Antimicrobial activity of catechol functionalized-chitosan versus Staphylococcus epidermidis

Accepted Manuscript Title: Antimicrobial activity of catechol functionalized-chitosan versus Staphylococcus epidermidis Authors: Andrea Amato, Luisa Maria Migneco, Andrea Martinelli, Loris Pietrelli, Antonella Piozzi, Iolanda Francolini PII: DOI: Reference:

S0144-8617(17)31106-2 https://doi.org/10.1016/j.carbpol.2017.09.073 CARP 12818

To appear in: Received date: Revised date: Accepted date:

27-7-2017 8-9-2017 22-9-2017

Please cite this article as: Amato, Andrea., Migneco, Luisa Maria., Martinelli, Andrea., Pietrelli, Loris., Piozzi, Antonella., & Francolini, Iolanda., Antimicrobial activity of catechol functionalized-chitosan versus Staphylococcus epidermidis.Carbohydrate Polymers https://doi.org/10.1016/j.carbpol.2017.09.073 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Antimicrobial activity of catechol functionalized-chitosan versus Staphylococcus epidermidis

Andrea Amatoa, Luisa Maria Mignecoa, Andrea Martinellia, Loris Pietrellib Antonella Piozzia, Iolanda Francolinia*

a

Department of Chemistry, Sapienza University of Rome, P. le Aldo Moro 5, 00185 Rome, Italy b

ENEA, C.R. Casaccia, Via Anguillarese 301, 00100 Rome, Italy

*Corresponding author:

Dr. Iolanda Francolini Sapienza University of Rome, Department of Chemistry, P. le Aldo Moro 5, 00185 Rome, Italy E-mail: [email protected] Tel: +39 06 4991 3162. Fax: +39 06 4991 3692

E-mail addresses:

[email protected] [email protected] [email protected] [email protected] [email protected]

1

Highlights

- New protein mussels mimics chitosan (CS) derivatives have been synthesized by reaction with a catechol-bearing molecule (hydrocaffeic acid, HCAF). - An efficient procedure for acetonide-protection of the catechol moiety of HCAF was set up. - The CS-HCAF conjugates showed good antioxidant activity confirming the efficacy of the adopted synthetic strategy. - Chitosan derivatives possessed enhanced antimicrobial activity versus Staphylococcus epidermidis compared to pure chitosan. - Commercial dressing coated with one of the CS-HCAF conjugates was able to significantly reduce S. epidermidis adhesion and prevent biofilm formation.

Abstract

Protein mussel-inspired adhesive polymers, characterized by the presence of catechol groups, possess superior muco-adhesive properties and have great potentiality in wound healing. Suitable materials for wound dressing should properly combine muco-adhesiveness and antimicrobial activity. In this work, catechol-functionalized chitosan was obtained by reaction with hydrocaffeic acid (HCAF), in order to investigate how catechol introduction at different content could affect the intrinsic antimicrobial activity of the polymer itself. Unexpectedly, an enhancement of chitosan antimicrobial activity was observed after catechol functionalization, with a fourfold reduction in the polymer minimum inhibitory concentration versus Staphylococcus epidermidis. Additionally, a commercial wound dressing coated with one of the synthesized CS-HCAF derivatives showed a significant reduction in the adhesion of S. epidermidis compared to the uncoated dressing (3-log reduction). The CS-HCAF derivatives also showed an interesting antioxidant property (EC50

2

ranging from 20 to 60 µg/mL), which further confirms the potentiality of these materials as wound dressings.

Keywords Chitosan; Hydrocaffeic acid; Bioadhesive polymers; Protein mussels mimics polymers; Wound dressings.

3

1.

Introduction

The development of bio-adhesive materials is an emerging topic in the biomaterial science because of their potential application as wound healing patches, tissue sealants, and hemostatic materials. Particularly, bio-adhesives are highly attractive for wound healing because they are able of adhering to body tissues in moist environments, are flexible, adapt to skin irregularities and can allow the controlled delivery of drugs (Padula et al., 2007). Among adhesive materials, the bio-inspired ones are those developed by applying principles used in nature to produce bio-adhesive molecules (Favi, Yi, Lenaghan, Xia, & Zhang, 2014). Indeed, adhesion to a surface by living organisms is generally mediated by the secretion of molecules endowed with reactive species able to establish strong chemical or physical bonds with the surface itself. A classical example of bio-adhesive molecules is represented by the set of proteins produced by marine mussels to anchor to foreign surfaces in seawater, in which the main component is the catecholic amino acid (Danner, Kan, Hammer, Israelachvili, & Waite, 2012). A number of bio-adhesives inspired by mussel adhesive proteins have been developed over the last decade (Forooshani & Lee, 2017). These materials have the common feature to display the catechol ortho-dihydroxyphenyl group, that is the side chain of 3,4dihydroxyphenylalanine (DOPA), known to be responsible for the superior adhesiveness of mussel proteins (Kaushik et al., 2015). Examples of polymeric protein mussel mimics are catechol derivedpolyethylenimine (Long, Xiao, & Cao, 2017; Kim et al., 2012; Lee et al., 2008), catechol derivedhyaluronic acid (Lih, Choi, Ahn, Joung, & Han, 2016; Lee et al., 2008) and catechol derivedchitosan (Ryu, Hong, & Lee, 2015; Ryu et al., 2011). Especially chitosan has attracted attention in the field of bio-adhesives because this polysaccharide possesses intrinsic muco-adhesive properties (Duttagupta, Jadhav, & Kadam, 2015). Very recently, partially crosslinked catechol-functionalized chitosan was used to coat needles to develop self-sealing haemostatic needles able to prevent bleeding following tissue puncture (Shin et al., 2017). Alongside this, chitosan has other important biological activities, specifically antimicrobial and hemostatic, which further justify the large use of 4

