Introduction

Fluorescent proteins, such as green fluorescent protein (GFP), are characterized by their unique β-barrel structure consisting of eleven β strands presented in a cylindrical shape1. This β-barrel encases a chromophore p-HBI (4-(p-hydroxybenzylidene)-5-imidazolinone) within the β-barrel through internal modification of the amino acid triad X-Y-G. Here, Y is tyrosine, G is glycine, and X is an amino acid that varies among different classes of fluorescent proteins2. This chromophore is shielded from the surrounding aqueous environment by the β-barrel, providing stability and a hydrophobic environment required for enabling fluorescence. This unique property of fluorescent proteins makes them an invaluable tool in biology for visualizing cellular processes and protein interactions3. Among fluorescent proteins, green-to-red photoconvertible variants form a subclass of β-barrel fluorescent proteins containing a green chromophore formed through internal modification of H-Y-G amino acid residues, where H is histidine. The exposure to UV light extends the conjugated π-electron system of the green chromophore after a photocleavage event, changing its chemical identity to (2-[(1E)-2-(5-imidazolyl)ethenyl]-4-(p-hydroxybenzylidene)-5-imidazolinone), which we abbreviate as IE-p-HBI. This results in an irreversible shift in fluorescence from green to red (Fig. S1)4. It is worth noting that the green-to-red transformation through main chain photocleavage in this family of proteins occurs within the β-barrel pocket, without the release or any significant shift in the position of the chromophore4. This allows photoconvertible proteins to provide precise spatial and temporal tracking of protein dynamics by altering fluorescence when exposed to specific wavelengths of light5.

The photocleavable fluorescent protein PhoCl1 belongs to a subclass of green-to-red photoconvertible fluorescent proteins and has characteristic green fluorescence conferred by the chromophore p-HBI6. Although similar to other green-to-red photoconvertible fluorescent proteins, it also undergoes a subsequent photodissociation from the β-barrel pocket, generating two species, (1) a β-barrel composed of 230 residues, and (2) a C-terminal peptide fragment (IE-p-HBI-NRVFTKYPR) of 9 amino acids that bear the red chromophore IE-p-HBI (Fig. 1A). The C-terminal peptide fragment IE-p-HBI-NRVFTKYPR will be referred to as CTPF. In contrast to other photoconvertible proteins, this photodissociation process has been reported to render PhoCl1 non-fluorescent by releasing the chromophore from the β-barrel’s hydrophobic environment to the aqueous environment (Fig. 1A)6.

Fig. 1: Photodissociation property of PhoCl1 and generation of red fluorescence through photoexposure to 405 nm light.
Fig. 1: Photodissociation property of PhoCl1 and generation of red fluorescence through photoexposure to 405 nm light.
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A Model of photodissociation of PhoCl1 with a built-in chromophore, p-HBI, within its barrel structure (represented as a cylinder with green emission), B Visual images of PhoCl1 acquired under a UV transilluminator (λ = 365 nm) at different timepoint of unexposed and exposed PhoCl1.Upon photoexposure PhoCl1 solution transitions from green to orange-red C Absorption spectra of PhoCl1 before (Green solid line) and after (Red solid line) exposure to 405 nm light for 12 h. D Fluorescence spectra of PhoCl1 before (Green solid line) and after (Red solid line) exposure to 405 nm light, and (E) The change in the fluorescence (λex = 400 nm, λem = 510 nm) of unexposed PhoCl1 (control) and 405 nm LED exposed PhoCl1 (photocleaved mixture) up to 6 h. Black circles are exposed PhoCl1 fluorescence decay, while the green solid line is unexposed PhoCl1 fluorescence. Relative Fluorescence Units (RFU) have been used as a measure of concentration.

In this work, we provide evidence that the photodissociated CTPF generates dim red fluorescence through aggregation. Dynamic light scattering experiments reveal that CTPF has a strong tendency to aggregate, indicating the formation of larger molecular assemblies. These experimental findings were also supplemented through molecular dynamics simulations to evaluate aggregation and QM/MM calculations to demonstrate that aggregation indeed plays a critical role in the observed red fluorescence.

Results and discussion

PhoCl1 displays the characteristic green light emission (green fluorescence) under the UV-transilluminator (λ = 365 nm) (Fig. 1B). We further subjected PhoCl1 to 405 nm LED (1.5 mWcm-2) for 6 h using our custom-designed light exposure setup (Fig. S2). Surprisingly, this photo exposure led to an unexpected outcome: the emergence of an orange-red colored solution, as seen in visual images at different timepoints (Fig. 1B). This observation is in contradiction with the previously reported findings, where an evident decrease and finally a loss in green fluorescence of PhoCl1 and its fusion partners was detected upon exposure to 405 nm light6,7,8. We characterized the spectral properties of unexposed and photoexposed PhoCl1 using UV-Visible spectroscopy. The samples containing the unexposed PhoCl1 displayed its characteristic absorption spectrum with a maximum at 485 nm (Fig. 1C, Green line)9.

