Nitration and Glycation Diminish the Î±-Synuclein Role in the

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Nitration and glycation diminish the #-synuclein role on the formation and scavenging of Cu -catalyzed reactive oxygen species 2+

Humberto Martinez Orozco, Laura Mariño, Ana Belén Uceda, Joaquín Ortega-Castro, Bartolomé Vilanova, Juan Frau, and Miquel Adrover ACS Chem. Neurosci., Just Accepted Manuscript • DOI: 10.1021/acschemneuro.9b00142 • Publication Date (Web): 11 Apr 2019 Downloaded from http://pubs.acs.org on April 12, 2019

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ACS Chemical Neuroscience

Nitration and glycation diminish the synuclein role on the formation and scavenging of Cu2+-catalyzed reactive oxygen species Humberto Martínez-Orozco$, Laura Mariño$, Ana Belén Uceda$, Joaquín Ortega-Castro, Bartolomé Vilanova, Juan Frau and Miquel Adrover*

Institut Universitari d'Investigació en Ciències de la Salut (IUNICS). Institut de Recerca en Ciències de la Salut (IdISBa). Departament de Química, Universitat de les Illes Balears, Ctra. Valldemossa km 7.5, E-07122 Palma de Mallorca, Spain.

$These

authors contributed equally to the work.

*Correspondence to: Miquel Adrover, University of Balearic Islands Ed. Mateu Orfila i Rotger Crta. Valldemossa, km 7.5 E-07122 Palma de Mallorca Illes Balears, Spain. Phone: +34 971 173491; Fax +34 971 173426; e-mail: [email protected]

KEYWORDS -Synuclein – Copper – Parkinson’s disease – Glycation – Nitration – Reactive oxygen species

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ABSTRACT Human -synuclein is a small monomeric protein (140 residues) essential to maintain the function of the dopaminergic neurons and the neuronal redox balance. However, it holds a dark side since it is able to clump inside the neurons forming insoluble aggregates known as Lewy bodies, which are considered the hallmark of Parkinson’s disease. Sporadic mutations and non-enzymatic post-translational modifications are well-known to stimulate the formation of Lewy bodies. Yet, the effect of non-enzymatic post-translational modifications on the function of -synuclein has been studied less intense. Therefore, here we study how nitration and glycation mediated by methylglyoxal affect the redox features of -synuclein. Both diminish the ability of synuclein

to

chelate

Cu2+,

except

when

Nε-(carboxyethyl)lysine

or

N ε-

(carboxymethyl)lysine (two advanced glycation end products highly prevalent in vivo) are formed. This results into a lower capacity to prevent the Cu-catalyzed ascorbic acid degradation and to delay the formation of H2O2. However, only methylglyoxal was able to abolish the ability of -synuclein to inhibit the free radical release. Both nitration and glycation enhanced the -synuclein availability to be damaged by O2·-, although glycation made -synuclein less reactive towards HO·. Our data represent the first report describing how non-enzymatic post-translational modifications might affect the redox function of -synuclein, thus contributing to a better understanding of its pathological implications.


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INTRODUCTION Human -synuclein (S) is a disordered protein mainly found in the neurons of the substantia nigra1 where it plays a key role in membrane trafficking and acts as a chaperone of synaptic SNAREs proteins.2 Endogenous S is also essential to maintain the neuronal redox balance between dopamine oxidation, a prominent source of reactive oxygen species (ROS) and the antioxidant enzyme response.3 In fact, snca -/neurons have an enhanced rate of ROS production and are more susceptible to oxidative4 and glycation5 damage. Therefore, S is able to inhibit apoptosis4 and prevent neurodegeneration.6,7 In contrast, S over-expression induces the production of ROS8,9 through the association of its oligomers to the cellular membrane.10 Soluble oligomers are also responsible for the development of Parkinson’s disease (PD).11 They evolve to amyloid fibrils, which clump into intraneuronal deposits known as Lewy bodies (LBs).12,13 LBs interfere with the trafficking in neurons, disrupt membranes and sequester proteins.14 Their formation, and the aggregation of S, is affected by: i) pathogenic mutations (e.g. A53T, A30P or E46K);15 ii) oxidative stress,16 which initiates the feed-forward cycle of oxidized S-induced stress3 detected in PD;17 iii) Cu/Fe homeostasis imbalance18 and the formation of Cu-S complexes;19,20 and iv) non-enzymatic phosphorylation23

post-translational or

glycation24-27)

modifications detected

on

(PTMs)

(i.e.

post-mortem

nitration,21,22 PD

brains.28-30

Nevertheless, it is still under debate whether PTMs are early or late events in PD pathogenesis, although the link between them is clear.28 Nitration is an irreversible PTM occurring in vivo due to the action of the nitrogen dioxide radical (NO2·) on the Tyr residues. It is found on S of aged brains, but especially in the S of people suffering PD. Consequently, nitrated S is considered as a clinical biomarker for the development of PD, but also for the neuronal vulnerability. The correlation between nitrated S and PD arises from the ability of nitration to alter the biological function of S. Nitration decreases the S binding to membranes, thus affecting the synaptic vesicle assembly. Moreover, nitrated S can activate microglia via the integrin receptor α5β1. This inflammatory response leads to the activation of astrocytes, and both cells form large amounts of ROS that can damage adjacent neurons. Furthermore, nitrated S induces proliferation and activity of specific effector T cells that contribute to the degeneration of dopaminergic neurons. In addition, it is also well known that nitration can stabilize partially folded conformations of S, which results into the inhibition of its fibrillation by stabilizing off-pathway toxic oligomers.22 Non-enzymatic glycosylation (also known as glycation) is another spontaneous agedependent PTM occurring on the protein nucleophilic side chains (mainly Lys and Arg) through their reaction with reducing sugars, or with their autoxidation by-products. It ends with the formation of a heterogeneous set of compounds (AGEs), which modify the physicochemical features of proteins and their function. Diabetes mellitus 3 ACS Paragon Plus Environment


increases 34-fold the accumulation of AGEs in brains, and it is able to accelerate the progression of both motor and cognitive signs in PD. Consequently, it is considered as an important risk factor for PD. In fact, large amounts of AGEs have been detected in the periphery of LB isolated from patients with PD. Hence, they trigger the LB formation as a result of their ability to crosslink monomeric S. In any case, AGEs enhance the toxicity of the S oligomers both in vivo and in vitro. In vivo glycation of S mainly occurs through the action of methylglyoxal (MG), a dicarbonyl metabolite formed during the intraneuronal glycolysis. Their levels are increased 2- to 5-fold in DM patients. MG reduces the ability of S to bind membranes, thus perturbing the physiological role of S on vesicular trafficking. Moreover, MG exacerbates S toxicity in human cell line and in patient-derived iPSCs. Upon knockdown of S in iPSCs, the toxicity of MG was abolished, thus confirming that cytotoxicity is associated with S. Consequently, MG reduces the motor performance of S expressing flies, and stimulate the loss of neuronal cells in mice.24-27 While the effect of nitration and glycation on the aggregation of S has been studied, there are no data reporting their effect on the S capacity to maintain the neuronal redox equilibrium,3,6,7 strongly unbalanced during PD.7,31,32 To get insights on this aspect, we have studied the effect of nitration and glycation (mediated by MG) on the capacity of S to scavenge ROS, and to inhibit their formation from Cu-catalyzed ascorbic acid (AA) degradation.33,34


