Stability Studies of New Caged bis-Deoxy-Coelenterazine Derivatives and Their Potential Use as Cellular pH Probes

Germano Giulian1*, Assunta Merolla2, Marco Paolino1, Annalisa Reale1, Mario Saletti1, Lluís Blancafort3, Andrea Cappelli1, Fabio Benfenati2,4, Fabrizia Cesca3,5

1 Department of Biotechnology, Chemistry and Pharmacy, University of Siena, via A. Moro 2, 53100 Siena, Italy

2 Center for Synaptic Neuroscience and Technology, Istituto Italiano di Tecnologia, 16132 Genova, Italy
3 Institut de Química Computacional i Catàlisi (IQCC), Departament de Química, Facultat de Ciències, Universitat de Girona, C/M. A. Capmany 69, 17003 Girona, Spain

4 IRCCS Ospedale Policlinico San Martino, Genova, Italy

5 Department of Life Sciences, University of Trieste, 34127 Trieste, Italy

This article has been accepted for publication and undergone full peer review but has not been through the copyediting, typesetting, pagination and proofreading process, which may lead to differences between this version and the Version of Record. Please cite this article as doi: 10.1111/PHP.13347
Corresponding author e-mail: [email protected] (Germano Giuliani)


The synthesis of new bis-deoxy-coelenterazine (1) derivatives bearing ester protective groups (acetate, propionate and butyrate esters) was accomplished. Moreover, their hydrolytic stability at room temperature was evaluated in dimethylsulfoxide (DMSO) as solvent, using the nuclear magnetic resonance (NMR) spectra of the key products at different time intervals. The results showed an increasing hydrolysis rate according to longest aliphatic chain, with a half-life of 24 days of the more stable acetate derivative (4a). Furthermore, the analysis of the experimental data revealed the greater stability of the enol tautomer in this aprotic polar solvent. This result was confirmed by theoretical calculations using the Density Functional Theory (DFT) approach, which gave us the opportunity to propose a detailed decomposition mechanism. Additionally, the derivatives obtained were tested by bioluminescence luciferase assays to evaluate their potential use as extra-cellular pH-sensitive reporter substrates of luciferase. The biological data support the idea that further structural modifications of these molecules may open promising perspectives in this field of research.

KEYWORDS Caged-Coelenterazine, bis-deoxy-Coelenterazine, Bioluminescence Luciferase assay, DMSO Stability, DFT, 2Y-System


Bioluminescence Imaging (BLI) has become a useful tool for real-time monitoring of biological events in living cells. BLI has been adopted in many biological applications including gene therapy, stem cell tracking, tumor growth assessment, drug development and protein-protein interactions.1-5 Most bioluminescence systems require a luciferase (enzyme), a luciferin (substrate) and molecular oxygen to produce visible light. The two main reporter assays used are Firefly luciferase (Fluc) and Renilla luciferase (Rluc). Rluc can catalyze the oxidation of its Coelenterazine (CTZ, 2) substrate to produce blue light with emission spectra in the range of 460-490 nm.6 As consequence of the high sensitivity, versatility and instrumental simplicity of this biotechnology,7-10 the interest of many researchers has grown in the past decades, but some drawbacks still limit its applicability, mainly related to its poor optical intensity and limited color variety. To overcome such restraints, the development of stable variants of luciferases obtained via random mutagenesis is a promising approach.11 However, the optical performances of these artificial luciferases greatly depend on the chemical structure of the substrate. For this reason, many luciferin analogues were synthetized to obtain improved photochemical properties such as a wider emissions range.12,13 These strategies have led to evident improvements in BLI technologies, however the molecules obtained still showed some inconveniences of the native CTZ related to their poor stability in aqueous solution (half-life of 15 minutes at 37° C) and the fast catalytic oxidation that generates rapidly decaying bioluminescence signals, which are not favorable for prolonged real-time imaging in vitro and in vivo.14,15 The development of CTZ cages has proven to be a brilliant strategy for solving both limitations. CTZ can be stabilized by introducing protective groups in different positions of the molecule, such as the 3- carbonyl of the imidazo-pyrazinone scaffold (3) or the 6-phenolic portion, thus temporarily limiting its interaction with Rluc.16,17 When activated by specific enzymes (or small molecule), caged luciferins can be converted into free substrates to generate the bioluminescent signal, thus leading to a longer half-life and less autoluminescence.18
In our previous work,19 we have synthesized a series of sulfur derivatives of bis-deoxy-CTZ with a higher luminescence signal and remarkable red-shifted emission compared to the original molecule. This bathochromic effect occurred in both chemiluminescence and bioluminescence, suggesting that it likely results from an intrinsic change in the electronic properties of the molecule, independently of how they are produced. Following these intriguing results, these sulfur derivatives were subjected to a

