Impact of the Molecular Environment on ThiolâEne Coupling For

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Impact of the Molecular Environment on Thiol-Ene Coupling For Bio-Functionalisation and Conjugation Burcu Colak, Júlio Cosme Santos Da Silva, Thereza A. Soares, and Julien Ezra Gautrot Bioconjugate Chem., Just Accepted Manuscript • DOI: 10.1021/acs.bioconjchem.6b00349 • Publication Date (Web): 11 Aug 2016 Downloaded from http://pubs.acs.org on August 16, 2016

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Bioconjugate Chemistry

Impact of the Molecular Environment on Thiol-Ene Coupling For Bio-Functionalisation and Conjugation Burcu Colak,1,2 Julio C. S. Da Silva,3 Thereza A. Soares3,4 and Julien E. Gautrot1,2* 1

Institute of Bioengineering and 2 School of Engineering and Materials Science, Queen Mary,

University of London, Mile End Road, London, E1 4NS, UK. 3

Departament of Fundamental Chemistry, CCEN, Federal University of Pernambuco, Cidade

Universitária, 50670-901, Recife, PE, Brazil. 4

Department of Chemistry, Umeå University, 90187 Umeå, Sweden.

* To whom correspondence should be addressed E-mail: [email protected].

ACS Paragon Plus Environment


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Abstract Thiol-ene radical coupling is increasingly used for the biofunctionalisation of biomaterials and the formation of 3D hydrogels enabling cell encapsulation. Indeed, thiol-ene chemistry presents interesting features that are particularly attractive for platforms requiring specific reactions of peptides or proteins, in particular in situ, during cell culture or encapsulation. Despite such interest, little is known about the factors impacting thiol-ene chemistry in situ, under biologically relevant conditions. Here we explore some of the molecular parameters controlling photoinitiated thiol-ene couplings with a series of alkenes and thiols, including peptides, in buffered conditions. 1H NMR and HPLC were used to quantify the efficiency of couplings and the impact of the pH of the buffer, as well as the molecular structure and local microenvironment close to alkenes and thiols to be coupled. Some of these observations are supported by molecular dynamics and quantum mechanics calculations. An important finding of our work is that the pKa of thiols (and its variation upon changes in molecular structure) have a striking impact on coupling efficiencies. Similarly, positively charged and aromatic amino acids are found to have some impact on thiol-ene couplings. Hence, our study demonstrates that molecular design should be carefully selected in order to achieve high biofunctionalisation levels in biomaterials with peptides or promote the efficient formation of peptide-based hydrogels.


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Introduction The biofunctionalisation of biomaterials is important to confer bioactive properties essential to their use in a wide range of applications, including medical implants,1, scaffolds

3

2

tissue engineering

cell culture platforms for cell expansion and in vitro assays,4 drug delivery systems

and imaging probes.5 Typically, proteins and peptides are adsorbed or coupled to biomaterials to alter cell phenotype or tissue response. In particular, cell culture in 3D hydrogels has recently received an increased attention due to the potential of such systems for the design of tissue engineering platforms and in vitro culture models.6 It is now well-established that cells interact with their microenvironment through a variety of parameters such as cell-cell interactions, adhesion to the matrix and sensing of its biochemical and physical properties as well as the presence of cytokines and growth factors.7 Including such parameters to the design of new biocompatible hydrogels that provide structural support but can be remodelled by cells as well as direct their phenotype is therefore particularly important. One essential element for such design is the ability to couple proteins or peptides to biomaterials in situ, in the presence of cells, in mild conditions and with non-toxic reagents, to maintain cell viability and preserve their phenotype. Such in situ biofunctionalisation can be applied for the design of 3D hydrogels encapsulating cells, for tissue engineering or as in vitro models,8-12 or the development of controlled dynamic cell-based assays.13, 14 Click chemistry has received much attention for the design of biofunctional platforms, owing to the efficiency of click couplings in mild conditions and at low concentrations (often desired due to the cost of proteins or peptides). In 2001, Sharpless et al. introduced the concept of click chemistry

15

This new concept referred to highly selective reactions, readily proceeding under

aerobic and ambient atmospheric conditions, not or little affected by molecular complexity and highly efficient, resulting in high yields and ease of isolation of products. A number of click chemistry systems have been developed and have had a considerable impact in the fields of chemistry, biochemistry and materials science. One of the most classic examples of click chemistry is the copper (I)-catalysed azide-alkyne cycloaddition,16, 17 but recently thiol-alkene (thiol-ene) and thiol-alkyne (thiol-yne)18-20 couplings have attracted some attention as they make use of readily available chemical functions present in the structure of polymers and materials as well as peptides and proteins.18-20 Thiol-ene chemistry has been studied since the 1900s and has



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been applied in various fields of contemporary chemistry, including organic synthesis, polymer chemistry, materials chemistry, surfac e functionalisation and bioconjugation.14,

21-24

The thiol-ene reaction proceeds via a two-step

radical process (see Figure 1a) in mild conditions and often quantitative yields. It requires only low concentrations of initiators, typically occurs fast and allows the simple isolation of products. The conditions required are so mild that even sunlight was reported to initiate free radical thiolene coupling (in the presence of a photoinitiator).25 In addition, a wide range of initiator systems, including many thermal and photoinitiators, have been developed.20, 26 Finally, oxygen and water inhibition, typically preventing conventional radical reactions and polymerisations, can often be neglected in thiol-ene reactions.25, 27 Thiol-ene chemistry has been applied in a wide range of systems,28 but in the field of polymer synthesis and functionalisation, has been particularly studied in organic solvents, in concentrated solutions or neat monomer systems.26,

29-31

Whereas these conditions are perfectly suited to

macromolecular design, they are often poorly adapted to the biofunctionalisation of biomaterials and the in situ preparation of hydrogels for 3D cell culture. Indeed, for such applications, low concentrations and reactions in buffers preserving the structure of proteins or the viability of cells are required. Therefore, it is important to study and understand the behaviour of thiol-ene chemistry in biologically relevant conditions and contexts. In organic solvents and neat conditions, the kinetics of thiol-ene couplings has been reported to proceed faster with electron-rich alkenes.28, 32 Slower reaction rates were observed with electron deficient molecules, but still allowed couplings to occur.25, 33 Other reactivity rules have been established, highlighting the importance of the respective rates of thiyl radical addition and transfer.34-36 For example, thiol-ene chemistry is thought to be low yielding with allylic and benzylic compounds, due to the hydrogen abstraction by thiyl radicals from the allylic and benzylic positions and therefore the formation of the stabilised radical species.34 Important differences were also reported between polymer-small molecule20 and polymer-polymer conjugation.37 For instance, Koo et al. noted that the efficiency of thiol-ene coupling between two macromolecules is reduced due to competitive bimolecular termination reactions.32 Recently, the robustness and simplicity of thiol-ene chemistry, the mild conditions required and the availability of alkene functions as side chains of macromolecules (including those not