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RESULTS AND DISCUSSION S is oxidized during the Cu2+-catalyzed AA degradation AA accumulates inside the neurons (~10mM) where it modulates the neurological transmission, improves the bioavailability of levodopa, and scavenges ROS protecting against excitotoxicity.32,35,36 However, under the altered Fe/Cu homeostasis observed in PD,37,38 free and/or aberrantly bound Cu catalyze the oxidation of AA producing ROS (Figure S1).37,39 We observed that recombinant S (Figure S2) decreased the rate of Cu-catalyzed AA degradation (Figure S3A) and diminished the formation of free ROS (Figure S3B), although suffering concomitant oxidations (Figure S4A). These mainly turned Met into Met-sulfoxide (Figure S4B), which essentially occurred on M1 and M5 as proven by their chemical shift perturbations (Figure 1), and those of the residues nearby (i.e. V3, F4, L8, S9). The appearance of weak

15N-HSQC

signals near the S129

and E130 cross-peaks indicated a low degree of oxidation of M127, whereas the peaks of D119 and D121, sensitive to M116 oxidation,40 remained unchanged indicating that M116 is less prone to be oxidized. 1H,1H-TOCSY spectra also evidenced a decrease in the intensity of the Pro Hδ-Hβ cross signals, which would indicate a faintly Cu/AAinduced Pro hydroxylation41 (Figure 1). Chemical shifts of other residues well-known to be sensitive to ROS (e.g. Lys, Phe or His)41,42 did not change during AA degradation (Figure S5). The di-Tyr crosslinking observed in LB from PD brains,43 became only relevant upon increasing the [Cu2+] and/or the [AA] (Figure S4C). In any case, the

15N-

HSQC spectra proved that oxidation had little impact on the averaged S conformation (Figure 1).

Figure 1. Overlay of the 15N-HSQC spectra (A) and the 1H,1H-TOCSY spectra (B) of S (80μM) in presence of AA (280μM) and Cu2+ (2.5μM) before (black) and after 24h of incubation at 25ºC (red). In the 15N-HSQC spectra, the peaks corresponding to M5 and other residues nearby are labelled in blue, the peaks of residues sensitive to M116 oxidation are labelled in black, and those sensitive to M127 oxidation are labelled in green.



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S diminishes the AA degradation rate via a Cu2+ chelating mechanism Since EDTA is also able to reduce the rate of AA degradation (Figure S3A), the formation of Cu-S complexes could be responsible for the diminished catalytic effect of Cu on AA degradation -Cu2+ binds S at V3-S9, at V48-V52 and at D119-A124 regions,19,20,44-47 whereas Cu+ mainly binds at M1, D2 and M5.20,48 Accordingly, when using Glu-C-digested S instead of full length, the rate of AA degradation increased (Table S1 and Figure S6). This can be attributed to the cleavage of the D119-A124 binding site (Figure S6A), but also because S fragments have a lower Cu affinity. Nitration of Y39, Y125, Y133 and Y136 modify the redox features of S The fact that nitration of S rapidly induces the neuron death49,50 led us to study whether this could be -at least partially- due to its effect on S redox capacity. S was incubated with tetranitromethane,51 which yielded monomeric nitrated S (S-NO2). Mass spectrometry (MS) proved the addition of four -NO2 groups on S, forming 3nitrotyrosine (3-NT) on Y39, Y125, Y133 and Y136, as suggested by its UV-Vis profile52 (Figure S7). Table 1. Thermodynamic parameters obtained from ITC experiments for the binding between Cu2+ and the different Ss. The thermodynamic data is obtained from the fitting of the molar enthalpy change versus the [S-X]/[Cu2+] to three different sequential binding sites. All ITC experiments were carried out in duplicate (n=2). Sa

S-NO2

S-MG

S-CML

S-CEL

4.0±0.2 (1.6)b; (0.67)c; (0.20)d; (0.11)e (6.6±0.3)·101 (13.3)b; (5.2)c; (50)d; (35)e

9.1±0.2

(1.9±0.2)·102

(3.7±1.6)·101

(5.3±0.8)·101

(7.7±0.1)·101

(1.5±0.3)·103

(2.4±0.4)·101

(2.5±0.1)·101

(9.9±0.6)·101

(4.0±0.5)·10-3

(6.1±0.5)·10-2

(3.4±0.8)·10-1

ΔHd1 (kcal/mol)

2.1±0.1 (0.006)c; (~1)d -3.2±0.1 (-9.9)c; (-7.7)e

-2.7±0.1

-9.0±0.5

-4.1±0.3

-4.3±0.2

ΔHd2 (kcal/mol) ΔHd3 (kcal/mol)

-2.3±0.4 (-55)c; (-1.1)e -3.7±0.3 (-25)c

-2.3±0.2

(3.5±0.8)·101

-2.9±0.3

-1.5±0.2

(-9.0±2.0)·10-1

-3.3±0.7

-3.1±0.1

-1.1±0.7

Kd1 (µM) Kd2 (µM) Kd3 (mM)

Values in italics correspond to bibliographic data. Ref. [46] cRef. [47] dRef. [44] eRef.[45] a

b

Nitration scarcely reduced the Cu2+/S affinity at the V3-S9 and V48-V52 binding sites (Table 1 and Figure S8A), likely due to the lack of Tyr nearby (Figure S9A). However, its effect on the D119-A124 region was remarkable (Kd3), which can be attributed to changes in the conformational population at the C-terminus induced by nitration of its Tyr.51 Accordingly, S-NO2 displayed a lower inhibitory effect on AA degradation than S (Figure 2A), but it did not enhance the overall formation of free ROS (Figure 2B). Although S does not affect the rate of O2·- production in neurons,53 it would have been interesting to study the effect of nitration on this process. 6 ACS Paragon Plus Environment

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Nonetheless, this was not possible because O2·- rapidly reacted with free AA (Figure S10), thus contributing to the AA autoxidation cycle.54 The rate of H2O2 formation was only slightly higher in presence of S-NO2 than with S (p0.01 between 2-15min), although its presence increased twice the final [H2O2]. This must be due to the ability of the S-MG/Cu2+ complex to direct the AA degradation towards the



formation of H2O2, which does not occur in the absence of Cu2+ (Figures 2C and S16A). The oxidation of H2O2 to form HO· was also inhibited by S-MG, but its potential was lower than that for S (Figures 2D and S16B). In any case, free HO· could not be trapped by S-MG (Figure 3B,E), which indicates that AGEs deplete the capacity of S to scavenge HO· and therefore, modify its ability to act as a neuronal sink of ROS.55 Nonetheless, this does not occur for all ROS since S-MG is a better scavenger of O2·than S (Figure 3F).