further investigation selecting derivative 3 as candidate for the synthesis of the corresponding caged compounds. As it can be noticed from the general structure of the caged bis-deoxy-CTZ 4 (see Chart 1), the only possible position to insert the protective groups is the carbonyl in C3 of the imidazo- pyrazinone scaffold. Therefore, we scheduled to synthetize a series of derivatives of 3, bearing ester protective groups (acetate, propionate and butyrate esters) in this position, aimed to evaluate their hydrolytic stability over time (at room temperature) in dimethylsulfoxyde (DMSO) as solvent. In fact, it is known that this aprotic polar solvent can enhance the photochemical reaction of the coelenterazine, but kinetic studies on their caged counterpart in this solvent is missing. We attempted to fill this gap by planning a protocol based on the 1H-NMR spectra of our derivatives dissolved in deuterated DMSO-d6. This procedure can give us additional information on the hydrolysis of the protective groups and the kinetic profile based on their different aliphatic chains. At the same time, we focused our attention on the cleavage of the protecting groups in a pH range from 7.5 to 6.0. Our intent was to determine their behavior in the hydrolysis of the ester group within this interval of pH. This investigation is related to the study of pathologies that cause small extracellular pH variations.
Brain cells are highly sensitive to a vast number of chemical factors; one of the most important intrinsic neuromodulator is the H+ ion that has a strong influence on neuronal function. Within the nervous system, the control of pH has particular relevance for synaptic transmission and the modulation of network excitability.20 pH affects neuronal activity, and neuronal activity itself can in turn generate sizeable shifts in intracellular pH.21 Tools to accurately measure H+ concentration are therefore important for understanding the role that pH plays during the evolution of both physiological and pathological network states. The past decade has witnessed an exponential increase in the use of optogenetics, which is based on the combination of optics and genetics, for treating neurological disorders. After the delivery of light sensitive opsin genes such as channelrhodopsin-2 (ChR2), halorhodopsin (NpHR) and others into the brain, excitation or inhibition of specific neurons in precise brain areas can be controlled by illumination at specific wavelengths with very high temporal and spatial resolution.22 The implementation of such techniques however requires specialized equipment, and the surgical implantation is very invasive. In order to eliminate the need for an external light source, a possibility is to make the animal produce endogenous light thanks to the action of light-producing proteins, i.e. luciferases. Renilla luciferase is the most used luciferase

because it is not ATP dependent and in general requires only molecular oxygen in addition to its substrate, coelenterazine (CTZ, 2), for luminescence. The limiting factors of RLuc are its rapid inactivation and low brightness. To improve RLuc characteristics, site directed mutagenesis of the RLuc gene and chemical synthesis of analogues of the coelenterazine were performed.11,19 Thanks to the optimization of the various luciferases and substrates, it is now possible to use a bioluminescence light source to activate opsins, as in the work of Tung and colleagues, where the activation of the inhibitory opsin eNpHR3.0 was driven by luciferase-emitted light, exploiting the bioluminescence resonance energy transfer (BRET) mechanism.23 In this work we engineered a number of pH- dependent coelenterazine variants, so that, if a caged-coelenterazine is selectively deprotected under pathological conditions, a bioluminescent probe of the disease can be obtained. In this way the protected molecule acts as a reservoir of the luminescent substrate until the pH decreases. From a chemical point of view, the hydrolysis yield of the ester group proposed in this work increases in a more acidic reaction environment, which promotes our derivatives as good candidates for the test.
<Chart 1>
Results and Discussion

The synthesis of the sulfur derivative 3 was achieved following our known procedure,19 reported in Scheme S1 and S2 of the Supporting Information. Particular attention was paid to handling the free compound due to its extreme sensitivity to air and light. We report here the final step of the synthesis of the caged bis-deoxy-coelenterazine 4a-c (Scheme 1), where the reaction of the suitable anhydride in presence of dry pyridine gave the protected substrates. As it can be seen in Scheme 1, the structure of 3 corresponds to the enol tautomer that is assumed to be the suitable isomer for this esterification. A theoretical investigation on the keto-enol tautomerism of 3 is reported later in the manuscript (see Figure 3).
<Scheme 1>
Kinetic studies