chitosan in drug delivery, tissue engineering and wound dressing. The amine groups of chitosan are responsible for all of these properties, especially for the antimicrobial activity (Li, Wu, & Zhao, 2016). The presence of amine groups also permits an easy chitosan functionalization. In this context, chitosan-catechol conjugates have been developed primarily by reacting amine groups with catechol-bearing molecules, through the formation of either amide bonds or catechol-amine adducts (Michael type or Schiff base) (Ghadban et al., 2016; Yavvari & Srivastava, 2015; Ryu et al., 2015; Zvarec, Purushotham, Masic, Ramanujan & Miserez, 2013). The effects of CS functionalization with catechol moieties on CS properties have been investigated, especially with regard to biocompatibility, tissue adhesion properties and mechanical strength (Ryu et al., 2015). However, there is a lack of information on how catechol derivatization of CS may affect the antimicrobial properties of the polymer itself. Indeed, since the primary mechanism of chitosan antimicrobial action is the interaction of its protonated amine groups with the anionic bacterial cell membrane (Carmona-Ribeiro & de Melo Carrasco, 2013), a weakening of the CS antimicrobial properties is possible as a consequence of the functionalization of the amine groups with catechol moieties. However, that is not a foregone conclusion because catechol compounds may have some antimicrobial activity, as demonstrated for hydroxytyrosol and caffeic acid (Jeong, Jeon, Lee, & Lee, 2009; Almajano, Carbò, Delgado & Gordon, 2007; Bisignano et al., 1998; Wahdan, 1998). The understanding of the effects of catechol groups on CS antimicrobial properties is of fundamental importance for application of catechol-chitosan conjugates as bioadhesive materials in wound dressing or tissue engineering. Indeed, in these applications, bacterial contamination often occurs with a consequent development of persistent infections (Percival et al., 2012; Percival, McCarty & Lipsky, 2015). Therefore, we planned experiments to investigate the antimicrobial activity of cathecol-chitosan derivatives at different catechol content against a strain of Staphylococcus epidermidis, an opportunistic pathogen involved in numerous nosocomial infections including those related to skin wounds and implanted medical devices.

5

Catechol-chitosan derivatives at different degrees of functionalization were synthesized by reacting CS amine groups with hydrocaffeic acid (HCAF, Figure 1) utilizing carbodiimide chemistry. HCAF belongs to the class of phenolic acids and can be extracted from different plants and fruits (Shahidi & Nacsk, 1995). In order to investigate the role of the catechol group in the antimicrobial activity, chitosan was also functionalized with phloretic acid (PHLO, Figure 1), a compound structurally similar to HCAF but with a phenolic group instead of a catechol one.

A

B CH3 * O HO

O

OH O

NH O

HO

x

*

O

NH2 OH

y

C

Figure 1. Chemical structure of phloretic acid (A), hydrocaffeic acid (B) and chitosan (C). In chitosan, x=0.85 and y=0.15.

The antimicrobial activity of the derivatives was investigated by the disk diffusion test and determination of the minimum inhibitory concentration. The ability of catechol-functionalized 6

chitosan in preventing bacterial adhesion was also investigated by layering the chitosan derivatives onto commercial dressings. Finally, since catechol bearing molecules are known to possess antioxidant properties, all of the obtained CS derivatives were also characterized in terms of antioxidant activity, by employing 2,2-Diphenyl-1-picrylhydrazyl (DPPH) as the free radical. The antioxidant activity, in combination with the antimicrobial one, may be relevant for application of the CS-derivatives in wound dressing, since free radicals, produced during the inflammatory response of the body to a pathogen, have been shown to favor diversity and adaptability in microbial biofilm communities (Boles & Singh, 2008).

2 Materials and Methods

2.1. Materials Chitosan

(CS,

degree

of

deacetylation

85%,

low

molecular

weight);

3-(4-

hydroxyphenyl)propionic acid (Phloretic acid, PHLO); 3,4-dihydroxyhydrocinnamic acid (Hydrocaffeic acid, HCAF); N-(3-dimethylaminopropyl)-N’-ethyl-carbodiimide hydrochloride (EDC), 2,2-Diphenyl-1-picrylhydrazyl (DPPH) were purchased from Sigma Aldrich as were all other solvents and reagents. DMSO D6100% were supplied from CIL (Cambridge Isotope Laboratories, Inc). All chemicals were of analytical grade and used as received.

2.2. CS functionalization with phloretic acid (PHLO) CS functionalization with PHLO was obtained via carbodiimide-mediated condensation between carboxyl groups of PHLO and amine groups of chitosan. Briefly, carboxylic group of PHLO was activated by adding EDC in stoichiometric ratio. Later, the activated PHLO was added to a 1% acetic acid aqueous solution of CS. Various CS/PHLO molar ratios (1/0.5, 1/1, 1/2) were employed. The mixture was stirred for 24 hr at room temperature. After reaction, the solution was dialyzed in water (membrane cutoff =35.000 Da) to remove acetic acid and low molecular weight 7

by-products. Finally, the polymer was recovered by freeze-drying. The obtained materials were named CS-PHLOx where x indicated the functionalization degree.

2.3. CS functionalization with hydrocaffeic acid (HCAF) To avoid by-product formation during CS functionalization, the HCAF catechol group was first protected by acetonide formation and then reacted with CS. Reaction pathway scheme for HCAF protection is shown in Figure 2.

O

O H2SO4

HO

+

OH

H3C OH

HO

O

CH3

reflux

HO

2h

HO

1

2 O

HO

CH3

O

DMP

O

O

O reflux

HO

2h

H3C

O

H3C

2

3 O

O

O H3C H3C

O

CH3

O CH3

LiOH (H2O)

OH

O CH3OH

3

H3C H3C

O

4

Figure 2. Reaction pathway scheme for protection of HCAF catechol group by acetonide formation.