Figure 2. Effect of the different Ss on the Cu-catalyzed AA degradation and on the formation of ROS. (A) Time-dependent AA (70μM) degradation at 25ºC measured by the decrease in its absorbance at 265nm in the presence of Cu2+ (2.5μM) alone (●) or in the presence of the different Ss. All these experiments were carried out in triplicate (n=3). (B) Time-dependent overall ROS formation monitored by the decrease in the fluorescence intensity of fluorescein (26μM; λexc 490nm). All the solutions were analyzed at 25ºC and contained AA (70μM) and Cu2+ (2.5μM) alone (●) or in the presence of the different Ss. The experimental data was smoothed using the negative exponential function in Sigmaplot. All these experiments were carried out in duplicate (n=2). (C) Time-dependent formation of H2O2 measured by the increase in resorufin fluorescence at 590nm. The reaction mixtures always contained AA (70μM) and Cu2+ (2.5μM) either alone (●) or in the presence of the different Ss. All these experiments were carried out in triplicate (n=3). (D) Time-dependent formation of HO· measured by the increase in the fluorescence of 3-CCA at 450nm (λexc 395nm). All these experiments were carried out in triplicate (n=3). The reaction mixtures always contained AA (70μM) and Cu2+ (2.5μM) either alone (●) or in the presence of the different Ss. The insert shows a zoom on the region framed in blue dashes. In all the experiments depicted in the panels A, B, C and D we used 10μM S (●), S-NO2 (●), S-MG (●), S-CEL (●) or S-CML (●).


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Figure 3. Effect of different PTMs on the readiness of S to interact with free radicals. (A) MALDI-TOF/TOF signal of S-NO2 incubated during 0min (▬) and 150min (▬) at 25ºC in buffer B1 with AA (70μM) and Cu2+ (2.5μM). (B) MALDI-TOF/TOF signal of S-MG incubated during 0min (▬) and 150min (▬) at 25ºC in buffer B1 with AA (70μM) and Cu2+ (2.5μM). (C) MALDITOF/TOF signal of S-CEL incubated during 0min (▬) and 150min (▬) at 25ºC in buffer B1 with AA (70μM) and Cu2+ (2.5μM). (D) MALDI-TOF/TOF signal of S-CML incubated during 0min (▬) and 150min (▬) at 25ºC in buffer B1 with AA (70μM) and Cu2+ (2.5μM). (E) UV-Vis spectra of the neocuproine-Cu+ complex formed from the hydroxylation of salicylic acid by HO· in the absence (▬) or in the presence of 10μM S (▬), S-NO2 (▬), S-MG (▬), S-CEL (▬) or S-CML (▬). (F) UV-Vis spectra of the formazan formed from NBT oxidation by free O2·- in the absence (▬) or in the presence of 10μM S (▬), S-NO2 (▬), S-MG (▬), S-CEL (▬) or S-CML (▬). All the experiments shown in panels A-E were carried out in duplicate (n=2), whereas those plotted in panel F were carried out in triplicate (n=3).

The formation of EDTA-like AGEs improves the redox protective potential of S The varied set of AGEs formed on S-MG and the heterogeneous glycation degree made impossible to distinguish the effect of each specific AGE on the redox features of S. To overcome this issue, and given that Nε-(carboxyethyl)lysine (CEL) and Nε(carboxymethyl)lysine (CML) have been detected on S deposits,25 we chemically 9 ACS Paragon Plus Environment


modified the 15 Lys of S to obtain homogeneously glycated S monomers with CEL (S-CEL) or with CML (S-CML) (Figures S17 and S18). The formation of CEL/CML on K6, K10 and/or K12 (Figure S15A) slightly affected the Cu2+-affinity at the V3-S9 region (Kd1). Moreover, both are responsible for the increased affinity at the V48-V52 binding region (Kd2) and/or the appearance of new ones (Table 1 and Figure S15B) as a result of their high chelating capacity.59,60 Therefore, CEL/CML notably increased the capacity of S to diminish the rate of the Cu2+-catalyzed AA degradation (Figure 2A). This explains that S-CEL and S-CML considerably slowed down the rate of H2O2 formation (Figure 2C), but only S-CEL was able to modify the final [H2O2], which was not due to a catalytic effect itself (Figure S19). Likewise S, S-CEL and S-CML also inhibited/delayed the formation of HO· (only the final [HO·] marginally increased in presence of S-CML; p=0.036 at 150min) (Figure 2D), although their own oxidative damage (Figure 3C,D) was much lower than that of S (Figure S4A). This can be attributed to a decrease in the capacity of S to trap HO· due to CML or CEL formation (Figure 3E), which must be an indirect effect since Lys seems not to take a direct role on this process (Figure 1B). The redox features of S-CEL and S-CML in comparison with those of S cannot only be attributed to the formation of CEL and CML. This is because N-methylglycine (a CML analogue) and N-methylalanine (a CEL analogue) did not affect the rate of the AA degradation (Figure S20A) and only marginally altered the rate of HO· formation (Figure S20B), while displaying a lower capacity to trap HO· and O2·- than S-CEL or S-CML (Figures S20C,D). Hence, CEL/CML formation at specific points along the S sequence is a key factor to explain the modified redox features of S. In addition, these results prove that S-MG might contain other AGEs different than CML or CEL capable to: i) accelerate the oxidation of AA; ii) catalyze oxidation of AA to form H2O2; and iii) stimulate the conversion of H2O2 to HO·.

Figure 4. Schematic representation of the effect of nitration and glycation on the ability of native S (underlined) to prevent AA degradation (blue), to inhibit the formation of H2O2 and HO· (red) and to scavenge HO· and O2·- (green).


