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The enzyme cyclic di-phosphoglycerate synthetase that is involved in the production of the osmolyte cyclic 2,3-diphosphoglycerate has been studied both biochemically and structurally. Cyclic 2,3-diphosphoglycerate is found exclusively in the hyperthermophilic archaeal methanogens, such as Methanothermus fervidus, Methanopyrus kandleri, and Methanothermobacter thermoautotrophicus. Its presence increases the thermostability of archaeal proteins and protects the DNA against oxidative damage caused by hydroxyl radicals. The cyclic 2,3-diphosphoglycerate synthetase enzyme has been crystallized and its structure solved to 1.7 Å resolution by experimental phasing. It has also been crystallized in complex with its substrate 2,3 diphosphoglycerate and the co-factor ADP and this structure has been solved to 2.2 Å resolution. The enzyme structure has two domains, the core domain shares some structural similarity with other NTP-dependent enzymes. A significant proportion of the structure, including a 127 amino acid N-terminal domain, has no structural similarity to other known enzyme structures. The structure of the complex shows a large conformational change that occurs in the enzyme during catalytic turnover. The reaction involves the transfer of the γ-phosphate group from ATP to the substrate 2,3 -diphosphoglycerate and the subsequent SN2 attack to form a phosphoanhydride. This results in the production of the unusual extremolyte cyclic 2,3 -diphosphoglycerate which has important industrial applications. [ABSTRACT FROM AUTHOR]

Glycosylation is the most prevalent protein post‐translational modification, with a quarter of glycosylated proteins having enzymatic properties. Yet, the full impact of glycosylation on the protein structure–function relationship, especially in enzymes, is still limited. Here, we show that glycosylation rigidifies the important commercial enzyme horseradish peroxidase (HRP), which in turn increases its turnover and stability. Circular dichroism spectroscopy revealed that glycosylation increased holo‐HRP's thermal stability and promoted significant helical structure in the absence of haem (apo‐HRP). Glycosylation also resulted in a 10‐fold increase in enzymatic turnover towards o‐phenylenediamine dihydrochloride when compared to its nonglycosylated form. Utilising a naturally occurring site‐specific probe of active site flexibility (Trp117) in combination with red‐edge excitation shift fluorescence spectroscopy, we found that glycosylation significantly rigidified the enzyme. In silico simulations confirmed that glycosylation largely decreased protein backbone flexibility, especially in regions close to the active site and the substrate access channel. Thus, our data show that glycosylation does not just have a passive effect on HRP stability but can exert long‐range effects that mediate the 'native' enzyme's activity and stability through changes in inherent dynamics. [ABSTRACT FROM AUTHOR]

Fluorescent proteins (FPs) are commonly used in pairs to monitor dynamic biomolecular events through changes in proximity via distance dependent processes such as Förster resonance energy transfer (FRET). The impact of FP association is assessed by predicting dimerization sites in silico and stabilizing the dimers by bio‐orthogonal covalent linkages. In each tested case dimerization changes inherent fluorescence, including FRET. GFP homodimers demonstrate synergistic behavior with the dimer being brighter than the sum of the monomers. The homodimer structure reveals the chromophores are close with favorable transition dipole alignments and a highly solvated interface. Heterodimerization (GFP with Venus) results in a complex with ≈87% FRET efficiency, significantly below the 99.7% efficiency predicted. A similar efficiency is observed when the wild‐type FPs are fused to a naturally occurring protein–protein interface system. GFP complexation with mCherry results in loss of mCherry fluorescence. Thus, simple assumptions used when monitoring interactions between proteins via FP FRET may not always hold true, especially under conditions whereby the protein–protein interactions promote FP interaction. [ABSTRACT FROM AUTHOR]

Protein oligomers are more common in nature than monomers, with dimers being the most prevalent final structural state observed in known structures. From a biological perspective, this makes sense as it conserves vital molecular resources that may be wasted simply by generating larger single polypeptide units, and allows new features such as cooperativity to emerge. Taking inspiration from nature, protein designers and engineers are now building artificial oligomeric complexes using a variety of approaches to generate new and useful supramolecular protein structures. Oligomerisation is thus offering a new approach to sample structure and function space not accessible through simply tinkering with monomeric proteins. [ABSTRACT FROM AUTHOR]

