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Lin, P.;  Yang, H.;  Nakata, E.;  Morii, T. Modulation of Enzyme Reactions on DNA Scaffold. Encyclopedia. Available online: https://encyclopedia.pub/entry/33194 (accessed on 19 June 2024).
Lin P,  Yang H,  Nakata E,  Morii T. Modulation of Enzyme Reactions on DNA Scaffold. Encyclopedia. Available at: https://encyclopedia.pub/entry/33194. Accessed June 19, 2024.
Lin, Peng, Hui Yang, Eiji Nakata, Takashi Morii. "Modulation of Enzyme Reactions on DNA Scaffold" Encyclopedia, https://encyclopedia.pub/entry/33194 (accessed June 19, 2024).
Lin, P.,  Yang, H.,  Nakata, E., & Morii, T. (2022, November 07). Modulation of Enzyme Reactions on DNA Scaffold. In Encyclopedia. https://encyclopedia.pub/entry/33194
Lin, Peng, et al. "Modulation of Enzyme Reactions on DNA Scaffold." Encyclopedia. Web. 07 November, 2022.
Modulation of Enzyme Reactions on DNA Scaffold
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Cells have developed intelligent systems to implement the complex and efficient enzyme cascade reactions via the strategies of organelles, bacterial microcompartments and enzyme complexes. The scaffolds such as the membrane or protein in the cell are believed to assist the co-localization of enzymes and enhance the enzymatic reactions. Inspired by nature, enzymes have been located on a wide variety of carriers, among which DNA scaffolds attract great interest for their programmability and addressability. Integrating these properties with the versatile DNA–protein conjugation methods enables the spatial arrangement of enzymes on the DNA scaffold with precise control over the interenzyme distance and enzyme stoichiometry.

DNA scaffold enzyme reaction catalytic enhancement

1. Introduction

Living organisms have evolved over millions of years to build a complex metabolic network containing thousands of enzymatic reactions for their survival [1]. Enzymes are spatially organized in the cell to implement specific sequential reactions via the strategies of compartmentalization [2]. The spatial organization of enzymes often relies on the specific scaffolds of proteins or the membrane to achieve the high efficiency and the specificity of enzymatic reactions [3]. Ribulose 1,5-bisphosphate carboxylase/oxygenase (RuBisCO) and carbonic anhydrase packed in the protein shell of carboxysome [4] and cytochrome P450 enzymes anchored on the membrane of endoplasmic reticulum [5] are the typical examples. In these compartments, the reactants in low concentration are believed to be effectively transferred through spatially arranged enzymes, thereby channeling metabolites to drive favorable reactions and preventing the toxic side reactions by intermediates [6]. It is the challenge to understand the efficient bioenergetic processes of nature and to construct human-engineered energy utilization systems [7].
To mimic and understand the natural systems, a wide variety of carriers has been built for the attachment of enzymes, as has been reviewed previously [8][9][10][11]. The further applications of the carriers such as protein, liposome or polymersome are challenged by the structural programmability of carriers and the spatial arrangement of enzymes [12]. These obstacles are tackled by means of DNA nanotechnology [13]. A typical example of DNA nanostructures, DNA origami, folds a long, single-stranded DNA into the predesigned, two-dimensional (2D) and three-dimensional (3D) DNA scaffolds with accurate addressability, providing the ideal templates for the assembly of enzymes [14][15]. Individual or multiple enzymes have been spatially arranged on the DNA scaffolds with precise control over the enzyme orientations, interenzyme distance and the stoichiometry of enzymes [16].
Interestingly, the catalytic enhancement of single type of enzyme assembled on the DNA scaffolds has been widely observed [17]. This phenomenon has been attributed to the substrate affinity to the negatively charged DNA scaffold surface by electrostatic interactions, lower local pH on the DNA scaffold surface, reduced adsorption of scaffolded enzymes on the reaction vessels or the ordered hydration layer attracted by the DNA surface [18]. However, the notion of whether these proposed mechanisms can apply in general for the catalytic enhancement of DNA-scaffolded enzymes remains controversial. Besides the single type of enzyme, the cascade reactions of multi-enzyme assembled on the DNA scaffold were also extensively studied [19][20]. While the close interenzyme distance, optimal spatial organization of enzymes and a confined DNA environment have been proposed to be the main factors enhancing the efficiency of enzyme cascade reactions on the DNA scaffold, the actual mechanisms remain to be elucidated [21].

2. Construction of the Artificial Enzymatic Reaction Systems Inspired by Nature

To implement the efficient enzyme cascade reactions for the biochemical transformation, cells utilize the strategy of compartmentalization by forming the membrane-bound organelles (e.g., mitochondria, chloroplasts and peroxisome), bacterial microcompartments and multi-enzyme complexes [22]. The spatial organization of enzymes on the specific scaffolds (e.g., proteins or membrane) exerts the high efficiency and specificity of cascade reactions by increasing the concentration of reactants, reducing the toxic intermediates and competing reactions and overcoming the unfavorable enzyme kinetics [23]. To mimic the natural systems, a wide ranges of materials have been applied to construct the artificial scaffolds for enzyme reactions in vitro [18].