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activated towards polymerisation or Michael additions) and that of thiols present in proteins and peptides, specifically through cysteine residues, have contributed to its popularity for biofunctionalisation and hydrogel formation, in buffers or physiological conditions. Thiol-ene chemistry was used for the biofunctionalisation of surfaces and polymer brushes, for example for the generation of protein patterns or the design of dynamic cell-based assays via in situ peptide coupling.14, 38 Hydrogels have also been produced via thiol-ene based crosslinking.21, 39, 40 In many of these systems, thiol-ene coupling is used to confer biofunctionality and bioactivity to the corresponding biomaterials, through the use of peptide sequences presenting cysteine residues.41-44 This strategy was applied to confer cell adhesion to biomaterials and culture platforms8, 14 and di-cysteine peptides were used to crosslink polymers and form enzymatically degradable hydrogel networks loaded with cells.45, 46 Despite such interest in applying thiol-ene coupling to biomaterials chemistry, little is known of the impact of structural parameters controlling the efficiency of thiol-ene coupling in buffered and physiological conditions. Compared to more traditional systems used in polymer functionalisation and the curing of resins, the impact of the chemistry of the biomaterials to be functionalised on thiol-ene coupling is not well established. Similarly, rules and guidelines controlling the reactivity of peptides via radical thiol-ene reactions have not been investigated systematically. In particular, the impact of changes in the molecular environment of the reactive centres, which have been directly implicated in the regulation of disulfide exchanges for example,47,

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has not been systematically studied but should contribute to simplify and

standardise the use of thiol-ene radical coupling for bioconjugation and biofunctionalisation of biomaterials. Such understanding will also be essential to control reaction conditions in biologically relevant contexts (for example, during cell encapsulation in 3D hydrogels, in vitro or for in vivo delivery) and to manage and address toxicity and mutagenicity concerns. In this report, we systematically study some of the reaction parameters (pH, concentration, type of medium) and structural factors (chemistry of polymer backbones, thiols and cysteine-bearing peptides) controlling radical thiol-ene coupling in biologically relevant buffers. To this aim, we developed simple methodologies, using NMR, HPLC and quantum computational chemistry, to quantify and rationalise reaction efficiencies. We identify important rules for the design of the chemical structure of thiol-ene based biomaterials.



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Figure 1. a) Thiol-ene reaction between pentenoic acid (A1) and the series of thiols (S1-S6). bd) 1H-NMR spectra of the alkene (d) and product peaks (b, c) of the A1-S1 reaction at different UV exposure times: 0 s (0 % conversion), 5 s (41 %), 10 s (68 %), 20 s (86 %) and 30 s (94 %). e) Impact of the time of UV exposure on the reaction conversion for A1 with a series of thiols.


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Results and discussion Design of a model system to study thiol-ene coupling. The reaction of a range of thiols (S1-5) with pentenoic acid A1, in equimolar concentrations, was used as a model system to monitor the coupling of thiols to alkenes in buffered conditions relevant to biofunctionalisation (see Figure 1a). These molecules were selected based on their solubility in PBS, simplicity of chemical structure and low molar masses. The efficiency of coupling was assessed via 1H NMR, via the integration of the peaks at 4.8-5.1 and 5.8-5.9 ppm, corresponding to olefinic protons, and that of the peaks at 1.5-1.68 and

2.54-2.61 ppm,

corresponding to the formation of thioethers (see Figure 1a-d). We first investigated the effect of UV exposure time on the efficiency of the thiol-ene reaction (Figure 1e). Reactions with thiols S1-5 all reached 92-100 % in 45 s of exposure to UV light, however, the reaction was sensitive to the molar mass of the reactants and associated steric hindrance, as the rates of coupling of PEG methyl ether thiol S5 and L-glutathione S3 were the slowest. For the rest of our studies, an exposure time of 60 s was used as conversions did not significantly improve between 45 s and this time point. In addition, we investigated the impact of the photoinitiator concentration and the intensity of the UV light on the extent of reaction (see Figures S1 and S2). We found that at an initiator concentration of 2 mol% (1 mM), conversions were already reaching a plateau. The UV exposure time had a clear impact on reaction conversions, with coupling efficiencies reaching a plateau near 10 mW/cm2. Impact of the pH of the buffer. Having determined optimised reaction conditions, the thiol-ene coupling of thiols S1, S4, S5 and S6 with alkene A1 was monitored next as a function of pH (Figure 2). Reactions were performed in PBS with 2 mol% of photoinitiator, a UV power of 17 mW/cm2 and 60 s of UV exposure time. The pH of the deuterated buffer was adjusted using NaOD and DCl and conversions were calculated based on 1H-NMR as in Figure 1. Reaction efficiencies were at their maximum (between 92 and 100 %) between pH 4 and 7, except for thiol S6, which showed a minor decrease to c.a. 84 %. The reaction deteriorates with increasing pH and coupling efficiencies fall to 0 % above pH 9 to 11, depending on the thiol. Indeed, in this pH range, the formation of increasing concentrations of thiolates prevents the formation of thiyl radicals and does not afford any notable coupling (Figure 2b).



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Figure 2. a) The impact of the buffer pH on the reaction conversion (60 s UV exposure) for A1 with thiols S1 and S4-S6. b) Impact of pH on thiol-ene reaction (left) and proposed hydrogen bonding in thiols S1 and S4-6. c-e) Optimised configurations of thiols S4 (c), S6 (d) and S1 (e).


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We hypothesized that differences in the pH at which reaction efficiencies rapidly collapse were attributable to differences in the pKa of the four thiols tested. Indeed, the pKas of these thiols should be influenced by the occurrence of intramolecular hydrogen bonds. Therefore, we calculated the pKa of thiols S1, S4, S6 and a diethylene oxide thiol modelling PEG S5 in the presence and absence of intramolecular hydrogen bonds. The cis and trans conformations of the respective thiols were used for the calculations. The cis and trans conformations differ with respect to the values (0 or 180 degrees) of the dihedral angles S-C-C-N in compounds (S1) and (S2), and S-C-C-O in compound (S3), respectively. The QM/MM calculations indicate that the cis conformation of cysteamine S4 is approximately 3.64 kcal/mol more stable than the trans conformation (Figure 2c). Likewise, the cis conformation of mercaptoethanol S6 is ca. 2.2 kcal/mol more stable than the trans conformation (Figure 2d). For both compounds the calculated pKa decreases upon the formation of an intramolecular hydrogen bond with the thiol group (Table 1). In the case of N-acetyl L-cysteine S1, the amide group is not a strong hydrogen bond acceptor, and thus was not found to form hydrogen bonds with the thiol group. However, the carboxylate group at pH ≥ 3 can form intramolecular hydrogen bonds with both the thiol and the amide groups when in the trans conformation (Figure 2e). The optimized geometries for S1 indicate that the formation of intramolecular hydrogen bonds between the carboxylate and the thiol and the amide groups (cis conformation) is about 5.5 kcal/mol-1 more stable than the trans conformation where the carboxylate makes one single hydrogen bond with the amide group (Figure 2e). However, the shorter length of the hydrogen bond formed between the carboxylic acid and the amide group (1.98 Å) compared to that formed with the thiol group (2.35 Å) contributes to weaken the impact of such conformation on the pKa of the molecule. Experimental measurements established pKa values of 9.5 and 8.0 for the thiol group in S1 and cysteine, respectively,49 in agreement with our measurements (comparing S1 and S4, Table 1). The thiol group in the diethylene oxide model compound of S5 was not found to form intramolecular hydrogen bonds. These findings support a direct correlation between the pKa calculated for the different thiols tested and the occurrence of a strong intramolecular hydrogen bond involving the thiol moiety (Table 1). Hence, the formation of hydrogen bonds involving the thiol group in S1, S4 and S6 lowers their respective pKas, albeit the effect is weaker for S1 (Table 1 and Figure 2c-e). These results suggest that



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thiols of non-terminal cysteine residues and, more generally, not in close proximity with proton donors, should react well via thiol-ene coupling in buffered conditions at physiological pH.