To study the decomposition kinetics of the acetate 4a, propionate 4b and butyrate 4c esters, we had to consider the two main pathways of decomposition: the hydrolysis of the protecting groups and the following chemiluminescent reaction of the free substrate. The product resulting from the luminescent reaction is the coelenteramide 5,24 while compound 3 was assumed to be the result of the deprotecting reaction. Therefore, these reactions can be monitored once the 1H-NMR spectra of the purified 3 and 5 are obtained.
<Figure 1>
In Fig. 1, the spectra of the pure 3, 4a and 5 dissolved in DMSO-d6 are compared to the spectra of the DMSO-d6 solution of 4a stored for 70 days at room temperature. This comparison allowed us to identify the indicative peaks of the chemical degradation reactions. As the first step in this direction, the two peaks (s, 8.66 ppm; s, 8.82 ppm) in the spectra of the acetyl derivative 4a decomposition (above in Fig. 1) were chosen as characteristic to describe the kinetic we want to study. These signals are assigned to the same aromatic proton bound respectively to C-5 of the pyrazine ring of 4a and C-6 of celenteramide 5 (see Chart 1). Proceeding over time, their mutable intensity can be correlated to the interconversion of the reagent 4a into the final product 5, therefore such signals are taken as reference for monitoring the long-term chemical stability of 4a. In the same Fig. 1, the spectra of 3 shows a displacement at 8.37 ppm of the C-5 proton peak with respect to the same signal of the protected substrate, which allows to evaluate the possible formation of free luciferin during the chemical degradations. As reported in literature,25 this behavior can be easily explained by the magnetic anisotropy effect of the adjacent ester carbonyl of 4a. At the same time, the aromatic signals comprised between 7.16-7.72 ppm of compound 4a and 3 are very similar. The shape and the chemical shifts of the peaks suggest which could be the conformation (enol tautomer) adopted by 3 in DMSO.
<Figure 2>

In Fig. 2a, the spectra of 4a obtained at different time intervals are shown. To study the kinetic rate of the hydrolysis reaction of 4a, we expected that the decreasing intensities of the selected peak over time are proportional to the remaining amount of the caged compound. Starting from the known initial concentration of 4a (11 mM; time = 0 h), the relative concentrations are reported in the graph in Fig. 2b. The curve shows how the hydrolysis of 4a proceed over time, where the concentration shows an exponential decrease in DMSO, taking 25° C as reference temperature for the following kinetic parameters determination. This tendency is typically associated with an apparent first-order kinetic reaction but the natural logarithm of concentration versus time should be calculated to confirm the experimental observation. These values are show in Fig. 2c, where the black dotted straight line is the calculated R-square linear regression of the data obtained, which seems to be in good agreement (R2 = 0.9917) with the behavior of a system that obeys a first-order rate law. Finally, the kinetic rate (k =
1.2 x 10-3 h-1) of the hydrolysis of 4a at 25° C, was determined as angular coefficient of the straight line obtained. Furthermore, from the analysis of the spectra reported in Fig. 2a we noticed that no trace of 3 was detected during the process, which is due to the different kinetics of the subsequent reactions. In fact, the hydrolysis of acetyl ester is slower than the chemiluminescence reaction, therefore the molecules of 3 obtained react promptly to give celenteramide 5 as the final product. To better visualize this interconversion, the relative percentage of compounds 4a and 5 is reported in Figure S1 of the Supporting Information, where the red line is relative to the decreasing value of 4a while the blue one is the increasing value of 5. The good correlation between these opposite trends suggests how faster the chemiluminescent reaction is compared to hydrolysis of the acetyl protecting group of 4a.
The same procedure was followed for the other compounds 4b and 4c. The comparison of 1H-NMR spectra and the concentrations over time for each derivative is reported in Figures S2-S5 of the Supporting Information. To proceed in our comparative study, we selected as reference the same aromatic peaks of the key compounds and similar starting concentrations of the caged derivatives dissolved in DMSO-d6. As reported in Figure S2, the hydrolysis of the propionate derivative 4b shows a different trend from what was obtained for the acetate derivative. The progress of the hydrolysis over time does not follow the exponential decrease previously observed and shows a linear regression that can be associated with an apparent zero-order kinetic law. This means that the