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CONCLUSIONS Our data constitutes the first experimental evidence showing that some of the PTMs found in LBs isolated from PD brains (i.e. glycation and nitration) affect the capacity of S to act as a fine-tuner of the neuronal redox balance. Nitration and MG-glycation reduced the S capacity to bind Cu2+, although glycation likely involved the appearance of new binding regions triggered by the formation of EDTA-like AGEs. This is supported by the fact that S-CEL and S-CML inhibited the Cu-catalyzed AA degradation and delayed the H2O2 formation better than S, while S-NO2 and S-MG displayed lower inhibition potential. In addition, S-NO2, S-MG and S-CEL were able to increase the [H2O2]. S-CEL and S-CML did not modify the capacity of S to inhibit the formation of HO·, which was clearly reduced and enhanced in the case of S-MG and S-NO2, respectively. In fact, MG-glycation was the only PTM that abolished the capacity of S to inhibit the free radical release. These PTMs also changed the capacity of S to trap or to be damaged by free ROS. AGEs formation but not nitration clearly reduced the capacity of S to trap HO·. Nevertheless, S-MG and S-NO2 displayed a better ability to trap O2·- than S, which was scarcely affected due to CEL or CML formation (Figure 4). In any case, it seems clear that the effect of PTMs on the redox function of S is linked to changes in its metal binding ability. This can merely occur as a result of the chemical modification of certain residues, or concomitant with subtle conformational changes induced by PTMs. For instance, we have recently proved that the averaged conformational ensemble of S-CEL resembles that of native S,61 thus conformational alterations are not required to modify the redox features of S. However, in the case of S-NO2, it seems that the structural rearrangement occurring at the C-terminal domain as a result of the Tyr nitration is responsible for the weakening of the affinity of D119-A124 towards Cu2+, although Tyr are not included within this binding stretch. Altogether, our data contribute to a better understanding of how PTMs occurring in vivo on S affect its physiological function.



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METHODS Chemicals and reagents All chemicals and reagents were analytical grade and they were purchased either from Sigma-Aldrich or from Acros Organics. Moreover, all of them were used as received without further purification, except the methylglyoxal (MG) solution at 40% (SigmaAldrich; M0252), which was purified by steam distillation. The collected fractions were then adjusted to pH 7.0 with a 0.1M NaOH solution, prior to its reaction with H2O2. Afterwards, the concentration of pure MG was indirectly quantified titrating the remaining H2O2 with KMnO4, according to Friedemann’s protocol.62 Collected fractions were frozen until use. All solutions used in the study were prepared by using milli-Q water. -Synuclein expression and purification E. coli BL21(DE3) cells were transformed with the pT7-7 plasmid containing the gene encoding for human -synuclein (S). Transformed cells were grown in sterilized Luria Bertani media (LB) (25g/l) containing ampicillin (100μg/ml) at 37ºC and 180rpm. At OD600nm=0.6-0.8

S

expression

was

induced

with

isopropyl-β-D-1-

thiogalactopyranoside (IPTG) (1mM) and further incubated for an additional 4h at 37ºC and 180rpm. Afterwards, cells were centrifuged (at 4ºC and 4000rpm during 15min) and the resulting pellet was resuspended in lysis buffer (10mM Tris-HCl, 1mM EDTA, 1mM PMSF, pH 8.0) and stirred for 1h at 4ºC. Cells were then lysed by three cycles of 2min sonication step. Cellular debris were removed by centrifugation (20min at 10000 x g and at 4ºC). Nucleic acids were also removed from the lysate by adding streptomycin sulfate (1% w/v) and stirring for 1h at 4ºC, followed by centrifugation during 30min (13500 x g at 4ºC). The obtained supernatant was then supplied by the slow addition of ammonium sulfate (up to 0.295 g/ml) and additionally stirred for 1h at 4ºC to induce the precipitation of S. Thereafter, the pellet was collected by centrifugation (13500 x g at 4ºC during 30min), dissolved in 10mM Tris-HCl at pH 7.4 (1/20th of the LB culture media) and filtered through a 0.22μm filter. The obtained solution was directly loaded onto an anion exchange column (GE Healthcare RESOURCETM Q; 6ml) using a GE ÄKTA Start FPLC. S was eluted with a NaCl gradient (0-600mM), using two different solutions (A: 10mM Tris-HCl at pH 7.4; B: 10mM Tris-HCl and 600mM NaCl at pH 7.4) at a flow rate of 4ml/min. The S fractions were pooled together and dialyzed extensively at 4ºC against 20mM sodium phosphate buffer containing 150mM NaCl (pH 7.4) (buffer B1).

The purity of the

obtained S was checked using MALDI-TOF/TOF (Figure S2A) and SDS-PAGE electrophoresis (Figure S2B) (>96%), whereas the exact molecular mass of the recombinant S (14451.231Da) was determined using a Q-Exactive Orbitrap highresolution mass spectrometer equipped with a heated electrospray ionization (HESI)


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probe (Thermo Fisher), through the deconvolution of the corresponding mass spectrum using Protein Deconvolution 3.0 software (Thermo) (Figure S2C). S concentration was measured by UV-Vis spectroscopy using a molar extinction coefficient estimated on the basis of amino acid content: εS_280nm=5960M-1·cm-1. Preparation and purification of monomeric nitrated -synuclein (S-NO2) Nitration of native S was carried out following a published protocol51 with minor modifications. Monomeric S (10μM) was incubated with 1.2mM tetranitromethane (TNM) (10% in ethanol) in degassed nitration buffer (NB; 0.1M Tris-HCl, 0.1M KCl at pH 8.0) during 3h at 30ºC. Afterwards, the excess of TNM was removed using a 5ml HiTrapTM desalting column, coupled to a GE ÄKTA Start FPLC and using milli-Q water as mobile phase. The fractions corresponding to the nitrated S (S-NO2) (Figure S7A) were pooled together and dialyzed at 4ºC against buffer B1. The formation of homogeneously nitrated monomeric S was confirmed by MALDI-TOF/TOF (Figure S6B) and its exact molecular mass was determined using a Q-Exactive Orbitrap highresolution mass spectrometer through the deconvolution of its mass spectrum (Figure S6C). S-NO2 displayed a molecular weight of 14638Da, which confirmed the addition of a –NO2 group on each one of the four Tyr that holds S (Y39, Y125, Y133 and Y136). This happened at ortho position,63 as suggests the band at 420nm in the UV-Vis spectrum of S-NO2 (Figure S7D), typical of 3-nitrotyrosine (3-NT) at neutral pH.64 Although it has been described that nitration of S yields a heterogeneous mixture of different polymeric and cross-linked species,21,51,52 the use of low concentrations of S (10μM) together with the fact that TNM is a Tyr specific reagent with a low di-Tyr crosslinking potential,63,65 precluded the formation of these species. This was confirmed by MALDI-TOF/TOF, size-exclusion chromatography and SDS-PAGE electrophoresis. Thus the produced S-NO2 was monomeric. Determination of S-NO2 concentration S-NO2 displays a different UV-Vis spectrum than that of native S (Figure S7D). This is mainly because 3-NT has strong absorption properties between 250-500nm.64 Consequently, the εS_280nm cannot be used to calculate the concentration of S-NO2. To determine the value of εS-NO2_280nm, different solutions containing S (2μM or 10μM) and TNM (0.24mM or 1.2mM) were incubated in buffer NB during 3h at 30ºC (the same conditions used to produce S-NO2). Afterwards, the resulting solutions were dialyzed extensively against buffer B1 and the corresponding UV-Vis spectra were acquired at 25ºC using a Shimadzu UV-2401-PC double beam spectrophotometer and quartz cells of 1cm path length. Reaction mixtures prepared in the absence of S but treated identically were used as spectral reference. From the Abs280nm and assuming that the concentration of monomeric S did not change (2 or 10μM), it was possible to estimate the εS-NO2_280nm value as 27702±55M-1·cm-1. 13 ACS Paragon Plus Environment


