Towards understanding the E. coli PNP binding mechanism and FRET absence between E. coli PNP and formycin A.

Biophysical Chemistry 230 (2017) 99–108

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Towards understanding the E. coli PNP binding mechanism and FRET absence between E. coli PNP and formycin A.

MARK

Małgorzata Prokopowicza,b,*, Bartosz Greńb, Joanna Cieślac, Borys Kierdaszukb a

Inter-Faculty Interdisciplinary Doctoral Studies in Natural Sciences and Mathematics, University of Warsaw, Stefana Banacha 2C, Warsaw 02-097, Poland Department of Biophysics, Institute of Experimental Physics, Faculty of Physics, University of Warsaw, Żwirki i Wigury 93, Warsaw 02-089, Poland c Department of Drug Technology and Biotechnology, Institute of Biotechnology Faculty of Chemistry, Warsaw University of Technology, Koszykowa 75, Warsaw 00-664, Poland b

H I G H L I G H T S

G RA P H I C A L AB S T R A C T

Incorporation of the dissociation • constant is necessary in FRET calculations.

aromatic amino acid at the position • 159 is the key residue in the nucleoside binding.

Tyr /Tyr inhibits or quenches FRET • from other Tyr to FA. probably enhances the binding • Tyr159 efficiency of FA. buffer at pH 8.3 seems to • Phosphate restore (partially) FRET in PNPF59Y−

*

FA complexes.

A R T I C L E I N F O

A B S T R A C T

Keywords: E. coli purine nucleoside phosphorylase Formycin A Fluorescence resonance energy transfer methodology Binding mechanism

The aim of this study is threefold: (1) augmentation of the knowledge of the E. coli PNP binding mechanism; (2) explanation of the previously observed ‘lack of FRET’ phenomenon and (3) an introduction of the correction (modified method) for FRET efficiency calculation in the PNP-FA complexes. We present fluorescence studies of the two E. coli PNP mutants (F159Y and F159A) with formycin A (FA), that indicate that the aromatic amino acid is indispensable in the nucleotide binding, additional hydroxyl group at position 159 probably enhances the strength of binding and that the amino acids pair 159–160 has a great impact on the spectroscopic properties of the enzyme. The experiments were carried out in hepes and phosphate buffers, at pH 7 and 8.3. Two methods, a conventional and a modified one, that utilizes the dissociation constant, for calculations of the energy transfer efficiency (E) and the acceptor-to-donor distance (r) between FA and the Tyr (energy donor) were employed. Total difference spectra were calculated for emission spectra (λex 280 nm, 295 nm, 305 nm and 313 nm) for all studied systems. Time-resolved techniques allowed to conclude the existence of a specific structure formed by amino acids at positions 159 and 160. The results showed an unexpected pattern change of FRET in the mutants, when compared to the wild type enzyme and a probable presence of a structure created between 159 and 160 residue, that might influence the binding efficiency. Additionally, we

abbreviations: Asp, aspartic acid; Arg, arginine; E, efficiency of FRET; f, fractional intensity; FA, formycin A; FRET, Fluorescence/Förster resonance energy transfer; H7, hepes buffer pH 7; H8, hepes buffer pH 8.3; Phe, phenylalanine; Pi, phosphate; PNs, purine nucleosides; PNP, purine nucleoside phosphorylase; PNPA, PNPF159A; PNPY, PNPF159Y; PNPWT, E. coli purine nucleoside phosphorylase; P7, phosphate buffer pH 7; P8.3, phosphate buffer pH 8.3; r, acceptor-to-donor distance; R0, Förster radius; τ, fluorescence lifetime; Tyr, tyrosine; Tyr*, excited state tyrosine; Tyr−, tyrosinate anion; Tyr−*, excited state tyrosinate anion * Corresponding author. E-mail address: [email protected] (M. Prokopowicz). http://dx.doi.org/10.1016/j.bpc.2017.09.001 Received 11 July 2017; Received in revised form 25 August 2017; Accepted 7 September 2017 Available online 19 September 2017 0301-4622/ © 2017 Elsevier B.V. All rights reserved.

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confirmed the indispensable role of the modification of the FRET efficiency (E) calculation on the fraction of enzyme saturation in PNP-FA systems.

1. Introduction

necessary [26,35]. This is a great advantage in such experiments, as it not only lowers the costs and experimental time, but allows simultaneous direct observation of the donor and acceptor fluorescence. On the other hand, the stoichiometry of the FA binding to E. coli PNP leads to the equilibrium state between free and bonded ligand. It is a cause of the concentration dependent efficiency values of the FRET in the enzyme-ligand pair such as PNP-FA, calculated with the conventional approach. Thus, as presented in this study, the real efficiency of the FRET should be calculated with the correction that takes into account a fraction of the fluorescent species that participates in the interaction. A similar method was used to calculate the transfer efficiency between the Gapped DNA and Rat polymerase β [27]. However, a detailed description of the method was not presented in the aforementioned work. This work is a continuation and an extension of a previously published study, where the lack of FRET was observed in complexes of PNP mutants with FA [28]. The current study is an attempt to explain this uncommon phenomenon in more detail. In particular, a series of emission spectra (enzymes, FA and complexes) in four different environments were registered. Subsequently, FRET efficiency and radii were computed using the collected data. Therefore, a two important aspects of this work emerge: 1) biological, that is focused on the characterization of the interactions between E. coli PNP and FA; 2) (bio) physical, which is concentrated on the ‘lack of FRET’ phenomenon.

