogging risk	37
	• Continuous operation	• TOF: 19 186 h^−1	• High capital cost

---

5. Conclusions and outlook

FAP-driven photodecarboxylation holds transformative potential for sustainable biofuel production, yet its practical deployment requires resolving inherent instabilities and catalytic inefficiencies. Key findings reveal that FAP inactivation stems primarily from photogenerated radicals and conformational disruption in organic solvents. The significant advancements include the following areas. (1) Radical suppression via anaerobic environments and green light reduces FAP photoinactivation. (2) The immobilization of FAP and catalysis in Pickering emulsion systems were proven to prevent the FAP protein from organic solvent damage, thereby enhancing the stability of FAP. (3) Engineering mutagenesis of FAP boosts short- and medium-chain fatty acid conversion. (4) The adoption of microfluidic systems was shown to enhance energy production efficiency via optimized light and mass transfer.

Beyond fuel production, FAP-catalyzed decarboxylation offers significant potential for synthesizing high-value chiral fine chemicals. As demonstrated in the deracemization of phosphinothricin (PPT), engineered FAP variants enable light-driven kinetic resolution with exceptional enantioselectivity (>95% ee), bypassing the need for cofactors or protective groups.⁹⁷ Furthermore, FAP's promiscuity allows decarboxylative functionalization of diverse α-functionalized carboxylic acids, including α-amino and α-hydroxy acids, for yielding enantiopure amines and alcohols, which are pivotal pharmaceutical intermediates.⁹⁸ The enzyme's ability to operate under mild, redox-neutral conditions and its compatibility with cascades (e.g., hydratase–FAP systems for chiral fatty alcohols) underscore its potential for sustainable synthesis of complex molecules.⁹⁹ For the removal of trans fatty acids in vegetable oil, Li et al. designed a V453E mutant and 99% conversion of elaidic acid was obtained.¹⁰⁰ Future efforts should focus on expanding FAP's substrate scope via directed evolution and leveraging its photophysical properties for spatiotemporal control in multi-step enzymatic syntheses.

However, critical gaps persist: the mechanistic interplay between FAD and amino acid radicals remains unclear, necessitating in situ spectroscopy studies. Nanozyme integration risks disrupting FAP's photocycle, demanding compatibility assessments. While microfluidic reactors show promise, scalability under continuous flow requires validation. The production potential of short-chain fatty acids derived from the fermentation of agricultural wastes and wastewater would be much higher than that of the long-chain fatty acids derived from waste oils. Despite the design of FAP mutants for short-chain fatty acid decarboxylation being promising, their conversion efficiencies still could not be satisfactory for industrial applications. Reducing the volume of the substrate-binding cavity and enhancing electrostatic interactions with short-chain fatty acids are crucial in addressing this issue.

Future work should prioritize combinatorial approaches involving directed evolution of FAP mutants to decarboxylate the short-chain fatty acids, nanozyme-functionalized carriers, and reactor modularity to achieve techno-economic viability. Additionally, life-cycle analysis should quantify the environmental benefits of FAP-based systems versus conventional hydroprocessing. To bridge the gap between bench-scale insights and scalable FAP-mediated biofuel production, the real-time structural dynamics and photoreactivity of FAP should be monitored and optimized. Specific probes integrated with synchrotron radiation small-angle X-ray scattering could be applied to determine the dynamic status of FAP.¹⁰¹ Meanwhile, an enzyme activity prediction model can be established with the aid of emerging machine learning algorithms to improve the photoreactivity through automatically modulating light intensity or replenishing a fresh enzyme for industrial applications.¹⁰² By addressing these challenges, FAP can transition from proof-of-concept to industrial biofuel production, aligning with global decarbonization goals. This review underscores the need for interdisciplinary innovation to unlock FAP's full potential in renewable aviation fuel ecosystems.

Conflicts of interest

There are no conflicts to declare.

Data availability

No primary research results, software or code have been included and no new data were generated or analysed as part of this review.

Supplementary information (SI) is available. See DOI: https://doi.org/10.1039/d5cc02448j.

Acknowledgements

This work was supported by the National Natural Science Foundation of China (no. 52376168 and 52021004), the Natural Science Foundation of Chongqing (no. CSTB2023NSCQ-JQX0005), and the Fundamental Research Funds for the Central Universities (no. 2024IAIS-ZD003).