Upon photo-exposure, a noticeable decrease in the absorption maximum of PhoCl1 was observed (Fig. 1C, Red line). The fluorescence spectrum of photoexposed PhoCl1 displayed a new distinct emission peak centered around 570 nm along with the characteristic emission maxima of undissociated PhoCl1 at 510 nm (Fig. 1D). Concurrently, the excitation scan (Fixed emission of 510 nm) of unexposed PhoCl1 showed an increasing trend in the intensity towards larger wavelengths (Fig. S3A), while the photoexposed PhoCl1 displayed the anticipated lower intensity due to photodissociation of PhoCl1 (Fig. S3B). We also recorded the excitation scan (Fixed emission at 570 nm) of the photoexposed PhoCl1, which displayed the presence of a new peak at 320 nm (Fig. S4). The kinetic studies of the photo-dissociation of PhoCl1 revealed notable changes in the absorption profile where an evident decrease in the intensity of 485 nm peak was observed (Fig. S5A, B). Similarly, a clear decrease in the characteristic green fluorescence at 510 nm was observed with time when excited with 400 nm light (Fig. S6A). While the unexposed PhoCl1 showed no decrease in green fluorescence at 510 nm (Fig. 1E). The fluorescence decay of PhoCl1 followed a first-order kinetics model, with a rate constant of ~0.73 h-1 (Fig. 1E). The RFU ratio between the red emission (570 nm) and green emission (510 nm) over time showed a marked increase which indicates the progress of the orange-red emission peak over the green emission peak (Fig. S6B). Additionally, we demonstrate that this could be accelerated by increasing the power density of the LEDs (Fig. S7). Furthermore, photodissociation yield of PhoCl1 has been determined to be 0.0024%. These taken together suggest the generation of orange-red fluorescence through photoexposure of PhoCl1.

To elucidate the origin of orange-red fluorescence after photoexposure of PhoCl1, we employed Ni-NTA chromatography to separate the photocleaved components (empty PhoCl1 barrel, CTPF, and intact PhoCl1). The PhoCl1 and empty-PhoCl1 barrel components possess an N-terminal 6xHis tag, which led us to hypothesize that it could be specifically bound to the Ni-NTA resin. In contrast, CTPF, which does not possess 6xHis tag, would have minimal affinity to the resin. This would enable CTPF to be effectively washed and subsequently collected for further characterization. Indeed, the collected supernatant, which corresponds to the fraction that dissociated from the Ni-NTA resin-bound 6xHis-PhoCl1 upon light irradiation, contained pure CTPF. The collected supernatant of the photoexposed PhoCl1 containing pure CTPF exhibited red fluorescence under the UV-transilluminator (λ = 365 nm) (Fig. 2A). Additionally, we corroborated the purity and molecular weight of the CTPF component by MALDI-TOF analysis. The MALDI-TOF mass spectrum highlighted the presence of a single prominent peak at m/z 1503.042, which closely matches the theoretically predicted mass of 1498.73 Da (Figs. 2B and S8). There are no other additional peaks in the higher molecular weight range between 4 and 30 kDa, indicating minimal contamination from empty PhoCl1 barrel and intact PhoCl1 (Figs. S9 and S10). The SDS-PAGE analysis served as a complementary experiment, confirming the purity of the CTPF with the absence of higher molecular weight bands that could arise from the empty PhoCl1 barrel and intact PhoCl1 (Fig. 2C, Lane 4). The MALDI-TOF and 12% SDS-PAGE gel results collectively support that the photocleaved CTPF is responsible for generating the red fluorescence.

Fig. 2: Characterization of the purified C-terminal peptide fragment (CTPF).
Fig. 2: Characterization of the purified C-terminal peptide fragment (CTPF).
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A Visual images of purified PhoCl1 (Left) and purified CTPF (Right) acquired under a UV transilluminator (λ = 365 nm), B Digitalized MALDI-TOF mass spectrum of the purified CTPF showing a prominent peak at 1503.042 m/z, corresponding to the expected molecular weight (1498.73 Da) of the peptide (Fig. S8) C 12% SDS-PAGE gel electrophoresis of unexposed PhoCl1, light-exposed PhoCl1, and pure CTPF. The purified CTPF lane shows the absence of both PhoCl1 and the PhoCl1 barrel, indicating the purity of CTPF. D Absorption spectrum of the purified CTPF, indicating the presence of a broad peak centered around 470 nm and 320 nm. E Emission spectrum (red line) of the purified CTPF using excitation wavelength of 470 nm indicates the presence of a sharp emission peak centered around 570 nm, while excitation spectrum (blue line) of purified CTPF using fixed emission of 570 nm indicated excitation peak centered around 480 nm and 320 nm and F Fluorescence lifetime of CTPF using an excitation wavelength of 485 nm was determined to be 0.34 ± 0.04 ns. The data was fitted using a biexponential decay model.

We characterized the spectral properties of CTPF through UV-Visible and fluorescence spectroscopy. The absorption spectrum of CTPF displayed a maximum of 470 nm and an additional peak at 320 nm (Fig. 2D). We recorded the emission spectrum of CTPF using 470 nm excitation wavelength, which exhibited a prominent emission peak at 570 nm (Fig. 2E). We also recorded the corresponding excitation spectrum (Fixed emission at 570 nm), which displayed two peaks around 480 and 320 nm. (Fig. 2E). CTPF emergence with time was measured by immobilizing PhoCl1 to Ni-NTA beads and observing the increase in red fluorescence of the supernatant at 570 nm, which followed a first-order rate kinetics with a rate constant of ~0.25 h-1 (Fig. S11). To compare the fluorescence properties of CTPF, with respect to established GFP-related chromophores, we determined the fluorescence lifetime and quantum yield of CTPF. The fluorescence lifetime (τ) of CTPF determined using a biexponential decay model was 0.34 ± 0.04 ns, compared to PhoCl1 and Rhodamine B, which exhibited lifetimes of 2.2 ± 0.04 ns and 0.86 ns, respectively (Figs. 2F and S12A, B)10. The low lifetime of CTPF is likely due to the rotational freedom near imidazolinone group (which is likely restricted when the chromophore is inside PhoCl1), facilitating nonradiative decay pathways11. The quantum yield of the CTPF was determined to be 0.23 ± 0.08%. A low brightness of 25.85 M−1 cm−1 for CTPF (38,000 M-1 cm-1 of EGFP) closely aligned with its dim visual appearance under the UV-transilluminator (λ = 365 nm)12. All these spectroscopic findings for CTPF closely align with those of synthetic Kaede chromophore analog. This synthetic chromophore corresponds to the photoconverted red chromophore of Kaede protein and resembles IE-pHBI chromophore present on CTPF13. A comparative summary of the spectroscopic properties of Kaede protein and CTPF is provided in Table S3.