The bright bioluminescence catalyzed by Photinus pyralis firefly luciferase (Fluc) enables a vast array of life science research such as bio imaging in live animals and sensitive in vitro diagnostics. The effectiveness of such applications is improved using engineered enzymes that to date have been constructed using amino acid substitutions. We describe ΔFlucs: consecutive single amino acid deletion mutants within six loop structures of the bright and thermostable ×11 Fluc. Deletion mutations are a promising avenue to explore new sequence and functional space and isolate novel mutant phenotypes. However, this method is often overlooked and to date there have been no surveys of the effects of consecutive single amino acid deletions in Fluc. We constructed a large semi-rational ΔFluc library and isolated significantly brighter enzymes after finding ×11 Fluc activity was largely tolerant to deletions. Targeting an 'omega-loop' motif (T352-G360) significantly enhanced activity, altered kinetics, reduced Km for D-luciferin, altered emission colors, and altered substrate specificity for redshifted analog DL-infraluciferin. Experimental and in silico analyses suggested remodeling of the Ω-loop impacts on active site hydrophobicity to increase light yields. This work demonstrates the further potential of deletion mutations, which can generate useful Fluc mutants and broaden the palette of the biomedical and biotechnological bioluminescence enzyme toolbox. [ABSTRACT FROM AUTHOR]

We report the site-specific coupling of single proteins to individual carbon nanotubes (CNTs) in solution and with single-molecule control. Using an orthogonal Click reaction, Green Fluorescent Protein (GFP) was engineered to contain a genetically encoded azide group and then bound to CNT ends in different configurations: in close proximity or at longer distances from the GFP's functional center. Atomic force microscopy and fluorescence analysis in solution and on surfaces at the single-protein level confirmed the importance of bioengineering optimal protein attachment sites to achieve direct protein-nanotube communication and bridging. [ABSTRACT FROM AUTHOR]

The Hbl toxin is a three-component haemolytic complex produced by Bacillus cereus sensu lato strains and implicated as a cause of diarrhoea in B. cereus food poisoning. While the structure of the HblB component of this toxin is known, the structures of the other components are unresolved. Here, we describe the expression of the recombinant HblL1 component and the elucidation of its structure to 1.36 Å. Like HblB, it is a member of the alpha-helical pore-forming toxin family. In comparison to other members of this group, it has an extended hydrophobic beta tongue region that may be involved in pore formation. Molecular docking was used to predict possible interactions between HblL1 and HblB, and suggests a head to tail dimer might form, burying the HblL1 beta tongue region. [ABSTRACT FROM AUTHOR]

Fluorescent proteins have revolutionized cell biology and cell imaging through their use as genetically encoded tags. Structural biology has been pivotal in understanding how their unique fluorescent properties manifest through the formation of the chromophore and how the spectral properties are tuned through interaction networks. This knowledge has in turn led to the construction of novel variants with new and improved properties. Here we describe the process by which fluorescent protein structures are determined, starting from recombinant protein production to structure determination by molecular replacement. We also describe how to incorporate and determine the structures of proteins containing non-natural amino acids. Recent advances in protein engineering have led to reprogramming of the genetic code to allow incorporation of new chemistry at designed residue positions, with fluorescent proteins being at the forefront of structural studies in this area. The impact of such new chemistry on protein structure is still limited; the accumulation of more protein structures will undoubtedly improve our understanding and ability to engineer proteins with new chemical functionality., (© 2023. The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature.)

Here we determined the structure of a cold active family IV esterase (EstN7) cloned from Bacillus cohnii strain N1. EstN7 is a dimer with a classical α/β hydrolase fold. It has an acidic surface that is thought to play a role in cold-adaption by retaining solvation under changed water solvent entropy at lower temperatures. The conformation of the functionally important cap region is significantly different to EstN7's closest relatives, forming a bridge-like structure with reduced helical content providing greater access to the active site through more than one substrate access tunnel. However, dynamics do not appear to play a major role in cold adaption. Molecular dynamics at different temperatures, rigidity analysis, normal mode analysis and geometric simulations of motion confirm the flexibility of the cap region but suggest that the rest of the protein is largely rigid. Rigidity analysis indicates the distribution of hydrophobic tethers is appropriate to colder conditions, where the hydrophobic effect is weaker than in mesophilic conditions due to reduced water entropy. Thus, it is likely that increased substrate accessibility and tolerance to changes in water entropy are important for of EstN7's cold adaptation rather than changes in dynamics.