hydrolysis of the propionate protecting group occurs independently of the reactant concentration. This unexpected result allows us to determine the kinetic rate of the chemical reaction through the slope of the straight line obtained, which gives a value of 12.4 x 10-3 mM h-1 at 25° C. The reasons for this kinetic behavior are difficult to identify, however we can exclude that the hydrolysis of 4b has reached some kind of equilibrium since the product 3 rapidly reacts in the chemiluminescence reaction.
In the same way, the hydrolytic kinetic parameters of the butyrate derivative 4c were achieved and the results are showed in Figure S4 of the Supporting Information. As it can be seen, the hydrolysis of 4c follows again an exponential decrease as observed with compound 4a, but the reaction is faster with a kinetic rate of 1.7 x 10-4 min-1 (10.2 x 10-3 h-1) at 25° C.
<Table 1>
The hydrolysis rates, half-life and kinetic order of compounds 4a-c hydrolysis are reported in Table 1. As it can be noted, while 4a and 4b show a similar half-life, the kinetic rate of compound 4c is about 10 times faster than the acetyl ester 4a, with a half-life of 3.8 days. Usually the hydrolytic stability of organic esters is related to many factors such as the mechanism involved in the reaction, the size and the resonance effect of the substituent groups, the solvent and the temperature at which the reaction occurs.26-28 In the present work, all the experiments were performed at the same temperature, in DMSO as solvent and the sulfur derivative 3 was the common substituent group, so our kinetic results cannot be ascribed to these variables. Therefore, only the different aliphatic chains should play a role on the kinetic variability observed. Taking into account the acid catalyzed hydrolysis mechanism reported in Chart 2, the partial positive charge of the intermediate I can be stabilized by the electron- donating inductive property of the adjacent methyl group of 4a. In the same way, a longer aliphatic chain increases this stabilizing effect, which brings to the faster hydrolysis of compounds 4b,c. In fact, the alkyl chain effect on the intermediate I can be considered the rate determination step of the hydrolysis mechanism, which promotes the formation of the carbocation as product of the reaction. Otherwise, taking the reaction of intermediate II as the key step of the hydrolytic kinetic (see Chart 2), the alkyl chain effect should act in the opposite manner stabilizing the tetrahedric intermediate II as reactant, so limiting the formation of the corresponding carboxylic acid and alcohol.

Instead, the effect of DMSO in the hydrolytic mechanism should be considered, as it is itself responsible for drastic changes compared to what has been observed in aqueous solutions.27 Yuan and coworkers29 report an opposite hydrolytic trend of derivative 4a dissolved in water, which is necessarily related to solvent change. Literature suggests that organic carboxylic acids show an increasing value of pKa (as example, acetic acid from 4.76 to 12.3) when dissolved in DMSO,30 which means the protonated form of the acids is more stable in this solvent. In our conditions, these altered properties can stabilize the carboxylic acids produced from the tetrahedric intermediate II (see Chart 2) and at the same time, decreasing the acidity of the solution (weaker catalytic condition), resulting in the slower kinetic rate observed for 4a. We may speculate that both factors should be considered to correctly interpret the different kinetic trends shown by derivatives 4a-c, however further work is necessary to verify this possibility.
<Chart 2>
Theoretical Calculations
As previously observed, another interesting information can be achieved from the spectra reported in Fig. 1: the good overlap of the aromatic signals (chemical displacement between 7.25-7.77 ppm) of compound 3 and those of the acetyl derivative 4a. This result supports the idea that the enol form of 3 in DMSO is more stable than the keto isomer (see Fig. 3). To verify this experimental observation, we performed computational calculations on both tautomers using the density functional theory (DFT) methodology, within CAM-B3LYP functional31 and 6-311G** basis set of the GAUSSIAN package (version 16).32 Before calculating the relative energies, the conformational analysis of the dihedral angle N7C8S13C14 was carried out. The energies of the conformers were calculated every 20° of rotation of the angle and the most stable geometries are reported in Fig. 3. Furthermore, explicit DMSO molecules were positioned near the 7H and hydroxyl proton to evaluate the possible formation of hydrogen bonds and their contribution to ground state energies. Following this approach, we fully optimized the geometries of the tautomers in DMSO as implicit solvent (ɛ = 46.826) by calculating the Gibbs free energy (at 298° K) for each conformation.
<Figure 3>

