Single-engineered-residue solvation perturbations regulate global protein architecture and function

Domain: biology
Published: 2026-03-10
Editorial outcome: major_revision

Abstract
Protein-water interactions fundamentally shape the structure, stability, dynamics, and functionality of proteins. However, the heterogeneous nature of the protein-water interface and the disparity in their dynamic interplay make it challenging to understand how local water perturbations influence protein structural dynamics over space and time. In this study, we introduce a photochromic molecule, spiropyran, to modify a specific residue of proteins, thereby achieving a reversible, residue-specific, and amplified perturbation on the hydrophobicity of protein surfaces. With the aid of controlled, amplified hydrophobic perturbations, we reveal that even residue-level changes in hydrophobicity induce significant global alterations in protein hydration patterns. These hydration shifts propagate in an amino acid sequence-dependent manner, initiating dramatic influences overall protein architecture and catalytic performance. Our findings establish that interfacial water networks not only capture the surface physicochemical patterns of proteins but also mediate the propagation of local perturbations into broader structural and functional fluctuations. By shifting the paradigm from “structure-function” to “structure-hydration-function”, our work provides innovative perspectives into understanding protein architecture and guiding future drug design strategies.

Introduction
Water serves as an active matrix within biological systems, providing a dynamic hydrogen bond network (HBN) that supports protein hydration1-3. This hydration water shell bridges the gap between the protein surface and bulk water4-6, therefore, it is closely implicated in protein folding and plays a critical role in their functions through molecular-level water-protein interactions7-9. For instance, protein hydration water can act as a lubricant, aiding the rearrangement of hydrogen bonds during protein conformational change, which facilitates the essential flexibility for protein folding10. Moreover, the cooperative effects of hydrophobic solvation and protein-protein interactions constitute key thermodynamic drivers for the formation of liquid-liquid phase separation and subsequent fibrils3. Consequently, fine-tuning hydration water presents significant potential for advancements in protein structural engineering and drug design.

The influence of local water perturbations on global protein structures is not yet well understood, partly due to the heterogeneous distribution of charge and hydrophobicity on protein surfaces11-13. Polar protein residues stabilize hydrogen-bond-ordered networks, while hydrophobic patches template stratified hydration layers with restricted rotational diffusion but enhanced lateral mobility14-16. These spatially segregated nanoscale patches, with distinct hydrogen bond lifetimes, interfacial viscosities, water dynamics, and proton-exchange rates, complicate the task of understanding how water networks span and reshape protein dynamics.

The issue of whether hydrophobic perturbations at individual residues can trigger long-range restructuring of the water layer and regulate the global protein structure remains a topic of contention. NMR and neutron scattering experiments17, as well as simulation studies, collectively suggest that localized mutations primarily affect the first-shell water (3-5 Å). These perturbations may potentially extend to 2-3 adjacent layers (10-15 Å) via HBNs. Nevertheless, recent evidence from femtosecond time-resolved fluorescence Stokes shift and anisotropy measurements indicated that water perturbations stemming from isolated mutations can exert global effects on solvation, which drives coupled protein side-chain motion5. Based on these findings, water perturbations from local residues are hypothesized to induce global conformational reconfiguration, but have not yet been fully verified. Such discrepancies may arise from small variations in hydrophobicity among amino acids (Kyte-Doolittle scale, -4.5 to 4.5)18, which obfuscate changes in solvation and present challenges for analysis.

In this study, we address the challenges of hydrophobicity changes at the single-residue level by engineering a specific protein residue with a photochromic molecule, spiropyran (Fig. 1a)19. The light-triggered transition between merocyanine (MC) and spiropyran (SP) isoforms results in a marked polarity shift that enhances the hydrophobicity disturbances at the modified residue. Both molecular dynamics (MD) simulations and terahertz (THz) spectroscopy results suggest that alterations in single-residue hydrophobicity not only disrupt local water configurations but also initiate a cascading reorganization of the entire hydration shell. This restructuring of the hydration shell progresses along amino acids in a sequence-dependent manner, triggering a cascading effect that influences global protein architecture and catalytic activities. Our research introduces a predictive “structure-hydration-function” framework that delineates the interplay between protein structure and hydration dynamics, which together determine functional outcomes. These insights lay the groundwork for the strategic design of protein engineering methodologies, presenting an avenue for fine-tuning biomolecular functionality.

Results
Residue-specific Engineering Alkaline Phosphatase (ALP) with Photochromic Molecule
Alkaline phosphatase (ALP, EC: 3.1.3.1) was chosen as a model protein to investigate the impact of local hydrophobicity due to its photostability under 450 nm illumination (Supplementary Figure 1-2). Loop regions, which are crucial for the structural integrity and flexibility of proteins20,21, are significantly affected by their interactions with water. Using the crystal structure of ALP (PDB 1ED9 [http://doi.org/10.2210/pdb1ED9/pdb]) as a guide22, specific residues in the loop region, located at varying distances from the catalytic site, were replaced with the unnatural amino acid p-azido-L-phenylalanine (pAzF). The selected sites were H372/872 (~5 Å from the catalytic site), V99/599 (~10 Å), E406/906 (~15 Å), and K185/685 (~25 Å) (Supplementary Figure 3-5)23. After comprehensive evaluations of catalytic activity and photo-stability, the E406/906M mutants (ALP-E15) were chosen for further studies (Supplementary Figure 6-7). To facilitate light-induced reversible and amplified perturbations in hydrophobicity, a dibenzocyclooctyne (DBCO)-functionalized SP (DBCO-SP) molecule (Supplementary Figure 8-10) with reversible photoisomerization (Supplementary Figure 11) and remarkable less hydrophobic-to-more hydrophobic transformation (Fig. 1b) was conjugated to the ALP-E15 through a click reaction (Fig. 1c). The successful conjugation of DBCO-SP to ALP was verified by mass spectra (Fig. 1d) and UV/vis spectra (Fig. 1e). Further quantitative analysis of the solvent-accessible surface area (SASA) indicated that the modified MC/SP accounts for ~4% of the total accessible protein surface (Supplementary Figure 12).

We then investigated the photoisomerization-induced changes of microenvironments on the engineered protein surface by measuring the fluorescence lifetime of MC. As a sensitive reporter for polarity (Supplementary Figure 13), MC exhibits site-dependent fluorescence lifetimes upon conjugating to different sites of ALP, indicating variations in local polarity microenvironment (Supplementary Figure 14-15). The fluorescence lifetimes of ALP-MC and its revertant state (RMC, regenerated by photoisomerization from SP to MC) were recorded by time-correlated single photon counting (TCSPC)24 (Fig. 1f). Statistical differences in fluorescence lifetime were observed between ALP-MC and ALP-RMC (Fig. 1g). In contrast, negligible variations occurred between free MC and RMC (Fig. 1g, Supplementary Figure 16), indicating that MC-SP isomerization specifically triggers rearrangements of the hydration shell that possibly leads to changes of protein conformation25.

Rearrangement of Hydrogen Bond Network and Global Protein Hydration
With the capability to enhance the hydrophobicity of a specific protein residue, we probed the photoisomerization-induced rearrangement of HBN within the hydration shell. We conducted MD simulations to investigate the perturbation of local hydrophobicity on the protein-water interface. The simulations were performed on a fully hydrated ALP based on a known structure (PDB ID: 1ED9 [http://doi.org/10.2210/pdb1ED9/pdb]) with MC and SP molecules modified at the mutation sites26.

We first evaluated the radial distribution function (RDF, (g(r)) of water between water and the modified residue, catalytic site, or the global ALP surface, in response to photoisomerization using the trajectories from the stable phase of each group (Figs. 2a-c and Supplementary Figure 17). At the modified residue, the hydrophilic MC shows a prominent g(r) peak in the 0-4 Å range (first hydration shell), presumably due to a denser, direct HBs between water and the polar groups of the MC probe (Fig. 2d and Supplementary Figure 18). In contrast, SP expels the first shell of water, creating a sub-nanoscale hydrophobic cavity. This leads to a gradual rise of the g(r) peak in the 4-8 Å range. Furthermore, the more mobile outer hydration layer shows higher sensitivity to distal perturbations, resulting in a concerted amplification of HBN reorganization27. Consequently, after photoisomerization to SP, the RDF of water around the distant catalytic site (15 Å from the modified site) also exhibits elevated g(r) peaks in the 3-8 Å range. While within 0-4 Å near the catalytic site, the g(r) peak is diminished in the ALP-SP group, which can be ascribed to a disturbance in dipole orientation coupled with a reduction in dielectric shielding28. For the global protein, the RDF across the entire ALP surface reveals only a minor decrease in the 3-8 Å range, suggesting that the averaging effect masks local depletion.

To define the spatial range of residue hydrophobicity effects on global water dynamics, we computed lifetime (τ HBA) of water-water (w-w) hydrogen bonds within successive hydration layers (1st: < 4 Å, 2nd: 4-6 Å, 3rd: 6-10 Å) around the modified residue29,30, distal catalytic site, and global ALP surface, respectively (Figs. 2e-g, Supplementary Figure 19). In the 1st and 2nd hydration shells, ALP-SP presented longer τ than ALP-MC in all these groups. This might be caused by the expansion of hydrophobic SP interfaces via ordered hydration cages that stabilize τ . In contrast, the competing hydrogen bonding between MC and water destabilizes the w-w HB, thereby shortening its lifetime (Fig. 2d). This effect decays over successive hydration layers in a distance-dependent way until reaching bulk water values after three layers. Besides, the most remarkable elongation in HB lifetime was also observed at the catalytic site for both SP and MC groups, which may be ascribed to the disordered dipole alignment in its electrically organized water network31. Therefore, we concluded that changes in the hydrophobicity of single residues can laterally propagate across the protein surface to remote sites, causing global hydration changes (Fig. 2h).

After confirming the stable w-w HB in the 1st hydration layer and the increased mobility of outer hydration layers upon photoisomerization to ALP-SP, we then calculated the water density fluctuations (WDFs) across protein surfaces to investigate the effect on distal residues. WDFs were defined as the probability of finding N water molecules in a 3.5 Å spherical probe volume centered on groups of each hydrophilic/hydrophobic residue (P(%))32. Accordingly, its function with the normalized water number fluctuation [<N2>–<N>2]/<N> (F ) acts as an indicator of water exchange dynamics at the residue-water interface. As illustrated in Fig. 2i and Supplementary Figure 20, both ALP-MC and ALP-SP have a bimodal F distribution, corresponding to hydrophilic (small F ) and hydrophobic (large F ) populations. Notably, in ALP-SP, both peaks shift toward small F values with higher P(%), indicating that enhanced local hydrophobicity leads to suppressed global water mobility. Paradoxically, this decreased global water mobility is likely due to the reorganization of water mediated by protein surface topology.

We then quantified the number of hydrogen-bonded (H-bonded) waters (N H-bonded water ) per solvent-accessible residue to explore the residue-specific physicochemical properties on the propagation induced by local perturbations. The difference value in H-bonded waters (ΔN H-bonded water ) for each residue, calculated by subtracting ALP-MC from ALP-SP (ALP-SP – ALP-MC), was then mapped onto the corresponding residues to provide a direct perspective on global hydration alterations (Fig. 2j). Highly perturbed patches (increased N red) and lowly perturbed patches (decreased N blue) discretely coexist across the protein surface, indicating that the hydration changes are heterogeneous and site-specific.

We next divided amino acids into four groups according to their physicochemical properties, including negatively charged (Asp, Glu), positively charged (Lys, Arg, His), polar uncharged residues (Ser, Thr, Asn, Gln, Cys, Tyr), and nonpolar residues (Ala, Val, Leu, Ile, Met, Pro, Phe, Trp, Gly). Then the ΔN per residue category (calculated as ALP-SP – ALP-MC) was subjected to normal distribution analysis (Fig. 2k-n, Supplementary Figure 21-22). Here, the mean ΔN (μ) represents the average net change per residue, whereas the standard deviation (σ) quantifies the degree of spatial heterogeneity in hydration dynamics in responses to photoisomerization. Notably, all types of residues, especially positively charged residues, showed positive changes in N (μ>0), indicating a global net increase in N upon photoisomerization to ALP-SP. The increase implies that changes in local hydrophobicity disrupt long-range water networks, freeing water molecules to enhance local entropy and drive global hydration restructuring. Moreover, negatively charged groups exhibited the greatest ΔN variability and the broadest frequency distribution (σ = 0.989), indicating the highest sensitivity to local perturbations. Positively charged and polar groups showed similar medium fluctuations and distribution widths. Conversely, nonpolar groups demonstrated the least fluctuations and the narrowest distribution with the highest frequency in ΔN = -0.5 to 0.5. This suggests that the propagation pathways of hydrophobicity perturbation are influenced by the electrostatic/polarity characteristics of different amino acids via HBN tuning33, in the sequence of negatively charged > positively charged ≈ polar (neutral) > nonpolar34. We concluded that the local photoisomerization-induced HBN disruption at the modified residue triggers a long-range reorganization of water across the protein surface, with propagation pathways modulated by the surface chemical microenvironment.

We next employed in situ Attenuated Total Reflection Terahertz (ATR-THz) spectroscopy to probe dynamic variations in protein hydration. The THz regime is exquisitely sensitive to perturbations in collective HBNs35, enabling direct detection of HB stretching (ν , 150-200 cm-¹) and librational modes (ν , 450-600 cm-¹) of hydration water36,37. Measurements were performed on protein film samples prepared at precisely controlled hydration levels38. Protein films were formed by drying solutions onto an IR-transparent diamond under gentle vacuum, followed by application of Tris-NaCl buffer to achieve target hydration levels (see SI for details). Significantly, stable intensity at 1550 cm-¹ (ΔR < 0.02; water-content internal standard) confirms constant water content after photoisomerization (Eq. S1-S2, Supplementary Figure 23).

To minimize bulk water interference and elucidate the reorganization of hydration water, THz spectra were measured across distinct hydration levels: low hydration state (~ 0.2 g H₂O/g protein), corresponding to a discontinuous sub-monolayer hydration regime on the protein surface; moderate hydration state (~ 0.7 g H₂O/g protein), which fully saturates the first hydration shell and partially populates the second shell; high hydration state (~ 1.2 g H₂O/g protein), approaching completion of the first three hydration shells9,39 (Fig. 3a-c and Supplementary figure 24-25, Supplementary Table S1). The single-beam ATR spectra of ALP-MC and ALP-SP at different hydration levels were converted to absorbance spectra by penetration depth correction40 (Fig. 3d). We then examined how hydration level governs the THz frequency shifts triggered by the ALP-MC to ALP-SP photoisomerization (Fig. 3e, f). In the fully hydrated group, the HB stretching band (ν ) was centered at ~ 130 cm-¹ for both ALP-MC and ALP-SP. In comparison, the librational band (ν ) shifted from 539 cm-¹ (ALP-MC) to 545 cm-¹ (ALP-SP). This small change is likely attributed to the buffered local disturbances by the HBN of bulk water. However, at reduced hydration level (~ 0.7 and 0.2 g/g), both modes exhibit larger blue-shifts after photoisomerization to ALP-SP with ν shifted from 586 to 612 cm-¹ (~ 0.7 g/g) and 584 to 614 cm-¹ (~ 0.2 g/g), while ν shifted from 170 to 195 cm-¹ (~ 0.7 g/g) and 165 to 196 cm-¹ (~ 0.2 g/g). These changes indicate the formation of a cooperatively rigidified HBN in the protein hydration layer that enhances the orientational order of water molecules41,42. Consistently, ATR-THz experiments with reversible ALP-RMC photo-switching reveal that the hydration layer is unable to fully revert to its initial state (Supplementary Figure 26). This indicates that a localized perturbation can induce a global and irreversible transition in protein hydration. Moreover, control experiments using pure water and native ALP ruled out possible contributions from light-induced heating effect on structural changes in the solvent or protein (Supplementary Figure 27-28). Consequently, the THz results demonstrated that a single-residue hydrophobicity change can induce HBN reorganization across global hydration. Moreover, this conversion is amplified under reduced hydration levels, hinting that the hydration layer may act as a critical regulator of biological interfaces.

To delineate the thermodynamic basis of light-induced hydration rearrangements between ALP-MC and ALP-SP, we used THz calorimetry to correlate spectral changes with entropy (ΔS) and enthalpy (ΔH). Absorption coefficients (α(ν)) and photoisomerization-induced differential spectra (Δα(ν) , ALP-SP minus ALP-MC) were calculated to isolate hydration-specific alterations (Eqs. 1-3, Supplementary Figure 29). The Δα(ν) was deconvolved into the hydrophobic “wrap” region (150-200 cm-¹, entropy-linked) and hydrophilic “bound” region (450-600 cm-¹, enthalpy-linked) (Supplementary Figure 30).

Spectral analysis of Δα(ν) spectra at different hydration levels revealed a negative Δα amplitude (Δα < 0), signifying a positive entropy change (ΔS > 0, Eq. 4 and S3, Supplementary Table S2) due to the release of loosely wrap-water from hydrophobic patches (Fig. 3g). Concurrently, a positive slope in the bound region (Δα /Δν > 0) indicated a negative enthalpy change (ΔH < 0, Eq. 5), reflecting enhanced rigid H-bond networks with hydrophilic residues. Consequently, the THz results demonstrated that a single-residue hydrophobicity change can induce HBN reorganization across global hydration by converting water from a loosely associated “wrap-water” state to a tightly bound, ordered configuration, thereby forming a rigid hydration layer. Furthermore, the observed entropy-enthalpy compensation minimizes the system's free energy, yielding a net favorable hydration contribution (ΔGhydration < 0, Eq. 6) that acts as the thermodynamic driver for conformational destabilization.

We then calculated THz spectra of global and residue-local hydration shells via auto-correlation functions of the time derivative of dipole based on MD simulations to distinguish the origin of THz peak shifts43, whether they arise from global hydration layer reorganization or localized hydrophobic changes by MC/SP photoisomerization. The raw MD-calculated spectra of bulk water without smoothing or signal extraction are provided to maintain statistical precision (Supplementary Figure 31). The employed qSPC/Fw water model exhibits a known force-field limitation that precludes the capture of the 200 cm-¹ peak44,45. Nonetheless, it qualitatively reproduces the librational band above ~300 cm-¹, showing agreement with the experimental spectrum of ambient water (Supplementary Figure 32). The known systematic blueshift does not impact the relative trends under discussion.

This allows for comparison of librational mode changes in hydration water after photoisomerization to ALP-SP. The THz spectral peak shifts are analyzed separately for the hydration layer around the modified-MC site and the global hydration layer of protein but excluded MC-modified sites. After photoisomerization from MC to SP, there is a blue shift of librational modes in both local hydration at the modification site (Fig. 3h) and the global protein hydration layer but excluded MC-modified sites (Fig. 3i). This confirms that the experimentally observed THz peak shifts originate primarily from hydration shell restructuring, rather than from site-specific vibrational amplification.

Global Protein Structural and Functional Changes Induced by the Rearrangement of HBN
We next probed the role of HBN rearrangement in mediating distal protein structural changes. After ensuring the systems were stable (Fig. 4a, Supplementary Figure 33)46, we scrutinized the differences in structural fluctuation of ALP-SP and ALP-MC through the root mean square fluctuation (RMSF) of protein backbone atoms. As illustrated in Fig. 4b, the difference in RMSF between ALP-MC and ALP-SP (ΔRMSF) indicates that ALP undergoes residue-specific conformational fluctuations. To depict the spatial distribution of these fluctuations, we mapped ΔRMSF values onto their corresponding residues. Notably, compared to ALP-MC, ALP-SP demonstrated enhanced fluctuations in regions containing SP modifications and in adjacent loop regions (Fig. 4c). The superimposition of simulated structures of ALP-MC and ALP-SP further validated these distinct conformational fluctuations and structural alternations (Fig. 4d). Specifically, we observed a rigidification in the secondary structure at residues 902-922 (SP modification site) and 823-835 (catalytic active site). Analysis of secondary structure evolution over the final 20 ns confirmed these differences persist (Supplementary Figure 34-36). The distance matrix (DM) analysis revealed an increased intramolecular distance in ALP-SP (Fig. 4e, f), suggesting reduced cooperativity among residues, which may lead to less compact global structures. Importantly, control simulations of the ALP-N mutant confirmed that the overall protein fold remains unperturbed by covalent modification alone, whereas the light-induced hydrophobic switch selectively enhances local flexibility and remodels the hydration shell (Supplementary Figure 37-38).