Page 14 of 30

Preparation and purification of monomeric -synuclein glycated with MG (SMG) Solutions of monomeric S (20μM) were incubated for 24h at 37ºC with MG (50mM) in buffer B1. Later on, the solutions were filtered through a 0.45μm membrane and purified by size-exclusion chromatography using a Superdex-75 HR 10/300 column (GE Healthcare) equilibrated with buffer B1. Samples were injected on an Äkta basic FPLC system at a flow rate of 0.5ml/min coupled to a UV/Vis detector (280nm). The fractions of the peak appearing at elution times similar to those corresponding to native S (Figure S13A), were pooled together and analyzed using MALDI-TOF/TOF and SDS-PAGE electrophoresis (Figure S13B,C). Both techniques evidenced that MG chemically modifies S mostly yielding monomeric S-MG. Its molecular weight was higher than that of native S, which can be ascribable to the addition of different MG moieties on S (likely on Lys side chains).27 Moreover, MALDI-TOF/TOF analysis revealed that S-MG includes a highly heterogeneous population of monomers with molecular weights ranking from ~15 to ~16.5kDa, attributable to different glycation degree and the different chemical nature of the advanced glycation end-products (AGEs) formed on S (Figure S13B). Determination of S-MG concentration AGEs arising from MG are able to absorb UV-Vis radiation,66 as proven by the fact that purified S-MG displays a different UV-Vis spectrum than that of S (Figure S13D). Therefore, the εS_280nm cannot be used to calculate the concentration of S-MG and consequently, the time-dependent variation of the molar extinction coefficient of SMG (εS-MGO_280nm) was determined. The UV-Vis spectra of a reaction mixture containing S (20μM) and MG (50mM) in buffer B1 were acquired during 24h (Figure S14A). The corresponding spectra of a solution containing MG (50mM) alone (Figure S14B) were subtracted to the formers, yielding the S-MG spectra at each incubation time (Figure S14C). From the ΔAbs280nm value and assuming that the concentration of monomeric S (20μM) remains constant during the glycation process (insoluble aggregates were not detected and the DLS autocorrelation function obtained at 0h of incubation overlapped with that obtained after 24h [Figure S14D]), the εS-MGO_280nm at each incubation time can be estimated. The obtained data was fitted to an exponential function (Figure S14E) and the relation between εS-MGO_280nm and the incubation time (h) was: εS-MGO_280nm= 5690 + 19517.9·(1-e-0.11t)

(1)

The UV-Vis absorption spectra were acquired using a Shimadzu UV-2401-PC double beam spectrophotometer thermostatted at 37ºC and quartz cells of 0.1cm path length. The buffer B1 background spectrum was used as spectral reference. The DLS


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autocorrelation functions were obtained using a DynoPro NanoStar at 37ºC, using 4μl disposable plastic cuvettes (WYATT). Preparation and purification of monomeric -synuclein homogeneously glycated with Nε-(carboxyethyl)lysine (S-CEL) or with Nε-(carboxymethyl)lysine (S-CML) Nε-(carboxyethyl)lysine (CEL) and Nε-(carboxymethyl)lysine (CML) are two of the most common AGEs found in S neuronal deposits.25 Therefore, CEL and CML were chemically synthetized on S, which allowed to obtain homogenously glycated samples, but also to specifically study the effect of both AGEs on S. Native S (200μM) was incubated in the presence of glyoxylic acid or pyruvic acid (50mM) in 150mM sodium phosphate buffer (pH 7.4) during 48h and 50ºC. The reaction was carried out in the presence of 75mM NaBH3CN, a reagent that selectively reduces the imine groups at neutral pH.67 After 48h of incubation, the reaction mixtures were dialyzed against buffer B1. MALDI-TOF/TOF analysis of the S incubated with pyruvic acid, showed a spectrum with a main signal centered at m/z~15613Da, although displaying a clear shoulder at m/z~15542Da (Figure S18A). The later matches with the molecular weight of a S with its 15 Lys modified by a carboxyethyl group, whereas the former would correspond to the same specie with an additional carboxyethyl group added either to the N-terminus or as a result of Nε-amine bi-alkylation68,69 (S-CEL). The S incubated with glyoxylic acid displayed a MALDI-TOF/TOF spectrum with a broad peak centered at ~15183Da (Figure S18B). However, its fine structure distribution offered information regarding the occurrence of the CML linkages on each S molecule. It was possible to detect the formation of S-CML10 (m/z ~15041Da), SCML11 (m/z ~15099Da), S-CML12 (m/z ~15156Da), S-CML13 (m/z ~15211Da), SCML14 (m/z ~15270Da) and S-CML15 (m/z ~15331Da), thus proving that the chemical synthesis of CML on S (S-CML) yields a slightly heterogeneous mixture of molecules with different glycation degree. The concentrations of S-CML or S-CEL were measured by UV-Vis spectroscopy. The addition of CML or CEL moieties on S does not involve the formation of additional chromophores, and they do not change the UV-Vis spectrum profile of native S (Figure S18C). Hence, the S-CML/S-CEL concentrations were measured using the εS_280nm=5960M-1·cm-1 estimated for native S. Enzymatic digestions and MALDI-TOF/TOF analysis Native S (150μM) was combined with endoproteinase Glu-C (0.1μg/μl) in buffer B1 and then incubated at 25ºC for 15h. The digested sample (1.5μl) was mixed with 1.5μl of a matrix solution (10μg of -cyano-4-hydroxycinnamic acid in a solution water:acetonitrile (70:30) containing 0.1% TFA). A 0.5μl aliquot of this mixture was 15 ACS Paragon Plus Environment