The carboxylic group of HCAF was first converted in the corresponding methyl ester by Fischer esterification. For this aim, few drops of concentrated H2SO4 were added to a solution of HCAF (0.290 g – 1.6 mmol) in methyl alcohol (MeOH, 25 ml), under nitrogen atmosphere in the dark. Reaction was carried out for 2h. After solvent evaporation, the residue was dissolved in ethylacetate (EtOAc) and washed with NaHCO3. Then, an extraction of the aqueous phase with 8

EtOAc for three times was performed. Organic extracts were washed with brine and dried over Na2SO4. Finally, solvent was removed under reduced pressure to give Product 2 (Figure 2) (yield = 95%).1H-NMR:(300 MHz, CDCl3) δ: 6.93 – 9.95 (m, 3H, ArH), 3.67 (m, 3H), 2.63 (m, 2H), 2.43 (m, 2H). In the second step, involving the formation of the acetonide protecting group, 2,2dimethoxypropane (0.33 g) and p-toluenesulfonic acid (pTsOH, 0.07 g) were added to the Product 2 dissolved in anhydrous chloroform. Reaction was carried out for 2 hr. Then, after neutralization of the mixture with NaHCO3 (saturated solution), the aqueous phase was extracted with anhydrous chloroform. The organic extracts were then washed with brine and dried over Na2SO4. After filtration, solvent was evaporated in vacuum and Product 3 (Figure 2) was isolated by column chromatography (yield = 60%). 1H-NMR:(300 MHz, CDCl3) δ: 6.93 – 9.95 (m, 3H, ArH), 3.67 (m, 3H), 2.63 (m, 2H), 2.43 (m, 2H), 1.65(s, 6H). In the final step, saponification of the Product 3 was performed. After dissolution of the Product 3 in MeOH (4 ml), an excess of LiOH (LiOH:Product 3 2:1 molar ratio) was added to the solution. The mixture was stirred for 24 hr at room temperature. After that time, pH of the mixture was adjusted to 5-6 by adding HCl 1M dropwise. Methanol was recovered in vacuum and solution and extracted with EtOAc (three times). Organic phases were collected and dried over Na2SO4. After filtration, the organic solvent was evaporated under vacuum to obtain the Product 4 (Figure 2) (yield = 95%).1H-NMR (300 MHz, CDCl3) δ : 6.93 – 9.95 (m, 3H, ArH), 2.63 (m, 2H), 2.43 (m, 2H), 1.65 (s, 6H). CS functionalization with acetonide-protected HCAF was carried out following a procedure similar to that described for PHLO. The carboxylic group of acetonide-protected HCAF was activated by adding EDC in stoichiometric ratio. Later, the activated HCAF was added to a 1% acetic acid aqueous solution of CS. Various CS/HCAF molar ratios (1/0.5, 1/1, 1/2) were employed. The mixture was stirred for 24 hr at room temperature. After reaction, the solution was dialyzed in water and the polymer was recovered by freeze-drying. Deprotection of the catechol group of HCAF functionalized-CS was performed by treating the polymer with HCL 1M for 18 hr at room

9

temperature. Deprotection was confirmed by 1H-NMR and UV spectroscopy. The obtained derivatives were named CS-HCAFx where x indicated the functionalization degree.

2.4 Evaluation of CS functionalization 1

H-NMR spectra were performed employing a Varian XL 300 instrument and D2O or

CD3COOD as solvents. Attenuated Total Reflection spectra (ATR) were acquired by a Nicolet 6700 FT-IR spectrometer equipped with a Golden Gate diamond single reflection device (Specac). Spectra were obtained by co-adding 200 interferograms in the range 4000–650 cm-1 at a resolution of 4 cm-1. UV–Vis spectroscopic analysis was performed by using a HP U2000 singular beam spectrophotometer working in the 190–1100 nm wavelength range and with a resolution of 0.004 nm.

2.5 Evaluation of the antioxidant activity of CS-derivatives DPPH (2,2-Diphenyl-1-picrylhydrazyl) free radical method is an antioxidant assay based on electron transfer. DPPH is a stable radical that turns from violet to yellow when scavenged. Reaction of DPPH with antioxidants lead to its reduction to DPPH-H. As a consequence, absorbance at 520 nm decreases. For each compound, different concentrations were tested (expressed as compound/DPPH molar ratio). CS and CS derivatives were dissolved in water (1% acetic acid) and added (4 mL) to a 1.5x10-4 mol/L methanol DPPH solution (10 mL). The antioxidant activity of pure PLHO and HCAF was also determined. The decrease in absorbance was determined at room temperature at 520 nm after 50 min (for CS and CS derivatives) or 30 min (for PHLO and HCAF), times in which a plateau was reached. The amount of DPPH was evaluated from a calibration curve. For each concentration, the residual DPPH as a function of compound/DPPH molar ratio was plotted. Scavenging activity was defined as the amount of compound needed for decreasing the initial DPPH concentration by 50% (Effective Concentration = EC50). 10

2.6 Evaluation of the antimicrobial activity of CS-derivatives. The antibacterial activity of CS-PHLOx and CS-HCAFx was assessed against S. epidermidis (ATCC 35984), an opportunistic pathogen often involved in nosocomial infections. The antimicrobial activity of pure CS and CS-derivatives was first evaluated by the disk diffusion test (Francolini, Piozzi & Donelli, 2014). A Tryptic soy agar Petri dish was inoculated with a bacterial inoculums at a 108 CFU/ml concentration (optical density = 0.125 at 550 nm). Then, cellulose disks impregnated with a solution of each sample at a 0,5 mg/mL concentration were placed onto the surface of the agar plate. Following incubation of the plates at 37 °C for 18 hr, the inhibition halo around each cellulose disk was measured. Then, the minimum inhibitory concentration (MIC) of pure CS, and CS-derivatives was assessed by the broth dilution test (Francolini, Piozzi & Donelli, 2014). Briefly, a bacterial inoculum at 1x106 CFU/mL in Muller-Hinton broth (M-H) with an optical density of 0.05 at 550 nm was first prepared. Then, a solution (1 mL) of either CS or CS derivatives at various concentrations (0.06 ÷ 2 mg/mL) was added to test tubes containing the bacterial inoculum (1 mL). A control tube with the bacterial inoculum (1 mL) and MH broth (1 mL) was also prepared. Control and test tubes were incubated at 37 °C for 24 hr. Following incubation, bacterial growth was determined by measuring the absorbance at 550 nm. Finally, the ability of CS-derivatives to prevent microbial adhesion was evaluated by coating commercial cellulose dressings with either pure CS or selected CS-derivatives. Briefly, samples of cellulose dressings (1x1 cm) were dipped into polymer aqueous solution (10 mg/mL, 500 µL) several times until complete absorption of the solution. After drying, the coated dressings were immersed in 1 mL S. epidermidis suspension in MH broth (0,5 MacFarland, optical density 0.125 at 550 nm). Following incubation at 37 °C for 24 h, bacteria adhered to the dressings were either detached by sonication to determine the number of colony forming units per surface area or fixed for Field Emission Scanning Electron Microscopy (FESEM) observations. 11