2.1. The Spatial Organization of Enzymes in Nature

Substrate channeling is the transportation of the intermediates from one enzyme active site to the next without the release into solution [23]. The typical example of substrate channeling is tryptophan synthase. This α2β2 tetrameric enzyme complex catalyzes a two-step cascade reaction that converts indole-3-glycerol-phosphate and serine to tryptophan. As the intermediate, indole is directly transferred through the 25 Å long hydrophobic tunnel bridging the two active sites [24]. Such hydrophobic tunnel is also found in carbamoyl–phosphate synthetase that channels carbamates to the neighboring active site. The substrate channeling couples the first hydrolysis reaction of glutamine and the successive reactions to maintain the proper stoichiometry despite of the three-order-of magnitude difference in Km for NH3 in the carbamate synthesis and that for glutamine in the first reaction [25]. Instead of using hydrophobic tunnel, the malate dehydrogenase–citrate synthase (MDH–CS) complex utilizes the electrostatic guidance to channel the metabolite oxaloacetate to achieve a high flux though the MDH–CS pair by overcoming the unfavorable kinetics of MDH forward reaction in the citric acid cycle [26].
Metabolons feature the dynamic assembly and disassembly of the enzyme complexes [27]. These highly transient protein assemblies are believed to allow the direct channeling of the intermediates from one enzyme to the next consecutive enzyme in the metabolic pathway. Purinosome [28], dhurrin metabolon [29] and glucosome [30] are the typical examples. While the spatiotemporal enzyme assemblies are proposed to enhance the metabolic flux and biochemical transformation, the study of the spatial organization of metabolon remains challenging due to the transient and complex interactions of proteins involved [27]. In 2008, the human purinosome was first identified, in which the ten chemical steps of de novo purine biosynthesis (DNPB) were catalyzed by six enzymes. Two human enzymes involved in DNPB were fused to either a green fluorescent protein (GFP) or an orange fluorescent protein (OFP) and transiently expressed in the purine-rich or purine-depleted cells. The cytoplasmic clusters formed by these two enzymes were observed by fluorescence microscopy in purine-depleted cells. This provided the evidence for the formation of the enzyme complex “purinosome” [31]. Afterward, the protein–protein interactions of purinosome and the regulation of its formation were extensively studied [32]. In 2020, Pareek et al. [33] used metabolomics and in situ three-dimensional sub-micrometer chemical imaging of single cells by gas cluster ion beam secondary ion mass spectrometry (GCIB-SIMS) to directly visualize DNPB by purinosome. It was proposed that purinosome consisted of at least nine enzymes that functioned synergistically to increase the pathway flux. Moreover, these DNPB metabolons were hypothesized to locate proximal to the mitochondria. In a recent study, Pedley et al. [34] found the heat shock protein 90 kDa (Hsp90) would help to regulate the physical properties of the purinosome and maintain the liquid-like state inside HeLa cells. This finding provides novel insights into how the liquid–liquid phase separation drives the formation of metabolon in the human cell.
As the best-studied bacterial microcompartments, carboxysomes play a central role in the carbon concentrating mechanism [4]. Two enzymes, RuBisCO and carbonic anhydrase (CA), are packed in the polyhedral protein shells of carboxysome. CA converts bicarbonate (HCO3) to CO2, which is subsequently consumed by RuBisCO in the carbon fixing reaction. The diffusion of negatively charged HCO3 into the protein shell is promoted by the positively charged pores of carboxysome, while the concentration of uncharged CO2 is increased in the compartment, resulting in the enhanced reactivity of RuBisCO [4]. Moreover, RuBisCO molecules have been hypothesized to form the condensate by the liquid–liquid phase separation, which provides the understanding of carboxysome biogenesis. The function of the protein shell of carboxysome in RuBisCO condensation remains to be elucidated [35].