Table 1: Calculated pKa values for thiols S1, S4, S6 and the model compound of thiol S5 in the presence or absence of intramolecular hydrogen bonds. The cis and trans conformationsa refer to the dihedral angles S-C-C-N in compounds S1 and S4, and S-C-C-O in compound S6, respectively. Thiol Compounds

pKa

Atomic charge (S-)

cis

Trans

Cis

Trans

Mercaptoethanol (S6)

8.2

10.1

-1.011

-1.117

Cysteamine (S4)

8.7

9.1

-1.036

-1.057

N-acetyl cysteamine (S1)

9.6

9.4

-1.087

-1.079

Moldel of PEG thiol (S5)b

9.0

-1.050

a

Intramolecular hydrogen bonds are formed in the cis conformation for S6 and S4 and in the trans conformation for S1. bIntramolecular hydrogen-bonds are absent in this compound. The pKa was estimated only for the lowest energy (trans) conformation of the compound.

Impact of the composition of the molecular environment. Next, the impact of other molecules present in the reaction medium was examined. We carried out thiol-ene reactions between S1 and A1 in the presence of amino acids and different electrolytes. Reaction mixtures containing S1 and A1 in deuterated PBS, with 2 mol% photoinitiator were spiked with a range of amino acids and electrolytes at two different molar ratios. The resulting solutions were then adjusted to pH 7 using NaOD or DCl, as free amino acids can change the pH of the resulting mixtures, and exposed to 60 s of UV light (17 mW/cm2, Figure 3). As before, 1H-NMR was used to study the efficiency of the reactions. Generally, thiolene coupling in the presence of a range of amino acids with neutral hydrophilic, hydrophobic or


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charged structures remained efficient, with most conversion levels between 73 and 93 %, confirming the very good tolerance of thiol-ene chemistry to a large range of functionalities. Amongst the neutral amino acids tested, proline, glycine and the hydrophobic phenylalanine and tyrosine did not show any significant decrease in coupling compared to the control (non-spiked) conditions. Efficiencies were more variable in the case of glutamine, serine and leucine, resulting in a modest (8-13 %) decrease in coupling, although not statistically significant. Although the reasons for this increase in variability are not clear, in the case of leucine it could arise from possible proton abstraction to form stable tertiary radicals. The presence of tryptophan also resulted in an increase in the variability of the coupling efficiencies measured (the average is reduced by 16 %, p = 0.170), potentially due the olefinic character of the double bond in the pyrrole ring and the possibility of addition of thiyl radicals to this moiety. Hence thiol-ene coupling may not be fully compatible with peptide sequences displaying high ratios of tryptophan residues compared to cysteines, although 10 % seems to be well tolerated. Amongst the charged amino acids tested, the negatively charged aspartic acid also resulted in an increase in the variability of the coupling efficiencies, with an overall 14 % decrease in coupling (n.s., p = 0.206), perhaps due to weak acid-base interactions with thiol moieties, resulting in some level of deprotonation. Three positively charged amino acids were tested: the primary amine lysine, the imidazole-derivative histidine and the guanidine derivative arginine (Figure 3b). Histidine did not affect the coupling of S1 to A1, possibly due to its low pKa and therefore neutral character at physiological pH. In contrast, lysine decreased the reaction efficiency by 19 % (p 0.031; the 15 % decrease by arginine was no significant, with a p value of 0.265), possibly through electrostatic stabilisation of the thiolate form of S1.



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Figure 3. a) Thiol-ene reaction between A1 and S1 in the presence of other amino acids and electrolytes. b) Reaction conversions (60 s UV exposure) of the A1-S1 reaction in the presence of a range of amino acids (aromatic, hydrophobic, positive, negative, neutral) and electrolytes. Blue represents 10 mol% (4.5 mM) and red 100 mol% (45 mM) of the added components with respect to the thiol:ene concentrations (1:1). See Table S1 for corresponding statistical analysis. n.s., non significant; *, p < 0.2; **, p < 0.05: ***, p < 0.01.


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Finally, the effect of different electrolytes, in particular different halides was examined. This was the result of our observation that allyl ammonium iodide does not significantly react via thiol-ene coupling (results not shown). Such behaviour led us to test whether bromides and iodides had an impact on thiol-ene coupling. Therefore, we added equimolar amounts of potassium bromide and potassium iodide to reaction mixtures of S1 and A1. We found that potassium iodide efficiently quenched the coupling (control KI 10 p 0.00123, control KI 100 p 0.0014), whereas the bromide electrolyte had a minor effect (84 % efficiency, n.s. p 0.434 Figure 3b). This is consistent with electron transfer and quenching phenomena observed with halides50 and the lower standard reduction potential of iodine atoms compared to bromine.51 Impact of the chemical structure close to the alkene moiety. Thiol-ene coupling having direct applications in the bioconjugation of proteins and peptides to polymers and small molecules presenting alkene functions41 and for the formation of degradable hydrogels,11, 52, 53 we next examined the role of the local chemical structure close to the alkene moiety. In addition to the negatively charged alkene A1, we tested the reactivity of N-acetyl Lcysteine S1 with neutral alkene amides A2 and A3 and positively charged alkenes A4-9 (see Figure 4). The neutral alkene amide A2 reacted with relatively high efficiency with S1 (73 %), but the allyl amide derivative A3 showed a 71 % drop in reactivity compared to that of A1. In addition, the allyl ammonium derivative A4 only reacted 4 % with S1. We proposed that the close proximity (one CH2 group only) of the alkene moiety and the nitrogen atom in A3 and A4 could account for the lack of reactivity of these two molecules with S1. Therefore we examined the coupling of S1 to A5 and A6, which display two and three methylene between the ammonium group and the alkene moiety, respectively. The reaction efficiency was restored to near completion in both cases (see Figure 4b). Similarly, polyelectrolyte A7, presenting olefinic ammonium residues showed poor reactivity in thiol-ene coupling with S1 (5 %, respectively), but polyelectrolytes A8 and A9 with extended aliphatic chains reacted well (> 99 %).



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Figure 4. a) Impact of alkene structure on the thiol-ene reaction with S1. b) Reaction conversion 60 s UV, photoinitiator 1 mM, A2-A6 or 300 s UV 2.5 mM, A7-A9) of S1 and the series of alkenes (A2-A9). See Table S2 for corresponding statistical analysis. c) Calculated Gibbs free energies of reactants, intermediates, transition states and products involved in the reaction of S1 with A4-6. d-f) Singly occupied molecular orbitals calculated for intermediates corresponding to A4 (d), A5 (e) and A6 (f).


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To gain further insight into the lack of reactivity of allylic ammoniums compared to longer olefinic derivatives, we have used density functional theory (DFT) to calculate the free energies and reaction profiles associated with the propagation and chain transfer steps for intermediates involved in the reaction of A4-6 to S1 (Figure 4c). The activation energy of the first step of the reaction, during which a carbon radical is formed, is nearly identical for all three compounds (activation barrier energies of 7.3, 7.6 and 6.7 kcal/mol corresponding to reactants A4, A5 and A6, respectively, see Table 2). The second step was found to be rate determining for the global reaction with calculated activation barrier energies of 19.3, 13.6 and 13.1 kcal/mol for intermediates associated with A4, A5 and A6, respectively (Table 2). Therefore, the kinetic discrimination due to the differential reactivity of the reactants occurs at the chain transfer step with an estimated free energy difference of 6.0 kcal/mol in favour of A5 and A6, compared to A4 (Figure 4c). In addition, the reaction profiles confirmed that the overall reaction is thermodynamically favourable for the conversion of all three reactants.