To further evaluate the role of specific MC/SP-surface interactions, we conducted MM/GBSA energy decomposition. The results show that interactions near modified MC/SP are mainly electrostatic, with negligible van der Waals contributions. Solvent water strongly screens these electrostatic effects beyond ~5 Å. Despite differing charge distributions, both isoforms exhibit highly similar interaction profiles with surface residues (Supplementary Figure 39). Therefore, the observed changes in flexibility and stability are more likely due to HBN reorganization instead of direct effect of the modification.

To corroborate the MD simulations, small-angle X-ray scattering (SAXS) experiments were conducted to investigate the structural dynamics47,48. A close alignment between the MD-calculated and experimental scattering profiles (χ2 <1.7) substantiates the accuracy of the simulations (Fig. 4g). Guinier analysis revealed an increase in Rg (36.93→38.63 Å; Fig. 4h) and Dmax (109→122 Å; Fig. 4i) from ALP-MC to ALP-SP (P < 0.01), indicating enhanced structural flexibility. The broadening of the Kratky plot peak was also observed in ALP-SP (Fig. 4j), a characteristic of global structural loosening that is not fully reversible upon relaxation back to the MC state (Supplementary Figure 40-41, Supplementary Table S3). Control experiments with unmodified ALP showed negligible changes in SAXS after photoisomerization (Supplementary Figure 42). Upon photoisomerization to ALP-SP, the thermal denaturation melting temperature also declined by 5 ℃, providing further evidence for the less structured state (Supplementary Figure 43). Collectively, the MD simulations, SAXS and melting temperature data suggest that light-induced hydrophobic perturbations at single residues induce not only local structural rigidity but also propagate to more distal regions, thereby augmenting global protein conformational flexibility.

Having confirmed that the hydrophobic changes of single residues can alter global protein structural dynamics via collective reorganization of HBN, we next investigated the enzymatic changes by interrogating hydrophilic (pNPP) and hydrophobic (4-MUP) substrates (Fig. 5a). For pNPP, after photoisomerization from ALP-MC to ALP-SP, Michaelis constant (Km) increased ca. 50% and catalytic constant (kcat) dropped ca. 37% (Fig. 5b). The reduced substrate affinity and catalytic activity suggest that HBN reorganization enhances interfacial water ordering that impedes the delivery of hydrophilic pNPP via solvation-coupled transport49. We further employed deuterated reagents in the pNPP assay to verify the influence of hydration on enzymatic activities. As illustrated in Fig. 5c, the reaction rates of both ALP-MC and ALP-SP were markedly suppressed in D₂O, attributable to the stronger deuterium bonding and more ordered HBN structure in D₂O. These require higher enthalpic energy to be disrupted, thereby hindering the access of pNPP to the catalytic pocket50. Moreover, the amplified rate difference between ALP-MC and ALP-SP in D₂O further confirmed that the regulation of enzyme function primarily stems from the hydration-layer HBN reorganization, rather than local electric field-induced water orientation. This is supported by the comparable dielectric constants of H₂O and D₂O, which yield consistent electric field effects.

To exclude nonspecific effects from the free photo isomers, we performed a control experiment in which unmodified ALP was mixed with SP. Illumination of this mixture resulted in negligible changes in enzymatic activity. This is in stark contrast to the significant effects observed with covalently conjugated ALP-SP, confirming that functional modulation arises from local perturbations of the hydration shell enabled by site-specific attachment rather than direct interactions with free photoisomers (Supplementary Figure 44).

Furthermore, to exclude potential interference from light-induced hydrophobic aggregation on enzymatic changes, OTG-supplemented experiments were designed. OTG suppressed the possible hydrophobic aggregation but did not reverse activity modulations, confirming that hydration effects primarily regulate enzymatic function (Supplementary Figure 45).

In contrast, the hydrolysis of hydrophobic substrate, 4-MUP, was less influenced by photoisomerization. The Km and kcat values remained unchanged in both H₂O and D₂O buffer (Fig. 5d-e). This is attributed to the hydrophobic nature of 4-MUP, which allows direct binding to the dehydrated active site, thereby bypassing solvent-mediated hydration processes. Collectively, these findings confirm that the reorganization of HBN can regulate catalytic efficiency through a structure-hydration-function coupling mechanism51.

To establish physiological generalizability, we introduced single-point mutations at a conserved surface residue (ALP E406/905) using amino acids with varying hydrophobicity (I > F > G > Q > K) (Supplementary Figure 46a). A clear correlation emerged between residue hydrophobicity and activity toward the hydrophilic substrate pNPP (activity: I < F < G < Q), while activity toward the hydrophobic substrate 4-MUP remained largely unchanged (Supplementary Figure 46b-c). The Lys (K) variant, likely modulating hydration via charge effects, was an exception. These results, alongside those from the SP/MC system, confirm that water-mediated allostery constitutes a general and tunable mechanism of protein regulation.

Methods
Mutant Alkaline Phosphatase (ALP) Expression and Purification. Alkaline phosphatase (ALP; EC 3.1.3.1) was selected as the model protein. The phoA gene (NCBI: NC_000913.3), encoding a C-terminal 6×His tag, was synthesized (GenScript) and cloned into pET28a (Novagen), yielding pET28a-phoA. To site-specifically incorporate p-azido-L-phenylalanine (pAzF) at defined distances from the catalytic center, amber (TAG) mutations were introduced at residues His872/372 (5 Å), Val599/99 (10 Å), Glu906/406 (15 Å), and Lys685/185 (25 Å) using site-directed mutagenesis kit (Mut Express® II Fast Mutagenesis Kit V2; Novartis). Plasmid and primer sequences are provided in Supplementary Data 1. Each TAG-mutant pET28a-phoA plasmid was co-transformed with pEVOL-pAzF (Addgene, plasmid no. 31186) into Escherichia coli BL21(DE3) (Novagen). Cells were cultured in LB medium with 50 µg/mL kanamycin (Sigma-Aldrich) and 34 µg/mL chloramphenicol (Sigma-Aldrich) at 37 °C. At OD ~0.4, expression was induced with 1 mM IPTG (Sigma-Aldrich), 1 mM pAzF, and 0.2% (w/v) L-arabinose (Sigma-Aldrich), followed by incubation at 25 °C for 16 h. ALP variants were purified sequentially by nickel-affinity chromatography (HisTrap HP column; Cytiva) and anion-exchange chromatography (Q Sepharose™ column, Cytiva). Purity and integrity were assessed by SDS-PAGE, BCA assay, and LC-MS (Thermo Fisher Scientific).

Synthesis of dibenzocyclooctyne (DBCO)-functionalized SP (DBCO-SP). The IM2 (1'-(2-hydroxyethyl)-3’, 3’-dimethylspiro [chromene-2, 2’-indoline] -6-sulfonate) was prepared according to previous literature52. DBCO-SP was synthesized via DCC/DMAP-mediated esterification of IM2 with DBCO-C6-acid. A 50 mL round-bottom flask containing DBCO-C6-acid (309.8 mg, 929.2 μmol, 1.20 equiv), DCC (319.5 mg, 1.55 mmol, 2.00 equiv), and DMAP (18.9 mg, 154.9 μmol, 0.20 equiv) in anhydrous DCM (20 mL) was purged with N₂. The mixture was stirred at 25 °C for 18 h to activate the carboxylic acid, then IM2 (300 mg, 774.3 μmol, 1.00 equiv) was added. After stirring at 25 °C for 16 h, the crude product was purified by preparative HPLC (C18 column, isocratic elution with 50:50 v/v CH₃CN/0.1% aq. NH₄OH) to afford DBCO-SP as a brown solid (175 mg, 32.2% yield). Structural confirmation was performed by ¹H NMR, (Bruker Avance III 400 MHz), and liquid chromatography-mass spectrometry (LC-MS; Thermo Fisher Scientific). Photoisomerization characterization was carried out using a UV-visible spectrophotometer (Tecan) as described in Supplementary Information.

Preparation of SP-modified ALP. Azide-functionalized mutant alkaline phosphatase (ALP-N , 20 μM) in 25 mM Tris-NaCl (pH 7.5) was reacted with DBCO-SP (10 mM in DMSO) at a 10:1 molar ratio (DBCO-SP: ALP-N ). The mixture was incubated at 25°C for 2 h or 4°C overnight with gentle agitation. Reaction products were purified using a 7 kDa MWCO spin desalting column (1,200 ×g, 2 min) to remove unreacted DBCO-SP and byproducts, followed by concentration via 30 kDa centrifugal filtration (6,000 ×g, 10 min). Mutant ALP and ALP-SP conjugates (1 mg/mL in 25 mM Tris-NaCl, pH 7.5) were buffer-exchanged and purified using a Superdex™ 200 Increase 10/300 GL column (Cytiva) equilibrated with 25 mM Tris-HCl, 100 mM NaCl (pH 7.5). The separation was performed at 25 °C with a flow rate of 0.5 mL/min, and fractions corresponding to monomeric conjugates were collected. Mass analysis was performed on a Q-Exactive Plus mass spectrometer (Thermo Fisher Scientific) coupled to an Acquity UPLC system (Waters). Deconvoluted masses were determined using BioPharma Finder 5.3 to confirm the conjugation.

Fluorescence lifetime measurement. Measurements were performed at 20°C using a MicroTime 200 system (PicoQuant) with 532-nm pulsed excitation (40 MHz; LDH-P-FA-530B laser). Fluorescence was collected through a 100×/1.45 NA oil immersion objective, filtered via 488/532 nm dichroic mirror and 582/64 nm bandpass filter (Chroma), and detected by single-photon avalanche diode (Excelitas SPCM-AQRH-14-TR). Mutant ALP variants (V10, E15, K25) were diluted to 100 pM in TAE buffer with oxygen scavenging system (2.5 mM protocatechuic acid, 25 nM protocatechuate-3,4-dioxygenase). Samples were deposited on Ni2+-functionalized coverslips for 10 min53,54. All measurements were performed in six independent experimental replicates, each prepared from separately diluted samples and measured on independently prepared coverslips. For photoconversion studies, the MC state was obtained following 1 h of dark adaptation, and the RMC state was generated via 450 nm illumination (10 min) followed by 2 h of subsequent dark adaptation. FLIM images were acquired by galvanometric scanning. Individual molecules were manually selected for lifetime extraction. Fluorescence decays were fitted to multi-exponential models using SymPhoTime 64 software.

Hydrated protein film preparation and THz spectroscopy measurements. Hydrated protein films with hydration levels ranging from 0.02 to 0.4 g H2O/g protein were prepared using a humidity-controlled method (Methods details are available in Supplementary Information). Terahertz (THz) spectra were recorded using a Fourier-transform infrared (FTIR) spectrometer (Vertex 80v; Bruker, Billerica, MA, USA) equipped with a deuterated L-alanine triglycine sulfate (DLaTGS) detector and a single-reflection attenuated total reflection (ATR) unit (diamond crystal, 45° incidence angle) in the sample compartment. The spectral range was set to 50-690 cm-1 with a resolution of 2 cm-1. Both the sample compartment and interferometer compartment were evacuated to a pressure of 2-3 mbar prior to measurement to minimize water vapor interference. Each spectrum was acquired as an average of 64 scans. All THz measurements were performed in three independent experimental replicates, each prepared from independently hydrated protein films and measured separately. The diamond ATR crystal was sequentially cleaned with ultrapure water (prepared using a Milli-Q Integral water purification system; MilliporeSigma), 0.5 M NaOH solution (Sigma-Aldrich), ethanol (Sigma-Aldrich) and dried under a stream of nitrogen gas (99.999% purity; Air Liquide) after each measurement.

THz Spectra and Data Analysis. The ATR absorption coefficient (α(ν)) was calculated using the Lambert-Beer law:
α(ν) = - 1/dp ln(I(ν)/I0(ν)) (1)
dp = λ / (2π√n2sample sin(θ)2-n2diamond) (2)
where I(ν) and I0(ν) denote the sample's and reference's frequency-dependent intensities. The theoretical upper limit for the penetration depth (dp) indicates how far the evanescent wave can penetrate the sample. The refractive index (ndiamond) of diamond is 2.38, while that of liquid samples (nsample) is set to 1.5. For simplicity, the refractive index of the sample was assumed to be the average of water at this frequency range. λ is the wavelength of the radiation, θ is the incident angle (here θ=4/π).

Difference spectra Δα(ν) were determined by subtracting the ALP-MC spectrum from ALP-SP spectrum, as shown in eq.3:
Δα(ν) = α(ν)ALP-SP - α(ν)ALP-MC (3)

Connecting Thermodynamics to Spectroscopy. Within the THz-calorimetry framework, the hydration entropy (ΔS) and enthalpy (ΔH) changes upon photoisomerization are deconvolved into hydrophobic (cavity-wrap) and hydrophilic (bound) contributions via linear correlations with their respective spectroscopic fingerprints.The correlations ΔS ∝ Δα and ΔH ∝ (Δα /Δν) were established using pre-calibrated conversion factors:
ΔShydration = ∆Scavity + ∆Sbound ≈ ΔSwrap = Δαwrap ΔS̅wrap (4)
ΔS̅wrap = -4.4 J∙mol-1K-1cm
ΔHhydration = ∆Hcavity + ∆Hbound ≈ ΔHbound = Δαbound /ΔνΔH̅bound (5)
ΔH̅bound = -320 kJ∙mol-1cm
∆Ghydration = ∆Gcavity + ∆Gbound ≈ ΔH - TΔS (6)
where Δαwrap and Δαbound /Δν are the spectroscopic amplitudes in the 150-200 cm-¹ regions and the slope in 450-600 cm-1 regions, respectively. ΔS̅wrap and ΔH̅bound are constant correlation factors that reported based on a large set of solutes.

SAXS Characterization. SAXS experiments were performed on the BL19U2 beamline of Shanghai Synchrotron Radiation Facility (SSRF). Using an X-ray wavelength (λ) of 1.033 Å, scattered intensities were recorded with a Pilatus 1M detector (DECTRIS Ltd). The sample-to-detector distance was set to 2.6 m, yielding a momentum transfer range (q = 4πsinθ/λ, where 2θ is the scattering angle) of 0.01-0.4 Å-1, corresponding to a spatial resolution of 10-600 Å. Samples (ALP-dark, ALP-vis, ALP-MC, and ALP-SP) were prepared as 1 mg/mL solutions (100 μL) and maintained at 20°C. Each sample was measured in three independent replicates using separately prepared aliquots across multiple beamtime sessions to ensure statistical robustness. For each sample and buffer, 20 frames were recorded to optimize signal-to-noise ratio. Two-dimensional scattering patterns were azimuthally averaged, solid-angle corrected, and normalized to transmitted beam intensity using BioXTAS RAW 1.2.1. Primary SAXS data analysis was performed with the ATSAS 3.0.3 suite, including Guinier analysis, pair distance distribution (P(r)) calculations, and Kratky plot assessment. Theoretical scattering profiles were generated from crystal structures (ALP-dark, ALP-vis) and MD-simulated structural ensembles (ALP-MC/ALP-SP). For MD-derived models, representative conformations were selected by structural clustering of trajectories, followed by energy minimization of each cluster centroid. Theoretical SAXS curves were computed and fitted against experimental data, with agreement quantified by the goodness-of-fit (χ²). Detailed acquisition and analysis parameters are summarized in Supplementary Table 4.

MD Model building. The initial model was derived from the RCSB PDB database with the PDB ID of 1ED9 [http://doi.org/10.2210/pdb1ED9/pdb]22. Referring to the research of Hou et al. on the catalytic reaction mechanism of ALP phosphorylase26, the Ser102 residue was modified to a deprotonated state. Meanwhile, the protein was protonated under the condition of pH = 7.5. The covalent modifications at the MC and SP mutation sites were accomplished by the CovCom program. The protein system was characterized by the FF14SB force field55. Regarding all non-standard residues, such as those with covalent modifications and the deprotonated SER, the Restrained Electrostatic Potential (RESP) charges were calculated via Gaussian56. Subsequently, the parameters were fitted using the GAFF2 force field57. For the Zn2+ and Mg2+ ions located at the active site, the LJ 12-6-4 non-bonding force field parameters, which were meticulously fine-tuned by Li et al.58,59, were adopted.

The solvation of the MC and SP models was carried out using the LEaP module. A periodic cubic box was constructed, and it was ensured that the distance between the box edges and the protein atoms was greater than 15 Å. The system was then solvated using the flexible fixed-charge qSPC/Fw water model45. Meanwhile, Na+ and Cl- ions were added to the box to ensure the electrical neutrality of the system. To ensure statistical robustness, four independent molecular dynamics simulations were performed, each initiated from a distinct initial configuration. Details of the initial simulation setup are provided in Supplementary Table S5.

MD simulation details. For both the MC and SP models, four-step minimization was performed using PMEMD60 to eliminate atomic clashes. In the first minimization step, the protein and the metal ions at the active site were fixed, while the water molecules and counter ions were optimized. In the second minimization step, the protein was fixed, and the water molecules, counter ions, and the metal ions at the active site were optimized. For the third and fourth minimization steps, no restrictions were imposed. Each of the four-step minimizations consisted of 2000 steps of conjugate-gradient optimization, followed by 6000 steps of steepest-descent optimization. Subsequently, the PMEMD.cuda was employed to perform system heating and density relaxation. During the heating stage, a Langevin thermostat was utilized within the NVT ensemble. With a timestep of 1 fs/step, the temperature was gradually increased to 300 K over a period of 100 ps. The density relaxation is also carried out four times, using the NPT ensemble. The time step is set to 1 fs/step, and each relaxation lasts for 50 ps to allow the system density to relax to 1.0 g/cm³. Finally, PMEMD.cuda was used to perform a 500 ns MD (molecular dynamics) simulation for each system under the NVT ensemble, with a timestep of 2 fs/step. Subsequently, for the conformation of each system in the dynamically stable stage, a 50 ps MD simulation with a timestep of 0.5 fs/step was carried out to calculate the hydrogen bond lifetime of the water outside the protein.

CovCom. CovCom mainly relies on BioPython61 and RDKit62. First, it uses BioPython to extract the residue atoms within 25 angstroms around the modification site and convert them into a format recognizable by RDKit. Next, RDKit was employed to splice the modified residues with the modified molecule, and a simple geometric optimization of the modified part is carried out based on the MMFF94 force field63. Finally, the atomic coordinates of the modified part are refined by simulated annealing to optimize the binding pose of the new molecule within the cavity formed by surrounding residue atoms within 25 Å. The program code is available at https://github.com/iawnix/CovCom.

MD simulation analysis. The RMSD (Root Mean Square Deviation) of the protein backbone (Cα, C, N, and O) is used to evaluate the kinetic stability of the two models. The dynamically stable stage (the last 20 ns) was selected for the analysis of RMSF (Root Mean Square Fluctuation), RDF (radial distribution function), and secondary structure. The analysis of RMSF and secondary structure was completed based on CPPTRAJ60. According to formula 7, the radial distribution functions of ALP phosphorylase (non-hydrogen atoms), mutation sites (non-hydrogen atoms), and the ions in active sites (Zn2+, Mg2+) with WAT@O in the system are calculated respectively. This process is completed based on MDAnalysis46.
gab(r) = ρab(r) / (Nb/V) (7)

Hydrogen bond analysis. The last 20 ns of the stable phase of the two systems is selected to count the number of hydrogen bonds between the mutation sites and water. The analysis of the hydrogen bond lifetime between water molecules in different water shell layers is derived from the kinetic sampling of the stable conformation with a time step of 0.5 fs for 50 ps. Following formula 8, the hydrogen bond lifetime (τ) is obtained through single-exponential fitting29, and this process is completed based on MD analysis.
C(t) = 〈h(0)∙h(t)〉 / 〈h〉 (8)

Infrared absorption spectroscopy analysis. For a system in thermodynamic equilibrium, according to the linear response theory64, the spectrum of the autocorrelation function of the total dipole moment derivative, μ̇(t), is proportional to the product of the infrared spectral coefficient and the refractive index44:
α(w)n(w) ∝ ∫dt eiwt 〈μ̇(0)μ̇(t)〉 (9)
μ̇(t) = ∑q i v⃗i (10)
Based on Equation (9-10), we calculated the infrared absorption spectra of the 9 Å hydration shell around the mutation site and the 9 Å hydration shell around the protein with the mutation site excluded for the two systems, respectively, to characterize the hydrogen bond network strength of hydration layers in different regions. The trajectories were derived from 50 ps molecular dynamics sampling with a 0.5 fs time step, performed on stable conformations.