2.2. Enzymatic Reactions on Various Carriers

The reactions of individual or multienzymes have been conducted on a wide range of carriers, such as metal–organic frameworks (MOFs) [37], hydrogel [38], graphene oxide [39], liposome [40], polymersome [41], proteins [42] and DNA nanostructures [43]. These artificial scaffolds are suggested to enhance the enzyme stability, reusability or catalytic ability, expanding the applications of enzyme in different fields such as the biosynthesis of value-added chemicals. These scaffolds are also applied to mimic and understand the natural systems such as complex metabolic pathways or molecular transport.
Yoshimoto et al. [44] entrapped glucose oxidase (GOx) and catalase into the liposome membrane. The hydrogen peroxide (H2O2) produced by GOx in the glucose oxidation reaction inside the liposome was decomposed by catalase. A remarkable protection effect of the liposome membrane on catalase activity inside the liposome was observed. The presence of outer membrane protein F (OmpF) enhanced the transport of glucose molecules from the exterior of the liposomes to the interior and increased the enzyme activity of GOx by 17 times compared with that of GOx encapsulated in the liposome in the absence of OmpF. This system demonstrates the typical example of the application of liposome as an enzyme carrier. Klermund et al. [41] compartmentalized the three-step reaction in a polymersome to synthesize CMP-N-acetylneuraminic acid (CMP-Neu5Ac). N-Acetylneuraminate lyase (NAL) and CMP-sialic acid synthetase (CSS) were inserted into polymersome, while N-Acyl-D-glucosamine 2-epimerase (AGE) with allosteric activator ATP was encapsulated inside the polymersome. Channel proteins OmpF enabled a selective mass transport across the polymer membrane. The incorporation of OmpF into the membrane restrained the cross-inhibitions in enzyme cascade reactions. The overall throughput was enhanced 2.2-fold compared to the reaction by free enzymes.

3. Catalytic Enhancement of Single Type of Enzyme Assembled on the DNA Scaffold

While the soft materials show their own advantages and the potential as the enzyme scaffolds, the further applications of carriers such as liposome, polymersome or protein face the difficulties in controlling the enzyme loading positions and stoichiometry. Therefore, the scaffolds that can overcome these challenges are required for the enzyme assembly in vitro. Given the predominant advantages of structural programmability and accurate addressability, DNA nanostructures are considered as the ideal platforms for the assembly of enzymes [46]. The reactions of a single type of enzyme assembled on the DNA scaffolds and the previously proposed mechanisms for the catalytic enhancement of enzyme by the DNA microenvironment have been reviewed. Understanding the origins of enhanced activity of DNA-scaffolded enzymes will expand their practical applications.

3.1. DNA Origami Scaffold

The past two decades have witnessed the rapid development of DNA origami and its applications [47]. In 2006, Rothemund [14] created DNA origami that folds a long, circular, single-stranded DNA template (7-kilobase) into desired two-dimensional (2D) shapes with the aid of over 200 short oligonucleotides (staple strands). Nonperiodic structures, such as square, rectangle, star and smiley face, were obtained. Featuring preparation simplicity, structural programmability and high folding yield, DNA origami plays an important role in the development of structural DNA nanotechnology. In 2009, Douglas et al. [15] extended the DNA origami method to build custom three-dimensional structures formed as pleated layers of DNA helices in the honeycomb lattices, providing a general route to the construction of complex 3D DNA nanostructures. By using the computer-aided design software for DNA origami nanostructures such as caDNAno, the staple sequences for folding newly designed DNA nanostructures are easily generated [48][49].

3.2. “Favorable Microenvironment” Provided by DNA

The enzyme scaffolded by DNA structures often displays enhanced activity and stability over its free form; however, the actual mechanisms for the higher catalytic ability are still under debate [66]. As studied previously, Glettenberg et al. [67] covalently conjugated peroxidase to different DNA oligonucleotides (ODN). The ODN markedly influenced the catalytic of tethered-enzyme in a DNA sequence-dependent manner. This phenomenon was attributed to the interactions such as hydrogen bonding or electrostatic contacts between ODN and the heme-containing catalyst. Rudiuk et al. [68] conjugated β-lactamase with a branched DNA complex constructed by four λ DNA (48.5 kbp), which underwent a dramatic and reversible higher-order structural transition regulated by spermine (SPM4+) and NaCl. The enhanced catalytic activity of enzyme was attributed to the “favorable microenvironment” composed of the giant and ordered DNA molecules. Then an interesting and important question arises: What is “favorable DNA microenvironment”, and what is the chemistry behind it?

3.3. Protection Effect Derived from DNA Scaffold

3.4. Substrate Affinity to the DNA Scaffold

3.5. Ordered Hydration Layer on the DNA Scaffold Surface

Zhao et al. [73] have observed 3- to 10-fold activity enhancements of five different enzymes, GOx, HRP, glucose-6-phosphate dehydrogenase (G6pDH), malic dehydrogenase (MDH) and lactic dehydrogenase (LDH), individually encapsulated in DNA cages. It was hypothesized that enzymes were stabilized by the highly ordered, hydrogen-bonded water environment formed by the negatively charged DNA cage surface, in which the stabilization of the hydrophobic interactions of a folded protein was induced by the solvent entropy penalty upon protein unfolding. The activity of encapsulated G6pDH reduced to approx. 25% activity in the presence of 1 M NaCl. This was suggested that Na+ would shield the negative charge on the DNA surface and disrupt the surface-bound hydration layer. However, the high concentration of salts containing the cations such as Na+, K+ and NH4+ also strongly inhibited the activity of free G6pDH. The DNA hydration layer may play an important role in the modulation of enzyme reactions on the DNA scaffold, but the stabilization of proteins may not be the general mechanism for enhancing the activity of the DNA-scaffolded enzymes.