Table 2 – Calculated free energies for propagation and chain-transfer processes involving Nacetyl-cysteine and alkenes A4, A5 and A6

Alkene Reactants

Propagation

Chaintransfer

∆G(kcal/mol)

∆G(kcal/mol)

TS1

Intermediate

TS2

Product

A4

0.0

7.3

0.7

19.3

-0.4

A5

0.0

7.6

2.0

13.6

-1.0

A6

0.0

6.7

2.3

13.1

-1.8



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It was previously postulated that the rate of chain transfer is correlated with the stability of the carbon-centered radical generated following propagation.35 Extensive computational calculations for thiol−ene reactions between methyl mercaptan and a series of alkenes confirmed the importance of the electron density near the alkene groups to modulate the stability of the carbon radical.30 Analysis of singly occupied molecular orbital (SOMO) of the corresponding transition states of the chain transfer gave further insight into the origin of the differences in activation energy calculated for the chain transfer. SOMOs calculated for the transition states of TS2-A4, TS2-A5 and TS2-A6 are presented in Figure 4d. Analysis of the SOMO for the transition states A4, A5 and A6 indicates a considerable overlap between the carbon centered radical and the alkyl electron-donor group adjacent. This overlap extends only up to the two closest alkyl group from the carbon-centred radical, with a low contribution of the nitrogen atom from +NR4 in A4 and A5. The +NR4 is an electron-withdrawing group, which in this case, can withdraw the electron density of the alkyl groups reducing their capacity to donate electron density for carboncentred radicals in the chain transfer step. The NBO partial atomic charges computed on the carbon-centred radical for the transition states of TS2-A5 (-0.41 e) and TS2-A6 (-0.42 e) are ca. 2 times more negative than for the transition state of TS2-A4 (-0.24 e), due to the donor effects of the longer alkyl chains. In addition, the increased distance from the positively charged ammonium decreases the electron withdrawing effect on the carbon radical centre, further promoting the stabilization of the radical and the corresponding transition state. The activation energy for addition of carbon-based radicals to alkenes is controlled by the reaction enthalpy and the charge transfer occurring in the corresponding transition state.54 Based on the structural similarity and comparable activation energies associated with radical formation for reactants A4, A5 and A6, we argue that charge transfer is the primary mechanism via which additional alkyl groups in A5 and A6 stabilize the carbon-based radical, with respect to A4. Impact of the chemical structure of cysteine-terminated peptides. The impact of the chemical structure in the vicinity of the thiol, specifically a cysteine within a peptide sequence, was investigated next, due to the potential importance of molecular design on reactivity and bioconjugation. In particular, we monitored the impact of aromatic amino acids, the vicinity of charges and the position of the cysteine residue on coupling efficiencies (see Figure 5). To do so, we used a combination of 1H NMR (to determine the extent of coupling) and


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HPLC (to explore the role of concentration and buffer on the coupling efficiency). We selected sequences of pentameric amino acid peptides for this study, with the ultimate residue being aromatic to allow UV monitoring via HPLC, and reacted them with alkene A1. We found that tyrosine in the sequence resulted in a decrease of the coupling efficiency compared to phenylalanine (Figure 5b), in slight contrast to our results on the reaction of A1 with S1 in the presence of single amino acids (Figure 3). Perhaps this is a result of the increased concentration of tyrosine in the reaction mixture with the present peptide (100 % compared to cysteine) as the single amino acid tyrosine was not sufficiently soluble to be tested at concentrations above that of 10 % of N-acetyl L-cysteine (so above 4.5 mM). However, the results showed that despite the larger molar mass of peptide GCGSF (compared to N-acetyl L-cysteine), high conversions (86 % by 1H NMR; 100 % by HPLC) could be reached. Hence peptide coupling via thiol-ene reaction can be efficient at moderate concentrations (45 mM) and in buffered conditions.



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Figure 5. a) The impact of the structure of peptides on thiol-ene coupling with A1. b) Sequence of peptides with cysteines in green, glycines and serines in blue, charged amino acids (arginine and aspartic acid) in red and aromatic (phenylalanine and tyrosine) amino acids in purple. c) Reaction conversions quantified by 1H-NMR for 4 peptide sequences (300 s UV exposure, 2.5 mM initiator, 45 mM peptide). d) Reaction conversions quantified by HPLC for 7 peptide sequences (300 s UV exposure, 2.5 mM initiator, 45 mM peptide). See Table S3 for corresponding statistical analysis. n.s., non significant; *, p < 0.2; **, p < 0.05: ***, p < 0.01. e)


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Reaction conversion and remaining reagents measured for the coupling of GCGSY and A1 at different concentrations (300 s UV exposure, 2.5 mM initiator; blue squares, product formed; blue circles, unreacted peptide). f) Reaction conversion and remaining reagents measured for the coupling of CRGSF/CGRSF and A1 at different concentrations (300 s UV exposure, 2.5 mM initiator). CRGSF (blue squares, product formed; blue circles, unreacted peptide) and CGRSF (red squares, product formed; red circles, unreacted peptide). Solid and dotted lines are only intended to guide the eye.

The position of the cysteine residue in the peptide sequence was also found to be important to improve thiol-ene coupling efficiency (Figure 5b). Two main cysteine-based sequences are commonly found in the literature to couple peptides via thiol-ene reaction or Michael addition: CGGXX55 and GCGXX

11, 42, 43, 53, 56

We explored the impact of the position of the cysteine on

the reaction efficiency and found a marked decrease in coupling when the cysteine residue was placed in the ultimate position (68 ± 2 % by HPLC p = 0.052). We propose that this is due to the local chemical structure of the resulting CGGSF peptide as the thiol finds itself two carbons away from a primary amine, as in cysteamine S4. Cysteamine S4 was found to quickly lose reactivity just above neutral pH (Figure 2a), due to a decrease in the pKa of its thiol group. Our results indicate that similar effects take place in the case of cysteine-terminated peptides, leading to a slight decrease in coupling efficiency in PBS (pH 7.4), highlighting that GCGXX type of sequences should be preferred for the design of peptide sequences for bioconjugation and biofunctionalisation. Neighbouring amino acids were also found to have a modest impact on thiol-ene efficiency (Figure 5b). The introduction of the negatively charged amino acid residue aspartic acid directly adjacent to the cysteine or separated from it by a glycine resulted in a slight increase in coupling efficiency (directly adjacent: 66 % by 1H NMR; 96 % by HPLC; separated by a glycine: 75 % by HPLC; p = 0.00524, comparing CDGSF and CGDSF). In contrast, the positively charged amino acid arginine had a negative effect on the thiol-ene reaction, whether when introduced directly next to the cysteine or separated from it by a glycine (64 % by 1H NMR for CRGSF; both CRGSF and CGRSF 58 % by HPLC). These results suggest that the local chemical structure around the thiol moiety plays a modest role on the thiol-ene coupling, but still impacts on its



