Kinetic mechanism of Renilla luciferase guides induced-fit engineering for improved bioluminescence

Kinetic mechanism of Renilla luciferase guides induced-fit engineering for improved bioluminescence | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Kinetic mechanism of Renilla luciferase guides induced-fit engineering for improved bioluminescence View ORCID Profile M. Toul , View ORCID Profile J. Horackova , View ORCID Profile A. Schenkmayerova , View ORCID Profile J. Planas-Iglesias , T. Landolt , View ORCID Profile J. Sucharitakul , View ORCID Profile Y. L. Janin , K. Prakinee , View ORCID Profile P. Chaiyen , View ORCID Profile S. Stavrakis , View ORCID Profile A. deMello , View ORCID Profile K. A. Johnson , View ORCID Profile J. Damborsky , View ORCID Profile M. Marek , View ORCID Profile D. Bednar , View ORCID Profile Z. Prokop doi: https://doi.org/10.1101/2025.09.16.675553 M. Toul 1 Loschmidt Laboratories, Department of Experimental Biology and RECETOX, Faculty of Science, Masaryk University , Kamenice 5, Bld. C13, 625 00 Brno, Czech Republic 2 International Clinical Research Center, St. Anne’s University Hospital , Pekarska 53, 656 91 Brno, Czech Republic Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for M. Toul J. Horackova 1 Loschmidt Laboratories, Department of Experimental Biology and RECETOX, Faculty of Science, Masaryk University , Kamenice 5, Bld. C13, 625 00 Brno, Czech Republic 2 International Clinical Research Center, St. Anne’s University Hospital , Pekarska 53, 656 91 Brno, Czech Republic Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for J. Horackova A. Schenkmayerova 1 Loschmidt Laboratories, Department of Experimental Biology and RECETOX, Faculty of Science, Masaryk University , Kamenice 5, Bld. C13, 625 00 Brno, Czech Republic 2 International Clinical Research Center, St. Anne’s University Hospital , Pekarska 53, 656 91 Brno, Czech Republic Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for A. Schenkmayerova J. Planas-Iglesias 1 Loschmidt Laboratories, Department of Experimental Biology and RECETOX, Faculty of Science, Masaryk University , Kamenice 5, Bld. C13, 625 00 Brno, Czech Republic 2 International Clinical Research Center, St. Anne’s University Hospital , Pekarska 53, 656 91 Brno, Czech Republic Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for J. Planas-Iglesias T. Landolt 3 Institute for Chemical and Bioengineering , ETH Zürich, Vladimir Prelog Weg 1, 8093 Zürich, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site J. Sucharitakul 4 Department of Biochemistry, Faculty of Dentistry, Chulalongkorn University , Bangkok 10300, Thailand Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for J. Sucharitakul Y. L. Janin 5 Structure et Instabilité des Génomes (StrInG), Muséum National d’Histoire Naturelle, INSERM, CNRS, Alliance Sorbonne Université , 75005 Paris, France Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Y. L. Janin K. Prakinee 6 School of Biomolecular Science and Engineering, Vidyasirimedhi Institute of Science and Technology (VISTEC) , Rayong 21210, Thailand Find this author on Google Scholar Find this author on PubMed Search for this author on this site P. Chaiyen 6 School of Biomolecular Science and Engineering, Vidyasirimedhi Institute of Science and Technology (VISTEC) , Rayong 21210, Thailand Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for P. Chaiyen S. Stavrakis 3 Institute for Chemical and Bioengineering , ETH Zürich, Vladimir Prelog Weg 1, 8093 Zürich, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for S. Stavrakis A. deMello 3 Institute for Chemical and Bioengineering , ETH Zürich, Vladimir Prelog Weg 1, 8093 Zürich, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for A. deMello K. A. Johnson 7 Institute for Cellular and Molecular Biology, Department of Molecular Biosciences, University of Texas , 2500 Speedway, Austin, TX, 78712, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for K. A. Johnson J. Damborsky 1 Loschmidt Laboratories, Department of Experimental Biology and RECETOX, Faculty of Science, Masaryk University , Kamenice 5, Bld. C13, 625 00 Brno, Czech Republic 2 International Clinical Research Center, St. Anne’s University Hospital , Pekarska 53, 656 91 Brno, Czech Republic Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for J. Damborsky M. Marek 1 Loschmidt Laboratories, Department of Experimental Biology and RECETOX, Faculty of Science, Masaryk University , Kamenice 5, Bld. C13, 625 00 Brno, Czech Republic 2 International Clinical Research Center, St. Anne’s University Hospital , Pekarska 53, 656 91 Brno, Czech Republic Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for M. Marek For correspondence: zbynek{at}chemi.muni.cz D. Bednar 1 Loschmidt Laboratories, Department of Experimental Biology and RECETOX, Faculty of Science, Masaryk University , Kamenice 5, Bld. C13, 625 00 Brno, Czech Republic 2 International Clinical Research Center, St. Anne’s University Hospital , Pekarska 53, 656 91 Brno, Czech Republic Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for D. Bednar For correspondence: zbynek{at}chemi.muni.cz Z. Prokop 1 Loschmidt Laboratories, Department of Experimental Biology and RECETOX, Faculty of Science, Masaryk University , Kamenice 5, Bld. C13, 625 00 Brno, Czech Republic 2 International Clinical Research Center, St. Anne’s University Hospital , Pekarska 53, 656 91 Brno, Czech Republic Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Z. Prokop For correspondence: zbynek{at}chemi.muni.cz Abstract Full Text Info/History Metrics Supplementary material Preview PDF Abstract Renilla luciferase (RLuc) remains one of the most popular bioluminescence reporters, but its molecular principle has yet to be fully understood. Here, we reveal a detailed kinetic mechanism of the RLuc catalytic cycle which uncovers multiple limiting factors: (i) an oxygen-induced irreversible inactivation, (ii) a low oxygen saturation, and (iii) rate-limiting induced-fit conformational dynamics coupled with the product release. Furthermore, we could determine the actual enzyme k cat value at all saturating substrates to be 22 s - 1 . This value is 5-fold higher than the previously reported apparent k cat values determined at physiological, non-saturating oxygen concentration. Our integrative analysis by transient kinetics, X-ray crystallography, and molecular dynamics linked the rate-limiting flexible enzyme opening to the dynamics of the loops surrounding the active site, which prompted targeted engineering of this limiting step by loop grafting. The resulting variant AncFT-L14 (AncFT7) showed a prolonged stable light emission thanks to the selectively improved induced-fit kinetics. Additional characterization of AncFT-L14 identified increased catalytic efficiency k cat / K m , product inhibition factor K p / K m , and a glow-type signal characteristic. Our results provide mechanistic details of RLuc catalysis and will govern future enzyme engineering to design the next generations of bioluminescence-based tools. Introduction Bioluminescence, a fascinating process of light emission by living organisms, plays a crucial role in nature. This includes prey or mate attraction, defense, camouflage, and communication 1 – 3 . Moreover, bioluminescent-based technologies have become increasingly important for many industrial applications over the past decades, spanning diagnostics, environment monitoring, high throughput drug screening, and gene expression and cellular interaction analysis 4 – 6 . Therefore, advances in deciphering and harnessing the bioluminescence principles not only improve our understanding of this natural process but also offer innovative solutions in diverse industrial fields. The emission of visible light by living organisms is achieved by their genetically encoded luciferase enzymes. Although individual organisms often possess sequentially and structurally unrelated luciferases employing very different substrates, their common feature is the oxidation of a substrate (‘luciferin’) using molecular oxygen, its conversion to a product (‘oxyluciferin’), and the release of a photon with a luciferase/luciferin-dependent emission wavelength ( Figure 1a ) 5 , 7 , 8 . Luciferase from the sea pansy Renilla reniformis , henceforth referred to as RLuc, is a 36 kDa cofactor-free monomeric enzyme, making it one of the most popular bioluminescence-based reporting systems. RLuc catalyzes the oxidative decarboxylation of coelenterazine (CTZ) to generate coelenteramide (CEI) while simultaneously releasing light with an emission maximum of ∼480 nm ( Figure 1b ) 9 , 10 . In 2006, Loening and coauthors reported an 8-point mutant variant of RLuc (RLuc8) with enhanced light output and serum inactivation resistance, making it a benchmark standard for many follow-up studies and RLuc-based assay development 11 . Download figure Open in new tab Figure 1: Overview of the luciferase-catalyzed reaction and bioluminescence production . ( a ) General scheme common for all luciferases, including the oxidation of the substrate (luciferin) into the product (oxyluciferin) and carbon dioxide (CO 2 ). Examples of specific substrate names and emission wavelengths for individual types of luciferases are stated above and below the reaction. ( b ) Renilla luciferase-catalyzed oxidative decarboxylation of coelenterazine (CTZ) into coelenteramide (CEI), CO 2 , and visible light emission with the maximum at ∼480 nm. The colored background corresponds to the color of the emitted light. In our recent analysis, we have identified that the dynamic behavior of RLuc8 is crucial for catalysis. This finding allowed us to generate a luciferase variant exhibiting highly stable light output with a 100-fold prolonged emission decay half-life 12 . This ‘glow-type’ variant, termed AncFT, was generated by transplanting a crucial fragment from RLuc8 to a reconstructed ancestral luciferase Anc HLD-RLuc , as reported by Chaloupkova et al. in 2019 13 . Furthermore, we have recently revealed the catalytic mechanism of RLuc8, identifying key enzyme residues involved in the reaction 14 . The mechanism also tracked individual chemical steps, capturing the substrate conversion from the enzyme-bound substrate state via multiple intermediates up to the enzyme-product state. However, despite such progress, no detailed mechanistic information was made available for the substrate binding and product release processes, i.e., the steps preceding and succeeding the described RLuc8 catalytic mechanism which covered only the oxidative decarboxylation chemistry steps. This gap currently represents one of the last missing puzzle pieces required to fully uncover the whole enzymatic