Finally, CTPF was lyophilized, and it retained its red fluorescence even after solubilization (Fig. S13A, B). The red emission from CTPF was monitored over 5 days, showing no significant change in fluorescence intensity over the observation period (5 days), indicating its stability. (Fig. S13C).

The emergence of red fluorescence in aqueous phase by CTPF bearing the chromophore IE-p-HBI is counter-intuitive as it should lose fluorescence through solvent-mediated non-radiative pathways once it is released from the hydrophobic environment of β-barrel. This observation led us to investigate the mechanisms responsible for the red fluorescence in CTPF. The major components of CTPF are the 9-amino-acid sequence (NRVFTKYPR) and an IE-p-HBI chromophore located at the N-terminus, composed of two five-membered aromatic rings and one six-membered ring (Fig. 1A). The IE-p-HBI chromophore likely requires exclusion of water from its immediate surroundings to maintain fluorescence, a characteristic that parallels its behavior within the hydrophobic core of the β-barrel14. We hypothesize that CTPF aggregates into clusters to minimize surrounding water molecules and achieve water exclusion15,16.

To validate our aggregation hypothesis, we conducted a comparative size analysis of both photoexposed and unexposed (control) PhoCl1 samples using dynamic light scattering (DLS). We propose that photocleavage of PhoCl1 at 405 nm would generate in situ CTPF, which, if unaggregated, would exhibit minimal changes in the observed size as compared to PhoCl1 (Barrel protein (Intact + Empty)). Conversely, aggregation would result in the presence of larger aggregates of CTPF in comparison to PhoCl1. The size analysis for the unexposed PhoCl1 at the onset of the experiment depicted an average hydrodynamic diameter of 7.2 ± 0.7 nm, which aligns with its expected dimensions of ~5 nm (Fig. 3A)17. Following light-induced photocleavage of PhoCl1 for 6 h, the mixture consisted of undissociated PhoCl1, empty PhoCl1 barrel, and CTPF. Along with the peak corresponding to undissociated PhoCl1 and empty PhoCl1 barrel of 5.6 ± 0.6 nm, an additional peak centered at 1224 ± 275 nm was also observed. The undissociated PhoCl1 and the empty barrel have comparable sizes, suggesting the appearance of the higher order species (>1000 nm) indicative of aggregation. In the unexposed PhoCl1 control, no size changes were detected up to 6 h, and it showed a dominant peak at 6.3 ± 0.1 nm corresponding to intact PhoCl1 and a low-intensity, broad peak between 100 and 1000 nm persisting throughout the experiment.

Fig. 3: Size analysis of in situ generated CTPF through photocleavage of PhoCl1 using dynamic light scattering (DLS).
Fig. 3: Size analysis of in situ generated CTPF through photocleavage of PhoCl1 using dynamic light scattering (DLS).
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A Size analysis of the unexposed and photoexposed PhoCl1 (6 h) samples to track in situ generation and aggregation of CTPF following photoexposure. The unexposed PhoCl1 showed a maximum and almost unaltered population (1–25 nm peak) throughout the experiment from 0 h (black solid line) to 6 h (green solid line). The photoexposed PhoCl1 displayed a decrease in the population of PhoCl1 (6–7 nm peak, red dashed line) following the photocleavage while the in situ generated CTPF displayed a size of 1224 ± 275 nm (red dashed line), indicative of aggregation. B Size analysis of pure CTPF is indicative of aggregates with a size range of 477 ± 79 nm.

The two main peaks (Barrel protein (Intact + Empty) < 25 nm) and aggregates (>100 nm) were further analyzed using ‘intensity ordered by area percent’, referred to as Iarea %. The data showed that the major population (83.4%) of the species present in the photocleaved PhoCl1 mixture corresponds to CTPF aggregates (1224 ± 275 nm). We further quantified the Iarea % over time, observing that light exposure led to an increase in Iarea% to ~80% within 6 h, whereas the unexposed control displayed a plateau with an Iarea% of ~25% (Fig. S14). Finally, we measured the hydrodynamic size of the purified CTPF, revealing a diameter of 477 ± 79 nm (Fig. 3B). These suggest the aggregation of CTPF when photo-released from PhoCl1. These molecules likely aggregate due to the hydrophobic nature of the chromophore while simultaneously minimizing the water content around the chromophore. Time-resolved anisotropy measurements revealed a rotational correlation time (θ) of CTPF (~0.1 ns) was lower than that of sfGFP (10.54 ns) (Fig. S15). This lower θ suggests greater rotational freedom of CTPF. In contrast, sfGFP, with its compact structure, displayed a higher θ, reflecting its rigid conformation. We also determined the rotational correlation time of ~0.1 ns for CTPF through the Perrin equation. The rotating hydrated unit size of CTPF was determined to be ~0.5 nm, aligning with established observation of small molecules18,19. We observed a r0 (fundamental anisotropy) value greater than 0.4, likely indicating light scattering contributions from the aggregates in addition to fluorescence20. Additionally, a non-zero residual anisotropy (r∞ ~ 0.1) in the case of CTPF also suggests potential restricted rotational motion of the CTPF molecules in the aggregate20,21. Together, the results suggest that CTPF aggregates with restricted chromophore rotation within the larger framework.