The results obtained are in good agreement with what was previously observed in the NMR spectra. In fact, comparing the energies of the two tautomers, a ∆G of -6.1 kcal/mol was obtained (see Fig. 3) which confirms that the enol conformation of 3 is more stable than the keto form. Analyzing the Maxwell-Boltzman distribution33 between conformers in thermal equilibrium condition (25° C), we obtain the 99.9% of possibility to find the molecule in the enol conformation.
Moreover, another signal appears in the NMR spectra reported in Fig. 1 as a broad peak at 12 ppm. The shape of this peak and its chemical shift can be assigned to carboxylic proton of the acetic acid obtained as byproduct of the decomposition reaction. Taken together, the results prompted us to propose a detailed mechanism of the chemiluminescent reaction in DMSO, as showed in Scheme 2.
<Scheme 2>
Biological assay
We subsequently focused our attention on the cleavage of the protecting groups, to determine the behavior of the three compounds at physiologically relevant pH values. To this aim, HEK293T cells were transfected with a construct carrying the Renilla luciferase sequence. After 24 h, cells were tripsinized and plated in a white 96-well plate. Bioluminescence intensity at 480 nm was measured using the TECAN 500F machine (Microplate Reader INFINITE F500 Tecan, Austria), up to 25 min after addition of the various molecules at 5 M and by setting the extracellular pH to 6.0, 7.4 and 8.0 (Fig. 4). Similar results were obtained by using the same molecules at 10 M (not shown). Bioluminescence intensity values measured 1 min after the addition of the three compounds were similar (Fig. 4A-C, graphs on the left), indicating that all reactions took place with a comparable efficiency. Bioluminescence data were therefore expressed as fold-change normalized to the values detected at t = 1 min, to highlight the sensitivity of the process (Fig. 4A-C, graphs on the right). Unlike the typical flash-like kinetics of coelenterazine,[19] with an immediate burst (few seconds) to peak emission rate followed by steady decay, the derivatives 4a-c showed slower kinetics: 4a induced bioluminescence peaked around 5 min, while for the other two molecules the peak was observed even later. Moreover, all the molecules generated higher bioluminescence at higher pH. Of note, bioluminescence at pH 8.0 did not further increase compared to that emitted at pH 7.4, probably because of saturation of the reaction. Thus, all the derivatives show maximal sensitivity in the range

between acidic (6.0) and physiological (7.4) pH conditions. 4a also gave the highest absolute bioluminescence values, which at the peak were about 10 times higher than the other molecules (Figure S6). In terms of sensitivity, the molecules 4b and 4c showed bigger fold changes than 4a (compare Figs. 4 B and C with A, graphs on the right). Overall, therefore, the 4a molecule showed the fastest and most sensitive pH-dependent bioluminescence emission, while 4b and 4c were characterized by delayed and more sustained emission.
<Figure 4>


In summary, by introducing ester protecting groups of different aliphatic chains into the carbonyl group of the bis-deoxy-CTZ 3, we synthesized new pro-substrates for Renilla luciferase. Through a protocol based on 1H-NMR spectra, the hydrolytic stability profiles (in DMSO-d6) of these molecules were evaluated, determining the kinetic rate, the half-life and kinetic order for each derivative. Furthermore, using the DFT modelling approach, an investigation on the relative energies of the tautomers of the free substrate 3 was performed and the computational results confirm the experimental observations previously obtained.
From the biological point of view the newly synthetized molecules showed appreciable pH sensitivity in the 6.0 to 7.4 range, with different kinetics. They therefore may represent useful tools to be exploited for non-invasive, in vivo chemogenetic applications, to be tuned to the specific pathological features of different brain diseases.

Supporting Information
Additional supporting information may be found online in the Supporting Information section at the end of the article:
Scheme S1:1(a) NBS, CH2Cl2, 0 °C, 3 h; (b) BINAP, Pd(OAc)2 (5 mol %), Cs2CO3, phenylboronic acid, Toluene-ethanol 1:1, reflux, 18 h; (c)Br2, Py, CHCl3, -10 °C to r.t., 2h; (d) benzenethiol, NaH, CH3CN dry, reflux, 6h.