The Stability of ALP enzymatic activity against blue light illumination. The enzymatic activity of alkaline phosphatase (ALP) against light illumination was determined using p-nitrophenyl phosphate (pNPP) as a substrate. The hydrolysis products of p-nitrophenol (pNP) were continuously monitored at 405 nm (ε = 18,300 M-¹cm-¹) on a Tecan plate reader. The reaction mixture consisted 50 nM ALP and 1 mM pNPP in assay buffer (25 mM Tris, 100 mM NaCl, pH 7.5). Before reaction, ALP was subjected to blue light illumination (450 nm) for different time intervals. The enzymatic activity of wild-type ALP and site-specific mutants were evaluated under identical conditions. Each measurement was performed in three independent experimental replicates, using separately prepared ALP samples.

Michaelis-Menten Kinetics Analysis. The Michaelis-Menten kinetics of MC-modified ALP were assessed under dark conditions and blue-light illumination using pNPP as a substrate. The reactions were initiated by incubating 50 nM enzyme in a reaction buffer (25 mM Tris, 100 mM NaCl, pH 7.5) with various concentrations pNPP (from 0 to 600 µM). The reaction rates were determined from the linear accumulation of pNP absorbance at 405 nm. The hydrolysis of 4-Methylumbelliferyl phosphate (4-MUP) at 25°C were monitored fluorometrically (λex = 360 nm, λem = 450 nm). The reactions were initiated by incubating 5 nM ALP-MC with various concentrations of 4-MUP (from 0-500 µM). The reaction rates were determined by examining the linear accumulation of 4-methylumbelliferone (4-MU) over a period of 30 minutes, at 1 min intervals. Fluorescence intensities were then converted into 4-MU concentrations through the standard curve. All kinetic measurements were performed in three independent replicates, each initiated from separately prepared reaction mixtures. kcat and KM values were obtained by fitting the Michaelis-Menten equation.