As for the determination of the number of adhered CFUs, the dressings were first washed twice with PBS (pH = 7.4), then inserted in vials containing 5 ml PBS and finally sonicated for 5 min at room temperature to detach adherent bacteria. After sonication, five serial dilutions in PBS were prepared and placed (10 µL) onto MH agar plates. After 18 hr incubation at 37°C, the colonies in the first dilution in which they were well separated were counted. Considering the dilution factor and the surface area of the treated dressings, the number of colony-forming units per sample surface (CFUs/cm2) was determined. Experiments were performed in n = 5 samples and data were reported as mean ± SD. Differences were considered significant for P values of < 0.05. As for FESEM observations, dressings

were washed twice with PBS, fixed with 2.5%

glutaraldehyde in 0.1 M PBS buffer at room temperature for 30 min, dehydrated through graded ethanol, treated with hexamethyldisilazane for 20 min and gold sputtered.

3. Results and Discussion

In the last decade, adhesive materials have been shown to hold great potentiality for tissue engineering, drug delivery and wound healing (Ryu, Hong & Lee, 2015). In all of these applications, much attention has been paid to chitosan, (Hurler & Skalko-Basnet, 2012; Peh, Khan & Ch'ng, 2000). . To increase mucoadhesivity, CS has been either blended with polymers (Bumgardner, Haggard & Jennings, 2017; Hoque, Prakash, Paramanandham, Shome & Haldar, 2017) or functionalized with catechol moieties to produce a protein mussels mimics polymer (Xu et al., 2017). Catechol-chitosan derivatives have shown to retain biocompatibility (Qiao et al., 2014) and to display superior mucoadhesive properties (Kim, Kim, Ryu, & Lee, 2015; Xu, Soliman, Barralet, & Cerruti, 2012) as well as greater tensile strength (Ryu, Jo, Koh, & Lee, 2014) compared to pure chitosan. However, limited information is available on the antimicrobial activity of cathecol-chitosan derivatives. Especially in wound healing applications, an intrinsic antimicrobial activity of the adhesive material is important because of the possible infection of the body site 12

interested by the application of the material (Percival et al., 2012; Percival, McCarty & Lipsky, 2015). Therefore, a balance between the adhesive properties and the antimicrobial activity should be accomplished for a successful application of catechol-functionalized chitosan as a bioadhesive. This study aims to investigate how catechol functionalization of chitosan may affect the antimicrobial activity of the polymer itself towards the opportunistic pathogen Staphylococcus epidermidis, often involved in skin wound infections. To this aim, chitosan was functionalized with hydrocaffeic acid (Figure 1) by amidation of CS amine groups. As negative control, chitosan was also functionalized with phloretic acid (Figure 1), a compound structurally similar to HCAF but with a phenolic group instead of a catechol one.

3.1 Determination of degree of CS functionalization Reactions involving catechol bearing molecules are always tricky because of the high reactivity of such compounds. In this work, we have set up a synthetic procedures to protect catechol group by acetonide formation before CS functionalization, providing a new protocol and good product yields. A qualitative estimation of chitosan functionalization was performed by IR-ATR spectroscopy. As an example, in Figure 3A the IR spectra for CS, CS-PHLO and CS-HCAF derivatives obtained at 1/1 molar ratio, are reported. In CS spectrum, signals at 3750 e 3000 cm-1, related to OH and NH group stretching, and those at 2920 e 2875 cm-1, assigned to C-H stretching of CH2 and CH3 groups, along with bending vibrations of the same groups present in the range at 1375- 1425 cm-1 were observed. Absorptions at 1680-1480 cm-1 were related to the stretching vibration of secondary amide C=O group, belonging to acetylated repetitive unit (amide I band, 1645cm-1, and secondary amine N-H bending, at 1599 cm-1). Finally, the C-O-H and C-O-C stretching signals were observed at 1150-1000 cm-1 and the glycosidic bond wagging (C-O-C) was observed at 895 cm-1.

13

Abs (U.A.)

Absorbance Ratio (A1640cm-1/A895cm-1)

A

CS-PHLO 1/1

CS-HCAF 1/1

CS

4000

18

B

16 14 12 10 8 6 4 2 0 CS

3600

3200

2800

2400 

2000

1600

1200

800

1/0.5 CS-HCAF

1/1

1/2

CS-PHLO

(cm-1)

Figure 3. IR spectra of pure CS, CS/PHLO 1/1 and CS/HCAF 1/1 (A); Absorbance ratio between the amide C=O signal (1640 cm-1) and the C-O-C signal (895 cm-1) for CS-PHLOs and CS-HCAFs (B).

In the spectra of CS derivatives, the success of functionalization was confirmed by the presence of the signals related to the aromatic ring of either PHLO or HCAF, specifically at 1514 cm-1 the C-C aromatic stretching and at 825 cm-1 the C-H aromatic bending. Additionally, the increase in the intensity of the signals in the 1680-1480 cm-1 region confirmed the amide bond formation. In order to evaluate the trend of the degree of CS functionalization as a function of the employed CS/PHLO or CS/HCAF molar ratio, the absorbance ratio between either the amide C=O signal (1640 cm-1) and the signal of the CS pyranose ring C-O-C (895 cm-1) was determined for each sample (Figure 3B). As it can be observed, in both series of derivatives the amidation degree increased with increasing CS/PHLO or CS/HCAF molar ratio. 1

H-NMR spectroscopy was employed to evaluate the CS functionalization degree. In Figure

4, as an example, the spectrum of CS-PHLO obtained at 1/1 molar ratio is reported and compared with that of pure chitosan. In the spectrum of CS, signals related to CH3 residue at 1.8 ppm and to H2, H3, H4, H5, H6 protons of CS backbone in the range 2.5-4.7 ppm were present. In the CS14

PHLO spectrum, new signals related to the aromatic protons of the phenolic moiety, in the range 7.5-6.5 ppm, were observed. In CS-HCAF samples having protected catechol group, the signal related to the six cyclic ketal protons was also observed (spectrum not shown). The CS functionalization degrees, calculated by the integral ratio between the signal at 1.8 ppm of the CH3 residue, and the signals of the aromatic protons, are reported in Table 1. In all cases, the functionalization degree increased with CS/PHLO or CS/HCAF molar ratio. A complete CS functionalization occurred only with PHLO. Probably, the increased steric hindrance of protectedHCAF reduced reactivity of the molecule.