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efficiency. The opposite effects of positively and negatively charged neighbouring amino acids imply that they could stabilise and destabilise thiolates, respectively. This hypothesis, based on our observations made at different pH (Figure 2), should result in the associated changes in reactivity, as observed in Figure 5b. To explore further what parameters impact on thiol-ene reaction, we examined the role of concentration on the efficiency of the coupling. This is particularly relevant to bioconjugation methods (in particular for coupling to proteins and antibodies), which can require low concentrations to be used. The concentrations of all reagents (including that of the photoinitiator) were reduced by identical factors by direct dilution of the most concentrated stock solution and the thiol-ene coupling of GCGSY to pentenoic acid A1 was monitored via HPLC. We found symmetric trends for the evolution of the starting peptide (unreacted) and that of the product, suggesting a loss of reactive radicals during the reaction at low concentrations. Below concentrations of 4.5 mM, no substantial product formation was observed. In addition, the sum of product and reagent concentrations only accounted for 90 % of the total thiol amounts initially used (Figure 5c). This suggests that, below a concentration of 4.5 mM of thiols, radical species are quenched by other species (or processes) present in the reaction milieu. In contrast, the changes in reagent and product concentrations observed with arginine-containing peptides were not symmetrical (Figure 5d). Hence whereas the product formation remained unchanged (between 56 and 67 %) over a range of starting peptide concentration of 4.5 to 45 mM, the relative concentration of starting peptide changed from 0 % (at high concentrations of 45 mM) to 61 % (for a starting concentration of peptide of 4.5 mM). Hence our results suggest that at high concentrations, a substantial fraction of the reacted peptides are not contributing to product formation and are presumably converted to other species via side-reactions. This could imply that radical-radical coupling reactions, which would be more sensitive to concentration than other side-reactions (as their rates depend on the square of the radical concentration), are frequent at high concentrations. Hence such bimolecular recombination was found to dominate termination in polymer-polymer couplings mediated by thiol-ene chemistry.32


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Figure 6. a) The impact of photoinitiator and peptide concentration (GCGSY) on the reagent conversion and product formation. Peptide concentrations were 4.5 mM (blue squares, product formed; blue circles, unreacted peptide) and 9 mM (red squares, product formed; red circles, unreacted peptide). Solid and dotted lines are only intended to guide the eye. b) The impact of buffer type and degassing on peptide (GCGSY) conversion and product formed. PBS (blue squares, product formed; blue circles, unreacted peptide), degassed PBS (red squares, product formed; red circles, unreacted peptide) and HEPES (green squares, product formed; green circles, unreacted peptide). Solid and dotted lines are only intended to guide the eye.



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Impact of competitive, radical quenching mechanisms. Thiol-ene couplings are thought to be relatively insensitive to the presence of oxygen as rate transfer of carbon-based radicals to thiols are typically one order of magnitude higher than their quenching by oxygen.57, 58 However, as the concentration of thiols decreases (for example in cases for which only low concentrations of reagents, proteins or peptides are practically usable), transfer rates may be dominated by oxygen quenching. Therefore, we tested whether such events significantly occur in peptide coupling via thiol-ene chemistry (Figure 6). We studied the impact of the concentration of photoinitiator on the reaction efficiency with the peptide GCGSY at two different concentrations (4.5 and 9 mM, Figure 6a), corresponding to 5- and 10-fold dilutions compared to experiments reported in Figure 5b. We found no effect of the photoinitiator concentration, within the range tested, on the consumption of thiols or the formation of product, although the latter levels were higher by c.a. 9 % at peptide concentrations of 9 mM. In addition, we carried out experiments at different thiol concentrations in degassed PBS buffer, to reduce the concentration of oxygen (Figure 6b). We found that although the levels of thiol consumption were comparable, the formation of product was significantly impaired (down to 47 %). These experiments suggest that oxygen is not the agent responsible for the quenching of radicals and the loss of product formation at higher concentrations (45 mM), but rather protects thiols against such reactions. In addition, our results imply that the reduced coupling observed at lower concentrations of thiols is not due to quenching by oxygen species but rather a simple reflection of reduced reaction rates. Therefore thiol-ene coupling displays some of the hallmarks of an oxygen-tolerant reaction, even at low reagent concentrations. Interestingly, we found that the choice of buffer is more important than reducing the concentration of oxygen. When couplings were carried out in HEPES buffer (pH 7, Figure 6b), we found a reduction in product formation and thiol consumption, perhaps indicating that tertiary amines from the buffer (present at 10 mM concentrations) can trap some of the radicals and reduce the reaction rate. Therefore, the presence of competing species other than oxygen appears as a limiting factor for thiol-ene coupling in buffers relevant to biofunctionalisation of biomaterials. In this respect, phosphate buffers appear as good media for thiol-ene based conjugation, biofunctionalisation and hydrogel formation.


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Conclusion Thiol-ene chemistry displays interesting features for the biofunctionalisation of biomaterials. The present study focused on the determination of some of the parameters controlling thiol-ene efficiency in buffered conditions relevant to the coupling of peptides and potentially proteins to biomaterials (surfaces or soluble polymeric materials). Our results demonstrate that the coupling of small molecules in buffered conditions occurs with high efficiency, although restricted by size, presumably via a diffusion-controlled mechanism. However, the pH of the environment and the pKa of the thiol to be coupled have dramatic effects on the efficiency of the reaction. Changes in the chemical composition of the reaction medium, the local structure of the alkene as well as the thiol, in particular in the case of cysteine-bearing peptides, also impact coupling efficiencies. These observations have important implications in the development of biofunctionalisation strategies, for example for the design of peptide sequences allowing the control of biochemical and physical properties of peptide-based biomaterials and hydrogels. Hence, for the loading of cells in 3D hydrogels, radical thiol-ene chemistry appears to be a promising and versatile tool as it can often be carried out in PBS, at neutral pH, and peptide sequences can be designed to increase the pKa of the thiols. However, HEPES should be avoided and other buffers may similarly interfere with the efficiency of the reaction. In addition, the presence of high concentrations of proteins, which may present amino acids interfering with such radical coupling should be avoided. Surprisingly, oxygen does not seem to be an issue and may even play a beneficial role in this reaction, although the underlying mechanism is unclear. In contrast, the use of thiol-ene chemistry for coupling of proteins and antibodies to surfaces and biomaterials remains challenging and our results indicate that the low thiol concentrations typically used when coupling such larger biomacromolecules would result in considerably lower rates of reactions. These issues should be addressed for thiol-ene coupling to be included in the toolbox of methodologies compatible with the precise, regioselective coupling of proteins and antibodies to biomaterials and biosensors.