The formation of protein complexes is central to biology, with oligomeric proteins more prevalent than monomers. The coupling of functionally and even structurally distinct protein units can lead to new functional properties not accessible by monomeric proteins alone. While such complexes are driven by evolutionally needs in biology, the ability to link normally functionally and structurally disparate proteins can lead to new emergent properties for use in synthetic biology and the nanosciences. Here we demonstrate how two disparate proteins, the haem binding helical bundle protein cytochrome b 562 and the β-barrel green fluorescent protein can be combined to form a heterodimer linked together by an unnatural triazole linkage. The complex was designed using computational docking approaches to predict compatible interfaces between the two proteins. Models of the complexes where then used to engineer residue coupling sites in each protein to link them together. Genetic code expansion was used to incorporate azide chemistry in cytochrome b 562 and alkyne chemistry in GFP so that a permanent triazole covalent linkage can be made between the two proteins. Two linkage sites with respect to GFP were sampled. Spectral analysis of the new heterodimer revealed that haem binding and fluorescent protein chromophore properties were retained. Functional coupling was confirmed through changes in GFP absorbance and fluorescence, with linkage site determining the extent of communication between the two proteins. We have thus shown here that is possible to design and build heterodimeric proteins that couple structurally and functionally disparate proteins to form a new complex with new functional properties., Competing Interests: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest., (Copyright © 2021 Johnson, Blaber, Evans, Worthy, Pope and Jones.)