3.6. Local pH Environment

3.7. General Factors for the Catalytic Enhancement of DNA-Scaffolded Enzymes

As reported previously [76], two enzymes with different pH preferences, xylose reductase (XR) and xylitol dehydrogenase (XDH), were individually assembled on the fully open state of a 3D DNA scaffold through the modular adaptor method [77][78][79][80][81][82] in high loading yields. XR was genetically fused to the modular adaptor ZF-SNAP to obtain ZS-XR. The zif268 bound with the specific DNA sequence, while SNAP-tag would react with benzylguanine incorporated in the DNA sequence to form the covalent linkage [78]. Similarly, XDH was fused to the C-terminal of modular adaptor Halo-GCN4 to construct enzyme HG-XDH. The Halo-tag substrate 5-chlorohexane (CH) was incorporated in the GCN4-binding DNA sequence. The catalytic enhancements were observed for both the DNA-scaffolded ZS-XR (sXR) and scaffolded HG-XDH (sXDH) over the respective free enzyme. In the enzyme reactions, XR converted xylose with the cofactor NADH to xylitol, while XDH produced xylulose from xylitol using NAD+. Such neutral or net negative charge of their substrates and cofactors indicated that the surface–substrate or –cofactor electrostatic attractive interaction could not account for the increase in activities of assembled enzymes. Instead, the large scaffold with high packing density of DNA helices improved the enzyme stability and reduced the adsorption of scaffolded enzymes to the reaction vessels, which could partly contribute to the catalytic enhancement of DNA-scaffolded XR or XDH. To assess the local pH environment of DNA scaffold, a dual-emission ratiometric pH indicator SNARF derivative was loaded on the DNA scaffold either facing near the surface or locating 6.7 nm away from the surface, which corresponded to the distance between the enzyme and the surface of the DNA scaffold. The local pH near the surface of the DNA scaffold or near the enzyme loaded position in the reaction buffer (pH 7.0) was deduced to be 6.2 or 6.5. Such local pH shifts would result in 25% enhancement of the catalytic activity for sXR and 30% reduction for sXDH since XR and XDH displayed the optimal pH at 6.0 and 8.0, respectively. Therefore, the postulated modulation of enzyme activity by the lower pH shift near the DNA scaffold surface unlikely explained the catalytic enhancements of both scaffolded enzymes [76].

3.8. Packed State of Enzymes on the DNA Scaffold

In cells, enzyme reactions were performed in highly packed conditions. To mimic this “cellular crowed environment” in vitro still remains challenging. Huyen et al. [83] applied the modular adaptor method to assemble the monomeric carbonic anhydrase (CA) on a DNA scaffold in the packed state or dispersed states. CA was genetically fused to the modular adaptor ZF-SNAP to obtain ZS-CA. In the packed state, the interenzyme distance between CA regions was less than 1 nm. The reactions of ZS-CA assembled on the DNA scaffold were performed with the substrate p-nitrophenyl acetate (p-NPA), p-nitrophenyl butyrate (p-NPB) or p-nitrophenyl valerate (p-NPV). Interestingly, the enzymatic reactions proceeded faster in the packed than in the dispersed state under same enzyme and substrate concentrations. Acceleration of the reactions in the packed assembly was more predominant for substrates with higher water-excluded volumes (higher hydrophobicity), in which the reactions were accelerated by 1.3-fold, 1.5-fold or 1.6-fold in the packed state over the dispersed state with p-NPA, p-NPB or p-NPV as the substrate. The entropic force of water increasing the local substrate concentration within the domain confined between enzyme surfaces was attributed to the acceleration of enzyme reactions in the packed assembly. The acceleration of the enzyme reaction in the packed state over the dispersed state was also observed for xylose reductase assembled on the same type of DNA scaffold. This system provides a reasonable molecular model of enzymes in a packed state inside the cell, such as the condensate in the liquid–liquid phase separation.