Scheme S2:1 (a) 11, HCl conc., H2O, EtOH, reflux 4 h.

Scheme S3: (a) phenylacetyl chloride, pyridine, CHCl3.
Figure S1. Qualitative comparison of kinetic curves related to the hydrolysis of 4a. The relative percentage of compounds 4a and final product 5 are show as decreasing value of 4a (orange line) and increasing value of 5 (blue line). According to the faster chemiluminescent reaction, compound 3 was not detected during the kinetic study.
Figure S2: 1H-NMR (400 MHz, DMSO-d6) spectra of compounds 3,5,4b and the spectra of 4b
(shown above in violet) obtained after 18 days at room temperature, are showed.

Figure S3: The chemical decomposition of 4b acquired by time-dependent 1H-NMR (DMSO-d6) spectra analysis at 25° C. In the graph on the right, the relative concentrations (expressed in mM) of compounds 4b obtained at different interval of time (h) are shown.
Figure S4: 1H-NMR (400 MHz, DMSO-d6) spectra of compounds 3,5,4c and the spectra of 4c
(shown above in violet) obtained after 4 days at room temperature, are showed.

Figure S5: The chemical decomposition of 4c acquired by time-dependent 1H-NMR (DMSO-d6) spectra analysis at 25° C. In the above graph, the relative concentrations (expressed in mM) of compounds 4c obtained at different interval of time (min) are shown. In the bottom graph, the natural logarithm of the concentrations versus time are reported.
Figure S6: Bioluminescence intensity from HEK293T cells transfected with RLuc8(S257G) at extracellular pH 7.4 at time t = 0 min and the bioluminescence intensity followed over time. HEK293T cells transfected with RLuc8(S257G) and exposed to extracellular pH 7.4 were incubated with 5 μM of 4a, 4b or 4c. The graph shows the time-course of bioluminescence values over 25 min. Data were analyzed by repeated measures ANOVA followed by the Tukey’s multiple comparison test. ***p<0.001, **p<0.01 4a versus 4b and 4c, *p<0.05 4a versus 4c. All data are expressed as means ± SEM of n=6 independent cell preparations.

Notes and References.
1. K. Pinel, J. Lacoste, G. Plane, M. Ventura and F. Couillaud, Gene Ther., 2014, 21, 434–439.

2. J. R. Pribaz, N. M. Bernthal, F. Billi, J. S. Cho, R. I. Ramos, Y. Guo, A. L. Cheung, K. P. Francis and L. S. Miller, J. Orthop. Res., 2012, 30, 335–340.
3. J. E. Kim, S. Kalimuthu and B. C. Ahn, Nucl. Med. Mol. Imaging, 2015, 49, 3–10.

4. Y. Inoue, F. Sheng, S. Kiryu, M. Watanabe, H. Ratanakanit, K. Izawa, A. Tojo and K. Ohtomo,
Mol. Imaging, 2011, 10, 377–385.

5. A. Dragulescu-Andrasi, C. T. Chan, A. De, T. F. Massoud and S. S. Gambhir, Proc. Natl. Acad. Sci. U. S. A., 2011, 108, 12060–12065.
6. T. Y. Jiang, L. P. Du and M. Y. Li, Photochem. Photobiol. Sci., 2016, 15, 466–480.

7. X. D. Tian, Z. Y. Li, C. W. Lau and J. Z. Lu, Anal. Chem., 2015, 87, 11325–11331.

8. B. R. Branchini, T. L. Southworth, D. M. Fontaine, D. Kohrt, F. S. Welcome, C. M. Florentine,
E. R. Henricks, D. B. DeBartolo, E. Michelini, L. Cevenini, A. Roda, M. J. Grossel, Anal. Biochem. 2017, 534, 36-39.
9. R. Nishihara, E. Hoshino, Y. Kakudate, S. Kishigami, N. Iwasawa, S. Sasaki, T. Nakajima, M. Sato, S. Nishiyama, D. Citterio, K. Suzuki, S. B. Kim, Bioconjugate Chem. 2018, 29, 1922-1931.
10. D. M. Mofford, S. T. Adams, G. S. K. K. Reddy, G. R. Reddy, S. C. Miller, J. Am. Chem. Soc.,
2015, 137, 8684–8687.