Purine nucleoside phosphorylases (PNPs, E.C. 2.4.2.1) – catalysts of the reversible purine nucleosides phosphorylation in vivo – play a significant role in PNs metabolic pathways. PNP catalyzes the reaction according to the following scheme [1]: β-nucleoside + Pi ⇔ purine base + α-D-pentose-1-phosphate PNP is an ubiquitous enzyme expressed in bacteria, parasites and mammals. Escherichia coli PNP (further also called PNPWT) is a highmolecular mass enzyme as it has been reported to be a hexamer (trimer of dimers) of identical subunits [2,3]. It differs in the ligand specificity from human PNP, a low-molecular PNP (trimer) [4–8]. Therefore, PNPWT is thought to become a suicide gene, a tool in a cancer therapy. First in-human studies show that PNPWT can be safely used as an activator of fludarabine, which leads to a tumor size decrease [9–14]. In spite of crucial differences in the sequence between low- and high-molecular PNPs and their interaction mechanism, their structure and arrangement of an active sites are very similar [15,16]. In addition, it has been shown that Phe200 in human PNP is a residue that takes an important part in nucleotide binding [16,17]. In PNPWT, an equivalent residue is a Phe159 [16], which is though to contribute to the purine ring binding. Additional, neighbouring Tyr160 residue might also participate in the binding via π–π interactions with Phe159. Therefore, in order to investigate the role of Phe159 and Tyr160 in PNPWT, we have examined two mutants – PNPF159Y (PNPY) and PNPF159A (PNPA) – in complexes with formycin A (FA). This approach might result in new insights into the mechanism of the interactions and indicate directions towards more efficient anti-tumor therapies. Formycin A (Fig. 1), as a fluorescent derivative of adenosine with a C–C glycosidic bond that PNP cannot cleave, is an excellent nucleoside binding model. It is a specific inhibitor of E. coli PNP, but not human, and possess significant antiviral activities [18–20]. Few works describes interactions of FA with E. coli PNP and their crystal structures [21,22]. According to the fluorescent studies, it was shown that in spite of N(1)H FA tautomer dominance in solution, PNP has a strong preference to N (2)-H tautomer binding [19,23,24]. This lies behind a mechanism of N (7) nucleoside protonation by Asp204 [25]. Moreover, crystal structure of PNP-FA-Pi complex indicated that only one monomer in dimer can adopt closed conformation, which is connected with the conformational change of H8 helix [22]. Despite of many published works about PNP catalysis and binding, their mechanisms are still not fully understood, which leaves a vast field for further investigations. The presence of 36 Tyr in E. coli PNP sequence, with Tyr160 being the closest tyrosine to the nucleoside binding site (ca. 8–10 Å) [26,28] and the red-shifted absorption spectrum of FA relative to PNPWT, enables the observation of fluorescence resonance energy transfer (FRET) in Tyr →FA direction [19,26]. Hence, in this work we have engaged the optical and time-resolved spectroscopy to study the in-solution interactions between E. coli PNP mutants with FA in the presence and absence of Pi, pH 7 and 8.3. Conducting experiment in alkaline conditions is an important aspect of this study, as cancerogenesis may lead to the alkalization of cells [29,30]. Fluorescence/Förster resonance energy transfer (FRET) is known to be a potent tool for studies of molecules interactions, protein folding, lateral diffusion and many other phenomena. It is applicable when distances between energy donors and acceptors are within the range from 10 Å to even 100 Å, depending on the fluorescent species [31–34]. FRET can be applied even to non-fluorescent species if fluorescent probes (dyes) are introduced into the studied system. In the presented case, however, both PNPWT (Tyr presence) and FA have fluorescent properties, therefore additional dyes are not

2. Materials and methods 2.1. Materials Solutions of formycin A, the wild type enzyme and enzyme mutants were expressed, purified and prepared according to Ref. [28]. 2.2. Steady-state absorption and fluorescence Ultraviolet absorption was monitored with a Varian (Australia) Cary 50 spectrophotometer, equipped with a cell compartment thermostatically stabilized at 25 °C, using 5 mm path-length cuvettes. Steadystate fluorescence emission and excitation spectra were measured with a Spex (USA) FluoroMax spectrofluorimeter at 10 °C and 25 °C, set up in the single photon counting mode, with 4-nm spectral resolution for excitation and emission. Samples were prepared in 500–570 μl of 50 mM hepes or phosphate buffers, both at pH 7 and pH 8.3 in 5 × 5 mm Suprasil cuvettes. Fluorescence emission (excited at λex 280 nm, 295 nm, 305 nm and 313 nm) spectra of enzymes, FA and their mixtures were recorded. Fluorescence spectra were corrected for the background emission of the buffers and for the inner filter effect (e.g. Eq.(1)). Concentrations of formycin A and enzymes were determined spectrophotometrically in each buffer at the absorption maxima at 294 nm and 278 nm, respectively. The utilized concentrations of FA were in a range of 1–27 μM. Concentrations of the enzymes were in a

Fig. 1. Two tautomers of formycin A in solution. Left N(1)-H, right - N(2)-H.

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range of 2–3 μM.

G = antilog10 ⎛ ⎝

δAex + δAem ⎞ 2 ⎠

resulting from the formation of their complex are visualized by total difference spectra. They are calculated by subtracting spectra of free FA and enzymes from the spectra of their mixtures. All fluorescence spectra were corrected for inner the filter effect (Eq. (1)).

(1)

where δAex and δAem are absorption changes at the excitation and emission wavelengths, respectively.

