**Preface to "Marine Natural Products with Antifouling Activity"**

In a highly competitive marine benthic environment, competition for available space is an important survival strategy for many species. Therefore, antifouling compounds are important components of an arsenal used by many benthic sessile organisms in order to prevent fouling by other organisms. Additionally, among marine microorganisms, a huge diversity of secondary metabolites are produced and have been disclosed to be bioactive against different components of biofouling communities. This Special Issue covers different aspects of the chemistry of marine-based antifouling compounds and their use in various antifouling applications. This collection of articles includes research publications on the discovery of new agents against marine biofouling, namely a butenolide analogue (Boc-butenolide) (1), elasnin (2), and polymethoxylated flavones and chalcones (3); a cyanobacterial polymer-based coating as an antibiotic-free alternative strategy to fight catheterassociated urinary tract infections (4); antimicrobial poly(lactic acid) (PLA) surfaces coated with βchitosan and β-chitooligosaccharides for food packaging apllicatins (5); insights into the molecular mechanisms underlying the synthesis, secretion, and curing of barnacle cyprid adhesive as potential molecular targets for the development of environmentally friendly antifouling compounds (6); and two review articles focusing on new potential marine microbial-derived antibiotics and biosurfactants against catheter-associated urinary tract infections (7) and recent strategies for the development of marine ecofriendly coatings (8).

This compilation of selected scientific publications highlights the interdisciplinarity underlying the theme of antifouling strategies, and their wide biotechnological applicability.

> **Tom Turk, Joana Reis Almeida** *Editors*

### *Article* **Synthetic Analogue of Butenolide as an Antifouling Agent**

**Ho Yin Chiang 1, Jinping Cheng 1, Xuan Liu 1, Chunfeng Ma 2and Pei-Yuan Qian 1,3,\***

	- Guangzhou 510000, China; msmcf@scut.edu.cn

**Abstract:** Butenolide derivatives have the potential to be effective and environmentally friendly antifouling agents. In the present study, a butenolide derivative was structurally modified into Boc-butenolide to increase its melting point and remove its foul smell. The structurally modified Bocbutenolide demonstrated similar antifouling capabilities to butenolide in larval settlement bioassays but with significantly lower toxicity at high concentrations. Release-rate measurements demonstrated that the antifouling compound Boc-butenolide could be released from polycaprolactone-polyurethane (PCL-PU)-based coatings to inhibit the attachment of foulers. The coating matrix was easily degraded in the marine environment. The performance of the Boc-butenolide antifouling coatings was further examined through a marine field test. The coverage of biofouler on the Boc-butenolide coatings was low after 2 months, indicating the antifouling potential of Boc-butenolide.

**Keywords:** antifouling compounds; structural optimisation; butenolide; larval attachment assay

#### **1. Introduction**

Since the prohibition of tributyltin in 2008, many studies have attempted to discover novel antifouling compounds from marine natural products [1]. Many bioactive marine natural products have been screened and tested in recent decades, and several reviews on marine natural products and their synthetic analogues as antifouling compounds have been published [2–5]. Although many potent antifouling compounds have been discovered, those compounds are rarely commercialised. The low supply of antifouling compounds has hindered the development of antifouling paints based on the marine natural products [2,3,6,7]. Two solutions to the problem exist. The first solution is to explore marine natural products from microorganisms, as microorganisms can produce a wide range of bioactive secondary metabolites [8–10]. The convenience of bacterial cultivation and the mass production of metabolites in a short period of time have benefits over the extraction of compounds from the microorganism [6,7,10]. The second solution involves structural optimisation using organic synthesis [6,11]. Secondary metabolites extracted from organisms are often complex in structure; thus, they can be difficult to synthesise effectively in large quantities for commercial-scale usage [6,11]. By studying the structure– activity relationship of bioactive compounds isolated from organisms, pharmacophores that are responsible for antifouling abilities can be identified [11]. Optimisation of the compound's structure is performed with the goal of increasing its potency, decreasing the toxicity of the original compound [12], improving other physical or chemical properties of the compound and simplifying the chemical structure for chemical synthesis.

The antifouling compound 5-octylfuran-2(5H)-one (butenolide) has a melting point at 23 ◦C, which causes it to change from solid form to liquid at high ambient temperatures. In the present study, a butenolide derivative was modified with a Boc-protecting-group at the terminal of the alkyl side chain, *tert*-butyl (5-(5-oxo-2,5-dihydrofuran-2-yl)pentyl)carbamate

**Citation:** Chiang, H.Y.; Cheng, J.; Liu, X.; Ma, C.; Qian, P.-Y. Synthetic Analogue of Butenolide as an Antifouling Agent. *Mar. Drugs* **2021**, *19*, 481. https://doi.org/10.3390/ md19090481

Academic Editors: Orazio Taglialatela-Scafati, Tom Turk and Joana Reis Almeida

Received: 21 June 2021 Accepted: 21 August 2021 Published: 25 August 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

(Boc-butenolide) (Figure 1). This modification aims to improve its environmental stability and to remove its foul smell by increasing its melting point from 23 ◦C to 132 ◦C. A higher melting temperature above ambient temperature could lead to a more precise control of the proportion of antifoulant added during coating formulation. The modified Boc-butenolide (Figure 1) was further characterized for its melting point, stability and antifouling bioactivity using anti-larval settlement bioassays with larvae of barnacles (*Amphibalanus amphitrite*) and tubeworms (*Hydroides elegans*) in the laboratory. The modified Boc-butenolide was further formulated into an antifouling paint using polymer matrix poly(ε-caprolactone) based polyurethane (PCL-PU). Release-rate measurements of Bocbutenolide and a marine field antifouling test were conducted to evaluate the feasibility of using Boc-butenolide as an active ingredient in antifouling coatings.

**Figure 1.** Chemical structure of *tert*-butyl (5-(5-oxo-2,5-dihydrofuran-2yl)pentyl)carbamate (Bocbutenolide).

#### **2. Results and Discussion**

*2.1. Stability and Solubility of Boc-Butenolide and Butenolide*

Figure 2a illustrates the measured concentrations of Boc-butenolide in ASW for 3 months. The concentration of Boc-butenolide measured at day 0 is around 64 ppm and dropped to approximately 50 ppm after 3 months in ASW. In the following anti-larval settlement bioassay experiments, the same nominal concentrations for each condition were used to compare their bioactivity. To understand the relationship between nominal concentration and soluble or working concentrations of the two compounds, the working concentrations for all nominal concentrations used in the experiments were tested using HPLC. Figure 2b shows that the measured working concentrations of Boc-butenolide are higher than all tested working concentrations for butenolide under all tested nominal concentrations (3.125, 6.25, 12.5, 25, 50, and 100 ppm). The average dissolution rate of Boc-butenolide is approximately 40%, while the average dissolution rate of butenolide is only around 10%.

**Figure 2.** Measured concentrations of Boc-butenolide in ASW for 3 months (**a**), and working concentrations of butenolide and Boc-butenolide (**b**) used in Figures 3 to 6.

#### *2.2. Antifouling Performance of Boc-Butenolide*

Figure 3 shows the settlement rate of *A. amphitrite* larvae treated with Boc-butenolide and butenolide. At nominal concentrations of 50 ppm to 100 ppm, both compounds inhibited the settlement of barnacle cyprids. Settlement rates between the two compounds

began to differ at 25 ppm. For Boc-butenolide, the cyprids showed some settlement at a rate of approximately 3%. The settlement rate continuously increased, reaching approximately 70% at a concentration of 3.125 ppm. For butenolide, the attachment of cyprids was inhibited between the concentrations of 6.25 ppm and 100 ppm, and approximately only 10% of the larvae settled in the treatment of 3.125 ppm.

**Figure 3.** *A. amphitrite* larval settlement rate after Boc-butenolide and butenolide treatments. Asterisks indicate a significant difference from the control with *p* < 0.05.

Figure 4 shows the mortality rate of *A. amphitrite* larvae treated with Boc-butenolide and butenolide. For both compounds, there was no significant difference in toxic effects compared with the control group between the nominal concentrations of 3.125 ppm and 12.5 ppm. A pronounced difference in toxic effects was observed at 25 ppm. A very low toxicity was observed for Boc-butenolide-treated larvae, i.e., the mortality rate was approximately 8%, compared with the obvious toxic effect observed for the butenolide-treated larvae, in which mortality rate was over 90% among the individuals at high concentrations. All cyprids died in both treatments with higher antifoulant concentrations (50 and 100 ppm), indicating the toxic effects of the antifouling compounds at high concentrations.

**Figure 4.** *A. amphitrite* larval mortality rate after Boc-butenolide and butenolide treatments. Asterisks indicate a significant difference from the control with *p* < 0.05.

The *H. elegans* larval settlement bioassay results were similar to those of the *A. amphitrite* larval settlement bioassay. No settlement of larvae was observed after treatments with Boc-butenolide at 50 and 100 ppm (Figure 5), whereas for butenolide treatments, effective inhibition of larval settlement was found even at a low concentration of 12.5 ppm. For Boc-butenolide treatments, a significant difference from the control group was still observed at 25 ppm, with a settlement rate of approximately 20%. An increase in settlement inhibition was observed from 3.125 ppm to 25 ppm. For butenolide treatments, a significant difference was found between treatment and control groups across all concentrations, with a settlement rate of approximately 50% at low concentrations of 3.125 and 6.25 ppm.

**Figure 5.** *H. elegans* larval settlement rate after Boc-butenolide and butenolide treatments. Asterisks indicate a significant difference from the control with *p* < 0.05.

In terms of the toxicity of the compounds (Figure 6), only Boc-butenolide treatments at 50 and 100 ppm showed significant toxicity on the *H. elegans* larvae. However, for butenolide treatments, toxic effects started to appear at 12.5 ppm, reaching a 100% mortality rate at concentrations above 50 ppm. Structural differences between Boc-butenolide and butenolide affect the pharmacokinetics of the two compounds, leading to differences in potency and toxicity towards the larvae.

#### *2.3. Release Rate of Boc-Butenolide from the Coatings*

Figure 7 shows the release rate of different concentrations of Boc-butenolide and butenolide for at least 90 d. Generally, the amount of compounds released from the coatings at a particular time is positively correlated with the initial concentration of the compounds in the coatings. The release rate could be controlled by changing the concentration of antifoulant in the coatings. The initial release rate of Boc-butenolide (150 μg/cm2/day) was much higher than that of butenolide (45 μg/cm2/day) for 10 wt% samples. A possible explanation is the hydrophilicity difference between Boc-butenolide and butenolide. A Boc-protecting-group was added to the side chain of butenolide. Thus, the Boc-butenolide was more hydrophilic with the added highly electronegative N and O atoms of the Boc group and easier to dissolve in seawater, thereby resulting in a high initial release rate. A huge decrease in the release of antifoulant from the coatings for both Boc-butenolide and butenolide was found after 1 month. Notably, the release of Boc-butenolide from the coatings for all four concentrations was much lower than that of butenolide after 1 month. This finding might be due to the fact that a steady release in the later stage cannot be supported after a huge initial release of Boc-butenolide.

**Figure 6.** *H. elegans* larval mortality rate after Boc-butenolide and butenolide treatments. Asterisks indicate a significant difference from the control with *p* < 0.05.

**Figure 7.** Release-rate measurement of (**a**) Boc-butenolide and (**b**) butenolide with PCL-PU matrix for 90 d.

PCL-PU polymer is biodegradable and environmentally friendly, and was applied as the polymer to develop the antifouling coating system in the present work. In the previous study, a PCL-PU/butenolide antifouling coating system was developed and test results showed that the polymer coating could be degraded in the sea [13]. In this experiment, PCL-PU/Boc-butenolide showed a large decrease in release rate when compared to that of PCL-PU/butenolide, which might be due to the compatibility of Boc-butenolide in PCL-PU and the relatively high solubility of Boc-butenolide in seawater. In future studies, more effort should be made in the improvement of the polymer structure so as to optimize the release performance of Boc-butenolide as an antifouling coating.

#### *2.4. Field-Test Performance*

A field test was conducted to evaluate the performance of the coatings in the marine environment. Figures 8 and 9 show the images and the relative coverage of the panels coated with different concentrations of Boc-butenolide or butenolide with PCL-PU as a polymer matrix. After 1 month of exposure, all of the coatings with antifoulants remained almost fouling-free, with less than 10% coverage. Approximately 80% biofilm coverage was observed on the control panels. All of the panels treated with the coatings showed good antifouling performances, indicating the effectiveness of Boc-butenolide and butenolide as antifoulants. This finding is consistent with the larval settlement results, suggesting that the compounds can prevent larval attachment.

**Figure 8.** Field test of various concentrations of Boc-butenolide and butenolide with PCL-PU matrix for 2 months. From left to the right, the control, 1, 2.5, 5 and 10 wt% of Boc-butenolide (**top**) or butenolide (**bottom**) are presented. The test was continued for 2 months and retrieved at monthly intervals.

In the second month's results, the coatings with higher concentrations of Boc-butenolide or butenolide showed better antifouling abilities (Figure 9). Performance levels of both Boc-butenolide and butenolide were similar. These results were consistent with the release rate and larval attachment results mentioned above. Although Boc-butenolide was less potent in preventing larval settlement at similar nominal concentrations in the laboratory studies, its huge amount of initial release helped to compensate for its lack of potency, resulting in an antifouling performance similar to that of butenolide in the field test.

The differences in effectiveness and toxicity between Boc-butenolide and butenolide are possibly due to their structural difference. Considering that a Boc group was added to the carbon side chain of butenolide in Boc-butenolide, its structure was large and bulky. Molecular weight and hydrophilicity can affect the pharmacokinetics of the compounds, especially regarding their absorption by organisms [14]. Generally, a large molecule containing electronegative atoms, such as N or O, is more hydrophilic and is difficult to be absorbed or become bioavailable to the organisms. Therefore, Boc-butenolide shows a lower effectiveness when compared with butenolide. The difference in effectiveness was obvious at 25 ppm. At the same time, as Boc-butenolide has a lower absorption or bioavailability, it also shows no or relatively low toxic effects to the treated larvae. The structural modification of butenolide changes the physical and chemical properties of the original compound. For instance, Boc-butenolide is odourless and with a lower toxicity towards the larvae compared with butenolide. Boc-butenolide's chemical stability is also increased, as the Boc group is stable towards most nucleophiles and bases. These changes in physical and chemical properties can be beneficial when designing effective antifouling coatings and also when considering the environmental impact for actual use.

From the release-rate results, an exponential decrease in Boc-butenolide release into the seawater throughout the test period was observed. This result may be due to the high hydrophilicity of Boc-butenolide, which allows it to dissolve more easily in the seawater and achieve a high release rate. The release rate can be improved through the structural modifications of Boc-butenolide. Changes in physical and chemical properties, such as the melting point and hydrophilicity, can be achieved by structural modification [12]. For instance, the water solubility of Boc-butenolide can be reduced by a slight structural modification. Another method for optimising the controlled release of Boc-butenolide in seawater is by developing a suitable polymer as a binder. For example, the antifouling

compound could chemically bind to the polymer chain, e.g., in tributyltin-SPC antifouling coatings [15,16]. The release of biocide would then be controlled by the hydrolysis of the polymer, thereby controlling the release rate of the biocide.

#### **3. Materials and Methods**

#### *3.1. Chemicals and Seawater*

All chemical reagents used in this study, unless otherwise specified, were purchased from Sigma Aldrich (St. Louis, MO, USA) and VWR chemicals (Haasrode, Belgium). 5-Octylfuran-2(5H)-one (butenolide) and *tert*-butyl (5-(5-oxo-2,5-dihydrofuran-2-yl)pentyl)carbamate (Bocbutenolide) with a purity of >99% were purchased from ChemPartner (Shanghai, China) and used as received. Acetonitrile and methanol used were of HPLC grade.

Seawater was collected using a pump at the Coastal Marine Laboratory of Hong Kong University of Science and Technology. Filtered seawater (FSW) was obtained by filtering seawater through a 0.22 μm filter membrane from Millipore (Merck KGaA, Darmstadt, Germany). Artificial seawater (ASW) was prepared according to ASTM D114198 standards (2013) [17].

The stock concentrations of butenolide and Boc-butenolide were made by dissolving 100 mg of butenolide or Boc-butenolide in 1 mL of DMSO to make a stock of 100 mg/mL, stored at −20 ◦C. The butenolide and Boc-butenolide samples used in the larval settlement and mortality experiments were prepared by serial dilution of stock concentrations of butenolide and Boc-butenolide using ASW as the diluent, and the DMSO content in final samples were lower than 0.5‰ *v*/*v*.

#### *3.2. Collection of Amphibalanus amphitrite Larvae Sample*

Adult *A. amphitrite* colonies were collected from Tso Wo Hang Pier (22◦23 32.1 N 114◦17 18.7 E), Hong Kong, from April 2018 to June 2018. The adults were kept in a water tank with running seawater at the Coastal Marine Laboratory (the Hong Kong University of Science and Technology) for no more than a week before experimental use. Adults were induced to hatch under light sources for 1 h; the larvae were obtained using a method described previously by Harder et al. [18]. The nauplii larvae newly released from the adults were reared on diatom *Chaetoceros gracilis* Schütt. The seawater culture medium was replaced daily with fresh FSW and algae. The nauplii reached the competent stage, known as cyprid, after 4 d of incubation at approximately 28 ◦C.

#### *3.3. Collection of Hydroides elegans Larvae Sample*

Adult *H. elegans* colonies were collected from a fish farm at Yung Shue O, Hong Kong (22◦24 N, 114◦21 E) from March 2019 to April 2019. The adults were kept in a water tank with running seawater at the Coastal Marine Laboratory of Hong Kong University of Science and Technology for no more than 3 d before experimental use. The larvae were collected according to the methods described by Qian and Pechenik [19]. The tube of the adults was gently cracked with forceps to release the gametes. Oocytes were then mixed with the sperm and transferred into a new container with 500 mL FSW for fertilisation. Larvae were reared on microalga *Isochrysis galbana* (Tahitian strain) after hatching. The seawater culture medium was replaced daily with fresh FSW and algae, and the trochophore-stage larvae reached the competent stage after 5 d of incubation at approximately 25 ◦C.

#### *3.4. Larvae Food and Cultivation*

The diet for *A. amphitrite* and *H. elegans* cultivated in this study comprised *C. gracilis* and *Isochrysis galbana*, respectively. In the laboratory, the algae were cultured with Guillard's f/2 medium. The f/2 medium was prepared by adding designated amounts of NaNO3, NaH2PO4 H2O, trace metal and vitamin solutions into autoclaved FSW [20]. Na2SiO3 9H2O was added for the cultivation of *C. gracilis*. Algal stocks were then added into the

culture medium in a 2 L Erlenmyer flask and subcultured bi-weekly. The cultures were bubbled and illuminated under 14 h/10 h light/dark cycle at 23 ◦C for incubation.

#### *3.5. Settlement Bioassay of A. amphitrite*

The test compounds were dissolved with a small amount of DMSO. The test compounds were used in six concentrations from 100 ppm to 3.125 ppm with 2-fold serial dilution. The same amount of DMSO was used as the negative control for all testing concentrations. Approximately 20 ± 2 individual *A. amphitrite* cyprids were placed into each well of the 24-well polystyrene culture plate containing 2 mL of FSW and were subjected to different treatments. For all treatments and controls, three replicates were performed. The plates were then incubated at 25 ◦C in darkness. After 48 h, the number of settled and swimming larvae were counted using a Leica MZ6 microscope, and possible toxic effects were noted.

#### *3.6. Settlement Bioassay of H. elegans*

The test compounds were dissolved with a small amount of DMSO. The test compounds were used at six different concentrations with a 2-fold serial dilution from 100 ppm to 3.125 ppm. The same amount of DMSO was used as the negative control for all testing concentrations. Approximately 10 ± 2 individual *H. elegans* larvae were placed into each well of a 24-well polystyrene culture plate that contained 2 mL of FSW with different concentrations of the test solution. Approximately 10−<sup>4</sup> molarity of 3-isobutyl-l-methylxanthine was added into each well as an inducer for the settlement of *H. elegans* larvae [7]. The plates were then incubated at 25 ◦C in darkness. After 24 h, the number of settled and swimming larvae were counted using a Leica MZ6 microscope, and possible toxic effects were noted.