A

B 2 5

1

4 6

Figure 4: 1H-NMR spectrum of CS-PHLO 1/1 (A), and pure chitosan (B).

The catechol function in CS-HCAF derivatives was retrieved by acidic treatment. The success of such a treatment was confirmed by both 1H-NMR spectroscopy by observing the disappearing of the signal related to methyl protons of cyclic ketal at 1.4 ppm (spectrum not shown) and UV-vis spectroscopy. In Figure 5, the UV spectra of CS-HCAF before and after protection of catechol moiety are reported. Particularly, HCAF displays π-π* transitions, related to the aromatic ring, around at 210 nm and in the 250-300 nm range (peak centered at 280 nm). In addition, a shoulder at 15

about 225 nm (n-π* transitions) is also present and attributed to the carbonyl group of the molecule. After protection of catechol groups through acetonide formation, a bathochromic effect of the peak centered at 280 nm was observed. That displacement was related to oxygen atom molecular orbital alignment with π system that leads to an energy reduction for the corresponding transition. The reduction of the band intensity same transition (hypocromic effect) wass related to a steric hindrance of the cyclic ketal greater than that of the catechol group. A displacement of the shoulder centered at 225 nm to higher wavelengths (from 225 to about 240 nm) was also observed and related to the carbonyl groups not linked to OH groups. After treatment with HCl 1 M for 18 hours, the de-protected HCAF spectrum became superimposable to that of HCAF, confirming the efficacy of the acidic treatment in the removal of acetonide group.

Protected CS-HCAF5 Deprotected CS-HCAF 5

Abs (AU)

HCAF

200

250

300 wavelength (nm)

350

400

Figure 5: UV spectra of pure HCAF and CS-HCAF5 protected and deprotected by acidic treatment.

3.2 Evaluation of antioxidant activity of CS-derivatives It is known that molecules displaying phenolic groups are endowed with radical scavenging properties. Therefore, an increase in the CS antioxidant activity is an indirect evidence of the success of CS functionalization. On the other hand, the antioxidant activity is very important in 16

many biological applications (Losada-Barreiro & Bravo-Diaz, 2017). In wound dressing, the use of an antioxidant material may contribute to the resolution of the inflammation process of the wound (Pohanka, 2013). Additionally, free radicals have been also shown to promote antibiotic resistance in biofilm-growing bacteria, as recently demonstrated in different biofilm communities (Boles & Singh, 2008). The antioxidant activity of CS-derivatives was evaluated by the DPPH method, which is widely used in the characterization of scavenging activity of phenolic and catechol compounds (Curcio et al., 2009). Firstly, the EC50 values for pure PHLO and HCAF were evaluated and resulted to be 5.4 mM and 0.02mM, respectively. As expected, HCAF possesses an antioxidant activity higher than PHLO and comparable to those of vitamin C (16.9 µM) and resveratrol (10.9 µM)(López-Burillo et al., 2003). Pure CS showed a high EC50 value (~10 mg/ml), consistent with data reported by Yen et al. (Yen, Yang & Mau, 2008), confirming the poor scavenging activity of this polysaccharide.

Table 1: Functionalization degree (FD), acronyms, EC50 values, inhibition halo (determined at 0.5 mg/mL polymer solution), and minimum inhibitory concentrations of the obtained CS-derivatives.

Polymer

CS/PHLO or CS/HCAF molar ratio

FD (%)

Acronym

EC50 (mg/mL)

Inhibition halo (mm)

MIC (mg/mL)

Chitosan

-

0

CS

11 ± 1

2±1

0.5

1/0.5

20

CS-PHLO20

6±1

0

>2

1/1

90a

CS-PHLO90

2±1

0

ND

1/2

97a

CS-PHLO97

NDb

ND

ND

1/0.5

5

CS-HCAF5

0.06 ± 0.01

6±1

ND

1/1

12

CS-HCAF12

0.045 ±0.005

ND

ND

1/2

20

CS-HCAF20

0.020 ± 0.005

4±1

0.125

CS-PHLO

CS-HCAF

17

a

Polymer was partial insoluble

b

ND = Not Determined

As far as the CS-derivatives are concerned, they resulted to be good scavengers, exerting their action depending on the degree of functionalization (Table 1). Indeed, the EC50 values of the CSderivatives decreased (higher activity) with the increase of the PHLO or HCAF content in the polymer. The best antioxidant properties were displayed by the CS-HCAF derivatives, for which EC50 values ranged from 20 to 60 µg/ml. That means, CS-HCAF derivatives possessed an antioxidant activity from 200 to 700 times greater than CS depending on the functionalization degree. This good antioxidant activity confirmed the efficacy of the proposed synthetic strategy.