2.5. Fluorescence lifetimes

2.3. Analysis of energy transfer parameters

Fluorescence lifetimes were measured using the time-correlated single-photon counting (TCSPC) technique with sub-nanosecond resolution using a commercial time-domain spectrofluorimeter (ChronosBH, ISS). A Ti:Sapphire laser (Mai-Tai, Spectra Physics) equipped with a third harmonic generator was used to excite samples at λex = 281 nm. Emission was observed through a band pass filter with the maximum of transmission at 310 nm. Temperature of the samples was stabilized at 25 °C. Time-resolved fluorescence data were fitted with sums of one, two and/or three exponential decays (Eq. (8)), convoluted with the instrument response function, using the ISS software, Vinci2. Concentrations of formycin A and enzymes were determined spectrophotometrically in each buffer at the absorption maxima at 294 nm and 278 nm, respectively. The utilized concentration of FA in PNPWT/ PNPY-FA complexes was equal to 10 μM and in PNPA-FA complexes to 2 μM. Concentrations of enzymes were in a range of 2–3 μM.

Energy transfer was quantified by calculating the Förster radius (R0, a distance where efficiency of the energy transfer equals to 50% [33]), the acceptor-to-donor distance r and the efficiency of the transfer E computed by the donor (PNP) emission quenching. The Förster radius was computed with Eq. (2) for each enzyme–ligand pair in all experimental systems.

R 0 = 0.211(κ 2×n×Q×J )1/6

(2)

where R0 is the Fö rster radius, κ is the orientation factor, arbitrary taken as 2/3, n is the refractive index (here 1.4), Q is the quantum yield of the donor (0.14) and J is the overlap integral given with the formula: 2

J=

∫0

∞

FD (λ )ϵA (λ ) λ4dλ

(3)

where FD(λ) is the fluorescence intensity of the donor, ϵA is the extinction coefficient of the acceptor and λ is wavelength expressed in nanometers. Missing blue parts of enzymes spectra were fitted if necessary and the overlapping parts of enzyme emission and ligand absorption spectra were calculated. Ligands absorption spectra were expressed in [M cm− 1 nm− 1] units and emission spectra were normalized. As it was mentioned before, two methods of calculating r and E were employed. In the conventional method the transfer efficiency (Ec) and the acceptor-to-donor distance (rc) were calculated using the following equations (Eqs. (4) and (5), respectively):

(4)

1/6 1 r = R 0 ⎛ −1⎞ ⎝E ⎠

(5)

where αi is the amplitude associated with the decay time τi. The fractional intensity of the decay fi is given by fi = αiτi/Σαiτi and the mean decay time can be calculated as < τ > =Σfiτi. The quality of the fit was evaluated by the structure observed in residuals plots and by the χR2 values. 3. Results

⎟

Spectral properties of E. coli PNP and FA are well described in literature [35], thus they will not be presented in this paper. However, it is worth mentioning that in the case of the PNPY mutant, after excitation at λex = 280 nm, the red-part of the emission is elevated relative to the PNPWT (Fig. 2). This effect can be attributed to the tyrosinate anion (Tyr−), that is formed due to the presence of two adjacent tyrosines at positions 159 and 160 (Fig. 2, insets) [28,36]. A change to other buffers did not influence the vibrational structure of the enzymes' spectra. The absorption maximum of enzymes is located at 280 nm, and the fluorescence maxima at 304 nm (PNPWT, PNPA) and 307 nm

where FMix and FD is the registered fluorescence intensity of the enzyme-ligand complexes and the donor alone in solution, respectively. However, the interaction between FA and the enzyme is not a simple dynamic process in solution but a more complicated process of association. Thus, a new approach to calculate values of E was introduced. In this approach dissociation constants, calculated for each experimental system, were used, and the interacting fraction of enzyme monomers α (Eq. (6)) was calculated:

α =

(8)

3.1. Fluorescent properties of mutants

F Ec = 1 − ⎛ Mix ⎞ ⎝ FD ⎠ ⎜

I (t ) = Σi αi exp(−t / τi )

1 ([L0]+[M0]+K d 2[M0] ([L0]+[M0]+K d )2 −4×[L0][M0] )

−

(6)

where [M0] is the total concentration of enzyme monomers, [L0] is the total concentration of the ligand (here FA) and Kd is the dissociation constant calculated from spectroscopic data (not shown). After α had been calculated, the energy transfer efficiency (Eα) was computed using the modified equation (Eq. (7)):

F 1 Eα = ⎛1 − Mix ⎞ FD ⎠ α ⎝ ⎜

⎟

(7)

As in the conventional approach, the acceptor-to-donor distance (rα), was calculated according to the Eq. (5), where modified Eα was utilized. Fig. 2. Fluorescence spectra of PNPY (Y), PNPA (A) and wild type PNP (WT) in hepes pH 7 at λex = 280 nm. The upper inset shows difference spectra between PNPY and PNPWT (—) and PNPA and PNPWT (- -). The lower inset presents emission spectra of PNPY at λex = 295 nm.

2.4. Total difference spectra Changes of fluorescence properties of both the enzyme and FA 101

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λex = 280 nm. A small increase of the emission intensity in the red-part of spectra is displayed. The maximum of spectra is shifted to 363 nm. Simultaneously, a considerable decrease of the enzyme emission is observed with the minimum shifted bathochromically(relative to hepes at pH 7) to 310 nm (Fig. 6A). Although identification of an isoemissive point is not straightforward, the most probable spot is around 337 nm. Difference spectra of PNPY-FA recorded at λex = 295 nm (Fig. 7A) testify to the quenching of the emission of both, the enzyme and FA. Well shaped vibrational structure can be seen, with minima positions corresponding to the N(1)-H FA tautomer. An increase of the FA emission is observed at λex = 313 nm (Fig. 8A), and it is several times higher than in the hepes buffer (Fig. 5A). A clear vibrational structure is displayed with three peaks at 331 nm, 347 nm, and 362 nm, the same as in hepes at pH 7. The inset in Fig. 8A displays a total difference emission spectra at λex = 305 nm. The presence of Pi changes their pattern by slight increase of the left-part of th FA emission difference spectra relative to the hepes buffer (Fig. 5A inset).