#### *3.7. Determination of Working Concentration and Stability Using High-Performance Liquid Chromatography (HPLC)*

The measurement and analysis of butenolide and Boc-butenolide were performed according to previous reports [13,21,22]. The preparation of nominal concentrations for butenolide and Boc-butenolide was described in the section of "Chemicals and seawater". The calibration standards were prepared by serial dilutions of stock concentrations of butenolide and Boc-butenolide using methanol as the diluent, and the DMSO content in all calibration standard samples was lower than 0.5‰ *v*/*v*. The stock concentrations of butenolide and Boc-butenolide were made by dissolving 500 mg of butenolide or Bocbutenolide in 1 mL of DMSO to make a stock of 500 mg/mL and stored at −20 ◦C. The working concentrations of butenolide and Boc-butenolide samples used in settlement bioassay were measured by reverse-phase HPLC using a Waters 2695 separation module coupled to a Waters 2669 photo-diode array (PDA) detector according to the peak area at 210 nm (Waters Corporation, Taunton, MA, USA). Identification of butenolide and Boc-butenolide was determined based on their retention times (butenolide, 11 ± 0.1 min; Boc-butenolide, 6.8 ± 0.1 min). The samples were tested with a 20 min gradient of 50–99% aqueous acetonitrile (ACN) containing 0.05% *v*/*v* trifluoroacetic acid (TFA) at a flow of 1 mL/min. The working concentrations of butenolide and Boc-butenolide were calculated according to their standard curves using peak areas plotted against known quantities of standards. The recoveries for butenolide and Boc-butenolide were 90.9% and 99.5%, respectively.

The stability of Boc-butenolide was measured by the concentration changes of Bocbutenolide in ASW throughout 3 months. The starting nominal concentration of Bocbutenolide was 200 ppm. At every time point, 5 mL of the solution was drawn and mixed with 10 mL dichloromethane (DCM). The DCM fraction with the analyte was dried under nitrogen gas, redissolved reconcentrated in 1 mL of methanol and subjected to above HPLC analysis.

#### *3.8. Preparation of Polymer/Antifoulant Coatings*

The polymer/antifoulant coatings were prepared using the solution casting method described by Ding et al. [23]. The coating was prepared by dissolving poly(ε-caprolactone) polyurethane (PCL-PU) [13] and butenolide or Boc-butenolide with different proportions (i.e., 95 wt% polymer and 5 wt% antifoulant for 5% antifoulant coating) in xylene, the mixture was then stirred vigorously until all solids dissolved, thereby forming a uniform solution. The coating solution was applied onto the surface of the panels, which were either epoxy panel (25 mm × 75 mm) for release rate determination or PVC panels (53 mm × 125 mm) for the field test. The panels were then placed under room temperature for 7 days until all solvents evaporated, and a continuous coating was formed.

#### *3.9. Determination of Antifoulant Release Rate from the Coatings*

The release rate of butenolide was determined by HPLC for quantification. The polymer/antifoulant coatings were prepared on an epoxy panel (25 mm × 75 mm) according to the above procedure. The coated panels were then placed into ASW. At certain time points (days 1, 8, 15, 22, 29, 50, 71 and 92 after immersion onto ASW), the panels were transferred into separate containers, which filled with 100 mL of fresh ASW. After 24 h immersion, the analyte in ASW was extracted with equal portion of dichloromethane (DCM). The DCM fraction with the analyte was dried under nitrogen gas, reconcentrated in 200 μL of methanol and subjected to HPLC analysis (Waters 2695, Taunton, MA, USA) using a reversed-phase system with a C18 column (Phenomenex Luna C18(2), 250 × 4.6 mm, 5 microns, Torrance, CA, USA) and a photodiode array detector (Waters 2998, Taunton, MA, USA) operated at 210 nm [13,21,22].

#### *3.10. Field Test*

The field tests were conducted in a fish farm in Yung Shue O, Hong Kong (114◦21 E, 22◦24 N) from January 2018 to March 2018. PVC panels (53 mm × 125 mm) covered with coatings were immersed in seawater at a depth of 1 m from the surface. The panels were retrieved once monthly, the dirt on the panels was removed by washing the panel gently with seawater before being photographed. The panels were placed back into the sea for monitoring. The antifouling potential of different panels were compared to determine the efficiency of the coatings. The estimation of the panel fouling coverage was achieved by ImageJ (National Institutes of Health, Bethesda, MD, USA) [24]. The percentage of area covered by foulers was calculated from the ratio of total fouling area to the panel area, in which the area was highlighted via the threshold function of ImageJ. IBM SPSS Statistics 22 was used for all statistical analyses. One-way ANOVA was used after initial analyses of heterogeneity and variance of the dataset with Levene's test followed by Tukey's post hoc test. Significance was defined as a *p*-value lower than 0.05.

#### **4. Conclusions**

The structure of butenolide was modified to Boc-butenolide to solve the problems of low melting point and smelliness of butenolide. *A. amphitrite* and *H. elagans* larval settlement bioassay results indicated that Boc-butenolide has similar antifouling ability against macrofoulers but with lower toxicity at high concentration. Boc-butenolide was released from the coatings and demonstrated antifouling ability for at least 2 months (as long as Boc-butenolide was released from coating). The release rate decreased with the increase in concentration of Boc-butenolide in the coatings. Our experiment demonstrated that Boc-butenolide exhibited good antifouling ability and could be a substitute compound for antifouling paints, and future efforts should focus on developing Boc-butenolide as a nontoxic antifouling compound with improved controlled release in the marine environment.

**Author Contributions:** Conceptualization and methodology, P.-Y.Q., H.Y.C., J.C. and C.M.; investigation, H.Y.C. and X.L.; formal analysis, H.Y.C. and J.C.; resources, P.-Y.Q. and C.M.; writing—original draft preparation, H.Y.C., J.C., X.L. and P.-Y.Q.; writing—review and editing, H.Y.C., J.C., X.L., C.M. and P.-Y.Q.; supervision and funding acquisition, P.-Y.Q. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research work was financially supported by the Hong Kong Branch of Southern Marine Science and Engineering Guangdong Laboratory (Guangzhou) (SMSEGL20SC01).

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The data presented in this study are available in the main text.

**Acknowledgments:** Special thanks to Lisa SOO at the Department of Ocean Science at The Hong Kong University of Science and Technology for her technical support in this project.

**Conflicts of Interest:** The authors declare the following competing interest: This work has been submitted for the U.S. provisional patent application (No. P1964US00).

#### **References**


## *Article* **Discovery of Antibiofilm Activity of Elasnin against Marine Biofilms and Its Application in the Marine Antifouling Coatings**

**Lexin Long 1,2, Ruojun Wang 2, Ho Yin Chiang 2, Wei Ding 3, Yong-Xin Li 4,\*, Feng Chen 5,\* and Pei-Yuan Qian 2,\***


**Abstract:** Biofilms are surface-attached multicellular communities that play critical roles in inducing biofouling and biocorrosion in the marine environment. Given the serious economic losses and problems caused by biofouling and biocorrosion, effective biofilm control strategies are highly sought after. In a screening program of antibiofilm compounds against marine biofilms, we discovered the potent biofilm inhibitory activity of elasnin. Elasnin effectively inhibited the biofilm formation of seven strains of bacteria isolated from marine biofilms. With high productivity, elasnin-based coatings were prepared in an easy and cost-effective way, which exhibited great performance in inhibiting the formation of multi-species biofilms and the attachment of large biofouling organisms in the marine environment. The 16S amplicon analysis and anti-larvae assay revealed that elasnin could prevent biofouling by the indirect impact of changed microbial composition of biofilms and direct inhibitory effect on larval settlement with low toxic effects. These findings indicated the potential application of elasnin in biofilm and biofouling control in the marine environment.

**Keywords:** elasnin; biofilms; marine; biofouling; natural products

#### **1. Introduction**

A biofilm is a microbial community attached to a surface [1]. It consists of microbial cells massed in the matrix of extracellular polymeric substances, which contain a large variety of biopolymers such as proteins, nucleic acids, lipids, and other substances [2]. Biofilms can be made up of a single microbial species or multiple species that colonize biotic or abiotic surfaces [3,4]. The elaborate biofilm architecture protects the microbes in biofilms and provides spatial proximity and internal homeostasis needed for growth and differentiation [3–5]. This composition makes microbial cells more resistant than their planktonic counterparts to diverse external insults such as antimicrobial treatment, poisons, protozoans, and host immunity [6,7]. For example, biofilms can render organisms 10- to 1000-fold less susceptible to antimicrobial agents; furthermore, organisms in multi-species biofilms are less susceptible to antimicrobial treatment than those in mono-species biofilms because of their complex interactions [6,8,9].

In the marine environment, biotic and abiotic surfaces are rapidly colonized by microorganisms and subsequent biofilm formation composed of bacteria, diatoms, fungi, unicellular algae, and protozoa [5,10], which creates a big problem for humans. Marine biofilms are critical in inducing biofouling and biocorrosion in the marine environment.

**Citation:** Long, L.; Wang, R.; Chiang, H.Y.; Ding, W.; Li, Y.-X.; Chen, F.; Qian, P.-Y. Discovery of Antibiofilm Activity of Elasnin against Marine Biofilms and Its Application in the Marine Antifouling Coatings. *Mar. Drugs* **2021**, *19*, 19. https://doi.org/ 10.3390/md19010019

Received: 27 November 2020 Accepted: 30 December 2020 Published: 5 January 2021

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Biofilms are important to habitat selection and settlement of many sessile marine organisms; for example, invertebrate larvae can distinguish biofilms composed of different microbial community structures to settle on or not [11–13]. The metabolites produced by microorganisms such as hydrogen sulfide, various acids, and ammonia destroy various materials. Every year, biofouling and biocorrosion contribute to enormous economic losses worldwide in industries, including heat exchange, oil and gas processing, storage and transportation, and drinking and wastewater industries [14–16]. Moreover, the persistence and transmission of harmful or pathogenic microorganisms and their genetic determinants within the marine biofilms pose a great threat to human beings [5].

Given the continuous increase in economic loss and the potential threats caused by the formation of marine biofilms, effective and economic control methods are necessary. At present, antibiofilm/antifouling coatings are the widely used and are an easy control method in the marine environment. However, the chemical inhibitors used in traditional coatings, such as amines, amides, organic tin, and cuprous oxide, are toxic and harmful, have no favorable environmental profile, and can be bioaccumulated. Increasing attention is focused on the use of natural products to develop effective and less hazardous coatings. Natural compounds with antibiofilm activities generated from the metabolic mechanism of microorganisms can be an ideal substitute for the traditional chemical biocides, presenting environmentally friendly properties such as low toxicity and biodegradability. However, insufficient productivity and difficulty in synthesis limit the development of naturally synthesized compounds [14,16,17].

Among the marine biofilms, as the early colonizers, bacteria are an important factor in determining the structure and function of a mature biofilm [5,14,18]. Therefore, bacteria from marine biofilms can be great targets for the discovery of antibiofilm compounds. In the present study, we used bacteria isolated from marine biofilms to screen for antibiofilm compounds, which led to the discovery of the potent antibiofilm activity of elasnin. With high productivity, elasnin-based coatings were consequently prepared, and their activities against natural multispecies marine biofilms were assessed in the field test.

#### **2. Results**

#### *2.1. Isolation and Identification of Biofilm Inhibition Compounds*

During our screening project, secondary metabolites produced by *Streptomyces mobaraensis* DSM 40847 (Figure 1b) exhibited strong biofilm inhibition activity against marine bacteria *Staphylococcus aureus* B04. Fractionation coupled with biofilm inhibition assay led to the identification of the main bioactive compound fraction 16 (Figure S1), which was produced at a high yield (approximately 332 mg/L), thereby achieving the maximum productivity after 2 days of incubation. Fraction 16 was subsequently purified using highperformance liquid chromatography (HPLC) and structurally characterized as a known compound elasnin using ultra-performance liquid chromatography-tandem mass spectrometry (UPLC–MS/MS) and nuclear magnetic resonance (NMR) spectroscopy (Figure 1, Supplementary Materials Figures S2 and S6 and Table S2).

Fraction 16: Appearance: colorless, viscous oil; UV (λmax): 291 nm; 1H NMR (500 MHz, DMSO-*d*6):δ0.80 (t, 3H), δ0.83 (t, 3H), δ0.86 (t, 3H), δ0.90 (t, 3H), 1.04~1.45 (overlapped, 18H), 1.61 (m, H, Ha-13), 1.86 (m, H, Hb-13), 2.32~2.47 (m, 6H), 3.87 (dd, 1H, *J* = 8.8, 5.7 Hz, H-6), 10.49 (s, 1H, 3-OH); 13C NMR (150 MHz, DMSO-*d*6): δ13.8, 13.8, 13.8, 13.8, 22.0, 22.1, 22.1, 22.1, 22.8, 22.8, 23.9, 27.6 (C-13), 28.9, 30.0, 30.5, 31.5, 39.8 (C-8), 52.8 (C-6), 103.1 (C-2), 114.2 (C-4), 154.5 (C-5), 163.6 (C-3), 163.8 (C-1), 206.8 (C-7); LC-ESI-MS: (*m/z*) [M + H]+ 393.3.

**Figure 1.** Elasnin was produced by *Streptomyces mobaraensis* DSM 40847 with high yield. (**a**) Mass spectra (ESI) and structure of elasnin; (**b**) growth of *S. mobaraensis* DSM 40847 on the GYM (Table S1) agar plate; (**c**) high-performance liquid chromatography (HPLC) analysis of the crude extracts of *S. mobaraensis* DSM 40847; (**d**) time course of the production of elasnin in AM4 medium at 30 ◦C.

#### *2.2. Elasnin Could Inhibit the Biofilm Formation of Multiple Strains of Bacteria Isolated from Marine Biofilms*

Ten strains isolated from marine biofilms—*Vibrio alginolyticus* B1, *Erythrobacter* sp. HKB8, *Ruegeria* B32, *S. aureus* B04, *S. hominis* N32, *S. arlettae* OM, *Microbacterium esteraromaticum* N22, *Idiomarina sediminum* N28, *Pseudoalteromonas* L001, and *Escherichia coli* N57—were used as targets in the minimum biofilm inhibitory concentration (MBIC) assay and minimal inhibitory concentration (MIC) assay. The MBIC is the lowest concentration of a compound that resulted in a certain reduction in the attached cells while the MIC is the lowest concentration needed for inhibiting the visible growth of planktonic cells. Among the ten strains, seven successfully formed biofilms during the test, and three (*V. alginolyticus* B1, *Erythrobacter* sp. HKB8, and *Ruegeria* B32) cannot form biofilms under the testing situation. For the seven biofilm-forming strains, the biofilms of four Gram-positive strains were sensitive to elasnin treatment with MBIC90 of 2.5 to 5 μg/mL and MBIC50 of 1.25 to 5 μg/mL, whereas the MBIC90 and MBIC50 of elasnin against Gram-negative strains ranged from 5 to 10 μg/mL and 1.25 to 10 μg/mL, respectively (Figure 2a). The MICs of nine strains were determined except for *Microbacterium esteraromaticum* N22 due to its inability to grow under the testing conditions. Elasnin inhibited the planktonic cells of *S. aureus* B04 and *Idiomarina sediminum* N28 from growing with MICs of 5 to 10 μg/mL while for other strains the MICs of elasnin were above 10 μg/mL (Figure 2b). Overall, comparing to antimicrobial activity, elasnin shows more significant efficiency in inhibiting biofilm formation.

**Figure 2.** Antibiofilm activities of elasnin against bacteria isolated from marine biofilms. (**a**) Minimum concentration needed for inhibiting biofilm formation; (**b**) summary of minimum biofilm inhibitory concentration (MBICs) and minimal inhibitory concentration (MICs) of elasnin against test strains.

#### *2.3. Preparation of Elasnin-Based Antibiofilm Coatings*

Given the high antibiofilm efficiency and high productivity, elasnin-based antibiofilm coatings were prepared and immersed in a fish farm to evaluate their efficiency against natural marine biofilms. In the present study, we used a crude extract of *S. mobaraensis* DSM40847 that contained high concentrations of elasnin (=336.64 mg/L in n-hexane, Figure S7) instead of pure elasnin. The extracts were mixed with polyurethane (polymer) on the basis of poly ε-caprolactone and applied directly on the surface of glass slides. The concentrations of the coatings were calculated on the basis of the percentage of crude extracts in total coatings (polymer and crude extracts) by weight. As such, other compounds in low amounts in the fractionated extract might affect the results of our field testing. However, their effect should be negligible because we did not detect any significant effects of the minor compounds on the crude extracts (Figure S7) and the crude extracts has the same biofilm inhibition efficiency as elasnin shows (MBIC of 0.8–4 and 4–20 μg/mL to *S. aureus* B04 and *E. coli* N57, respectively).

#### *2.4. Release Rate of Elasnin from Antibiofilm Coatings*

The release rate of elasnin from the coatings was dependent on time and concentration during the 4 weeks observation (Figure 3c). In general, the release of elasnin was maintained at a low rate throughout the period; the higher the concentration, the faster the release of elasnin into the artificial seawater. The highest release rate of approximately 5 μg day−<sup>1</sup> cm−<sup>2</sup> occurred in the second week for the concentration of 10 wt%; for other concentrations, the maximum release rate was approximately 4 μg day−<sup>1</sup> cm−<sup>2</sup> in the first week. The release rate decreased over time and depended on the total amount of elasnin remaining in the coatings. After immersion for 4 weeks, the release rate dropped to approximately 1 μg day−<sup>1</sup> cm−<sup>2</sup> for the concentrations of 10 wt% and 5 wt% and 0.5 μg day−<sup>1</sup> cm−<sup>2</sup> for 1.5 wt% and 2.5 wt%.

**Figure 3.** Antibiofilm (weeks 2 and 3) and antifouling (week 4) performance of elasnin-based antibiofilm coatings. (**a**) Confocal laser scanning microscopy (CLSM) images (weeks 2 and 3) of the coated surface (week 4); (**b**) Biomass of biofilms observed by CLSM. (**c**) Monitoring of the release rate of elasnin into the artificial seawater; Biomass is calculated using Comstat 2.1 on the basis of the CLSM images and values that are significantly different among elasnin-based antibiofilm coatings, and the control groups are indicated by asterisks: \* for *p* < 0.05 and \*\* for *p* < 0.01.

#### *2.5. Elasnin-Based Coatings Inhibited the Formation of Multi-Species Biofilms and the Attachment of Large Biofouling Organisms in the Marine Environment*

The performance of the antibiofilm coatings was assayed every week from the second to the fourth week by direct and confocal laser scanning microscopy (CLSM) observation (Figure 3a). Based on the quantitative analysis of CLSM images, the average biofilm biomass on the slides without elasnin was 116.44 μm3 μm−<sup>2</sup> in the second week and 259.95 μm3 μm−<sup>2</sup> in the third week, whereas the average biomass of biofilms measured on

5 wt% and 10 wt% coating slides was less than 0.1 μm3 μm−<sup>2</sup> in the second week and less than 120 μm3 μm−<sup>2</sup> in the third week. For coatings with low concentrations (1.5 wt% and 2.5 wt%), no significant differences were observed with regard to average biomass (61.97 and 84.73 μm3 μm−2, respectively) in the second week, but the biomass was significantly lower than that in the control (259.95 μm3 μm−2) in the third week, with an average biomass of approximately 125 and 145 μm3 μm−2, respectively (Figure 3b). In the fourth week, slides coated with low concentrations of elasnin (1.5 wt%, 2.5 wt%, and control) were fouled by large marine organisms, whereas those coated with high concentrations of elasnin exhibited an anti-macrofouling activity and almost no larval settlement, except for a small area near the edges because of the edge effects commonly found on testing panels. Elasnin-based antibiofilm coatings inhibited the biofilm formation of multiple bacterial species in the first 2 weeks. However, after immersion for 4 weeks, the glass slides coated with low concentrations of elasnin were eventually covered with large biofouling organisms probably because of the low releasing rate of the elasnin after 3 weeks.

#### *2.6. Elasnin Changed the Microbial Community Structure of Natural Marine Biofilms*

Considering that the number of biofilms developed by the end of the second week was limited, and macrofoulers had overgrown by the end of the fourth week, only the 3 week-old biofilms developed on 10 wt% coatings and those on the control glass slides (coated with poly ε-caprolactone-based polyurethane only) were selected for 16S amplicon analysis to determine the changes in biofilm microbial community triggered by elasnin. A total of 3,000,000 16S rRNA gene sequences (500,000 per sample) were classified into 31 phyla (*Proteobacteria* were classified down to the class level). The microbial composition of the biofilms differed between the 10 wt% coatings and the control slides, as confirmed by alpha- and beta-diversity analysis. In the Bray–Curtis dissimilarity (beta-diversity) dendrogram (Figure 4a), the control and treatment groups were clustered separately on the basis of the differences in microbial abundance among the samples; the observed operational taxonomic units (OTUs) and Shannon diversity for the treated biofilm were significantly lower than those in the control group (Figure 4b), indicating that species richness and diversity in the treated biofilms were reduced.