pathway. Indeed, the previously identified significance of RLuc8 flexibility makes the exploration of these ligand transport steps paramount to allow even more targeted luciferase engineering. In this study, we elucidate a detailed kinetic pathway of RLuc8, describing the catalytic cycle from the initial enzyme-substrate collision up to the final product release. Our analysis thoroughly investigates all the elementary rate constants and the corresponding thermodynamic parameters involved. The outcome underscores the significance of the conformational dynamics, particularly highlighting a rate-limiting induced-fit step crucial to the overall catalytic process. Targeting this step presents a promising avenue to enhance the effectiveness of existing luciferases. Accordingly, the combination of this kinetic study with the insights from crystallographic and computational analyses provides a deeper understanding of the luciferase molecular principle. We leverage it to address and engineer the limiting induced-fit transition, resulting in a luciferase variant with superior bioluminescence properties compared to previously reported Renilla -type luciferases 11 – 16 . Results Oxygen-dependent kinetics provides updated luciferase steady-state parameters Kinetic analyses of all kinds of luciferases reported up to date mostly rely on varying only the luciferin substrate without assuming the effect of the oxygen co-substrate 11 – 18 . To gain a more detailed insight into the kinetics of RLuc8 catalysis, we systematically collected datasets varying both the CTZ and the oxygen substrates and globally analyzed these data using numerical simulations ( Figure 2 ). The obtained parameters ( Figure 2 , Figure S1 , Table SI ) were consistent with previously reported values 11 – 15 but further identified independent substrate binding sites and a very high value of Michaelis constant towards oxygen ( K m,O2 ) of 719 µM, substantially above the concentration of dissolved oxygen under physiological conditions (∼250 µM). This fact shifts the real turnover number k cat to more than 20 s - 1 , which is considerably higher than the apparent k cat,app value of ∼4–5 s - 1 reported up until now when only saturating CTZ concentration but physiological (i.e., not-saturating) oxygen concentration was taken into account ( Table SI ) 11 , 12 , 14 . Such findings indicated that increasing affinity towards oxygen may be an appealing strategy to enhance the luciferase performance. Redesigning the enzyme by engineering its affinity to oxygen to reach saturation even at physiological concentrations would yield up to 5-fold increased catalytic efficiency. Download figure Open in new tab Figure 2: Two-substrate oxygen-dependent steady-state kinetic analysis of the Renilla luciferase RLuc8. ( a– e ) Cumulative luminescence progress curves upon mixing RLuc8 with varying coelenterazine (CTZ) concentrations (yellow to dark blue curves) and fixed oxygen (O 2 ) concentrations. The resulting concentrations after the mixing were 8 nM RLuc8 and 0.2–15 µM CTZ. The resulting O 2 concentration is stated for each panel (a)–(e) in the top left corner of the graph. All the experiments were performed in 100 mM potassium phosphate buffer pH 7.5 at 37 °C. The data are shown as averages of 6 technical replicates and black solid lines represent the best fit according to the kinetic pathway represented in (f). ( f ) Extended steady-state kinetic pathway for independent binding of CTZ and O 2 and their conversion by the RLuc8 enzyme (E), leading to the final product (P) or the enzyme-inactive state (E’.X). The analysis allowed the determination of Michaelis constants ( K m ) for both substrates, turnover number ( k cat ), enzyme inactivation rate constant ( k inact ), and product inhibition equilibrium constant ( K p ). In contrast, the global analysis additionally detected slow irreversible inactivation of RLuc8 ( k inact = 0.015 s - 1 ) at high oxygen concentrations ( Figure 2 , Figure S1 , Table SI ). This was consistent with previous studies reporting a complete abolishment of other CTZ-utilizing luciferases after a defined number of catalytic cycles 19 – 22 . Herein identified pronounced inactivation at elevated oxygen levels points towards the involvement of oxygen radical species that have been recently proven to be involved in luciferase catalysis 14 . Linking these two events together via our observations provides a deeper understanding of this unusual, previously reported inactivation phenomenon. At the same time, this finding contradicts the aforementioned engineering strategy of increasing RLuc8 affinity towards oxygen. Although lowering the K m,O2 value would improve the luciferase catalytic efficiency, saturating the enzyme with oxygen would simultaneously intensify its inactivation, overall leading to an inefficient bioluminescence reporter. Altogether, the refined steady-state kinetic analysis provided useful new insights into the RLuc8 mechanism and identified critical weak spots. Yet, further detailed analysis was required to spotlight the true real limitation to be targeted by protein engineering. Transient kinetics reveals a detailed kinetic pathway and the rate-limiting step of the luciferase We achieved a comprehensive understanding of the kinetic pathway of the RLuc8 catalytic cycle via rigorous pre-steady-state global analysis of 28 datasets, including anaerobic substrate binding, product binding, time-resolved spectral, oxygen-dependent, and temperature-dependent transient kinetic experiments ( Figure 3 ). The data were collected by conventional stopped-flow and quench-flow devices but additionally, using an advanced continuous-flow microfluidic chip reported previously 23 . This device enabled high-throughput data collection with minimal sample consumption and rapid heat transfer, ideal for the systematic collection of transient kinetic and thermodynamic data. Analyzing all the data globally using a numerical simulation approach 24 , 25 allowed us to derive an extended detailed kinetic pathway ( Figure 4 , Figure S2 , Table SII , Supplementary Note 1 ) which identified multiple key features of the RLuc8 catalysis: (i) oxygen co-substrate can bind to any of the enzyme conformational states (either E, E.CTZ, or E*.CTZ); (ii) oxygen binding is independent of the CTZ substrate binding and/or the enzyme conformational change as the affinities towards the corresponding species ( K d,O2 ) are identical; (iii) both the CTZ substrate binding and the CEI product release occur via the induced-fit mechanism and the conformational transition, indicating substantial importance of protein dynamics in accordance with the previously reported data 12 ; (iv) initial binding/release of CTZ/CEI ( k ±1 / k ±5 ) is considerably faster compared to the coupled enzyme conformational change (closing/opening; k ±2 / k ±4 ); and namely (v) although the chemical substrate-to-product conversion step ( k +3 ) is slow, the real limitation lies in the follow-up step of the enzyme conformational change ( k +4 ) preceding the product release. Download figure Open in new tab Figure 3: Pre-steady-state transient kinetic data collected for the Renilla luciferase RLuc8 and their global numerical analysis. A collection of 28 datasets combines oxygen-dependent steady-state kinetics ( a ), anaerobic substrate binding kinetics ( b ), product release kinetics ( c ), multiple turnover kinetics ( d ), single turnover kinetics ( e ), oxygen-dependent burst kinetics ( f ), and on-chip temperature-dependent kinetics ( g ). The color of datapoints corresponds to the measured signal which is noted in the Y-axes and illustrated in the adjacent pictograms together with the experimental setup. Black solid lines represent the best fit of the global analysis yielding the extended kinetic pathway and rate/equilibrium constants displayed in Figure 4 . Experimental conditions (concentrations, temperature, buffer, replicates, etc.) are provided in the Online Methods section for each experiment individually. Temperatures and oxygen concentrations are also stated above each dataset. CTZ = coelenterazine; CEI = coelenteramide; Trp = tryptophan; SVD = singular value decomposition. Download figure Open in new tab Figure 4: Extended kinetic mechanism of the Renilla luciferase RLuc8. Key conformational changes during the catalysis, allowing the enzyme to switch between the open and the closed states, are highlighted with purple and blue colors, respectively. The rate-limiting step of the product-bound state conformational change is highlighted in red. ( a ) Graphical illustration of the uncovered enzyme (E; purple/blue blob) kinetic pathway describing the coelenterazine (CTZ; green hexagon) and oxygen (O 2 ; grey oval) substrates conversion into the coelenteramide (CEI; brown hexagon) product. The pathway includes independent binding of the substrates and critical conformational changes during the substrate binding and product release processes. Note that the carbon dioxide product is not illustrated in the pathway as it was not monitored in any of the experiments and thus no kinetic information is available. E = RLuc8 enzyme; CTZ = coelenterazine substrate; O 2 = oxygen co-substrate; CEI = coelenteramide product; E.CTZ and E*.CTZ = enzyme-coelenterazine complex in two different conformations; E.O 2 = enzyme-oxygen complex; E.CTZ.O2 and E*.CTZ.O2 = ternary enzyme-coelenterazine-oxygen complex in two different conformations; E.CEI and E*.CEI = enzyme-coelenteramide complex in two different conformations. ( b ) Reaction coordinate illustrating energy barriers of transitions between individual steps of the newly extended kinetic pathway, including the rate-limiting step marked with the red dashed arrow. ( c ) Values and standard errors of individual rate and equilibrium constants and activation energies of the newly extended kinetic pathway. The notion of the parameters is per those provided in (a). The reported values correspond to reaction kinetics at 15 °C in 100 mM potassium phosphate buffer pH 7.5. The last outcome of transient kinetics, pinpointing the rate-limiting conformational step, has delineated another focal point for subsequent engineering studies. It extended the conclusion of applying solely the steady-state analysis which suggested the modulation of the weak oxygen binding ( K d,O2 ). Notably, engineering the rate-limiting conformational dynamics does not appear to yield any conflicting side effects compared to irreversible inactivation complications associated with focusing on increasing oxygen affinity. Since we previously identified that the active-site loops of Renilla -type luciferases were responsible for extensive