(PNPY). Absorption and emission maxima of FA are located at 295 nm and 340 nm, respectively. 3.2. PNPY-FA complexes in hepes Total difference spectra of PNPY-FA complexes in the hepes buffer reflect different behaviour that strongly depends on pH of the environment. 3.2.1. pH 7 Although the enzyme emission considerably drops down during titration with FA, after excitation at λex = 280 nm, only a small gain of the FA emission is observed in the hepes buffer at pH 7 (Fig. 3A). Red parts of the total difference spectra are positive with the maximum shifted from 340 nm (emission maximum of free FA) to 347 nm. This is in slight contrast to the previous experiments [28]. However, alterations in a biological samples may occur. These results are the average of several repetitions. Simultaneously, a potential isoemissive point at 326 nm is perturbed (Fig. 3A) and a small drift of the minimum (304–306 nm) is observed with concomitant FA concentration increase. Excitation at 295 nm corresponds to the FA and Tyr− excitation maximum. For the PNPWT-FA complexes, excitation at this wavelength (not shown) causes a considerable increase of the FA emission spectrum. However, in the PNPY-FA complexes (Fig. 4A) mostly quenching of the FA emission is observed. A vibrational structure, with peaks at 331 nm, 347 nm, and 363 nm, can be seen in the spectra. The intensities of the difference spectra oscillate around zero (Fig. 4A). After excitation at λex = 313 nm (Fig. 5A), characteristic absorption maximum of the N(2)-H tautomer, where Tyr is not excited and Tyr− has low absorption [37,38], a positive total difference emission spectra were calculated. FA binding to the active centre enhances the rigidity of its environment and minimizes the solvent effects, thus it might increase the emission intensity, as it has been reported for the wild type enzyme [35]. However, the FA emission increase is not as significant as that observed for the WT PNP. Excitation at λex = 305 nm (inset Fig. 5 A) results in both positive and negative total difference spectra, which values are close to zero.

3.2.5. pH 8.3 After excitation at λex = 280 nm an enhanced increase of the FA emission intensity is displayed (Fig. 6B) relative to the previously described conditions. With a concomitant increase of the FA concentration an increase of its emission and the decrease of the Tyr emission is observed. A minimum of the enzyme emission quenching is motile and shifts from 303 nm to 306 nm for the highest FA concentration. An isoemissive point, however diffused, might be placed at 323 nm (similar to Fig. 3A) and is bathochromically shifted with the respect to the isoemissive point of PNPWT-FA complexes (315 nm) [35]. The maximum of the total difference spectra is flattened and placed at 355 nm, without a characteristic vibrational structure. Difference spectra of PNPY-FA recorded at λex = 295 nm, unlike at pH 7, testify to an increase of the FA fluorescence (Fig. 7B). The spectra are smooth with no clearly marked peaks and the maximum at 346 nm. Spectra recorded after excitation at λex = 313 nm (Fig. 8B) are positive and ca. two times higher than in pH 7 (Fig. 8A) and three times

3.2.2. pH 8.3 Calculated difference spectra at λex = 280 nm are negative in the whole range (Fig. 3B). The minimum of the enzyme quenching is slightly red-shifted from 306 nm to 308 nm relative to the minimum at pH 7 (Fig. 3A) and its small drift (2–3 nm) is observed. Total difference spectra are smooth, without any visible vibrational structure that could point to interactions with a specific tautomer. However, the extent of the enzyme quenching is comparable with that observed at pH 7. One should notice that the quenching of the emission at λex = 295 nm is much stronger in hepes at pH 8.3 (Fig. 4B) than at pH 7 (Fig. 4A). Total difference spectra (Fig. 4B) reflect a reciprocal of the shape of the FA emission spectrum in solution with minima at 326 nm, 339 nm and 355 nm. Positive total difference emission spectra were calculated at λex = 313 nm (Fig. 5B). They are characterized by a visible vibrational structure with three sharp peaks at 331 nm, 346 nm, and 360 nm, that are assigned to the N(2)-H tautomer. Nevertheless, their intensities are slightly lower than in pH 7. Excitation at λex = 305 nm (Inset Fig. 5B) results in the negative difference spectra. 3.2.3. PNPY-FA in phosphate buffer Incorporation of Pi to the environment changes the pattern of total difference spectra of PNPY-FA complexes, mostly at pH 8.3, when compared to the difference spectra collected in the hepes buffers. Fig. 3. Total difference emission spectra of PNPY-FA complexes with increasing FA concentration after excitation at λex = 280 nm. (A): total difference spectra in 50 mM hepes pH 7; (B): total difference spectra in 50 mM hepes pH 8.3.

3.2.4. pH 7 Fig. 6A presents a total difference emission spectra recorded at 102

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Fig. 4. Total difference emission spectra of PNPY-FA complexes with increasing FA concentration after excitation at λex = 295 nm. (A): total difference spectra in 50 mM hepes pH 7; (B): total difference spectra in 50 mM hepes pH 8.3.

Fig. 6. Total difference emission spectra of PNPY-FA complexes with increasing FA concentration after excitation at λex = 280 nm. (A): total difference spectra in 50 mM phosphate buffer at pH 7; (B): total difference spectra in 50 mM phosphate buffer at pH 8.3.

Fig. 5. Total difference emission spectra of PNPY-FA complexes with the increasing FA concentration after excitation at λex = 313 nm. (A): total difference spectra in 50 mM hepes pH 7; (B): total difference spectra in 50 mM hepes at pH 8.3. The insets display total difference spectra after excitation at λex = 305 nm in the reference buffer.