References

1. O. C. Anika, S. G. Nnabuife, A. Bello, E. R. Okoroafor, B. Kuang and R. Villa, Prospects of low and zero-carbon renewable fuels in 1.5-degree net zero emission actualisation by 2050: a critical review, Carbon Capture Sci. Technol., 2022, 5, 100072.
2. M. R. Allen, D. J. Frame, P. Friedlingstein, N. P. Gillett, G. Grassi, J. M. Gregory, W. Hare, J. House, C. Huntingford, S. Jenkins, C. D. Jones, R. Knutti, J. A. Lowe, H. D. Matthews, M. Meinshausen, N. Meinshausen, G. P. Peters, G.-K. Plattner, S. Raper, J. Rogelj, P. A. Stott, S. Solomon, T. F. Stocker, A. J. Weaver and K. Zickfeld, Geological Net Zero and the need for disaggregated accounting for carbon sinks, Nature, 2025, 638, 343–350.
3. M. K. Awasthi, S. Sarsaiya, A. Patel, A. Juneja, R. P. Singh, B. Yan, S. K. Awasthi, A. Jain, T. Liu, Y. Duan, A. Pandey, Z. Zhang and M. J. Taherzadeh, Refining biomass residues for sustainable energy and bio-products: an assessment of technology, its importance, and strategic applications in circular bio-economy, Renewable Sustainable Energy Rev., 2020, 127, 109876.
4. A. B. Nasrin, A. A. Abdul Raman, N. A. Bukhari, M. A. Sukiran, A. Buthiyappan, V. Subramaniam, A. A. Aziz and S. K. Loh, A critical analysis on biogas production and utilisation potential from palm oil mill effluent, J. Cleaner Prod., 2022, 361, 132040.
5. Y. Liu, P. Cruz-Morales, A. Zargar, M. S. Belcher, B. Pang, E. Englund, Q. Dan, K. Yin and J. D. Keasling, Biofuels for a sustainable future, Cell, 2021, 184, 1636–1647.
6. J. Zhang, M. S. Webber, Y. Pu, Z. Li, X. Meng, M. L. Stone, B. Wei, X. Wang, S. Yuan, B. Klein, B. Seemala, C. E. Wyman, K. K. Ramasamy, M. Thorson, M. H. Langholtz, J. S. Heyne, A. Koishybay, S. Adhikari, S. Cao, A. Sutton, G. A. Tuskan, Y. Román-Leshkov, A. J. Ragauskas, T. Ling and B. H. Davison, Sustainable aviation fuels from biomass and biowaste via bio- and chemo-catalytic conversion: catalysis, process challenges, and opportunities, Green Energy Environ., 2024, 10, 1210–1234.
7. J. Yang, Z. Xin, Q. He, K. Corscadden and H. Niu, An overview on performance characteristics of bio-jet fuels, Fuel, 2019, 237, 916–936.
8. IATA, Connecting primary data from airlines to CO₂ emissions calculations, Available from https://www.iata.org/en/services/data/environment-sustainability/co2-connect/, (accessed Mach 8, 2025).
9. IEA, Aviation – Tracking report, Available from https://www.iea.org/energy-system/transport/aviation, (accessed March 2, 2025).
10. W. Grimme and M. Braun, Estimation of potential hydrogen demand and CO₂ mitigation in global passenger air transport by the year 2050, Transp. Res. Proc., 2022, 65, 24–33.
11. IATA, Sustainability: SAF outlook and SAF registry, Available from https://www.iata.org/en/iata-repository/pressroom/presentations/sustainability-saf-outlook-registry-gmd-2024/, (accessed March 6, 2025).
12. C. Bergero, G. Gosnell, D. Gielen, S. Kang, M. Bazilian and S. J. Davis, Pathways to net-zero emissions from aviation, Nat. Sustain., 2023, 6, 404–414.
13. D. R. Vardon, B. J. Sherbacow, K. Guan, J. S. Heyne and Z. Abdullah, Realizing “net-zero-carbon” sustainable aviation fuel, Joule, 2022, 6, 16–21.