Page 16 of 30

spotted onto a steel target plate (MTP 384), air-dried, and subjected to mass determination using a Bruker Autoflex III MALDI-TOF/TOF spectrometer equipped with a 200-MHz smart-beam pulsed N2 laser (λ 337nm). The IS1 and IS2 voltages were 19kV and 16.65kV respectively, and the lens voltage was 8.2kV. Measurements were performed using a positive reflector mode with matrix suppression below 400Da. External calibration was performed using a standard peptide mixture. Mass spectra were

matched

against

Swiss-Prot

databases

using

PeptideMass-ExPASy.

The

identification of three fragments confirmed the enzymatic digestion of S (Table S1). Isothermal titration calorimetry (ITC) measurements ITC experiments were carried out to determine the dissociation constants (Kd) and the enthalpy changes (ΔH) of the binding between Cu2+ and the different -synucleins (Ss; i.e. native, S-NO2, S-CEL, S-CML or S-MG). In all experiments, the sample cell (190μl) was filled with a monomeric protein solution (150μM) prepared in buffer B1 and stirred at 250rpm, while a solution containing CuCl2 (3mM) and glycine (6mM) prepared in the same buffer was loaded in the injection syringe. Solutions were degassed before use. Cu2+ solution was titrated into the sample cell as a sequence of 34 injections of 1.43μl each. The time between successive injections was 400s. Raw data corresponding to the heating rate (μcal/s) was integrated to obtain the observed molar enthalpy change. CuCl2 solution was titrated against the buffer and subtracted from the raw data before the model fitting to nullify the heat of dilution. The molar enthalpy change versus the [S-X]/[Cu2+] ratio was analyzed using several models: i) two different independent binding sites; ii) two different sequential binding sites; iii) three different sequential binding sites. The later was the model providing lower residuals and additionally, it gave the dissociation constants more similar to the ones already reported for S-Cu2+ binding process (Table 1 and Figure S8). ITC measurements were carried out in duplicate on a Nano-ITC (TA instruments©) at 25ºC, and data acquisition and analysis were performed using the NanoAnalyze software (TA Instruments). Study of ascorbic acid oxidation Ascorbic acid (AA) is a highly concentrated and an essential antioxidant in neurons.32 However, in presence of some metal cations it degrades with the concomitant formation of reactive oxygen species (ROS).39 To study the effect of the different Ss on the AA degradation, several stock solutions were prepared: i) 100μM CuCl2 in milli-Q water, prepared from a more concentrated solution (20mM CuCl2 in 40mM glycine, a weak Cu2+ chelator). This protocol was carried out to avoid the formation of insoluble Cu(OH)2 when adding Cu2+ to the buffer B1;70 ii) 3mM AA in milli-Q water. This stock solution was prepared freshly every 24h and kept on ice until use; iii) 10mM EDTA in milli-Q water; iv) 10mM N-methyl-glycine (a CML analogue) in milli-Q water; and v)


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10mM N-methyl-DL-alanine (a CEL analogue) in milli-Q water. These stock solutions were combined with buffer B1 to prepare reaction mixtures that always contained 70μM AA and 2.5μM Cu2+, in the absence or in the presence of ~5μM EDTA, 5/10μM of the different Ss as well as of the digested S, 160μM N-methyl-glycine or 160μM Nmethyl-DL-alanine. Solutions containing 70μM AA or 2.5μM Cu2+ alone were also used as controls. The temporal variation in the AA concentration was measured at 256nm during 150min, using a 1cm quart cell and a Shimadzu UV-2401-PC double beam spectrophotometer.

The

absorbance

of

buffer

B1

was

subtracted

from

the

measurements. Experiments were run in triplicate at 25ºC. Study of total ROS formation from Cu-AA degradation AA in presence of Cu2+ degrades towards the formation of different ROS [39], which formation can be monitored studying the decrease in fluorescence of fluorescein.71,72 A fluorescein stock solution (2mM) was prepared in buffer B1 and added to a final concentration of 26μM to reaction mixtures that contained AA (70μM) alone or in the presence of 2.5μM Cu2+. The AA/Cu2+ mixtures were also studied in the presence of 2.5/10μM S. The temporal variation of the fluorescence signal at 518nm (λexc 490nm) was followed during 150min. All the experiments were run in duplicated at 25ºC using a Varian Cary Eclipse fluorescence spectrophotometer and quartz cells of 1cm path length. NMR study of S oxidation Reaction mixtures containing S (10μM), Cu2+ (2.5μM) and AA (70μM) or H2O2 (5mM), were prepared in buffer B1 in presence of 10% (v/v) D2O. These mixtures were incubated at 25ºC and their 1H-NMR spectra were acquired each hour during 24h. The reaction mixture containing H2O2 was used as control to prove the formation of Met-sulphoxide73 on the Met of S during its incubation with AA and Cu2+.

15N/13C

isotope-enriched S (80μM) was also incubated with Cu2+ (2.5μM) and AA (280μM) in buffer B1 in presence of 10% (v/v) D2O. 2D 1H,15N-HSQC spectra were acquired at 25ºC each hour of incubation during 24h. 2D 1H,1H-TOCSY spectra were also acquired in the absence of AA/Cu2+ and in the presence of AA/Cu2+ after 24h of incubation. The spectral assignment of S at 25ºC was obtained from the one deposited at this temperature (Biological Magnetic Resonance Bank code 18857), and confirmed using triple

resonance

HNCACB,

CBCACONH,

HNCO,

HCACO

and

HCCH-TOCSY

experiments (Bruker standard pulse sequences). In all experiments, water suppression was achieved by the Watergate pulse sequence.74 An exponential line broadening of 1.5Hz was applied to the 1H-NMR spectra before Fourier transformation. Chemical shifts were referenced to water at 25ºC. All the experiments were recorded at 25ºC on a 600-MHz Bruker Avance III spectrometer, equipped with a 5-m