Quantification and statistical analysis. Data are presented as mean  standard deviation (SD). Sample sizes (n), representing independent experiments, are indicated in the figure legends. Statistical analyses were performed using GraphPad Prism 9.5.1. Comparisons across multiple groups were assessed using one-way or two-way analysis of variance (ANOVA), as appropriate. Exact P values are provided in the figures. A P value < 0.05 was considered statistically significant.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In this manuscript by Liu et al, the authors use molecular dynamics simulations and terahertz spectroscopy to investigate how changes between two isoforms of a photochromic molecule that is attached to a single protein residue influence global protein hydration, as well as catalytic activity. The authors specifically investigate changes in hydration water structure associated with merocyanine and spiropyran, which are more hydrophilic and hydrophobic isoforms, respectively, to determine how varying local chemical properties near two residues can influence global water hydration. The authors claim that these changes manifest as long-range structural reconfigurations that impact catalytic activity. This conclusion is certainly interesting, although I do not believe that the results in their current form support this broad conclusion. My primary concern is with the limited statistical analysis of the results as described in my first comment below. I list my specific comments below.\n(1) No analysis of uncertainty or error is provided for any of the simulation results in the manuscript, which coupled with the small variations in simulation observables makes it difficult to determine if the conclusions based on the simulations are robust. For example, differences between the RDFs in Fig. 2a-c are extremely small, and without error bars in the hydrogen bond lifetimes in Figs. 2e-g it is not possible to tell if there are meaningful trends. I recommend that the authors perform replicate simulations to support uncertainty quantification, analyze and report uncertainty for all reported simulation quantities in Figs. 2, 3, and 4, and then discuss uncertainty when reporting their results. This comment is particularly important to address with respect to the data presented as RDFs, hydrogen bond lifetimes, and protein RMSDs, all of which appear quite similar to each other and hence need rigorous validation of potential differences to support the conclusions in the text.\n(2) Similar issues with large uncertainty are present for some experimental results. The authors should comment on the statistical significance of the data in Fig. 1g, where ALP-MC and RMC appear within error despite being describe as different in the text. Likewise, are the Rg values obtained from the data in Fig. 4h statistically significant?\n(3) The MD simulations focus entirely on the role of water, but the MC and SP isoforms have substantially different charge distributions and potentially different conformations. Do MC/SP themselves interact with surface-exposed amino acids differently? Such interactions might impact protein conformation without impacting water structure.\n(4) Some issues of clarity:\n- Pg. 4 – the distances listed after each mutation presumably refers to the distance from the catalytic cite, but this isn’t clear in the main text.\n- Fig. 1a – it is unclear what the numerical values (-17.7 D, -4.3 D) in the figure refer to.\n- Fig. 1b – both measured contact angles are <90 degrees and would correspond to hydrophilic surfaces. The caption should clarify that SP is more hydrophobic than MC, not that the surface itself is hydrophobic.\n- The authors refer to hydration layers (<4 A, etc). How are these distances computed – that is, what reference atoms are used when calculating this threshold distance, and what defines the distal catalytic site vs. modified residue? For the latter, are the atoms in MC/SP included? What happens if water molecules diffuse outside of this range within the 50 ps of sampling, are they still included?", "reviewer_2": "In this work by Liu et al. the authors engineer a photochromic molecule (merocyanine) to the peptide alkaline phosphatase (ALP) with the goal to create a light-induced reversible perturbation of the hydrophobicity of the protein surface. The authors use state-of-the-art methods to engineer and isolate the modified protein. The methods to modify and isolate the modified protein are scientifically sound. The authors convincingly show the conformational change of the merocyanine/spiropyran (MC/SP) while attached to the protein by fluorescence lifetime measurements. The authors then use experimental techniques, including Far-IR/THz ATR FTIR spectroscopy alongside molecular dynamics simulations to investigate the effect of the hydrophobicity change on the protein hydration.\nHere are my comments; I suggest the authors make revisions prior to publication.\n- On p. 5 the authors state that the modification occupies only 2.2 ‰ residues. It would perhaps help to state how much of the surface is altered, not the percentage of amino acid in total.\n- On p. 5 and Fig. 6 it is shown that the ALP-MC and ALP-RMC show significant deviations in the fluorescence lifetime data. The authors claim this suggests that the isomerization alters the local microenvironment on the protein surface. Why is that the most likely option? Would it not be possible (if not more likely) that the protein to transition into a new configuration? In the cited study [25] Ma et al. do not discuss MC as a probe for polarity shifts, viscosity etc. I find this section too vague and suggest an overhaul.\n- The authors use Far-IR (THz) ATR measurements of the hydrated sample to probe the spectroscopic signatures in the two conformations of the surface-attached photochromic molecule. On p. 11 the authors describe the “deconvolution method” to describe their ATR spectra. They use a sum of two damped harmonic osciallators. I would suggest adding the equations to the supplementary. The results of the fits are shown in Fig S19. The authors then directly attribute the amplitudes (denoted as “intensities” in the text) to a bound and wrap water fraction. A justification for this fit procedure is missing. This is crucial as this is the fundament for further discussion of the data. Studies by the Havenith group found that these spectroscopic regions are correlated for a “bound” and “wrap” water fraction. However, this correlation was not found by an intensity comparison of a damped harmonic oscillator model, but they demonstrated changes in the slope in the 400 and 600 cm-1 region after subtraction of corresponding bulk water contribution, which is not performed here. I suggest to carefully check if this procedure can be applied here as well.\n- In Fig. 3e) how were the error bars calculated? Are the errors calculated from the fit? Were the ATR measurements repeated/reproduced? I would expect that water content varies significantly between samples and thus could potentially contribute to a systematic error. Another important factor here is temperature. It is well known that the librational band of water has a strong temperature dependence. With increasing temperature, the librational band red shifts and decreases in amplitude. Following the analysis of the authors, this would also lead to a similar change in the Ibound/ Iwrap (which measures the intensity of the bands at ~150 and ~550 cm-1 ). Considering that it is not implausible to assume that some heating is induced during the sample illumination, this could potentially lead to these amplitude changes as well. Furthermore, it is reasonable that due to e.g. evaporation of water the overall water content could be altered. The authors need to convincingly clarify how (if) the temperature was controlled. In Fig. 1e) the authors demonstrate that the change from MC to SP is (at least to some degree) reversible. It would be reassuring to see the reversibility also here for the Far-IR ATR experiments.\n- On p. 12 the authors state that the MD-calculated spectra cannot reproduce the low-frequency modes of water below 300 cm-1 due to force-field limitations. It would be insightful to also have a clearer picture on the HB-stretch region, but I understand that the cost for such simulation would be large. It is, however, necessary to clarify the limitations of the current MD model regarding the librational band. Surely it was previously discussed in the literature to what extent the force field has an influence on the accuracy of the librational band?\n- On p. 15 the authors present SAXS data. For the ALP-SP upon photoirradiation the data shows a broadening of the Kratky plot peak (structural loosening) and a reduction of the thermal denaturation melting temperature by 5 °C. The authors state that this suggests an augmentation of the global protein conformational flexibility via collective reorganization of the hydrogen bond network. Further they find changes in the enzymatic activity. It is unclear to me how the finding of a change in flexibility or enzymatic activity can only be explained by a change in the HBN? How can the authors exclude a direct influence of the added SP/MC on the protein? The data clearly shows that the photoisomerization is not fully reversible if attached to the protein. Does this not show that the added SP/MC clearly directly (and perhaps even irreversibly) reacts with the protein itself?\nAs a last note, I do not agree with the authors on using the term “butterfly effect”. In simple words it refers to the phenomenon that in a dynamic system small changes of initial conditions can result in large differences in a later state. It is connected to a dynamic system and describes the unpredictability of small changes over time. A change of hydrophobicity might lead to long-range differences but this does not describe the evolution in time.", "reviewer_3": "The manuscript by Liu and co-workers deals with an important issue in molecular biophysics, i.e. the impact changes in hydration water has on macromolecular structure, dynamics and function. The authors address this subject by examining a surface modified version of the protein alkaline phosphatase (ALP) by a combined computational and experimental approach, notably by MD simulations and THz spectroscopy, respectively. The surface modification consisted in attaching two copies of the photochromic molecule spiropyran (SP; hydrophilic) to two surface amino acid residues that, upon illumination with blue light, undergo reversible isomerization to the more hydrophobic molecule merocyanine (MC). The authors show that changes in hydrophobicity induced by switching from one to the other molecule has effects on hydration-water hydrogen-bonding connectivity and dynamics, even farther from the site of surface modification. Furthermore, evidence is provided for the modification of protein structure and activity upon switching.\nThe reviewer has no direct expertise in MD simulations and THz spectroscopy and cannot judge if they are well executed or not. The results obtained on SP and on MC differ markedly, strongly suggesting there is indeed an effect of changes in hydrophobicity of the attached tags on water and protein structure and dynamics. The reviewer’s main concern is the suggested relevance of the results obtained with two copies of the very large tag attached to an unmodified protein in a functional and natural context. Therefore, in the reviewer’s view, the conclusion conveyed in the title (and given in the abstract) does not seem to be justified: the reader might underestimate the level of perturbation by the tags and mistakenly believe the level of ‘solvent perturbation’ is merely ‘enhanced’, as the authors write. How can the authors exclude that the solvent perturbation by switching of the tag isn’t merely an enhancement but of a different nature, and therefore not physiologically relevant ?\nThe reviewer has the following detailed comments in numerical order:\n1) Page (p) 2, line (l) 8: ‘… we reveal that even subtle, residue level changes in hydrophobicity …’. See comment above. The changes by these two large tags do not appear to be subtle and should not be mistaken by the reader to be comparable to those at play when a single residue is exchanged against another.\n2) P 2, l 12: ‘Our findings establish that interfacial water networks not only capture the surface physicochemical patterns of proteins, …’. The generalization of the author’s finding using large tags does not seem to justify a generalization to proteins.\n3) P3, beginning of second paragraph: it is not clear exactly to which statement in the previous paragraph the ‘however’ refers to.\n4) P3, beginning of third paragraph: Nickels et al (reference 17) did use neutron spectroscopy rather than ‘neutron scattering simulations’.\n5) P3, 3rd paragraph, l 7: Please specify to what experiments ‘femtosecond fluorescence’ refers to.\n6) P4, 2nd paragraph: The ‘butterfly’ effect is mentioned several times in the manuscript, but the analogy not explained.\n7) P4, 2nd paragraph: ‘structure-hydration-function’ appeared as ‘structure-function-hydration’ earlier in the manuscript.\n8) P4, second-to-last line: It should be explained why methionine mutations are mentioned here. Also, the length in brackets should be defined here already. Later in the manuscript, it becomes clear that the distance to the active site is mentioned.\n9) P5, second line: it should be rationalized why two and not only one tag is attached. Also, what is the extension of each tag and at what distance to the protein surface is the isomerizing group located ?\n10) P5, second paragraph: from which group(s) does the measured fluorescence originate, from the tags ?\n11) P5, second paragraph, description of fig. 1g: The fluorescence of ALP-MC and ALP-RMC are shown to differ. How does from this observation follow the conclusion that ‘isomerization specifically alters the local microenvironment’ ? Aren’t ALP-MC and ALP-RMC in the same isomeric state ?\n12) P7, 2nd paragraph, l 5: ‘… direct H-bonding with MC …’. It is not clear which group/molecule is H-bonding with MC. Water ?\n13) P7, 2nd paragraph, l 11: ’15 Å from the modified site’; How is this site defined; the point of attachment at the protein surface ? Or the very end of the tag ?\n14) P7, 3rd paragraph, l 4: it is unclear what is hydrogen bonding with what. Something seems to be missing in the sentence, the ‘modified residue’ does not connect.\n15) P9, third-to-last lien: what is meant with ‘initial disruption ?\n16) P11, l 1: Please define ATR-THz\n17) P11, second paragraph: How is fully hydrated (100 %) defined ? To connect with the literature, it would be useful to convert the 50% and 10% hydration samples into g water / g protein and indicate to how many hydration layers each value corresponds. In the end of p 11, the definition is vague: ‘(50% and 10%), containing 1-2 hydration layers waters.’\n18) P12, 1st paragraph: what is meant by ‘converting wrap- and bound-water’ ?\n19) P13: what does the hydration level of 10% make it ‘critical’ ?\n20) P14: The RMSD of backbone atoms in ALP-SP and ALP-MC are computed and compared. How do they compare to those of unmodified ALP-E15 ? Are there any changes in RMSD that originate from the surface modification and not from the change in hydrophobicity upon isomerization ?\n21) P14, fourth-to-last-line: Control experiments with unmodified ALP-E15 are described. Why do the authors refer to photoisomerization for this sample ?\n22) P18, last line: ‘Our findings reveal a hydration heterogeneity in governing protein dynamics, …’ What does this mean ?"}, "author_response": "Reply to Reviewers’ Comments\nResponse to Reviewer: #1\nPlease accept our heartfelt thanks for your thorough consideration and constructive\nsuggestions. We have conducted additional control experiments and refined the\nlanguage.\nPoint-by-point response to your comments is as follows.\n(1) No analysis of uncertainty or error is provided for any of the simulation results in\nthe manuscript, which coupled with the small variations in simulation observables\nmakes it difficult to determine if the conclusions based on the simulations are robust.\nFor example, differences between the RDFs in Fig. 2a-c are extremely small, and\nwithout error bars in the hydrogen bond lifetimes in Figs. 2e-g it is not possible to tell\nif there are meaningful trends. I recommend that the authors perform replicate\nsimulations to support uncertainty quantification, analyze and report uncertainty for\nwhen reporting their results. This comment is particularly important to address with\nrespect to the data presented as RDFs, hydrogen bond lifetimes, and protein RMSDs,\nall of which appear quite similar to each other and hence need rigorous validation of\npotential differences to support the conclusions in the text.\nReply: We thank the reviewer for figuring out the critical need for uncertainty\nquantification.\nIn response, we performed three additional independent MD simulations for each\nsystem (ALP-MC and ALP-SP) (Figs. S30 and R1). All data in the revised Figs. 2 and\n4f, including RDF, hydrogen-bond lifetimes, and protein RMSDs and RMSFs, involved\nerror bars and statistical analysis.\nIn Fig.2a-c, we conducted RDF analysis from four replicate MD simulations. The\nresults indicated that even after data averaging, RDF curves for ALP-MC and ALP-SP\nshowed similar significant differences. To quantify this difference, further statistical\nanalysis was performed on the characteristic peaks. The findings demonstrated\nsignificant statistical differences in the RDF values at modification sites, catalytic\nactivity centers, and global protein hydration layer (Fig. S16).\nFor hydrogen bond lifetimes in Fig. 2e-g, we performed additional uncertainty\n1\nanalysis that revealed statistically significant differences between the two isoforms.\nIn\nparticular, changes were observed for first and second hydration layers at the\nmodification site (p < 0.05), as well as more pronounced differences at the catalytic site\n(p < 0.001) and across global protein’s first hydration shell (p < 0.01). Moreover, water\ndensity fluctuations and number of H-bonded water per residue from four replicate\nsimulations are consistent, conforming the reliability of our conclusions (Figs. S19 and\nS21). These updates are incorporated into the revised manuscript.\nFig. S30 Backbone RMSD evolution during MD simulation relative to the initial\nstructure. Four independent replicate simulations for (a) ALP-MC and (b) ALP-SP.\nFig. R1 RMSF of protein backbone atoms (C, C , N, O) between ALP-MC and\nα\nALP-SP as a function of residue sequence number. Shaded areas indicate mean ±\nSD from four independent simulations; solid circles denote residues where the mean\nRMSF difference between the compared states exceeds 0.2 Å.\ncontaining uncertainty test. The g(r) peaks of water oxygen atoms around (a) the\nmodified residue, (b) the catalytic site, and (c) the entire protein surface for ALP-MC\nand ALP-SP. Data points represent mean ± SD calculated from four independent MD\nsimulations. Statistical significance was assessed using two-way analysis of variance\n(ANOVA). *P < 0.05, **P < 0.01, ***P < 0.001; ns, no significance.\nFig. S19 Replicated probability distribution of the group-specific water density\nfluctuations ([<N2-<N>2]/<N>) calculated within spherical volumes around each\nsurface residue group of ALP-MC and ALP-SP.\nH-bonded water\n(top) and the corresponding frequency distributions of ΔN (bottom)\nH-bonded water\nbetween ALP-MC and ALP-SP systems containing uncertainty test. Data are\nstratified by residue type: (a, e, i) negatively charged residues; (b, f, j) positively\ncharged residues; (c, g, k) polar (neutral) residues; (d, h, l) nonpolar residues.\n(2) Similar issues with large uncertainty are present for some experimental results. The\nauthors should comment on the statistical significance of the data in Fig. 1g, where\nALP-MC and RMC appear within error despite being describe as different in the text.\nLikewise, are the Rg values obtained from the data in Fig. 4h statistically significant?\nReply: Thank you for your insightful comments.\nTo evaluate the statistical significance of fluorescence lifetimes in Fig. 1g, we\nperformed two-way analysis of variance (ANOVA) on the datasets. ANOVA indicated\nthat ALP-MC and ALP-RMC are significantly different (p value of 0.0068), and Fig.\n1g was updated. We also applied the same rigorous statistics to analyze the data for\nradius of gyration (Rg) and maximum particle dimension (D ). Obtained results\nmax\nconfirmed the statistical significance of differences between SAXS-derived structural\nparameters of ALP-MC and ALP-SP (p < 0.01, Fig. S37). Meanwhile, the difference\nbetween ALP-MC and ALP-RMC is less pronounced (p < 0.05, Fig. S37), indicating\nonly partial recovery of protein conformation.\nRevised text: “A statistically differences in fluorescence lifetime between ALP-MC and\nALP-RMC were observed (p < 0.01) (Fig. 1g).” (Page 5, Lines 17-18, in red)\n“Guinier analysis revealed an increase in R (36.93→38.63 Å; Fig. 4h) and D\ng max\n(109→122 Å; Fig. 4i) from ALP-MC to ALP-SP (p < 0.01), indicating enhanced\nstructural flexibility. The broadening of the Kratky plot peak was also observed in ALP-\nSP, a characteristic of global structural loosening that is not fully reversible upon\nrelaxation back to the MC state (Fig. S36-S37, Table S2).” (Page 16, Lines 6-9, in red)\n“Quantification and statistical analysis. Data are presented as mean ± standard\ndeviation (SD). Sample sizes (n), representing independent experiments, are indicated\nin the figure legends. Statistical analyses were performed using GraphPad Prism 9.5.1.\nComparisons across multiple groups were assessed using one-way or two-way analysis\nof variance (ANOVA), as appropriate. Exact P values are reported in the figures or\nfigure legends. A P value < 0.05 was considered statistically significant.” (Page 29,\nLines 2-7, in red)\nuncertain test. Comparison of (a) radius of gyration (Rg) and (b) maximum particle\ndimension (D ) among ALP-MC, ALP-SP and ALP-RMC. Data are presented as\nmax\nmean ± SD (n = 3 per group). Statistical significance was assessed using one-way\nANOVA. ns, not significant; *P < 0.05; **P < 0.01.\n(3) The MD simulations focus entirely on the role of water, but the MC and SP isoforms\nhave substantially different charge distributions and potentially different conformations.\nDo MC/SP themselves interact with surface-exposed amino acids differently? Such\ninteractions might impact protein conformation without impacting water structure.\nReply: Thank you for raising this important issue.\nWe performed a quantitative energy decomposition analysis using MM/GBSA to\nevaluate whether MC and SP isoforms interact differently with surface-exposed amino\nacids, and if such interactions independently affect protein conformation. This approach\nallowed us to calculate three key interaction energy components—vacuum electrostatic\nenergy, solvated electrostatic energy, and van der Waals energy—to quantify the\ninfluence of modified MC/SP (position 406) on the rest of protein.\nIn this analysis, MC/SP modification at the mutation site was treated as a “ligand”\nwithin a protein-ligand binding free energy framework. Specifically, the solvated\nbinding free energy between ligand (MC/SP) and the receptor (the remainder of the\nprotein) was decomposed into five distinct components: internal energy terms, vacuum\nelectrostatic energy, van der Waals energy, polar solvation energy and non-polar\nsolvation energy (see details in the derivation below). Through this per-residue energy\ndecomposition, we further resolved the differences of interaction energies between\nThe results are shown in Fig. S40 (where distance is defined between the C atom\nα\nof MC/SP molecule and the C atoms of other amino acid residues).\nAs shown in Fig.\nα\nS40a (ALP-MC) and b (ALP-SP), van der Waals interaction energy contributions from\nthe mutation site to other residues were minor in both isoforms (overall < 5 kcal/mol),\nwhile electrostatic interactions being the dominant form. Notably, although both\nisoforms exhibit long-range electrostatic interactions in vacuum (energy contributions\nexceeding 10 kcal/mol at distances up to ~20 Å from the mutation site), the shielding\neffect of solvent water is profound. The electrostatic energy decays rapidly with\ndistance, becoming negligible beyond ~5 Å, with only residues directly covalently\nlinked to the mutation site retaining strong electrostatic interactions.\nThus, despite the inherent differences in charge distribution between MC and SP\nisoforms, their interaction patterns with surface-exposed amino acids—the relative\nstrength of van der Waals/electrostatic contributions and distance dependence of these\ninteractions—are highly consistent. This suggests that the difference in charge\ndistribution between isoforms does not lead to significant differences in their\ninteractions with surface-exposed amino acids. Furthermore, as the dominant\ninteraction (electrostatic) is effectively screened by solvent water and van der Waals\ninteractions are intrinsically weak, the strength of interactions between the MC/SP and\nother surface residues is unlikely to induce significant conformational changes in either\nisoform.