References

  1. Hossain, G.S.; Nadarajan, S.P.; Zhang, L.; Ng, T.K.; Foo, J.L.; Ling, H.; Choi, W.J.; Chang, M.W. Rewriting the metabolic blueprint: Advances in pathway diversification in microorganisms. Front. Microbiol. 2018, 9, 155.
  2. Chen, A.H.; Silver, P.A. Designing biological compartmentalization. Trends Cell Biol. 2012, 22, 662–670.
  3. Agapakis, C.M.; Boyle, P.M.; Silver, P.A. Natural strategies for the spatial optimization of metabolism in synthetic biology. Nat. Chem. Biol. 2012, 8, 527–535.
  4. Bonacci, W.; Teng, P.K.; Afonso, B.; Niederholtmeyer, H.; Grob, P.; Silver, P.A.; Savage, D.F. Modularity of a carbon-fixing protein organelle. Proc. Natl. Acad. Sci. USA 2012, 109, 478–483.
  5. Šrejber, M.; Navrátilová, V.; Paloncýová, M.; Bazgier, V.; Berka, K.; Anzenbacher, P.; Otyepka, M. Membrane-attached mammalian cytochromes P450: An overview of the membrane’s effects on structure, drug binding, and interactions with redox partners. J. Inorg. Biochem. 2018, 183, 117–136.
  6. Kastritis, P.L.; Gavin, A.C. Enzymatic complexes across scales. Essays Biochem. 2018, 62, 501–514.
  7. Zhang, Y.H.P. Substrate channeling and enzyme complexes for biotechnological applications. Biotechnol. Adv. 2011, 29, 715–725.
  8. Seo, M.J.; Schmidt-Dannert, C. Organizing multi-enzyme systems into programmable materials for biocatalysis. Catalysts 2021, 11, 409.
  9. Rabe, K.S.; Müller, J.; Skoupi, M.; Niemeyer, C.M. Cascades in compartments: En route to machine-assisted biotechnology. Angew. Chem. Int. Ed. 2017, 56, 13574–13589.
  10. Küchler, A.; Yoshimoto, M.; Luginbühl, S.; Mavelli, F.; Walde, P. Enzymatic reactions in confined environments. Nat. Nanotechnol. 2016, 11, 409–420.
  11. Vázquez-González, M.; Wang, C.; Willner, I. Biocatalytic cascades operating on macromolecular scaffolds and in confined environments. Nat. Catal. 2020, 3, 256–273.
  12. Dubey, N.C.; Tripathi, B.P. Nature inspired multienzyme immobilization: Strategies and concepts. ACS Appl. Bio Mater. 2021, 4, 1077–1114.
  13. Madsen, M.; Gothelf, K.V. Chemistries for DNA nanotechnology. Chem. Rev. 2019, 119, 6384–6458.
  14. Rothemund, P.W. Folding DNA to create nanoscale shapes and patterns. Nature 2006, 440, 297–302.
  15. Douglas, S.M.; Dietz, H.; Liedl, T.; Högberg, B.; Graf, F.; Shih, W.M. Self-assembly of DNA into nanoscale three-dimensional shapes. Nature 2009, 459, 414–418.
  16. Hong, F.; Zhang, F.; Liu, Y.; Yan, H. DNA origami: Scaffolds for creating higher order structures. Chem. Rev. 2017, 117, 12584–12640.
  17. Fu, J.; Wang, Z.; Liang, X.H.; Oh, S.W.; Iago-McRae, E.S.; Zhang, T. DNA-scaffolded proximity assembly and confinement of multienzyme reactions. Top. Curr. Chem. 2020, 378, 38.
  18. Du, P.; Xu, S.; Xu, Z.K.; Wang, Z.G. Bioinspired Self-Assembling Materials for Modulating Enzyme Functions. Adv. Funct. Mater. 2021, 31, 2104819.
  19. Jaekel, A.; Stegemann, P.; Saccà, B. Manipulating enzymes properties with DNA nanostructures. Molecules 2019, 24, 3694.
  20. Zhang, Y.; Ge, J.; Liu, Z. Enhanced activity of immobilized or chemically modified enzymes. ACS Catal. 2015, 5, 4503–4513.
  21. Linko, V.; Nummelin, S.; Aarnos, L.; Tapio, K.; Toppari, J.J.; Kostiainen, M.A. DNA-based enzyme reactors and systems. Nanomaterials 2016, 6, 139.
  22. Schmitt, D.L.; An, S. Spatial organization of metabolic enzyme complexes in cells. Biochemistry 2017, 56, 3184–3196.
  23. Wheeldon, I.; Minteer, S.D.; Banta, S.; Barton, S.C.; Atanassov, P.; Sigman, M. Substrate channelling as an approach to cascade reactions. Nat. Chem. 2016, 8, 299–309.