14. N. A. Huq, G. R. Hafenstine, X. Huo, H. Nguyen, S. M. Tifft, D. R. Conklin, D. Stück, J. Stunkel, Z. Yang, J. S. Heyne, M. R. Wiatrowski, Y. Zhang, L. Tao, J. Zhu, C. S. McEnally, E. D. Christensen, C. Hays, K. M. Van Allsburg, K. A. Unocic, H. M. Meyer, Z. Abdullah and D. R. Vardon, Toward net-zero sustainable aviation fuel with wet waste–derived volatile fatty acids, Proc. Natl. Acad. Sci. U. S. A., 2021, 118, e2023008118.
15. M. L. Stone, M. S. Webber, W. P. Mounfield, III, D. C. Bell, E. Christensen, A. R. C. Morais, Y. Li, E. M. Anderson, J. S. Heyne, G. T. Beckham and Y. Román-Leshkov, Continuous hydrodeoxygenation of lignin to jet-range aromatic hydrocarbons, Joule, 2022, 6, 2324–2337.
16. IEA, Net zero roadmap: a global pathway to keep the 1.5 °C goal in reach, Available from https://www.iea.org/reports/net-zero-roadmap-a-global-pathway-to-keep-the-15-0c-goal-in-reach#overview, (accessed March 1, 2025).
17. X. Guo, A. Xia, W. Zhang, Y. Huang, X. Zhu, X. Zhu and Q. Liao, Photoenzymatic decarboxylation: a promising way to produce sustainable aviation fuels and fine chemicals, Bioresour. Technol., 2023, 367, 128232.
18. A. H. Bhatt, Y. Zhang, A. Milbrandt, E. Newes, K. Moriarty, B. Klein and L. Tao, Evaluation of performance variables to accelerate the deployment of sustainable aviation fuels at a regional scale, Energy Convers. Manage., 2023, 275, 116441.
19. M. Wang, R. Dewil, K. Maniatis, J. Wheeldon, T. Tan, J. Baeyens and Y. Fang, Biomass-derived aviation fuels: challenges and perspective, Prog. Energy Combust. Sci., 2019, 74, 31–49.
20. D. Sorigué, B. Légeret, S. Cuiné, S. Blangy, S. Moulin, E. Billon, P. Richaud, S. Brugière, Y. Couté, D. Nurizzo, P. Müller, K. Brettel, D. Pignol, P. Arnoux, Y. Li-Beisson, G. Peltier and F. Beisson, An algal photoenzyme converts fatty acids to hydrocarbons, Science, 2017, 357, 903–907.
21. M. M. E. Huijbers, W. Zhang, F. Tonin and F. Hollmann, Light-driven enzymatic decarboxylation of fatty acids, Angew. Chem., Int. Ed., 2018, 57, 13648–13651.
22. D. Sorigué, K. Hadjidemetriou, S. Blangy, G. Gotthard, A. Bonvalet, N. Coquelle, P. Samire, A. Aleksandrov, L. Antonucci, A. Benachir, S. Boutet, M. Byrdin, M. Cammarata, S. Carbajo, S. Cuiné, R. B. Doak, L. Foucar, A. Gorel, M. Grünbein, E. Hartmann, R. Hienerwadel, M. Hilpert, M. Kloos, T. J. Lane, B. Légeret, P. Legrand, Y. Li-Beisson, S. L. Y. Moulin, D. Nurizzo, G. Peltier, G. Schirò, R. L. Shoeman, M. Sliwa, X. Solinas, B. Zhuang, T. R. M. Barends, J. P. Colletier, M. Joffre, A. Royant, C. Berthomieu, M. Weik, T. Domratcheva, K. Brettel, M. H. Vos, I. Schlichting, P. Arnoux, P. Müller and F. Beisson, Mechanism and dynamics of fatty acid photodecarboxylase, Science, 2021, 372, eabd5687.
23. P. Hong, L.-C. Ning, A.-A. Wu, K. Tan and X. Lu, Insights into the mechanism of fatty acid photodecarboxylase: a theoretical investigation, Chem. Phys. Lett., 2021, 771, 138550.
24. G. Londi, G. Salvadori, P. Mazzeo, L. Cupellini and B. Mennucci, Protein-driven electron-transfer process in a fatty acid photodecarboxylase, JACS Au, 2025, 5, 158–168.
25. D. J. Heyes, B. Lakavath, S. J. O. Hardman, M. Sakuma, T. M. Hedison and N. S. Scrutton, Photochemical Mechanism of Light-Driven Fatty Acid Photodecarboxylase, ACS Catal., 2020, 10, 6691–6696.
26. W. Zhang, M. Ma, M. M. E. Huijbers, G. A. Filonenko, E. A. Pidko, M. van Schie, S. de Boer, B. O. Burek, J. Z. Bloh, W. J. H. van Berkel, W. A. Smith and F. Hollmann, Hydrocarbon Synthesis via Photoenzymatic Decarboxylation of Carboxylic Acids, J. Am. Chem. Soc., 2019, 141, 3116–3120.