We adopted a molecular dynamics (MD) simulation approach to validate whether this aggregation of CTPF (IE-p-HBI-NRVFTKYPR) is involved in the emission of red fluorescence. We conducted two independent simulations of 600 ns consisting of 15 CTPF molecules, randomly dispersed in solution, with distinct initial velocities. We calculated the interaction energy among all chromophores in the system, which reflects the total energetic contribution from all sorts of non-covalent interactions between multiple chromophore entities within the system. The data indicated that as CTPF molecules aggregate, the total energy of the system decreases and stabilizes upon complete aggregation, observed around 400 ns of simulation (Fig. 4A). The aggregation pattern reveals that the chromophore undergoes hydrophobic clustering ~200 ns onwards, shielding itself from water and facilitating the formation of aggregates (Fig. 4B and Supplementary Video S1). The decomposition of the total energy into the Lennard-Jones and electrostatic components indicates that the aggregation is largely driven by Lennard-Jones interactions between the chromophores (Fig. S16). This is evident as only Lennard-Jones energy decreases and stabilizes as the aggregation proceeds. We suspected the involvement of π-π stacking interactions among the chromophores during aggregation. We calculated the dihedral angle between the tyrosine ring and five-membered imidazolinone ring present within the chromophore. We observed that the chromophores attained nearly planar conformations (dihedral angle 180°), one of the essential conditions for showing fluorescence (Fig. S17)22. Additionally, the ‘solvent-accessible surface area’ (SASA) of the chromophore decreases consistently throughout the simulation trajectory, starting from the dispersed state, as aggregation progresses. Once aggregation is complete, the SASA stabilizes and fluctuates around its minimum value (Fig. S18). This further suggests that aggregation is predominantly driven by hydrophobic interactions between the chromophores. A similar aggregation behavior was observed in the simulation using 30 randomly dispersed CTPF molecules, where the interaction energy of the system decreases and stabilizes ~100 ns onwards (Fig. S19A). The simulation trajectory showed that the decrease in the system’s energy is accompanied by the aggregation of the chromophore, leading to the exclusion of water molecules around the chromophore (Fig. S19B).

Fig. 4: Computational validation of the proposed aggregation of the CTPF.
Fig. 4: Computational validation of the proposed aggregation of the CTPF.
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A Time-dependent variation of total interaction energy (Coulomb and van der Waals) among all chromophores along the molecular dynamics trajectory. The energy scaled relative to initial separated state. B Snapshots of chromophore aggregation at different time points along the molecular dynamics simulation trajectory, illustrating the structural changes as interaction energy decreases. The chromophore is shown in red using a van der Waals representation, while the peptide is represented in purple C Variation in the hydration number or the number of water molecules (Nwater) around the chromophores in the 15 CTPF molecules. The x-axis represents the chromophores present in the 15 CTPF molecules used to simulate the aggregation. The chromophores highlighted with red colors were selected for the QM/MM calculations on the basis of variable water content. D Excitation and E Emission spectrum acquired through QM/MM of a select chromophore with hydration no. 8, indicated in red solid line, with an overlay of the experimental spectral scan (black solid line).

Furthermore, we calculated the “hydration number,” i.e., the number of water molecules surrounding the chromophore in the stable aggregated cluster. The simulations revealed fluctuations in the hydration number of the aggregated chromophore ranging from 4 to 14 which is less than the hydration number (~30) of the chromophore in isolated state (Fig. 4C). This decreased hydration number is critical, as water molecules are known to significantly influence the chromophore’s spectroscopic behavior, by modulating its electronic properties. Further, to get insight into the chromophore’s spectroscopic properties, we performed quantum mechanics/molecular mechanics (QM/MM) simulations on the most stable chromophore conformation obtained after the 400 ns trajectory. We selected five representative chromophores with varying water content (3, 5, 6, 8, and 11 water molecules) for QM/MM calculations. In each QM/MM simulation (i) chromophore and adjoining amino acid asparagine (Asn) were treated in the quantum mechanical (QM) region (ii) residues within 3 Å of the chromophore comprised of the active region, including water (iii) remaining atoms were modeled using molecular mechanics (MM). The excitation and emission profiles were performed using the B3LYP functional and cc-PVDZ basis set on this selected group of chromophores23,24. These calculations revealed distinct absorption peaks between 460 and 490 nm, which closely aligns with the experimentally observed absorption peak at 470 nm (Figs. 4D and S20A). For the emission profiles, we found emission maxima between 540 and 590 nm, which varied based on the presence of water molecules surrounding the chromophore (Fig. 4E and S20B). This hydration-dependent behavior indicates that the local environment is crucial in influencing the chromophore’s emissive property. The computationally predicted emission peaks closely align with the experimentally measured emission peaks centered around 570 nm, providing strong validation for our model. The close agreement between the computational and experimental spectral absorption and emission wavelengths also suggests that aggregation plays a role in the emergence of the red fluorescence of the CTPF.

We assessed how various physicochemical factors have an impact on the red emission of CTPF. The temperature-dependent fluorescence measurements indicate a decrease in the CTPF emission intensity with an increase in temperature from 25 °C to 65 °C (Fig. S21). We also examined the effect of salt concentration, specifically NaCl, on the fluorescence of CTPF. An increase in CTPF emission intensity was observed while increasing the NaCl concentration from 150 mM to 2 M (Fig. S22). We monitored the CTPF emission intensity with increasing guanidinium chloride, which indicated no significant increase in the emission intensity at the highest concentration of 6 M (Fig. S23). Finally, we observed the change in the fluorescence property of CTPF with different concentrations of surfactant CTAB (Fig. S24). An increase in CTPF emission intensity was observed with the progressive increase in CTPF concentration (Fig. S25). The effect of pH on the fluorescence intensity was measured relative to pH 8.0, at which most of the investigations of CTPF were performed. There is an almost negligible effect on the fluorescence of CTPF either at neutral condition (pH 7.0) or basic conditions from pH 9.0 to pH 11.0, while a decrease in the fluorescence intensity couple with a shift in wavelength to a shorter wavelength (broad peak between 530 and 570 nm) at acidic pH 5.0 (Fig. S26). This shift is likely due to chromophore protonation, a phenomenon observed in similar photoconvertible fluorescent proteins like DendFP and Dendra225.