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Materials and methods Materials PEG methyl ether thiol (average Mn 1,000), N-acetyl L-cysteine (99%), cysteamine hydrochloride (97%), L-glutathione reduced (98%), ethanethiol (97%), 2-mercaptoethanol (99%),

4-pentenoic

acid

(97%),

Irgacure

2959

(2-hydroxy-4'-(2-hydroxyethoxy)-2-

methylpropiophenone, (98%), phosphate buffered saline tablet, allylamine (98%), triethylamine (99%), diethylamine (99.5%), 4-pentenoyl chloride (98%), allyl bromide (97%), 5-bromo-1pentene (95%),

4-bromo-1-butene (97%), acetic anhydride (98%), 2-(dimethylamino)ethyl

methacrylate (contains 700-1000 ppm monomethyl ether hydroquinone as inhibitor, 98%), ethyl α-bromoisobutyrate (98%), ethanol (99.8%), 2,2'-bipyridine (>99%), copper (I) chloride (>99.995% trace metals basis), deuterium oxide (99.9% atom % D), deuterium chloride solution (37 wt% in deuterium oxide, 99% atom % D), sodium deuteroxide (40% in deuterium oxide 99 atom % D), hydrochloric acid (37%), anhydrous magnesium sulphate (99.5%), sodium bicarbonate (99.7%), sodium chloride (99.5%), dichloromethane (99.5%), anhydrous dichloromethane (99.8%), methanol (HPLC 99.9%), tetrahydrofuran (99.9%), ethyl acetate (99.7%), diethyl ether (99%), petroleum ether 40-60°C, acetonitrile (HPLC 99.9%), heptane (HPLC 99%), dimethylformamide (pharmaceutical secondary standard) and silica gel were purchased from Sigma Aldrich. Custom peptides GCGSY, GCGSF, CGGSF, CRGSF, CGRSF, CDGSF and CGDSF (95 %) were purchased from Proteogenix, France. Synthesis of N,N-diethylpent-4-enamide (A2) Diethylamine (1 eq., 0.0253 mol, 1.85 g, 2.62 mL) was dissolved in anhydrous dichloromethane (7 mL) and the flask was placed in an ice bath. Triethylamine (1 eq., 2.56 g, 1.86 mL) was added, then pentenoyl chloride (1 eq., 3 g, 2.79 mL) dropwise with vigorous stirring. The content turned brown and contained salts. Dichloromethane (100 mL) was added to the mixture and extracted from hydrochloric acid (0.5 M, 30 mL three times), saturated sodium bicarbonate (30 mL three times) and brine (30 mL three times). The aqueous layers were washed with dichloromethane and recovered. The organic phases were combined, dried over magnesium sulphate and evaporated to yield a brown oil 3.02 g, 99 % pure. 1H NMR (400 MHz; CDCl3) δ 1-1.1 (6H, dt), 2.32 (4H, m), 3.2-3.3 (4H, dq) 4.9-5


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(2H, m) and 5.7-5.8 (1H, m) (Figure S7). v/cm-1 ~2900 (w, C-H), 1643 (s, C=O) and 1432 (m, C=C) (Figure S8). Synthesis of N-allylacetamide (A3) Allylamine (1 eq., 0.0294 mol, 1.678 g, 2.20 mL) and triethylamine (1 eq., 0.0294 mol, 2.97 g, 4.099 mL) were dissolved in tetrahydrofuran (10 mL), then acetic anhydride (5 eq., 0.147 mol, 15 g, 13.89 mL) was slowly added with stirring and the mixture was heated to 50 ºC for 24h resulting a brown solution. The solvent was evaporated and a column was performed in petroleum ether: ethyl acetate (1:1), then graduated to ethyl acetate. The solvent was evaporated to yield a brown oil 3.458 g, 75 % pure (impurity is acetic acid). 1H NMR (400 MHz; D2O) δ 2.0 (3H, s), 3.7 (2H, d), 5-5.1 (2H, m) and 5.7-5.8 (1H. m) (Figure S7). v/cm-1 ~2900 (w, C-H), 1641 (s, C=O), 1548 (m, C=C) and 1284 (m, C-N) (Figure S8). Synthesis of N,N,N-triehtyl-N-prop-2-en-1-ammonium bromide (A4), N,N,N-triehtyl-N-but-3-en1-ammonium bromide (A5) and N,N,N-triehtyl-N-pent-4-en-1-ammonium bromide (A6). Triethylamine (1.25 eq., 0.0310 mol) was dissolved in diethyl ether (5.5 mL), then was added allyl bromide (1 eq., 0.0248 mol) or 4-bromo-1-butene (1 eq.) or 5-bromo-1-pentene (1 eq.) , the reaction was heated 50 ºC for 17h. The solution was evaporated using pressure. Diethyl ether was added and the solution was dried twice more to yield a white powder (A4) 99 % pure, yellow oil (A5 and A6) 99 % pure. N,N,N-triethylprop-2-en-1-ammonium (A4) 1H NMR (400 MHz; D2O) δ 1.3 (9H, t), 3.3 (6H, q), 3.8 (2H, d), 5.6-5.7 (2H, m) and 5.8-5.9 (1H, m) (Figure S7). v/cm-1 ~2900 (w, C-H), 1398 (m, C=C) and 1170 (m, C-N) (Figure S8). N,N,N-triethylbut3-en-1-ammonium (A5) 1H NMR (400 MHz; D2O) δ 1.2 (9H, t), 2.4 (2H, dt), 3.1 (2H, m), 3.2 (6H, q), 5.1-5.2 (2H, m) and 5.6-5.7 (1H, m) (Figure S7). v/cm-1 ~2900 (w, C-H), 1398 (m, C=C) and 1170 (m, C-N) (Figure S8). N,N,N-triethylpent-4-en-1-ammonium (A6) 1H NMR (400 MHz; D2O) δ 1.2 (9H, t), 1.7 (2H, tt), 2.1 (2H, dt), 3-3.1 (2H, m), 3.2 (6H, q), 3.8 (2H, d), 5-5.1 (2H, m) and 5.7-5.8 (1H, m) (Figure S7). v/cm-1 ~2900 (w, C-H), 1403 (m, C=C) and 1178 (m, C-N) (Figure S8). Synthesis of PDMAEMA A solution with ethanol: deionised water (1:4) was prepared and degassed for 30 min. 2(dimethylamino)ethyl methacrylate (0.0954 mol) was weighed into a flask containing ethyl α-



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bromoisobutyrate (0.00037 mol), dissolved in ethanol-water (7.5 mL) and degassed for 30 min. Into a second flask was weighed 2,2'-bipyridine (0.00019 mol), dissolved in ethanol-water solution (7.5 mL) and degassed for 30 min. To the 2,2'-bipyridine solution was added copper (I) chloride (0.00019 mol), the brown solution was sonicated for 10 minutes. The catalyst solution was transferred to the monomer solution and the reaction was stirred under inert atmosphere for 5 h, 50 ºC. The ethanol was evaporated and the water freeze dried. The remaining solid was dissolved in tetrahydrofuran and added to silica gel (20 g), agitated for 1 h, filtered and concentrated using a rotary evaporator, then precipitated in heptane, filtered and dried under reduced pressure, 98 % pure. PDMAEMA, GPC, Mn 67926, PD 1.6 (Figure S12). 1H NMR (400 MHz; D2O) δ 0.75-1.2 (3H, m), 1.7-2.0 (2H, m), 2.3 (6H, s), 2.6-2.8 (2H, m) and 4.1 (2H, m) (Figure S9). v/cm-1 ~2900 (w, C-H), 1720 (s, C=O), 1261 (m, C-N) and 1100 (s, C-O) (Figure S10). Functionalisation of PDMAEMA to PDMAEMA propene (A7), PDMAEMA butene (A8), PDMAEMA pentene (A9). PDMAEMA (1eq., 1 g, 0.0064 mol) was dissolved in dimethylformamide (10 mL), then was added allyl bromide (5eq., 0.031 mol), 4-bromo-1-butene or 5-bromo-1-pentene (2.5eq., 0.015 mol) and the reaction was heated overnight at 70 ºC. Precipitated in diethyl ether and the remaining solid was dissolved in methanol and precipitated in diethyl ether, the recovered polymer was precipitated from methanol twice more. The polymer was recovered and dried under reduced pressure, A7 and A9 is 97 % and A8 98 % pure. PDMAEMA propene (A7) 1H NMR (400 MHz; D2O) δ 0.80-1.3 (3H, m), 1.8-2.1 (2H, m), 3.2 (6H, s), 3.6-3.9 (2H, m), 4.0-4.1 (2H, m), 4.4-4.6 (2H, m) 5.7-5.9 (2H, m) and 6.0-6.2 (1H, m) (Figure S9). v/cm-1 ~2900 (w, CH), 1727 (s, C=O), 1660 (s, C=C), 1272 (m, C-N) and 1150 (s, C-O) (Figure S10). PDMAEMA butene (A8) 1H NMR (400 MHz; D2O) δ 0.75-1.2 (3H, m), 1.8-2.1 (2H, m), 2.6-2.7 (2H, m), 3.2 (6H, s), 3.4-3.6 (2H, m), 3.7-4.0 (2H, m), 4.3-4.6 (2H, m) 5.3-5.4 (2H, m) and 5.8-5.9 (1H, m) (Figure S9). v/cm-1 ~2900 (w, C-H), 1726 (s, C=O), 1646 (w, C=C), 1247 (m, C-N) and 1149 (s, C-O) (Figure S10).PDMAEMA pentene (A9) 1H NMR (400 MHz; D2O) δ 0.85-1.4 (3H, m), 1.92.1 (2H, m), 2.1-2.3 (4H, m), 3.3 (6H, s), 3.4-3.5 (2H, m), 3.7-4.0 (2H, m), 4.3-4.6 (2H, m) 5.05.2 (2H, m) and 5.8-6.0 (1H, m) (Figure S9). v/cm-1 ~2900 (w, C-H), 1727 (s, C=O), 1643 (w, C=C), 1243 (m, C-N) and 1137 (s, C-O) (Figure S10).