27. A. Aleksandrov, A. Bonvalet, P. Müller, D. Sorigué, F. Beisson, L. Antonucci, X. Solinas, M. Joffre and M. H. Vos, Catalytic Mechanism of Fatty Acid Photodecarboxylase: On the Detection and Stability of the Initial Carbonyloxy Radical Intermediate, Angew. Chem., Int. Ed., 2024, 63, e202401376.
28. T. M. Hedison, D. J. Heyes and N. S. Scrutton, Making molecules with photodecarboxylases: a great start or a false dawn?, Curr. Res. Chem. Biol., 2022, 2, 100017.
29. B.-S. Chen, Y.-Y. Zeng, L. Liu, L. Chen, P. Duan, R. Luque, R. Ge and W. Zhang, Advances in catalytic decarboxylation of bioderived fatty acids to diesel-range alkanes, Renewable Sustainable Energy Rev., 2022, 158, 112178.
30. W. Z. Ng, E.-S. Chan, W. Gourich, C. W. Ooi, B. T. Tey and C. P. Song, Perspective on enzymatic production of renewable hydrocarbon fuel using algal fatty acid photodecarboxylase from Chlorella variabilis NC64A: potentials and limitations, Renewable Sustainable Energy Rev., 2023, 184, 113548.
31. Y. Sui, X. Guo, R. Zhou, Z. Fu, Y. Chai, A. Xia and W. Zhao, Photoenzymatic decarboxylation to produce hydrocarbon fuels: a critical review, Mol. Biotechnol., 2023, 67, 2866–2880.
32. R. Wu, X. Li, L. Wang and D. Zhong, Ultrafast dynamics and catalytic mechanism of fatty acid photodecarboxylase, Angew. Chem., Int. Ed., 2022, 61, e202209180.
33. B. Lakavath, T. M. Hedison, D. J. Heyes, M. Shanmugam, M. Sakuma, R. Hoeven, V. Tilakaratna and N. S. Scrutton, Radical-based photoinactivation of fatty acid photodecarboxylases, Anal. Biochem., 2020, 600, 113749.
34. C. Xue, G. Li, Q. Zheng, X. Gu, Q. Shi, Y. Su, Q. Chu, X. Yuan, Z. Bao, J. Lu and L. Li, Tryptophan metabolism in health and disease, Cell Metab., 2023, 35, 1304–1326.
35. Q. Huang, Q. Huang, R. A. Pinto, K. Griebenow, R. Schweitzer-Stenner and W. J. Weber, Inactivation of horseradish peroxidase by phenoxyl radical attack, J. Am. Chem. Soc., 2005, 127, 1431–1437.
36. X. Guo, A. Xia, W. Zhang, F. Li, Y. Huang, X. Zhu, X. Zhu and Q. Liao, Anaerobic environment as an efficient approach to improve the photostability of fatty acid photodecarboxylase, Chin. Chem. Lett., 2023, 34, 107875.
37. F. Li, A. Xia, X. Guo, Y. Huang, X. Zhu, W. Zhang, R. Chen and Q. Liao, Photo-driven enzymatic decarboxylation of fatty acids for bio-aviation fuels production in a continuous microfluidic reactor, Renewable Sustainable Energy Rev., 2023, 183, 113507.
38. A. Xia, X. Guo, Y. Chai, W. Zhang, Y. Huang, X. Zhu, X. Zhu and Q. Liao, Green light enhanced the photostability and catalytic performance of fatty acid photodecarboxylase, Chem. Commun., 2023, 59, 6674–6677.
39. D. R. Cardoso, P. Homem-de-Mello, K. Olsen, A. B. F. da Silva, D. W. Franco and L. H. Skibsted, Deactivation of triplet-excited riboflavin by purine derivatives: important role of uric acid in light-induced oxidation of milk sensitized by riboflavin, J. Agric. Food Chem., 2005, 53, 3679–3684.
40. S. Zhang, J. Chen, M.-L. Lian, W.-S. Yang and X. Chen, An engineered, self-propelled nanozyme as reactive oxygen species scavenger, Chem. Eng. J., 2022, 446, 136794.
41. Y. Dai, Y. Ding and L. Li, Nanozymes for regulation of reactive oxygen species and disease therapy, Chin. Chem. Lett., 2021, 32, 2715–2728.