protein dynamics 12 , their engineering was selected as the most promising approach to target the limiting conformational step. Rational loop grafting generates a variant with favorable kinetic and bioluminescent properties Because flexible loops were identified as the most promising candidate elements to be engineered, we decided to exchange the loops between the herein deeply characterized extant luciferase RLuc8 and the homologous reconstructed luciferases Anc HLD-RLuc and AncFT, which we reported previously 12 , 13 . The loops were grafted from one scaffold to another rationally, employing the recently developed web tool LoopGrafter which takes into account loop geometries, lengths, dynamics, and energies 26 , 27 . LoopGrafter automatically identified the correlation motions of the loop L9 (grafted previously) and the loop L14 (grafted in this study). Using RLuc8 as a template scaffold and grafting its loop L14 into AncFT via LoopGrafter resulted in a variant AncFT-L14 (also termed AncFT7) with remarkably improved properties ( Figure 5 ). The circular dichroism spectrum ( Figure 5b ) remained similar (i.e., the enzyme was properly folded), its emission spectrum was comparable to AncFT ( Figure 5c ), and the melting temperature stayed higher than for RLuc8 ( Figure 5d , Figure S3 ), but catalytic parameters and light-emission properties of AncFT-L14 improved substantially. Catalytic efficiency k cat / K m of AncFT-L14 exceeded the values for both RLuc8 and AncFT and also the original ancestral enzyme Anc HLD-RLuc . Importantly, its product inhibition factor K p / K m improved significantly compared to the original variants ( Figure 5e , Figure S4 ). Such an increase indicated that we might have altered the targeted rate-limiting step of the product release process, explaining the observed increased preference for the substrate over the product ( K p / K m ). Crucially, the combination of catalytic efficiency and inhibition factors (i.e., k cat / K m × K p / K m ), herein termed as ‘glow-type effectivity’, was enhanced nearly 4-fold compared to AncFT, more than 20-fold compared to RLuc8, and more than 700,000-fold compared to Anc HLD-RLuc ( Figure 5e ). As a result, the newly constructed variant AncFT-L14 exhibited greater light emission stability in cell lysates compared to the already partially optimized variant AncFT ( Figure 5f ). Such an outcome makes AncFT-L14 an excellent candidate for bioluminescence assays requiring prolonged or continuous luminescence data collection. Download figure Open in new tab Figure 5: Sequence alignment and experimental characterization of selected luciferases. The comparison includes the 8-point mutant of the modern Renilla luciferase (RLuc8) 11 , reconstructed ancestral luciferase (Anc HLD-RLuc ) 13 , fragment-transplanted luciferase (AncFT) 12 , and the herein reported newly generated luciferase (AncFT-L14). ( a ) Multiple sequence alignment with highlighted regions (orange and purple rectangles with black outlines, respectively) that were rationally grafted to generate the novel luciferase AncFT-L14 with enhanced bioluminescence properties. Background of each residue corresponds to the Clustal X coloring scheme. ( b ) Circular dichroism spectra of all the tested variants. The experiment was measured in 100 mM potassium phosphate buffer pH 7.5 at room temperature and the data are presented as the average of 5 technical replicates. (c) Bioluminescence emission spectra of the characterized variants. No spectrum was collected for Anc HLD-RLuc due to the insufficient intensity of its light emission. The experiment was measured in 100 mM potassium phosphate buffer pH 7.5 at room temperature and the data are presented as the average of 3 technical replicates. (d) Thermal stability analysis determining onset temperatures T onset and melting temperatures T m for all the characterized luciferases. The experiment was measured in 100 mM potassium phosphate buffer pH 7.5 with the temperature ramp of 1 °C/min from 20 to 90 °C and the values are presented as best fit values ± standard errors. (e) Comparison of enzymology parameters, including the catalytic efficiency k cat / K m , product inhibition factor K p / K m , and ‘glow-type’ effectivity (product of the two), demonstrating the superiority of AncFT-L14 compared to other tested variants. The experiment was measured in 100 mM potassium phosphate buffer pH 7.5 at 37 °C and the values are presented as best fit values ± standard errors. ( f ) Bioluminescence decay measurements in cell lysates revealing the enhanced light emission stability of the ‘glow-type’ luciferase AncFT-L14 compared to AncFT and namely to RLuc8. No decay curve was collected for Anc HLD-RLuc due to the insufficient intensity of its light emission. The experiment was measured in 100 mM potassium phosphate buffer pH 7.5 at 37 °C and the data are presented as the average of 3 technical replicates. Integrative analysis explains the molecular basis of the luciferase improvement In order to gain deeper insights into the molecular basis of the improvement achieved by grafting the loop L14 from RLuc8 to AncFT, we subjected AncFT-L14 to a detailed analysis combining transient kinetics ( Figure 6 ), structural biology ( Figure 7 ), and in silico modeling ( Figure 8 ). Kinetic characterization revealed that grafting the loop L14 indeed affected the protein conformational flexibility by substantially improving the induced-fit step kinetics and resulting equilibrium ( Figure 6a , Figures S5 and S6 , Table SIII , Supplementary Note 2 ). Interestingly, the engineering had only a minor effect on the first step of the induced-fit binding mechanism, i.e., the initial enzyme-ligand collision ( K 1 ). On the contrary, the second step of the enzyme-ligand complex conformational change ( K 2 ) yielded highly favorable equilibrium and kinetics for AncFT-L14, concluding that we altered the targeted conformational flexibility nearly exclusively. Selective engineering of the induced-fit/conformational step is a challenging task rarely reported in the literature 28 – 32 which highlights the potential of the loop grafting strategy and the LoopGrafter tool 26 to yield innovative biocatalysts with favorable properties. Download figure Open in new tab Figure 6: Transient kinetic analysis of the induced-fit transport process steps for characterized variants Anc HLD-RLuc , AncFT, AncFT-L14, and RLuc8 . ( a ) Illustrative representation of the two steps involved in the induced-fit binding mechanism and the influence of the corresponding equilibrium constants on each of these steps. E.O 2 = enzyme-oxygen complex; CTZ = coelenterazine substrate; E.CTZ.O 2 and E*.CTZ.O 2 = ternary enzyme-coelenterazine-oxygen complex in two different conformations. The nomenclature and illustrations correspond to symbols used in Figure 3 . ( b ) Comparison of equilibrium constants characterizing individual steps of the induced-fit binding mechanism. K 2 value is not provided for Anc HLD-RLuc due to its rigid simple one-step binding and no conformational change observed upon the initial collision. Note the logarithmic scale of the y-axis. The experiment was measured in 100 mM potassium phosphate buffer pH 7.5 at varying temperatures (9 to 22 °C) and the values are presented as best fit values ± standard errors. Download figure Open in new tab Figure 7: Co-crystal structure of AncFT-L14 luciferase complexed with azacoelenterazine (azaCTZ). ( a ) Cartoon representation of the overall structure of AncFT-L14 (PDB ID: 7OME) enzyme (green) with bound azaCTZ (yellow spheres). The L9 loop is colored in violet, and the L14 loop is colored in orange. ( b ) 2Fo-Fc electron density (contour level 1.5 σ) for azaCTZ (yellow sticks) and selected residues (green sticks) at the active site pocket. ( c ) Close-up view of the azaCTZ binding in the AncFT-L14 active site. Residues creating the active site in green stick representation, azaCTZ in yellow stick representation, and key hydrogen bonds are shown as dashed yellow lines. ( d ) 2Fo-Fc electron density (contour level 1.5 σ) for the L14 loop (orange sticks) and a partial sequence (V 147 IESW) of the L9 loop (violet sticks). The azaCTZ in yellow sphere representation. ( e ) Close-up view of the molecular contacts between the L9 (violet) and L14 (orange) loops. ( f ) Structure-based partial sequence alignment of AncFT, AncFT-L14, and RLuc8. Secondary structure elements are shown above the alignment. The L14 loop sequence transplanted from RLuc8 into AncFT, leading to the creation of AncFT-L14, is labeled with an orange box. ( g - j ) Structural comparisons of the L9 and L14 loops conformations in AncFT (PDB ID: 7QXR) (g), AncFT-L14 (PDB ID: 7OME) (h), and RLuc8 (PDB ID: 2PSF) (i), and their corresponding superposition (j). Note that the L14 loop in AncFT-L14 adopts a unique conformation. Download figure Open in new tab Figure 8: Computational simulations of enzyme-ligand complexes of RLuc8, AncFT-L14, and Anc HLD-RLuc . ( a ) The potential of mean force used to pull coelenterazine (CTZ) into the active site of the luciferases obtained by adaptive steered molecular dynamics (ASMD). ( b ) A flipped CTZ end-pose (yellow sticks) from simulation of binding to Anc HLD-RLuc (red cartoon). The crystallographic ligand obtained for AncFT-L14 is shown as a reference (translucent gray sticks, PDB ID 7OME). ( c ) The equilibrium probabilities of coelenteramide (CEI) macrostates based on the CEI-active site distance (‘bound’, ‘tunnel’, ‘unbound’) derived from the adaptive sampling of CEI unbinding. The simulations were started from the crystal-like CEI pose for each enzyme plus from the flipped pose in Anc HLD-RLuc shown in (b). The error bars indicate the standard error of the equilibrium probability from bootstrapping a random 80 % of the data 100 times. ( d ) Analysis of access tunnel opening potential during the adaptive sampling of CEI unbinding. The macrostates (shown as bubbles) were constructed based on the width of the tunnel mouth; their equilibrium probability is written as a percentage inside the bubbles and reflected in their size. ( e – f ) The interaction energies of CEI per residue of the ‘bound’ CEI macrostate (e) and the ‘tunnel’ macrostate (f) from adaptive sampling with RLuc8 and AncFT-L14. For clarity, only attractive interactions are shown (less than −1.0 kcal.mol - 1 for the ‘bound’ states and less than −0.5 kcal.mol - 1 for the ‘tunnel’ states). The error bars indicate the standard error. ( g ) The cap domains of RLuc8, AncFT-L14, and Anc HLD-RLuc with mapped root-mean-square fluctuations (RMSF) of Cα atoms in the ‘unbound’ states from adaptive sampling. Next, we determined a co-crystal