13C, 15N, 1H

triple


resonance cryoprobe. All spectra were processed using NMRPipe/NMRDraw75 and analyzed by Xeasy/Cara76 and Sparky software.77 Fluorescence study of the di-tyrosine formation The effect of ROS on Tyr residues in S might involve radical coupling to form dityrosine linkages along a dimerization process.43,78 Hence, the Tyr-Tyr crosslinking was studied on reaction mixtures containing native S or S-NO2 (10μM), Cu2+ (2.5 or 10μM) and AA (70μM), which were prepared in buffer B1 and incubated at 25ºC during 150min. The formation of dityrosine involves the loss of the Tyr fluorescence signal (λmax_em ~305nm) concomitant with an increase in the fluorescence emission intensity between 405-410nm.43 Consequently, the fluorescent spectra of the different reaction mixtures were acquired between 375 and 525nm (λexc 325nm) each 5min using a Varian Cary Eclipse fluorescence spectrophotometer and quartz cells of 1cm path length. Experiments were performed in duplicate. Study of superoxide anion (O2·-) formation Nitroblue tetrazolium (NBT) was used to monitor the time-dependent formation of O2·during the Cu-catalyzed AA degradation. Once O2·- is formed rapidly reacts with NBT to yield formazan, which increases the absorbance of the overall solution at 560nm.79,80 A reaction mixture containing AA (70μM), Cu2+ (2.5μM) and NBT (50μM) was prepared in 10mM sodium phosphate buffer at pH 7.4. The UV-Vis spectra of this reaction mixture were recorded at 25ºC and at different incubation times using a 1cm plastic cell and a Shimadzu UV-2401-PC double beam spectrophotometer. The absorbance of the phosphate buffer was subtracted from the measurements. Experiments were run in duplicate. Study of hydrogen peroxide (H2O2) formation The effect of the different Ss on the H2O2 formation was studied using the red peroxidase kit assay (MAK165; Sigma-Aldrich). Red peroxidase substrate (RS) is a non-fluorescent compound that reacts with H2O2 (1:1) in the presence of horseradish peroxidase (HRP) to form the highly fluorescent resorufin. According to the protocol provided by Sigma-Aldrich, RS was reconstituted with 250μl of DMSO to prepare the RS stock solution, whereas HRP was reconstituted with 1ml of the assay buffer (20U/ml). A solution containing 50μl of the RS stock solution, 200μl of the HRP stock solution and 4.75ml of the assay buffer (master mix; MX) was prepared. The timedependent H2O2 formation was measured on 50μl of samples containing AA, Cu2+ and the different Ss, previously mixed with 50μl of MX solution. In the final reaction mixture, the concentration of AA was always of 70μM, that of Cu2+ was 2.5μM and the concentration of the different Ss was 5/10μM. In all cases, AA was the final reagent added before starting the fluorescence measurements, recorded during 200min. Assay buffer was used as background fluorescence and subtracted from the data. Control 18 ACS Paragon Plus Environment

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experiments without Cu2+ and without AA were also done. The concentrations of H2O2 in the reaction mixtures were determined from a standard calibration curve, obtained using freshly prepared H2O2 solutions with concentrations of 0, 0.5, 2, 4, 10, 20, 30 and 50μM (prepared from a 20mM H2O2 stock solution). 50μl of the different H2O2 solutions were mixed with 50μl of the MX solution, incubated 30min at 25ºC and afterwards, the corresponding fluorescence intensities were recorded. All fluorescence measurements were done in triplicate at 25ºC on a Varian Cary Eclipse fluorescence spectrophotometer using 96-well plates (λexc 540nm; λem 590nm). Study of hydroxyl radical (HO·) formation The effect of the different Ss on the HO· formation was studied using the coumarin-3carboxylic acid (3-CCA). In the presence of HO·, 3-CCA is rapidly oxidized to the fluorescent 7-hydroxy-coumarin-3-carboxylic acid (7-OH-CCA) (λmax_exc 395nm; λmax_em 450nm), thus allowing to record the time-dependent HO· formation.81 A 3-CCA stock solution (20mM) was prepared in 20mM sodium phosphate buffer (pH 9.0), and added to a final concentration of 100μM to reaction mixtures that always contained 70μM AA and 2.5μM Cu2+. These reaction mixtures were prepared in the absence or in the presence of 5μM EDTA, 5/10μM of the different Ss, 160μM Nε-methyl-glycine or 160μM Nε-methyl-DL-alanine. Solutions containing 70μM AA or 2.5μM Cu2+ alone were also used as controls. The temporal variation of the fluorescence signal at 450nm was followed during 150min. All the experiments were run in triplicate or higher at 25ºC using a Varian Cary Eclipse fluorescence spectrophotometer and quartz cells of 1cm path length. Scavenging capacity of AA and the different Ss on the O2·The effect of glycation and nitration on the ability of S to trap the O2·- was studied using the phenazine-methosulfate (PMS)-NADH method.82 The O2·- produced through the PMS-NADH method rapidly reacts with NBT to yield formazan. Therefore, if the different Ss are able to scavenger O2·-, the formation of formazan will decrease and consequently, the absorbance of the overall solution at 560nm. Initially, stock solutions of PMS (30μM), NADH (338μM) and NBT (200μM) were prepared in 10mM sodium phosphate buffer (pH 7.4) and stored at 4ºC until use. Afterwards, reaction mixtures containing 84.5μM NADH and 7.5μM PMS, in the absence or in the presence of 10μM of S, S-NO2, S-MG, S-CEL, S-CML, Nε-methyl-glycine (160μM) or Nεmethyl-DL-alanine (160μM) or AA (5, 30, 100, 200 or 300μM), were prepared in 100mM sodium phosphate buffer (pH 7.4) and incubated during 5min at room temperature. Subsequently, NBT was added to a final concentration of 50μM and incubated 5min at room temperature. The UV-Vis spectra of the different reaction mixtures were acquired at 25ºC using a 1cm plastic cell and a Shimadzu UV-2401-PC