In summary, our experimental and computational analysis validate the aggregation of CTPF, which in turn plays a crucial role in the emergence of red fluorescence following PhoCl1 photoexposure. These findings highlight the behavior of the photodissociated chromophore-containing peptide (Fig. 5)6. The observation that photocleavage of PhoCl1 results in red fluorescence due to aggregation of CTPF provides critical insights into the fate of photodissociated fragments in engineered fluorescent proteins. This is particularly important because many optogenetic tools rely on predictable post-cleavage behavior, often assuming the photofragments to be inert. Our findings reveal that the chromophore fragment can persist and undergo aggregation, leading to a new spectral output. While we do not propose CTPF as a direct tool for high-resolution bioimaging due to its aggregation-prone nature, the generation of red-emitting aggregates from a photocleaved product opens up avenues for designing triggered fluorescence turn-on systems. Future efforts may focus on (1) engineering the chromophore-containing peptide to control its aggregation, (2) leveraging this behavior for designing sensors based on aggregation-induced emission (AIE), and (3) using the aggregation of photocleaved peptides as a tunable parameter for material assembly and/or fluorescence tagging.

Fig. 5: PhoCl1 photoexposure: aggregation-induced red fluorescence.
Fig. 5: PhoCl1 photoexposure: aggregation-induced red fluorescence.
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A Current model depicting the loss of PhoCl1 fluorescence after 405 nm induced photocleavage event of PhoCl1 and B Expanded model indicating aggregation of CTPF units after 405 nm induced photocleavage event of PhoCl1. This aggregation results in the emission of red fluorescence. Illustration was created using the Inkscape open-source vector graphic editor.

Conclusion

PhoCl1, a photocleavable fluorescent protein, which upon 405 nm photoexposure splits into two fragments- an empty β-barrel and a 9-amino acid C-terminal peptide fragment containing chromophore. The β-barrel turns non-fluorescent when the chromophore along with C-terminal peptide fragment (CTPF), detaches from it. The detachment of CTPF was deemed to render the PhoCl1 non-fluorescent. However, to the best of our knowledge, we have provided evidence for the generation of red fluorescence from the detached CTPF. We demonstrate that the released chromophore, in the form of a C-terminal peptide fragment (CTPF), generates dim red fluorescence through aggregation. We further characterized the red fluorescence through various spectroscopic techniques and determined the excitation and emission maxima to be 470 nm and 570 nm, respectively. We determined the quantum yield of CTPF to be ~0.23% which was corroborated through the visual dim appearance upon excitation in the UV-transilluminator. Dynamic light scattering data showed that CTPF forms aggregates, which we propose plays a key role in red fluorescence by preventing water molecule interference. To gain deeper insights into the fluorescence generation mechanism, we conducted molecular dynamics simulations which support the experimental finding of aggregation. To further evaluate the generation of red fluorescence, we performed quantum mechanics/molecular mechanics (QM/MM) simulations, which corroborate the experimental observations of excitation and emission maxima.

Materials and methods

DNA purification kits and enzymes

Plasmid purification kits, PCR purification kits, and Ni-NTA agarose were purchased from Qiagen. The restriction enzymes BamHI, NdeI, and Dpn1 were acquired from New England Biolabs (NEB). PrimeSTAR GXL DNA polymerase was acquired from DSS Takara India Pvt. Ltd.

Chemicals and reagents

Tris base, 2-mercaptoethanol, kanamycin sulfate, nickel sulfate, sodium chloride, acetic acid, methanol, glycerol, and, bacterial growth media- Luria Bertani (LB) broth (Miller), LB agar (Miller) were purchased from Sisco Research Laboratory (SRL) Pvt. Ltd. Cytidine-5′-triphosphate (CTP) was procured from Cayman Chemical (USA). Phenylmethylsulfonyl fluoride (PMSF) and disodium dihydrogen ethylenediaminetetraacetate dihydrate (EDTA) were acquired from Tokyo Chemical Industry (TCI) Co. Ltd. Isopropyl-ß-D-1-thiogalactopyranoside (IPTG), low EEO agarose, and imidazole were obtained from HiMedia Laboratories.

Consumables

Ten-kilodalton MWCO Amicon® centrifugal filters were acquired from Merck. SnakeSkinTM dialysis tubing (10 kDa MWCO, 0.22 mm Internal Diameter) was acquired from Thermofisher Scientific. UV LEDs (400–405 nm, VAOL-5GUV0T4) were purchased from VCC Optoelectronics. Costar® 96-well UV-transparent microplates and Costar® fluorescence black microplates were purchased from Corning Incorporated.

Plasmids

pET-PhoCl1-6xHis was a gift from Robert Campbell (Addgene plasmid #164033; http://n2t.net/addgene:164033; RRID: Addgene_164033).

Cloning of 6xHis-PhoCl1

We cloned an N-terminal 6xHis- PhoCl1 into the pET28a(+) vector using restriction-free cloning, inserting it between the NdeI and BamHI sites26. To achieve this, we PCR amplified the PhoCl1 gene along with an N-terminal 6xHis tag sequence from a plasmid containing the pET-PhoCl1-6xHis plasmid (Addgene Plasmid #164033). We added overhangs complementary to NdeI and BamHI restriction sites of pET28a(+) vector in an additional round of PCR to facilitate the insertion of the N-terminal 6xHis-PhoCl1 gene into the pET28a(+) vector. We did the whole plasmid amplification in another round of PCR using pET28a(+) as a template and 6xHis-PhoCl1 amplicon as a megaprimer in 1:5 ratio. The parent plasmid in the PCR products was digested with Dpn1 enzyme and transformed into Escherichia coli DH5α chemical competent cells. The cloned plasmid vectors were purified, and the sequence was confirmed through Sanger sequencing (Barcode Biosciences, India).

Tables S1 and S2 list the nucleotide/amino acid sequences and primers used for the cloning in this study, respectively.