42. Y. Liu, Y. Cheng, H. Zhang, M. Zhou, Y. Yu, S. Lin, B. Jiang, X. Zhao, L. Miao, C.-W. Wei, Q. Liu, Y.-W. Lin, Y. Du, C. J. Butch and H. Wei, Integrated cascade nanozyme catalyzes in vivo ROS scavenging for anti-inflammatory therapy, Sci. Adv., 2020, 6, eabb2695.
43. H. Wei and E. Wang, Nanomaterials with enzyme-like characteristics (nanozymes): next-generation artificial enzymes, Chem. Soc. Rev., 2013, 42, 6060–6093.
44. S. Zhou, H. Cai, X. He, Z. Tang and S. Lu, Enzyme-mimetic antioxidant nanomaterials for ROS scavenging: design, classification, and biological applications, Coord. Chem. Rev., 2024, 500, 215536.
45. C. Korsvik, S. Patil, S. Seal and W. T. Self, Superoxide dismutase mimetic properties exhibited by vacancy engineered ceria nanoparticles, Chem. Commun., 2007, 1056–1058,  10.1039/B615134E.
46. F. Li, A. Xia, X. Guo, Y. Huang, X. Zhu, X. Zhu and Q. Liao, Immobilization of fatty acid photodecarboxylase in magnetic nickel ferrite nanoparticle, Bioresour. Technol., 2023, 385, 129374.
47. M. Zhang, L. Wei, H. Chen, Z. Du, B. P. Binks and H. Yang, Compartmentalized droplets for continuous flow liquid–liquid interface catalysis, J. Am. Chem. Soc., 2016, 138, 10173–10183.
48. S. Cao, Y. Long, S. Xiao, Y. Deng, L. Ma, M. Adeli, L. Qiu, C. Cheng and C. Zhao, Reactive oxygen nanobiocatalysts: activity-mechanism disclosures, catalytic center evolutions, and changing states, Chem. Soc. Rev., 2023, 52, 6838–6881.
49. L. L. Dugan, J. K. Gabrielsen, S. P. Yu, T.-S. Lin and D. W. Choi, Buckminsterfullerenol free radical scavengers reduce excitotoxic and apoptotic death of cultured cortical neurons, Neurobiol. Dis., 1996, 3, 129–135.
50. X. Jia, E. Wang and J. Wang, Rational design of nanozymes for engineered cascade catalytic cancer therapy, Chem. Rev., 2025, 125, 2908–2952.
51. K. N. Ingenbosch, J. C. Vieyto-Nuñez, Y. B. Ruiz-Blanco, C. Mayer, K. Hoffmann-Jacobsen and E. Sanchez-Garcia, Effect of organic solvents on the structure and activity of a minimal lipase, J. Org. Chem., 2022, 87, 1669–1678.
52. J. Liang, J. Ruan, B. Njegic, A. Rawal, J. Scott, J. Xu, C. Boyer and K. Liang, Insight into bioactivity of in-situ trapped enzyme-covalent-organic frameworks, Angew. Chem., Int. Ed., 2023, 62, e202303001.
53. F. Secundo, Conformational changes of enzymes upon immobilisation, Chem. Soc. Rev., 2013, 42, 6250–6261.
54. S. Simić, M. Jakštaitė, W. T. S. Huck, C. K. Winkler and W. Kroutil, Strategies for transferring photobiocatalysis to continuous flow exemplified by photodecarboxylation of fatty acids, ACS Catal., 2022, 12, 14040–14049.
55. S. N. Chanquia, F. V. Benfeldt, N. Petrovai, P. Santner, F. Hollmann, B. E. Eser and S. Kara, Immobilization and application of fatty acid photodecarboxylase in deep eutectic solvents, ChemBioChem, 2022, 23, e202200482.
56. Y. Ma, X. Zhang, W. Zhang, P. Li, Y. Li, F. Hollmann and Y. Wang, Photoenzymatic production of next generation biofuels from natural triglycerides combining a hydrolase and a photodecarboxylase, ChemPhotoChem, 2020, 4, 39–44.
57. X. Guo, A. Xia, F. Li, Y. Huang, X. Zhu, W. Zhang, X. Zhu and Q. Liao, Photoenzymatic decarboxylation to produce renewable hydrocarbon fuels: a comparison between whole-cell and broken-cell biocatalysts, Energy Convers. Manage., 2022, 255, 115311.