  24. Maria-Solano, M.A.; Iglesias-Fernández, J.; Osuna, S. Deciphering the allosterically driven conformational ensemble in tryptophan synthase evolution. J. Am. Chem. Soc. 2019, 141, 13049–13056.
  25. Miles, E.W.; Rhee, S.; Davies, D.R. The molecular basis of substrate channeling. J. Biol. Chem. 1999, 274, 12193–12196.
  26. Wu, F.; Minteer, S. Krebs cycle metabolon: Structural evidence of substrate channeling revealed by cross-linking and mass spectrometry. Angew. Chem. Int. Ed. 2015, 54, 1851–1854.
  27. Sweetlove, L.J.; Fernie, A.R. The role of dynamic enzyme assemblies and substrate channelling in metabolic regulation. Nat. Commun. 2018, 9, 2136.
  28. Pedley, A.M.; Benkovic, S.J. A new view into the regulation of purine metabolism: The purinosome. Trends Biochem Sci. 2017, 42, 141–154.
  29. Kotopka, B.J.; Smolke, C.D. Production of the cyanogenic glycoside dhurrin in yeast. Metab. Eng. Commun. 2019, 9, e00092.
  30. Araiza-Olivera, D.; Chiquete-Felix, N.; Rosas-Lemus, M.; Sampedro, J.G.; Peña, A.; Mujica, A.; Uribe-Carvajal, S. A glycolytic metabolon in S accharomyces cerevisiae is stabilized by F-actin. FEBS J. 2013, 280, 3887–3905.
  31. An, S.; Kumar, R.; Sheets, E.D.; Benkovic, S.J. Reversible compartmentalization of de novo purine biosynthetic complexes in living cells. Science 2008, 320, 103–106.
  32. Pedley, A.M.; Pareek, V.; Benkovic, S.J. The Purinosome: A Case Study for a Mammalian Metabolon. Annu. Rev. Biochem. 2022, 91, 89–106.
  33. Pareek, V.; Tian, H.; Winograd, N.; Benkovic, S.J. Metabolomics and mass spectrometry imaging reveal channeled de novo purine synthesis in cells. Science 2020, 368, 283–290.
  34. Pedley, A.M.; Boylan, J.P.; Chan, C.Y.; Kennedy, E.L.; Kyoung, M.; Benkovic, S.J. Purine biosynthetic enzymes assemble into liquid-like condensates dependent on the activity of chaperone protein HSP90. J. Biol. Chem. 2022, 298, 101845.
  35. Wang, H.; Yan, X.; Aigner, H.; Bracher, A.; Nguyen, N.D.; Hee, W.Y.; Long, B.M.; Price, G.D.; Hartl, F.U.; Hayer-Hartl, M. Rubisco condensate formation by CcmM in β-carboxysome biogenesis. Nature 2019, 566, 131–135.
  36. Liu, L.N. Distribution and dynamics of electron transport complexes in cyanobacterial thylakoid membranes. Biochim. Biophys. Acta Bioenerg. 2016, 1857, 256–265.
  37. Liang, W.; Xu, H.; Carraro, F.; Maddigan, N.K.; Li, Q.; Bell, S.G.; Huang, D.M.; Tarzia, A.; Solomon, M.B.; Amenitsch, H.; et al. Enhanced activity of enzymes encapsulated in hydrophilic metal–organic frameworks. J. Am. Chem. Soc. 2019, 141, 2348–2355.
  38. Yue, L.; Wang, S.; Wulf, V.; Willner, I. Stiffness-switchable DNA-based constitutional dynamic network hydrogels for self-healing and matrix-guided controlled chemical processes. Nat. Commun. 2019, 10, 4774.
  39. Lin, P.; Zhang, Y.; Ren, H.; Wang, Y.; Wang, S.; Fang, B. Assembly of graphene oxide-formate dehydrogenase composites by nickel-coordination with enhanced stability and reusability. Eng. Life Sci. 2018, 18, 326–333.
  40. Walde, P.; Ichikawa, S. Enzymes inside lipid vesicles: Preparation, reactivity and applications. Biomol. Eng. 2001, 18, 143–177.
  41. Klermund, L.; Poschenrieder, S.T.; Castiglione, K. Biocatalysis in polymersomes: Improving multienzyme cascades with incompatible reaction steps by compartmentalization. ACS Catal. 2017, 7, 3900–3904.
  42. Zhang, G.; Quin, M.B.; Schmidt-Dannert, C. Self-assembling protein scaffold system for easy in vitro coimmobilization of biocatalytic cascade enzymes. ACS Catal. 2018, 8, 5611–5620.
  43. Rajendran, A.; Nakata, E.; Nakano, S.; Morii, T. Nucleic-Acid-Templated Enzyme Cascades. ChemBioChem 2017, 18, 696–716.