structure of AncFT-L14 complexed with the substrate analog azacoelenterazine (azaCTZ) ( Figure 7 , Table SIV , Supplementary Note 3 ) 14 , 33 . The structure was solved at 1.5 Å resolution and allowed us to elucidate the consequences of grafting the L14 loop. We observed that the overall binding mode and molecular contacts of azaCTZ in AncFT-L14 are very similar, if not identical, to the previously characterized AncFT luciferase ( Figure S7 ) 14 . However, careful inspection revealed an atypical conformation of the grafted L14 loop in AncFT-L14, distinguishing it from AncFT and RLuc8 ( Figure 7d–j ). Specifically, the transplanted sequence element L 223 VKGGKP is in proximity to the bound substrate analog (azaCTZ) but they do not make extensive physical contacts. Instead, the L14 loop makes numerous, mostly non-polar and hydrophobic contacts with the neighboring L9 loop. We previously demonstrated that the unique structural dynamics of the L9 loop has a dramatic effect on the enzyme performance 12 . Therefore, the improvement of the AncFT-L14 catalytic efficiency and bioluminescence properties may result from the structural complementarity and interface plasticity between the L9 and L14 loops. We think that their coordinated motions can be crucial for the substrate and product (un)loading during catalysis via enzyme conformational changes identified by kinetic analysis. Additionally, the structure obtained in this work determined at the higher resolution (1.5 Å) revealed some new molecular details about the luciferase catalysis. Notably, two water molecules are unambiguously resolved in the putative dioxygen-binding site ( Figure 7b-c ). The presence of these water molecules at the bottom of the active site pocket, close to the reactive C2 carbon atom of native CTZ, mimics the binding of dioxygen in a Michaelis enzyme-substrate complex ( Supplementary Note 3 ). Our structural observations thus highlight the plausible involvement of D118, N51, and W119 in the positioning of a co-substrate molecule (dioxygen) such that it can be directly attacked by the C2 carbon of activated CTZ. Moreover, hydrogen bonding is observed between the side chain of W119 and a carbonyl group of azaCTZ, evidencing an additional anchoring point for the stabilization of the enzyme-substrate complex ( Figure 7c , Figure S8 ). Collectively, crystallographic observations suggest that the improved catalysis of AncFT-L14 is not associated with an altered substrate positioning or transient state formation but because of an optimized geometry of flexible loops, including the transplanted loop L14. Alteration of the protein dynamics and ligand transport efficiency were further analyzed by molecular docking ( Figure S9 – S11 , Table SV ) and by simulating the CTZ substrate binding and the CEI product release using adaptive steered molecular dynamics (ASMD) and adaptive sampling molecular dynamics, respectively ( Figure 8 , Supplementary Note 4 and 5 ). Substrate binding was easiest when the CTZ starting pose adapted the crystal-like orientation for both AncFT-L14 and RLuc8 but binding to AncFT-L14 required 7 kcal.mol - 1 less work ( Figure 8a , Figures S12 and S13 ), in accordance with the higher affinity determined by kinetic analysis ( Figure 6 ). Product release analysis provided comparable trends for AncFT-L14 and RLuc8 by energetically favoring the unbound state of the product ( Figure 8c , Table SVI and SVII ) while the transport process steps, characterized by the clustered macrostates ‘bound’, ‘tunnel’, and ‘unbound’, differed notably ( Supplementary Note 5 , Figure S14–S17 ). RLuc8 showed more pronounced contacts with the access tunnel-lining residues of the dynamic loop L9 and α4 fragment ( Figure 8f , Figure S19 ). This element was moving to the side during the CEI release ( Figure 8g ), overall resulting in a conformation with a widely open tunnel in which RLuc8 remained a majority of the time ( Figure 8d ). In contrast, AncFT-L14 interacted on average more strongly with the active site residues compared to RLuc8 ( Figure 8e , Figure S18 ) and preferred the catalytically competent conformation with the closed tunnel ( Figure 8d ). This result is in agreement with our previous observation, supporting the interpretation that we selectively targeted the induced-fit conformational step by grafting the loop L14 from RLuc8 to AncFT 27 . The simulation for the original ancestral protein Anc HLD-RLuc yielded an unexpected outcome when compared to all other luciferases ( Supplementary Note 4 and 5 ). The ASMD simulation of substrate binding provided the most favorable results for the flipped, non-productive CTZ orientation ( Figure 8b ). Moreover, pulling CTZ into the active site by ASMD required twice as much energy in comparison to AncFT-L14 and RLuc8 ( Figure 8a ). Similarly, simulations of the CEI product release – in the orientation captured with RLuc8 – yielded significantly lower affinity for Anc HLD-RLuc by exhibiting a 90-fold lower probability of the ‘bound’ state compared to the ‘unbound’ state ( Figure 8c , Table SVI ). Strikingly, starting the CEI product release simulation from the flipped orientation, obtained by ASMD with CTZ ( Figure 8b ), resulted in a completely different outcome ( Figure 8 , Figure S17 and S20 , Table SVI and SVII ). The ‘bound’ state became significantly more probable compared to any other simulated protein and ligand, indicating high stability of the Anc HLD-RLuc .CEI complex in this flipped orientation ( Figure 8c , Table SVI and SVII ). Such a tight non-productive binding explains the substantial product inhibition observed for Anc HLD-RLuc previously 12 . Furthermore, the product inhibition has been linked to the bioluminescence flash. Therefore, the strong preference of AncFT-L14 towards the properly oriented substrate/product, with no signs of the flipped orientation, explains the mechanism behind generating this new variant with highly stable, ‘glow-type’ bioluminescence emission. Discussion and Conclusions In addition to being remarkable light-emitting machines in nature, luciferases have become invaluable tools in research and clinical diagnostics as bioreporters and bioimaging agents 4 – 6 . Despite their enduring popularity for over half a century, our understanding of their structure-function relationships and detailed molecular mechanisms has only recently begun to unfold. Renilla -type luciferase catalytic mechanism was revealed only in 2023 14 and although it provided extremely useful information on snapshots following the substrate-to-product conversion inside the enzyme active site, the elementary steps of the substrate binding and the product release processes remained elusive. Here, we tackled the problem by extending steady-state kinetic data with oxygen-dependent, temperature-dependent, and spectral pre-steady-state datasets followed by their complex global analysis using numerical integration. Our results revealed an unusually high value of the Michaelis constant for the oxygen co-substrate, markedly exceeding the concentration of dissolved oxygen at physiological conditions. These results demonstrate that the enzyme was never fully saturated by both the substrates in the previous kinetic studies, and so the reported k cat values are only apparent k cat,app parameters at the saturating CTZ substrate, but non-saturating oxygen concentrations 11 , 12 , 14 . The real k cat value is 21.9 s - 1 . This result was verified by using real k cat to back-calculate k cat,app at the physiological dissolved oxygen concentration (∼ 250 µM). This resulted in obtaining a value of 5.7 s - 1 which well approximates the previously reported values (4.5–5.1 s - 1 ) 11 , 12 , 14 . Employing high concentrations of dissolved oxygen allowed us to resolve a slow irreversible inactivation step which has never been observed for RLuc8. On the other hand, similar behavior and enzyme inactivation after a given number of turnover cycles was previously reported for native RLuc 20 , NanoKAZ/NanoLuc 21 , and Gaussia luciferase (GLuc) 19 , validating our outcome. We suggest that the consensus-based mutagenesis of RLuc, leading to a more resistant variant RLuc8 11 , also decreased its susceptibility to inactivation so it was undetectable under physiological conditions. The fact that the inactivation is more pronounced with increasing oxygen concentration clearly points towards an oxygen-induced mechanism of inactivation which is also in accordance with the previously reported 17 Da adducts detected for inactivated GLuc 22 . These observations are in line with the recently revealed catalytic mechanism of CTZ bioluminescence proceeding via highly reactive oxygen radical intermediates 14 , likely the source of enzyme covalent modification and inactivation. The luciferases have apparently evolved to critically balance their affinities towards oxygen to yield optimal performance. Low affinity results in the undersaturation of the enzyme, preventing it from reaching its full catalytic performance and light emission. Conversely, high affinity leads to the excessive formation of oxygen radicals gradually inactivating the enzyme. This inactivation adversely impacts bioimaging applications with extended acquisition times. The transient kinetic analysis identified another constraint of the RLuc8 catalysis – its conformational change during the product release is the rate-limiting step of the overall enzymatic cycle. Targeting selectively this conformational step is a difficult task because it requires altering the protein dynamics without impairing the chemical transformation and ligand collision steps. This is evidenced by only very few examples of such engineering work reported in the scientific literature 28 – 32 . We addressed this issue by grafting multiple mobile loops from the modern luciferase protein into the reconstructed ancestral luciferase using the LoopGrafter tool 26 , 27 . Detailed analysis by kinetics, X-ray crystallography, and molecular dynamics confirmed that engineering of the protein via the grafting approach yielded the desired improvement of the conformational step by altering the protein flexibility around the access tunnel region. Interestingly, the first step of the induced-fit binding mechanism, i.e., the initial protein-substrate collision, remained nearly unaffected. The detailed comparison of individual luciferase variants by molecular modeling has yielded an additional view of the structure-function relationship. The simulations identified high energy barriers for ligand binding by Anc HLD-RLuc , in accordance with exceptionally low affinities observed by transient kinetics and the lack of successful ligand co-crystallization