58. C. K. Winkler, S. Simić, V. Jurkaš, S. Bierbaumer, L. Schmermund, S. Poschenrieder, S. A. Berger, E. Kulterer, R. Kourist and W. Kroutil, Accelerated reaction engineering of photo(bio)catalytic reactions through parallelization with an open-source photoreactor, ChemPhotoChem, 2021, 5, 957–965.
59. Y.-Y. Zeng, L. Liu, B.-S. Chen and W. Zhang, Light-driven enzymatic decarboxylation of dicarboxylic acids, ChemistryOpen, 2021, 10, 553–559.
60. W. Xu, K. Mou, H. Zhou, J. Xu and Q. Wu, Transformation of triolein to biogasoline by photo-chemo-biocatalysis, Green Chem., 2022, 24, 6589–6598.
61. Y. Ma, X. Zhong, B. Wu, D. Lan, H. Zhang, F. Hollmann and Y. Wang, A photodecarboxylase from Micractinium conductrix active on medium and short-chain fatty acids, Chin. J. Catal., 2023, 44, 160–170.
62. M. Amer, E. Z. Wojcik, C. Sun, R. Hoeven, J. M. X. Hughes, M. Faulkner, I. S. Yunus, S. Tait, L. O. Johannissen, S. J. O. Hardman, D. J. Heyes, G.-Q. Chen, M. H. Smith, P. R. Jones, H. S. Toogood and N. S. Scrutton, Low carbon strategies for sustainable bio-alkane gas production and renewable energy, Energy Environ. Sci., 2020, 13, 1818–1831.
63. H. T. Duong, Y. Wu, A. Sutor, B. O. Burek, F. Hollmann and J. Z. Bloh, Intensification of photobiocatalytic decarboxylation of fatty acids for the production of biodiesel, ChemSusChem, 2021, 14, 1053–1056.
64. L. A. D. Benincá, A. S. França, G. C. Brêda, R. A. C. Leão, R. V. Almeida, F. Hollmann and R. O. M. A. de Souza, Continuous-flow CvFAP photodecarboxylation of palmitic acid under environmentally friendly conditions, Mol. Catal., 2022, 528, 112469.
65. F. Cheng, M.-Y. Li, D.-J. Wei, X.-J. Zhang, D.-X. Jia, Z.-Q. Liu and Y.-G. Zheng, Enabling biocatalysis in high-concentration organic cosolvent by enzyme gate engineering, Biotechnol. Bioeng., 2022, 119, 845–856.
66. Y. Zhao, Y. Kondo, Y. Kuwahara, K. Mori and H. Yamashita, Two-phase reaction system for efficient photocatalytic production of hydrogen peroxide, Appl. Catal., B, 2024, 351, 123945.
67. L. Chen, S. Zhang, X. Liu and X. Ge, Recent advances in water-mediated multiphase catalysis, Curr. Opin. Colloid Interface Sci., 2023, 65, 101691.
68. X. Guan, Z. Wan and T. Ngai, Reaction-controlled microgel-lipase co-stabilized compartmentalized emulsion for recyclable biodiesel production, Aggregate, 2023, 4, e300.
69. L. Ni, C. Yu, Q. Wei, D. Liu and J. Qiu, Pickering emulsion catalysis: interfacial chemistry, catalyst design, challenges, and perspectives, Angew. Chem., Int. Ed., 2022, 61, e202115885.
70. K. Li, H. Zou, R. Ettelaie, J. Zhang and H. Yang, Spatial localization of two enzymes at Pickering emulsion droplet interfaces for cascade reactions, Angew. Chem., Int. Ed., 2023, 62, e202300794.
71. H. Zou, H. Shi, S. Hao, Y. Hao, J. Yang, X. Tian and H. Yang, Boosting catalytic selectivity through a precise spatial control of catalysts at Pickering droplet interfaces, J. Am. Chem. Soc., 2023, 145, 2511–2522.
72. W. Wei, R. Ettelaie, X. Zhang, M. Fan, Y. Dong, Z. Li and H. Yang, Co-compartmentalization of enzymes and cofactors within Pickering emulsion droplets for continuous-flow catalysis, Angew. Chem., Int. Ed., 2022, 61, e202211912.
73. Y. Han, H. Jiang, C. Huang, X. Wu, Y. Ouyang, H. Chen, D. Lan, Y. Wang, B. Zheng and J. Xia, Enzymatic interfacial conversion of acylglycerols in Pickering emulsions stabilized by hydrogel microparticles, J. Colloid Interface Sci., 2024, 661, 228–236.