  44. Yoshimoto, M.; Wang, S.; Fukunaga, K.; Fournier, D.; Walde, P.; Kuboi, R.; Nakao, K. Novel immobilized liposomal glucose oxidase system using the channel protein OmpF and catalase. Biotechnol. Bioeng. 2005, 90, 231–238.
  45. Frey, R.; Mantri, S.; Rocca, M.; Hilvert, D. Bottom-up construction of a primordial carboxysome mimic. J. Am. Chem. Soc. 2016, 138, 10072–10075.
  46. Tapio, K.; Bald, I. The potential of DNA origami to build multifunctional materials. Multifunct. Mater. 2020, 3, 032001.
  47. Wang, P.; Meyer, T.A.; Pan, V.; Dutta, P.K.; Ke, Y. The beauty and utility of DNA origami. Chem 2017, 2, 359–382.
  48. Castro, C.E.; Kilchherr, F.; Kim, D.N.; Shiao, E.L.; Wauer, T.; Wortmann, P.; Bathe, M.; Dietz, H. A primer to scaffolded DNA origami. Nat. Methods. 2011, 8, 221–229.
  49. Douglas, S.M.; Marblestone, A.H.; Teerapittayanon, S.; Vazquez, A.; Church, G.M.; Shih, W.M. Rapid prototyping of 3D DNA-origami shapes with caDNAno. Nucleic Acids Res. 2009, 37, 5001–5006.
  50. Andersen, E.S.; Dong, M.; Nielsen, M.M.; Jahn, K.; Subramani, R.; Mamdouh, W.; Golas, M.M.; Sander, B.; Stark, H.; Oliveira, C.L.P.; et al. Self-assembly of a nanoscale DNA box with a controllable lid. Nature 2009, 459, 73–76.
  51. Han, D.; Pal, S.; Nangreave, J.; Deng, Z.; Liu, Y.; Yan, H. DNA origami with complex curvatures in three-dimensional space. Science 2011, 332, 342–346.
  52. Veneziano, R.; Ratanalert, S.; Zhang, K.; Zhang, F.; Yan, H.; Chiu, W.; Bathe, M. Designer nanoscale DNA assemblies programmed from the top down. Science 2016, 352, 1534.
  53. Kong, G.; Xiong, M.; Liu, L.; Hu, L.; Meng, H.M.; Ke, G.; Zhang, X.B.; Tan, W. DNA origami-based protein networks: From basic construction to emerging applications. Chem. Soc. Rev. 2021, 50, 1846–1873.
  54. DeLuca, M.; Shi, Z.; Castro, C.E.; Arya, G. Dynamic DNA nanotechnology: Toward functional nanoscale devices. Nanoscale Horiz. 2020, 5, 182–201.
  55. Zhang, D.Y.; Seelig, G. Dynamic DNA nanotechnology using strand-displacement reactions. Nat. Chem. 2011, 3, 103–113.
  56. Rangel, A.E.; Hariri, A.A.; Eisenstein, M.; Soh, H.T. Engineering aptamer switches for multifunctional stimulus-responsive nanosystems. Adv. Mater. 2020, 32, 2003704.
  57. Turek, V.A.; Chikkaraddy, R.; Cormier, S.; Stockham, B.; Ding, T.; Keyser, U.F.; Baumberg, J.J. Thermo-Responsive Actuation of a DNA Origami Flexor. Adv. Funct. Mater. 2018, 28, 1706410.
  58. Ijäs, H.; Hakaste, I.; Shen, B.; Kostiainen, M.A.; Linko, V. Reconfigurable DNA origami nanocapsule for pH-controlled encapsulation and display of cargo. ACS Nano 2019, 13, 5959–5967.
  59. Marras, A.E.; Shi, Z.; Lindell III, M.G.; Patton, R.A.; Huang, C.M.; Zhou, L.; Su, H.J.; Arya, G.; Castro, C.E. Cation-activated avidity for rapid reconfiguration of DNA nanodevices. ACS Nano 2018, 12, 9484–9494.
  60. Kopperger, E.; List, J.; Madhira, S.; Rothfischer, F.; Lamb, D.C.; Simmel, F.C. A self-assembled nanoscale robotic arm controlled by electric fields. Science 2018, 359, 296–301.
  61. Ijäs, H.; Nummelin, S.; Shen, B.; Kostiainen, M.A.; Linko, V. Dynamic DNA origami devices: From strand-displacement reactions to external-stimuli responsive systems. Int. J. Mol. Sci. 2018, 19, 2114.
  62. Chandrasekaran, A.R.; Anderson, N.; Kizer, M.; Halvorsen, K.; Wang, X. Beyond the fold: Emerging biological applications of DNA origami. ChemBioChem 2016, 17, 1081–1089.
  63. McCluskey, J.B.; Clark, D.S.; Glover, D.J. Functional Applications of Nucleic Acid–Protein Hybrid Nanostructures. Trends Biotechnol. 2020, 38, 976–989.
  64. Jiao, Y.; Shang, Y.; Li, N.; Ding, B. DNA-based enzymatic systems and their applications. Iscience 2022, 25, 104018.
  65. Jiang, Q.; Liu, S.; Liu, J.; Wang, Z.G.; Ding, B. Rationally designed DNA-origami nanomaterials for drug delivery in vivo. Adv. Mater. 2019, 31, 1804785.