Overexpression of 6xHisPhoCl1 protein

The plasmid pET-6xHis-PhoCl1 was transformed into E. coli BL21(DE3) chemical competent cells. One percent of the overnight grown cultures of E. coli BL21(DE3) cells transformed with above-mentioned plasmids were used to inoculate LB media supplemented with 50 µg/mL of kanamycin sulfate. The resulting secondary culture was then incubated at 37 °C and 180 rpm in an incubator shaker. Once the optical density (O.D.) measured at 600 nm of the culture reached 0.6, induction was performed by adding 1 mM IPTG to trigger overexpression. Following induction, the temperature was reduced to 18 °C and maintained for 18 h. Subsequently, the cells were harvested by centrifugation at 6000 RPM at 4 °C for 10 min. The resulting cell pellets were flash-frozen and stored at −80 °C for further use.

Ni-NTA affinity purification of 6xHis-PhoCl1

The cells were resuspended in a lysis buffer consisting of 100 mM Tris-HCl, pH 8.0, 300 mM NaCl, 10 mM imidazole, 0.1 mM PMSF, and 0.025% 2-mercaptoethanol. Ultrasonication was utilized for cell disruption, using 1 s ON and 3 s OFF cycles, at 60% amplification for a total time of 20 min. After sonication, the lysed cells were centrifuged at 14,000 rpm for 30 min to separate the cell debris from the supernatant. The clarified lysate was filtered through a 0.22 µm syringe-driven filter and loaded onto a pre-equilibrated 2 mL Ni-NTA gravity column. The equilibration buffer comprised of 10 mM imidazole, 100 mM Tris-HCl, pH 8.0, 300 mM NaCl, 0.1 mM PMSF, and 0.025% 2-mercaptoethanol for all proteins.

The purity of eluted fractions containing the desired proteins was verified through 12% SDS-PAGE gel electrophoresis. The pure protein fractions were combined and dialyzed using SnakeSkinTM dialysis tubing against 50 mM Tris-HCl, pH 8.0, and 300 mM NaCl.

405 nm light source setup

We custom-assembled a 405 nm light source using UV LEDs (400–405 nm, VAOL-5GUV0T4) to expose the photocleavable protein PhoCl1 and its fusion constructs. The setup involved connecting the LED’s positive terminal to a 470 Ω resistor in series, with both terminals mounted on a breadboard and linked to a DC power supply (5 V). The schematic and full setup details are shown in Fig. S2. To facilitate photoexposure the breadboard-mounted LED was placed in an inverted position inside the microcentrifuge tube containing the PhoCl1.

Isolation of pure CTPF through 405 nm induced photocleavage of PhoCl1

The purified 6xHis-PhoCl1 was immobilized on pre-equilibrated Ni-NTA affinity resin till full saturation. The equilibration buffer was composed of 50 mM Tris-HCl, pH 8.0, and 300 mM NaCl. The resin was washed thoroughly to remove the unbound protein. The Ni-NTA resin immobilized with 6xHis-PhoCl1 was resuspended again in equilibration buffer and transferred to 2 mL microcentrifuge tubes (MCTs). Once the resins settled properly (~7.5 mm bead height), the excess buffer was removed, leaving behind ~350 µL of it. The MCTs were exposed to 405 nm light (1.5 mW cm-2) using our custom-built LED setup for 12 h (Fig. S2). Following photoexposure the supernatant containing the photo-released CTPF was collected. The isolated CTPF was lastly passed through a small amount of Ni-NTA resin in a gravity column to remove any fusion contamination if present.

The purified CTPF was analyzed for its molecular weight and purity using MALDI-TOF spectrometry. We used 12% SDS-PAGE gel electrophoresis to check for any higher molecular weight protein bands in the purified CTPF.

The purified PhoCl1 and CTPF concentration was measured using the Bicinchoninic Acid (BCA) assay.

Photodissociation yield (φdissociation) of PhoCl1 calculation

Using the power density (4 mW/cm2) of the 405 nm LED and the tube radius equal to 0.6 cm, we calculated the total energy delivered to PhoCl1 over a 2-h period (corresponding to the linear portion of the graph). Given the energy of a single photon, we determined the total number of photons emitted. To estimate the total number of PhoCl1 molecules dissociated in 2 h, we multiplied the initial number of PhoCl1 molecules by the photodissociation fraction, which was derived from the decrease in fluorescence observed over this period. We have assumed that all photons contribute exclusively to the photodissociation process. We calculated the photodissociation yield based on

$${\varphi }_{{dissociation}}=\,\frac{{Number}\,{of}\,{dissociated}\,{molecules}\,}{{Total}\,{number}\,{of}\,{photons}\,{absorbed}}$$

Matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF MS)

We photoexposed the Ni-NTA immobilized 6xHis-PhoCl1 equilibrated with 50 mM (NH4)2CO3 buffer to collect CTPF. Ammonium bicarbonate buffer was used to avoid the interference of tris buffer in MALDI analysis. The CTPF peptide and α-Cyano-4-hydroxycinnamic acid (CHCA) were mixed in 4:1 ratio, drop-casted on the MALDI target plate, and dried at room temperature. The MALDI-TOF spectra were recorded on the Bruker Autoflex MALDI-TOF instrument in the linear positive mode from m/z 1000 to 4000. We additionally recorded the spectrum in the range m/z 4000 to 30,000 to check for the presence of higher molecular weight protein contamination. The matrix used in this case was sinapinic acid.

UV-Visible absorption, fluorescence emission, and excitation spectra measurements

UV-Vis absorption and fluorescence emission spectra were recorded using a BioTek Cytation 5 multimode plate reader. For the absorption measurements, 150 µL each of 6x-His PhoCl1 (30 µM) and CTPF (30 µM) were placed in a 96-well UV-transparent microplate. Absorbance was measured across a wavelength range of 300 to 700 nm with a 5 nm step size.