Fig. 7. Total difference emission spectra of PNPY-FA complexes with increasing FA concentration after excitation at λex = 295 nm. (A): total difference spectra in 50 mM phosphate buffer at pH 7; (B): total difference spectra in 50 mM phosphate buffer at pH 8.3.

higher than in hepes at pH 8.3 (Fig. 5B). The inset in Fig. 8B displays a total difference emission spectra at λex = 305 nm. The presence of Pi in pH 8.3 leads to the increase of the FA emission intensity relative to the hepes (both pH) and Pi buffer at pH 7.

3.3. PNPA-FA complexes Total difference spectra obtained from PNPA-FA complexes (Supplementary material, SFig.1–SFig.4) are characterized by weak quenching of the enzyme's fluorescence in each buffer after λex = 280 nm. However, in pH 7 (H7, P7), the increase of FA emission 103

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of 0.48 in P7 at 25 °C (Table 2). On the other hand, introduction of α in calculations of FRET parameters for the PNPFY-FA and PNPWT-FA complexes has a visible positive impact the FRET efficiency and a acceptor-to-donor distance values in each experimental system. An interesting observation is that in all experimental systems E value is higher in PNPY-FA than in PNPWT-FA complexes, with the highest computed efficiency in P7 at 10 °C. Although the transfer efficiency is quite high in both types of complexes (with PNPY and PNPWT), virtual radii are located in the range of 16–19 Å, which is almost twice a distance between the closest Tyr and FA in PNPWT [21] and almost four times that distance in PNPY (for explanation see 4). Values of r resemble distances of other than Tyr159/160 tyrosine residues that are located within a distance of ca. 13–18 Å from FA. One should notice the dependence of E on temperature (Tables 1 and 2). With the increase of temperature, values of (E) for PNPWT and PNPY complexes decrease. In contrast, the opposite behaviour is observed for PNPA. 3.5. Fluorescence lifetimes 3.5.1. Temperature 25 °C Table 3 presents the fluorescence lifetimes (τi) with their fractional intensities (fi) and the average fluorescence lifetimes (τav) of enzymes and their complexes (Mix) in each experimental system. Three decay components are necessary to describe fluorescence kinetics of the PNPY mutant whereas only two components are found in the case of PNPA and PNPWT. The longest average fluorescence lifetime was registered for the PNPA mutant and the shortest one for the PNPY mutant. The longest components of the fluorescence decays of PNPWT and PNPY are very similar (ca. 2.75 ns), however with lower fractional intensity (fi) in the case of PNPY mutant. Corresponding component of the PNPA mutant (2.8–3 ns) is longer than for PNPWT and PNPY and simultaneously has higher f than PNPY (but similar to PNPWT) in all buffers. The short component of the PNPWT fluorescence decay comprises of 0.12–0.15 of the total intensity fraction and varies between 0.23 and 0.37 ns, similarly to the previous results [35]. Similar fractional intensity was registered for the PNPA mutant, however, its short component is longer than in PNPWT and varies between 0.37 and 0.77 ns. The corresponding component in the PNPY mutants constitutes ca. 0.1 of total intensity and varies between 0.2 and 0.26 ns. Moreover, the appearance of the additional 3rd component (1.08–1.3 ns) with the fractional intensity similar to the shortest one was registered. After FA addition, the behaviour of the average fluorescence lifetime time has changed. The shortest τav represents the PNPWT and the longest one — the PNPA mutant. The longest component of the fluorescence lifetimes of PNPY-FA complexes in the hepes buffers is the same as for the enzyme itself, however with a significantly lower fractional intensity (in all buffers). On the other hand, an increase of this component in Pi buffers is observed relative to the PNPY solely in solution. The 3rd component has slightly decreased in both hepes buffers while after Pi addition it has raised. The 2nd (shortest) component always

Fig. 8. Total difference emission spectra of PNPFY-FA complexes with increasing FA concentration after excitation at λex = 313 nm. (A): total difference spectra in 50 mM phosphate buffer at pH 7; (B): total difference spectra in 50 mM phosphate buffer at pH 8.3. The insets display total difference spectra after excitation at λex = 305 nm in the reference buffer.

is stronger than in PNPY-FA complexes, while in hepes pH 8.3 quenching of FA is observed. The excitation at λex = 313 nm, hepes both pH, results in difference spectra oscillating around zero. Addition of Pi results in increase of the difference spectra with the concomitant FA concentration increase. 3.4. FRET parameters FRET efficiency (E) and a acceptor-to-donor distance (r) were calculated at two temperatures for each of the complexes (PNPWT/PNPY/ PNPA-FA). Although it is quite artificial to describe the observed effects in terms of FRET parameters, as the total difference spectra do not testify to its occurrence, the parameters can be useful as a measure of the strength of the interaction (enzyme fluorescence quenching). Table 1 presents averaged values of the transfer efficiency and virtual radii calculated with the new approach that utilizes the α factor. Table 2 summarizes averaged values of the transfer efficiency and virtual radii calculated with the conventional approach. Values of the FRET parameters for PNPA-FA complexes are insufficient to confirm a binding of FA to the active site. Moreover, in the hepes buffers they do not display noticeable differences between the conventional and the new approach. However, addition of Pi at both pH noticeably enhances the FRET efficiency that reaches the highest value

Table 1 Acceptor-to-donor distance in Å and FRET efficiency calculated from Eqs. (5) and (7), respectively, for the mutants and the wild type PNP in four different environments at two different temperatures. PNPY