attempts. Unexpectedly, the simulations favored the non-productive flipped orientation of the CTZ substrate and the CEI product inside the active site of the ancestral protein. Once CEI is bound in this orientation, a significant amount of energy is required to expel it from the active site, which is in agreement with the substantial product inhibition reported for Anc HLD-RLuc previously 12 . The new insights gained from molecular dynamics raise a question about the catalytic mechanism of Anc HLD-RLuc . We hypothesize that the enzyme may bind the CTZ substrate inside its active site in the non-productive orientation but provides enough of a hydrophobic environment for the spontaneous CTZ decomposition associated with weak luminescence, leading to the very low catalytic activity measured. This hydrophobic site is required for efficient light emission to avoid luminescence quenching by water molecules and to minimize thermal dissipation by restricting CTZ degrees of freedom 34 , 35 . Similar chemiluminescence-like CTZ decomposition leading to a weak emission has been described in the presence of albumin 36 , 37 as well as in aprotic solvents such as DMSO or DMF 14 , 38 – 40 . The mechanism based on the ‘scaffolding and desolvation’ effect, followed by a spontaneous substrate conversion inside the hydrophobic environment, could represent the point of evolutionary divergence. We speculate that such a phenomenon would serve as the very first stage of the bioluminescence activity emergence from different enzyme classes, as observed for multiple luciferase enzymes across diverse kingdoms of life 13 , 41 – 47 . In conclusion, this study describes an extended molecular basis for the Renilla luciferase catalysis with the full kinetic pathway. It identifies critical limitations in the catalytic cycle whose elimination promises the development of industrially applicable variants surpassing the enzymes used in state-of-the-art bioluminescence assays. This is demonstrated by rationally engineering the variant AncFT-L14 (AncFT7) with high catalytic efficiency and stable glow-type bioluminescence. The follow-up integrative analysis provides an even more robust exploration of the luciferase function at the molecular level, validating the proposed mechanism and the loop grafting strategy towards glow-type bioluminescence. Our engineered enzymes exhibits substantially more stable signal than the natural enzyme while being less prone to inactivation, paving the way for diverse bioluminescence applications requiring a long-lasting and stable light emission signal. Materials and Methods Luciferin and aza-luciferin molecules The coelenterazine (CTZ) luciferin was purchased from P-LAB (cat. no. R4094.3). Substrate analogue azacoelenterazine (azaCTZ) was synthesized as described previously 14 . Loop transplantation to create AncFT-L14 The mutant construct was generated using standard PCR-based nested protocols and inserted into the pET21b expression vector between NdeI and BamHI sites. Briefly, the L14 loop exchange mutant was designed based on sequence and structural comparison. The correlated motions of loop L9 and loop L14 were identified using the in-house web-based tool LoopGrafter 26 , 27 . The RLuc L14 loop sequence L 235 VKGGKP was introduced instead of the corresponding element encompassing I 223 KGDGPE sequence in AncFT 12 to create the AncFT-L14 (also termed AncFT7) mutant. Loop transplantation was carried out in two-step PCR using Phusion polymerase (NEB, UK) according to the manufacturer’s protocol. The list of the used primers is available in Table I . After the mutagenesis reaction, the original template was removed by DpnI (NEB, USA) treatment (2 h at 37°C), followed by the DpnI inactivation (20 min at 80°C). The resulting plasmids were transformed into chemocompetent E. coli DH5α cells, plated on LB-agar (tryptone 20 g.L - 1 , yeast extract 10 g.L - 1 , NaCl g.L - 1 , agar g.L - 1 ) containing ampicillin (100 µg.mL - 1 ), and incubated at 37 °C overnight. Plasmids were isolated from three randomly selected colonies, and error-free sequences were verified by DNA sequencing (Eurofins Genomics, Germany). View this table: View inline View popup Download powerpoint Table I: Nucleotide primers used in PCR mutagenesis to generate AncFT-L14. Overproduction and purification of Anc HLD-RLuc , AncFT, AncFT-L14, and RLuc8 Overexpression of proteins from pET21b plasmid (amp R ) was carried out in E. coli BL21 cells (NEB, USA) cultivated in LB medium supplemented with ampicillin (100 μg.mL - 1 ). Once the culture OD 600 reached ∼0.6, protein production was induced at 20 °C by adding IPTG to a final concentration of 0.5 mM. The cells were then harvested after 16 h incubation by centrifugation (4,000 rpm, 20 min, 4 °C), and the cell pellets were resuspended in 20 mM potassium phosphate buffer pH 7.5 containing 500 mM NaCl and 10 mM imidazole and stored at −80°C. Prior to purification, the thawed cell mass was disrupted by sonication using Sonic Dismembrator Model 705 (Fisher Scientific, USA). Lysates were clarified by centrifugation (14,000 rpm, 1 h, 4 °C) using a Sigma 6-16K centrifuge (SciQuip, UK) equipped with a 12166 rotor. Supernatants containing the recombinant His-tagged proteins at their C-terminal ends were metal-affinity purified using Ni-NTA Superflow Cartridge 5mL (Qiagen, Germany) installed on FPLC system (Bio-Rad Laboratories, USA) and equilibrated with the 20 mM potassium phosphate buffer pH 7.5 containing 500 mM NaCl and 10 mM imidazole. Proteins were eluted with imidazole gradient and monomers were separated using gel permeation chromatography on Äkta FPLC (GE Healthcare, Sweden) equipped with HiLoad TM 16/600 Superdex TM 200 pg column (GE Healthcare, Sweden) and equilibrated with 10 mM Tris-HCl buffer pH 7.5 containing 50 mM NaCl. Purified proteins were concentrated to final concentrations using Centrifugal Filter Units Amicon R Ultra-15 Ultracel R -10K (Merck Millipore Ltd., Ireland). The purity of proteins was verified on SDS-PAGE. The concentration of protein samples was measured using DeNovix R DS-11 Spectrophotometer (DeNovix Inc., USA). Circular dichroism spectroscopy Circular dichroism spectra were recorded at 20 °C using the Chirascan spectropolarimeter (Applied Photophysics, UK). The signal was measured from 185 to 260 nm at a rate of 100 nm.min - 1 , with 1.0 s integration time, and 1 nm bandwidth in a 0.1 cm quartz cuvette. The spectrum was measured in five consecutive technical replicates and then averaged and corrected for the ellipticity of the buffer. The circular dichroism signal was expressed as mean residue ellipticity Θ MRE , calculated using Equation 1 where Θ obs is the measured ellipticity in degrees, M W is the protein molecular weight in kDa, n is the number of protein residues, l is the cell path length in cm, and c is the protein concentration in mg.mL - 1 . Thermal stability measurements Thermal unfolding was studied using NanoDSF Prometheus (NanoTemper, Germany) by monitoring tryptophan fluorescence over the temperature range of 20 to 90 °C, at a heating rate of 1 °C/min. The onset and melting temperatures ( T onset and T m , respectively) were evaluated directly by the manufacturer-provided software ThermControl v2.0.2. Bioluminescence emission spectral analysis Bioluminescence emission spectra were measured at ambient temperature using the spectrofluorometer FluoroMax-4 (HORIBA, Japan). 1 mL of an enzyme solution in 100 mM potassium phosphate buffer pH 7.5 was added into a quartz cuvette and the enzymatic reaction was initiated by manual injection of 50 µL of ethanolic solution of coelenterazine followed by a quick mixing. The acquisition of luminescence emission spectra started after 10 seconds from the addition of the coelenterazine substrate. The resulting concentration of an enzyme in the reaction mixture ranged from 0.7 to 1.0 µM while the concentration of coelenterazine was approximately 30 µM. The data were collected for wavelengths from 350 to 700 nm with a step of 2 nm and each spectrum was measured in triplicates. The obtained spectrum was normalized relative to the value of the highest peak for each of the enzymes individually and the shape and positions of the luminescence emission peaks were compared. Bioluminescence emission decay and stability NIH/3T3 mouse fibroblast cells (ATCC ® CRL-1658™) were transfected according to the manufacturer protocol using Lipofectamine 2000 (Thermo Fisher, USA) with pcDNA3.1(+) plasmids containing the genes of luciferases codon-optimized for expression in mammalian cells (Gene Art, Thermo Fisher, USA). Cells were lysed 24 hours after the transfection, and the bioluminescence signal in the lysate was measured with a microplate reader FLUOstar Omega (BMG Labtech, Germany) using the commercial Renilla Luciferase Assay System (Promega, USA) and also using an in-house prepared assay mixture composed of 100 mM potassium phosphate buffer pH 7.5 with 4.5 µM coelenterazine (final concentration). Cells transfected with an empty pcDNA3.1(+) plasmid were used as a negative control. The measurements were done in three independent replicates. Conventional steady-state kinetic experiments (physiological oxygen atmosphere) Solid coelenterazine was dissolved in ice-cold ethanol and stored under a nitrogen atmosphere in dark glass vials at –20 °C. Before each measurement, the concentration and maintained quality of the ethanol stock solutions were verified spectrophotometrically. A buffer solution of coelenterazine was prepared by mixing an appropriate volume of the ethanol stock solution with 100 mM phosphate buffer pH 7.5 immediately before the measurement. Tested luciferase enzyme samples were diluted in the same 100 mM phosphate buffer pH 7.5. The enzymatic reaction was initiated inside the microplate reader FLUOstar OPTIMA (BMG Labtech, Germany) by the automatic addition of 225 µL buffer solution of coelenterazine into a microplate well containing 25 µL of enzyme solution. The resulting bioluminescence activity traces were collected at 37 °C until the luminescence intensity decreased to less than 0.5 % of its maximal measured value to ensure the full conversion of coelenterazine. Each luminescence trace was measured in 3 repetitions. The concentration of coelenterazine in the final reaction mixture ranged from 0.2 to 7.9 µM while the concentration of luciferase enzymes ranged from 17 to 640 nM. Oxygen-dependent kinetic experiments 100 mM potassium phosphate buffer pH 7.5 and water solutions were incubated in the anaerobic glovebox Belle MR2 (Belle Technology, UK) overnight while stirring to remove dissolved oxygen. The glovebox was filled with ultra-high purity (UHP) nitrogen atmosphere containing a residual oxygen concentration of 2.4 ppm, as determined by the oxygen meter O2M-3 (Belle Technology, UK). A bottle with lyophilized evacuated (oxygen-free) RLuc8 protein was put into the anaerobic glovebox as well, it was dissolved with anaerobic water and incubated for at least one hour to remove possible traces of dissolved oxygen. Finally, a nitrogen-flushed aliquot of CTZ dissolved in pure ethanol was put into the glovebox, incubated for approximately 20 minutes to remove dissolved oxygen, and then closed to prevent further evaporation of the sample. Aerobic buffer solutions of controlled oxygen concentrations were prepared by filling 100 mM potassium phosphate buffer pH 7.5 into glass tonometers and flushing/bubbling the solution with a gas mixture containing different ratios of oxygen and nitrogen for at least one hour. Afterward, the tonometers were hermetically closed, put into the anaerobic glovebox, and kept closed before loading into the Stopped-Flow instrument to prevent leakage of oxygen into the glovebox. Oxygen concentration in the tonometer solutions was calculated based on Henry’s Law using known experimental temperature and pressure. CTZ stock solution was diluted with 100 mM potassium phosphate buffer pH 7.5 and then manually premixed anaerobically with the RLuc8 enzyme to reach the desired concentration of both components. The premixed RLuc8.CTZ complex was loaded into the first port of the Stopped-Flow instrument KinetAsyst SF-61DX2 (TgK Scientific, UK) while the second one was filled with the aerobic buffer from the prepared tonometer. The two solutions were rapidly mixed in a 1:1 ratio and the enzymatic reaction was followed by an increase of the luminescence signal over 60 seconds at 37 °C. The port filling was repeated for 10 different CTZ concentrations and 5 different oxygen concentrations in a fully combinatorial manner (50 different conditions measured in total). The resulting concentrations during the experiments were 8 nM RLuc8, 0.2–15 µM CTZ, and 61–829 µM oxygen. By applying logarithmic distribution of data points collection (high frequency at the beginning and lower frequency at the end of the collection), both pre-steady-state and steady-state phases of the reaction could be accurately monitored. Steady-state kinetic data analysis The obtained kinetic curves were integrated in order to obtain changes of cumulative luminescence over time that are proportional to the product formation increase with time. The resulting curves were analyzed by applying an updated protocol employing global numerical fitting of raw kinetic data using KinTek Explorer 10 (KinTek Corporation, USA). The software allows for the input of a given kinetic model via a simple text description, and the program then derives the differential equations needed for numerical integration automatically. Numerical integration of rate equations searching a set of kinetic parameters that produce a minimum χ 2 value was performed using the Bulirsch–Stoer algorithm with adaptive step size, and nonlinear regression to fit data was based on the Levenberg– Marquardt method. Residuals were normalized by sigma value for each data point. The standard error (S.E.) was calculated from the covariance matrix during nonlinear regression. In addition to S.E. values, a more rigorous analysis of the variation of the kinetic parameters was accomplished by confidence contour analysis using FitSpace Explorer (KinTek Corporation, USA). In these analyses, the lower and upper limits for each parameter were derived from the confidence contour obtained from setting χ 2 threshold at 0.95. In the case of oxygen-dependent data, a general random order binding two-substrate steady-state model with irreversible enzyme inactivation ( Figure 9 ) was used to obtain realistic values of turnover number k cat , Michaelis constants K m,CTZ and K m,O2 , product inhibition constant K p , and inactivation rate constant k inact while in the case of ‘conventional’ (physiological oxygen atmosphere) experiments, a simpler minimal steady-state model ( Figure 10 ) was used to derive k cat , K m,CTZ , and K p . A conservative estimate for diffusion-limited substrates and product binding k +1 , k +2 , and k -4 (100 μM - 1 .s - 1 ) was used as a fixed value to mimic the rapid equilibrium assumption. Download figure Open in new tab Figure 9: Kinetic scheme of the random order two-substrate binding steady-state model. Except for the turnover number k cat and Michaelis constant K m for both substrates, the model also includes an irreversible enzyme inhibition and product inhibition, described by k inact and K p , respectively. Calculation of the steady-state kinetic parameters from the respective elementary rate constants is provided below the kinetic scheme. Download figure Open in new tab Figure 10: Kinetic scheme of the simple minimal steady-state model. The model is capable of accurately determining values of the turnover number k cat , Michaelis constant for the coelenterazine (CTZ) substrate K m,CTZ , and product inhibition constant K p . Calculation of the steady-state kinetic parameters from the respective elementary rate constants is provided below the kinetic scheme. Tryptophan fluorescence and luminescence stopped-flow kinetic experiments Transient kinetic traces upon the coelenterazine (CTZ) substrate conversion by the RLuc8 enzyme were monitored after rapidly mixing the components using the Stopped-Flow SFM 3000 mixing system equipped with a xenon arc lamp light source and the MOS-500 spectrometer (BioLogic, France). In each case, the reaction was initiated by mixing 75 µL of the enzyme solution with 75 µL of the CTZ solution with a total flow rate of 13 mL.min - 1 and then monitored in two modes using two different sources of the optical signal: (i) native tryptophan fluorescence and (ii) bioluminescence. Quenching of the tryptophan fluorescence, caused by the CTZ substrate binding in the active site, was monitored using the 340 ± 13 nm band-pass emission filter after the excitation at 295 nm. Bioluminescence signal changes, direct output of the luciferase reaction upon the substrate conversion, were monitored without applying any emission filter and no external excitation light source. The experiment was performed in 100 mM potassium phosphate buffer pH 7.5 at either 15 °C or 37 °C in three different experimental setups: (i) multiple turnover (excess of the substrate) varying the CTZ concentration; (ii) multiple turnover varying the RLuc8 concentration; and (iii) single turnover experiment (excess of the enzyme). Each kinetic trace was collected in 6 consecutive technical replicates and then averaged. The resulting concentrations after mixing were 2 µM RLuc8 and 1.3–64 µM CTZ in the CTZ-varying multiple turnover experiment, 15 µM CTZ and 1.8–94 µM RLuc8 in the RLuc8-varying experiment, and 50 µM RLuc8 and 24 µM CTZ in the single turnover experiment. Product fluorescence stopped-flow kinetic experiments Transient kinetic traces upon the coelenteramide (CEI) product binding into the RLuc8 enzyme active site were monitored after rapidly mixing the components using the Stopped-Flow SFM 3000 mixing system equipped with a xenon arc lamp light source and the MOS-500 spectrometer (BioLogic, France). The measurement was initiated by mixing 75 µL of the enzyme solution with 75 µL of the CEI solution with a total flow rate of 13 mL.min - 1 and then monitored by the change of the product fluorescence upon binding using the 375 nm long-pass emission filter after the excitation at 330 nm. The experiment was performed in 100 mM potassium phosphate buffer pH 7.5 at 15 °C and each kinetic trace was collected in 6 consecutive technical replicates and then averaged. The resulting concentration of CEI after mixing was 10 µM while the concentration of RLuc8 varied from 1.2 to 342 µM to generate a concentration series. Anaerobic tryptophan and substrate fluorescence stopped-flow kinetic experiments 100 mM potassium phosphate buffer pH 7.5 and water solutions were incubated in the anaerobic glovebox Belle MR2 (Belle Technology, UK) overnight while stirring to remove dissolved oxygen. The glovebox was filled with ultra-high purity (UHP) nitrogen atmosphere containing a residual oxygen concentration of 2.2 ppm, as determined by the oxygen meter O2M-3 (Belle Technology, UK). A bottle with lyophilized evacuated (oxygen-free) RLuc8 protein was put into the anaerobic glovebox as well, it was dissolved with anaerobic water and incubated for at least one hour to remove possible traces of dissolved oxygen. Finally, a nitrogen-flushed aliquot of CTZ dissolved in pure ethanol was put into the glovebox, incubated for approximately 20 minutes to remove dissolved oxygen, and then closed to prevent further evaporation of the sample. CTZ stock solution and reconstituted RLuc8 solution were then diluted with the anaerobic 100 mM potassium phosphate buffer pH 7.5 to the desired concentrations to generate the initial samples for the stopped-flow experiments. The solutions were subsequently loaded into the Stopped-Flow instrument KinetAsyst SF-61DX2 (TgK Scientific, UK) that was placed in the same anaerobic glovebox Belle MR2 (Belle Technology, UK). The experiment was initiated by rapidly mixing the components in a 1:1 ratio and then monitored in two modes using two different sources of the optical signal: (i) native tryptophan fluorescence and (ii) native substrate fluorescence. Quenching of the tryptophan fluorescence, caused by the CTZ substrate binding in the active site, was monitored using the 313–430 nm band-pass emission filter after the excitation at 295 nm. The increase of the substrate fluorescence, caused by binding into the rigidifying hydrophobic active site of the enzyme, was monitored using the 445 nm long-pass emission filter after the excitation at 415 nm. The experiment was performed in 100 mM potassium phosphate buffer pH 7.5 at either 25 °C or 7 °C and each kinetic trace was collected in 6 consecutive technical replicates and then averaged. The resulting concentration of RLuc8 after mixing was 2 µM while the concentration of CTZ varied from 0.4 to 74 µM to generate a concentration series. Time-resolved spectral absorbance stopped-flow kinetic experiments Transient kinetic traces upon the coelenterazine (CTZ) substrate conversion/binding by the RLuc8 enzyme were monitored after rapidly mixing the components using the Stopped-Flow instrument KinetAsyst SF-61DX2 (TgK Scientific, UK). The measurement was initiated by mixing the enzyme and the CTZ substrate solution in a 1:1 ratio and then monitored by the change of absorbance spectra over time using the