\nDeduction process\nFor the complex formed by the ligand and receptor, the solvated binding free\nenergy is defined as:\nΔG =ΔG +ΔG -(ΔG +ΔG ) (1)\nbind, solv bind, vac com, solv rep, solv lig, solv\nHere, ΔG -(ΔG +ΔG ) can be decomposed into ΔG +ΔG\ncom, solv rep, solv lig, solv pol nonpol\ncomponents. ΔG is calculated using the Generalized Born (GB) model, while\npol\n7\nΔG is given by:\nnonpol\nΔG =γSASA+β (2)\nnonpol\nThe term ΔG in equation (1) is:\nbind, vac\nΔG =ΔG -(ΔG +ΔG ) (3)\nbind,vac com,vac rep,vac lig,vac\nNeglecting entropic effects,ΔG becomes:\nbind,vac\nΔG =E -(E +E ) (4)\nbind,vac MM,com,vac MM,rep,vac MM,lig,vac\nThe molecular energy in vacuum, E , is:\nMM,vac\nE =E +E +E (5)\nMM,vac int,vac vdW,vac ele,vac\nThe internal energy of the complex E can be decomposed into contributions\nint,com,vac\nfrom the receptor (E ), the ligand (E ), and an internal energy term\nint,rep,vac int,lig,vac\nbetween the receptor and ligand (E ). For non-covalent complexes, this cross\nint,rep-lig\nterm is zero, but it is non-zero in our covalent system.\nThe electrostatic energy of the complex, E , is decomposed as:\nele,com,vac\nj∈M N-1 M-1\ni∈N N M\nE =∑ ∑E +∑ ∑E +∑ ∑E (6)\nele, com,vac ij ih jh\ni i j\nj h≠i h≠j\nThe first term represents the non-bonded electrostatic interaction between the\nreceptor and ligand, the second term represents intra-receptor interactions, and the third\nterm represents intra-ligand interactions. Thus, the final result of equation (4) yields\nthree key interaction components between the receptor and ligand: the electrostatic\ninteraction energy, the van der Waals interaction energy, and the internal energy:\nthree key interaction components between the receptor and ligand: the electrostatic\ninteraction energy, the van der Waals interaction energy, and the internal\nenergy:\nΔG =ΔE +ΔE +ΔE (7)\nbind,vac int,rep-lig ele,interation vdW,interation\nConsequently, the total solvated binding free energy is:\nΔG =ΔE +ΔE +ΔE +ΔE +(γSASA+β) (8)\nbind,solv int,rep-lig ele,interation vdW,interation GB\nFor per-residue energy decomposition, the interaction energy between a single\nreceptor residue and the ligand (MC/SP) is partitioned, with one part assigned to residue\nand the other to the ligand. Based on this, the three target energy terms in our study are\nexplicitly defined as:\n8\nΔE\nE ele,vac = ele,interation\nΔE +ΔE\nE ele,solv = ele,interation GB\nΔE\nE vdW = vdW,interation\nRevised text: Fig. S40 and the corresponding methodological details were supplied to\nthe Supporting Information, specifically in Section 4.4. Additionally, we have included\na discussion of the energy decomposition analysis and its implications in the main text.\n“To further evaluate the role of specific MC/SP-surface interactions, we conducted\nMM/GBSA energy decomposition. The results show that interactions near modified\nMC/SP are mainly electrostatic, with negligible van der Waals contributions. Solvent\nwater strongly screens these electrostatic effects beyond ~5 Å. Despite differing charge\ndistributions, both isoforms exhibit highly similar interaction profiles with surface\nresidues (Fig. S40). Therefore, the observed changes in flexibility and stability are more\nlikely due to HBN reorganization instead of direct effect of the modification.” (page 15,\nline 23-29, in red)\nat the 406 mutation site. Average interaction energies (E , E , E ) between\nvdW ele, vac ele, solv\nMC/SP modification and all other residues are plotted as a function of C -C distance.\nα α\nResults are shown for (a) the ALP-MC and (b) ALP-SP isoforms. Panels (c-h) show the\nenergy decomposition of interactions between the mutated MC/SP site and the\nsurrounding protein residues: (c, d) electrostatic interactions in solvent water; (e, f)\nelectrostatic interactions in vacuum; (g, h) van der Waals contributions.\n10\n(4) Some issues of clarity:\n- Pg. 4 – the distances listed after each mutation presumably refers to the distance\nfrom the catalytic cite, but this isn’t clear in the main text.\nReply: We have revised the text to improve clarity.\nRevised text: “Using the crystal structure of ALP (PDB:1ED9) as a guide22,\nspecific residues in the loop region, located at varying distances from the catalytic site,\nwere replaced with the unnatural amino acid p-azido-L-phenylalanine (pAzF). The\nselected sites were H372/872 (~5 Å from the catalytic site), V99/599 (~10 Å),\nE406/906\n(~15 Å), and K185/685 (~25 Å)”. (Page 4, line 28, in red)\n- Fig. 1a – it is unclear what the numerical values (-17.7 D, -4.3 D) in the figure refer\nto.\nReply: The numerical values denote the molecular dipole moments for the MC and SP\nstates, which are ~17.7 Debye and ~4.3 Debye, respectively. This definition has been\nexplicitly added in Fig.1a and corresponding caption.\nRevised text: “(a) Reversible photoisomerization between merocyanine (MC) and\nspiropyran (SP) accompanied by a large shift in molecular dipole moment (μ) (in\nDebye).” (Page 6, caption in Fig. 1a, in red)\n- Fig. 1b – both measured contact angles are <90 degrees and would correspond to\nhydrophilic surfaces. The caption should clarify that SP is more hydrophobic than MC,\nnot that the surface itself is hydrophobic.\nReply: Thank you for your correction. We have revised the labels of Fig. 1b to\nclarify that SP is relatively more hydrophobic than MC, with MC labeled as “less\nhydrophobic” and SP as “more hydrophobic”.\n- The authors refer to hydration layers (<4 A, etc). How are these distances computed –\nthat is, what reference atoms are used when calculating this threshold distance, and\nwhat defines the distal catalytic site vs. modified residue? For the latter, are the atoms\nin MC/SP included? What happens if water molecules diffuse outside of this range\n11\nwithin the 50 ps of sampling, are they still included?\nReply: We thank the reviewer for these insightful questions and are happy to clarify\nthe definitions and analysis procedures.\n1. Definition of the Hydration Layer\nThe definition of hydration layer was based on the position of the first minimum\nin the protein–water radial distribution function, g(r), which demarcates the first\nhydration shell. The reference atoms used for the g(r) calculation were selected as\nfollows:\nFor the entire ALP protein: all protein atoms and the Zn–Zn–Mg ions.\nFor the distal catalytic site: the Zn–Zn–Mg ion cluster.\nFor the modified residue site: .the atoms of the mutated amino acid backbone and\nside chain, including the MC/SP modifier atoms.\n2. Treatment of Water Molecules in Dynamic Analyses\nThe handling of water molecules was adapted to each specific calculation.\nFor g(r) analysis: as a statistical ensemble property, all water molecules in the\nsystem were included regardless of temporary diffusion.\nFor hydrogen bond lifetime analysis: the initial set of water molecules within the\ndefined cutoff (based on the first frame) was tracked throughout the entire 50 ps\nsampling period, even if they diffused out of the region.\nFor IR spectral calculations: only water molecules that remained persistently\nwithin the defined region during the entire sampling window were considered, ensuring\nthat the spectrum reflects the dynamics of a stably bound hydration layer.\nPlease accept our heartfelt thanks for your thorough consideration and constructive\nsuggestions. We have conducted additional control experiments and refined the\nlanguage.\nPoint-by-point response to your comments is as follows.\n(1) On p. 5 the authors state that the modification occupies only 2.2 ‰ residues. It\nwould perhaps help to state how much of the surface is altered, not the percentage of\namino acid in total.\nReply: To address this concern, we performed a detailed analysis of the solvent-\naccessible surface area (SASA) to accurately evaluate the extent of surface modification.\nThe results show that the modified MC/SP accounts for approximately 4% of total\nSASA (see Fig. R2).\nFollowing your suggestion, the original description based on the percentage of total\namino acid residues has been removed from the manuscript.\nFig. R2 Proportion of solvent-accessible surface area (SASA) of modified MC/SP\nmolecules vs. total protein surface area. Molecular dynamics (MD) simulations were\nused to quantify this proportion, revealing the contribution of modified MC (a) and SP\n(b) molecules to the protein’s surface properties.\n(2) On p. 5 and Fig. 6 it is shown that the ALP-MC and ALP-RMC show significant\n13\ndeviations in the fluorescence lifetime data. The authors claim this suggests that the\nisomerization alters the local microenvironment on the protein surface. Why is that the\nmost likely option? Would it not be possible (if not more likely) that the protein to\ntransition into a new configuration? In the cited study [25] Ma et al. do not discuss MC\nas a probe for polarity shifts, viscosity etc. I find this section too vague and suggest an\noverhaul.\nReply: We thank the reviewer for this insightful comment. We agree that protein\nconformational changes could contribute to the observed fluorescence lifetime\ndifferences between ALP-MC and ALP-RMC. Our data indicate that the changes of\nfluorescence lifetime are attributed to the hydration-shell reorganization that leads to\nprotein conformational fluctuations.\nSingle-residue photoisomerization initially perturbs the local hydration layer and\ntriggers a reorganization of global hydration water, which subsequently induces modest\nconformational fluctuations of the protein. These structural responses further reshape\nthe hydration shell, forming synergistic feedback that alters the protein surface\nmicroenvironment (discussed in Fig. 4).\nSeveral observations support this interpretation: (1) the fluorescence lifetime of\nMC is highly sensitive to solvent polarity (Fig. S13); (2) MC conjugated at different\nALP surface residues (V10, E15, and K25) exhibits distinct fluorescence lifetimes (Fig.\nS14), indicating its sensitivity to site-specific environments; (3) the fluorescence\nlifetime difference between free MC and regenerated free MC in aqueous solution is\nmuch less pronounced whereas statistically significant changes occur only upon\ncovalent conjugation to ALP (Fig. 1g), suggesting that MC-SP-MC isomerization\nspecifically perturbs the protein-associated hydration shell that possibly leads to\nchanges of protein conformation.\nWe characterized protein conformational changes using MD simulations and\nsmall-angle X-ray scattering (SAXS) (Fig. 4). These analyses reveal that after\nphotoisomerization to SP, the protein show increased side-chain fluctuations and a more\nloosely packed protein surface with incomplete recovery to the original state. This\nbehavior is consistent with non-catastrophic conformational changes rather than a\n14\ndominant transition to a distinct global protein configuration.\nWe have clarified this point in the revised manuscript.\nIn response to your suggestion, we replaced Ref. 25 with a more appropriate\ncitation that explicitly establishes MC-like fluorophores as probes for local polarity,\nviscosity, and hydration dynamics (Chem. Commun. 2021, 57, 13313-13316).\nRevised text: “We then investigated the photoisomerization-induced changes of\nmicroenvironments on the engineered protein surface by measuring the fluorescence\nlifetime of MC. As a sensitive reporter for polarity (Fig. S12), MC exhibits site-\ndependent fluorescence lifetimes upon conjugating to different sites of ALP, indicating\nvariations in local polarity microenvironment (Figs. S13-S14). The fluorescence\nlifetimes of ALP-MC and its revertant state (RMC, regenerated by photoisomerization\nfrom SP to MC) were recorded by time-correlated single photon counting\n(TCSPC)24(Fig. 1f). Statistically differences in fluorescence lifetime were observed\nbetween ALP-MC and ALP-RMC (p < 0.01) (Fig. 1g). In contrast, negligible variations\noccurred between free MC and RMC (Fig. 1g, Fig. S15). This indicates that MC-SP\nisomerization specifically triggers rearrangements of the hydration shell that possibly\nleads to changes of protein conformation.25” (Page 5，line 12-21, in red)\n(3) The authors use Far-IR (THz) ATR measurements of the hydrated sample to probe\nthe spectroscopic signatures in the two conformations of the surface-attached\nphotochromic molecule. On p. 11 the authors describe the “deconvolution method” to\ndescribe their ATR spectra. They use a sum of two damped harmonic osciallators. I\nwould suggest adding the equations to the supplementary. The results of the fits are\nshown in Fig S19. The authors then directly attribute the amplitudes (denoted as\n“intensities” in the text) to a bound and wrap water fraction. A justification for this fit\nprocedure is missing. This is crucial as this is the fundament for further discussion of\nthe data. Studies by the Havenith group found that these spectroscopic regions are\ncorrelated for a “bound” and “wrap” water fraction. However, this correlation was not\nfound by an intensity comparison of a damped harmonic oscillator model, but they\ndemonstrated changes in the slope in the 400 and 600 cm-1 region after subtraction of\n15\ncorresponding bulk water contribution, which is not performed here. I suggest to\ncarefully check if this procedure can be applied here as well.\nReply: We thank the reviewer for this insightful comment. We fully agree that a clear\nmethodological justification is essential, as the interpretation of ATR- THz spectra lay\nthe foundation for subsequent discussion.\nFollowing the reviewer’s suggestion, we now include the damped harmonic\noscillator (DHO) model equations in the Supplementary Information (Section 4.2.3,\nPage 7) for spectral deconvolution, together with detailed definitions of all fitting\nparameters1, 2,\n3 aων2\nΔα(ν)=∑\ni i\n2\ni=1 ω2\nν2ω2+π2(ν2+ i -ν2)\ni i 4π2\nwhere α, ω, and ν correspond to the amplitude, width, and the central frequency of the\ni i i\ni-th resonance, the unperturbed central frequency is given as ν =√ν2+ ω i 2 .\nThe model\n0,i i 4π2\nformulation is adapted from established THz calorimetry and hydration spectroscopy\nframeworks (Havenith et al., 2017; Pezzotti et al., 2023).\nWe have refined the physical basis of our analysis. Rather than directly attributing\nDHO amplitudes to bound and wrap water fractions, we now adopt difference\nabsorption coefficient spectra (Δα = α – α ), an approach widely used by the\nALP-SP ALP-MC\nHavenith group, to suppress bulk water and protein core contributions and isolate\nhydration-layer-specific signatures (Fig. 3g; Figs. S26-S27; Table S1).\nWithin this framework, bound and wrap water contributions are inferred primarily\nfrom slope changes in the 400-600 cm⁻¹ region and amplitude variations in the 150-200\ncm⁻¹ region, respectively. The DHO model is used to parameterize systematic spectral\nchanges rather than as a direct physical proxy.\nWe revised Fig. 3 and the corresponding text accordingly and derived\nthermodynamic signatures (ΔS > 0, ΔH < 0) consistent with a hydration-driven\ninterfacial reorganization upon photoisomerization. These revisions ensure that the\ndeconvolution procedure is physically justified, reproducible, and aligned with\nestablished THz spectroscopy methodologies.\nrearrangements between ALP-MC and ALP-SP, we used THz calorimetry to correlate\nspectral changes with entropy (ΔS) and enthalpy (ΔH). Absorption coefficients (α(ν))\nand photoisomerization-induced differential spectra (Δα(ν) , ALP-SP minus ALP-\nSP-MC\nMC) were calculated to isolate hydration-specific alterations (Eqs. 1-3, Fig. S26). The\nΔα(ν) was deconvolved into the hydrophobic “wrap” region (150-200 cm-¹,\nSP-MC\nentropy-linked) and hydrophilic “bound” region (450-600 cm-¹, enthalpy-linked) (Fig.\nS27).\nSpectral analysis of Δα(ν) spectra at different hydration levels all revealed a\nSP-MC\nnegative Δα amplitude (Δα < 0), signifying a positive entropy change (ΔS > 0,\nwrap wrap\nEq. 4 and S3, Table S1) due to the release of loosely wrap-water from hydrophobic\npatches (Fig. 3g). Concurrently, a positive slope in the bound region (Δα /Δν > 0)\nbound\nindicated a negative enthalpy change (ΔH < 0, Eq. 5), reflecting enhanced rigid H-\nbond networks with hydrophilic residues. Consequently, the THz results demonstrated\nthat a single-residue hydrophobicity change can induce HBN reorganization across\nglobal hydration by converting water from a loosely associated \"wrap-water\" state to\na tightly bound, ordered configuration, thereby forming a rigid hydration layer.\nFurthermore, the observed entropy-enthalpy compensation minimizes the system's free\nenergy, yielding a net favorable hydration contribution (ΔGhydration < 0, Eq. 6) that acts\nas the thermodynamic driver for conformational destabilization.” (Page 12, line 10-29,\nin red)\nFig. S26. ATR-THz absorption coefficient (α) spectra for ALP-MC and ALP-SP at\n17\nhydration levels spanning 10% to 100% (w/w).\nFig. S27 Differential absorption spectra (Δα) across hydration levels. (a-c)\nExperimental (circles) and fitted (lines) Δα(ν) spectra under full (100% w/w), partial\n(50% w/w) and critical (10% w/w) hydration level. Deconvoluted components are\nshown as shaded bands.\nTable S1 Damped harmonic oscillator fit results of hydrogen-bond stretching and\nin-/out-of-plane librational bands from differential absorption spectra (Δα) at\ndifferent hydration levels (100%, 50%, 10% w/w).\nSample a ν (cm-1) a ν (cm-1) a ν (cm-1)\n1 1 2 2 3 3\n100% -489 179.86 -1285 319.91 -1357 437.55\n50% -720 178.27 -2252 338.56 -4200 481.95\n10% -1614 176.67 -4204 325.85 -9628 499.78\n(4) In Fig. 3e) how were the error bars calculated? Are the errors calculated from the\nfit? Were the ATR measurements repeated/reproduced? I would expect that water\ncontent varies significantly between samples and thus could potentially contribute to a\nsystematic error. Another important factor here is temperature. It is well known that the\nlibrational band of water has a strong temperature dependence. With increasing\ntemperature, the librational band red shifts and decreases in amplitude. Following the\nanalysis of the authors, this would also lead to a similar change in the I / I (which\nbound wrap\nmeasures the intensity of the bands at ~150 and ~550 cm-1). Considering that it is not\nimplausible to assume that some heating is induced during the sample illumination, this\ncould potentially lead to these amplitude changes as well. Furthermore, it is reasonable\nthat due to e.g. evaporation of water the overall water content could be altered. The\n18\nauthors need to convincingly clarify how (if) the temperature was controlled. In Fig. 1e)\nthe authors demonstrate that the change from MC to SP is (at least to some degree)\nreversible. It would be reassuring to see the reversibility also here for the Far-IR ATR\nexperiments.\nReply: We thank the reviewer for these insightful and critical questions.\nError bars and reproducibility. The error bars in Fig. 3e represent the standard\ndeviation obtained from three independent ATR-THz measurements, each performed\non separated hydrated protein film (Fig. R3). The reported errors reflect the natural\nvariability in sample preparation, hydration level, and film uniformity, instead of fitting\nuncertainty. All ATR measurements were repeated and reproduced under identical\nexperimental and analysis protocols, ensuring that the reported errors reflect genuine\nexperimental variability rather than fitting artifacts or analysis inconsistencies.\nFig. R3 ATR-THz spectroscopy probes the hydration dynamic changes between\nALP-MC and ALP-SP. ATR-THz spectroscopy reveals hydration dynamics of photo-\nswitched albumin. ATR-THz absorption spectra of ALP-MC and ALP-SP states at\nhydration levels of 100% (a), 50% (b), and 10% (w/w) (c). Data are from three\nindependent experiments.\nControl for water content. We fully agree that both water content and temperature are\ncritical experimental variables.\nTo exclude water-content induced artifacts, we implemented quantitative hydration\ncontrol and verification procedures (detailed in Supporting Information Section 4.2.2).\nHydration levels were directly quantified using a hydration ratio (R ), defined as the\nhyd\nintegrated intensity ratio of the water-sensitive O–H stretching band (~3500 cm⁻¹) to\nthe hydration-insensitive protein amide II band (~1550 cm⁻¹) (Fig. S22a). The mean\ndifference in hydration ratio between ALP-MC and ALP-SP states (ΔR was < 0.02),\nhyd\n19\nwell below the commonly accepted equivalence threshold of 0.05, confirming\ncomparable hydration states during analysis.\nTo further minimize variability and prevent evaporation, all samples were prepared\nunder controlled humidity conditions with precise water addition and were hermetically\nsealed during measurements. Consistent O–H stretching band intensities across all\nreplicates and experimental stages confirm the absence of detectable water loss or drift\n(Fig. S22b).\nFig. S22 Exclude possible changes water content of hydrated protein film induced\nby illumination and evaporation. (a) FTIR spectra of hydrated protein films after\nphotoisomerization. Red arrows denote the O-H stretching band of water (~3500 cm-1)\nand amide II band (~1550cm-1). (b) Relative water content (normalized to amide II) for\nALP-MC and ALP-SP across hydration levels (10%, 50%, and 100%). The data shown\nare presented as the mean  SD (n=3).\nTemperature Stabilization. We agree with this point regarding the strong temperature\ndependence of the water librational band. Although the ATR-THz setup does not\ninclude active temperature control, we explicitly quantified possible photo-induced\ntemperature rise with a micro-thermocouple. Under the same illumination conditions\nfor MC/SP transition (450 nm, 10 mW cm⁻²), temperature measurements revealed a\nmaximum increase of < 1 °C (Fig. S24).\nTo assess the influence of such minor heating on ATR-THz spectra, we performed\ncontrol experiments with pure water and wild-type ALP in Tris buffer under same\nillumination conditions. As shown in Fig. S25, illumination-induced temperature rise\ndid not produce detectable changes in the librational band or overall THz absorbance.\nenvironment (20 °C), eliminating external thermal fluctuations. Taken together, these\ncontrols demonstrate that the spectral changes reported in Fig. 3 arise from\nphotoisomerization-induced interfacial hydration rearrangements, rather than thermal\nartifacts.\nFig. S24 Exclude possible light-induced heating. (a) Temperature changes of a 10 µL\nsolution under 450 nm illumination were monitored with a micro-thermocouple. (b)\nTemperature changes over different illumination times (0-20 min) and c corresponding\npost-illumination temperature measurements (0-30 min).\nFig. S25 The Effect of illumination on THz absorbance spectra. Spectra changes of\n(a) pure water and (b) wild type ALP in pure water after exposure to 450 nm light for\n10 min. Data are from three independent experiments\nReversibility for the Far-IR ATR experiments. We appreciate the reviewer’s\nsuggestion to examine reversibility in the ATR-THz measurements.\nThe apparent difference between Fig. 1e and Fig. 3 arises from the fundamentally\ndifferent molecular mechanisms probed by the two techniques. UV-vis spectroscopy\n(Fig. 1e) monitors molecular-level reversibility of the MC-SP photoisomerization.\nIn\ncontrast, ATR-THz spectroscopy probes the collective hydration-protein system. Our\ndata indicated that while the photo switching is reversible, it triggers irreversible global\nchanges, including loss of enzymatic activity (Fig. R4), persistent alterations in\nhydration hydrogen-bond lifetimes (Fig. S13), and conformational changes detected by\nSAXS (Fig. S36, Table S2).\nConsistently, repeated photo-switching cycles in ATR-THz experiments reveal that\nthe hydration-layer signatures do not fully revert to their initial state (Fig. S23). This\nlack of reversibility is therefore not an experimental limitation, but a key finding of this\nwork that localized hydrophobicity perturbation can drive a global, irreversible\ntransition in protein hydration and structure. This irreversibility suggests that the newly\nformed interfacial water-protein interactions stabilize a thermodynamically favored\nstate that is no longer accessible to the original functional conformation.\nRevised text: “The broadening of the Kratky plot peak was also observed in ALP-SP, a\ncharacteristic of global structural loosening that is not fully reversible upon relaxation\nback to the MC state (Fig. S36-S37, Table S2)”. (Page 16, line 6-9, in red)\n“Consistently, ATR-THz experiments with reversible ALP-RMC photo-switching\nreveal that the hydration layer is unable to fully revert to its initial state (Fig. S23). This\nindicates that a localized perturbation can induce a global and irreversible transition\nin protein hydration.” (Page 12, line 1-3, in red)\nFig. R4 Photo-controlled enzyme activity. A comparison of the catalytic kinetics\nbetween ALP-MC (dark) and ALP-SP (post-illumination at 450 nm) over 60 minutes.\nmodified-ALP variants. (a) Fluorescence lifetime images of ALP-E15-MC/-RMC.\nScale bar: 2 μm. Color scale represents lifetime values (right). (b) Ensemble-averaged\nTCSPC decay curves for ALP-E15-MC and ALP-E15-RMC. (c) Corresponding\nlifetime histograms with Gaussian fits for E15 variants. Mean lifetime (τ ± σ) and\nmolecule count (N) annotated.