**Figure 4.** Composition analysis of biofilms grown on the control slides (without crude extracts) and treatment slides (with 10% CR coatings). (**a**) Similarity comparison of microbial compositions among biofilms on control slides (C-1,2,3) and 10 wt% elasnin-based coatings (E10-1,2,3) based on the beta-diversity (Bray–Curtis) at the phylum level. (**b**) Alpha-diversity of biofilms at the phylum level. The difference between the two types of biofilms is calculated using Student's t-test and is indicated by an asterisk: \* for *p* < 0.05.

#### *2.7. Elasnin Inhibited the Larval Settlement of Balanus Amphitrite with a Low Toxic Effect*

The antilarval settlement activity of elasnin was measured using *B. amphitrite*, and its possible toxic effects were preliminarily assessed by the mortality rate. When the larvae were exposed to elasnin for 24 h, the larval settlement was significantly inhibited

at concentrations above 12.5 μg/mL. No increase in mortality rate was observed at a concentration range from 6.25 to 50 μg/mL compared with the control groups (Figure 5a). After 48 h exposure to elasnin, the settlement inhibition was slightly reduced for the concentration of 25 μg/mL, whereas for the concentration of 50 μg/mL, the inhibitory effect was significant. An increased mortality rate of around 65% and reduced vitality of larva were observed at concentrations of 50 μg/mL after 48 h exposure, but no significant changes were exhibited in the mortality rate for other concentrations of the group compared with the control (Figure 5b).

**Figure 5.** Effect of elasnin on the percentage of larval settlement and mortality rate of *B. amphitrite* after treatment for (**a**) 24 h and (**b**) 48 h. The differences between the control and treatment groups are calculated using Student's t-test and are indicated by asterisks: \* for *p* < 0.05, \*\* for *p* < 0.01, and \*\*\* for *p* < 0.001.

#### **3. Discussion**

Here, we used bacteria isolated from marine biofilms to screen for the antibiofilm compounds that targeted the marine biofilms, which led to the discovery of the antibiofilm activity of elasnin. Elasnin-based coatings were then prepared with simple and costeffective methods, and its efficiency against multi-species biofilms was tested in the natural marine environment.

The predictive validity of the screening assay is an important determinant of the success of drug discovery [19,20]. Marine biofilms are complex mixed-species microbial communities with specific intraspecies and interspecies communication and interaction [21,22]. Therefore, the simulation of the marine biofilm formation is difficult under laboratory conditions. In accelerating and simplifying the screening for antibiofilm compounds against marine biofilms with high predictive validity, bacteria isolated from marine biofilms were selected as the target for the bioassay. Consequently, the potential inhibitory activity of elasnin against marine biofilms was discovered during screening. Seven out of ten strains of bacteria isolated from marine biofilms had successfully formed biofilms under the testing situation, and elasnin showed great inhibiting efficiency against all biofilm-forming strains. The inhibiting efficiency of elasnin-based coatings against natural marine biofilms was then validated in the field test, which indicated a great predictive validity of our screening assays.

Elasnin was first discovered in 1978 as a new elastase inhibitor with low toxicity in mice and high selectivity for human granulocyte elastase [23], and its antimicrobial and antibiofilm activities have never been discovered. Based on the test results in the present study, elasnin not only inhibited the biofilm formation of both mono- and multispecies biofilms but also inhibited the settlement of large biofouling organisms. The 16S amplicon analysis revealed that the species richness and diversity of biofilms on the elasnin-based coatings were reduced, which might indirectly inhibit the settlement of biofouling organisms because the changed microbial compositions of the biofilms might not be suitable for them to settle on. In addition, the antilarval-settlement assay showed that elasnin could inhibit larval settlement, which had low toxicity. Considering the effective concentration of elasnin and the water mobility in the marine environment, the toxic effect of elasnin should be negligible, although a high mortality rate was recorded under the high concentration after 48 h exposure. However, the present study only superficially assessed the potential toxic effect of elasnin by the mortality rate of the larva, and its environmental impact should be further explored.

In addition, the wide-type strain *S. mobaraensis* DSM 40847 produced elasnin in substantial quantity (0.33 g/L), which was considered as a new industrial-producing strain of antibiofilm agents. Low product yield has always been a limitation for the development and commercial applications of newly discovered biologically active compounds. The fermentation of this bacterium would address the supply problem that often limited natural products from biotechnical development, leading to the late-stage development of elasnin. Subsequently, elasnin-based coatings were easily prepared with low expenditures. During a 4 week observation, the coatings (10 wt%) inhibited the biofilm formation in the first 2 weeks and the larval settlement in the last 2 weeks. Coatings began to lose their effectiveness after the third week in the field probably because of the reduced release rate of elasnin. Since 2008, when tributyltin (TBT) was restricted by the implementation of the International Maritime Organization Treaty on biocides, the development of efficient and environmentally friendly surface coatings became a hot topic. Collectively, originating from nature, elasnin showed great antibiofilm and antifouling activities with low toxic effects; combined with the low cost of supply, elasnin could provide a new selection for the development of antibiofilm and antifouling materials.

In the present study, we identified a potent antibiofilm compound, namely, elasnin, from the strain *S. mobaraensis* DSM 40847 in the course of our screening program using bacteria isolated from marine biofilms. With high productivity, elasnin-based antibiofilm coatings were easily prepared, which presented a favorable performance in inhibiting the biofilm formation and attachment of macro-foulers in the marine environment. The 16S amplicon and antilarval settlement assays revealed that the antifouling performance of elasnin-based coatings might be caused by the indirect effect of elasnin on biofilm's microbial compositions and its direct inhibitory effect on the larval settlement. With low toxicity, high efficiency, and high productivity, elasnin showed great potential in the applications of biofilm and biofouling control in the marine environment.

#### **4. Materials and Methods**

#### *4.1. Strains, Culture Media, and Chemicals*

*Streptomyces mobaraensis* DSM 40847 was purchased from the German Collection of Microorganisms and Cell Cultures (DSMZ, Braunschweig, Germany). The marine bacteria *V. alginolyticus* B1, *Erythrobacter* sp. HKB8, *Ruegeria* B32, *S. aureus* B04, *S. hominis* N32, *S. arlettae* OM, *M. esteraromaticum* N22, *I. sediminum* N28, *Pseudoalteromonas* L001, and *E. coli* N57 were isolated from marine biofilms and obtained from the culture collection of our laboratory [24]. Soybean powder was purchased from Wugumf, Shenzhen, China. Soluble starch was purchased from Affymetrix, Santa Clara, CA, USA. Magnesium sulfate hydrate was purchased from Riedel-de-Haën, Seelze, Germany. Bacteriological peptone was obtained from Oxoid, Milan, Italy. Mueller-Hinton broth (MHB) was purchased from Fluka Chemie AG, Buchs, Switzerland. Phosphate-buffered saline (PBS) and 3- (4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) were purchased from Thermo Fisher Scientific Inc., San Jose, CA, USA. Lysogeny broth (LB), glucose, and 1-butanol were purchased from VWR International Ltd., Leicestershire, UK. All other chemicals were supplied by Sigma-Aldrich Corporation, Saint Louis, MO, USA.

#### *4.2. Bioactive Compound Isolation and Identification*

Stock cultures of *S. mobaraensis* DSM 40847 were inoculated into 50 mL of AM4, AM5, and AM6 media (Table S1) containing glass beads (to break up globular colonies) and incubated at 30 ◦C on a rotary shaker (170 rpm). The culture broth was extracted with 1-butanol on days 3, 5, and 7. The crude extracts were dissolved in DMSO before storage and bioassay. Pure compounds were isolated by reversed-phase HPLC (Waters 2695, Milford, MA, USA) using a semi-prep C18 column (10 × 250 mm) that was eluted with a 55 min gradient of 5–95% aqueous acetonitrile containing 0.05% trifluoroacetic acid at a flow rate of 3 mL/min. The structure of elasnin was elucidated through NMR analysis of 1H, 1H-1H-COSY, 1H-13C-HSQC, and 1H-13C-HMBC NMR spectra recorded on a Bruker AV500 NMR spectrometer (Bruker, Billerica, MA, USA) and 13C-NMR spectra obtained with the Bruker DRX600 NMR Spectrometer (Bruker, Billerica, MA, USA) using dimethyl sulfoxide-d6 ( 1H-NMR DMSO-d6: δH = 2.50 ppm; DMSO-d6: δC = 39.50 ppm).

#### *4.3. Productivity Monitoring and Extraction Efficiency Comparison*

A stock culture of *S. mobaraensis* DSM 40847 was incubated in the AM4 medium. One milliliter of culture broth was collected every 12 h and extracted with 1 mL of 1-butanol, ethyl acetate, or n-hexane. The solvent for extraction was then removed by evaporation. The crude extract was dissolved in methanol and quantified through HPLC analysis with a Phenomenex Luna C18 column. The peak of elasnin was identified from the retention time, and its concentration was calculated on the basis of an established standard curve (Figure S9).

#### *4.4. MBIC Assay and MIC Assay Against Marine Bacteria*

MBICs were determined as previously described [25,26]. In brief, an overnight culture of test strains was diluted into approximately 107 CFU/mL with LB and 0.5% glucose and treated with various concentrations of elasnin (or only media for control) in 96-well cell culture plates. Then, the plates were incubated at 37 ◦C for 24 h and rinsed two times with 1 × PBS to remove non-adhering and planktonic cells. After rinsing, an MTT staining assay was conducted to measure viable cells in the biofilms because MTT could react with activated succinate dehydrogenase in the mitochondria of viable cells to form blue-violet formazan, which could be read at 570 nm after dissolving in DMSO. The MBIC50 and MBIC90 were defined as the lowest concentration needed for inhibiting 50% and 90% of biofilm formation individually. The biofilm inhibition efficiency was calculated using the following equation: Biofilm inhibition (%) = (OD570nm of test compound) / (OD570nm of control) × 100%. The experiments were performed in triplicate and repeated three times.

MICs were determined with test strains according to the Clinical and Laboratory Standards Institute guideline CLSI M100 (2018). Briefly, a 10<sup>5</sup> CFU/mL overnight culture of test strains was inoculated into MHB and treated with elasnin (or only media for control) at a series of concentrations. After incubation for 24 h, the minimum concentrations at which no bacterial growth was visible were recorded as the MICs. The experiments were performed in triplicate and repeated twice; vancomycin and kanamycin were used as a positive control in the experiments.

#### *4.5. Elasnin-Based Antibiofilm Coating Preparation*

A 4 L culture broth of *S. mobaraensis* DSM 40847 (incubated as previously described) was extracted using n-hexane to obtain a sufficient amount of high-elasnin-content crude extracts. The elasnin-based antibiofilm coatings were prepared following the same procedures as those described by Ma et al. (2017). For the 10 wt% coatings, polymer (0.90 g, 90 wt%) and crude extracts (0.10 g, 10 wt%) were dissolved by vigorously stirring xylene and tetrahydrofuran (v:v = 1:2) at 25 ◦C. After mixing, a glass slide was coated with the solution and left to dry at room temperature for a week to remove the solvent. The same procedure was used in preparing coatings with different concentrations of crude extracts.

#### *4.6. Field Test and Release Rate Determination*

Coated glass slides were submerged in seawater at a fish farm in Yung Shue O, Hong Kong (114◦21 E, 22◦24 N) for 2 to 4 weeks. Afterward, the glass slides were retrieved and transported back to the laboratory in a cooler with in situ seawater and were washed two times using an autoclave and 0.22 μm filtered seawater (FSW) to remove loosely attached particles and cells. The slides were then stained using the FilmTracer™ LIVE⁄DEAD Biofilm Viability kit and investigated under CLSM (Zeiss LSM710, Carl Zeiss, Oberkochen, Germany). Moreover, the release rate of elasnin was determined by measuring its concentration using HPLC under static conditions. The coated panels were immersed in 100 mL of sterilized artificial seawater held in a measuring container. Ten milliliters of seawater was collected after immersion for 24 h, and elasnin was extracted with the same volume of dichloromethane, which was then removed under nitrogen gas. After drying, the extract was resuspended in 100 mL of methanol and underwent HPLC analysis. The release rate was measured every week for 4 straight weeks, and each concentration was tested in duplicate.

#### *4.7. DNA Extraction, 16S rRNA Gene Sequencing, and Analyses*

Biofilm samples on the coated slide surface were collected with autoclaved cotton and stored in DNA storage buffer (10 mM Tris-HCl; 0.5 mM EDTA, pH 8.0) at −80 ◦C. Before the extraction, samples were vortexed several times to release the microbial cells into the DNA storage buffer. All the samples were then subjected to centrifugation at 10,000 rpm for 1 min, and the supernatant was discarded. After continuous treatment with 10 mg/mL of lysozyme and 20 mg/mL of proteinase K, DNA was extracted from the treated microbial cells with a microbial genomic DNA extraction kit (Tiangen Biotech, Beijing, China) following the manufacturer's protocol.

The quality of DNA samples was controlled using NanoDrop (which tested DNA purity, OD260/OD280) and agarose gel electrophoresis (which tested DNA degradation and potential contamination). The hypervariable V3-V4 region (forward primer: 5 -CCTAYGGGRBGCASCAG-3 ; reverse primer: 5 -GGACTACNNGGGTATCTAAT-3 ) of prokaryotic 16S rRNA genes was used to amplify DNA from biofilms by polymerase chain reaction (PCR). The PCR products were purified before library construction and sequenced at Novogene (Beijing, China) on the NovaSeq 6000 System. The read length was 250 bp, and each pair of reads had a 50 bp overlapping region. The paired-end reads were subjected to quality control using the NGS QC Toolkit (version 2.0, The National Institute of Plant Genome Research, New Delhi, India) [27]. The 16S rRNA gene amplicon data were analyzed using the software package QIIME2 and then merged using Q2\_manifest\_maker.py in QIIME2 [28]. The low-quality reads and chimeras were removed using dada2 commands in QIIME2. A total of 500,000 filtered reads for each sample were selected to normalize the uneven sequencing depth. OTUs were classified de novo from the pooled reads at 97% sequence similarity using a classifier trained by the Naive Bayes method. Representative sequences were then recovered using the feature-classifier classify-sklearn script in QIIME2. The alpha-diversity analyses (observed OTUs and Shannon diversity) were performed using the script "qiime diversity alpha" in QIIME2. Beta-diversity based on the Bray–Curtis distances was conducted by the cluster analysis in the software PAST (version 3.0) [29]. Furthermore, the taxonomic structure was drawn in Excel wo (Office 365 MSO 64-bit) on the basis of the relative abundance.

#### *4.8. Antilarval-Settlement Assay*

The direct antilarval-settlement assay was conducted using cyprids of the barnacle *B. amphitrite* Darwin as described previously [30–33]. In brief, adult *B. amphitrite* (Darwin) were collected from the intertidal zone in Pak Sha Wan, Hong Kong (22◦19 N, 114◦16 E) and raised to competence for experiments. Elasnin was dissolved in DMSO and diluted into four concentrations from 50 to 6.25 μg/mL with a twofold serial dilution. DMSO was used as a negative control. About 10 ± 2 *B. amphitrite* cyprids were inoculated into

each well of a 24-well polystyrene culture plate that contained 2 mL of 0.22 μm FSW with different treatments. For all treatments and controls, three replicates were performed. The plates were then incubated at 25 ◦C in darkness. After 24 and 48 h, the number of settled and swimming larvae was counted using a Leica MZ6 microscope (Leica Microsystems, Wetzlar, Germany), and possible toxic effects were also noted.

#### *4.9. Statistical Analyses*

Statistical analyses for all data were performed using the GraphPad Prism 8.0.2 software (San Diego, CA, USA). The composition of the biofilm on the coatings was compared with that in the control groups using Student's t-test.

#### **5. Patents**

The authors declare the following competing interests: This work has been submitted for the U.S. Patent Application (No. 16999437) and Chinese Patent application (No. 202010850564.X).

**Supplementary Materials:** The following are available online at https://www.mdpi.com/1660-3 397/19/1/19/s1, Table S1: Media used in this study, Figure S1: Bioactivities of crude extract of the secondary metabolites produced by *S. mobaraensis* DSM 40847 (incubated with AM4 media and extracted with 1-butanol) and 20 fractions of it. Fraction 17 and 15 are the analogs of franction 16 (elasnin), Figure S2: 13C-NMR analysis of bioactive fraction 16 (Elasnin), Figure S3: 1H-NMR analysis of bioactive fraction 16 (Elasnin), Figure S4: 1H-1H COSY of bioactive fraction 16 (Elasnin), Figure S5: 1H-13C HSQC of bioactive fraction 16 (Elasnin), Figure S6: 1H-13C HMBC of bioactive fraction 16 (Elasnin), Table S2 13C-NMR (150 MHz, DMSO-d6), 1H-NMR (500 MHz, DMSO-d6) and HMBC correlations of compound Fraction 16 (DMSO-d6) and comparisons between Elasnin (Omura, Nakagawa et al. 1979), Figure S7: HPLC profile of high-elasnin-content crude extracts and productivity of crude extracts/elasnin by using different extraction solvent, Figure S8: Microbial compositions of biofilms on control slides (C-1,2,3) and 10 wt% elasnin-based (E10-1,2,3) coatings at the genus level, Figure S9: Stand curve of elasnin acquired by HPLC.

**Author Contributions:** L.L. contributed to the experiment design, performed all the experiments, interpreted the data, and prepared the manuscript. R.W. performed the experiments of DNA extraction, 16S amplicon analysis, wrote related methods in the manuscript, and commented on the manuscript. H.Y.C. performed the antilarval-settlement assay, assisted in the coating preparation and field test, and commented on the manuscript. R.W. and W.D. isolated the bacteria from marine biofilms. Y.-X.L. designed and supervised this study, discussed the results and implications at all stages, and edited the manuscript. F.C. and P.-Y.Q. supervised this study, gave technical support and conceptual advice, and edited the manuscript. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was financially supported by the National Key R&D Program of China (2018YFA0903200), China Ocean Mineral Resources Research and Development Association (COM-RRDA17SC01), the Hong Kong Branch of Southern Marine Science and Engineering Guangdong Laboratory (Guangzhou) (SMSEGL20SC01 and GML2019ZD0409) and a CRF grant from HKSAR government (C6026-19G-A).

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The data presented in this study are available in the main text and the supplementary materials of this article.

**Acknowledgments:** Special thanks to Weipeng Zhang of the College of Marine Life Science at Ocean University for helping to isolate the bacteria from marine biofilms and Rui Feng of the Division of Life Science at the Hong Kong University of Science and Technology for assisting in doing NMR analysis.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


## *Article* **Flavonoid Glycosides with a Triazole Moiety for Marine Antifouling Applications: Synthesis and Biological Activity Evaluation**

**Daniela Pereira 1,2,†, Catarina Gonçalves 2,†, Beatriz T. Martins 1, Andreia Palmeira 1,2, Vitor Vasconcelos 2,3, Madalena Pinto 1,2, Joana R. Almeida 2,\*, Marta Correia-da-Silva 1,2,\* and Honorina Cidade 1,2**


**Abstract:** Over the last decades, antifouling coatings containing biocidal compounds as active ingredients were used to prevent biofouling, and eco-friendly alternatives are needed. Previous research from our group showed that polymethoxylated chalcones and glycosylated flavones obtained by synthesis displayed antifouling activity with low toxicity. In this work, ten new polymethoxylated flavones and chalcones were synthesized for the first time, including eight with a triazole moiety. Eight known flavones and chalcones were also synthesized and tested in order to construct a quantitative structure-activity relationship (QSAR) model for these compounds. Three different antifouling profiles were found: three compounds (**1b**, **11a** and **11b**) exhibited anti-settlement activity against a macrofouling species (*Mytilus galloprovincialis*), two compounds (**6a** and **6b**) exhibited inhibitory activity against the biofilm-forming marine bacteria *Roseobacter litoralis* and one compound (**7b**) exhibited activity against both mussel larvae and microalgae *Navicula* sp. Hydrogen bonding acceptor ability of the molecule was the most significant descriptor contributing positively to the mussel larvae anti-settlement activity and, in fact, the triazolyl glycosylated chalcone 7b was the most potent compound against this species. The most promising compounds were not toxic to *Artemia salina*, highlighting the importance of pursuing the development of new synthetic antifouling agents as an ecofriendly and sustainable alternative for the marine industry.