  66. Tan, Y.Q.; Xue, B.; Yew, W.S. Genetically Encodable Scaffolds for Optimizing Enzyme Function. Molecules 2021, 26, 1389.
  67. Glettenberg, M.; Niemeyer, C.M. Tuning of peroxidase activity by covalently tethered DNA oligonucleotides. Bioconjugate Chem. 2009, 20, 969–975.
  68. Rudiuk, S.; Venancio-Marques, A.; Baigl, D. Enhancement and Modulation of Enzymatic Activity through Higher-Order Structural Changes of Giant DNA–Protein Multibranch Conjugates. Angew. Chem. Int. Ed. 2012, 51, 12694–12698.
  69. Timm, C.; Niemeyer, C.M. Assembly and purification of enzyme-functionalized DNA origami structures. Angew. Chem. Int. Ed. 2015, 54, 6745–6750.
  70. Kim, S.H.; Kim, K.R.; Ahn, D.R.; Lee, J.E.; Yang, E.G.; Kim, S.Y. Reversible regulation of enzyme activity by pH-responsive encapsulation in DNA nanocages. ACS Nano 2017, 11, 9352–9359.
  71. Lin, J.L.; Wheeldon, I. Kinetic enhancements in DNA–enzyme nanostructures mimic the sabatier principle. ACS Catal. 2013, 3, 560–564.
  72. Kosinski, R.; Perez, J.M.; Schöneweiß, E.C.; Ruiz-Blanco, Y.B.; Ponzo, I.; Bravo-Rodriguez, K.; Erkelenz, M.; Schlücker, S.; Uhlenbrock, G.; Sanchez-Garcia, E.; et al. The role of DNA nanostructures in the catalytic properties of an allosterically regulated protease. Sci. Adv. 2022, 8, eabk0425.
  73. Zhao, Z.; Fu, J.; Dhakal, S.; Johnson-Buck, A.; Liu, M.; Zhang, T.; Woodbury, N.W.; Liu, Y.; Walter, N.G.; Yan, H. Nanocaged enzymes with enhanced catalytic activity and increased stability against protease digestion. Nat. Commun. 2016, 7, 10619.
  74. Zhang, Y.; Tsitkov, S.; Hess, H. Proximity does not contribute to activity enhancement in the glucose oxidase–horseradish peroxidase cascade. Nat. Commun. 2016, 7, 13982.
  75. Xiong, Y.; Huang, J.; Wang, S.T.; Zafar, S.; Gang, O. Local environment affects the activity of enzymes on a 3D molecular scaffold. ACS Nano 2020, 14, 14646–14654.
  76. Lin, P.; Dinh, H.; Morita, Y.; Zhang, Z.; Nakata, E.; Kinoshita, M.; Morii, T. Evaluation of the role of the DNA surface for enhancing the activity of scaffolded enzymes. Chem. Commun. 2021, 57, 3925–3928.
  77. Nakata, E.; Dinh, H.; Ngo, T.A.; Saimura, M.; Morii, T. A modular zinc finger adaptor accelerates the covalent linkage of proteins at specific locations on DNA nanoscaffolds. Chem. Commun. 2015, 51, 1016–1019.
  78. Ngo, T.A.; Nakata, E.; Saimura, M.; Morii, T. Spatially organized enzymes drive cofactor-coupled cascade reactions. J. Am. Chem. Soc. 2016, 138, 3012–3021.
  79. Nguyen, T.M.; Nakata, E.; Saimura, M.; Dinh, H.; Morii, T. Design of modular protein tags for orthogonal covalent bond formation at specific DNA sequences. J. Am. Chem. Soc. 2017, 139, 8487–8496.
  80. Ngo, T.A.; Dinh, H.; Nguyen, T.M.; Liew, F.F.; Nakata, E.; Morii, T. Protein adaptors assemble functional proteins on DNA scaffolds. Chem. Commun. 2019, 55, 12428–12446.
  81. Nguyen, T.M.; Nakata, E.; Zhang, Z.; Saimura, M.; Dinh, H.; Morii, T. Rational design of a DNA sequence-specific modular protein tag by tuning the alkylation kinetics. Chem. Sci. 2019, 10, 9315–9325.
  82. Zhang, Z.; Nakata, E.; Dinh, H.; Saimura, M.; Rajendran, A.; Matsuda, K.; Morii, T. Tuning the Reactivity of a Substrate for SNAP-Tag Expands Its Application for Recognition-Driven DNA-Protein Conjugation. Chem. Eur. J. 2021, 27, 18118–18128.
  83. Dinh, H.; Nakata, E.; Mutsuda-Zapater, K.; Saimura, M.; Kinoshita, M.; Morii, T. Enhanced enzymatic activity exerted by a packed assembly of a single type of enzyme. Chem. Sci. 2020, 11, 9088–9100.
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