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Thiol-ene reaction for NMR monitoring. Deuterated-PBS was prepared (1 PBS tablet per 200 mL deuterium oxide) in a volumetric flask. This solution was used for the thiol-ene experiments. Thiol (45.4 µmol, 45 mM) and alkene (45.4 µmol, 45 mM) 1:1 ratio were dissolved in deuterated-PBS (1 mL). Irgacure 2959 solution was prepared in methanol (0.198 M, 0.0444 g/mL). To thiol-ene solution (0.5 mL) was added 2 mol% (1 mM, 5 µL, for all small molecules) or 5 mol% (2.5 mM, 12.5 µL, for polymers or peptides) photoinitiator in methanol (stock solution concentration: 0.198 M). The samples were irradiated with UV (17 mW, 350-500 nm) for a known amount of time (60 s for small molecules, power 1.02 J/cm2; 300 s for polymers and peptides, power 5.10 J/cm2) and analysed by 1H NMR. The conversion was calculated through the consumption of the alkene peak with respect to the formation of the two product peaks, the average of the two calculations were taken to obtain the average percentage conversion, (1H NMR for the thiol-ene products, starting from various thiols, alkenes and peptides are presented in Figures S3, S6 and S11). Certain variables controlling the thiol-ene reaction were investigated; UV irradiation time, photoinitiator concentration, pH of the solution and intensity of UV irradiation with a series of thiols and alkenes. The pH of the solutions were adjusted using the Hanna instruments pH meter. Thiol-ene reaction for HPLC monitoring. The peptides were dissolved in PBS at known concentrations (45.4 µmol, 45 mM) then diluted to known concentrations and measured using HPLC connected to a UV detector (275 nm for tyrosine containing sequences and 254 nm for phenylalanine containing sequences) for the peptide calibration. Atlantis T3 3 μm 4.6x150 mm column was used with a solvent gradient; from 0-20 min acetonitrile: water 2:98 to 15:85, 20-25 min 30:70, 25-35 min 100:0, 35-40 min 2:98. Reaction mixtures containing peptide: pentenoic acid (A1) 1:1 and photoinitiator 5 mol% (2.5 mM) were prepared and reacted using UV for 300 s then diluted and measured using HPLC for the reaction calibration. Irgacure 2959 solution was prepared in methanol (0.198 M, 0.0444 g/mL). For the low concentration reaction, mixtures containing peptide: pentenoic acid (A1) 1:1 and photoinitiator 5 mol% (2.5 mM) were prepared, diluted to known concentrations then



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reacted using UV for 300s. HPLC traces were analysed for the product formation and unreacted peptide via a calibration, typical traces are shown in Figures S13, S14 and S15.

Instrumentation. 1

H NMR was produced using Bruker AV 400 and AVIII 400. Abbreviations for the peaks: s,

singlet; d, doublet; t, triplet; q, quartet; dt, doublet of triplets; dq, doublet of quartets; tt, triplet of triplets and m, multiplet. ATR-FTIR were produced using a Bruker Tensor 27 spectrometer equipped with a MCT detector, results were acquired at a resolution of 16 cm-1 and a total of 128 scans per run in the region of 600-4000 cm-1. Abbreviations for the peaks: s, strong; m, medium and w, weak. The UV light used to initiate reactions was the Omnicure series 1500. The radiometer used to measure the UV light intensity was from International light technologies, ILT 1400-A radiometer photometer. HPLC analysis was conducted using a Waters separations module instrument equipped with a 2489 UV/vis detector, a reversed-phase C18 Atlantis T3 3 μm 4.6x150 mm column, a reversed-phase C18 Atlantis T3 5 μm 3x50 mm guard column and a water/acetonitrile gradient. All samples were filtered through 0.2 μm supor membrane pore before analysis. GPC analysis were measured using an Agilent 1260 Infinity system equipped with an refractive index and variable wavelength detector, 2 PLgel 5 μm mixed-C column (300x7.5 mm), a PLgel 5 mm guard column (50x7.5 mm) operated in DMF with NH4BF4 (5 mM). The instrument was calibrated with poly(methyl methacrylate) standards (550 to 46890 g/mol). All samples were filtered through 0.2 μm nylon 66 before analysis. Statistical analysis. All data were analysed pairwise by Tukey's test and increased variation in data was determined by * p < 0.2 and significance was determined by ** p < 0.05: *** p < 0.01. A full summary of statistical analysis is provided in the Tables S1-S3. Computational details. All quantum chemical and QM/MM calculations were carried out using the C.01 version of the Gaussian 09 program. In the study of the reaction mechanism, the geometry optimizations were carried out using the hybrid GGA B3LYP exchange-correlation functional.59, 60 The 6-311G(d,p) basis set functions was used to treat H, C, N, O and S atoms. Open-shell species were described