3.3 Evaluation of antimicrobial activity of CS derivatives The antimicrobial activity of CS-derivatives was tested versus S. epidermidis. Firstly, the disk diffusion test on agar plates was performed to determine the inhibition halo around disks impregnated with the samples (Figure 6A). The diameters of the observed inhibition halos are reported in Table 1. The small halo produced by CS (2 mm) is presumably due to the poor diffusion of the polymer in the agar medium, while the absence of inhibition halo around the disks impregnated with the CS-PHLO samples indicates the antimicrobial inefficiency of these samples. This is probably due to the reduction in the content of the CS NH2 groups after functionalization with PHLO, being the NH2 groups responsible for CS antimicrobial activity (Zheng & Zhu, 2003). In contrast, all of the CS-HCAFs displayed an inhibition halo. Interestingly, the CS-HCAF samples having protected catechol groups did not show inhibition halo (Figure 6), This behavior could be related to the poor solubility of samples having protected catechol groups compared to free CSHCAF samples. Indeed, it is known that catechol functionalization of chitosan dramatically increases water solubility of the polymer up to 60 mg/mL also at neutral pH, unlike pure chitosan

18

(Kim, Ryu, Lee, & Lee, 2013). An intrinsic activity of free catechol moiety could also contribute to HCAF antimicrobial activity. To better investigate this phenomenon, the MICs of the derivatives were also determined and compared with those of pure CS. In these experiments, only CS-PHLO20 and CS-HCAF20 were considered because having the same degree of functionalization. In addition, the CS-PHLO samples at higher functionalization degrees resulted poorly water soluble. CS-PHLO20 was not active at the concentrations tested in our experiments (Table 1). In contrast, CS-HCAF20 showed a good antimicrobial activity with a MIC value significantly lower than that of CS. This increase in the antimicrobial activity of CS evidenced a synergistic action between chitosan and HCAF against S. epidermidis.

A CS-HCAF5 p*

CS-HCAF5 dep**

CS-PHLO20

CS

CS-PHLO90

19

B

Figure 6: (A) Inhibition halos around disks impregnated with either CS or the different CSderivatives (p*= Protected catechol moiety; dep**= Deprotected catechol moiety); (B) Number of S. epidermidis CFUs adhered per cm2 of dressing, uncoated or coated with the different CSderivatives. Asterisks indicate significant differences between trial types, *p<0.05.

Although chitosan has been already functionalized with antioxidant phenolic compounds (Curcio et al., 2009; Bozic, Gorgieva & Kokol, 2012; Brzonova, Steiner, Zankel, Nyanhongo & Guebitz, 2011; Soliman, Zhang, Merle, Cerruti & Barralet, 2014), the correlation between the antimicrobial activity and the presence of catechol moieties able to increase chitosan solubility is novel. Indeed, the good solubility of the cathecol-CS conjugates could permit to the chitosan chains to better diffuse in water and interact more efficiently with the bacterial cell membrane. Additionally, catechol moieties could have an intrinsic antimicrobial activity, as reported for some catechol bearing natural extracts (Bisignano et al., 1999). The mechanism is still unclear. However, it seems to be related to the ability of catechols to induce a permeability change of membranes which was attributed by fatty acids changes (Sousa, Guebitz & Kokol, 2009; Park, Ko & Kim, 2001; Ramos, Duque, Huertas & Haidour, 1995). Alongside this, Schweigert et al. (2001) reported that catechols 20

can act both as antioxidants, preventing lipid peroxidation, and as pro-oxidants, damaging macromolecules incuding DNA and proteins.

3.4 Evaluation of the antiadhesiveness properties of CS derivatives To evaluate the possible activity of CS derivatives in preventing microbial adhesion and biofilm formation, commercial cellulose dressings were first coated with CS-HCAF and CS-PHLO derivatives and then left in contact for 24 hr with a S. epidermidis suspension. After incubation, bacteria adhered to the dressing surface were either fixed for SEM observation or detached by sonication and counted. In Figure 6B, the number of CFU adherent per dressing surface unit (CFU /cm2) for the samples after 1 day of incubation is reported. Uncoated and CS-coated dressings were used as controls. As it can be noted, bacterial adhesiveness on the dressing coated with pure CS was significantly lower compared to the uncoated dressing (ca. 2-log reduction in bacterial adhesion compared to the uncoated dressing, p<0.01), confirming the good CS antimicrobial activity. Dressings coated with CS-PHLO20 had a lower ability to prevent microbial adhesion compared to the CS-coated dressing, in accordance with both the MIC results and inhibition halos. On the contrary, dressings coated with CS-HCAF20 showed superior anti-adhesiveness properties than CS (1-log reduction, p<0.05). These findings were confirmed by SEM observations (Figure 7).

A

B

21

C

D

E

F

Figure 7. SEM micrographs of uncoated dressing (A,B), CS-coated dressing (C,D,E) and CSHCAF20 coated dressing (F) after 24 hr-incubation with S. epidermidis. White arrows indicate the presence of single cells.

Specifically, a heavy colonization of the surface of the uncoated dressing was observed with the presence of both single colonies (Figure 7A) and large biofilm structures (Figure 7B). The CScoated dressing surface was highly rough (Figure 7C) presumably related to the deposition of CS nano- and micro-aggregates on the dressing surface, induced by solvent evaporation. S. epidermidis was able to adhere to the CS-coated dressing (Figure 7D) and form large bacterial aggregates (Figure 7E). The roughness of CS-HCAF20-coated dressing surface was significantly lower (Figure 7F) than that of the CS-coated dressing, suggesting a CS-HCAF20 better affinity to the dressing 22

presumably thanks to the catecholic moieties. The CS-HCAF20-coated surface was essentially free from S. epidermidis colonization, with just few sporadic adhering cells (Figure 7F).

4. Conclusions This research was focused on the investigation of the antimicrobial activity of catecholfunctionalized chitosan for potential wound dressing application. Chitosan, largely employed for this purpose, was functionalized with hydrocaffeic acid, bearing a catechol group, via carbodiimidemediated condensation. An easy and efficient procedure for acetonide-protection of the catechol moiety of hydrocaffeic acid was set up. The CS-HCAF derivatives resulted to be the best systems in terms of combined antimicrobial and antioxidant activity. Particularly, CS-HCAF20 displayed a fourfold lower MIC than CS and an EC50 700 times lower than CS. The higher antimicrobial activity of CS-HCAF20 resulted in a reduction of S. epidermidis adhesion on commercial dressings compared to CS. Taken as a whole, these findings suggest how the introduction of catechol moieties into chitosan may represent a way not only to increase muco-adhesiveness of the polymer but also to enhance its antimicrobial and antioxidant activity, all important features for application in wound healing.

Funding This work was supported by the Sapienza University of Rome, through a grant to I.F.

Conflicts of Interest The authors declare no conflict of interest.