double beam spectrophotometer. The absorbance of the phosphate buffer was subtracted from the measurements. Experiments were run in triplicate or higher. Scavenging capacity of the different Ss on the HO· The cupric reducing antioxidant capacity (CUPRAC) method was applied to analyze the effect of glycation and nitration on the ability of S to trap the HO·.83,84 CUPRAC involves the formation of HO· as a result of the reaction between Fe2+ and H2O2. After this reaction is completed, the latter is degraded using catalase to avoid chemical interferences during the determination. HO· can hydroxylate salicylic acid, which is able to reduce the neocuproine-Cu2+ to neocuproine-Cu+ (yellow; λabs_max 450nm). If the different Ss are able to trap HO·, this would avoid the hydroxylation of salicylic acid and consequently, the formation of the neocuproine-Cu+ complex. Initially, stock solutions of salicylic acid (10mM), FeCl2 (20mM in milli-Q water containing 40μl of HCl 1M for each ml of solution), EDTA (20mM), H2O2 (10mM), CuCl2 (10mM) and AcNH4 (1M) were prepared in milli-Q water. Additionally, neocuproine (7.5mM) was prepared in ethanol, whereas a catalase solution (298U/ml) was prepared in buffer B1. All stock solutions were stored at 4ºC in dark until use. Initially, reaction mixtures containing 0.5mM salicylic acid, 0.5mM Fe2+, 0.5mM EDTA and 0.5mM H2O2, in the absence or in the presence of S (10μM), S-NO2 (10μM), S-MG (10μM), S-CEL (10μM), S-CML (10μM), Nε-methyl-glycine (160μM) or Nε-methyl-DL-alanine (160μM), were prepared in B1 and incubated during 10min at 37ºC and at 500rpm. Afterwards, catalase was added to a final concentration of 15U/ml and further incubated 30min under the same conditions. Later on, 100μl of the resulting mixtures were diluted in 1ml of milliQ water containing 1mM of Cu2+, 0.75mM of neocuproine and 0.2M of AcNH4. These solutions were incubated during 2.5h at room temperature and their UV-Vis spectrum were recorded at 25ºC using a 1cm quartz cell and a Shimadzu UV-2401-PC double beam spectrophotometer. The absorbance of a 1ml mixture containing 1μM of Cu2+, 0.75μM of neocuproine, 0.2M of AcNH4 and 100μl of milli-Q water were subtracted from the measurements. Experiments were run in duplicate. MALDI-TOF/TOF MS study of the effect of reactive oxygen species (ROS) on the molecular weight of the different Ss To analyze whether the different Ss could act as ROS scavengers, the time-dependent changes in their molecular mass were monitored using a Bruker Autoflex III MALDITOF/TOF spectrometer equipped with a 200-MHz smart-beam pulsed N2 laser (λ 337nm). Aliquots of 1μl of reaction mixtures containing 10μM S (native, S-NO2, SMG, S-CML or S-CEL), 70μM AA and 2.5μM Cu2+ prepared in buffer B1, were taken after 0 and 150min of incubation (25ºC) and supplemented by TFA (0.2% v/v). Samples were then combined with 1μl of matrix solution (10μg of -cyano-4hydroxycinnamic acid in a solution water:acetonitrile (70:30) containing 0.1% TFA),


Page 20 of 30

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and a 0.5μl aliquot of this mixture was spotted onto a steel target plate (MTP 384), airdried and subjected to mass determination. The IS1 and IS2 voltages were 20kV and 18.5kV respectively, and the lens voltage was 7.5kV. Measurements were performed using a positive reflector mode with matrix suppression below 400Da. The spectra were calibrated externally using a protein calibration standard (3600-17000Da) from Bruker. The experiments were performed in duplicate. Statistical analysis Experimental values in the different graphs are expressed as mean +/- standard deviation. Comparisons between different data groups (formation of H2O2 in presence of S vs. in presence of S-NO2; formation of H2O2 in presence of S-MG vs. in the absence of protein; formation of HO· in presence of S vs. in presence of S-CML) were analyzed using the independent-samples t-test. A two-tailed p-value less than 0.05 were considered statistically significant. Statistical analyses were performed using SPSS version 23.0 (SPSS Inc., Chicago, IL, USA).



Page 22 of 30

ASSOCIATED CONTENT Supporting Information The Supporting Information is available free of charge on the ACS Publications website at DOI: XXX Electrophoretic and spectroscopic characterization of recombinant native S, S-NO2, S-MG, S-CML and S-CEL. Absorbance and fluorescence studies on the effect of S, S-NO2, S-MG, S-CML and S-CEL on the AA degradation, on the overall formation of ROS, and on the specific formation of H2O2 and HO·. Absorbance studies on the ability of S, S-NO2, S-MG, S-CML and S-CEL to scavenger HO· and O2·-. Spectroscopic studies of the effect of ROS on the oxidation state of S. Spectroscopic studies of the effect of Glu-C digested S on the AA degradation. ITC studies on the biding of S, S-NO2, S-MG, S-CML or S-CEL with Cu2+.

ABBREVIATIONS S, -synuclein; LBs, Lewy bodies; PD, Parkinson’s disease; PTMs, non-enzymatic post-translational methylglyoxal;

modifications;

CEL,

iPSCs,

induced

Nε-(carboxyethyl)lysine;

CML,

pluripotent

stem

cells;

Nε-(carboxymethyl)lysine;

MG, AA,

ascorbic acid; ROS, reactive oxygen species; S-NO2, monomeric nitrated -synuclein; S-MG,

mixture

of

-synuclein

monomers

heterogeneously

glycated

with

methylglyoxal; S-CEL, monomeric -synuclein where all Lys residues have been replaced by CEL; S-CML, monomeric -synuclein where all Lys residues have been replaced by CML.

AUTHOR INFORMATION Corresponding Author *Phone: +34 971 173491. E-mail: [email protected] ORCID Miquel Adrover: 0000-0002-4211-9013 Joaquin Ortega-Castro: 0000-0001-8131-0315 Author Contributions H.M.-O. performed the UV-Vis and the fluorescence studies. L.M. acquired, processed and analyzed the NMR data and carried out the protein production and purification. A.B.U. acquired and analyzed the mass spectroscopy data. H.M.-O. and J.O.-C. carried out the nitration and the glycation of S and purified the modified proteins. B.V. carried out the ITC measurements and the interpretation of the derived data. J.F. and M.A. conceived and designed the experiments. M.A., L.M. and J.F. wrote the manuscript. H. M.-O., L. M. and A. U. contributed equally.


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Notes The authors declare no competing financial interest.

ACKNOWLEDGMENTS The authors are grateful for the excellent technical assistance from the Serveis Cientificotècnics at the UIB, especially to Dr. Gabriel Martorell for his generous help with NMR measurements and to Dr. Rosa Gomila for her aid with the MALDI-TOF set up and analysis. We are also grateful to the Dr. Pilar Sanchis for her help in the statistical analysis. We thank Dr. Kris Pauwels for providing the S plasmid (originally obtained from Dr. Daniel Otzen [Aarhus University]), for carefully proof-reading the manuscript and for reviewing its scientific content. H.M. wishes to acknowledge the “Fundación Carolina” for his fellowship (2017). L.M. wants to thank MINECO for the FPU PhD grant FPU14/01131. This work was funded by the Spanish Government in the framework of the Project CTQ2014-55835-R.



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