**Keywords:** flavonoids; synthesis; click chemistry; biofouling; antifouling; eco-friendly alternatives

#### **1. Introduction**

Marine biofouling, resulting from the accumulation of marine micro and macroorganisms on submerged surfaces, has been a huge problem for maritime industries, causing several technical and economic problems, including corrosion of materials and the increase in fuel consumption. Moreover, marine biofouling is associated with environmental and health problems, due to an increase in gas emissions and the spread of invasive species [1,2].

Biocidal paints containing organotin compounds, namely tributyltin (TBT), were widely used for decades in the maritime industry to prevent biofouling. However, due to their negative effect on the environment and on live organisms, these substances were completely banned in 2008 by the international maritime organization [3]. Since then, some booster biocides, such as Irgarol 1051 or Sea-nine 211, in combination with copper, have been used; nevertheless, even these compounds have demonstrated toxicity on living organisms.

**Citation:** Pereira, D.; Gonçalves, C.; Martins, B.T.; Palmeira, A.; Vasconcelos, V.; Pinto, M.; Almeida, J.R.; Correia-da-Silva, M.; Cidade, H. Flavonoid Glycosides with a Triazole Moiety for Marine Antifouling Applications: Synthesis and Biological Activity Evaluation. *Mar. Drugs* **2021**, *19*, 5. https:// dx.doi.org/10.3390/md19010005

Received: 5 December 2020 Accepted: 21 December 2020 Published: 24 December 2020

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https://creativecommons.org/ licenses/by/4.0/).

Therefore, it is imperative to find new antifouling (AF) compounds with environmentally safe characteristics [4–6]. Several non-toxic marine natural products with AF activity have been reported; among them, some flavonoids presented potential AF activity and low toxicity, suggesting their potential as new lead compounds for the development of new AF agents [7].

Previous works from our group reported some glycosylated flavones [8] and chalcones [9] with potential AF activity. Interestingly, when comparing the anti-settlement activity against *Mytilus galloprovincialis* of previously described chalcones, it seemed that the presence of a polymethoxylated B-ring could be important for this activity [9]. Moreover, the introduction of a triazole moiety is associated with an increase in AF activity [10]. In fact, over the last decade, there has been a great interest in the synthesis of 1,2,3-triazoles due to the fact of these moieties behaved as more than passive linkers. They carried favorable physicochemical properties, showing importance to biological activity [11,12]. This approach has been used to generate a vast array of compounds with biological potential [13–16], namely with AF activity [10,17,18]. Moreover, some antimicrobial agents are based on nitrogen heterocycles, including the triazole-based biocides fluconazole and itraconazole, which suggest their potential to act as AF agents [10].

Based on this, the present work aims to synthesize new potential AF polymethoxylated chalcone and flavone derivatives with glycosyl groups incorporating a 1,2,3-triazole moiety using a click chemistry approach. The potential of synthesized compounds as benign AF agents was assessed against the adhesive larvae of the macrofouling mussel *Mytilus galloprovincialis* and the biofilm-forming marine bacteria *Cobetia marina*, *Vibrio harveyi*, *Pseudoalteromonas atlantica*, *Halomonas aquamarina* and *Roseobacter litoralis.* The most promising compounds were submitted to complementary assays to evaluate their viability as AF agents, including the evaluation of possible mechanisms of action related with adhesion and neurotransmission pathways. These compounds were also tested for anti-microalgal activity towards *Navicula* sp. and general ecotoxicity using nauplii of the marine shrimp *Artemia salina*.

#### **2. Results and Discussion**

#### *2.1. Synthesis and Structure Elucidation*

A series of four glycosylated flavones and four glycosylated chalcones bearing a 1,2,3 triazole moiety was synthesized. To prepare glycosylated flavones (Scheme 1), flavones **1a** and **1b**, used as building blocks, were synthesized by the Mentzer synthesis, through direct thermal cyclocondensation of phloroglucinol and β-ketoesters, with good yields, as described by Seijas et al. [19]. However, instead of a microwave (MW) irradiation, the synthesis of flavones **1a** and **1b** was performed in a muffle furnace. After, the propargylation of flavones **1a** and **1b** was achieved with propargyl bromide, giving rise to flavones **2a** and **2b** with 66% and 55% yield, respectively. Copper(I)-catalysed azide alkyne cycloaddition (CuAAC), commonly referred as click chemistry, was developed by the Sharpless and Meldal groups in 2002, and is the most useful reaction for the regioselective synthesis of 1,4-disubstituted-1,2,3-triazole ring [20,21]. This involves a reaction of a terminal alkyne and an aliphatic azide using copper (I) as a catalyst in low-time and mild conditions, with high yields and few by-products [20–22]. Therefore, the incorporation of the triazole linked glycosidic moiety in flavones **2a** and **2b** was accomplished by CuAAC under MW irradiation, giving rise to flavones **3a**, **3b**, **4a** and **4b** with 49–82% yield (Scheme 1).

**Scheme 1.** Synthesis of flavones **1a**–**1b, 2a**–**2b**, **3a**–**3b** and **4a**–**4b**. (i) 240 ◦C, 60–80 min, 74–77%; (ii) Cs2CO3, tetrabutylammonium bromide (TBAB), acetone, 60 ◦C, 6 h, 55–66%; (iii) Sodium ascorbate, CuSO4.5H2O, tetrahydrofuran (THF):water, microwave (MW), 30 min, 49–82%; (iv) NaN3, acetone:water, r.t., 3 h, 72%.

The first step in the synthetic process to obtain glycosylated chalcones (Scheme 2) was the propargylation of 2,4-dihydroxyacetophenone with propargyl bromide. As for the synthesis of flavones **2a**–**2b**, firstly this reaction was accomplished with propargyl bromide, in the presence of anhydrous Cs2CO3 and tetrabutylammonium bromide (TBAB). Nevertheless, in addition to the desired 4-*O*-monosubstituted acetophenone (**5**), the 2,4-disubstituted acetophenone was obtained. Therefore, this reaction was performed in the presence of anhydrous K2CO3, as described by Zhao et al. [23], with slight modifications, and the 4-*O*-monosubstituted acetophenone (**5**) was successfully obtained as expected, with a 76% yield. Afterwards, the base-catalysed aldol reaction of this propargylated acetophenone with benzaldehydes afforded chalcones **6a** and **6b** with moderate yields, which were subsequently submitted to MW assisted CuAAC with azide sugar derivatives, affording triazole linked glycosylated chalcones **7a**, **7b**, **8a** and **8b** with 45–65% yield.

In order to perform structure–activity relationship studies, structure related nonglycosylated chalcones were also synthesized (Scheme 3). Firstly 2,4-dihydroxyacetophenone was protected with methoxymethyl chloride affording **9** with 84% yield. Chalcones **10a** and **10b** were prepared by base-catalyzed aldol reaction of **9** and 3,4-dimethoxy- and 3,4,5 trimethoxybenzaldehyde with 33% and 47% yield, respectively, as described before [24,25], with slight modifications. Chalcones **11a** and **11b** were obtained with moderate yields by deprotection of methoxymethyl group at C-4 of intermediate chalcones **10a** and **10b**, as described by Loureiro et al. [26].

The 2,3,4,6-tetra-*O*-acetyl-β-D-glucopyranosyl azide (**12**), used as a building block for the synthesis of glycosylated derivatives **3a**, **3b**, **7a** and **7b**, was synthesized from 2,3,4,6-tetra-*O*-acetyl-α-D-glucopyranosyl bromide and sodium azide, as described by Adesoye et al. [27], with 72% yield.

**Scheme 2.** Synthesis of chalcones **6a**–**6b**, **7a**–**7b** and **8a**–**8b**. (i) K2CO3, acetone, 60 ◦C, 1 h, 76%; (ii) 40% NaOH, methanol, microwave (MW), 3 h, 41–43%; (iii) Sodium ascorbate, CuSO4.5H2O, tetrahydrofuran (THF):water, microwave (MW), 1 h, 45–65%; (iv) NaN3, acetone:water, r.t., 3 h, 72%.

**Scheme 3.** Synthesis of chalcones **10a**–**10b** and **11a**–**11b**. (i) K2CO3, acetone, 60 ◦C, 1 h, 84%; (ii) 40% NaOH, methanol, microwave (MW), 4 h, 33–47%; (iii) p-Toluenesulfonic acid (PTSA), methanol, 50 ◦C, 5 h, 24–31%.

The newly synthesized compounds, **2a**, **2b**, **3a**, **3b**, **4a**, **4b**, **7a**, **7b**, **8a** and **8b** were characterized by high resolution mass spectrometry (HRMS) and nuclear magnetic resonance (NMR). The coupling constants of the vinylic system (JH<sup>α</sup>-H<sup>β</sup> = 15.5–15.3 Hz) confirm the (*E*)-configuration for all synthesized chalcones. The NMR spectra of the newly synthesized compounds **3a**, **3b**, **4a**, **4b**, **7a**, **7b**, **8a** and **8b** showed characteristic signals for the flavone scaffold and chalcone precursors. Additionally, signals of a triazole ring (δH-3 8.61–7.73 s, δC2 144.1–141.6 and δC3 125.2–121.5) and a glycosyl moiety were observed. The position of the triazole ring on these compounds was evidenced by the correlation

found in the heteronuclear multiple bond correlation (HMBC) spectra between the proton signals of H-1 and the carbon signals of C-2 and C-3.

#### *2.2. Mussel (Mytilus galloprovincialis) Larvae Anti-Settlement Activity*

Mussels are one of the main macrofouling organisms present on ships and submerged maritime structures worldwide; thus, they are a target species used in settlement inhibition bioassays [28,29]. Due to the presence of a muscular sensory foot, mussel plantigrade larvae are highly specialized in adhesion to the submerged surfaces and the fixation is made through the production of byssal threads [30], which constitutes the endpoint of this bioassay. Therefore, for the evaluation of the AF activity of the compounds towards macrofouling species, the ability of the synthetized flavonoids to inhibit the settlement of *Mytilus galloprovincialis* larvae at 50 μM was assessed. In this screening bioassay, in addition to glycosylated flavones **3a**, **3b**, **4a** and **4b** and chalcones **7a**, **7b**, **8a** and **8b**, non-glycosylated flavones **1a**–**b** and **2a**–**b** and chalcones **6a**–**b**, **10a**–**b** and **11a**–**b** were tested in order to perform SAR studies. Results showed that among 18 tested flavonoids (10 chalcones and 8 flavones), seven chalcones (**6a**, **6b**, **7b**, **8a**, **8b**, **11a** and **11b**) and only three flavones (**1b**, **4a** and **4b**) presented a percentage of settlement ≤ 40%, suggesting that chalcone scaffold seems to be more promising for anti-settlement activity. These 10 compounds were further selected for dose–response studies in order to determine LC50/EC50 values.

Among these, three chalcones (**7b**, **11a**, **11b**) and one flavone (**1b**) revealed effective anti-settlement activity (EC50 < 25 μg/mL), with triazolyl glycosylated chalcone **7b** being the most potent (EC50 = 3.28 <sup>μ</sup>M; 2.43 <sup>μ</sup>g·mL−1), showing the highest therapeutic ratio (> 60.98) (Table 1).


**Table 1.** Antifouling (AF) effectiveness and toxicity parameters of flavones **1b**, **4a** and **4b** and chalcones **6a**, **6b**, **7b**, **8a**, **8b**, **11a** and **11b** towards mussel plantigrade larvae.

EC50: minimum concentration that inhibited 50% of larval settlement; LC50: median lethal dose; LC50/EC50: therapeutic ratio; CI: confidence interval. EC50 are recommend to be less than 25 μg/mL and therapeutic ratio higher than 15 for effective AF agents [31].

#### *2.3. Quantitative Structure—Activity Relationship*

Quantitative structure–activity relationship (QSAR) studies have been used for several years to point out small molecules' properties that are relevant for activity, and to forecast the activity of new compounds [32]. Therefore, a QSAR model was built to highlight the descriptors that are being relevant for anti-settlement activity against *M. galloprovincialis* plantigrades of the tested flavonoids. In this work, a 2D-QSAR model was elaborated using the Comprehensive Descriptors for Structural and Statistical Analysis (CODESSA 2.7.2) software package, which calculates approximately 500 descriptors. The heuristic method performs a pre-selection of descriptors by eliminating descriptors that are not available for each structure, that have a small variation in magnitude, that are correlated pairwise, and that have no statistical significance. The heuristic method is a very useful method for searching the best set of descriptors, without restrictions on the data set size [33].

The correlation coefficient (R2), squared standard error (S2), and Fisher's value (F) were used to evaluate the validity of regression equation [34]. As the rules of QSAR establish that there must be one descriptor for each five molecules used to build the model [34], three descriptors were used to build the QSAR equation. The multilinear regression analysis using Heuristic method for 15 compounds in the three-descriptor model is shown in Figure 1. The compounds are uniformly distributed around the regression line (Figure 1), which suggests that the obtained model has satisfactory predictive ability.

**Figure 1.** Quantitative structure-activity relationship (QSAR) model obtained with the heuristic method for 15 molecules with the CODESSA software (R2 = 0.7945, F = 14.18, S<sup>2</sup> = 0.0243). X, ΔX and *t*-test are the regression coefficient of the linear model, standard errors of the regression coefficient, and the t significance coefficient of the determination, respectively. AF = antifouling activity.

The best QSAR equation had a R<sup>2</sup> of 0.7945, Fisher value of 14.18, and S<sup>2</sup> of 0.0243, which reveals that the proposed model has statistical validity [35]. The R2 is higher than 0.6, which is an indicator of a good fit to the regression line [36], representing close to 80% of the total variance in AF activity shown by the test compounds. The QSAR model is significant at a 95% level, as shown by the Fisher F-test (F = 14.18), which is higher than the tabulated value (3.59), as desired for a statistically significant model [35]. The squared standard deviation S2 is small and close to zero (s<sup>2</sup> = 0.0243), proving that the model is significant and has low variation about the regression line [37]. The reliability of the resulting QSAR model was explored using two different types of validation criteria: external validation by using a test set and internal validation by leave-one-out (LOO) cross-validation [38]. The model was able to predict the activity of an external test set with an average difference of 0.19 from the experimental value [39]. Moreover, the cross-validated R2 (Q2 = 0.5953) from the LOO internal validation process is higher than 0.5 and smaller than the overall R2, as expected, and the difference between R<sup>2</sup> and Q2 is lower than 0.3, which indicates that the model does not suffer from overfitting [40].

By interpreting the molecular descriptors in the regression model (Figure 1), it is possible to have some insight into structural characteristics that are likely to be responsible for AF activity of the studied compounds. There are three descriptors included in the regression model, which proved to be important features and provide statistically significant contributions to the QSAR equation.

As indicated by the higher t-test value, hydrogen bonding acceptor ability of the molecule (HACA1) is a charged partial surface area (CPSA) descriptor that appeared as the most significant descriptor for the obtained QSAR model, contributing positively to the AF activity [41]. HACA1 is determined by the equation:

$$\text{HACA1} = \sum\_{\mathbf{A}} \mathbf{S}\_{\mathbf{A}} \mid \mathbf{A} \in \mathbb{X}\_{\text{H}-\text{accept}} \tag{1}$$

where *SA* stands for solvent-accessible surface area of H-bonding acceptor atoms, selected by threshold charge. This descriptor proves the importance of the hydrogen bonding acceptor properties for the activity of the test compounds [42].

The topological descriptor average complementary information content of order 2 (CIC2) descriptor is predicted as being negatively implied in the AF activity of the test compounds [41]. The CIC2 descriptor represents the difference between the maximum possible complexity of a molecule and its real topological information. It belongs to the multi-graph information content indices and it describes neighborhood symmetry of second order [43]. The constitutional descriptor number of triple bonds is also responsible for a decrease in activity.

The molecular descriptors used in the QSAR model demonstrate that the mechanism underlying the AF activity of flavonoids is mainly related to their HACA1, and it may be prejudiced by topological CIC2 and by the presence of triple bonds. Interestingly, the triazolyl glycosylated chalcone **7b**, with the most promising anti-settlement activity, is one of the compounds with more hydrogen-bonding acceptors. In contrast, propargylated flavones **2a** and **2b** had a percentage of settlement higher than 40% at 50 μM, and therefore were not selected for dose–response studies and for the determination of the LC50/EC50 values. Moreover, propargylated chalcones **6a** and **6b** showed the lowest activity. Overall, the examination of the molecular descriptors reported in this work can lead to a better understanding of the relation between the structure and AF activity of flavonoids.

#### *2.4. Biofilm-Forming Marine Bacteria Growth Inhibitory Activity*

Although the macrofouling species represent the most problematic component of fouling in terms of biomass and negative repercussions, the first micro-colonizers are also of extreme importance, since they represent the basis of the fouling community, and ultimately, they may modulate the colonization of further species by inducing or inhibiting species adhesion via biochemical cues [44]. Thus, synthesized flavonoids were further evaluated for their ability to inhibit the growth of five marine biofilm-forming bacteria, *Vibrio harveyi, Cobetia marina, Halomonas aquamarina, Pseudoalteromonas atlantica* and *Roseobacter litoralis*.

Results showed that only the bacterial growth of *Roseobacter litoralis* was meaningfully compromised by tested compounds, with significant inhibitory activity for propargylated chalcones **6a** and **6b** (Figure 2). These compounds were selected for concentration–response analysis (Figure 3).

**Figure 2.** Bacterial growth inhibition screening of flavonoid derivatives (15 μM) towards five biofilm-forming marine bacteria: *Vibrio harveyi*, *Cobetia marina*, *Halomonas aquamarina*, *Pseudoalteromonas atlantica* and *Roseobacter litoralis*. B: Negative control with 1% dimethyl sulfoxide (DMSO); C: positive control with penicillin–streptomycin–neomycinstabilized solution. \* indicates significant differences at *p* < 0.05 (Dunnett test), against the negative control (B).

**Figure 3.** Concentration–response growth inhibition activity of compounds **6a** and **6b** towards *Roseobacter litoralis*. B: DMSO negative control; C: positive control with penicillin–streptomycin–neomycin-stabilized solution.

Compounds **6a** and **6b** presented low anti-bacterial activity towards *R. litoralis* with EC30 values of 135 and 83.5 μM, respectively.

#### *2.5. Biofilm—Forming Marine Diatoms Growth Inhibitory Activity*

The most promising compounds regarding anti-settlement activity (**1b**, **7b**, **11a**, **11b**) were further evaluated for their ability to inhibit the growth of the biofilm-forming

microalgae *Navicula* sp. This marine diatom is a major biofouling species that very effectively colonizes submerged surfaces by secreting adhesive extracellular polymer substances (EPS), and thus is a good representative of fouling microalgae.

Only triazolyl glycosylated chalcone **7b** showed significant inhibitory activity with the concentration–response analyses revealing an EC50 value of 41.76 <sup>μ</sup>M; 30.94 <sup>μ</sup>g·mL<sup>−</sup>1, suggesting the ability of this compound to act also as a promising AF agent against microfouling species.

#### *2.6. In Vitro Acetylcholinesterase (AChE) and Tyrosinase (Tyr) Activities*

The identification of the mechanism of action associated with AF activity remains a challenge for the scientific community. According to Qian et al. (2013) antifoulants appear to affect settlement through distinct pathways, which can be classified roughly into several categories such as inhibitors of ion channel function, inhibitors of quorum sensing, blockers of neurotransmission or inhibitors of adhesive production or release [45]. Moreover, some specific target molecules in fouling organisms have been determined, such as AChE, which seems to be involved in cholinergic neural signaling during the settlement [46]. It is known that the commercial booster biocide Sea-Nine 211 acts by this mechanism [47,48], as well as two natural compounds isolated from marine organisms, territrem A and pulmonarin [49,50]. For this reason, the ability of the most promising compounds to modulate the activity of AChE was evaluated (**1b**, **7b**, **11a** and **11b**). AChE activity was significantly induced for chalcones **7b** and **11b** (Figure 4). Induced AChE activity has been described as an exposure effect that is in some cases associated with apoptosis [51], and thus the specific target behind these compounds' bioactivity should be further explored in future work.