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using an unrestricted treatment. Transition states were obtained using the Synchronous TransitGuided Quasi-Newton (STQN) method developed by Schlegel and co-workers.61 For chain transfer step, the transition states were found using the QST3 method.62 Transition states were characterized as having a single imaginary frequency whose vibrational modes correspond to formation of RS–C bond (in the first step) and a concerted process in the second step, in which the S-H bond is breaking as the H-C bond is forming. Cartesian atomic coordinates for reactant, intermediate and transition states obtained along the reaction coordinate are available upon request. The calculations of the pKa of cysteamine (S3), mercaptoethanol (S6), N-acetyl L-cysteine (S1) and PEG methyl ether thiol (S5) was carried out using a computational approach based on the linear relationship between atomic charges of the anionic form of the thiolates and their respective experimental pKa values.63 Different protocols to compute the atomic charges and to treat solvent effects were previously validated.63 For the cases where the atomic charges were calculated using the ChelpG method64 at the M06-2X/6-311/CPCM level of theory, starting from structures optimized at the same level of theory, the pKa was calculated from the linear relationship given by equation 1:

pKa=−18.923qChelpG (S − )−10.970

(Equation 1)

where qChelpG(S-) is the partial atomic charge of the S- species in the anionic form of thiolates molecules. We have modified the original computational protocol proposed by Monard and coworkers,63 to include explicit water molecules around the thiol-containing compounds during the calculation of the atomic charges. This was required to take into account the effects of solventassociated charges and polarization on the stabilisation of thiol group and the conformation of the molecule. Hence, geometry optimization was performed for the thiol compounds at the M062X/6-311/CPCM level of theory, and atomic charges were obtained from the optimized geometries using the ChelpG method. The resulting configurations and atomic parameters were used as initial configuration for Monte Carlo simulations in the NVT ensemble, which provided representative configurations of the solvent molecules around the thiol compounds. Classical Monte Carlo simulations were carried out using an intermolecular pair interaction potential containing the 12-6 Lennard-Jones and Coulomb terms. The parameters used in the LennardJones term were taken from OPLSAA force field.65 Based on the last configuration obtained



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from the MC simulations, water molecules within a sphere of 6 Angstrom of radius from the thiols were selected. MC-generated configurations were geometry-optimized via Quantum Mechanics/Molecular Mechanics (QM/MM) using the ONIOM method,66 as implemented in the Gaussian09 software via the scheme (M06-2x/6-311G:UFF). The QM/MM calculations were performed for the neutral thiol species, which were subsequently used for the ChelpG charge calculations upon removal of the proton of the thiol group. These calculations were carried out at the M06-2X/BS2/CPCM level of theory, in which BS2 indicates that the 6-311G basis set were used to treat thiol molecules and the 3-21G basis set to describe the water molecules. Long-range electrostatic interactions were treated via the use of CPCM method67 with a dielectric constant ε = 78.0. These geometries and atomic charges were used to estimate the pKa of the thiol group using equation 1. Values of atomic charge of S- and pKa calculated for the four thiols are reported in Table S1.

Author Information Corresponding Author. *E-mail: [email protected]. Notes. The authors declare no competing financial interest.

Acknowledgment B.C. thanks Queen Mary, University of London for her PhD studentship. JCSS is the recipient of a PNPD/CAPES postdoctoral fellowship. TAS is the recipient of a visiting professorship from the Umeå Centre for Microbial Research (UCMR) Linnaeus Program. Dr Helena Azevedo and Daniela S. Ferreira are acknowledged for help and discussion with peptide design and characterisation via HPLC. The authors are grateful to the Brazilian funding agencies FACEPE and CAPES (BioComp Grant), to the Royal Society (Newton International Exchange Scheme, NI140090) and to the Swedish Foundation for International Cooperation in Research and Higher Education (STINT, IG2011-2048) for financial support.

Supporting Information The Supporting Information is available free of charge on the ACS Publications website at DOI: .Additional experimental results (additional kinetics, NMR, FTIR, GPC, HPLC, optimised geometries of thiols and results of statistical analysis).


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TOC Figure. Impact of the molecular environment on thiol-ene chemistry.



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Table of content 90x35mm (300 x 300 DPI)


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Proposed cover 139x138mm (300 x 300 DPI)



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Figure 1. a) Thiol-ene reaction between pentenoic acid (A1) and the series of thiols (S1-S6). b-d) 1H-NMR spectra of the alkene (d) and product peaks (b, c) of the A1-S1 reaction at different UV exposure times: 0 s (0 % conversion), 5 s (41 %), 10 s (68 %), 20 s (86 %) and 30 s (94 %). e) Impact of the time of UV exposure on the reaction conversion for A1 with a series of thiols. Figure 1 121x242mm (300 x 300 DPI)


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Figure 2. a) The impact of the buffer pH on the reaction conversion (60 s UV exposure) for A1 with thiols S1 and S4-S6. b) Impact of pH on thiol-ene reaction (left) and proposed hydrogen bonding in thiols S1 and S46. c-e) Optimised configurations of thiols S4 (c), S6 (d) and S1 (e). Figure 2 162x173mm (300 x 300 DPI)



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Figure 3. a) Thiol-ene reaction between A1 and S1 in the presence of other amino acids and electrolytes. b) Reaction conversions (60 s UV exposure) of the A1-S1 reaction in the presence of a range of amino acids (aromatic, hydrophobic, positive, negative, neutral) and electrolytes. Blue represents 10 mol% (4.5 mM) and red 100 mol% (45 mM) of the added components with respect to the thiol:ene concentrations (1:1). See Table S1 for corresponding statistical analysis. n.s., non significant; *, p < 0.2; **, p < 0.05: ***, p < 0.01. Figure 3 190x160mm (300 x 300 DPI)


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Figure 4. a) Impact of alkene structure on the thiol-ene reaction with S1. b) Reaction conversion 60 s UV, photoinitiator 1 mM, A2-A6 or 300 s UV 2.5 mM, A7-A9) of S1 and the series of alkenes (A2-A9). See Table S2 for corresponding statistical analysis. c) Calculated Gibbs free energies of reactants, intermediates, transition states and products involved in the reaction of S1 with A4-6. d-f) Singly occupied molecular orbitals calculated for intermediates corresponding to A4 (d), A5 (e) and A6 (f). Figure 4 195x290mm (300 x 300 DPI)



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Figure 5. a) The impact of the structure of peptides on thiol-ene coupling with A1. b) Sequence of peptides with cysteines in green, glycines and serines in blue, charged amino acids (arginine and aspartic acid) in red and aromatic (phenylalanine and tyrosine) amino acids in purple. c) Reaction conversions quantified by 1HNMR for 4 peptide sequences (300 s UV exposure, 2.5 mM initiator, 45 mM peptide). d) Reaction conversions quantified by HPLC for 7 peptide sequences (300 s UV exposure, 2.5 mM initiator, 45 mM peptide). See Table S3 for corresponding statistical analysis. n.s., non significant; *, p < 0.2; **, p < 0.05: ***, p < 0.01. e) Reaction conversion and remaining reagents measured for the coupling of GCGSY and A1 at different concentrations (300 s UV exposure, 2.5 mM initiator; blue squares, product formed; blue circles, unreacted peptide). f) Reaction conversion and remaining reagents measured for the coupling of CRGSF/CGRSF and A1 at different concentrations (300 s UV exposure, 2.5 mM initiator). CRGSF (blue squares, product formed; blue circles, unreacted peptide) and CGRSF (red squares, product formed; red circles, unreacted peptide). Solid and dotted lines are only intended to guide the eye. Figure 5 197x234mm (300 x 300 DPI)


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Figure 6. a) The impact of photoinitiator and peptide concentration (GCGSY) on the reagent conversion and product formation. Peptide concentrations were 4.5 mM (blue squares, product formed; blue circles, unreacted peptide) and 9 mM (red squares, product formed; red circles, unreacted peptide). Solid and dotted lines are only intended to guide the eye. b) The impact of buffer type and degassing on peptide (GCGSY) conversion and product formed. PBS (blue squares, product formed; blue circles, unreacted peptide), degassed PBS (red squares, product formed; red circles, unreacted peptide) and HEPES (green squares, product formed; green circles, unreacted peptide). Solid and dotted lines are only intended to guide the eye. Figure 6 103x180mm (300 x 300 DPI)


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