\nFig. S36 SAXS analysis of ALP-MC, ALP-SP and ALP-RMC. (a) Experimental\nSAXS profiles, displayed as logI(q) versus the scattering vector q (q = 4 πsinθ/λ). (b)\nThe linear region of the Guinier plot, from which Rg and I(0) can be derived. (c)\nDistance probability functions, p(r) of the three samples. (d) Scattering curve of the\nthree samples shown in dimensionless Kratky plots ((qRg)2I(q)/I(0) versus qRg). The\ndata shown in (a-d) are presented as the mean ± SD from 3 independent experiments.\nSP and ALP-RMC.\nSample Rg (Å)a D (Å)b\nmax\nALP-MC 36.83 ± 0.11 109 ± 0.05\nALP-SP 38.61 ± 0.10 124 ± 0.04\nALP-RMC 37.98 ± 0.05 117 ± 0.08\nTable notes:\na. The radius of gyration (Rg) was determined from the Guinier approximation in the\nlow-q region (q*Rg < 1.3).\nb. The maximum particle dimension (D ) was estimated from the pair-distance\nmax\ndistribution function, p(r).\nFig. S23 ATR-THz spectroscopy assesses the hydration dynamics across a\ncomplete MC→SP→RMC photoisomerization cycle.\n(5) On p. 12 the authors state that the MD-calculated spectra cannot reproduce the low-\nfrequency modes of water below 300 cm-1 due to force-field limitations. It would be\ninsightful to also have a clearer picture on the HB-stretch region, but I understand that\nthe cost for such simulation would be large. It is, however, necessary to clarify the\nlimitations of the current MD model regarding the librational band. Surely it was\npreviously discussed in the literature to what extent the force field has an influence on\nthe accuracy of the librational band?\nReply: We thank the reviewer for highlighting the force field limitations in reproducing\nthe librational band and low-frequency modes. We agree that the limitations of classical\nMD force fields in reproducing the librational band and other low-frequency water\nmodes should be clearly stated and contextualized within prior literature.\nTo this end, we systematically evaluated five widely used AMBER-supported\nwater models (TIP3P, rigid SPC/E, SPC/Fw, qSPC/Fw, and OPC3-pol), using PBE-\nbased DFT calculations as a reference for vibrational behavior. This comparison\nconfirms a well-established consensus in the literature: the librational band of water is\nhighly sensitive to force-field choice, particularly to molecular rigidity, flexibility, and\nthe inclusion of polarization effects.\nAs shown in Fig. S28, rigid water models such as TIP3P and rigid SPC/E, which\nconstrain intramolecular geometry, fail to reproduce the full vibrational spectrum of\nwater. These models systematically yield blue-shifted and overly narrow features,\nespecially in the O–H stretching and H–O–H bending regions, and consequently\nprovide an inadequate description of librational dynamics. Flexible fixed-charge\nmodels (SPC/Fw and qSPC/Fw) improve the description by allowing intramolecular\ndegrees of freedom and partial inclusion of nuclear quantum effects, but still exhibit\ndeviations in the librational band due to the absence of explicit electronic polarization.\nPolarizable water models are known to provide a more accurate representation of\nvibrational spectra, including librational modes. However, as previously reported3, 4,\nsuch models (e.g., OPC3-pol) may introduce additional artifacts, such as over-amplified\nlibrational peaks associated with Drude oscillators. These trade-offs have been\nextensively discussed in prior studies and remain an active area of methodological\ndevelopment.\nBased on this evaluation, we selected qSPC/Fw as a practical compromise between\ncomputational efficiency and spectral fidelity. While this model does not reproduce the\nabsolute peak positions or detailed line shapes of the librational band with quantitative\naccuracy (Fig. S29), it reliably captures relative spectral shifts and hydrogen-bond\nnetwork dynamics in response to perturbations. Importantly, the focus of our MD\nanalysis is on comparative, photoisomerization-induced changes rather than absolute\nreproduction of experimental THz spectra.\nIn the revised manuscript, we now explicitly acknowledge these limitations and\nclarify that interpretations of the librational band are made with these established\nconstraints in mind. Corresponding discussions and additional comparative data have\nbeen added to the Supporting Information (new Figs. S28 and S29), providing\ntransparency regarding the force-field dependence and the scope of validity of the MD\nresults.\nRevised text: “The employed SPC/E water model exhibits a known force-field\nlimitation that precludes the capture of the 200 cm-¹ peak44. Nonetheless, it qualitatively\nreproduces the librational band above ~300 cm-¹, showing agreement with the\nexperimental spectrum of ambient water (Fig. S29). The known systematic blueshift\ndoes not impact the relative trends under discussion.” (Page 13, line 6-10, in red).\nFig. S28 Comparison of the full MD-calculated infrared (IR) spectra across six\nwater models with the standard experimental IR spectrum of bulk H₂O. Spectra\nwere computed as the Fourier transform of the dipole gradient-velocity autocorrelation\nfunction for six AMBER-supported classical water models (TIP3P, rigid SPC/E,\nSPC/Fw, quantum-corrected flexible qSPC/Fw, polarizable OPC3-pol) and reference\ndata from PBE-DFT ab initio molecular dynamics (AIMD).\nspectra. Calculated THz spectra of bulk H₂O using five AMBER-compatible water\nmodels (TIP3P, rigid SPC/E, SPC/Fw, qSPC/Fw, OPC3-pol) and PBE-DFT-based ab\ninitio molecular dynamics (AIMD) simulations, compared to the experimental\nspectrum.\n(6) On p. 15 the authors present SAXS data. For the ALP-SP upon photoirradiation the\ndata shows a broadening of the Kratky plot peak (structural loosening) and a reduction\nof the thermal denaturation melting temperature by 5 °C. The authors state that this\nsuggests an augmentation of the global protein conformational flexibility via collective\nreorganization of the hydrogen bond network. Further they find changes in the\nenzymatic activity. It is unclear to me how the finding of a change in flexibility or\nenzymatic activity can only be explained by a change in the HBN? How can the authors\nexclude a direct influence of the added SP/MC on the protein? The data clearly shows\nthat the photoisomerization is not fully reversible if attached to the protein. Does this\nnot show that the added SP/MC clearly directly (and perhaps even irreversibly) reacts\nwith the protein itself?\nReply: We thank the reviewer for raising this important point. We therefore address this\nconcern explicitly below.\nExclusion of direct MC/SP-protein interactions\nTo assess whether the MC and SP isoforms perturb protein structure differently\nthrough direct interactions with surface residues, we performed a quantitative\nMM/GBSA energy decomposition analysis. In this framework, the MC/SP moiety at\n27\nposition 406 was treated as a “ligand”, and the remainder of the protein as the “receptor”.\nThe total solvated interaction free energy was decomposed into vacuum\nelectrostatic, van der Waals, polar solvation, and nonpolar solvation contributions. Per-\nresidue energy decomposition was further carried out to resolve distance-dependent\ninteraction profiles (Fig. S40 and the corresponding methodological details were\nsupplied to the Supporting Information, specifically in Section 4.4).\nSeveral key observations emerge: 1) For both ALP-MC and ALP-SP, van der\nWaals interaction energies between the photo switch and other residues are weak,\nremaining below 5 kcal mol-1 at all distances (Fig. S40a, b). 2) Although long-range\nelectrostatic interactions appear notable in vacuum (>10 kcal mol-¹ within ~20 Å), these\ninteractions are strongly screened by water. In the solvated state, electrostatic\ncontributions decay rapidly and become negligible beyond ~5 Å, with only covalently\nadjacent residues retaining appreciable interactions. 3) Crucially, the magnitude, spatial\ndecay, and relative contributions of van der Waals and electrostatic interactions are\nhighly similar for MC and SP isoforms, despite their different charge distributions.\nThese results demonstrate that the direct physical interactions between the MC/SP\nmoiety and surface-exposed amino acids are both weak and nearly identical for the two\nisomeric states. As such, they are insufficient to account for the observed differences in\nglobal flexibility, thermal stability, or enzymatic activity.\nReversible MC/SP photoisomerization on the protein\nTo address the concern regarding irreversible chemical interactions between\nisomers and ALP, we performed repeated photo-switching cycles on ALP-MC.\nAs shown by the UV-Vis spectra (Fig. R5), MC ↔ SP photoisomerization remains\nfully reversible upon covalent conjugation to ALP and can be cycled many times\nwithout loss of switching efficiency. This rules out possible irreversible photochemical\nreactions between the MC/SP moieties and the protein backbone or side chains,\nconfirming that the isomers themselves remain chemically reversible.\nThe observed irreversibility in some outcomes (e.g., activity loss or hydration\nsignatures), however, do not response to the photoisomerization itself, but the effect of\n28\nphotoisomerization on HBN and global conformational changes.\nIrreversible Conformational changes and Enzymatic Outcomes\nHaving excluded direct interactions between modified MC/SP and protein, we\nattributed the broadened Kratky peak and reduced melting temperature to a hydration-\nmediated effect. The reversible local photoisomerization event alters surface\nhydrophobicity and interfacial hydration, triggering a collective reorganization of the\nhydration HBN. This reorganization propagates across the protein-water interface,\nleading to changes in global conformational flexibility and function.\nConsistent with this interpretation, the photo switching itself is reversible, whereas\nthe hydration-coupled structural relaxation of the protein is not fully reversible,\nreflecting a transition to a new, metastable conformational ensemble.\nIn summary, our additional reversibility experiments and quantitative energy\ndecomposition analysis demonstrate that:1) the MC/SP moiety does not irreversibly\nreact with the protein; 2) direct MC/SP-protein interactions are weak, solvent-shielded,\nand nearly identical for both isomers; 3) the observed SAXS and activity changes are\nbest explained by a hydration-mediated, collective mechanism rather than by direct\nperturbation from the attached photo-switch.\nRevised text: We have added Fig. S23 and the corresponding methodological details\nto the Supporting Information, specifically in Section 4.4. Additionally, we have\nincluded a discussion of the energy decomposition analysis and its implications in the\nmain text.\n“To further evaluate the role of specific MC/SP-surface interactions, we conducted\nMM/GBSA energy decomposition. The results show that interactions near modified\nMC/SP are mainly electrostatic, with negligible van der Waals contributions. Solvent\nwater strongly screens these electrostatic effects beyond ~5 Å. Despite differing charge\ndistributions, both isoforms exhibit highly similar interaction profiles with surface\nresidues (Fig. S40). Therefore, the observed changes in flexibility and stability are more\nlikely due to HBN reorganization instead direct effect of the modification.” (page 15,\nline 23-29, in red)\nUV/vis absorption spectra of ALP-MC and ALP-SP. (b)Reversible spectra switching of\nthe chromophore between its MC and SP forms, triggered by visible light illumination\nand incubation in the dark, respectively.\nat 406 mutation site. Average interaction energies (E , E , E ) between\nvdW ele, vac ele, solv\nMC/SP modificaton and all other residues are plotted as a function of Cα-Cα distance.\nResults are shown for (a) the ALP-MC and (b) ALP-SP isoforms. Panels (c-h) show the\nenergy decomposition of interactions between the mutated MC/SP site and the\nsurrounding protein residues: (c, d) electrostatic interactions in solvent water; (e, f)\nelectrostatic interactions in vacuum; (g, h) van der Waals contributions.\n(7) As a last note, I do not agree with the authors on using the term “butterfly effect”.\nIn simple words it refers to the phenomenon that in a dynamic system small changes of\n31\ninitial conditions can result in large differences in a later state. It is connected to a\ndynamic system and describes the unpredictability of small changes over time. A\nchange of hydrophobicity might lead to long-range differences but this does not\ndescribe the evolution in time.\nReply: Thank you for this precise comment. We fully agree that the term “butterfly\neffect” has a strict definition in nonlinear dynamics. We have removed the term\n“butterfly effect” throughout the manuscript.\nPlease accept our heartfelt thanks for your thorough consideration and constructive\nsuggestions. We have conducted additional control experiments and refined the\nlanguage.\nPoint-by-point response to your comments is as follows.\n(1) Page (p) 2, line (l) 8: ‘… we reveal that even subtle, residue level changes in\nhydrophobicity …’. See comment above. The changes by these two large tags do not\nappear to be subtle and should not be mistaken by the reader to be comparable to those\nat play when a single residue is exchanged against another.\nReply: We agree that the perturbation from the SP/MC tags is not “subtle”. We removed\n“subtle” in the revised manuscript (e.g., Page 2, Line 8; Abstract).\nWe would like to emphasize that core concept of this study is not to mimic\nphysiological mutations, but rather to employ a pronounced, photo-controllable\nhydrophobicity switch as a tool to probe residue-specific solvation effects in situ. This\napproach allows us to impose a well-defined and reversible local hydrophobicity\nperturbation at a specific site, which would be difficult to achieve using conventional\nmutagenesis.\nTo better reflect the scope and intent of the study and to avoid overstating its\nphysiological relevance, we have revised the manuscript title to: “Single-Engineered-\nResidue Solvation Perturbations Regulate Global Protein Architecture and Function”.\n(2). P 2, l 12: ‘Our findings establish that interfacial water networks not only capture\nthe surface physicochemical patterns of proteins, …’. The generalization of the author’s\nfinding using large tags does not seem to justify a generalization to proteins.\nReply: We thank the reviewer for this important comment.\nIn order to tone down the biological significance of our findings, and the title of\nthe revised manuscript is changed to “Single-Engineered-Residue Solvation\nPerturbations Regulate Global Protein Architecture and Function”\nWe also performed additional experiments to prove that the hydration shifts of\nsingle native residue could also alter global protein enzymatic reaction. The glutamic\n33\nacid (E) at residue E406 (equivalent to E905 in the full-length construct) of wild ALP\nwas replaced by a series of natural amino acids spanning a hydrophobicity gradient (I >\nF > G > Q > K) (Fig. S43a). We then tested the enzymatic activity of these mutated\nALP using hydrophilic substrate pNPP or hydrophobic substrate 4-MUP.\nFor hydrophilic substrate pNPP, the enzymatic activities correlated strongly with\nside-chain hydrophobicity (I < F < G < Q), while for hydrophobic substrate 4-MUP,\nenzymatic activities remained largely unchanged with one exception of K (Fig. S43b-\nc). We argued that the exception of K mutation was possibly attributed to a distinct\nelectrostatic mechanism instead of hydration effect.\nThese results from unmodified protein, further confirmed our conclusion that\nphysiologically relevant changes in surface hydrophobicity of single amino acid could\nmodulate enzyme function, presumably through reorganization of the hydration shell.\nThe corresponding discussion has been incorporated into the main text.\nRevised text: “To establish physiological generalizability, we introduced single-\npoint mutations at a conserved surface residue (ALP E406/905) using amino acids with\nvarying hydrophobicity (I > F > G > Q > K) (Fig. S43a). A clear correlation emerged\nbetween residue hydrophobicity and activity toward the hydrophilic substrate pNPP\n(activity: I < F < G < Q), while activity toward the hydrophobic substrate 4-MUP\nremained largely unchanged (Fig. S43b-c). The Lys (K) variant, likely modulating\nhydration via charge effects, was an exception. These results, alongside those from the\nSP/MC system, confirm that water-mediated allostery constitutes a general and tunable\nmechanism of protein regulation.” (Page 18, line 20-28, in red).\n(a) Schematic of the ALP mutation site and Alphafold 3-predicted structures of mutant\nproteins compared with the wild-type (WT) enzyme. The mutated residues span a\nhydrophobicity gradient: I > F > G > Q ~ E (WT) > K. (b,c) Enzymatic activities of the\nvariants were measured with (b) hydrophilic pNPP and (c) hydrophobic 4-MUP as\nsubstrates. Data are presented as mean ± SD (n = 3). Statistical significance was\ndetermined by one-way ANOVA. ns, no significance; ***P < 0.001; ****P < 0.0001.\n(3). P3, beginning of second paragraph: it is not clear exactly to which statement in the\nprevious paragraph the ‘however’ refers to.\nReply: We thank the reviewer.\nTo correct this, we have removed “however” from this sentence.\n(4). P3, beginning of third paragraph: Nickels et al (reference 17) did use neutron\nspectroscopy rather than ‘neutron scattering simulations.\nReply: We thank the reviewer for correcting this inaccurate description. We have\ncorrected in the revised manuscript text.\nRevised text: “NMR and neutron scattering experiments17, as well as simulation studies,\ncollectively suggest that localized mutations primarily affect the first-shell water (3-5\nÅ).” (Page 3, line 24-25, in red)\n35\n(5) P3, 3rd paragraph, l 7: Please specify to what experiments ‘femtosecond\nfluorescence’ refers to.\nReply: The term “femtosecond fluorescence” refers specifically to the femtosecond\ntime-resolved fluorescence Stokes shift and anisotropy measurements5.\nRevised text: “Nevertheless, recent evidence from femtosecond time-resolved\nfluorescence Stokes shift and anisotropy measurements indicated that water\nperturbations stemming from isolated mutations can exert global effects on solvation,\nwhich drives coupled protein side-chain motion”. (Page 3, line 27-28，in red)\n(6) P4, 2nd paragraph: The ‘butterfly’ effect is mentioned several times in the\nmanuscript, but the analogy not explained.\nReply: We thank the reviewer for this comment. We agree that the “butterfly effect”\nanalogy is not sufficiently defined and may cause ambiguity in a scientific context. All\nreferences to the “butterfly effect” have therefore been removed from the manuscript\n(pages 2, 5, and 19).\n(7) P4, 2nd paragraph: ‘structure-hydration-function’ appeared as ‘structure-function-\nhydration’ earlier in the manuscript.\nReply: We thank the reviewer for spotting this inconsistency. It has been corrected as\n“structure-hydration-function” throughout the text.\n(8) P4, second-to-last line: It should be explained why methionine mutations are\nmentioned here. Also, the length in brackets should be defined here already. Later in\nthe manuscript, it becomes clear that the distance to the active site is mentioned.\nReply: We thank the reviewer for these important clarifications. To resolve the\nambiguities, we have revised the text as follows.\nThe “M” notation (e.g., H872M) was site-directed modification and not indicative\nof methionine mutation. This misleading notation has been removed; we now explicitly\nstate the incorporation of pAzF at the specified position.\nThe values in brackets (e.g., 5 Å, 10 Å) represent the approximate distance\n36\nbetween the modification site and the catalytic center, and this definition has been added\nat the first mention of these values to ensure clarity.\nRevised text: \"Using the crystal structure of ALP (PDB:1ED9) as a guide22, specific\nresidues in the loop region, located at varying distances from the catalytic site,\nwere replaced with the unnatural amino acid p-azido-L-phenylalanine (pAzF). The\nselected sites were H372/872 (~5 Å from the catalytic site), V99/599 (~10 Å), E406/906 (~15 Å),\nand K185/685 (~25 Å).\"(Page 4, line 28, in red)\n(9) P5, second line: it should be rationalized why two and not only one tag is attached.\nAlso, what is the extension of each tag and at what distance to the protein surface is the\nisomerizing group located?\nReply: The use of two molecular tags is dictated by the homodimeric structure of\nalkaline phosphatase (ALP). Each monomer contains an engineered modification site,\nresulting in the attachment of one tag per monomer (two tags per functional dimer).\nRegarding the probe geometry, the fully extended lengths of the MC and SP\nisomers are approximately 14.9 Å and 14.2 Å, respectively (Fig. R6a,d). Due to linker\nflexibility, the isomerizing group samples a range of distances from the protein surface.\nMolecular dynamics simulations indicate its effective distance (relative to the C of the\nα\nmodified residue) is predominantly within 8.5-12.5 Å (Fig. R6b,e), with stable\npositioning during the simulated trajectory (Fig. R6c,f).\nmodification targets. (a,d) Schematic definition of distances between the\nisomerization site of MC/SP and the modification site DBCO. (b,e) Representative\nsnapshots from equilibrated MD simulations, illustrating the spatial distance between\nthe isomerization site of MC (b) or SP (e) and the C atom of the modified residue.\nα\n(c,f)\nTemporal evolution of distances between the isomerization site of MC (c) or SP (f) and\nthe C atom of the modified residue over a 20 ns trajectory after MD simulation\nα\nequilibration.\n(10) P5, second paragraph: from which group(s) does the measured fluorescence\noriginate, from the tags?\nReply: The fluorescence arises from MC chromophore. SP form is non-fluorescent.\nThe principle of design is to compare the lifetime of the initial MC state to that of the\nreverted MC state (after SP-to-MC isomerization). We speculated that fluorescence\nlifetime change is a reporter of the local microenvironment at the engineered protein\nsurface that associated with global hydration HBN rearrangement and conformational\nchanges.\n(11) P5, second paragraph, description of fig. 1g: The fluorescence of ALP-MC and\nALP-RMC are shown to differ. How does from this observation follow the conclusion\nthat ‘isomerization specifically alters the local microenvironment’? Aren’t ALP-\nMC and ALP-RMC in the same isomeric state ?\nReply: We agree with this reviewer that both ALP-MC and ALP-RMC are same MC\nisomeric state (RMC is regenerated via MC→SP→MC photoisomerization). However,\nthe critical difference lies not in their final chemical state, but in their history of\nformation.\nALP-MC: MC bound to the engineered ALP surface. It reflects the “native”\nmicroenvironment of binding site, where the hydration shell, local protein conformation,\nand HBNs are “stable”.\nALP-RMC: MC regenerated in situ, following a two-step transition of surface-\nconjugated MC: first, 450 nm irradiation induces photoisomerization of open MC to\nclose SP, and subsequent dark-state reversion restores the open MC form.\nThe close to open switch imposes transient local perturbation on neighboring\nprotein side chains and the first hydration shell, triggering long-lived rearrangement of\nlocal protein conformation and/or water HBN. Consequently, although ALP-RMC and\nALP-MC are chemically identical (open MC) structure, they locate in distinct\nmicroenvironment. Therefore, we speculated that the significant difference in\nfluorescence lifetime between ALP-MC and ALP-RMC (p<0.01, Fig. 1g) arises\nspecifically from the in-situ MC→SP→MC photoisomerization process on the protein\nsurface.\nThis speculation is confirmed by two key observations: (1) control experiment\nwith free MC and RMC showing no lifetime difference rigorously, thus excludes\ninherent photophysical changes of the probe itself (Fig. 1g, S15); (2) the well-\nestablished sensitivity of MC’s fluorescence lifetime to local polarity and viscosity\n(Figs. S12-S14) confirms its role as a reliable reporter of the local microenvironment6-\n8. Therefore, the observed lifetime shift reflects isomerization-triggered rearrangements\nof protein microenvironment, including hydration shell reordering and local\nconformational changes.\nRevised text: “We then investigated the photoisomerization-induced changes of\nmicroenvironments on the engineered protein surface by measuring the fluorescence\nlifetime of MC. As a sensitive reporter for the heterogeneous protein surface\nmicroenvironment, MC exhibits site-dependent fluorescence lifetimes that reflect\nvariations in local hydration polarity and viscosity (Figs. S12-S14). The fluorescence\nlifetimes of ALP-MC and its revertant state (RMC, regenerated by photoisomerization\nfrom SP to MC) were recorded by time-correlated single photon counting (TCSPC)24\n(Fig. 1f). Statistically differences in fluorescence lifetime were observed between ALP-\nMC and ALP-RMC (p < 0.01) (Fig. 1g), In contrast, negligible variations occurred\nbetween free MC and RMC (Fig. 1g, Fig. S15). This indicates that MC-SP isomerization\nspecifically triggers rearrangements of the hydration shell that possibly leads to\nchanges of protein conformation.” (Page 5, line 12-21, in red)\n12) P7, 2nd paragraph, l 5: ‘… direct H-bonding with MC …’. It is not clear which\ngroup/molecule is H-bonding with MC Water?