A well-known pathway in the production of biological adhesives of mussels is the 3,4-dihydroxyphenyl-L-alanine (L-DOPA) metabolism that functions in the production of DOPA-containing mussel byssal plaques by the action of Tyr that catalyses the conversion of DOPA precursor into DOPA residues [46,52]. Considering this, the most promising compounds in the inhibition of mussel adhesion were tested for their ability to inhibit Tyr (Figure 5). Results show that flavone **1b** is able to significantly decrease Tyr activity at all the concentrations tested, reaching 23.5% of inhibition at 100 μM. Therefore, the inhibition of this enzyme, with a crucial effect in the formation of mussel adhesive, could be one of the mechanisms involved in the inhibition of the mussel settlement. This also highlights a specific AF mode of action related with mussel adhesion and explains the absence of activity against bacteria and diatoms.

**Figure 5.** Tyr activity of the most promising compounds **1b, 7b**, **11a** and **11b**. B: Dimethyl sulfoxide (DMSO) (1% in water). C: Kojic acid (1.4 mM, water). \* indicates significant differences at *p* < 0.05 (Dunnett test), against the negative control (B).

#### *2.7. Environmental Fate Parameters: Artemia Salina Ecotoxicity Bioassay*

Ecotoxicity assays carried out on non-target organisms aim to understand how tested compounds can affect sensitive non-target organisms and influence the health status of the surrounding ecosystem [53]. *Artemia salina* is a species of small crustaceans that live in salty marine environments and are used as test organisms because of their easy culture, short generation time, cosmopolitan distribution and commercial availability of their eggs in latent form [54].

Ecotoxicity results showed that the most promising compounds **1b**, **7b**, **11a** and **11b** are non-toxic to *Artemia salina* (less than 10% mortality) at both concentrations tested (25 and 50 μM) (Figure 6), in contrast to the commercial AF agent ECONEA® which was previously shown by our group to cause 100% lethality at the same concentrations and conditions [9]. These results suggest that any of the tested compounds could be a good alternative, being more environmentally compatible antifoulants.

**Figure 6.** Mortality rate of *Artemia salina* nauplii after 48 h of exposure to compounds **1b**, **7b**, **11a** and **11b**, B: DMSO (1% in filtered seawater). C: K2Cr2O7 (13.6 μM, filtered seawater).

#### **3. Materials and Methods**

#### *3.1. Synthesis and Structure Elucidation of Chalcones and Flavones*

MW reactions were performed using a glassware setup for atmospheric pressure reactions and a 100 mL Teflon reactor (internal reaction temperature measurements with a fiber-optic probe sensor) and were carried out using an Ethos MicroSYNTH 1600 Microwave Labstation from Milestone (Thermo Unicam, Oeiras, Portugal). The reactions were monitored by analytical thin-layer chromatography (TLC) Macherey-Nagel Silica gel 60 F254 (Macherey-Nagel, Dueren, Germany), Purifications of compounds were carried out by flash chromatography using Macherey-Nagel silica gel 60 (0.04–0.063 mm) (Macherey-Nagel, Dueren, Germany), preparative TLC using Macherey-Nagel silica gel 60 (GF254) (Macherey-Nagel, Dueren, Germany) plates and crystallization. Melting points were obtained in a Köfler microscope (Wagner and Munz, Munich, Germany) and are uncorrected. 1H and 13C NMR spectra were taken in CDCl3 or DMSO-d6 at room temperature, on Bruker Avance 300 and 500 instruments (Bruker Biosciences Corporation, Billerica, MA, USA) (300.13 MHz or 500 MHz for 1H and 75.47 or 120 MHz for 13C). Chemical shifts are expressed in δ (ppm) values relative to tetramethylsilane (TMS) as an internal reference; 13C NMR assignments were made by 2D (HSQC and HMBC) NMR experiments (longrange 13C-1H coupling constants were optimized to 7 Hz). HRMS mass spectra of compounds **2a**, **2b**, **3a**, **3b**, **4a**, **4b**, **7a**, **7b** and **8b** were performed on an APEXQe FT-ICR MS (Bruker Daltonics, Billerica, MA) equipped with a 7T actively shielded magnet, at C.A.C.T.I.—University of Vigo, Spain. Ions were generated using a Combi MALDIelectrospray ionization (ESI) source. HRMS mass spectrometry of compound **8a** was performed on an LTQ OrbitrapTM XL hybrid mass spectrometer (Thermo Fischer Scientific, Bremen, Germany) controlled by LTQ Tune Plus 2.5.5 and Xcalibur 2.1.0. at CEMUP— University of Porto, Portugal. Phloroglucinol, ethyl 3,4-dimethoxybenzoylacetate, ethyl 3,4,5-trimethoxybenzoylacetate, 2,4-dihydroxyacetophenone and 2,3,4,6-tetra-*O*-acetylα-D-glucopyranosyl bromide were purchased from Sigma Aldrich (St. Louis, MO, USA). 3,4-Dimethoxybenzaldehyde and 3,4,5-trimethoxybenzaldehyde were purchased from Acros Organics (Janssen Pharmaceuticalaan, Geel, Belgium). 2-Azidoethyl-2,3,4,6-tetra-*O*acetyl-β-D-glucopyranoside was purchased from Synthose (Concord, ON, Canada).

#### 3.1.1. Synthesis of Flavones **1a** and **1b**

A mixture of phloroglucinol (0.175 g, 1.39 mmol) and ethyl 3,4-dimethoxybenzoylacetate (0.700 g, 2.78 mmol) or ethyl 3,4,5-trimethoxybenzoylacetate (0.739 g, 2.78 mmol) was heated at 240 ◦C in muffle furnace (Thermo Fisher Scientific, Oeiras, Portugal) for 60–100 min. Afterwards, the crude mixture was dissolved in 10% NaOH (20 mL) and washed with diethyl ether (2 × 20 mL), and the product was precipitated by adding 37% HCl. The solid was filtered and washed with water, and the flavones **1a** and **1b** were obtained with 74% and 77% yields, respectively. The structure elucidation of compounds **1a** and **2b** was established by 1H and 13C NMR techniques and data were in accordance with previously reported results [19].

#### 3.1.2. Synthesis of 7-*O*-Propargylflavones **2a** and **2b**

To a solution of **1a** (0.200 g, 0.64 mmol) or **1b** (0.200 g, 0.58 mmol), cesium carbonate (0.207 g, 0.64 mmol or 0.189 g, 0.58 mmol), tetrabutylammonium bromide (TBAB) (0.205 g, 0.64 mmol or 0.187 g, 0.58 mmol) in anhydrous acetone (20 mL), and propargyl bromide solution, 80 wt.% in toluene (0.071 mL, 0.64 mmol or 0.065 mL, 0.58 mmol), were added. The mixture was refluxed at 60 ◦C during 6 h and filtered. The filtrate was evaporated under reduced pressure and purification was carried out by flash column chromatography (SiO2; n-hexane: ethyl acetate), 8:2) followed by crystallization in acetone.

2-(3,4-dimethoxyphenyl)-5-hydroxy-7-(prop-2-yn-1-yloxy)-4H-chromen-4-one (**2a**). Light yellow solid; Yield: 66%; m.p.: 222–224 ◦C (acetone); 1H NMR (DMSO-d6, 500 MHz), δ: 12.93 (1H, s, OH-5), 7.71 (1H, dd, J = 8.5 and 2.2 Hz, H-6 ), 7.57 (1H, d, J = 2.2 Hz, H-2 ), 7.14 (1H, d, J = 8.7 Hz, H-5 ), 7.03 (1H, s, H-3), 6.87 (1H, d, J = 2.2 Hz, H-8), 6.44 (1H, d,

J = 2.3 Hz, H-6), 4.94 (2H, d, J = 2.5 Hz, H-1), 3.88 (3H, s, 3 -OCH3), 3.85 (3H, s, 4 - OCH3), 3.66 (1H, t, J = 2.4 Hz, H-3). 13C NMR (DMSO-d6, 120 MHz) δ: 182.3 (C4), 164.0 (C2), 163.2 (C7), 161.3 (C5), 157.3 (C8a), 152.5 (C4 ), 149.2 (C3 ), 122.9 (C1 ), 120.5 (C6 ), 111.9 (C5 ), 109.6 (C2 ), 105.3 (C4a), 104.3 (C3), 98.8 (C6), 94.0 (C8), 79.2 (C3), 78.6 (C2), 56.4 (C1), 56.1, 56.0 (3 -OCH3 and 4 -OCH3). HRMS (ESI+) *m*/*z* calcd for C20H17O6 [M + H+] 353.10196, found 353.10174.

5-hydroxy-7-(prop-2-yn-1-yloxy)-2-(3,4,5-trimethoxyphenyl)-4H-chromen-4-one (**2b**). Light yellow solid; Yield: 55%; m.p.: 211–213 ◦C (acetone); 1H NMR (DMSO-d6, 300.13 MHz), δ: 12.87 (1H, s, OH-5), 7.36 (2H, s, H-2 and H-6 ), 7.16 (1H, s, H-3), 6.91 (1H, d, J = 2.3 Hz, H-8), 6.46 (1H, d, J = 2.3 Hz, H-6), 4.96 (2H, d, J = 2.4 Hz, H-1), 3.91 (6H, s, 3 - OCH3 and 5 -OCH3), 3.76 (3H, s, 4 - OCH3), 3.70 (1H, t, J = 2.4 Hz, H-3). 13C NMR (DMSO-d6, 75.47 MHz) <sup>δ</sup>: 182.2 (C4), 163.4 (C2), 163.1 (C7), 161.2 (C5), 157.1 (C8a), 153.3 (C3 and C5 ), 140.9 (C4 ), 125.8 (C1 ), 105.3 (C3), 105.2 (C4a), 104.2 (C2 and C6 ), 98.7 (C6), 93.9 (C8), 79.1 (C2), 78.4 (C3), 60.3 (4 -OCH3), 56.4 (C1, 3 -OCH3 and 5 -OCH3). HRMS (ESI+) *m*/*z* calcd for C21H19O7 [M + H+] 383.11253, found 383.11233.

#### 3.1.3. Synthesis of Flavone-Triazolyl-Glycosides **3a** and **3b**

To a solution of **2a** (0.100 g, 0.28 mmol) or **2b** (0.100 g, 0.27 mmol) and 2,3,4,6 tetra-*O*-acetyl-β-D-glucopyranosyl azide (0.106 g, 0.28 mmol or 0.102 g, 0.27 mmol) in THF/water solvent mixture (2:1; 30 mL), sodium ascorbate (0.225 g, 1.14 mmol or 0.216 g, 1.09 mmol) and copper(II) sulphate pentahydrate (0.142 g, 0.57 mmol or 0.136 g, 0.54 mmol) were added. The reaction vessel was sealed and the mixture was kept stirring and heated for 30 min at 70 ◦C under MW irradiation of 500 W. After cooling, the reaction mixture was filtered and concentrated under reduced pressure. The water suspension was extracted with ethyl acetate (2 × 20 mL), and the combined organic layers were dried over anhydrous sodium sulphate, evaporated under reduced pressure, and then purified by crystallization in acetone.

(*2R*,*3R*,*4S*,*5R*,*6R*)-2-(acetoxymethyl)-6-(4-(((2-(3,4-dimethoxyphenyl)-5-hydroxy-4-oxo-4Hchromen-7-yl)oxy)methyl)-1H-1,2,3-triazol-1-yl)tetrahydro-2H-pyran-3,4,5-triyl triacetate (**3a**). Light yellow solid; Yield: 82%; m.p.: 143–145 ◦C (acetone); 1H NMR (DMSO-d6, 500 MHz), δ: 12.93 (1H, s, OH-5), 8.61 (1H, s, H-3), 7.71 (1H, dd, J = 8.5 and 2.1 Hz, H-6 ), 7.60 (1H, d, J = 2.2 Hz, H-2 ), 7.15 (1H, d, J = 8.7 Hz, H-5 ), 7.05 (1H, s, H-3), 6.94 (1H, d, J = 2.2 Hz, H-8), 6.47 (1H, d, J = 2.2 Hz, H-6), 6.39 (1H, d, J = 9.2 Hz, H-1), 5.68 (1H, t, J = 9.4 Hz), 5.56 (1H, t, J = 9.5 Hz), 5.19 (1H, t, J = 9.8 Hz) (H-2, H-3, H-4), 5.32 (2H, s, H-1), 4.39–4.36 (1H, m, H-5), 4.15–4.07 (2H, m, H-6), 3.86 (3H, s, 4 -OCH3), 3.83 (3H, s, 3 -OCH3), 2.03, 1.99, 1.96, 1.77 (12H, s, 2, 3, 4, -COCH3). 13C NMR (DMSO-d6, 120 MHz) δ: 182.1 (C4), 170.1, 169.6, 169.4, 168.5 (2, 3, 4, 6-COCH3), 163.7 (C2 and C7), 161.2 (C5), 158.0 (C8a), 152.3 (C4 ), 149.1 (C3 ), 142.7 (C2), 124.0 (C3), 122.7 (C1 ), 120.2 (C6 ), 111.7 (C5 ), 109.5 (C2 ), 105.0 (C3), 104.1 (C4a), 98.7 (C6), 93.7 (C8), 83.9 (C1), 73.3 (C5), 72.1, 70.1, 67.5 (C2, C3, C4), 61.8 (C6), 61.7 (C1), 55.9, 55.8 (3 -OCH3 and 4 -OCH3), 20.5, 20.4, 20.3, 19.9 (2, 3, 4, 6-COCH3). HRMS (ESI+) *m*/*z* calcd for C34H36N3O15 [M + H+] 726.21409, found 726.21380.

(*2R*,*3R*,*4S*,*5R*,*6R*)-2-(acetoxymethyl)-6-(4-(((5-hydroxy-4-oxo-2-(3,4,5-trimethoxyphenyl)-4Hchromen-7-yl)oxy)methyl)-1H-1,2,3-triazol-1-yl)tetrahydro-2H-pyran-3,4,5-triyl triacetate (**3b**). Yellow solid; Yield: 79%; m.p.: 169–171 ◦C (acetone); 1H NMR (DMSO-d6, 300.13 MHz), δ: 12.85 (1H, s, OH-5), 8.61 (1H, s, H-3), 7.38 (2H, s, H-2 and H-6 ), 7.17 (1H, s, H-3), 7.00 (1H, d, J = 2.2 Hz, H-8), 6.49 (1H, d, J = 2.2 Hz, H-6), 6.39 (1H, d, J = 9.1 Hz, H-1), 5.68 (1H, t, J = 9.3 Hz), 5.56 (1H, t, J = 9.4 Hz), 5.19 (1H, t, J = 9.7 Hz) (H-2, H-3, H-4), 5.33 (2H, s, H-1), 4.40–4.36 (1H, m, H-5), 4.17–4.06 (2H, m, H-6), 3.91 (6H, s, 3 -OCH3 and 5 -OCH3), 3.76 (3H, s, 4 -OCH3), 2.03, 2.00, 1.96, 1.77 (12H, s, 2, 3, 4, 6-COCH3). 13C NMR (DMSO-d6, 75.47 MHz) <sup>δ</sup>: 182.2 (C4), 170.1, 169.6, 169.4, 168.5 (2, 3, 4, 6-COCH3), 163.8 (C7), 163.4 (C2), 161.2 (C5), 157.3 (C8a), 153.3 (C3 and C5 ), 142.7 (C2), 140.9 (C4 ), 125.8 (C1 ), 123.9 (C3), 105.1 (C3 and C4a), 104.2 (C2 and C6 ), 98.7 (C6),

93.9 (C8), 83.9 (C1), 73.3 (C5), 72.1, 70.1, 67.5 (C2, C3, C4), 61.8 (C1 and C6), 60.3 (4 -OCH3), 56.3 (3 -OCH3 and 5 -OCH3), 20.5, 20.4, 20.3, 19.9 (2, 3, 4, 6-COCH3). HRMS (ESI+) *m*/*z* calcd for C35H38N3O16 [M + H+] 756.22466, found 756.22445.

#### 3.1.4. Synthesis of Flavone-Triazolyl-Glycosides **4a** and **4b**

To a solution of **2a** (0.090 g, 0.26 mmol) or **2b** (0.100 g, 0.26 mmol) and 2-azidoethyl-2,3,4,6-tetra-*O*-acetyl-β-D-glucopyranoside (0.109 g, 0.26 mmol) in tetrahydrofuran/water solvent mixture (2:1; 30 mL), sodium ascorbate (0.207 g, 1.05 mmol) and copper(II) sulphate pentahydrate (0.131 g, 0.52 mmol) were added. The reaction vessel was sealed and the mixture was kept stirring and heated for 30 min at 70 ◦C under MW irradiation of 500 W. After cooling, the reaction mixture was filtered and concentrated under reduced pressure. The water suspension was extracted with ethyl acetate (2 × 20 mL), and the combined organic layers were dried over anhydrous sodium sulphate, evaporated under reduced pressure, and then purified by crystallization in ethyl acetate/n-hexane (**4a**) or by flash column chromatography (SiO2; n-hexane: ethyl acetate, 3:7) (**4b**).

(*2R*,*3R*,*4S*,*5R*,*6R*)-2-(acetoxymethyl)-6-(2-(4-(((2-(3,4-dimethoxyphenyl)-5-hydroxy-4-oxo-4H-chromen-7-yl)oxy)methyl)-1H-1,2,3-triazol-1-yl)ethoxy)tetrahydro-2H-pyran-3,4,5-triyl triacetate (**4a**). Light brown solid; Yield: 60%; m.p.: 99–100 ◦C (n-hexane: ethyl acetate); 1H NMR (DMSO-d6, 300.13 MHz), δ: 12.94 (1H, s, OH-5), 8.15 (1H, s, H-3), 7.72 (1H, dd, J = 8.5, 1.8 Hz, H-6 ), 7.59 (1H, d, J = 1.9 Hz, H-2 ), 7.14 (1H, d, J = 8.6 Hz, H-5 ), 7.05 (1H, s, H-3), 6.96 (1H, d, J = 2.1 Hz, H-8), 6.47 (1H, d, J = 2.1 Hz, H-6), 5.28 (2H, s, H-1), 5.23 (1H, t, J = 9.6 Hz), 4.90 (1H, t, J = 9.7 Hz), 4.77–4.71 (1H, m) (H-2, H-3, H-4), 4.72 (1H, d, J = 8.1 Hz, H-1), 4.60–4.57 (2H, m, H-6), 4.19–4.01 (5H, m, H-1, H-2, H-5), 3.89 (3H, s, 3 -OCH3), 3.86 (3H, s, 4 -OCH3), 2.02, 1.98, 1.92, 1.89 (12H, s, 2, 3, 4, 6-COCH3). 13C NMR (DMSO-d6, 75.47 MHz) δ: 182.1 (C4), 170.1, 169.6, 169.3, 169.0 (2, 3, 4, 6-COCH3), 163.9 (C7), 163.7 (C2), 161.2 (C5), 157.2 (C8a), 152.3 (C4 ), 149.0 (C3 ), 141.7 (C2), 125.2 (C3), 122.8 (C1 ), 120.2 (C6 ), 111.7 (C5 ), 109.5 (C2 ), 105.0 (C4a), 104.1 (C3), 99.2 (C1), 98.6 (C6), 93.5 (C8), 71.9, 70.7, 68.1 (C2, C3, C4), 70.6, 67.4, 61.7 (C1, C2, C5), 61.9 (C1), 55.9, 55.8 (3 -OCH3 and 4 -OCH3), 49.4 (C6), 20.5, 20.4, 20.3, 20.3 (2, 3, 4, 6-COCH3). HRMS (ESI+) *m*/*z* calcd for C36H40N3O16 [M + H+] 770.24031, found 770. 23792.