For fluorescence emission spectra, 150 µL each of CTPF (30 µM) was loaded into a 96-well black polystyrene microplate. The measurements were done using 470 nm excitation and emission from 520-700 nm. The fluorescence spectrum of the control (30 µM 6xHis-PhoCl1) was recorded using 400 nm excitation and an emission range of 450-700 nm. Additionally, we measured the excitation spectra of CTPF at fixed emission wavelengths of 510 nm and 570 nm. For PhoCl1 dissociation measurement, PhoCl1 was exposed to 405 nm LED, and the 510 nm RFU upon 400 nm excitation was taken at different time points and plotted. The fit was done using the first-order rate kinetics model: [A] = [A]0e−kt. Here, [A] represents the PhoCl1 concentration, [A]0 is the initial PhoCl1 concentration, k is the rate constant, and t is time. Relative Fluorescence Units (RFU) have been used as a measure of concentration.

For CTPF emergence with time, the beads were immobilized with PhoCl1 and exposed to light (Power density- 4 mWcm-2), and supernatant RFU at 570 nm upon 470 nm excitation was taken at different timepoints and plotted. The fit was done using first-order rate kinetics model: [P] = [A]0(1-e-kt).

Here, [P] represents the CTPF concentration, [A]0 is the initial PhoCl1 concentration, k is the rate constant, and t is time. Relative Fluorescence Units (RFU) have been used as a measure of concentration.

Fluorescence lifetime measurements

The fluorescence lifetime in the time domain was determined using a home-built confocal microscope integrated with a lifetime measurement system (Nanoplasmonics lab, IIT Gandhinagar). The system was equipped with LDH-D-C-485 nm pulsed LASER source (PicoQuant GmbH) for the fluorophore excitation. The fluorescence decay was detected by a Single-Photon Avalanche Diode (SPAD) solid-state detector (Excelitas Technologies®) connected to a TimeHarp 260 board (Time-Correlated Single Photon Counting (TCSPC) and Multi-Channel Scaling (MCS) board. The instrument response function (IRF) is recorded via the synchronization (sync) signal from the laser driver module to the TimeHarp 260 photon counting card, based on the set sync rate. For fitting, we utilize SymPhoTime 64 software, which loads the IRF values directly from the .ptu file associated with each sample. The photoluminescence (PL) decay is fitted using the pre-set models (bi-exponential) in the SymPhoTime 64 software, iterating until convergence. CTPF and 6xHis-PhoCl1 (10 µL) were added on a glass slide and placed on the sample stage of the confocal microscope. The protein sample was excited with a 485 nm pulse laser operating with a repetition rate of 8 MHz while the fluorescence decay was captured. Before the experiment, the laser power (100 µW) was measured at the objective. The following formula has been used to get the average decay time:

$$\uptau =\frac{{\uptau }_{1}{A}_{1}+{\uptau }_{2}{A}_{2}}{{A}_{1}+{A}_{2}}$$

where τ1 is a fluorescence lifetime of the first decay component, τ2 is a fluorescence lifetime of the second decay component, A1 is amplitude of the first decay component and A2 is amplitude of the second decay component.

Quantum yield measurement of CTPF

UV-Vis absorption spectra were measured using a JASCO UV-Vis spectrophotometer (V-750), while fluorescence emission studies were carried out with a Horiba-Jobin-Yvon Fluorolog-3 spectrofluorometer. Fluorescence spectra were recorded in quartz cuvettes with a 1 mm path length and slit widths of 1–1.5 nm (same for both, excitation and emission), using a peptide concentration of ∼40 μM. The excitation wavelength for the emission spectra was set to the absorption maxima of the sample. Fluorescein (quantum yield = 0.95 in 0.1 M NaOH)27 was used as the reference for quantum yield (QY) determination, which was calculated using the following equation:

$${{{\rm{Q}}}}_{{{{\rm{S}}}}}={{{\rm{Q}}}}_{{{{\rm{R}}}}}\frac{{{{{\rm{I}}}}_{{{\rm{S}}}}}}{{{{\rm{I}}}}_{{{{\rm{R}}}}}}\frac{{{{\mathrm{OD}}}}_{{{{\rm{R}}}}}}{{{{\mathrm{OD}}}}_{{{{\rm{S}}}}}}\frac{{\upeta }_{{{{\rm{S}}}}}^{2}}{{\upeta }_{{{{\rm{R}}}}}^{2}}$$

Here, Q represents the quantum yield, I is the integrated intensity, OD is the optical density, and η is the refractive index. The subscript R refers to the reference fluorophore (fluorescein) with a known quantum yield, while S represents the sample fluorophore (CTPF).

Brightness determination

The molar extinction coefficient (ε) of CTPF is measured experimentally using Beer-Lambert’s law. The brightness was calculated using the following equation:

$${{{\mathrm{Brightness}}}}=\upepsilon * {{\mathrm{Quantum\; Yield}}}$$

Dynamic light scattering measurements

Particle size analysis of concentrated solution of PhoCl1 (1.5 mg/mL) and its photocleaved mixture was conducted using Malvern Zetasizer Dynamic Light Scattering (DLS) instrument. The PhoCl1 solution was exposed to 405 nm light for 6 h, and the DLS measurements were done at time points of 2, 4, and 6 h using 1 mL of sample. For each time point, the particle size of unexposed PhoCl1 was also measured as a control.

Fluorescence anisotropy measurements

These experiments were conducted at the IIT, Bombay Central Instrumentation Facility. The lyophilized powder of CTPF was sent via courier and resolubilized prior to the start of the experiment. Anisotropy measurements were performed using DeltaFlex TCSPC Lifetime Fluorimeter. The excitation and emission wavelengths were set to 470 nm and 570 nm, respectively. The fluorescence intensity was recorded at both parallel (\(I\)∥) and perpendicular (\(I\)⊥) orientations to the plane of polarized light used for excitation. This was used to calculate the change in anisotropy (r) vs time (t). This decay curve was fit with r(t) = r0e(−t/θ), where θ is the rotational correlation coefficient. As a control, a similar experiment was performed with sfGFP with an excitation and emission wavelength of 488 nm and 510 nm.