11. a) O. Shimomura, Y. Kishi, S. Inouye, Biochem. J. 1993, 296, 549-551. b) A.M. Loening, A.M. Wu, S.S. Gambhir, Nat. Methods, 2007, 4, 641–643.

12. R. Nishihara, H. Suzuki, E. Hoshino, S. Suganuma, M. Sato, T. Saitoh, S. Nishiyama, N. Iwasawa, D. Citterio, K. Suzuki, Chem. Commun. 2015, 51, 391−394.
13. T. Kuchimaru, S. Iwano, M. Kiyama, S. Mitsumata, T. Kadonosono, H. Niwa, S. Maki, S. Kizaka-Kondoh, Nat. Commun. 2016, 7, 11856.
14. L. J. Kricka, Anal. Chem., 1993, 65, 460–462.

15. M. Mirasoli, E. Michelini, Anal. Bioanal. Chem., 2014, 406, 5529–5530.

16. W. W. Lorenz, R. O. McCann, M. Longiaru, M. J. Cormier, Proc. Natl. Acad. Sci. U. S. A., 1991,
88, 4438–4442.

17. E. P. Coutant, Y. L. Janin, Chem. – Eur. J., 2015, 21, 17158–17171.

18. M. Otto-Duessel, V. Khankaldyyan, I. Gonzalez-Gomez, M. C. Jensen, W. E. Laug, M. Rosol,
Mol. Imaging, 2006, 5, 57–64.

19. G. Giuliani, P. Molinari, G. Ferretti, A. Cappelli, M. Anzini, S. Vomero, T. Costa, Tetrahedron Lett., 2012, 53, 5114–5118.
20. a) P. Drapeau, D.A. Nachshen, J. Gen. Physiol. 1988, 91, 305-315. b) J.S. Tabb, P.E. Kish, R. Van Dyke, T. Ueda, J. Biol. Chem. 1992, 267, 15412-15418. c) C.G. Dulla, P. Dobelis, T. Pearson, B.G. Frenguelli, K.J. Staley, S.A. Masino, Neuron, 2005, 48, 1011-1023.
21. a) Z. Ahmed, J.A. Connor, J. Gen. Physiol. 1980, 75, 403-426. b) Z.Q. Xiong, P. Saggau, J.L. Stringer, J. Neurosci. 2000, 20, 1290-1296.
22. K. Deisseroth, Nat. Neuroscience 2015, 18, 1213-1225.

23. a) J.K. Tung, C.A. Gutekunst, R.E. Gross, Scientific Report, 2015, 5, 14366. b) J.K. Tung, F.H. Shiu, K. Ding, R.E. Gross, Neurobiol. Dis. 2018, 109, 1-10.
24. J. Lee, Photochem. Photobiol. 2017, 93, 389-404.

25. R. J. Abraham, M. Mobli, R. J. Smith, Magn. Reson. Chem. 2003, 41, 26–36.

26. R. Gomez-Bombardelli, E. Calle, J. Casado, J. Org. Chem. 2013, 78, 6880−6889.

27. R. K. Wolford. J. Phys. Chem., 1963, 67, 632.

28. P. G. Takis, K. D. Papavasileiou, L. D. Peristeras, G. C. Boulougouris, V. S. Melissas, A. N. Troganis, Phys. Chem. Chem. Phys., 2017, 19, 13710–13722.
29. M. Yuan, X. Ma, T. Jiang, Y. Gao, Y. Cui, C. Zhang, X. Yang, Y. Huang, L. Du, I. Yampolsky, M. Li, Org. Biomol. Chem., 2017, 15, 10238–10244.
30. Y. Fu, L. Liu, R. Li, R. Liu, Q. Guo, J. Am. Chem. Soc., 2004, 126, 814-822.

31. a) A. D. Becke, J. Chem. Phys. 1993, 98, 5648–5652; b) C. Lee, W. Yang, R. G. Parr, Phys. Rev.
1988, 37, 785–789.