(*2R*,*3R*,*4S*,*5R*,*6R*)-2-(acetoxymethyl)-6-(2-(4-(((5-hydroxy-4-oxo-2-(3,4,5-trimethoxyphenyl)-4Hchromen-7-yl)oxy)methyl)-1H-1,2,3-triazol-1-yl)ethoxy)tetrahydro-2H-pyran-3,4,5-triyl triacetate (**4b**). Light yellow solid; Yield: 49%; m.p.: 96–98 ◦C (n-hexane: ethyl acetate); 1H NMR (DMSO-d6, 300.13 MHz), δ: 12.86 (1H, s, OH-5), 8.15 (1H, s, H-3), 7.38 (2H, s, H-2 and H-6 ), 7.17 (1H, s, H-3), 7.02 (1H, d, J = 2.2 Hz, H-8), 6.49 (1H, d, J = 2.2 Hz, H-6), 5.29 (2H, s, H-1), 5.23 (1H, t, J = 9.5 Hz), 4.90 (1H, t, J = 9.7 Hz), 4.74 (1H, t, J = 8.8 Hz) (H-2, H-3, H-4), 4.84 (1H, d, J = 8.0, H-1), 4.60–4.56 (2H, m, H-6), 4.21–4.01 (5H, m, H-5, H-1 and H-2), 3.91 (6H, s, 3 -OCH3 and 5 -OCH3), 3.76 (3H, s, 4 -OCH3), 2.02, 1.98, 1.92, 1.89 (12H, s, 2, 3, 4, 6-COCH3). 13C NMR (DMSO-d6, 75.47 MHz) δ: 182.2 (C4), 170.1, 169.6, 169.3, 169.0 (2, 3, 4, 6-COCH3), 164.0 (C7), 163.4 (C2), 161.1 (C5), 157.3 (C8a), 153.3 (C3 and C5 ), 141.6 (C2), 140.9 (C4 ), 125.8 (C1 ), 125.2 (C3), 105.3 (C4a), 105.1 (C3), 104.2 (C2 and C6 ), 99.2 (C1), 98.7 (C6), 93.7 (C8), 71.9, 70.7, 70.6, 68.1 (C2, C3, C4 and C5), 67.4, 61.7 (C1, C2), 62.0 (C1), 60.3 (4 -OCH3), 56.4 (3 and 5 -OCH3), 49.4 (C6), 20.5, 20.4, 20.3, 20.3 (2, 3, 4, 6-COCH3). HRMS (ESI+) *m*/*z* calcd for C37H42N3O17 [M + H+] 800.25087, found 800.24829.

#### 3.1.5. Synthesis of Propargyloxyacetophenone **5**

To a solution of 2,4-dihydroxyacetophenone (1.00 g, 6.57 mmol) and potassium carbonate (0.91 g, 6.57 mmol) in anhydrous acetone (20 mL), propargyl bromide solution, 80 wt.% in toluene (0.73 mL, 6.57 mmol) was added. The mixture was refluxed at 60 ◦C during 3 h. Then, the reaction mixture was filtered, evaporated under reduced pressure and purified by flash column chromatography (SiO2; n-hexane: ethyl acetate), 9:1), giving rise to **5** with

76% yield. The structure elucidation of compound **5** was established by 1H and 13C NMR techniques and data were in accordance with previously reported results [23].

#### 3.1.6. Synthesis of Propargyloxychalcones **6a** and **6b**

To a solution of **5** (0.350 g, 1.84 mmol) in methanol (20 mL) was added a solution of 40% NaOH in methanol, until pH 14, under stirring. Afterwards, 3,4-dimethoxybenzaldehyde (0.612 g, 3.68 mmol) or 3,4,5-trimethoxybenzaldehyde (0.772 g, 3.68 mmol) was slowly added to the reaction mixture. The reaction was submitted to MW irradiation at 180 W at 70 ◦C for 4 h. After, a solution of 10% HCl was added until pH 5, and the obtained solid was filtered, washed with water, and purified by crystallization with methanol, giving rise to chalcone **6a** and **6b** with 41% and 43% yield, respectively. The structure elucidation of compounds **6a** and **6b** was established by 1H and 13C NMR techniques and data were in accordance with previously reported results [23].

#### 3.1.7. Synthesis of Chalcone-Triazolyl-Glycosides **7a** and **7b**

To a solution of **6a** (0.140 g, 0.41 mmol) or **6b** (0.200 g, 0.54 mmol) and 2,3,4,6-tetra-*O*-acetyl-β-D-glucopyranosyl azide (0.309 g, 0.82 mmol or 0.405 g, 1.09 mmol) in tetrahydrofuran (THF)/water solvent mixture (2:1; 30 mL), sodium ascorbate (0.328 g, 1.66 mmol or 0.430 g, 2.17 mmol) and copper(II) sulphate pentahydrate (0.207 g, 0.83 mmol or 0.271 g, 1.09 mmol) were added. The reaction mixture was heated for 1 h at 70 ◦C under MW irradiation of 250 W with agitation. After cooling, the reaction mixture was filtered and concentrated under reduced pressure. The water suspension was extracted with ethyl acetate (2 × 20 mL) and the combined organic layers were washed with water (1 × 20 mL), dried over anhydrous sodium sulphate, evaporated under reduced pressure, and then purified by crystallization with methanol.

(*2R*,*3R*,*4S*,*5R*,*6R*)-2-(acetoxymethyl)-6-(4-((4-(€-3-(3,4-dimethoxyphenyl)acryloyl)-3- hydroxyphenoxy)methyl)-1H-1,2,3-triazol-1-yl)tetrahydro-2H-pyran-3,4,5-triyl triacetate (**7a**). Yellow solid; yield: 51%; m.p.: 169–171 ◦C (methanol); 1H NMR (CDCl3, 300.13 MHz), δ: 13.49 (1H, s, OH-2 ), 7.89 (1H, s, H-3), 7.86 (1H, d, J = 9.3 Hz, H-6 ), 7.84 (1H, d, J = 15.5 Hz, H-β), 7.43 (1H, d, J = 15.5 Hz, H-α), 7.26–7.22 (1H, m, H-6), 7.16 (1H, d, J= 1.9 Hz, H-2), 6.90 (1H, d, J= 8.3 Hz, H-5), 6.58–6.55 (2H, m, H-3 and H-5 ), 5.90 (1H, d, J= 9.2 Hz, H-1), 5.48–5.38 (2H, m), 5.27–5.18 (1H, m) (H-2, H-3,H-4), 5.24 (2H, s, H-1), 4.33–4.11 (2H, m, H-6), 4.04–3.98 (1H, m, H-5), 3.96 (3H, s, 3-OCH3), 3.93 (3H, s, 4-OCH3), 2.07, 2.06, 2.02, 1.86 (12H, s, 2, 3, 4, 6-COCH3). 13C NMR (CDCl3, 75.47 MHz) δ: 192.0 (CO), 170.6, 170.0, 169.5, 169.1 (2, 3, 4, 6-COCH3), 166.6 (C2 ), 164.5 (C4 ), 151.8 (C3), 149.4 (C4) 145.0 (Cβ), 144.1 (C2), 131.4 (C6 ), 127.8 (C1), 123.6 (C6), 121.5 (C3), 118.0 (Cα), 114.7 (C1 ), 111.3 (C5), 110.3 (C2), 107.9 (C5 ), 102.3 (C3 ), 85.9 (C1), 75.3 (C5), 72.7, 70.4, 67.8 (C2, C3, C4), 62.0 (C1), 61.6 (C6), 56.2 (3-OCH3), 56.1 (4-OCH3), 20.8, 20.7, 20.6, 20.3 (2, 3, 4, 6-COCH3). HRMS (ESI+) *m*/*z* calcd for C34H38N3O14 [M + H+] 712.23483, found 712.23337.

(*2R*,*3R*,*4S*,*5R*,*6R*)-2-(acetoxymethyl)-6-(4-((3-hydroxy-4-((*E*)-3-(3,4,5-trimethoxyphenyl) acryloyl)phenoxy)methyl)-1H-1,2,3-triazol-1-yl)tetrahydro-2H-pyran-3,4,5-triyl triacetate (**7b**). Yellow solid; yield: 57%; m.p.: 127–128 ◦C (quamarnol); 1H NMR (CDCl3, 300.13 MHz), δ: 13.41 (1H, s, OH-2 ), 7.89 (1H, s, H-3), 7.87 (1H, d, J = 8.8 Hz, H-6 ), 7.81 (1H, d, J = 15.4 Hz, H-β), 7.45 (1H, d, J = 15.4 Hz, H-α), 6.87 (2H, s, H-2 and H-6), 6.60–6.56 (2H, m, H-3 and H-5 ), 5.91–5.88 (1H, m, H-1), 5.48–5.39 (2H, m), 5.27–5.21 (1H, m) (H-2, H-3, H-4), 5.25 (2H, s, H-1), 4.34–4.13 (2H, m, H-6), 4.03–4.00 (1H, m, H-5), 3.93 (6H, s, 3-OCH3 and 5-OCH3), 3.91 (3H, s, 4-OCH3), 2.08, 2.07, 2.02, 1.87 (12H, s, 2, 3, 4, 6- COCH3). 13C NMR (CDCl3, 75.47 MHz) δ: 191.9 (CO), 170.6, 170.0, 169.5, 169.1 (2, 3, 4, 6-COCH3), 166.6 (C2 ), 164.6 (C4 ), 153.7 (C3 and C5), 145.0 (Cβ), 144.1 (C2), 140.8 (C4), 131.5 (C6 ), 130.3 (C1), 121.5 (C3), 119.5 (Cα), 114.7 (C1 ), 108.0 (C5 ), 105.9 (C2 and C6), 102.3 (C3 ), 86.0 (C1), 75.4 (C5), 72.7, 70.4, 67.8 (C2, C3, C4), 62.1 (C1), 61.6 (C6),

#### 61.2 (4-OCH3), 56.4 (3-OCH3 and 5-OCH3), 20.8, 20.7, 20.6, 20.3 (2, 3, 4, 6-COCH3). HRMS (ESI+) *m*/*z* calcd for C35H40N3O15 [M + H+] 742.24539, found 742.24366.

#### 3.1.8. Synthesis of Chalcone-Triazolyl-Glycosides **8a** and **8b**

To a solution of **6a** (0.050 g, 0.15 mmol) or **6b** (0.2500 g, 0.68 mmol) and 2-azidoethyl-2,3,4,6-tetra-*O*-acetyl-β-D-glucopyranoside (0.123 g, 0.30 mmol or 0.566 g, 1.36 mmol) in THF/water solvent mixture (2:1; 30 mL), sodium ascorbate (0.117 g, 0.59 mmol or 0.538 g, 2.71 mmol) and copper(II) sulphate pentahydrate (0.074 g, 0.30 mmol or 0.339 g, 1.36 mmol) were added. The reaction mixture was heated for 1 h at 70 ◦C under MW irradiation of 250 W with agitation. After cooling, the reaction mixture was filtered and the THF in the filtrate was evaporated under reduced pressure. Then, the water suspension was extracted with ethyl acetate (2 × 20 mL) and the combined organic layers were washed with water (1 × 20 mL), dried over anhydrous sodium sulphate, concentrated under reduced pressure, and then purified by preparative TLC (SiO2; n-hexane: ethyl acetate, 2:8) (**8a**) or flash column chromatography (SiO2; n-hexane: ethyl acetate, 5:5) (**8b**).

(*2R*,*3R*,*4S*,*5R*,*6R*)-2-(acetoxymethyl)-6-(2-(4-((4-((E)-3-(3,4-dimethoxyphenyl)acryloyl)-3 hydroxyphenoxy)methyl)-1H-1,2,3-triazol-1-yl)ethoxy)tetrahydro-2H-pyran-3,4,5-triyl triacetate (**8a**). Yellow solid; yield: 65%; m.p.: 77–79 ◦C (n-hexane: ethyl acetate); 1H NMR (CDCl3, 300.13 MHz), δ: 13.49 (1H, s, OH-2 ), 7.87 (1H, d, J = 9.7 Hz, H-6 ), 7.85 (1H, d, J = 15.4 Hz, H-β), 7.73 (1H, s, H-3), 7.44 (1H, d, J = 15.4 Hz, H-α), 7.25 (1H, dd, J = 8.1, 1.8, H-6), 7.16 (1H, d, J = 1.8, H-2), 6.91 (1H, d, J = 8.4, H-5), 6.60–6.56 (2H, m, H-3 and H-5 ), 5.23 (2H, s, H-1), 5.18 (1H, t, J = 9.4), 5.06 (1H, t, J = 9.6), 5.03–4.97 (1H, m) (H-2, H-3, H-4), 4.68–4.52 (2H, m, H-6), 4.48 (1H, d, J = 7.9, H-1), 4.27–4.21 (2H, m), 4.15–4.10 (1H, m), 3.90–3.81 (1H, m) (H-1, H-2), 3.96 (3H, s, 3-OCH3), 3.94 (3H, s, 4-OCH3), 3.72–3.66 (1H, m, H-5), 2.08, 2.01, 1.99, 1.95 (12H, s, 2, 3, 4, 6-COCH3). 13C NMR (CDCl3, 75.47 MHz) δ: 192.0 (CO), 170.7, 170.3, 169.6, 169.5 (2, 3, 4, 6-COCH3), 166.6 (C2 ), 164.6 (C4 ), 151.8 (C4), 149.4 (C3) 144.9 (Cβ), 143.1 (C2), 131.5 (C6 ), 127.9 (C1), 124.6 (C3), 123.6 (C6), 118.1 (Cα), 114.7 (C1 ), 111.3 (C5), 110.3 (C2), 107.9 (C5 ), 102.3 (C3 ), 100.6 (C1), 72.1 (C5), 72.6, 71.1, 68.3 (C2, C3and C4), 67.8, 61.8 (C1 and C2), 62.1 (C1), 56.2 (3 -OCH3), 56.1 (4 -OCH3), 50.3 (C6), 20.9, 20.7, 20.7, 20.7 (2, 3, 4, 6-COCH3). HRMS (ESI+) *m*/*z* calcd for C36H42N3O15 [M + H+] 756.261044, found 756.26373.

(*2R*,*3R*,*4S*,*5S*,*6S*)-2-(acetoxymethyl)-6-(2-(4-((3-hydroxy-4-((*E*)-3-(3,4,5-trimethoxyphenyl) acryloyl)phenoxy)methyl)-1H-1,2,3-triazol-1-yl)ethoxy)tetrahydro-2H-pyran-3,4,5-triyl triacetate (**8b**). Yellow solid; yield: 45%; m.p.: 78–81 ◦C (n-hexane: ethyl acetate); 1H NMR (CDCl3, 300.13 MHz), δ: 13.41 (1H, s, OH-2 ), 7.87 (1H, d, J = 9.8 Hz, H-6 ), 7.81 (1H, d, J = 15.4 Hz, H-β), 7.74 (1H, s, H-3), 7.46 (1H, d, J = 15.4 Hz, H-α), 6.87 (2H, s, H-2 and H-6), 6.61–6.57 (2H, m, H-3 and H-5 ), 5.24 (2H, s, H-1), 5.18 (1H, t, J= 9.4), 5.07 (1H, t, J = 9.6), 4.99 (1H, t, J = 8.7) (H-2, H-3, H-4), 4.68–4.52 (2H, m, H-6), 4.48 (1H, d, J = 7.9, H-1), 4.28–4.21 (2H, m), 4.13–4.08 (2H, m) (H-1 and H-2), 3.93 (3H, s, 4-OCH3), 3.91 (6H, s, 3-OCH3 and 5-OCH3), 3.72–3.66 (1H, m, H-5), 2.04, 2.02, 1.99, 1.95 (12H, s, 2, 3, 4, 6-COCH3). 13C NMR (CDCl3, 75.47 MHz) δ: 191.9 (CO), 170.7, 170.3, 169.6, 169.5 (2, 3, 4, 6-COCH3), 166.7 (C2 ), 164.8 (C4 ), 153.6 (C3 and C5), 144.9 (Cβ), 143.0 (C2), 140.7 (C4), 131.5 (C6 ), 130.4 (C1), 124.7 (C3), 119.6 (Cα), 114.6 (C1 ), 108.0 (C5 ), 105.9 (C2 and C6), 102.3 (C3 ), 100.6 (C1), 72.1 (C5), 72.6, 71.1, 68.3 (C2, C3 and C4), 67.8, 61.9 (C1 and C2), 62.1 (C1), 60.5 (4-OCH3), 56.4 (3-OCH3 and 5-OCH3), 50.3 (C6), 21.2, 20.9, 20.7, 20.7 (2, 3, 4, 6-COCH3). HRMS (ESI+) *m*/*z* calcd for C37H44N3O16 [M + H+] 786.27161, found 786.26915.

#### 3.1.9. Synthesis of Acetophenone **9**

To a solution of 2,4-dihydroxyacetophenone (1.00 g, 6.57 mmol) and potassium carbonate (2.73 g, 19.72 mmol) in anhydrous acetone (20 mL), chloromethyl methyl ether (0.749 mL, 9.86 mmol) was added and the mixture refluxed for 1 h at 60 ◦C. Then, the reaction

mixture was filtered, evaporated under reduced pressure and purified by flash column chromatography (SiO2; n-hexane: ethyl acetate, 9:1), giving rise to **9** with 84% yield. The structure elucidation of compound **9** was established by 1H and 13C NMR techniques and data were in accordance with previously reported results [55].

#### 3.1.10. Synthesis of Chalcones **10a** and **10b**

To a solution of **9** (0.500 g, 2.55 mmol) in methanol (20 mL) a solution of 40% NaOH in methanol was added until pH 14, under stirring. Then, a solution of 3,4 dimethoxybenzaldehyde (0.847 g, 5.10 mmol) or 3,4,5-trimethoxybenzaldehyde (1.00 g, 5.10 mmol) in methanol was slowly added to the reaction mixture. The reaction was submitted to MW irradiation at 180 W at 70 ◦C for 4 h. Then, a solution of 10% HCl was added until pH 5, and the obtained solid was filtered and washed with water and purified by crystallization with methanol, giving rise to **10a** and **10b** with 33% and 47% yield, respectively. The structure elucidation of both compounds was established by 1H and 13C NMR techniques and data of **10a** were in accordance with previously reported results [24]. Although the synthesis of compound **10b** has been previously reported [25], the NMR data are described here for the first tim€(*E*)-1-(2-hydroxy-4-(methoxymethoxy)phenyl)-3- (3,4,5-trimethoxyphenyl)prop-2-en-1-one (**10b**). Yellow solid; yield: 47%; m.p.: 132–134 ◦C (methanol); 1H NMR (CDCl3, 500 MHz), δ: 13.29 (1H, s, OH-2 ), 7.85 (1H, d, J = 9.1 Hz, H-6 ), 7.81 (1H, d, J = 15.4 Hz, H-β), 7.45 (1H, d, J = 15.4 Hz, H- α) 6.87 (2H, s, H-2 and H-6), 6.65 (1H, d, J = 2.4 Hz, H-3 ), 6.60 (1H, dd, J = 9.0,2.5 Hz, H-5 ), 5.23 (2H, s, H-1), 3.93 (6H, s, 3-OCH3 and 5-OCH3), 3.91 (3H, s, 4-OCH3), 3.49 (3H, s, H-2). 13C NMR (CDCl3, 120 MHz) δ: 192.0 (CO), 166.4 (C2 ), 163.8 (C4 ), 153.6 (C3 and C5), 144.9 (Cβ), 140.7 (C4), 131.4 (C6 ), 130.4 (C1), 119.5 (Cα), 115.0 (C1 ), 108.4 (C5 ), 105.9 (C2 and C6), 104.1 (C3 ), 94.2 (C1), 61.2 (4-OCH3), 56.6 (C2), 56.4 (3-OCH3 and 5-OCH3).

#### 3.1.11. Synthesis of Chalcones **11a** and **11b**

To a solution of **10a** (0.200 g, 0.58 mmol) or **10b** (0.250 g, 0.67 mmol) in methanol (10 mL), p-toluenesulfonic acid monohydrate (0.110 g, 0.58 mmol or 0.127 g, 0.67 mmol) was added. The reaction was submitted to conventional heating at 50 ◦C for 5 h. After the addiction of 10 mL of water, methanol was evaporated, and the aqueous solution was extracted with ethyl acetate (2 × 20 mL). The organic phase was washed with water (1 × 20 mL), dried over anhydrous sodium sulphate and concentrated under reduced pressure, giving rise to an orange solid. The crude product was purified by flash column chromatography (n-hexane: ethyl acetate, 8:2) (**11a**) or crystallization with chloroform (**11b**), giving rise to chalcone **11a** and **11b** with 24% and 31% yield, respectively. The structure elucidation of compounds **11a** and **11b** was established by 1H and 13C NMR techniques and data were in accordance with previously reported results [56,57].

#### 3.1.12. Synthesis of 2,3,4,6-Tetra-*O*-Acetyl-*β*-D-Glucopyranosyl Azide (**12**)

To a solution of 2,3,4,6-tetra-*O*-acetyl-α-D-glucopyranosyl bromide (1.00 g, 2.43 mmol) in acetone: water (9 mL, 2:1), sodium azide (0.197 g, 3.04 mmol) was added and the reaction mixture stirred at room temperature for 3 h. After, the acetone was evaporated under reduced pressure and a white solid was filtered. Crystallization of the solid with ethanol afforded 2,3,4,6-tetra-O-acetyl-*α*-D-glucopyranosyl azide **12** with 72% yield. The structure elucidation was established by 1H and 13C NMR techniques and data were in accordance with previously reported results [27].