KinetaScan CCD Detector (TgK Scientific, UK). The measured spectral changes originated from (i) CTZ/CEI (un)binding by the enzyme and (ii) CTZ-to-CEI conversion. The experiment was performed in 100 mM potassium phosphate buffer pH 7.5 at either 7 °C or 37 °C in two different experimental setups: (i) standard aerobic mixing monitoring the whole catalytic cycle; (ii) anaerobic mixing (details on anaerobization of the samples provided in the previous section) to dissect only the initial kinetic phases of the substrate binding with no follow-up conversion. Each kinetic trace was collected in 6 consecutive technical replicates and then averaged. The resulting concentrations after mixing were 74 µM RLuc8 and 64 µM CTZ in the aerobic experiments and 25 µM RLuc8 and 64 µM CTZ in the anaerobic experiment. Temperature-dependent rapid-mixing microfluidic chip kinetic experiments Transient kinetic traces upon the coelenterazine (CTZ) substrate binding by luciferase enzymes (Anc HLD-RLuc , AncFT, AncFT-L14, and RLuc8) at different temperatures were collected after rapidly mixing the components using the advanced continuous-flow microfluidic chip allowing high-throughput data collection with minimal sample consumption and rapid heat-transfer in the setup described and published previously 23 . The measurement was initiated by mixing the luciferase enzyme solution with the CTZ solution at the total flow rate of 2.1 µL.min - 1 and then monitored by the change of the substrate fluorescence upon binding using the 488 nm long-pass emission filter after the excitation at 405 nm. The experiment was performed in 100 mM potassium phosphate buffer pH 7.5 at temperatures varying from 9 to 22 °C for AncFT, AncFT-L14, and RLuc8 and from 22 to 49 °C for Anc HLD-RLuc . The resulting concentration of CTZ after mixing was kept constant at 30 µM while the concentrations of the luciferase enzymes varied from 7 to 67 µM for Anc HLD-RLuc , from 10 to 94 µM for AncFT, from 6 to 57 µM for AncFT-L14, and from 7 to 76 µM for RLuc8. Global numerical analysis of transient kinetic data All the transient kinetic curves obtained by the methods described in the corresponding sections above were directly analyzed globally by numerical fitting using KinTek Explorer 10 (KinTek Corporation, USA). The software allows for the input of a given kinetic model via a simple text description, and the program then derives the differential equations needed for numerical integration automatically. Numerical integration of rate equations searching a set of kinetic parameters that produce a minimum χ 2 value was performed using the Bulirsch–Stoer algorithm with adaptive step size, and nonlinear regression to fit data was based on the Levenberg–Marquardt method. Residuals were normalized by sigma value for each data point. The standard error (S.E.) was calculated from the covariance matrix during nonlinear regression. In addition to S.E. values, a more rigorous analysis of the variation of the kinetic parameters was accomplished by confidence contour analysis using FitSpace Explorer (KinTek Corporation, USA). In these analyses, the lower and upper limits for each parameter were derived from the confidence contour obtained from setting χ 2 threshold at 0.95. During the analysis procedure, multiple kinetic schemes were tested until the minimal pathway accurately accounting for all the collected data was identified together with the corresponding rate constants and activation energies. The whole process of data fitting from starting kinetic curves and defined observables up to the final kinetic scheme is described more in detail in Supplementary Note 1 . Co-crystallization assays Crystallization experiments were done at 20 °C using the hanging-drop vapor-diffusion method in EasyXtal 15-well plates (Qiagen, Germany) with drops equilibrated against 500 μL of reservoir solution. For co-crystallization, AncFT-L14 enzyme concentrated to ∼9 mg.mL - 1 was mixed with azaCTZ 14 in a 1:2 molar enzyme-ligand ratio. Crystals were obtained after mixing 1 μL of enzyme-ligand mixture with 1 μL of the precipitant solution consisting of 0.1 M SPG (succinic acid, sodium phosphate monobasic monohydrate, and glycine) buffer pH 9 and 25 % PEG 1500. The crystals were harvested after 5 to 7 days of incubation. All crystals were fished out, cryo-protected in the corresponding reservoir solution supplemented with 20 % glycerol, and cryo-cooled in liquid nitrogen for X-ray data collection. Diffraction data collection and data processing X-ray data were collected at PXIII beamline at SLS Synchrotron (Villigen, Switzerland) at the wavelength of 0.999 Å using a Pilatus 2M-F detector. The data were processed using XDS 48 , and Aimless 49 was used for data merging. Initial phases were solved by molecular replacement using Phaser 50 implemented in Phenix 51 with AncFT (PDB ID: 7QXR) 14 employed as a search model. The refinement was carried out in cycles of automated refinement in phenix.refine program 51 and manual model building in Coot 52 . The final models were validated using tools provided by Coot 52 and MolProbity 53 . Visualizations of structural data were created using PyMOL 2.0 (Schrödinger LLC, USA). Structural superposition was carried out using the secondary structure matching (SSM) superimpose tool in the Coot 52 . Atomic coordinates and structure factors of the enzyme-ligand complex were deposited in the Protein Data Bank ( www.wwpdb.org ) 54 under the PDB code 7OME. Preparation of ligands and enzyme structures for in silico simulations The structures of CTZ (substrate) and CEI (product) were prepared using Avogadro 1.2.0 software 55 : the multiplicity of the bonds was edited to match the keto forms, all missing hydrogens were added, and the structures were minimized by the steepest descent algorithm in the Auto Optimize tool of Avogadro, using the Universal Force Field (UFF). For the molecular docking, the restrained electrostatic potential (RESP) charges of the ligands were derived by the RESP ESP charge Derive (R.E.D.) Server Development 2.0 56 . Next, the AutoDock atom types were added, and PDBQT files were generated by MGLTools 57 , 58 . For the molecular dynamics simulations, the antechamber module of AmberTools16 59 was used to calculate the charges for the ligands, add the atom types of the Amber force field, and compile them in a PREPI parameters file. Also, the parmchk2 tool from AmberTools16 was used to create additional FRCMOD parameter files to compensate for any missing parameters. The crystal structures of RLuc8, AncFT-L14, and Anc HLD-RLuc (chains A of PDB IDs 2PSF, 7OME, and 6G75, respectively) were downloaded from the RCSB Protein Data Bank 54 and stripped of all HETATM records (non-protein atoms). The structures were aligned to chain A of 2PSF in PyMOL 2.5.4 (Schrödinger LLC, USA) and protonated with H++ web server v. 4.0 60 , 61 , using pH = 7.5, salinity = 0.1 M, internal dielectric = 10, and external dielectric = 80 as parameters. For the subsequent docking, AutoDock atom types and Gasteiger charges were added to the enzymes by MGLTools 57 , 58 , and the corresponding PDBQT files were generated. Molecular docking The AutoDock Vina 1.1.2 62 software was used for molecular docking. For docking to the active site, the docking grid was specified as an x = 33.00 Å, y = 33.38 Å, z = 32.62 Å sized box, with a center in x = 47.83, y = 21.98, z = 12.41 Å, covering the catalytic pocket and access tunnels. The flag -- exhaustiveness = 100 was used to sample the possible conformational space thoroughly. The number of output conformations of the docked ligand was set to 10. The results were analyzed in PyMOL 2.5.4 (Schrödinger LLC, USA). System preparation and equilibration for ASMD RLuc8, AncFT-L14, and Anc HLD-RLuc structures were prepared as described above. Then, the crystallographic water molecules were added back into the systems, except those overlapping with the protein or ligand. Next, histidine residues were renamed according to their protonation state (HID – Nδ protonated, HIE – Nε protonated, HIP – both Nδ and Nε protonated). The ligand was prepared as described in the corresponding section above. Twelve systems were prepared for the CTZ binding analysis: four different CTZ poses were placed at the tunnel mouth of each studied enzyme using PyMOL (Schrödinger LLC, USA). The tLEaP module of AmberTools16 59 was used to neutralize the systems with Cl − and Na + ions, import the ff14SB force field 63 to describe the protein and the PREPI file parameters to describe the ligand, add a truncated octahedral box of TIP3P water molecules 64 to the distance of 10 Å from any atom in the system, and generate the topology file, coordinate file, and PDB file. The system equilibration was carried out with the PMEMD.CUDA 65 – 67 module of Amber 16 59 . Five minimization steps and twelve steps of equilibration dynamics were performed. The first four minimization steps comprised 2,500 cycles of the steepest descent algorithm followed by 7,500 cycles of the conjugate gradient algorithm, while harmonic restraints gradually decreased. The restraints were applied as follows: 500 kcal.mol - 1 .Å - 2 on all heavy atoms of the protein and ligand, and then 500, 125, and 25 kcal.mol - 1 .Å - 2 on protein backbone atoms and ligand heavy atoms. The fifth step comprised 5,000 cycles of the steepest descent algorithm followed by 15,000 cycles of the conjugate gradient algorithm without restraint. The equilibration MD simulations consisted of twelve steps: (i) The first step involved 20 ps of gradual heating from 0 to 298 K at constant volume using Langevin dynamics, with harmonic restraints of 200 kcal.mol - 1 .Å - 2 on all heavy atoms of the protein and ligand, (ii) ten steps of 400 ps equilibration Langevin dynamics each at a constant temperature of 298 K and a constant pressure of 1 bar with decreasing harmonic restraints of 150, 100, 75, 50, 25, 15, 10, 5, 1, and 0.5 kcal.mol - 1 .