32. Gaussian 16, Revision C.01, M. J. Frisch, G. W. Trucks, H. B. Schlegel, G. E. Scuseria, M. A. Robb, J. R. Cheeseman, G. Scalmani, V. Barone, G. A. Petersson, H. Nakatsuji, X. Li, M. Caricato, A. V. Marenich, J. Bloino, B. G. Janesko, R. Gomperts, B. Mennucci, H. P. Hratchian,
J. V. Ortiz, A. F. Izmaylov, J. L. Sonnenberg, D. Williams-Young, F. Ding, F. Lipparini, F. Egidi, J. Goings, B. Peng, A. Petrone, T. Henderson, D. Ranasinghe, V. G. Zakrzewski, J. Gao,
N. Rega, G. Zheng, W. Liang, M. Hada, M. Ehara, K. Toyota, R. Fukuda, J. Hasegawa, M. Ishida, T. Nakajima, Y. Honda, O. Kitao, H. Nakai, T. Vreven, K. Throssell, J. A. Montgomery, Jr., J. E. Peralta, F. Ogliaro, M. J. Bearpark, J. J. Heyd, E. N. Brothers, K. N. Kudin, V. N. Staroverov, T. A. Keith, R. Kobayashi, J. Normand, K. Raghavachari, A. P. Rendell, J. C. Burant,
S. S. Iyengar, J. Tomasi, M. Cossi, J. M. Millam, M. Klene, C. Adamo, R. Cammi, J. W. Ochterski, R. L. Martin, K. Morokuma, O. Farkas, J. B. Foresman, and D. J. Fox, Gaussian, Inc., Wallingford CT, 2016.
33. S. Glen. “Maxwell-Boltzmann Distribution” From


Figure Captions

Chart 1. bis-Deoxy-coelenterazine (1), coelenterazine (2), the thio-derivative synthetized (3), generic caged bis-deoxy-coelenterazine structure reported in this work (4) and the corresponding thio- coelenteramide (5).
Chart 2. A generic mechanism of acid hydrolysis of esters; stability of intermediate I plays a relevant role on the kinetic rate of the reaction.
Scheme 1. Reagents: (a) (RCO)2O, anhydrous pyridine.
Scheme 2. Decomposition mechanism of 4a in DMSO at room temperature. The initial hydrolysis is followed by the chemiluminescent reaction of 3 via dioxetanone intermediate 7,24 to give the final product coelenteramide 5.
Figure 1. 1H-NMR (400 MHz, DMSO-d6) spectra of compounds 3, 4a, and 5 compared with the spectra of 4a (shown above in violet) obtained after stored the compound in solution for 70 days at room temperature, are showed. The peaks corresponding to water (δ = 3.3 ppm) and DMSO (δ = 2.5 ppm) are marked with an asterisk.
Figure 2. (a) The chemical decomposition of 4a acquired by time-dependent 1H-NMR (DMSO-d6) spectra analysis at room temperature. (b) In the graph, the relative concentrations (expressed in mM) of compounds 4a obtained at different intervals of time (h) are showed. (c) The natural logarithm of concentrations versus time is reported, the straight line (black dotted) is expected for a first-order reaction.
Figure 3. Calculated energy gap between the tautomers of 3. The hydrogen bond (value in Angstrom) with DMSO is highlighted in purple.
Figure 4. Bioluminescence intensity from HEK293T cells transfected with RLuc8(S257G) at various extracellular pH conditions. HEK293T cells transfected with RLuc8(S257G) and exposed to extracellular pH 6.0 (black lines), 7.4 (red lines) and 8.0 (green lines) were incubated with 5 μM of compound 4a (data in panel A), 4b (data in panel B) or 4c (data in panel C) at time t = 0 min and the bioluminescence intensity followed over time. Left: Bioluminescence intensity values measured 1 min after the addition of the 4a (A), 4b (B) and 4c (C) compounds. Data were analyzed by one-way Kruskal-Wallis test; n.s., p>0.05. Right: Time-course of the bioluminescence data expressed as fold- change with respect to the bioluminescence value detected at t = 1 min. Data were analyzed by repeated measures ANOVA followed by the Tukey’s multiple comparison test. Panel A: *p<0.05 for

both pH 7.4 and pH 8.0 versus pH 6.0 at 5-15 min, for pH 8.0 only at 20 min. Panel B: *p<0.05 for both pH 7.4 and pH 8.0 versus pH 6.0 at 20-25 min, for pH 8.0 only at 15 min. Panel C: *p<0.05 for pH 7.4 and pH 8.0 versus Coelenterazine  pH 6.0 at all indicated points. All data are expressed as means ± SEM of n=6 independent cell preparations.