Fluorescent proteins (FPs) are commonly used in pairs to monitor dynamic biomolecular events through changes in proximity via distance dependent processes such as Förster resonance energy transfer (FRET). The impact of FP association is assessed by predicting dimerization sites in silico and stabilizing the dimers by bio-orthogonal covalent linkages. In each tested case dimerization changes inherent fluorescence, including FRET. GFP homodimers demonstrate synergistic behavior with the dimer being brighter than the sum of the monomers. The homodimer structure reveals the chromophores are close with favorable transition dipole alignments and a highly solvated interface. Heterodimerization (GFP with Venus) results in a complex with ≈87% FRET efficiency, significantly below the 99.7% efficiency predicted. A similar efficiency is observed when the wild-type FPs are fused to a naturally occurring protein-protein interface system. GFP complexation with mCherry results in loss of mCherry fluorescence. Thus, simple assumptions used when monitoring interactions between proteins via FP FRET may not always hold true, especially under conditions whereby the protein-protein interactions promote FP interaction., Competing Interests: The authors declare no conflict of interest. 1a) E. A.Rodriguez, R. E.Campbell, J. Y.Lin, M. Z.Lin, A.Miyawaki, A. E.Palmer, X.Shu, J.Zhang, R. Y.Tsien, Trends Biochem. Sci.2017, 42, 111;27814948b) S.Duwe, P.Dedecker, Curr. Opin. Biotechnol.2019, 58, 183.311706102a) N. C.Shaner, P. A.Steinbach, R. Y.Tsien, Nat. Methods2005, 2, 905;16299475b) R. H.Newman, M. D.Fosbrink, J.Zhang, Chem. Rev.2011, 111, 3614;21456512c) A.Ibraheem, R. E.Campbell, Curr. Opin. Chem. Biol.2010, 14, 30.199134533A.Miyawaki, D. M.Shcherbakova, V. V.Verkhusha, Curr. Opin. Struct. Biol.2012, 22, 679.230000314R. Y.Tsien, Annu. Rev. Biochem.1998, 67, 509.97594965M. V.Matz, A. F.Fradkov, Y. A.Labas, A. P.Savitsky, A. G.Zaraisky, M. L.Markelov, S. A.Lukyanov, Nat. Biotechnol.1999, 17, 969.105046966P. J.Cranfill, B. R.Sell, M. A.Baird, J. R.Allen, Z.Lavagnino, H. M.de Gruiter, G. J.Kremers, M. W.Davidson, A.Ustione, D. W.Piston, Nat. Methods2016, 13, 557.272402577D. A.Zacharias, J. D.Violin, A. C.Newton, R. Y.Tsien, Science2002, 296, 913.119885768T.Förster, Ann. Phys.1948, 437, 55.9B.Hellenkamp, S.Schmid, O.Doroshenko, O.Opanasyuk, R.Kuhnemuth, S. R.Adariani, B.Ambrose, M.Aznauryan, A.Barth, V.Birkedal, M. E.Bowen, H.Chen, T.Cordes, T.Eilert, C.Fijen, C.Gebhardt, M.Gotz, G.Gouridis, E.Gratton, T.Ha, P.Hao, C. A.Hanke, A.Hartmann, J.Hendrix, L. L.Hildebrandt, V.Hirschfeld, J.Hohlbein, B.Hua, C. G.Hubner, E.Kallis, et al., Nat. Methods2018, 15, 669.3017125210a) S. C.Alford, Y.Ding, T.Simmen, R. E.Campbell, ACS Synth. Biol.2012, 1, 569;23656278b) X. X.Zhou, H. K.Chung, A. J.Lam, M. Z.Lin, Science2012, 338, 810.2313933511H. L.Worthy, H. S.Auhim, W. D.Jamieson, J. R.Pope, A.Wall, R.Batchelor, R. L.Johnson, D. W.Watkins, P.Rizkallah, O. K.Castell, D. D.Jones, Commun. Chem.2019, 2, 83.12P.Trigo‐Mourino, T.Thestrup, O.Griesbeck, C.Griesinger, S.Becker, Sci. Adv.2019, 5, eaaw4988.3145708813M. D.Wiens, Y.Shen, X.Li, M. A.Salem, N.Smisdom, W.Zhang, A.Brown, R. E.Campbell, ChemBioChem2016, 17, 2361.2778139414A. J.Lam, F.St‐Pierre, Y.Gong, J. D.Marshall, P. J.Cranfill, M. A.Baird, M. R.McKeown, J.Wiedenmann, M. W.Davidson, M. J.Schnitzer, R. Y.Tsien, M. Z.Lin, Nat. Methods2012, 9, 1005.2296124515L. H.Lindenburg, A. M.Hessels, E. H.Ebberink, R.Arts, M.Merkx, ACS Chem. Biol.2013, 8, 2133.2396215616M.Mastop, D. S.Bindels, N. C.Shaner, M.Postma, T. W. J.GadellaJr., J.Goedhart, Sci. Rep.2017, 7, 11999.2893189817S. C.Reddington, P. J.Rizkallah, P. D.Watson, R.Pearson, E. M.Tippmann, D. D.Jones, Angew. Chem., Int. Ed.2013, 52, 5974.18S. C.Reddington, E. M.Tippmann, D. D.Jones, Chem. Commun.2012, 48, 8419.19J. A.Arpino, P. J.Rizkallah, D. D.Jones, PLoS One2012, 7, e47132.2307755520E. K.Bomati, J. E.Haley, J. P.Noel, D. D.Deheyn, Sci. Rep.2014, 4, 5469.2496892121a) K.Brejc, T. K.Sixma, P. A.Kitts, S. R.Kain, R. Y.Tsien, M.Ormo, S. J.Remington, Proc. Natl. Acad. Sci. U. S. A.1997, 94, 2306;9122190b) M. H.Seifert, D.Ksiazek, M. K.Azim, P.Smialowski, N.Budisa, T. A.Holak, J. Am. Chem. Soc.2002, 