74. C. Bai, H. Jiang, M. Deng, Q. Gong, P. Li and W. Liu, An efficient hydrophobic modification strategy for enhancing collagen stability: effect of lauric acidic N-hydroxysulfosuccinimide ester on the structure and properties of fish collagen, Polym. Degrad. Stabil., 2025, 232, 111127.
75. S. Mortazavi and H. Aghaei, Make proper surfaces for immobilization of enzymes: immobilization of lipase and α-amylase on modified Na-sepiolite, Int. J. Biol. Macromol., 2020, 164, 1–12.
76. G. Xu, Y. Xu, A. Li, T. Chen and J. Liu, Enzymatic bioactivity investigation of glucose oxidase modified with hydrophilic or hydrophobic polymers via in situ RAFT polymerization, J. Polym. Sci., Part A: Polym. Chem., 2017, 55, 1289–1293.
77. C. Aselmeyer, B. Légeret, A. Bénarouche, D. Sorigué, G. Parsiegla, F. Beisson and F. Carrière, Fatty acid photodecarboxylase is an interfacial enzyme that binds to lipid–water interfaces to access its insoluble substrate, Biochemistry, 2021, 60, 3200–3212.
78. L. Shi, J. Zhang, M. Zhao, S. Tang, X. Cheng, W. Zhang, W. Li, X. Liu, H. Peng and Q. Wang, Effects of polyethylene glycol on the surface of nanoparticles for targeted drug delivery, Nanoscale, 2021, 13, 10748–10764.
79. F. Zha, J. Rao and B. Chen, Modification of pulse proteins for improved functionality and flavor profile: a comprehensive review, Compr. Rev. Food Sci. Food Saf., 2021, 20, 3036–3060.
80. S. N. Chanquia, J. P. Bittner, P. Santner, L. K. Szabó, J. S. Madsen, M. L. Øhlenschlæger, A. G. Sarvari, A. Merrild, K. G. Fonss, D. Jaron, L. Lutz, S. Kara and B. E. Eser, Active-site mutagenesis of fatty acid photodecarboxylase: experimental and computational insight into substrate chain-length specificity, ACS Catal., 2024, 14, 15837–15849.
81. Z. Qin, Y. Zhou, Z. Li, M. Höhne, U. T. Bornscheuer and S. Wu, Production of biobased ethylbenzene by cascade biocatalysis with an engineered photodecarboxylase, Angew. Chem., Int. Ed., 2024, 63, e202314566.
82. W. Xu, Y. Chen, D. Li, Z. Wang, J. Xu and Q. Wu, Rational design of fatty acid photodecarboxylase enables the efficient decarboxylation of medium- and short-chain fatty acids for the production of gasoline bio-alkanes, Mol. Catal., 2022, 524, 112261.
83. Y. Sun, E. Calderini and R. Kourist, A reconstructed common ancestor of the fatty acid photo-decarboxylase clade shows photo-decarboxylation activity and increased thermostability, ChemBioChem, 2021, 22, 1833–1840.
84. R. Furukawa, W. Toma, K. Yamazaki and S. Akanuma, Ancestral sequence reconstruction produces thermally stable enzymes with mesophilic enzyme-like catalytic properties, Sci. Rep., 2020, 10, 15493.
85. V. A. Risso, J. M. Sanchez-Ruiz and S. B. Ozkan, Biotechnological and protein-engineering implications of ancestral protein resurrection, Curr. Opin. Struct. Biol., 2018, 51, 106–115.
86. L. Zhou, C. Tao, X. Shen, X. Sun, J. Wang and Q. Yuan, Unlocking the potential of enzyme engineering via rational computational design strategies, Biotechnol. Adv., 2024, 73, 108376.
87. Y. Zeng, X. Yin, L. Liu, W. Zhang and B. Chen, Comparative characterization and physiological function of putative fatty acid photodecarboxylases, Mol. Catal., 2022, 532, 112717.