The rotational correlation time obtained from the time-resolved TCSPC data was further correlated with that obtained from the Perrin equation using the fluorescence lifetime of CTPF.

$${r}_{{ss}}=\frac{{r}_{0}}{1+\frac{\tau }{\theta }}$$

Here rss is the steady state anisotropy, which is calculated from the following equation using time-dependent intensity I(t) decay.

$${r}_{{ss}}=\frac{{\int }_{0}^{{\infty }}r\left(t\right).I\left(t\right){dt}}{{\int }_{0}^{{\infty }}I\left(t\right){dt}}$$

Here \(I\left(t\right)=I{||}+2{{{\rm{I}}}}\)⊥

Molecular dynamics simulations

Preparation of the CTPF peptide-chromophore conjugate system. The initial conformation of the peptide part of CTPF (NVRFTKYPR) was generated using the tLEaP module of AmberTools28. The conformation of the chromophoric part IE-p-HBI was derived from the crystal structure of PhoCl1 (PDB: 7DNA)7, and conjugated N-terminal to the NVRFTKYPR peptide in PDB format.

To obtain charge on the chromophore, we utilized the restricted electrostatic potential (RESP) method, we employed the RED module with the HF/6-311 G* basis set29. For charge calculations, the terminal end of IE-p-HBI was capped with N-methyl group to mimic the peptide-bound state. The total charge of p-HBI was set to -1, while the N-methyl charge was set to 0. Subsequently, ‘General AMBER Force Field’ (GAFF) was used to generate force field parameters of the chromophore IE-p-HBI using the calculated RESP charges30,31. The initial conformation and force field of the conjugated system (IE-p-HBI and peptide) were generated using XLeap (AmberTools)28, combining GAFF (p-HBI) and Amberff14sb force fields for peptide32.

The AMBER-formatted force field and coordinates were converted to GROMACS format using a Python script33. All simulations were performed using GROMACS-2020.234.

Simulation details

CTPF was placed in a cubic box of 80 Å dimension. To eliminate any steric contact from the system, an energy minimization in vacuum was performed using the steepest descent method for 10,000 steps with the tolerance force 0.001 kJ mol-1. The system was solvated using the TIP3P water model followed by the addition of 150 mM NaCl ions to neutralize the system35,36. A second energy minimization was performed using the steepest descent method for 5000 steps with a tolerance force of 0.001 kJ mol−1. Thereafter, the system was heated to 27 °C using the Berendsen thermostat for 10 ns, during which the heavy atoms of the solute were position restrained37. To achieve a perfectly equilibrated system, annealing was performed for 100 ns. The annealing steps involved heating the system to 50 °C in 3 steps and then cooling it back to 27 °C in 3 steps. Subsequently, a 100 ns constant temperature and pressure simulation was performed at 27 °C and 1 atm pressure using a V-scale thermostat and Berendsen barostat, respectively, followed by a 600 ns constant volume and temperature simulation at 300 K using Nose-Hoover thermostat37,38,39,40. During the simulation, the bonds were constrained using the LINC algorithm41. The electrostatic effects were treated with the PME method using the 10 Å distance cut-off for long-range interaction.

To enhance statistical significance, two independent simulations (production run) of 600 ns were performed using a different initial structure and velocities. Further, to investigate chromophore’s aggregation, total interaction energy calculations between the chromophores were performed for all independent trajectories over 600 ns. The energy fluctuations converged after 400 ns, indicating complete aggregation. The resulting aggregated structures were then analyzed using QM/MM simulations to determine the chromophore’s spectroscopic characteristics.

QM/MM calculation

The spectroscopic properties of the chromophore were determined through QM/MM calculations on chromophore aggregates. The final aggregated conformation was selected based on the most stable energies obtained from molecular dynamics (MD) simulations. Subsequently, water molecules within a 3.5 Å radius of the chromophore were calculated to assess the hydration shell. Based on the hydration number, we have selected the four chromophores for the QM/MM calculation.

In each QM/MM simulation, (i) chromophore and adjoining amino acid asparagine (Asn) were treated in the quantum mechanical (QM) region, (ii) residues within 3 Å of the chromophore comprised of the active region, including water, (iii) remaining atoms were modeled using molecular mechanics (MM). The QM region is treated using B3LYP functional and cc-PVDZ basis set, while the charges and van der Waals parameters of MM region are taken from the Amber force field. The electrostatic at the QM//MM boundary is treated using the Electrostatic Embedding model42,43.

Initially, the aggregated conformation was optimized at the ground-state geometry level using the aforementioned parameters, followed by the calculation of the Hessian matrix to determine the vibrational levels of the system in its ground state. Subsequently, vertical excitation calculations were performed using Time-Dependent Density Functional Theory (TD-DFT) to identify the maximum excitation energy44. Thereafter, the excited-state geometry was optimized, and Hessian matrix at excited geometry was calculated to generate the excited-state vibrational energy levels. Vertical emission calculations were conducted to investigate fluorescence properties. All the QM/MM calculations are performed using the ORCA-5.0.0 package45.

Statistics and reproducibility

All experiments were carried out independently three times unless otherwise mentioned. Data analyses and plotting were carried out using XMGrace, Microsoft Excel, and Origin plotting suit. Data were represented as mean ± standard deviation. We also performed Student’s t-test (two-tailed) wherever applicable (significant if p < 0.05). All the MD simulation data were analyzed by using the various GROMACS modules and home-written code. The structural analysis of the protein data bank (PDB) files was done using the visualization software “UCSF Chimera” and Visual Molecular Dynamics (VMD). Figures were prepared and finalized using Microsoft PowerPoint and Inkscape open-source vector graphic editor.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.