\nReply: The H-bonding refers to water molecules forming direct hydrogen bonds with\nthe polar groups of MC moiety (e.g., the phenolate O and N), rather with protein. This\nhas been clarified in the revised text.\nRevised text: \"...presumably due to a denser, direct HBs between water and the\npolar groups of the MC probe.\" (Page 7, line 12-13, in red)\n13) P7, 2nd paragraph, l 11: ’15 Å from the modified site’; How is this site defined;\nthe point of attachment at the protein surface? Or the very end of the tag?\nReply: The “modified site” is defined as the covalent attachment point on the protein\nsurface (the C atom of the pAzF residue), not the distal end of the molecular tag. This\nα\nensures a consistent distance metric across all analyses, and the text has been clarified\nin the revised manuscript.\nRevised text: “Using the crystal structure of ALP (PDB:1ED9) as a guide22,\nspecific residues in the loop region, located at varying distances from the catalytic site,\nwere replaced with the unnatural amino acid p-azido-L-phenylalanine (pAzF). The\nselected sites were H872 (~5 Å from the catalytic site), V599 (~10 Å), E906 (~15 Å),\nand K685 (~25 Å) (Fig. S3-S5)”. (Page 4, line 28, in red)\n40\n(14) P7, 3rd paragraph, l 4: it is unclear what is hydrogen bonding with what. Something\nseems to be missing in the sentence, the ‘modified residue’ does not connect.\nReply: We have revised the sentence to clarify two key points. First, the hydrogen\nbonds here are ‘water-water hydrogen bonds’; second, the ‘successive hydration layers’\nare defined as the layers ‘around the modified residue, the distal catalytic site, and the\nglobal ALP surface’ to establish a clear connection between hydration layers and the\nmodified residue.\nTo clarify, we have revised the relevant sentences in the manuscript as follows:\n“To define the spatial range of residue hydrophobicity effects on global water\ndynamics, we computed the lifetime (τ ) of water-water hydrogen bonds within\nHB\nsuccessive hydration layers (1st: < 4 Å, 2nd: 4-6 Å, 3rd: 6-10 Å) around the modified\nresidue29,30, distal catalytic site, and global ALP surface, respectively (Figs. 2e-g, Fig.\nS18).” (Page 7, line26-28, in red)\n(15) P9, third-to-last line: what is meant with ‘initial disruption?\nReply: “Initial disruption” refers to the local perturbation of HBN and hydration shell\nat the modified residue, which is directly triggered by MC to SP photoisomerization.\nAs detailed in the preceding paragraphs and Figure 2, this photoisomerization\nswitches the probe from MC to SP state. This switch acts as the “initial disruption” by\ndramatically altering the local polarity, and then recruiting (MC) or expelling (SP) water\nmolecules in the first hydration shell. Consequently, disrupting the existing local water\nHBN.\nRevised text: \"...the local photoisomerization-induced HBN disruption at the\nmodified residue triggers a long-range reorganization...\" (Page 9, line 25, in red)\n(16) P11, l 1: Please define ATR-THz\nReply: The term was defined in the revised manuscript as follows:\nRevised text: “We next employed in situ Attenuated Total Reflection Terahertz\n(ATR-THz) spectroscopy to probe dynamic variations in protein hydration.” (Page 11,\nline 1, in red)\n41\n17) P11, second paragraph: How is fully hydrated (100 %) defined ? To connect with\nthe literature, it would be useful to convert the 50% and 10% hydration samples into g\nwater / g protein and indicate to how many hydration layers each value corresponds. In\nthe end of p 11, the definition is vague: ‘(50% and 10%), containing 1-2 hydration\nlayers waters.’\nReply: We thank the reviewer for raising this important point. In response, we have\nrevised the manuscript to define all hydration levels quantitatively using the standard\nmetric of grams of water per gram of protein (H O/protein, g/g) and to relate these\n2\nvalues to hydration-layer coverage commonly reported in the literature.\nSpecifically, fully hydrated (100%) corresponds to 0.4 g H O/g protein, where the\n2\nfirst hydration shell is saturated with incipient bulk-like water9. The intermediate\nhydration level (50%) corresponds to ~0.2 g H O/g protein, extending beyond the first\n2\nhydration shell and partially populating the second. The lowest hydration level (10%)\ncorresponds to ~0.04 g H O/g protein, which completes the first hydration shell and is\n2\ngenerally regarded as the critical hydration required to maintain native protein\nstructure10. The ambiguous description of “1-2 hydration layers” on page 11 has been\nreplaced with these quantitative definitions in the revised manuscript.\nRevised text：“To minimize bulk water interference and extrude the reorganization of\nhydration water, THz spectra were measured across distinct hydration levels: fully\nhydrated (0.4 g H O/g protein, ~100%), where the first hydration shell is saturated and\n2\nbulk-like water emerges; partially dehydrated (0.2 g H O/g protein, ~50%), which\n2\nextends into the second hydration layer; and critically hydrated (0.04 g H O/g protein,\n2\n~10%), which predominantly completes the first hydration shell and is essential for\nmaintaining native protein structural integrity (Fig. 3a-c).”(Page 11, line 12-17，in\nred)\n“However, at reduced hydration level (50% and 10%), both modes exhibit larger\nblue-shifts after photoisomerization to ALP-SP with ν shifted from 586 to 612 cm-¹\nLib\n(50%) and 584 to 614 cm-¹ (10%), while ν shiftedfrom 170 to 195 cm-¹ (50%) and\nHB\n42\n165 to 196 cm-¹ (10%).” (Page 11, line 25-28，in red)\n18) P12, 1st paragraph: what is meant by ‘converting wrap- and bound-water’ ?\nReply: We thank the reviewer for this question. The phrase describes a light-induced\nreorganization in the hydration layer, driven by the increased hydrophobicity of ALP-\nSP.\nAccording to established protein hydration terminology11, 12,13, “wrap-water”\ndenotes mobile water that loosely associated with hydrophobic surfaces, while “bound-\nwater” refers to water molecules that tightly hydrogen-bonded to polar/charged groups,\nforming a more ordered network.\nThe MC-to-SP photoisomerization reduces the population of mobile wrap-water,\npromoting a conversion toward a more ordered, bound-water state. This reorganization\nis directly supported by the blue-shifted THz librational and hydrogen-bond stretching\nmodes, which indicate water rigidification and enhanced H-bond network cooperativity.\nRevised text: “To delineate the thermodynamic basis of light-induced hydration\nrearrangements between ALP-MC and ALP-SP, we used THz calorimetry to correlate\nspectral changes with entropy (ΔS) and enthalpy (ΔH). Absorption coefficients (α(ν))\nand photoisomerization-induced differential spectra (Δα(ν) , ALP-SP minus ALP-\nSP-MC\nMC) were calculated to isolate hydration-specific alterations (Eqs. 1-3, Fig. S26). The\nΔα(ν) was deconvolved into the hydrophobic “wrap” region (150-200 cm-¹,\nSP-MC\nentropy-linked) and hydrophilic “bound” region (450-600 cm-¹, enthalpy-linked) (Fig.\nS27).” (page 12, line 10-17, in red)\n“Consequently, the THz results demonstrated that a single-residue hydrophobicity\nchange can induce HBN reorganization across global hydration by converting water\nfrom a loosely associated \"wrap-water\" state to a tightly bound, ordered configuration,\nthereby forming a rigid hydration layer.” (page 12, line 23-26, in red)\n(19) P13: what does the hydration level of 10% make it ‘critical’ ?\nReply: The designation of ~10% hydration as “critical” is well-established in the\nliterature and refers to the minimal water content required to maintain the structural\n43\nintegrity and dynamical coupling essential for protein function14-16. At this threshold,\nwater molecules are predominantly confined to the first hydration shell, forming strong,\nspecific interactions (e.g., hydrogen bonds and electrostatic forces) with polar and\ncharged groups on the protein surface. This tightly bound water is structurally vital,\nstabilizing native protein conformation and facilitating the slow, retarded dynamics that\nare strongly coupled to protein surface motions17, 18.\nAs hydration increases beyond this level, water begins to populate the second and\nsubsequent hydration shells, which are weakly bound and exhibit dynamics\nprogressively approaching those of bulk water. The ~10% level thus marks a critical\ntransition: it ensures the first hydration shell is complete, a prerequisite for sustaining\nthe anharmonic fluctuations and collective motions necessary for enzymatic activity.\nBelow this critical threshold, both water and protein dynamics become severely\nrestricted, leading to a loss of biological function and risking irreversible structural\nchanges. Therefore, the 10% hydration level in our study represents this functionally\nand dynamically critical hydration state.\nRevised text: “…critically hydrated (0.04 g H O/g protein, ~10%), which\n2\npredominantly completes the first hydration shell and is essential for maintaining native\nprotein structural integrity 9,39(Fig. 3a-c)”. (Page 11, line 15-17, in red)\n(20) P14: The RMSD of backbone atoms in ALP-SP and ALP-MC are computed and\ncompared. How do they compare to those of unmodified ALP-E15? Are there any\nchanges in RMSD that originate from the surface modification and not from the change\nin hydrophobicity upon isomerization?\nReply: We thank the reviewer for raising this essential point, which allows us to clearly\nseparate the effect of the covalent modification from that of the photo-regulated\nhydrophobic switch.\nTo address this concern, we performed additional MD simulations of the\nunmodified parent protein ALP-E15 (carrying only the pAzF mutation; denoted ALP-N )\n3\nunder the same conditions as ALP-MC and ALP-SP.\nBackbone RMSD analysis reveals no significant differences among ALP-N , ALP-\n3\nthat covalent modification does not perturb the overall structural framework. In contrast,\nthe RMSF profiles show almost identical patterns for ALP-N and ALP-MC, while\nrearrangement driven by its increased hydrophobicity (Fig. S35). The merged\nsecondary structure maps further confirmed that all three systems maintain an identical\nsecondary structure composition.\nTogether, these results establish that the conformational changes observed in our\nstudy arise from the light-controlled hydrophobicity switch (MC → SP), rather than\nfrom the static presence of the surface modification.\nRevised text: “Importantly, control simulations of the ALP-N mutant confirmed\n3\nthat the overall protein fold remains unperturbed by covalent modification alone,\nwhereas the light-induced hydrophobic switch selectively enhances local flexibility and\nremodels the hydration shell (Fig. S34-S35).” (Page 15, line 19-23, in red)\nFig. S34 Backbone RMSD evolution during MD simulation relative to the initial\nstructure for ALP-N (grey), ALP-MC (fuchsia) and ALP-SP (blue).\n3\nsquare fluctuation (RMSF) profiles of the backbone atoms across the protein sequence\nfor ALP-N , ALP-MC and ALP-SP. Shaded areas indicate mean ± s.d. from four\n3\nindependent simulations; solid circles denote residues where the mean RMSF\ndifference between the compared states exceeds 0.2 Å. (b, d, f) Structural superposition\nof the corresponding states shown in (a, c, e), with ALP-N in grey, ALP-MC in fuchsia\n3\nand ALP-SP in blue.\n21) P14, fourth-to-last-line: Control experiments with unmodified ALP-E15 are\ndescribed. Why do the authors refer to photoisomerization for this sample?\nReply: We thank the reviewer for this critical observation.\nWe agree that our phrasing was not precise. The control experiments with\nunmodified ALP-E15 were designed to test whether free MC/SP molecules in solution\ncould influence enzyme activity via non-covalent interactions. The absence of\nsignificant changes in this control confirmed that the pronounced effects observed in\nthe ALP-SP conjugate do not stem from direct interactions with the photo isomers.\nInstead, it supports our conclusion that the effect is mediated by a local perturbation of\nthe hydration shell, which requires the precise spatial control afforded by covalent\nattachment of the photo isomers. We have amended the text to reflect this logic.\nperformed a control experiment in which unmodified ALP was mixed with SP.\nIllumination of this mixture resulted in negligible changes in enzymatic activity. This is\nin stark contrast to the significant effects observed with covalently conjugated ALP-SP,\nconfirming that functional modulation arises from local perturbations of the hydration\nshell enabled by site-specific attachment rather than direct interactions with free\nphotoisomers (Fig. S41)” (Page 18, line 1-7, in red)\n22) P18, last line: ‘Our findings reveal a hydration heterogeneity in governing protein\ndynamics, …’ What does this mean?\nReply: We thank the reviewer for raising this point. This statement encapsulates our\ncentral finding that hydration water is not a passive solvent, but rather an active and\nstructurally heterogeneous mediator of protein dynamics.\nIn this work, “hydration heterogeneity” refers to heterogeneous structural and\ndynamic properties of water molecules at the protein-water interface, arising from the\nchemical and topological heterogeneity of the protein surface. Our results show that\nthese interfacial waters sense and encode local surface properties—such as the\nhydrophobicity change from MC to SP—and subsequently propagate these local effects\nthrough the hydration network. This leads to hierarchical changes in protein dynamics,\nstarting from altered residue-specific hydration, propagating to intermediate loop\nmotions, and ultimately affecting global functional states. Thus, protein function is\nregulated not only by its structure but also by the physical chemistry of its hydration\nlayer.\n(1) van der Post, S. T.; Tielrooij, K. J.; Hunger, J.; Backus, E. H.; Bakker, H. J. Femtosecond study of the\neffects of ions and hydrophobes on the dynamics of water. Faraday Discuss 2013, 160, 171-189;\ndiscussion 207-124.\n(2) Chakraborty, S.; Bhattacharya, I.; Mitra, R. K. Solvation plays a key role in antioxidant-mediated\nattenuation of elevated creatinine level: An in vitro spectroscopic investigation. J Phys Chem B 2023,\n127 (40), 8576-8585.\n(3) Sega, M.; Schroder, C. Dielectric and terahertz spectroscopy of polarizable and nonpolarizable water\nmodels: A comparative study. J Phys Chem A 2015, 119 (9), 1539-1547.\n(4) Sappl, M.; Szabadi, A.; Honegger, P.; Konig, F.; Steinhauser, O.; Schroder, C. Insights from the\nabsorption coefficient for the development of polarizable (multipole) force fields. Molecules 2025, 30\n(14).\n(5) Qin, Y. Z.; Wang, L. J.; Zhong, D. P. Dynamics and mechanism of ultrafast water-protein interactions.\nP Natl Acad Sci USA 2016, 113 (30), 8424-8429.\n(6) Bai, Y. L.; Huang, Y. A.; Wan, W.; Jin, W. H.; Shen, D.; Lyu, H. C.; Zeng, L. G.; Liu, Y. Derivatizing\nmerocyanine dyes to balance their polarity and viscosity sensitivities for protein aggregation detection.\nChem Commun 2021, 57 (98), 13313-13316.\n(7) Mandal, D.; Pal, S. K.; Sukul, D.; Bhattacharyya, K. Photophysical processes of merocyanine 540 in\nsolutions and in organized media. J Phys Chem A 1999, 103 (41), 8156-8159.\n(8) MacNevin, C. J.; Gremyachinskiy, D.; Hsu, C. W.; Li, L.; Rougie, M.; Davis, T. T.; Hahn, K. M.\nEnvironment-sensing merocyanine dyes for live cell imaging applications. Bioconjugate Chem 2013, 24\n(2), 215-223.\n(9) Tompa, K.; Bánki, P.; Bokor, M.; Kamasa, P.; György, L.; Tompa, P. Interfacial water at protein surfaces:\nWide-line nmr and dsc characterization of hydration in ubiquitin solutions. Biophys J 2009, 96 (7), 2789-\n2798.\n(10) Sirotkin, V. A.; Khadiullina, A. V. Hydration of proteins: Excess partial enthalpies of water and\nproteins. J Phys Chem B 2011, 115 (50), 15110-15118.\n(11) Chandler, D. Interfaces and the driving force of hydrophobic assembly. Nature 2005, 437 (7059),\n640-647.\n(12) Das Mahanta, D.; Brown, D. R.; Pezzotti, S.; Han, S. G.; Schwaab, G.; Shell, M. S.; Havenith, M.\nLocal\nsolvation structures govern the mixing thermodynamics of glycerol-water solutions. Chem Sci 2023, 14\n(26), 7381-7392.\n(13) Pezzotti, S.; Sebastiani, F.; van Dam, E. P.; Ramos, S.; Nibali, V. C.; Schwaab, G.; Havenith, M.\nSpectroscopic fingerprints of cavity formation and solute insertion as a measure of hydration entropic\nloss and enthalpic gain. Angew Chem Int Edit 2022, 61 (29).\n(14) Frauenfelder, H.; Fenimore, P. W.; McMahon, B. H. Hydration, slaving and protein function. Biophys\nChem 2002, 98 (1-2), 35-48.\n(15) Hishida, M.; Kaneko, A.; Yamamura, Y.; Saito, K. Contrasting changes in strongly and weakly bound\nhydration water of a protein upon denaturation. J Phys Chem B 2023, 127 (28), 6296-6305.\n(16) Wang, S.; Gao, J.; Chu, X. K. Residue-specific structural and dynamical coupling of protein and\nhydration water revealed by molecular dynamics simulations. Biomolecules 2025, 15 (5).\n(17) Wallace, V. P.; Ferachou, D.; Ke, P.; Day, K.; Uddin, S.; Casas-Finet, J.; Van Der Walle, C. F.; Falconer,\nR. J.; Zeitler, J. A. Modulation of the hydration water around monoclonal antibodies on addition of\nexcipients detected by terahertz time-domain spectroscopy. J Pharm Sci-Us 2015, 104 (12), 4025-4033.\n(18) Born, B.; Weingärtner, H.; Bründermann, E.; Havenith, M. Solvation dynamics of model peptides\nprobed by terahertz spectroscopy. Observation of the onset of collective network motions. J Am Chem\nSoc 2009, 131 (10), 3752-3755.\n(1) The authors clearly indicate the number of replicates in the captions of figures, but\nto help with clarity I suggest also adding this information to appropriate points of the\nMethods (e.g., indicating in the methods that there were four independent simulations,\npresumably initiated from unique initial configurations etc).\nReply: We thank the reviewer for this helpful suggestion. We have now explicitly\nindicated the number of independent replicates or simulations in the Methods, in\naddition to the figure captions, and clarified that each was initiated from distinct initial\nconfigurations or separately prepared samples to ensure robustness and reproducibility.\nRevised text: “To ensure statistical robustness, four independent molecular\ndynamics simulations were performed, each initiated from a distinct initial\nconfiguration.” (Page 26, Lines 41-43, in red)\n“All measurements were performed in six independent experimental replicates, each\nprepared from separately diluted samples and measured on independently prepared\ncoverslips.” (Page 24, Lines 40-42, in red)\n“All THz measurements were performed in three independent experimental replicates,\neach prepared from independently hydrated protein films and measured separately.”\n(Page 25, Lines 14-16, in red)\n“Each sample was measured in three independent replicates using separately prepared\naliquots across multiple beamtime sessions to ensure statistical robustness.” (Page 26,\nLines 12-14, in red)\n“Each measurement was performed in three independent experimental replicates, using\nseparately prepared ALP samples.” (Page 28, Lines 24-25, in red)\n“All kinetic measurements were performed in three independent replicates, each\ninitiated from separately prepared reaction mixtures. k and K values were obtained\ncat M\nby fitting the Michaelis-Menten equation.” (Page 28, Lines 37-38, in red)\n(2) I am a little confused by the exact water force field that was used after reviewing\nthe response to Reviewer 2. The authors refer to a SPCE flexible force field, which I\ntook to be SPC/fw, but the response suggests that they used q-SPC/fw. Please clarify\n1\nthe exact force field used in the Methods. I suggest also using the conventional literature\nterm for it as opposed to SPCE flexible force field to align with the names in revised\nFigures S29/S30.\nReply: We thank the reviewer for pointing out this ambiguity and apologize for the\nconfusion caused by the inconsistent terminology in the previous version.\nWe clarify that the flexible fixed-charge qSPC/Fw water model was used\nconsistently throughout all molecular dynamic simulations. The term “SPCE flexible\nforce field” has been replaced by conventional designation qSPC/Fw (J. Chem. Phys.\n2006, 125, 184507), in agreement with the nomenclature used in original Figures S29\nand S30.\nThe Methods section and the corresponding discussion in the main text have been\nrevised to ensure a clear and consistent description of the water model employed.\nRevised text: “The employed qSPC/Fw water model exhibits a known force-field\nlimitation that precludes the capture of the 200 cm-¹ peak.” (Page 13, Lines 6-7, in\nred)\n“A periodic cubic box was constructed, and it was ensured that the distance\nbetween the box edges and the protein atoms was greater than 15 Å. The system was\nthen solvated using the flexible fixed-charge qSPC/Fw water model45.” (Page 26, Lines\n34-35, in red)\n(3) Fig. R2 of the rebuttal, which is presented in response to a comment to Reviewer 2,\nis (to me) a helpful depiction of the area occupied by MC/SP. It appears that this figure\nwas only created for the response document—I suggest also adding it to the revised SI.\nReply: We thank the reviewer for this helpful suggestion. In response, we have\nincorporated Fig. R2 into the Supplementary Information and added a corresponding\ndiscussion to the main text to describe the surface area occupied by the MC/SP\nmodification.\nRevised text: “Further quantitative analysis of the solvent-accessible surface area\n(SASA) indicated that the modified MC/SP accounts for ~4% of the total accessible\nsurface (Fig. S12).” (Page 5, Lines 11-13, in red)\nmolecules vs. total protein surface area. Molecular dynamics (MD) simulations were\nused to quantify this proportion, revealing the contribution of modified MC (a) and SP\n(b) molecules to the protein’s surface properties.\nI have reviewed the revised manuscript and the authors response to my questions. The\nauthors have satisfactorily addressed all issues raised in my original review. The\nrevisions are thorough, and the manuscript is now ready for publication.\nWe thank the reviewer for all the comments and suggestions that help us improve\nour work.\nThe authors adequately addressed all issues this referee raised, except for one,\nwhich I think is incorrectly addressed and needs to be corrected and clarified before\npublication. In particular, the relation between a given hydration level in g water / g\nprotein and the number of hydration layers seems to be off by an order of magnitude.\nFor a medium sized protein (like myoglobin, maltose binding protein, RNAse), 0.4 g\nwater / g protein is typically referred to as corresponding to one complete first hydration\nlayer [1–3]. A complete first hydration layer is generally necessary and sufficient for a\nprotein to be fully active. Below about 0.2 g water / g protein, myoglobin has been\nshown not to be active any more [4].\nThe following text in the revision thus needs to be revised again:\n‘THz spectra were measured across distinct hydration levels: fully hydrated (0.4g\nH O/g protein, ~100%), where the first hydration shell is saturated and bulk-like water\n2\nemerges; partially dehydrated (0.2 g H O/g protein, ~50%), which extends into the\n2\nsecond hydration layer; and critically hydrated (0.04 g H O/g protein, ~10%), which\n2\npredominantly completes the first hydration shell and is essential for maintaining native\nprotein structural integrity9,39 (Fig. 3a-c).’\nIn particular, it is not correct to state that a hydration level of 0.04 g H O/g protein\n2\ncorresponds to a critical hydration level with a complete first hydration shell. To the\ncontrary, the literature points to insufficient and inactive proteins at that hydration level.\nFurthermore, the 10%-level is inconsistently defined in the main text and the SI, as in\nthe SI it is written that ‘The resulting films retained 0.1-0.15 g H₂O/g protein,\ncorresponding to ca. ~10% water content.\nOverall, it still remains unclear how the percentage levels are defined. What does\n100% correspond to? Why not skip the percentage level and only state the three\nmeasured hydration levels in g H₂O/g protein ?\nIt is crucial to consistently define the hydration levels and to translate them into\nhydration-layer coverage that is consistent with values and definitions given in the\nliterature.\n5\n[1] Rupley, J.A., and G. Careri. 1991. 'Protein Hydration and Function.' in Anfinsen\nC.B., Edsall J.T., Richards F.M. and Eisenberg D.S. (eds.), Advances in Protein\nChemistry (Academic Press: New York).\n[2] Bon, C., M. S. Lehmann, and C. Wilkinson. 1999. 'Quasi-Laue neutron-diffraction\nstudy of the water arrangement in crystals of triclinic hen egg-white lysozyme', Acta\nCrystallogr D Biol Crystallogr, 55: 978-87.\n[3] Fichou, Y., M. Heyden, G. Zaccai, M. Weik, and D. J. Tobias. 2015. 'Molecular\nDynamics Simulations of a Powder Model of the Intrinsically Disordered Protein Tau',\nJ Phys Chem B, 119: 12580-9.\n[4] Kleinert, T., W. Doster, F. Post, and M. Settles. 1993. \"Hydration effects on protein\nfunction: the kinetics of ligand binding to myoglobin.\" In Proceedings of the Italian\nPhysical Society, edited by M.U. Palma, M.B. Palma-Vittorelli and F. Parak, 127-30.\nBologna.\nReply: We thank the referee for figuring this issue. We agree that our previous revision\ncontained an incorrect conversion between hydration level (g H₂O/g protein) and\nhydration-layer coverage, leading to misleading statements regarding to “critical\nhydration”.