H7 H8 P7 P8

Eα 10 °C 0.77 0.76 0.81 0.79

PNPA

25 °C 0.75 0.68 0.75 0.73

rα 10 °C 16.9 17 16.2 16.5

25 °C 17 18.2 17 17.4

Eα 10 °C 0.08 0.09 0.36 0.14

PNPWT

25 °C 0.14 0.07 0.48 0.27

104

rα 10 °C 31 30.3 22.6 27.7

25 °C 28 32.2 20.9 24.4

Eα 10 °C 0.73 0.66 0.7 0.78

25 °C 0.63 0.65 0.68 0.7

rα 10 °C 17.45 18.7 17.84 16.7

25 °C 19.06 18.5 18.2 18

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Table 2 Acceptor-to-donor distance in Å and FRET efficiency calculated from Eqs. (5) and (4), respectively, for the mutants and the wild type PNP in four different environments at two different temperatures. PNPY

H7 H8 P7 P8

PNPA

Ec 10 °C 0.45 0.47 0.53 0.47

25 °C 0.41 0.41 0.49 0.31

rc 10 °C 21.3 21.1 20.3 21.08

25 °C 21.9 21.6 20.85 23.5

PNPWT

Ec 10 °C 0.07 0.09 0.16 0.09

25 °C 0.09 0.05 0.11 0.07

rc 10 °C 31.5 30.5 27.3 29.9

25 °C 30.4 33.5 29.02 31.4

Ec 10 °C 0.46 0.45 0.51 0.43

25 °C 0.39 0.39 0.4 0.35

rc 10 °C 21.13 21.65 20.5 21.67

25 °C 22.1 22.3 22.01 22.9

increases. The two decay components of the PNPA mutant, independently of the environment, always decline, with a simultaneous increase of the fractional intensity of the short component. It is worth noticing that the most significant reduction of f is observed in H8 and P7, where the strongest enzyme quenching is observed (Supplementary material SFig. 1B and SFig. 3A). After FA addition to the PNPWT a significant decrease of both, τ and f of the long component, is observed. The strongest fluorescence lifetime quenching is observed in the P7 buffer and the lowest value of f is calculated for the P8 buffer. The length of the short component increases in all examined buffers.

PNPWT/PNPY-FA or PNPA-FA complexes, respectively. The tendency of fractional intensities is similar in all types of complexes, as f decreases for the long component and increases for the short one.

3.5.2. Temperature 10 °C At lower temperature (Table 4) a change of the fluorescence lifetimes of each enzyme is observed. For both, the PNPY mutant and PNPWT, a surprising decrease of long component τ is observed. Moreover, an increase of the fractional intensity of PNPY is registered, while for PNPWT f remains nearly constant. On the other hand, both τi of PNPA are noticeably enhanced while the corresponding fractional intensity f is preserved. The significant augmentation of the PNPA average fluorescence lifetime is observed whereas the τav of PNPWT has slightly decreased or remained constant in hepes and Pi buffers respectively. At the lower temperature the disappearance of the 3rd component and the decrease of τav in most of the buffers is observed for the PNPY mutant. The increase of the PNPA and PNPWT average fluorescence lifetimes (in general) with concomitant FA addition is registered at 25 °C, suggesting that collisional quenching takes part in both complexes. On the other hand, the PNPY τav is visibly shorter at lower temperature than at 25 °C. FA addition to the enzymes provokes the decline of the long component whereas it causes the increase or decrease of the short one in

4.1. Spectra

4. Discussion The experiments brought out numerous facts regarding the energy transfer in the studied complexes, which indicate a potential solution of the discussed problem. They clearly show that F159A substitution is responsible for the essential loss of FA binding to the active centre. Results obtained for PNPY-FA complexes are more ambiguous, however, they suggest positive impact of Tyr159 on FA binding.

Low enzyme quenching and the lack of preferences in the PNPA-FA complexes testify to the collisional quenching and thus that the aromatic amino acid at 159 position is essential for the efficient FA binding to the active site. Results obtained for PNPY-FA complexes are not trivial to interpret. The extent of the Tyr emission quenching in all experimental systems suggests a strong FA association to the active site. However, the lack or a minute FA fluorescence gain complicates the analysis of the results. The starting point for a further discussion is the existence of an additional fluorescent species — a tyrosinate anion (Tyr−/Tyr−*) in the Tyr159–Tyr160 pair [28,36]. As a result of the bathochromic shift of Tyr− emission relative to Tyr and its low quantum yield [36], the spectral overlap of the donoracceptor (Tyr−– FA) pair in the complexes is virtually gone, which might explain the lack of energy transfer from 159/160 tyrosine residue to the FA after excitation. A minor gain of the FA fluorescence (in some cases) after excitation at λex = 280 nm is insufficient to reliably indicate FRET. The increase of the FA fluorescence might be also a result

Table 3 Fluorescence lifetimes [ns] and their fractional intensity decays at 25 °C in four environments. Samples were excited at λex = 281 nm. The emitted light was filtered with a band pass filter with the transmission maximum at 310 nm. According to the literature, the binding stoichiometry is two or three FA molecules per a PNP hexamer [21,22]. PNPY