88. K. Myrtollari, E. Calderini, D. Kracher, T. Schöngaßner, S. Galušić, A. Slavica, A. Taden, D. Mokos, A. Schrüfer, G. Wirnsberger, K. Gruber, B. Daniel and R. Kourist, Stability increase of phenolic acid decarboxylase by a combination of protein and solvent engineering unlocks applications at elevated temperatures, ACS Sustainable Chem. Eng., 2024, 12, 3575–3584.
89. Y. Bao, Y. Xu and X. Huang, Focused rational iterative site-specific mutagenesis (FRISM): a powerful method for enzyme engineering, Mol. Catal., 2024, 553, 113755.
90. J. Zhou, F. Hollmann, Q. He, W. Chen, Y. Ma and Y. Wang, Continuous fatty acid decarboxylation using an immobilized photodecarboxylase in a membrane reactor, ChemSusChem, 2024, 17, e202301326.
91. G. Ginni, S. Kavitha, R. K. Yukesh, B. K. Shashi, S. K. Adish, M. Rajkumar, K. Gopalakrishnan, P. Arivalagan, T. L. C. Nguyen and J. B. Rajesh, Valorization of agricultural residues: different biorefinery routes, J. Environ. Chem. Eng., 2021, 9, 105435.
92. Q. Xu, X. Liu, D. Wang, Y. Liu, Q. Wang, B.-J. Ni, X. Li, Q. Yang and H. Li, Enhanced short-chain fatty acids production from waste activated sludge by sophorolipid: performance, mechanism, and implication, Bioresour. Technol., 2019, 284, 456–465.
93. W. Fang, P. Zhang, T. Zhang, D. C. Requeson and M. Poser, Upgrading volatile fatty acids production through anaerobic co-fermentation of mushroom residue and sewage sludge: performance evaluation and kinetic analysis, J. Environ. Manage., 2019, 241, 612–618.
94. F. Li, A. Xia, X. Guo, W. Zhang, Y. Huang, X. Zhu, X. Zhu and Q. Liao, Continuous hydrocarbon fuels production by photoenzymatic decarboxylation of free fatty acids from waste oils, J. Environ. Chem. Eng., 2023, 11, 110748.
95. R. S. Capaz, J. A. Posada, P. Osseweijer and J. E. A. Seabra, The carbon footprint of alternative jet fuels produced in Brazil: exploring different approaches, Resour., Conserv. Recycl., 2021, 166, 105260.
96. L. Jing, H. M. El-Houjeiri, J.-C. Monfort, J. Littlefield, A. Al-Qahtani, Y. Dixit, R. L. Speth, A. R. Brandt, M. S. Masnadi, H. L. MacLean, W. Peltier, D. Gordon and J. A. Bergerson, Understanding variability in petroleum jet fuel life cycle greenhouse gas emissions to inform aviation decarbonization, Nat. Commun., 2022, 13, 7853.
97. F. Cheng, H. Li, D.-Y. Wu, J.-M. Li, Y. Fan, Y.-P. Xue and Y.-G. Zheng, Light-driven deracemization of phosphinothricin by engineered fatty acid photodecarboxylase on a gram scale, Green Chem., 2020, 22, 6815–6818.
98. J. Xu, Y. Hu, J. Fan, M. Arkin, D. Li, Y. Peng, W. Xu, X. Lin and Q. Wu, Light-driven kinetic resolution of α-functionalized carboxylic acids enabled by an engineered fatty acid photodecarboxylase, Angew. Chem., Int. Ed., 2019, 58, 8474–8478.
99. W. Zhang, J.-H. Lee, S. H. H. Younes, F. Tonin, P.-L. Hagedoorn, H. Pichler, Y. Baeg, J.-B. Park, R. Kourist and F. Hollmann, Photobiocatalytic synthesis of chiral secondary fatty alcohols from renewable unsaturated fatty acids, Nat. Commun., 2020, 11, 2258.
100. D. Li, T. Han, J. Xue, W. Xu, J. Xu and Q. Wu, Engineering fatty acid photodecarboxylase to enable highly selective decarboxylation of trans fatty acids, Angew. Chem., Int. Ed., 2021, 60, 20695–20699.
101. T. C. Pochapsky, A dynamic understanding of cytochrome P450 structure and function through solution NMR, Curr. Opin. Biotechnol, 2021, 69, 35–42.
102. S. Mazurenko, Z. Prokop and J. Damborsky, Machine Learning in enzyme engineering, ACS Catal., 2020, 10, 1210–1223.

---