#### *3.2. Mussel (Mytilus galloprovincialis) Larvae Anti-Settlement Activity*

Mussel (*Mytilus galloprovincialis*) plantigrades were collected in juvenile aggregates during low neap tides at Memória beach, Matosinhos, Portugal (41◦13 59 N; 8◦43 28 W). In laboratory, mussel plantigrade larvae (0.5–2 mm) were isolated in a binocular magnifier (Olympus SZX2-ILLT, Tokyo, Japan) to a petri dish with filtered seawater, and those with functional foot and competent exploring behaviour were selected for the bioassays. The flavonoids were screened at 50 μM in 24-well microplates with 4 well replicates per condition and 5 larvae per well, for 15 h, in the darkness at 18 ± 1 ◦C, following Almeida et al. (2015) [58]. Test solutions were obtained by dilution of the compounds stock solutions (50 mM) in DMSO and prepared with filtered seawater. All bioassays included a negative control with DMSO and a positive control with CuSO4, a potent AF agent. After the exposure period, the anti-settlement activity was determined by the presence/absence of attached byssal threads produced by each individual larvae.

All compounds that caused more than 60% of settlement inhibition (≤40% of settlement) in the screening bioassay were considered active and selected for the determination of the semi-maximum response concentration that inhibited 50% of larval settlement (EC50), at compounds concentrations of 3.12, 6.25, 12.5, 25, 50, 100, 200 μM.

#### *3.3. Quantitative Structure–Activity Relationship*

The eighteen flavonoid derivatives (**1a**, **1b**, **2a**, **2b**, **3a**, **3b**, **4a**, **4b** from Scheme 1; **6a**, **6b**, **7a**, **7b**, **8a**, **8b** from Scheme 2; and **10a**, **10b**, **11a**, **11b** from Scheme 3) were used to build a QSAR model using the experimental data obtained from the mussel (*Mytilus galloprovincialis*) larvae anti-settlement activity in vivo bioassay (AF activity = log(100/%settlement). AF activity was selected as a dependent variable in the QSAR analysis. The 18 molecules were randomly distributed into a training set (15 molecules) and a test set (3 molecules). CODESSA software (version 2.7.10, University of Florida, Gainesville, FL, USA) was used to calculate more than 500 constitutional, topological, geometrical, electrostatic, quantum-chemical and thermodynamical molecular descriptors [59]. The heuristic multilinear regression methodology was chosen to perform a complete search for the best multilinear correlations with a multitude of descriptors of the training set [60]. The 2D-QSAR model with the best square of the correlation coefficient (R2), F-test (F), and squared standard error (S2) was selected. The final model was further validated using the test set and leave-one-out (LOO) internal validation.

#### *3.4. Inhibitory Activity against Biofilm-Forming Marine Bacteria Growth*

For anti-bacterial screening, five strains of marine biofilm-forming bacteria from the Spanish Type Culture Collection (CECT): *Cobetia marina* CECT 4278, *Vibrio harveyi* CECT 525, *Halomonas aquamarina* CECT 5000, *Pseudoalteromonas atlantica* CECT 570, and *Roseobacter litoralis* CECT 5395 were used. Bacteria were inoculated and incubated for 24 h at 26 ◦C in marine broth (Difco) at an initial density of 0.1 (OD600) in 96 well flat-bottom microtiter plates and exposed to the test compounds at 15 μM. Test solutions were obtained by dilution of the compounds stock solutions (50 mM) in DMSO. Bacterial growth inhibition in the presence of the compounds was determined in quadruplicate at 600 nm using a microplate reader (Biotek Synergy HT, Vermont, USA). Negative and positive controls used were a solution of marine broth with DMSO, and a solution of marine broth with penicillin–streptomycin–neomycin, respectively. Compounds exerting a significant antibacterial activity (Dunnet test, *p* < 0.05) in the screening bioassays were selected for the determination of the effective inhibitory concentration (EC50).

#### *3.5. Inhibitory Activity against Biofilm-Forming Marine Diatom Growth*

The anti-microalgal activity of the most promising compounds was also evaluated against a benthic marine diatom, *Navicula* sp., purchased from the (Telde, Gran Canaria) Spanish Collection of Algae (BEA). Diatom cells were inoculated in f/2 medium (Sigma) at an initial concentration of 2–4 × 106 cells mL−<sup>1</sup> and grown in 96-well flat-bottom microtiter plates for 10 days at 20 ◦C. *Navicula* growth inhibition in the presence of each compound at 15 μM was determined in quadruplicate and quantified based on the difference in cell densities among the treatments, and cells were counted using a Neubauer counting chamber. A positive control with cycloheximide (3.55 μM) and a negative control with f/2 medium 0.1% DMSO were included. Compounds that showed significant inhibitory

activity in the screening assay (Dunnet test, *p* < 0.05) were selected for further determination of their effective inhibitory concentrations (EC50).

#### *3.6. In Vitro Acetylcholinesterase (AChE) and Tyrosinase (Tyr) Activities*

The ability of the most promising compounds to inhibit AChE and Tyr was tested to assess their potential mode of action related with neurotransmission disruption or impairment of adhesive metabolism pathways, respectively.

AChE activity was evaluated using Electrophorus electric AChE Type V-S (SIGMA C2888, E.C. 3.1.1.7), according to Ellman et al. (1961) [61] with some modifications [58,62]. Reaction solution containing 1 M phosphate buffer pH 7.2, 10 mM dithiobisnitrobenzoate (DTNB) (acid dithiobisnitrobenzoate and sodium hydrogen carbonate in phosphate buffer) and 0.075 M acetylcholine iodide was added to pure AChE enzyme (0.25 U/mL) and each test compound (final concentration of 25, 50 and 100 μM, 1% DMSO) in quadruplicate. All tests included a positive control with eserine (200 μM, water) and a negative control with 1% DMSO in water. The optical density was measured at 412 nm in a microplate reader (Biotek Synergy HT, Winooski, Vermont, USA) during 5 min at 25 ◦C.

Tyr inhibition assay was performed using *Agaricus bisporus* Tyr (EC1.14.18.1) according to Adhikari et al. (2008) [63] with some modifications [8]. The enzymatic reaction follows the catalytic conversion of L-Dopa to dopaquinone and the formation of dopachrome by measuring the absorbance at 475 nm. Briefly, Tyr (25 U/mL) was added to 50 mM phosphate buffer pH 6.5 and the tested compounds at 25, 50 and 100 μM (final concentrations, 1% DMSO). The enzymatic activity was triggered by the addition of L-dopa (25 mM). Kojic acid (1.4 mM, water) was included as positive control and 1% DMSO in water as negative control.

#### *3.7. Environmental Fate Parameters: Artemia Salina Ecotoxicity Bioassay*

The brine shrimp (*Artemia salina*) nauplii lethality test was used to determine the toxicity of promising AF compounds to non-target organisms [64]. *Artemia salina* eggs were allowed to hatch in seawater for 48 h at 25 ◦C. Bioassays were performed in 96-wells microplates with 15–20 nauplii per well and 200 μL of the compounds test solution. Compounds were tested at final concentrations of 25 and 50 μM (filtered seawater with 1% DMSO). All tests included K2Cr2O7 (13.6 μM) as positive control and DMSO (1%) as negative control. Bioassays were run in the dark at 25 ◦C, and the percentage of mortality was determined after 48 h of exposure.

#### *3.8. Statistical Analysis*

Datasets from anti-settlement, antibacterial and anti-microalgal bioassay, and determination of AChE and Tyr activities, were analysed by one-way analysis of variance (ANOVA) followed by a multi-comparisons Dunnett's test against negative control (*p* < 0.05). For the AF bioassays, the half maximum response concentration (EC50) values for each compound, when applicable, were calculated using Probit regression analysis. Significance was considered at *p* < 0.01, and 95% lower and upper confidence limits (95%LCL; UCL). The software IBM SPSS Statistics 26 (Armonk, New York, USA) was used for statistical analysis.

#### **4. Conclusions**

In this study, eight new triazole-flavonoid hybrids were synthesized using the click chemistry approach. From the series of synthesized compounds, flavone **1b** and chalcones **7b**, **11a** and **11b** showed significant anti-settlement activity towards the macrofouling species *Mytilus galloprovincialis* adhesive larvae. Regarding the compounds' structures, HACA1 was the most significant descriptor for the obtained QSAR model, contributing positively to the AF activity. Particularly, triazolyl glycosylated chalcone **7b**, with a high number of hydrogen-bonding acceptors, showed the most effective anti-settlement activity (EC50 = 3.28 <sup>μ</sup>M; 2.43 <sup>μ</sup>g·mL−1) with the highest therapeutic ratio (LC50/EC50 > 60.98), exhibiting also a significant inhibitory activity against the marine diatom *Navicula* sp. (EC50 = 41.76 <sup>μ</sup>M; 30.94 <sup>μ</sup>g·mL−1), suggesting potential in the suppression of biofouling colonization succession. Flavone **1b**, which was effective against the settlement of mussel larvae, also showed capacity to inhibit the activity of Tyr, which might explain the specific AF activity against mussel larvae. Ecotoxicity studies on the non-target species *Artemia salina* revealed that the flavonoids **1b**, **7b**, **11a** and **11b** did not show ecotoxicity to the nauplii of this sensitive crustacean, even at 50 μM, a concentration much higher than their EC50. These results disclosed synthetic flavonoids, particularly a new chalcone incorporating a 1,2,3- triazole ring (**7b**), with potential to be a good environmentally compatible alternative to the majority of the antifoulants in use. Flavonoids are ubiquitous in Nature and, therefore, they come with the advantage that they have been selected during evolution to have high specificity, high efficiency and some might be potential nontoxic inhibitors of fouling. Natural compounds are usually biodegradable, not leaving residue in the environment, and are thus considered one of the most promising alternatives to the biocides in use. However, the yields of natural compounds from marine organisms are generally poor, hindering their development as AF agents. Moreover, optimizing a micro-organism for enhanced production of antifoulant is generally laborious and time consuming. Synthesis of nature-like antifoulants seems to be a more sustainable way to create an opportunity to produce commercial supplies for the antifouling industry.

**Author Contributions:** Conceptualization, H.C.; methodology, J.R.A.; M.C.-d.-S.; and H.C.; compounds synthesis, D.P.; B.T.M.; compounds structure elucidation, D.P.; quantitative structureactivity relationship studies, A.P.; in vivo AF bioassays, enzymatic determinations and ecotoxicological bioassays, C.G.; J.R.A.; resources, M.P.; V.V.; writing—original draft preparation, D.P.; C.G.; writing—review and editing, D.P.; C.G.; J.R.A.; M.C.-d.-S.; H.C.; M.P.; V.V.; supervision, J.R.A.; M.C.-d.-S.; H.C.; project administration, J.R.A. and M.C.-d.-S.; funding acquisition, V.V., J.R.A. and M.C.-d.-S. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was supported by national funds through the FCT—Foundation for Science and Technology within the scope of UIDB/04423/2020 and UIDP/04423/2020 and under the projects: NASCEM-PTDC/BTA-BTA/31422/2017 (POCI-01-0145-FEDER-031422), and PTDC/AAG-TEC/0739/2014 (POCI-01-0145-FEDER-016793; Project 9471-RIDTI) co-financed by FCT, PORTUGAL2020 COMPETE 2020, through the European Regional Development Fund (ERDF); and by the structured program of R&D&I ATLANTIDA (reference NORTE-01-0145-FEDER-000040), supported by the North Portugal Regional Operational Programme (NORTE2020), through the ERDF. D. P. and C. G. acknowledge the grants they received (SFRH/BD/147207/2019 and NASCEM/BI-Lic/2019-53, respectively).

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Data is contained within the article.

**Acknowledgments:** The authors thank Sara Cravo for all the technical and scientific support.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


## *Article* **Natural Cyanobacterial Polymer-Based Coating as a Preventive Strategy to Avoid Catheter-Associated Urinary Tract Infections**

**Bruna Costa 1,2, Rita Mota 1,3, Paula Tamagnini 1,3,4, M. Cristina L. Martins 1,2,5 and Fabíola Costa 1,2,\***


Received: 17 April 2020; Accepted: 24 May 2020; Published: 26 May 2020

**Abstract:** Catheter-associated urinary tract infections (CAUTIs) represent about 40% of all healthcare-associated infections. Herein, the authors report the further development of an infection preventive anti-adhesive coating (CyanoCoating) meant for urinary catheters, and based on a natural polymer released by a marine cyanobacterium. CyanoCoating performance was assessed against relevant CAUTI etiological agents, namely *Escherichia coli*, *Proteus mirabilis*, *Klebsiella pneumoniae,* methicillin resistant *Staphylococcus aureus* (MRSA), and *Candida albicans* in the presence of culture medium or artificial urine, and under biofilm promoting settings. CyanoCoating displayed a broad anti-adhesive efficiency against all the uropathogens tested (68–95%), even in the presence of artificial urine (58–100%) with exception of *P. mirabilis* in the latter condition. Under biofilm-promoting settings, CyanoCoating reduced biofilm formation by *E. coli*, *P. mirabilis,* and *C. albicans* (30–60%). In addition, CyanoCoating prevented large crystals encrustation, and its sterilization with ethylene oxide did not impact the coating stability. Therefore, CyanoCoating constitutes a step forward for the implementation of antibiotic-free alternative strategies to fight CAUTIs.

**Keywords:** cyanobacteria; uropathogens; anti-adhesive coating; urinary catheters; surface modification; catheter-associated urinary tract infections

#### **1. Introduction**

Urinary catheters are the most common indwelling device, with 15–25% of hospitalized patients undergoing catheterization [1]. More than 30 million urinary catheters are used per year to manage urinary incontinence and urinary retention, during and/or after surgical practices in the USA only [2]. Infection is the main concern associated with the use of catheters (either long- or short-term). Catheter-associated urinary tract infections (CAUTIs) account for approximately 40% of all healthcare-associated infections; therefore, are associated to major economic burden (\$1000 per treatment of CAUTI in USA) [3]. This problem is rising together with bacterial antibiotic resistance, which is considered by the World Health Organization (WHO) as one of the most severe health threats around the world [4]. CAUTI establishment is related with the impairment of the natural defense systems of the healthy urological mucosa. When the use of a catheter is required, the natural flush of bacteria by micturition is hampered [5]. Moreover, damage to the inner walls of the urinary system breaches the natural protection against bacterial adhesion, which, adding to the presence of a foreign material and a compromised immune system, contributes to the establishment of CAUTIs. CAUTIs arise from cross contamination derived from the patient's normal fecal flora or from the healthcare personnel handling [6]. These infections are always associated with the occurrence of microbial biofilms, being the most prevalent Gram-negative bacteria, such as *Escherichia coli*, *Klebsiella pneumoniae*, *Proteus mirabilis,* and *Pseudomonas aeruginosa*, Gram-positive *Staphylococcus aureus*(including methicillin-resistant strains), and yeasts—particularly *Candida* species, etiological agents that are particularly well adapted to the urinary tract microenvironment [7].

CAUTIs are a major cause of catheter encrustation, which is promoted by urease-positive pathogens, such as *P. mirabilis*, *P. aeruginosa,* and *K. pneumoniae* [8]. Urease catalyzes the hydrolysis of urea into ammonia and carbamate, which in turn increases the urine pH promoting the formation of crystals [9]. The formation of biofilm itself may also promote catheter occlusion by the large amount of mucoid material produced (e.g., by *P. aeruginosa, K. pneumoniae*) or by the emergence of hyphae (e.g., *C. albicans*). Other CAUTI associated complications include bladder stones, septicemia, endotoxic shock, and pyelonephritis contributing to patients' suffering, and frequently worsening other concomitant chronic pathologies [10]. In this way, new strategies are needed to optimize patient safety, control costs, and to reduce bacterial resistance. The current materials used to produce catheters include polyurethanes (PUs), silicone, polytetrafluoroethylene (PTFE), polyvinylchloride (PVC), and latex rubber [8]. PUs are among the best choices for biomedical applications due to their mechanical properties, namely durability, elasticity, fatigue resistance, and compliance [8]. The advantage of using PUs instead of silicone for urinary catheters is that PUs originate catheters with larger internal diameters (due to thinner walls) that are less prone to occlusion, and soften within the patient's body, becoming more comfortable [8,11].

The most promising approach to improve urinary catheter safety is to alter its surface to avoid biofilm formation preventing the consequent infection [12–15]. For the development of anti-adhesive surfaces, natural polymers, such as hyaluronic acid and heparin, can be used [15–17]. Polysaccharides from marine sources, such as alginate, ulvan, agarose, and carrageenans have also been reported as possible alternatives [16,18,19]. Previously, Costa et al. [20] developed CyanoCoating, a coating based on a well-characterized extracellular polymer produced by a marine cyanobacterium [21]. These authors demonstrated that CyanoCoating has anti-adhesive properties against *S. aureus*, *S. epidermidis*, *P. aeruginosa*, and *E. coli*, and is biocompatible, having the potential to be applied to a wide range of medical devices, including blood contacting materials [20].

The present study is aimed at evaluating CyanoCoating capability to endure urinary catheter specifications (urine, uropathogens, and sterilization). Moreover, the absence of contaminants in the raw biological material was confirmed. Overall, the results obtained highlight the translational potential of CyanoCoating to mitigate challenges imposed by CAUTIs.

#### **2. Results**

#### *2.1. Biopolymer Regulatory Compliance Assessment: Metal and Microbial Contamination*

The extracellular cyanobacterial polymer, mainly of heteropolysaccharidic nature, used to prepare the CyanoCoating is a new material not yet described on pharmacopeia, thus, metal and microbial contamination was addressed. The Inductively Coupled Plasma–Atomic Emission Spectrometry (ICP–AES) results showed that the isolated biopolymer was not contaminated with arsenic (As), cadmium (Cd), lead (Pb), or mercury (Hg) (Supplementary Table S1). Moreover, the microbiological

assays showed that the biopolymer was not contaminated with bacteria or fungi, even before the autoclave sterilization process, as no colony-forming units (CFUs) where observed up to 5 days.

#### *2.2. CyanoCoating Surface Characterization*

CyanoCoating was previously characterized in terms of thickness and wettability [20]. However, since surface topography is known to impact biofilm development, this parameter was evaluated by atomic force microscopy (AFM). CyanoCoating and medical grade polyurethane (PU) were covalently bound through a polydopamine (pDA) layer to gold (Au) substrates, as previously described [20]. The deposition of either the pDA + CyanoCoating or the control pDA + PU increased significantly surface roughness of Au substrates, as depicted in Figure 1A. CyanoCoating exhibited a smoother surface in comparison with PU, as demonstrated by the decrease of the average roughness (Ra) (Figure 1A1) and the root mean square roughness (Rq) (Figure 1A2). Representative AFM three-dimensional (3D) images of the threes surfaces can be observed in Figure 1B.

**Figure 1.** Characterization, by atomic force microscopy (AFM), of the surface roughness of gold substrates (Au), Au substrates coated with a polydopamine (pDA) layer plus polyurethane (PU), or Au substrates coated with a pDA layer plus CyanoCoating. (**A1**) Average roughness (Ra), (**A2**) Root mean square roughness (Rq) and (**B**) AFM three-dimensional (3D) surface images. Statistical analysis was performed by non-parametric Kruskal–Wallis analysis and statistical differences are indicated with \* (*p* < 0.05) and \*\*\*\* (*p* < 0.001).

#### *2.3. CyanoCoating Biological Performance*

#### 2.3.1. Microbial Adhesion Assays

As the anti-adhesive performance of CyanoCoating was previously evaluated against *Escherichia coli* and *Pseudomonas aeruginosa* [20], herein, we focused on other relevant uropathogens for catheter-associated urinary tract infections (CAUTIs): *Proteus mirabilis*, *Klebsiella pneumoniae*, methicillin resistant *Staphylococcus aureus* (MRSA), and *Candida albicans* [22], according to ISO 22196:2007 [23]. Overall, the results obtained (Figure 2 and Figure S1) showed that microbial adhesion to CyanoCoating was significantly lower than the adhesion to medical grade PU, ranging from 68 ± 28% (*P. mirabilis*) to 95 ± 48% (*K. pneumoniae*). Importantly, CyanoCoating was efficient in preventing *S. aureus* (MRSA) adhesion (80 ± 27%), a microorganism that is very difficult to eradicate. Moreover, CyanoCoating could also reduce in 69 ± 19% the adhesion of the yeast *C. albicans* (responsible for 10–15% of CAUTIs [24]).