H7

τav H8

τav P7

τav P8

τav

Mix

τi 2.8 0.26 1.2

fi 0.72 0.12 0.16

τi 2.8 0.4 1.16

0.75 0.1 0.15

2.7 0.32 0.95

0.75 0.11 0.14

2.9 0.39 1.16

0.76 0.12 0.12

2.85 0.455 1.57

2.21 2.75 0.2 1.08

2.25

τi 3 0.61

0.35 0.3 0.35

2.8 0.72

0.25 0.4 0.35

3 0.77

0.25 0.48 0.27

2.85 0.37

fi 0.85 0.15

τi 2.85 0.53

0.8 0.2

2.6 0.56

0.8 0.2

2.75 0.58

0.88 0.12

2.65 0.38

PNPWT fi 0.67 0.33

τi 2.7 0.37

0.37 0.63

2.7 0.312

0.4 0.6

2.8 0.36

0.78 0.22

2.75 0.23

2.06

2.38

1.26

1.37

Mix

2.58

1.35

2.25 2.7 0.24 1.3

fi 0.25 0.45 0.3 1.23

2.2 2.75 0.23 1.14

PNPA

2.15

105

τi 2.05 0.6

0.85 0.15

1.9 0.46

0.85 0.15

1.6 0.51

0.88 0.12

2 0.6

0.5 0.5 1.2

2.43

2.45

fi 0.4 0.6 1.18

2.34

1.41

2.54

fi 0.87 0.13 2.4

1.31

2.54

Mix

0.45 0.55 1 0.33 0.67 1.06

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Table 4 Fluorescence lifetimes [ns] and their fractional intensity decays at 10 °C in four environments. Samples were excited at λex = 281 nm. The emitted light was filtered with a band pass filter with the transmission maximum at 310 nm. According to the literature, the binding stoichiometry is two or three FA molecules per a PNP hexamer [21,22]. PNPY

H7 τav H8 τav P7 τav P8 τav

Mix

τi 2.6 0.34

fi 0.85 0.15

τi 2 0.66

0.8 0.2

1.7 0.68

0.8 0.2

1.35 0.65

0.9 0.1

1.7 0.43

2.26 2.2 0.49

2.1

τi 3.75 1.1

0.30 0.7

3.35 0.97

0.30 0.7

3.5 1

0.55 0.45

3.3 0.78

fi 0.8 0.2

τi 3.65 0.67

0.8 0.2

3.4 0.79

0.8 0.2

3.3 0.7

0.85 0.15

3.2 0.56

PNPWT fi 0.35 0.65

τi 2.45 0.21

0.65 0.35

2.45 0.33

0.6 0.4

2.65 0.28

0.7 0.3

2.65 0.24

1.7

2.87

0.86

1.13

Mix

3.2

1

2.02 2.3 0.2

fi 0.35 0.65 1.13

1.86 2.4 0.48

PNPA

τi 1.7 0.33

0.9 0.1

1.95 0.48

0.9 0.1

1.75 0.56

0.9 0.1

1.8 0.56

2.42

0.85 0.15 1.74

2.43

2.4

fi 0.9 0.1 1.54

2.23

2.26

2.92

fi 0.9 0.1 2.24

2.48

3

Mix

0.73 0.27 1.44 0.75 0.25 1.44

ligand is bound to the active centre it has an access to the donor's energy. Therefore, the real extent of fluorescence quenching is higher than it would result from simple donor–acceptor efficiency equation (Eq. (4)). Therefore, the Eα values, in contrary to the Ec, becomes independent of the FA concentration.

of the enhanced rigidity of the enzyme environment [31,35]. According to the previous results [28], the arrangement of the 159–160 pair that promotes or inhibits FRET from Tyr160 is possible. The most recent crystallographic studies [21] show that the arrangement of FA-Phe159-Tyr160 (in PNPWT crystal) actually resembles the one that inhibits FRET from Tyr160. However, (see FRET parameters), our results suggest that energy is passed to the FA not solely from Tyr160, as it was suspected [35], but also from other tyrosine residues. Probable FRET restoration in the presence od Pi at pH 8.3 (Fig. 6B), might be explained by a negative charge compensation. At pH 8.3, the Pi shifts its equilibrium in favor to HPO 24−, which is incorporated into the active site [39]. Simultaneous increase of the amount of Tyr− and FA−, is followed by increase of the local negative charge. A protonation of one of Tyr159/160 becomes indispensable in a such situation, provokes the appearance of FRET (Fig. 6B) and stabilizes the environment (Fig. 7B). Presented data indicate that Tyr− has a major impact on the energy transfer from tyrosines other than just Tyr160, in spite of its small fluorescence quantum yield and only a minute contribution to the emitting molecules. Nevertheless, in some circumstances, a modified local density of optical states (here: a change form Tyr* to Tyr−*) might lead to the photonic background modification, which in turn might indirectly influence FRET [40].

4.3. Fluorescence lifetimes 4.3.1. Enzymes The aromatic pair of 159–160 amino acids has a noticeable impact on the average Tyr fluorescence lifetime. Fractional intensity values of PNPY confirm the presence of Tyr−* formation that depletes the population of Tyr*. Moreover, the registered 3rd decay component (ca. 1.1) probably belongs to the Tyr−*, however for now it is not certain. The unusual decrease of the fluorescence lifetimes (of PNPWT/ PNPY at 10 °C) has also been observed in the cytochrome c [43]. This suggest a presence of an additional structure. In particular, the presence of aromatic molecules in a close proximity might result in higher order molecular couplings (e.g., an excimer formation) [31], which probably occurs in PNPWT and PNPY. That newly created structure is destabilized when the temperature increases leading to the restoration of the Tyr fluorescence lifetime. Moreover, the disappearance of the third component at 10 °C might also confirm the formation of an additional structure. On the contrary, increased τ of PNPA (relative to the 25 °C) is a result of diminution of enzyme's dynamic motions.