\nIn particular, we acknowledge that hydration levels on the order of 0.04 g H₂O/g\nprotein are far below completion of the first hydration shell, and are corresponding to\ninsufficient hydration and loss of protein activity, in agreement with Refs. 1-4 cited by\nthe referee. All such interpretations have been removed.\nTo establish a quantitative evaluation of hydration assignments, we first evaluated\nthe hydration-shell coverage using MD simulations of ALP. Specifically, we quantified\nthe amount of water associated with successive hydration shells, defined as 0-4 Å, 0-6\nÅ, and 0-9 Å from the protein surface.\nThe simulations yield water-to-protein mass ratios of approximately 0.51, 0.83,\nand 1.36 g H₂O /g protein for saturation of 1st, 1st + 2nd, 1st + 2nd + 3rd, respectively (Fig.\nS24, Table S1). These values are in good agreement with the commonly cited literature\nrange of ~0.4-0.5 g H₂O/g protein for completion of the first hydration shell in medium-\n6\nsized globular proteins, with modest system-specific variation arising from protein size,\nsurface chemistry, and topology.\nTable S1. Experimental hydration levels and corresponding hydration shell coverage\nderived from molecular dynamics simulations.\nSimulated Experimental\nHydration shell Experimental Experimental\nMass ratio hydration level\nregime Sample condition hydration shell coverage\n(g H₂O / g protein) (g H₂O / g protein)\n1st 0.51 ± 0.005 Vacuum-Dried Film ~ 0.2 Partial 1st shell\n1st + 2nd 0.83 ± 0.008 +0.5 μL buffer ~ 0.7 Fully 1st + partial 2nd shell\n1st + 2nd + 3rd 1.36 ± 0.012 +1 μL buffer ~ 1.2 Nearly 1st - 3rd shell\nThe absolute hydration levels of the protein films were quantified using ATR-THz\nspectroscopy. Protein films were prepared by depositing 10 µL of concentrated ALP-\nMC solution (100 mg mL-1 in 2.5 mM Tris, 10 mM NaCl, pH 7.5) onto an IR diamond\nwindow and drying to yield a protein mass of 1 mg. Extended vacuum drying (~ 1 mbar,\n20 min) produced a spectroscopically dry reference state, characterized by a negligible\nabsorption feature at 550 cm-1.\nControlled rehydration was then performed by adding defined volumes of Tris-\nNaCl buffer (0.15, 0.5, and 1.0 µL), yielding well-defined hydration levels of 0.15,\n0.2,\nand 1.5 g H₂O / g protein, respectively (Fig. S25a). These reference states enabled\ncalibration of characteristic THz absorption features as a function of known water\ncontent. By matching the spectra of mildly vacuum-dried films (~ 3 mbar, 1 min) to this\ncalibration curve (Fig. S25b), their residual hydration was determined to be ~0.2 g H₂O\n/ g protein. Using the same calibration, buffer additions of 0.5 µL and 1.0 µL correspond\nto hydration levels of approximately 0.7 and 1.2 g H₂O / g protein, respectively.\nOn this basis, the experimentally accessed hydration states can be consistently\nassigned as follows:\n(i) ~ 0.2 g H₂O/g protein corresponds to a discontinuous sub-monolayer hydration\nregime, well below completion of the first hydration shell1-3;\n7\n(ii) ~ 0.7 g H₂O/g protein corresponds to full coverage of the first hydration shell with\npartial population of the second;\n(iii) ~ 1.2 g H₂O/g protein approaches saturation of the first two shells and substantially\npopulates the third hydration layer.\nTo avoid ambiguity, we have removed percentage-based hydration descriptors\nfrom both the main text and the Supplementary Information. All hydration levels are\nnow reported exclusively in absolute units (g H₂O/g protein) and explicitly linked to\nhydration-layer coverage in a manner consistent with established definitions in the\nliterature. Fig. 3 and its caption have been revised accordingly, and Figs. S25 and S26\nhave been added to the Supplementary Information.\nRevised text: “To minimize bulk water interference and extrude the reorganization of\nhydration water, THz spectra were measured across distinct hydration levels: low\nhydration state (~ 0.2 g H₂O/g protein), corresponding to a discontinuous sub-\nmonolayer hydration regime on the protein surface; moderate hydration state (~ 0.7 g\nH₂O/g protein), which fully saturates the first hydration shell and partially populates\nthe second shell; high hydration state (~ 1.2 g H₂O/g protein), approaching completion\nof the first three hydration shells9,39(Fig. 3a-c and Fig. S25-S26, Table S1).” (Page 11,\nLines 12-17, in red)\n“However, at reduced hydration level (~ 0.7 and 0.2 g/g), both modes exhibit larger\nblue-shifts after photoisomerization to ALP-SP with ν shifted from 586 to 612 cm-¹ (~\nLib\n0.7 g/g) and 584 to 614 cm-¹ (~ 0.2 g/g), while ν shiftedfrom 170 to 195 cm-¹ (~ 0.7\nHB\ng/g) and 165 to 196 cm-¹ (~ 0.2 g/g).” (Page 11, Lines 25-28, in red)\nmolecular dynamics simulations. The mass of water per gram of protein (g H₂O / g\nprotein) within the 1st, 1st + 2nd, and 1st + 2nd + 3rd hydration shells was quantified via\nmolecular dynamics simulations of ALP-MC (a) and ALP-SP (b). Data are presented as\nmean ± SD (n = 4).\nFig. S25 Quantification of water content in protein hydration films by ATR-THz\nspectroscopy. (a) ATR-THz absorption spectra of protein films after complete vacuum\ndrying (1 mbar, 20 min) and following addition of defined buffer volumes (0.15, 0.2\nand 1.5 μL). (b) Comparison with hydration films prepared under mild vacuum\nconditions (3 mbar, 1 min), enabling estimation of water content (~ 0.2 g/g for vacuum-\ndried films and ~ 1.2 g/g after addition of 1.0 μL buffer). Shaded regions represent the\nstandard deviation from three independent experiments.\nReference\n(1) Rupley, J. A.; Careri, G. Protein hydration and function. Adv Protein Chem 1991,\n41, 37-172.\n(2) Bon, C.; Lehmann, M. S.; Wilkinson, C. Quasi-laue neutron-diffraction study of the\nwater arrangement in crystals of triclinic hen egg-white lysozyme. Acta Crystallogr D\nBiol Crystallogr 1999, 55 (Pt 5), 978-987.\n(3) Kuntz, I. D., Jr.; Kauzmann, W. Hydration of proteins and polypeptides. Adv Protein\nChem 1974, 28, 239-345.\n9"}, {"round": 2, "reviewer_comments": {"reviewer_1": "In this revision, the authors have carefully addressed my main prior points related to statistical significance through appropriate replicate simulations and statistical analysis, which (for the most part; see below) is clearly explained. The authors’ response to my comments and those of the other reviewers have raised a few remaining minor points that require clarification, but otherwise I believe the manuscript is substantially improved. Remaining points are:\n(1) The authors clearly indicate the number of replicates in the captions of figures, but to help with clarity I suggest also adding this information to appropriate points of the Methods (e.g., indicating in the methods that there were four independent simulations, presumably initiated from unique initial configurations etc).\n(2) I am a little confused by the exact water force field that was used after reviewing the response to Reviewer 2. The authors refer to a SPCE flexible force field, which I took to be SPC/fw, but the response suggests that they used q-SPC/fw. Please clarify the exact force field used in the Methods. I suggest also using the conventional literature term for it as opposed to SPCE flexible force field to align with the names in revised Figures S29/S30.\n(3) Fig. R2 of the rebuttal, which is presented in response to a comment to Reviewer 2, is (to me) a helpful depiction of the area occupied by MC/SP. It appears that this figure was only created for the response document – I suggest also adding it to the revised SI.", "reviewer_2": "I have reviewed the revised manuscript and the authors response to my questions. The authors have satisfactorily addressed all issues raised in my original review. The revisions are thorough, and the manuscript is now ready for publication.", "reviewer_3": "Liu and co-workers carefully revised their manuscript, responding to the many detailed points raised by three referees.\nImportantly, they have carried out additional simulations and experiments on protein mutants that corroborate their conclusions drawn in the originally submitted manuscript.\nThe authors adequately addressed all issues this referee raised, except for one, which I think is incorrectly addressed and needs to be corrected and clarified before publication. In particular, the relation between a given hydration level in g water / g protein and the number of hydration layers seems to be off by an order of magnitude. For a medium sized protein (like myoglobin, maltose binding protein, RNAse), 0.4 g water / g protein is typically referred to as corresponding to one complete first hydration layer [1 – 3]. A complete first hydration layer is generally necessary and sufficient for a protein to be fully active. Below about 0.2 g water / g protein, myoglobin has been shown not to be active any more [4].\nThe following text in the revision thus needs to be revised again:\n‘THz spectra were measured across distinct hydration levels: fully hydrated (0.4g H2O/g protein, ~100%), where the first hydration shell is saturated and bulk-like water emerges; partially dehydrated (0.2 g H2O/g protein, ~50%), which extends into the second hydration layer; and critically hydrated (0.04 g H2O/g protein, ~10%), which predominantly completes the first hydration shell and is essential for maintaining native protein structural integrity9,39 (Fig. 3a-c).’\nIn particular, it is not correct to state that a hydration level of 0.04 g H2O/g protein corresponds to a critical hydration level with a complete first hydration shell. To the contrary, the literature points to insufficient and inactive proteins at that hydration level.\nFurthermore, the 10%-level is inconsistently defined in the main text and the SI, as in the SI it is written that ‘The resulting films retained 0.1-0.15 g H₂O/g protein, corresponding to ca. ~10% water content.\nOverall, it still remains unclear how the percentage levels are defined. What does 100% correspond to ? Why not skip the percentage level and only state the three measured hydration levels in g H₂O/g protein ?\nIt is crucial to consistently define the hydration levels and to translate them into hydration-layer coverage that is consistent with values and definitions given in the literature.\n[1] Rupley, J.A., and G. Careri. 1991. 'Protein Hydration and Function.' in Anfinsen C.B., Edsall J.T., Richards F.M. and Eisenberg D.S. (eds.), Advances in Protein Chemistry (Academic Press: New York).\n[2] Bon, C., M. S. Lehmann, and C. Wilkinson. 1999. 'Quasi-Laue neutron-diffraction study of the water arrangement in crystals of triclinic hen egg-white lysozyme', Acta Crystallogr D Biol Crystallogr, 55: 978-87.\n[3] Fichou, Y., M. Heyden, G. Zaccai, M. Weik, and D. J. Tobias. 2015. 'Molecular Dynamics Simulations of a Powder Model of the Intrinsically Disordered Protein Tau', J Phys Chem B, 119: 12580-9.\n[4] Kleinert, T., W. Doster, F. Post, and M. Settles. 1993. \"Hydration effects on protein function: the kinetics of ligand binding to myoglobin.\" In Proceedings of the Italian Physical Society, edited by M.U. Palma, M.B. Palma-Vittorelli and F. Parak, 127-30. Bologna."}, "author_response": "Reviewer #1 (Remarks to the Author):\n(1) The authors clearly indicate the number of replicates in the captions of figures, but\nto help with clarity I suggest also adding this information to appropriate points of the\nMethods (e.g., indicating in the methods that there were four independent simulations,\npresumably initiated from unique initial configurations etc).\nReply: We thank the reviewer for this helpful suggestion. We have now explicitly\nindicated the number of independent replicates or simulations in the Methods, in\naddition to the figure captions, and clarified that each was initiated from distinct initial\nconfigurations or separately prepared samples to ensure robustness and reproducibility.\nRevised text: “To ensure statistical robustness, four independent molecular\ndynamics simulations were performed, each initiated from a distinct initial\nconfiguration.” (Page 26, Lines 41-43, in red)\n“All measurements were performed in six independent experimental replicates, each\nprepared from separately diluted samples and measured on independently prepared\ncoverslips.” (Page 24, Lines 40-42, in red)\n“All THz measurements were performed in three independent experimental replicates,\neach prepared from independently hydrated protein films and measured separately.”\n(Page 25, Lines 14-16, in red)\n“Each sample was measured in three independent replicates using separately prepared\naliquots across multiple beamtime sessions to ensure statistical robustness.” (Page 26,\nLines 12-14, in red)\n“Each measurement was performed in three independent experimental replicates, using\nseparately prepared ALP samples.” (Page 28, Lines 24-25, in red)\n“All kinetic measurements were performed in three independent replicates, each\ninitiated from separately prepared reaction mixtures. k and K values were obtained\ncat M\nby fitting the Michaelis-Menten equation.” (Page 28, Lines 37-38, in red)\n(2) I am a little confused by the exact water force field that was used after reviewing\nthe response to Reviewer 2. The authors refer to a SPCE flexible force field, which I\ntook to be SPC/fw, but the response suggests that they used q-SPC/fw. Please clarify\n1\nthe exact force field used in the Methods. I suggest also using the conventional literature\nterm for it as opposed to SPCE flexible force field to align with the names in revised\nFigures S29/S30.\nReply: We thank the reviewer for pointing out this ambiguity and apologize for the\nconfusion caused by the inconsistent terminology in the previous version.\nWe clarify that the flexible fixed-charge qSPC/Fw water model was used\nconsistently throughout all molecular dynamic simulations. The term “SPCE flexible\nforce field” has been replaced by conventional designation qSPC/Fw (J. Chem. Phys.\n2006, 125, 184507), in agreement with the nomenclature used in original Figures S29\nand S30.\nThe Methods section and the corresponding discussion in the main text have been\nrevised to ensure a clear and consistent description of the water model employed.\nRevised text: “The employed qSPC/Fw water model exhibits a known force-field\nlimitation that precludes the capture of the 200 cm-¹ peak.” (Page 13, Lines 6-7, in\nred)\n“A periodic cubic box was constructed, and it was ensured that the distance\nbetween the box edges and the protein atoms was greater than 15 Å. The system was\nthen solvated using the flexible fixed-charge qSPC/Fw water model45.” (Page 26, Lines\n34-35, in red)\n(3) Fig. R2 of the rebuttal, which is presented in response to a comment to Reviewer 2,\nis (to me) a helpful depiction of the area occupied by MC/SP. It appears that this figure\nwas only created for the response document—I suggest also adding it to the revised SI.\nReply: We thank the reviewer for this helpful suggestion. In response, we have\nincorporated Fig. R2 into the Supplementary Information and added a corresponding\ndiscussion to the main text to describe the surface area occupied by the MC/SP\nmodification.\nRevised text: “Further quantitative analysis of the solvent-accessible surface area\n(SASA) indicated that the modified MC/SP accounts for ~4% of the total accessible\nsurface (Fig. S12).” (Page 5, Lines 11-13, in red)\nmolecules vs. total protein surface area. Molecular dynamics (MD) simulations were\nused to quantify this proportion, revealing the contribution of modified MC (a) and SP\n(b) molecules to the protein’s surface properties.\nI have reviewed the revised manuscript and the authors response to my questions. The\nauthors have satisfactorily addressed all issues raised in my original review. The\nrevisions are thorough, and the manuscript is now ready for publication.\nWe thank the reviewer for all the comments and suggestions that help us improve\nour work.\nThe authors adequately addressed all issues this referee raised, except for one,\nwhich I think is incorrectly addressed and needs to be corrected and clarified before\npublication. In particular, the relation between a given hydration level in g water / g\nprotein and the number of hydration layers seems to be off by an order of magnitude.\nFor a medium sized protein (like myoglobin, maltose binding protein, RNAse), 0.4 g\nwater / g protein is typically referred to as corresponding to one complete first hydration\nlayer [1–3]. A complete first hydration layer is generally necessary and sufficient for a\nprotein to be fully active. Below about 0.2 g water / g protein, myoglobin has been\nshown not to be active any more [4].\nThe following text in the revision thus needs to be revised again:\n‘THz spectra were measured across distinct hydration levels: fully hydrated (0.4g\nH O/g protein, ~100%), where the first hydration shell is saturated and bulk-like water\n2\nemerges; partially dehydrated (0.2 g H O/g protein, ~50%), which extends into the\n2\nsecond hydration layer; and critically hydrated (0.04 g H O/g protein, ~10%), which\n2\npredominantly completes the first hydration shell and is essential for maintaining native\nprotein structural integrity9,39 (Fig. 3a-c).’\nIn particular, it is not correct to state that a hydration level of 0.04 g H O/g protein\n2\ncorresponds to a critical hydration level with a complete first hydration shell. To the\ncontrary, the literature points to insufficient and inactive proteins at that hydration level.\nFurthermore, the 10%-level is inconsistently defined in the main text and the SI, as in\nthe SI it is written that ‘The resulting films retained 0.1-0.15 g H₂O/g protein,\ncorresponding to ca. ~10% water content.\nOverall, it still remains unclear how the percentage levels are defined. What does\n100% correspond to? Why not skip the percentage level and only state the three\nmeasured hydration levels in g H₂O/g protein ?\nIt is crucial to consistently define the hydration levels and to translate them into\nhydration-layer coverage that is consistent with values and definitions given in the\nliterature.\n5\n[1] Rupley, J.A., and G. Careri. 1991. 'Protein Hydration and Function.' in Anfinsen\nC.B., Edsall J.T., Richards F.M. and Eisenberg D.S. (eds.), Advances in Protein\nChemistry (Academic Press: New York).\n[2] Bon, C., M. S. Lehmann, and C. Wilkinson. 1999. 'Quasi-Laue neutron-diffraction\nstudy of the water arrangement in crystals of triclinic hen egg-white lysozyme', Acta\nCrystallogr D Biol Crystallogr, 55: 978-87.\n[3] Fichou, Y., M. Heyden, G. Zaccai, M. Weik, and D. J. Tobias. 2015. 'Molecular\nDynamics Simulations of a Powder Model of the Intrinsically Disordered Protein Tau',\nJ Phys Chem B, 119: 12580-9.\n[4] Kleinert, T., W. Doster, F. Post, and M. Settles. 1993. \"Hydration effects on protein\nfunction: the kinetics of ligand binding to myoglobin.\" In Proceedings of the Italian\nPhysical Society, edited by M.U. Palma, M.B. Palma-Vittorelli and F. Parak, 127-30.\nBologna.\nReply: We thank the referee for figuring this issue. We agree that our previous revision\ncontained an incorrect conversion between hydration level (g H₂O/g protein) and\nhydration-layer coverage, leading to misleading statements regarding to “critical\nhydration”.\nIn particular, we acknowledge that hydration levels on the order of 0.04 g H₂O/g\nprotein are far below completion of the first hydration shell, and are corresponding to\ninsufficient hydration and loss of protein activity, in agreement with Refs. 1-4 cited by\nthe referee. All such interpretations have been removed.\nTo establish a quantitative evaluation of hydration assignments, we first evaluated\nthe hydration-shell coverage using MD simulations of ALP. Specifically, we quantified\nthe amount of water associated with successive hydration shells, defined as 0-4 Å, 0-6\nÅ, and 0-9 Å from the protein surface.\nThe simulations yield water-to-protein mass ratios of approximately 0.51, 0.83,\nand 1.36 g H₂O /g protein for saturation of 1st, 1st + 2nd, 1st + 2nd + 3rd, respectively (Fig.\nS24, Table S1). These values are in good agreement with the commonly cited literature\nrange of ~0.4-0.5 g H₂O/g protein for completion of the first hydration shell in medium-\n6\nsized globular proteins, with modest system-specific variation arising from protein size,\nsurface chemistry, and topology.\nTable S1. Experimental hydration levels and corresponding hydration shell coverage\nderived from molecular dynamics simulations.\nSimulated Experimental\nHydration shell Experimental Experimental\nMass ratio hydration level\nregime Sample condition hydration shell coverage\n(g H₂O / g protein) (g H₂O / g protein)\n1st 0.51 ± 0.005 Vacuum-Dried Film ~ 0.2 Partial 1st shell\n1st + 2nd 0.83 ± 0.008 +0.5 μL buffer ~ 0.7 Fully 1st + partial 2nd shell\n1st + 2nd + 3rd 1.36 ± 0.012 +1 μL buffer ~ 1.2 Nearly 1st - 3rd shell\nThe absolute hydration levels of the protein films were quantified using ATR-THz\nspectroscopy. Protein films were prepared by depositing 10 µL of concentrated ALP-\nMC solution (100 mg mL-1 in 2.5 mM Tris, 10 mM NaCl, pH 7.5) onto an IR diamond\nwindow and drying to yield a protein mass of 1 mg. Extended vacuum drying (~ 1 mbar,\n20 min) produced a spectroscopically dry reference state, characterized by a negligible\nabsorption feature at 550 cm-1.\nControlled rehydration was then performed by adding defined volumes of Tris-\nNaCl buffer (0.15, 0.5, and 1.0 µL), yielding well-defined hydration levels of 0.15,\n0.2,\nand 1.5 g H₂O / g protein, respectively (Fig. S25a). These reference states enabled\ncalibration of characteristic THz absorption features as a function of known water\ncontent. By matching the spectra of mildly vacuum-dried films (~ 3 mbar, 1 min) to this\ncalibration curve (Fig. S25b), their residual hydration was determined to be ~0.2 g H₂O\n/ g protein. Using the same calibration, buffer additions of 0.5 µL and 1.0 µL correspond\nto hydration levels of approximately 0.7 and 1.2 g H₂O / g protein, respectively.\nOn this basis, the experimentally accessed hydration states can be consistently\nassigned as follows:\n(i) ~ 0.2 g H₂O/g protein corresponds to a discontinuous sub-monolayer hydration\nregime, well below completion of the first hydration shell1-3;\n7\n(ii) ~ 0.7 g H₂O/g protein corresponds to full coverage of the first hydration shell with\npartial population of the second;\n(iii) ~ 1.2 g H₂O/g protein approaches saturation of the first two shells and substantially\npopulates the third hydration layer.\nTo avoid ambiguity, we have removed percentage-based hydration descriptors\nfrom both the main text and the Supplementary Information. All hydration levels are\nnow reported exclusively in absolute units (g H₂O/g protein) and explicitly linked to\nhydration-layer coverage in a manner consistent with established definitions in the\nliterature. Fig. 3 and its caption have been revised accordingly, and Figs. S25 and S26\nhave been added to the Supplementary Information.\nRevised text: “To minimize bulk water interference and extrude the reorganization of\nhydration water, THz spectra were measured across distinct hydration levels: low\nhydration state (~ 0.2 g H₂O/g protein), corresponding to a discontinuous sub-\nmonolayer hydration regime on the protein surface; moderate hydration state (~ 0.7 g\nH₂O/g protein), which fully saturates the first hydration shell and partially populates\nthe second shell; high hydration state (~ 1.2 g H₂O/g protein), approaching completion\nof the first three hydration shells9,39(Fig. 3a-c and Fig. S25-S26, Table S1).” (Page 11,\nLines 12-17, in red)\n“However, at reduced hydration level (~ 0.7 and 0.2 g/g), both modes exhibit larger\nblue-shifts after photoisomerization to ALP-SP with ν shifted from 586 to 612 cm-¹ (~\nLib\n0.7 g/g) and 584 to 614 cm-¹ (~ 0.2 g/g), while ν shiftedfrom 170 to 195 cm-¹ (~ 0.7\nHB\ng/g) and 165 to 196 cm-¹ (~ 0.2 g/g).” (Page 11, Lines 25-28, in red)\nmolecular dynamics simulations. The mass of water per gram of protein (g H₂O / g\nprotein) within the 1st, 1st + 2nd, and 1st + 2nd + 3rd hydration shells was quantified via\nmolecular dynamics simulations of ALP-MC (a) and ALP-SP (b). Data are presented as\nmean ± SD (n = 4).\nFig. S25 Quantification of water content in protein hydration films by ATR-THz\nspectroscopy. (a) ATR-THz absorption spectra of protein films after complete vacuum\ndrying (1 mbar, 20 min) and following addition of defined buffer volumes (0.15, 0.2\nand 1.5 μL). (b) Comparison with hydration films prepared under mild vacuum\nconditions (3 mbar, 1 min), enabling estimation of water content (~ 0.2 g/g for vacuum-\ndried films and ~ 1.2 g/g after addition of 1.0 μL buffer). Shaded regions represent the\nstandard deviation from three independent experiments.\nReference\n(1) Rupley, J. A.; Careri, G. Protein hydration and function. Adv Protein Chem 1991,\n41, 37-172.\n(2) Bon, C.; Lehmann, M. S.; Wilkinson, C. Quasi-laue neutron-diffraction study of the\nwater arrangement in crystals of triclinic hen egg-white lysozyme. Acta Crystallogr D\nBiol Crystallogr 1999, 55 (Pt 5), 978-987.\n(3) Kuntz, I. D., Jr.; Kauzmann, W. Hydration of proteins and polypeptides. Adv Protein\nChem 1974, 28, 239-345.\n9"}, {"round": 3, "reviewer_comments": {"reviewer_3": "The authors have addressed the remaining issue of hydration levels adequately and now state them consistently and correctly with respect to the vast literature."}, "author_response": ""}]

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