Å - 2 on protein backbone and ligand heavy atoms, and (iii) the last step involving 400 ps of equilibration dynamics at a constant temperature of 298 K and a constant pressure of 1 bar with no restraint. The simulations employed periodic boundary conditions based on the particle mesh Ewald method 68 for treatment of the long-range interactions beyond the 10 Å cut-off, the SHAKE algorithm 69 to constrain the bonds that involve hydrogen atoms, the Berendsen barostat 70 at 1 bar, and the Langevin temperature equilibration using a collision frequency of 1 ps - 1 . After the equilibration, the number of Cl − and Na + ions needed to reach 0.1 M salinity was calculated using the average volume of the system in the last equilibration step. The whole process was repeated, from the tLEaP step, to correct the number of the added ions. Adaptive steered molecular dynamics The CTZ binding trajectories were calculated with adaptive steered molecular dynamics (ASMD). The ASMD method applies constant external force on two atoms in the simulated systems. By pulling two atoms together, we simulated ligand binding. During ASMD, several parallel simulations were started from the same state. The simulation ran in stages where the distance between the selected atoms changed by 2 Å per stage. At the end of each stage, the parallel simulations were collected, analyzed, and the Jarzynski average 71 , 72 was calculated throughout the stage. The trajectory with its work value closest to the Jarzynski average was selected, and the state at the end of this trajectory was used as the starting point for the next stage. We used the default values for setting up ASMD from the respective Amber tutorial and the ASMD publication 73 . The simulations were run with 25 parallel MDs, steered by 2 Å stages of distance increments, with a velocity of 10 Å.ns - 1 and a force of 7.2 N. The rest of the MD settings were set as in the last equilibration step. To simulate CTZ binding, we selected the N1-nitrogen of CTZ and the Cγ atom of D120 residue as steering atoms (D120 in RLuc8 corresponds to D118 in the ancestral variants). The initial distance between the steering atoms in the equilibrated systems was measured using PyMOL (Schrödinger LLC, USA). The selected steering atoms were pulled together to the distance of 4.4 Å, as observed in the crystal structure PDB ID 7QXR (azaCTZ-bound AncFT luciferase). System preparation and equilibration for adaptive sampling The adaptive sampling method was used to simulate product release from the studied enzymes with a thorough sampling of the unbinding process. The systems for adaptive sampling were built using the crystal structures of RLuc8, AncFT-L14, and Anc HLD-RLuc . Three systems were generated with CEI placed in the active site, as seen in PDB ID 7QXQ (AncFT + CEI). An additional system was prepared for AncHLD-RLuc with a flipped CEI, as observed in the adaptive steered MD simulation of CTZ binding. The following steps were performed with the High Throughput Molecular Dynamics (HTMD) 74 scripts. The protein structures were protonated with PROPKA 2.0 75 at pH 7.5. Each system was solvated in a cubic water box of TIP3P 64 water molecules with the edges at least 10 Å away from the protein by the solvate module of HTMD. Cl − and Na + ions were added to neutralize the charge of the protein and get a final salt concentration of 0.1 M. The topology of the system was built using amber.build module of HTMD, with the ff14SB 63 Amber force field and the previously compiled PREPI and FRCMOD parameter files for the ligands. The systems were equilibrated using the equilibration_v2 module of HTMD3 74 . The system was first minimized using a conjugate-gradient method for 500 steps. Then the system was heated to 298 K and minimized as follows: (i) 500 steps (2 ps) of NVT thermalization with the Berendsen barostat with 1 kcal.mol - 1 .Å - 2 constraints on all heavy atoms of the protein, (ii) 1,250,000 steps (5 ns) of NPT equilibration with Langevin thermostat and same constraints, and (iii) 1,250,000 steps (5 ns) of NPT equilibration with the Langevin thermostat without any constraints. During the equilibration simulations, holonomic constraints were applied on all hydrogen-heavy atom bond terms, and the mass of the hydrogen atoms was scaled with factor 4, enabling time steps of 4 fs 76 – 79 . The simulations employed periodic boundary conditions, using the particle mesh Ewald method to treat interactions beyond the 9 Å cut-off. The 1-4 electrostatic interactions were scaled with a factor of 0.8333, and the smoothing and switching of van der Waals interaction was performed for a cut-off of 7.5 Å 78 . Adaptive sampling HTMD3 74 was used to perform adaptive sampling of the conformations of the systems. Production MD runs of 50 ns were started using the equilibrated systems, employing the same settings as in the last equilibration step. The trajectories were saved every 0.1 ns. Adaptive sampling was performed using the distance of the N1-nitrogen of CEI and the Cγ atom of catalytic residue D120 as the adaptive metric (residue D120 in RLuc corresponds to D118 in the ancestral variants), and time-lagged independent component analysis (tICA) 80 projection in 1 dimension. 20 epochs of 10 parallel MDs were performed, corresponding to a cumulative time of 10 µs per system. Markov state model construction The simulations were made into a simulation list using HTMD3 74 ; the water and ions were filtered out, and unsuccessful simulations shorter than 50 ns were omitted. Such filtered trajectories were combined, which resulted in 10 µs of cumulative simulation time for each of the five systems. CEI release was studied based on the distance of the N1-nitrogen of CEI and the Cγ atom of the D120 residue (D120 in RLuc corresponds to D118 in the ancestral variants). The data were clustered using the MiniBatchKmeans algorithm to 1,000 clusters. Markov state models (MSM) with three macrostates were constructed at a lag time of 17 ns for each system with crystal-like CEI and at 15 ns for the flipped CEI unbinding. The three macrostates were labeled based on the CEI location as ‘bound’, ‘tunnel’, and ‘unbound’. The opening of the access tunnel (active site opening caused by the conformational change of the α4 helix) was studied based on the distance of the Cα atoms of I159/V185 residues in RLuc8, I157/L183 in AncFT-L14, and A158/L184 in Anc HLD-RLuc . The data were clustered using the MiniBatchKmeans algorithm to 1,000 clusters. MSMs with two macrostates for each system were constructed at a lag time of 17 ns for RLuc8, 20 ns for AncFT-L14 and Anc HLD-RLuc with crystal-like CEI, and 15 ns for Anc HLD- RLuc with flipped CEI. The Chapman-Kolmogorov test was performed to ensure the models are Markovian and describe the data well ( Figure S14 and S15 ). The states were visualized in VMD 1.9.3 81 , and the statistics of the ligand-active site distance were calculated (mean distance, standard error S.E., minimum, and maximum). The trajectory was saved for each model and visualized in PyMOL (Schrödinger LLC, USA). Kinetics analysis of molecular dynamics simulations Kinetic values (MFPT on/off, k on , k off , k off / k on , Δ G 0 , and K d ) were calculated by the kinetics module of HTMD3 74 between the source (‘unbound’ state) and sink (‘bound’ state). Also, the equilibrium probability of each macrostate (population of a specific conformation observed during the simulation) was calculated and visualized. Finally, bootstrapping of the kinetics calculation was performed, using randomly selected 80 % of the data, run 100 times. The kinetic values were then averaged, and the standard errors were determined. Binding energy calculation using MM/GBSA The molecular mechanics/generalized Born surface area (MM/GBSA) 82 , 83 method was applied to calculate CEI binding free energy (Δ G bind ) in the enzymes and the respective residue-by-residue interactions. Ante-MMPBSA.py module of AmberTools 16 59 was used to remove the solvent and ions from the original topology files of each system, define the Born radii as mbondi3, and generate the corresponding topology files for the complex, receptor, and ligand. The Δ G bind of the ligand in each structure was calculated with the MMPBSA.py program 82 . The generalized Born method was used ( &gb namelist) with the implicit generalized Born solvent model ( igb=8 ) and salt concentration 0.1 M ( saltcon=0.1 ). The solvent-accessible surface area was computed with an LCPO algorithm 84 . Decomposition of the interactions ( &decomp namelist) was generated per residue ( idecomp=2 ), with discrimination of all types of energy contributions ( dec_verbose=0 ). The MM/GBSA energies were calculated on 1,000 snapshots of each enzyme’s ‘bound’ and ‘tunnel’ macrostates. Data availability Atomic coordinates and structural factors have been saved in the Protein Data bank ( www.wwpdb.org ) 54 under the PDB ID accession code 7OME . Molecular dynamic simulations were deposited to the Zenodo repository with the assigned DOI of 10.5281/zenodo.1551624 . Competing interests Kenneth A. Johnson is the president of KinTek Corporation which develops and distributes the KinTek Explorer software used in this study. Jiri Damborsky and Zbynek Prokop are co-founders of the biotechnology spin-off Enantis. The mutants presented in this manuscript are covered in the submitted patent application PCT/EP2022/072839. Acknowledgements This research was supported by the European Union’s Horizon 2020 Research and Innovation Programme under grant agreement Nos. 857560 and 101136607, and by the project National Institute for Neurology Research (No. LX22NPO5107 MEYS), financed by the European Union – Next Generation EU. The authors thank the RECETOX Research Infrastructure (No. LM2023069) financed by the Ministry of Education, Youth, and Sports. Computational resources were provided by the e-INFRA CZ and ELIXIR-CZ (Nos. 90254 and LM2023055), supported by the Ministry of Education, Youth and Sports of the Czech Republic. The work on this paper was supported by the Czech Science Foundation (GA22-09853S and GX25-17329X). Martin Toul was supported by the Brno Ph.D. Talent scholarship (JCMM & Brno City Municipality) and the Fulbright Postgraduate Scholarship (Fulbright Commission). The crystallographic data were collected at the PXIII beamline at the Swiss Light Source (SLS) in Villigen (Switzerland). We are grateful to the members of the SLS synchrotron for the use of their beamline and help during data collection. Funder Information Declared European Union , 857560 , 101136607 , 90254 The Ministry of Education, Youth and Sports , LX22NPO5107 , LM2023069 , LM2023055 Czech Science Foundation, https://ror.org/01pv73b02 , GA22-09853S , GX25-17329X Footnotes ↵ # Joined first authors References 1. ↵ Schramm , S. & Weiß , D . Bioluminescence – The vibrant glow of nature and its chemical mechanisms . ChemBioChem 25 , e202400106 ( 2024 ). OpenUrl PubMed 2. ↵ Letendre , F. , Twardowski , M. , Blackburn , A. , Poulin , C. & Latz , M. I . A review of mechanically stimulated bioluminescence of marine plankton and its applications . Front. Mar. Sci . 10 , 1299602 ( 2024 ). OpenUrl 3. ↵ Collins , S. B. & Bracken-Grissom , H. D . The language of light: a review of bioluminescence in deep-sea decapod shrimps . Biol. Rev . 99 , 1806 – 1830 ( 2024 ). OpenUrl 4. ↵ Syed , A. J. & Anderson , J. C . 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