References

23

Almajano, M. P., Carbò, R., Delgado, M. E., Gordon, M. H. (2007). Effect of pH on the antimicrobial activity and oxidative stability of oil-in-water emulsions containing caffeic acid. Journal of Food Science, 72, 258-263. Bisignano, G., Tomaino, A., Lo Cascio, R., Crisafi, G., Uccella, N., Saija, A. (1999). On the invitro antimicrobial activity of oleuropein and hydroxytyrosol. Journal of Pharmacy and Pharmacology, 51, 971-4. Boles, B. R., Singh P. K. (2008). Endogenous oxidative stress produces diversity and adaptability in biofilm communities. Proceedings of the National Academy of Sciences of the USA, 105, 12503-8. Bozic, M., Gorgieva, S., Kokol, V. (2012). Laccase-mediated functionalization of chitosan by caffeic and gallic acids for modulating antioxidant and antimicrobial properties. Carbohydrate Polymers, 87, 2388-2398. Brzonova, I., Steiner, W., Zankel, A., Nyanhongo, G. S., Guebitz, G. M. (2011). Enzymatic synthesis of catechol and hydroxyl-carboxic acid functionalized chitosan microspheres for iron overload therapy. European Journal of Pharmaceutics & Biopharmaceutics,79(2), 294-303. Bumgardner, J. D., Haggard, W. O., Jennings, J. A. (2017). Evaluation of a chitosan-polyethylene glycol paste as a local antibiotic delivery device. World Journal of Orthopedics, 8, 130-141. Carmona-Ribeiro, A. M., de Melo Carrasco, L. D. (2013). Cationic antimicrobial polymers and their assemblies. International Journal of Molecular Science, 14, 9906–9946. Curcio, M., Puoci, F., Iemma, F., Parisi, O. I., Cirillo, G., Spizzirri, U. G., Picci, N. (2009). Covalent insertion of antioxidant molecules on chitosan by a free radical grafting procedure. Journal of Agricultural& Food Chemistry, 57(13), 5933-8. Danner, E. W., Kan, Y., Hammer, M. U., Israelachvili, J. N., Waite, J. H. (2012). Adhesion of mussel foot protein Mefp-5 to Mica: an underwater superglue. Biochemistry, 51, 6511–6518. Duttagupta, D. S., Jadhav, V. M., Kadam, V. J. (2015). Chitosan: a propitious biopolymer for drug delivery. Current Drug Delivery, 12, 369-81.

24

Favi, P. M., Yi, S., Lenaghan, S. C., Xia, L., Zhang, M. (2014). Inspiration from the natural world: from bio-adhesives to bio-inspired adhesives. Journal of Adhesion Science and Technology, 28, 290–319. Forooshani, P. K., Lee, B. P. (2017). Recent approaches in designing bioadhesive materials inspired by mussel adhesive protein. Journal of Polymer Science Part A: Polymer Chemistry, 55, 9-33. Francolini, I., Piozzi, A., Donelli, G. (2014). Efficacy evaluation of antimicrobial drug-releasing polymer matrices. Methods in Molecular Biology, 1147, 215–225. Ghadban, A., Ahmed, A. S., Ping, Y., Ramos, R., Arfin, N., Cantaert, B., Ramanujan, R. V., Miserez, A. (2016). Bioinspired pH and Magnetic Responsive Catechol-Functionalized Chitosan Hydrogels with Tunable Elastic Properties. Chemical Communications, 52, 697-700. Hoque, J., Prakash, R. G., Paramanandham, K., Shome, B. R., Haldar, J. (2017). Biocompatible injectable hydrogel with potent wound healing and antibacterial properties. Molecular Pharmaceutics, 4, 1218-1230. Hurler, J., Skalko-Basnet, N. (2012). Potentials of chitosan-based delivery systems in wound therapy: bioadhesion study. Journal of Functional Biomaterials, 3, 37-48. Jeong, E. Y., Jeon, J. H., Lee, C. H., Lee, H. S. (2009). Antimicrobial activity of catechol isolated from Diospyros kaki Thunb. roots and its derivatives toward intestinal bacteria. Food Chemistry, 115, 1006-1010. Kaushik, N. K., Kaushik, N., Pardeshi, S., Sharma, J. P., Lee, S. H., Choi, E. H. (2015). Biomedical and clinical importance of mussel-inspired polymers and materials. Marine Drugs, 13, 6792-6817. Kim, E., Song, I. T., Lee, S., Kim, J.-S., Lee, H., Jang, J.-H. (2012). Drawing sticky adenoassociated viruses on surfaces for spatially patterned gene expression. Angewandte Chemie International Edition, 51(23), 5598–5601. Kim, K., Kim, K., Ryu, J. H., Lee, H. (2015). Chitosan-catechol: a polymer with long-lasting mucoadhesive properties. Biomaterials, 52, 161-170.

25

Kim, K., Ryu, J. H., Lee, D. Y., Lee, H. (2013). Bio-inspired catechol conjugation converts waterinsoluble chitosan into a highly water-soluble, adhesive chitosan derivative for hydrogels and LbL assembly. Biomaterials Science, 1, 783-790. Lee, H., Lee, Y., Statz, A. R., Rho, J., Park, T. G., Messersmith, P. B. (2008). Substrateindependent layer-by-layer assembly by using mussel-adhesive-inspired polymers. Advanced Materials, 20(9), 1619-1623. Lee, Y., Lee, H., Kim, Y. B., Kim, J., Hyeon, T., Park, H., Messersmith, P. B., Park, T. G. (2008). Bioinspired surface immobilization of hyaluronic acid on monodisperse magnetite nanocrystals for targeted cancer imaging. Advanced Materials, 20(21), 4154-4157. Li, J., Wu, Y., Zhao, L. (2016). Antibacterial activity and mechanism of chitosan with ultra high molecular weight. Carbohydrate Polymers, 148, 200-5. Lih, E., Choi, S. G., Ahn, D. J., Joung, Y. K., Han, D. K. (2016). Optimal conjugation of catechol group onto hyaluronic acid in coronary stent substrate coating for the prevention of restenosis. Journal of Tissue Engineering, 7, 1-11. Long, Y., Xiao, L., Cao, Q. (2017). Co-polymerization of catechol and polyethylenimine on magnetic nanoparticles for efficient selective removal of anionic dyes from water. Powder Technology, 310, 24–34. López-Burillo, S., Tan, D. X., Mayo, J. C., Sainz, R. M., Manchester, L. C., Reiter, R. J. (2003). Melatonin, xanthurenic acid, resveratrol, EGCG, vitamin C and alpha-lipoic acid differentially reduce oxidative DNA damage induced by Fenton reagents: a study of their individual and synergistic actions. Journal of Pineal Research, 34(4), 269-77. Losada-Barreiro, S., Bravo-Díaz, C. (2017). Free radicals and polyphenols: The redox chemistry of neurodegenerative diseases. European Journal of Medicinal Chemistry, 133, 379-402. Padula, C., Nicoli, S., Aversa, V., Colombo, P., Falson, F., Pirot, F., Santi, P. (2007). Bioadhesive film for dermal and transdermal drug delivery. European Journal of Dermatology, 17, 309-12.