4.2. FRET parameters Large values of r (Table 1) for PNPWT/PNPY-FA complexes strongly suggest that energy transfer is passed (also) from other (than Tyr159/ 160) tyrosines located in the enzymes. This lies in a contrary to the previous hypothesis [35]. Each tyrosine has a specific probability to donate excitation energy that depends on its orientation and a distance from an FA molecule [31–33]. In a proximity of FA, there are several tyrosines in a distance shorter than R0. Therefore, inhibiting position solely is insufficient to explain the absence of FRET. Higher values of E and r for PNPY-FA complexes relative to PNPWTFA complexes (Tables 1 and 2) suggest a positive impact of Tyr159 on the binding efficiency. An additional hydroxyl group might tighten the FA bonding by forming HB with the nucleoside proton or can engage H+–π interaction (if Tyr is in the neutral form) [41,42]. Tyr160 could also form a putative hydrogen bond with e.g., Arg217 Nη2 as a cause of a closed conformation state (predicted by VMD programme) [22]. Comparison of the efficiency transfers calculated by both methods: the one using α (Table 1, Supplementary material SFig. 5A) and the conventional one (Table 2, Supplementary material SFig. 5B), clearly shows that a correction of the dissociation constant Kd must be taken into account. Calculation of energy transfer in the enzyme-ligands systems is more complicated as a cause of association process and the fact that energy donors are hidden within protein structure. Only after

4.3.2. Complexes The decrease of fluorescence lifetimes long components, together with f at both temperatures in PNPA-FA complexes is in a good agreement with the mechanism of the collisional quenching. Results obtained for PNPWT-FA complexes also suggest that collisional quenching takes part in the enzyme fluorescence decrease. As mentioned before, crystallographic studies show that in PNPWT-FA-Pi crystal, only two/three of six active sites are in the closed conformation [21,22]. Therefore, the presence of dynamic equilibrium of dissociation-association processes of FA molecule is possible, causing the effect of mixed quenching. On the other hand, preservation of the longest τ values and essential f decrease in the PNPY-FA complexes relative to the PNPWT in the 25 °C, strongly suggest the presence of static quenching solely. Hence, it seems that the F159Y mutation has strong impact on the FA binding. However, with the temperature decrease, situation turns into probably mixed quenching (as in PNPWT). This ambiguous and not trivial to interpret dependence might be in accordance with aforementioned newly created structure in 159–160 pair (see Section 4.3.1). This new ‘molecule species' might change local conformation and impede the FA binding, therefore causing the increase in collisional quenching. Nevertheless, one should notice that the 106

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FA binding still seems to be stronger in PNPY than in PNPWT. This would suggest that the 159–160 pair has a great impact on the enzyme binding and spectral properties. 5. Conclusions In a view of the above, it seems that the observed FRET in the PNPWT-FA complexes is an averaged process of the emission quenching of several Tyr residues and in the PNPY-FA complexes one (or all) of the following takes place: (1) aforementioned inhibiting position arrests the energy transfer from Tyr160; (2) the newly created system of the 159–160 tyrosines pair and the presence of Tyr− somehow inhibit FRET from other tyrosines; (3) the π–π interactions of FA-Phe159-Tyr160 in the wild type enzyme might lead to higher order multipole couplings or exchange interactions e.g., Dexter electron transfer [44]. The FA and Tyr159 (Tyr159−) are in the distance that allows for the orbitals to overlap (Dexter transfer). In assumption that the FA* can transfer an electron to the Tyr− and create Tyr−* we would observe the FA emission quenching or bathochromic isoemissive point shift in the respect to the PNPY-FA complexes. Both of these are observed. To resume: 1. The dissociation constant should be incorporated in FRET parameters calculations for enzyme-ligand complexes; 2. The PNPA mutant is not able to efficiently bind FA alike with and without Pi; 3. The 159–160 amino acids pair has a paramount impact on spectroscopic properties of E. coli PNP; 4. The aromatic amino acid at the position 159 is a key residue for the nucleoside binding; 5. An additional Tyr−/Tyr*− might inhibit the energy transfer from other Tyr to FA or be responsible for Dexter transfer in the FA*→ Tyr− direction; 6. The additional Tyr159 probably enhances the binding efficiency of FA; 7. The phosphate buffer at pH 8.3 seems to restore (partially) FRET in PNPF59Y-FA complexes; Acknowledgments These studies were supported by the Polish Ministry of Scientific Research and Higher Education (grant No. NN202105536). The experiments were carried out at the Department of Biophysics, Institute of Experimental Physics, Faculty of Physics, University of Warsaw. This article is a part of the doctoral dissertation of Malgorzata Prokopowicz orcid="0000-0002-0160-9884". References [1] M.D. Erion, J.D. Stoeckler, W.C. Guida, R.L. Walter, S.E. Ealick, Purine nucleoside phosphorylase. 2, Catalytic mechanism. Biochem. 36 (1997) 11735–11748. [2] J. Tebbe, A. Bzowska, B. Wielgus-Kutrowska, W. Schröder, Z. Kazimierczuk, D. Shugar, W. Saenger, G. Koellner, Crystal structure of the purine nucleoside phosphorylase (PNP) from Cellulomonas sp. and its implication for the mechanism of trimeric PNPs, J. Mol. Biol. 294 (1999) 1239–1255. [3] M.J. Pugmire, S.E. Ealick, Structural analyses reveal two distinct families of nucleoside phosphorylases, Biochem. J. 361 (2002) 1–25. [4] B.K. Kim, S. Cha, R.E. Parks, Purine nucleoside phosphorylase from human erythrocytes. I. Purification and properties, J. Biol. Chem. 243 (1968) 1763–1770. [5] T.A. Krenitsky, Purine nucleoside phosphorylase: kinetic mechanism, and specificity, Mol. Pharm. 3 (1967) 526–536. [6] B.C. Robertson, P.A. Hoffee, Purification and properties of purine nucleoside phosphorylase from Salmonella typhimurium, J. Biol. Chem. 248 (1973) 2040–2043. [7] K.F. Jensen, P. Nygaard, Purine nucleoside phosphorylase from Escherichia coli and Salmonella typhimurium. Purification and some properties, Eur. J. Biochem. 51 (1975) 253–265. [8] E.M. Bennett, C. Li, P.W. Allan, W.B. Parker, S.E. Ealick, Structural basis for substrate specificity of Escherichia coli purine nucleoside phosphorylase, J. Biol. Chem. 278 (47) (2003) 47110–47118.

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