124, 7932.1209533722a) A. M.Hartley, H. L.Worthy, S. C.Reddington, P. J.Rizkallah, D. D.Jones, Chem. Sci.2016, 7, 6484;28451106b) A.Shinobu, N.Agmon, J. Chem. Theory Comput.2017, 13, 353;28068768c) A.Shinobu, G. J.Palm, A. J.Schierbeek, N.Agmon, J. Am. Chem. Soc.2010, 132, 11093.2069867523D.Kozakov, D. R.Hall, B.Xia, K. A.Porter, D.Padhorny, C.Yueh, D.Beglov, S.Vajda, Nat. Protoc.2017, 12, 255.2807987924T.Ansbacher, H. K.Srivastava, T.Stein, R.Baer, M.Merkx, A.Shurki, Phys. Chem. Chem. Phys.2012, 14, 4109.2233109925A.Kyrychenko, M. V.Rodnin, C.Ghatak, A. S.Ladokhin, Anal. Biochem.2017, 522, 1.2810816826T. J.Lambert, Nat. Methods2019, 16, 277.3088641227B. T.Andrews, A. R.Schoenfish, M.Roy, G.Waldo, P. A.Jennings, J. Mol. Biol.2007, 373, 476.1782271428B.Manavalan, S.Basith, Y. M.Choi, G.Lee, S.Choi, PLoS One2010, 5, e15782.2120342229a) X.Shu, N. C.Shaner, C. A.Yarbrough, R. Y.Tsien, S. J.Remington, Biochemistry2006, 45, 9639;16893165b) N. C.Shaner, R. E.Campbell, P. A.Steinbach, B. N.Giepmans, A. E.Palmer, R. Y.Tsien, Nat. Biotechnol.2004, 22, 1567.1555804730a) B. T.Bajar, E. S.Wang, S.Zhang, M. Z.Lin, J.Chu, Sensors2016, 16, 1488;b) D.Shcherbo, E. A.Souslova, J.Goedhart, T. V.Chepurnykh, A.Gaintzeva, ShemiakinaII, T. W.Gadella, S.Lukyanov, D. M.Chudakov, BMC Biotechnol.2009, 9, 24;19321010c) G. N.van der Krogt, J.Ogink, B.Ponsioen, K.Jalink, PLoS One2008, 3, e1916.1838268731B. M. C.Cloin, E.de Zitter, D.Salas, V.Gielen, G. E.Folkers, M.Mikhaylova, M.Bergeler, B.Krajnik, J.Harvey, C. C.Hoogenraad, L.van Meervelt, P.Dedecker, L. C.Kapitein, Proc. Natl. Acad. Sci. U. S. A.2017, 114, 7013.2863028632F. V.Subach, V. N.Malashkevich, W. D.Zencheck, H.Xiao, G. S.Filonov, S. C.Almo, V. V.Verkhusha, Proc. Natl. Acad. Sci. U. S. A.2009, 106, 21097.1993403633F. V.Subach, V. V.Verkhusha, Chem. Rev.2012, 112, 4308.2255923234A. C.Stiel, M.Andresen, H.Bock, M.Hilbert, J.Schilde, A.Schonle, C.Eggeling, A.Egner, S. W.Hell, S.Jakobs, Biophys. J.2008, 95, 2989.1865822135Collaborative Computational Project Number 4 , Acta Crystallogr., Sect. D: Biol. Crystallogr.1994, 50, 760.1529937436A. J.McCoy, R. W.Grosse‐Kunstleve, P. D.Adams, M. D.Winn, L. C.Storoni, R. J.Read, J. Appl. Crystallogr.2007, 40, 658.1946184037P.Emsley, K.Cowtan, Acta Crystallogr., Sect. D: Biol. Crystallogr.2004, 60, 2126.1557276538G. N.Murshudov, A. A.Vagin, E. J.Dodson, Acta Crystallogr., Sect. D: Biol. Crystallogr.1997, 53, 240.1529992639D. D.Fernandes, J.Bamrah, S.Kailasam, G. W.Gomes, Y.Li, H. J.Wieden, C. C.Gradinaru, Sci. Rep.2017, 7, 13063.29026195, (© 2020 The Authors. Published by Wiley‐VCH GmbH.)

Functional integration of proteins with carbon-based nanomaterials such as nanotubes holds great promise in emerging electronic and optoelectronic applications. Control over protein attachment poses a major challenge for consistent and useful device fabrication, especially when utilizing single/few molecule properties. Here, we exploit genetically encoded phenyl azide photochemistry to define the direct covalent attachment of four different proteins, including the fluorescent protein GFP and a β-lactamase binding protein (BBP), to carbon nanotube side walls. AFM showed that on attachment BBP could still recognize and bind additional protein components. Single molecule fluorescence revealed that on attachment to SWCNTs function was retained and there was feedback to GFP in terms of fluorescence intensity and improved resistance to photobleaching; GFP is fluorescent for much longer on attachment. The site of attachment proved important in terms of electronic impact on GFP function, with the attachment site furthest from the chromophore having the larger effect on fluorescence. Our approach provides a versatile and general method for generating intimate protein-CNT hybrid bioconjugates. It can be potentially applied to any protein of choice; the attachment position and thus interface characteristics with the CNT can easily be changed by simply placing the phenyl azide chemistry at different residues by gene mutagenesis. Thus, our approach will allow consistent construction and modulate functional coupling through changing the protein attachment position.