26

Park, S.-H., Ko, Y.-J., Kim, C.-K. (2001). Toxic Effects of Catechol and 4-Chlorobenzoate Stresses on Bacterial Cells. The Journal of Microbiology, 39(3), 206-212. Peh, K., Khan, T., Ch'ng, H. (2000). Mechanical, bioadhesive strength and biological evaluations of chitosan films for wound dressing. Journal of Pharmacy & Pharmaceutical Sciences, 3, 303-11. Percival, S. L., Hill, K. E., Williams, D. W., Hooper, S. J., Thomas, D. W., Costerton, J. W. (2012). A review of the scientific evidence for biofilms in wounds. Wound Repair and Regeneration, 20, 647–657. Percival, S. L., McCarty, S. M., Lipsky, B. (2015). Biofilms and wounds: an overview of the evidence. Advances in Wound Care, 4, 373–381. Pohanka, M. (2013). Role of oxidative stress in infectious disease. A review. Folia Microbiologica, 58, 503-513. Qiao, H., Sun, M., Su, Z., Xie, Y., Chen, M., Zong, L., Gao, Y., Li, H., Qi, J., Zhao, Q., Gu, X., Ping, Q. (2014). Kidney-specific drug delivery system for renal fibrosis based on coordinationdriven assembly of catechol-derived chitosan. Biomaterials, 35, 7157-7171. Ramos, J. L., Duque, E., Huertas, M. J., , Haidour A. (1995). Isolation and expansion of the catabolic potential of a Pseudomonas putida strain able to grow in the presence of high concentrations of aromatic hydrocarbons. Journal of Bacteriology, 177, 3911-3916. Ryu, J. H., Jo, S., Koh, M. Y., Lee, H. (2014). Bio-inspired, water soluble to insoluble self conversion for flexible, biocompatible, transparent, catecholamine polysaccharide thin films. Advanced Functional Materials, 24, 7709-7716. Ryu, J. H., Lee, Y., Kong, W. H., Kim, T. G., Park, T. G., Lee, H. (2011). Catechol-functionalized chitosan/pluronic hydrogels for tissue adhesives and hemostatic materials. Biomacromolecules, 12(7), 2653-9. Ryu, J.H., Hong, S., Lee, H. (2015). Bio-inspired adhesive catechol-conjugated chitosan for biomedical applications: a mini review. Acta Biomaterialia, 27, 101-115.

27

Schweigert, N., Zehnder, A. J., Eggen, R.I. (2001). Chemical properties of catechols and their molecular modes of toxic action in cells, from microorganisms to mammals. Environmental Microbiology, 3(2), 81-91. Shahidi, F., Nacsk, M. (1995). Food phenolics: Sources, chemistry, effects, and application. Technomic Publishing Company Incorporated Lancaster, 27, 245-278. Shin, M., Park, S. G., Oh, B. C., Kim, K., Jo, S., Lee, M. S., Oh, S. S., Hong, S. H., Shin, E. C., Kim, K. S., Kang, S. W., Lee, H. (2017). Complete prevention of blood loss with self-sealing haemostatic needles. Nature Materials, 16(1), 147-152. Soliman, G. M., Zhang, Y. L., Merle, G., Cerruti, M., Barralet, J. (2014). Hydrocaffeic acidchitosan nanoparticles with enhanced stability, mucoadhesion and permeation properties. European Journal of Pharmaceutics & Biopharmaceutics, 88(3), 1026-37. Sousa, F., Guebitz, G. M., Kokol, V. (2009). Antimicrobial and antioxidant properties of chitosan enzymatically functionalized with flavonoids. Process Biochemistry, 44, 749-756. Wahdan, H. A. (1998). Causes of the antimicrobial activity of honey. Infection, 26(1), 26-31. Xu, J., Soliman, G. M., Barralet, J., Cerruti, M. (2012). Mollusk glue inspired mucoadhesives for biomedical applications. Langmuir, 28, 14010-14017. Xu, J., Tam, M., Samaei, S., Lerouge, S., Barralet, J., Stevenson, M. M., Cerruti, M. (2017). Mucoadhesive chitosan hydrogels as rectal drug delivery vessels to treat ulcerative colitis. Acta Biomaterialia, 48, 247-257. Yavvari, P. S., Srivastava, A. (2015). Robust, Self-Healing Hydrogels from Catechol Rich Polymers. Journal of Materials Chemistry B, 3, 899-910. Yen, M., Yang, J., Mau, J. (2008). Antioxidant properties of chitosan from crab shell. Carbohydrate Polymers, 74, 840-44. Zheng, L-Y., Zhu, J-F. (2003). Study on antimicrobial activity of chitosan with different molecular weights. Carbohydrate Polymers, 54(4), 527–530.

28

Zvarec, O., Purushotham, S., Masic, A., Ramanujan, R. V., Miserez, A. (2013). CatecholFunctionalized Chitosan/Iron Oxide Nanoparticle Composite Inspired by Mussel Thread Coating and Squid Beak Interfacial Chemistry. Langmuir, 29, 10899-10906.

29