**Figure 2.** CyanoCoating anti-adhesive performance compared to medical grade polyurethane (PU). The coatings were tested against the uropathogens mentioned above each graph using the respective growth medium; see Materials and Methods. Data represent mean ± Standard deviation (n = 9). The assay was performed according to ISO 22196. Statistical analysis was performed by non-parametric Kruskal–Wallis analysis and statistical differences are indicated with \* (*p* < 0.05), \*\*\* (*p* < 0.005) and \*\*\*\* (*p* < 0.001).

#### 2.3.2. Microbial Adhesion Assays with Artificial Urine

The anti-adhesive performance of CyanoCoating was subsequently assessed with artificial urine medium (AUM) against *E. coli*, *P. mirabilis, K. pneumoniae, S. aureus*(MRSA) and *C. albicans*, also according to ISO 22196:2007. In the presence of AUM, CyanoCoating significantly reduced the adhesion of most of the uropathogens compared to PU. For the Gram-negative *E. coli* and *K. pneumoniae* a reduction of 65 ± 28% and 98 ± 54%, respectively, was observed, while for the Gram-positive *S. aureus* (MRSA) and the yeast *C. albicans*, a striking 95 ± 34% and 100% reduction, respectively, was observed (Figure 3 and Figure S2).

**Figure 3.** CyanoCoating anti-adhesive performance compared to medical grade polyurethane (PU) with artificial urine medium. The coating was tested against the uropathogens mentioned above each graph. Data represent mean ± Standard deviation (n = 9). The assay was performed according to ISO 22196. Statistical analysis was performed by non-parametric Kruskal–Wallis analysis and statistical differences are indicated with \* (*p* < 0.05) and \*\*\*\* (*p* < 0.001).

#### 2.3.3. Biofilm Formation

In order to evaluate CyanoCoating effectiveness in preventing biofilm formation, a biofilm assay was performed according to Costa et al., [25]. After 24 h, the number of CFUs detached from the

surfaces by sonication were determined. The efficiency of the sonication process was verified by observing the surfaces using inverted fluorescence microscopy. A reduction trend on biofilm formation was observed for *E. coli* (39 ± 10%), *P. mirabilis* (39 ± 15%) and *C. albicans* (60 ± 30%) on CyanoCoating samples compared to the control PU, while for *K. pneumoniae* and *S. aureus* MRSA no significant differences were observed (Figure 4).

**Figure 4.** Effect of CyanoCoating on the prevention of biofilm formation compared to medical grade polyurethane (PU), by measuring the bacteria detached from the surfaces. Data represent mean ± Standard deviation (n = 9). Statistical analysis was performed by Mann–Whitney test (*t*-test) analysis and statistical differences are indicated with \* (*p* < 0.05) and \*\* (*p* < 0.01).

#### *2.4. Encrustation Development*

Salts deposition on top of CyanoCoating, was evaluated by scanning electron microscopy (SEM) and energy-dispersive X-ray spectroscopy (EDS), after incubation with supplemented artificial urine medium (AUS). This urine is supplemented with urease and ovalbumine to promote an encrustation

environment [26,27]. SEM micrographs (Figure 5A, left panel) show the clean surfaces of the CyanoCoating and the control PU at the initial time points (before the immersion in AUS). Seven days after the immersion, it was possible to observe salt deposition on top of the samples (Figure 5A, right panel), particularly in the PU surface where agglomerates of larger crystals are clearly visible. The EDS spectra (Figure 5B) indicate the presence of elements that could suggest the formation of struvite (NH4MgPO4·6H2O), brushite (CaHPO4·2H2O), or hydroxyapatite (Ca5(PO4)3(OH)) in the surface of the samples immersed in AUS (Figure 5B, lower panel). Higher amounts of magnesium (Mg) and phosphorus (P) were found on PU samples suggesting accumulation of struvite, whereas the presence of calcium (Ca) and P on CyanoCoating suggests accumulation of brushite or hydroxyapatite. Before immersion in AUS, EDS spectra of PU and CyanoCoating samples only present silicon (Si), gold (Au) and carbon (C), the expected elements of the substrates, and palladium (Pd) from the SEM analysis sputtering.

**Figure 5.** Encrustation development on CyanoCoating compared to medical grade polyurethane (PU).

(**A**) SEM micrographs of the coatings before and after 7 days immersion in supplemented artificial urine medium (AUS). Magnification 30×. (**B**) Energy-dispersive X-ray spectroscopy (EDS) spectra of the selected areas on each coating surfaces before and after 7 days of immersion in AUS. **a** to **e** correspond to the areas highlighted in the SEM micrographs above.

#### *2.5. CyanoCoating Stability After Sterilization*

To evaluate the stability of CyanoCoating after sterilization by ethylene oxide (EO), the most common industrial sterilization technique for medical devices [28] and compatible with most of the biomaterials used in their manufacture, samples were characterized both physically (water contact angle measurements) and biologically (anti-adhesive performance). Our results revealed that EO sterilization did not significantly alter CyanoCoating wettability, compared to unsterilized samples, and samples submitted to the regular laboratorial ethanol-based disinfection protocol (Figure 6). Similarly, the anti-adhesive performance of CyanoCoating after EO sterilization was not altered, compared to samples submitted to the ethanol-based disinfection protocol, using *E. coli* or *P. mirabilis* as model bacteria (Figure 7).

**Figure 6.** Surface characterization, by water contact angle (captive bubble method), of CyanoCoating samples without sterilization, after sterilization with ethanol 70% (*v*/*v*) or with ethylene oxide (EO). Data represent mean ± Standard deviation (n = 9).

**Figure 7.** CyanoCoating anti-adhesive performance after sterilization with ethanol 70% (*v*/*v*) or ethylene oxide (EO). Data represent mean ± Standard deviation (n = 9). Adhesion of *Escherichia coli* and *Proteus mirabilis* in Tryptic Soy Broth (TSB). The assay was performed according to ISO 22196.

#### **3. Discussion**

Among all healthcare-associated infections, catheter-associated urinary tract infections (CAUTIs) are recognized as the most prevalent worldwide [29]. In this work, we explore the possibility of a previously developed anti-adhesive coating, CyanoCoating [20], to endure urinary catheters specifications.

Concerning the quality of the raw material (cyanobacterial extracellular polymer) used to produce CyanoCoating, the absence of fungi and bacteria indicate that all steps performed from the cell cultures to the polymer extraction ensured a high purity level of the product, fulfilling the quality requirements suggested by pharmacopeia, and the regulations imposed by healthcare authorities.

In the previous work, the broad-spectrum activity of CyanoCoating was assessed against relevant etiological agents responsible for medical devices-associated infections, including the uropathogens *Escherichia coli* and *Pseudomonas aeruginosa* (reducing bacterial adhesion by at least 80%) [20]. Here, the potential of CyanoCoating for CAUTIs mitigation was assessed against other relevant uropathogens, namely *Proteus mirabilis*, *Klebsiella pneumoniae*, methicillin-resistant *Staphylococcus aureus* (MRSA), and the yeast *Candida albicans* [30–32]. Overall, CyanoCoating greatly impaired the adhesion of the tested microorganisms (ranging from 68 ± 28% to 95 ± 48%). Considering that CyanoCoating is highly hydrophilic and exhibits a smoother topography compared to polyurethane (as visible in the AFM images) the hypothesis of an anti-adhesive mechanism of action is the most plausible. It is known that highly hydrophilic surfaces prevent the adsorption of proteins/cells due to the establishment of a hydration layer formed by well-structured water molecules linked to the surface by hydrogen bonds that works as a physical barrier [33]. In addition, the lack of bactericidal activity previously reported [20] reinforce our hypothesis. Similar results were obtained by other authors, using poly(ethylene glycol) (PEG) [34] or sulfobetaine methacrylate (SBMA) [35] anti-adhesive synthetic coatings onto PU or silicone surfaces, with *E. coli* and *S. epidermidis* or *P. aeruginosa* and *S. aureus* only. Our results demonstrate that CyanoCoating is effective against a broader range of microorganisms, including urease-positive bacteria and yeasts (this work and [20]).

To better mimic the in vivo environment that bacteria encounter in the urinary tract [32,36], artificial urine medium was used for the in vitro adhesion assays. The microorganisms were chosen since *E. coli* is the most prevalent in CAUTIs, *P. mirabilis* is responsible for the most severe cases, *C. albicans* causes 10–15% of these infections and the other bacteria are also relevant [22,37]. In the presence of artificial urine medium, the overall microbial adhesion to CyanoCoating and PU surfaces was significantly lower than with culture medium, in particular for the Gram-negative bacteria *E. coli*, *P. mirabilis,* and *K. pneumoniae*. This result can be associated to the media composition; culture media promote bacterial growth and biofilm formation mechanisms since they contain glucose as a carbon source, in contrast with the artificial urine medium. Nevertheless, CyanoCoating performance was much better than PU against all the microorganisms tested, in particular for *K. pneumoniae* and *C. albicans*. In addition, we demonstrated that the efficiency of CyanoCoating was not negatively affected by clinically relevant sterilization procedures such as ethylene oxide (EO).

The efficiency of CyanoCoating on preventing biofilm formation was assessed against all uropathogens mentioned above. This method counts the CFUs originated after detachment of the biofilm by sonication instead of other indirect methods commonly used, e.g. the resazurin assay that assess the metabolic activity of bacteria in biofilms [38] or the canonical crystal violet assay that stains the extracellular matrix [38]. This last method cannot be used here due to the heteropolysaccharidic nature of the polymer used to generate the CyanoCoating [21]. Biofilm formation was significantly impaired for *E. coli*, *P. mirabilis,* and *C. albicans* ranging from 39 ± 10% to 60 ± 30%, suggesting that CyanoCoating hinders biofilm formation against a broad-spectrum of microorganisms, even for the difficult to eradicate fungi *C. albicans* [39]. Our results reinforce the strategy of using natural polymers to prevent biofilm formation, as reported by others, e.g., the use of carboxymethyl chitosan to coat medical grade silicone and that reduced biofilm formation by Gram-negative bacteria [37], or the low-molecular weight chitosan hydrogels used to coat polystyrene microplates that avoid biofilm formation by *Candida* spp. [39]. Current technologies in the market are based on the release of antimicrobial agents by the coating, such as antiseptics or antibiotics, to inhibit the colonization of the catheters. However, in spite of the broad-spectrum activity, these coatings exhaust their antimicrobial activities over long periods, are associated with toxicity and contribute for the development of antimicrobial resistance [2,40]. Having in mind the goal of developing an antibiotic-free coating, CyanoCoating may be combined with bactericidal compounds, such as antimicrobial peptides, that can be either immobilized or delivered [41,42].

Another critical aspect on indwelling urinary catheters is the mineral deposition on their surfaces. Frequently urinary catheters become blocked by hard mineral deposits, resulting in urine leakage, discomfort to the patient, and even catheter encrustation. In the worst-case scenario, the encrustation can only be solved by removing the catheter, which may cause trauma to the urethra [26,43]. The encrustation is exacerbated by the presence of urease positive pathogens, such as *P. mirabilis* [26]. Therefore, we challenge CyanoCoating with artificial urine medium supplemented with urease, which also contains albumin that mimics the bacterial and cellular debris that infected urine frequently contains [26]. The energy-dispersive X-ray spectroscopy (EDS) results clearly indicated the presence of Ca, P, Mg, and O that could suggest struvite (NH4MgPO4·6H2O), brushite (CaHPO4·2H2O) or hydroxyapatite (Ca5(PO4)3(OH)) formation. However, while on the control PU surface big rectangular shaped crystals protruded from the surface suggesting the formation of struvite [44], on the CyanoCoating individual crystallites with powdery appearance and smaller in size were formed, which is consistent with brushite or hydroxyapatite [45,46]. All together, these results show that CyanoCoating is less prone to encrustation, and therefore less prone to promote catheter blockage.

#### **4. Materials and Methods**

#### *4.1. Cyanobacterium Growth Conditions and Biopolymer Isolation*

The unicellular cyanobacterium *Crocosphaera chwakensis* CCY0110 [47] (previously identified as *Cyanothece* sp. CCY 0110; Culture Collection of Yerseke, The Netherlands; kindly provided by Lucas Stal) was grown in 2 L bioreactors with ASNIII medium, at conditions previously described [20,21]. Cells were grown until an optical density at 730 nm of approximately 2.5–3.5 and the extracellular biopolymer was isolated as previously described [20].

#### *4.2. Biopolymer Contaminants*

#### 4.2.1. Assessment of Metal Contaminants

To assess the putative contamination of the cyanobacterial polymer with heavy metals, the presence of arsenic (As), cadmium (Cd), lead (Pb), and mercury (Hg) was evaluated. For this purpose, aqueous polymer solutions 0.5% (*w*/*v*) were prepared and mineralized using 5% HNO3 (*v*/*v*). Then, an Inductively Coupled Plasma–Atomic Emission Spectrometer (ICP–AES) (Ultima, Jobin Yvon), equipped with a 40.68 MHz RF generator and a Czerny-Turner monochromator with 1.00 m) was used for metals quantification.

#### 4.2.2. Polymer Microbiological Control

To assess the microbiological quality of the raw material, polymer bioburden (contamination with bacteria or fungi) was evaluated by microbiological assays as recommended by Portuguese Pharmacopeia [48]. To perform the assays, 10 mL of polymer solution 1% (*w*/*v*) were filtered by a 0.45 μm filter (Merck). Then, the filter was cut into halves and each part was placed on top of either Tryptic Soy Agar (TSA) plates or Sabouraud Dextrose Agar (SDA). After 24 h incubation period at 37 ◦C, the number of colonies-forming units (CFUs) were counted. Two replicates of each condition were performed.

#### *4.3. CyanoCoating Development*

CyanoCoating was prepared as previously reported by [20]. Briefly, gold substrates (Au) were cleaned for 5 min, with "piranha" solution (7 parts of sulfuric acid (95%, *v*/*v* (BDH Prolabo): 3 parts of hydrogen peroxide (Merck), (CAUTION: this solution reacts violently with organic solvents and should be handled with care). Then, substrates were immersed in freshly prepared dopamine solution (2-(3,4-Dihydroxyphenyl)ethylamine hydrochloride, Sigma-Aldrich) (2 mg/mL in 10 mM TrisHCl pH 8.5) to allow formation of a polydopamine (pDA) layer on top of the Au substrate [20]. Subsequently, the polymer solution at 0.5% (*w*/*v*) was spin-coated (model WS-650-23, Laurell Technologies Corporation, North Wales, PA, USA) at 9000 rpm for 1 min on top of pDA-coated Au substrate. As control samples, medical grade polyurethane (PU) surfaces were prepared by similarly spin-coating the PU (Pellethane 2363 80 AE; Velox) solution at 0.1% (*w*/*v*) in tetrahydrofuran (Merck), on top of pDA-coated Au substrate [20].

#### *4.4. CyanoCoating Surface Characterization by Atomic Force Microscopy (AFM)*

Atomic Force Microscopy (AFM) images were obtained using a PicoPlus 5500 controller (Keysight Technologies, Santa Rosa, CA, USA). The images of gold substrate were performed in Tapping Mode, in air using a bar-shaped cantilever with a spring constant (k) in the range of 1–5 N/m (AppNano, Mountain View, CA, USA). The images on polyurethane (PU) and CyanoCoating were obtained in Contact Mode, in air, using a triangular shape cantilever V-shaped cantilever with a spring constant k = 0.085 N/m (Hydra-All-G, AppNano, Mountain View, CA, USA). The scan speed was set at 1.0 l/s, for both AFM modes. The scan size was 5 <sup>×</sup> 5 <sup>μ</sup>m2. The software used to obtain the images was the PicoView 1.2 (Keysight Technologies, Santa Rosa, CA, USA). The WSxM5.0 software (Nanotec Electronica, Feldkirchen, Germany) was used to perform the roughness surface measurements [49].

#### *4.5. CyanoCoating Biological Performance Evaluation*

#### 4.5.1. Microbial Strains, Media, and Growth Conditions

*P. mirabilis* (clinical isolate provided by Faculdade de Medicina Dentária, Universidade do Porto) was grown on cystine-lactose-lectrolyte-deficient agar (CLED agar) (Merck) and tryptic soya broth (TSB) (Merck). *E. coli* (ATCC 25922) and *S. aureus* MRSA (ATCC 33591), obtained from the American Type Culture Collection (ATCC), were grown on tryptic soya agar (TSA) (Merck) and TSB (Merck). *K. pneumoniae* (clinical isolate provided by Centro Hospitalar do Porto) was grown on TSA and Todd Hewitt Broth (THB). *C. albicans* (DSM 1386), obtained from the German Collection of Microorganisms and Cell Cultures GmbH (DSM), was grown on Sabouraud Dextrose Agar (SDA) (Merck) and Sabouraud Dextrose Broth (SDB) (Merck). The initial microbial inoculum was adjusted in TSB for *E. coli* and *S. aureus*, in THB for *K. pneumoniae*, or in SDB for *C. albicans*, according to OD600nm measurement and subsequently confirmed by count of CFUs.

#### 4.5.2. Microbial Adhesion Assays

Microbial adhesion assays were performed using *P. mirabilis*, *K. pneumoniae*, *S. aureus* MRSA and *C. albicans* according to ISO 22196:2007 (Plastics—Measurement of antibacterial activity on plastics surfaces) [23]. For CyanoCoating and PU disinfection, samples were immersed subsequently for 15 min, twice in ethanol 70% (Merck) and twice in filtered type II water (0.22 μm syringe filter), being dried with argon stream in a flow hood, and then transferred to a 24-well plate. Then, a 5 μL inoculum drop (1.8 <sup>×</sup> 10<sup>6</sup> CFUs/mL) was placed on top of the samples and then covered with a previously sterilized polypropylene (PP) coverslip (Ø 9 mm), using the method described above. Samples were incubated for 24 h at 37 ◦C in moisturized condition. After 24 h, samples were rinsed with Phosphate Buffered Saline (PBS) three times. Adhered bacteria or fungi were fixed with paraformaldehyde 4% (*v*/*v*) in PBS, for 30 min at room temperature (RT). After rinsing with PBS three times, samples were stained with 4 ,6-diamidino-2-phenylindole (DAPI) (0.1 μg/mL) for 30 min at RT, protected from light. Afterwards, samples were rinsed with PBS and transferred to an uncoated 24-well μ-plate (#82406, IBIDI, Gräfelfing, Germany) with the surface facing the bottom. Results represent average of three independent assays, with three replicates per sample.

High-content screening microscope (IN Cell Analyzer 2000, GE Healthcare, Chicago, IL, USA) with a Nikon 20× / 0.95 NA Plan Apo objective (binning 1 × 1), using a charge-coupled device (CCD) Camera (CoolSNAP K4) was used to observe samples from microbial adhesion assays. Image field of view (FOV) x-y for this objective is 0.8 × 0.8 cm. Moreover, 9 FOV per sample were acquired spanning an area of 5.76 cm2. The excitation and emission filters used were DAPI (excitation: 365 nm; emission: 420 nm). On-the-fly deconvolution was performed. The number of adherent bacteria were quantified using the ImageJ software, and values were converted to bacteria per mm2.

Adhesion reduction percentages were calculated according to the formula: [number of adhered bacteria per mm<sup>2</sup> on CyanoCoating <sup>×</sup> 100]/[number of adhered bacteria per mm<sup>2</sup> on PU]. The standard deviations were calculated considering error propagation of the measurements uncertainties.

#### 4.5.3. Antimicrobial Adhesion Assays in the Presence of Artificial Urine Medium

To better simulate the conditions of microbial adhesion inside urinary tract, the anti-adhesive performance of CyanoCoating against *E. coli, P. mirabilis, K. pneumoniae, S. aureus MRSA,* and *C. albicans* was performed as explained previously (see Section 4.5.2.), but using artificial urine medium prepared according to Brooks et al. [32] (composition: Supplementary Table S2) to adjust initial inoculum. After 24 h incubation period, samples were processed, as described in Section 4.5.2., the number of adherent bacteria were quantified using the ImageJ software, and values were converted to bacteria per mm2. Results represent average of three independent assays, with three replicates per sample. The adhesion reduction percentages and respective standard deviations were calculated, as described in Section 4.5.2.

#### 4.5.4. Biofilm Formation Assessment

*E. coli, P. mirabilis, K. pneumoniae, S. aureus* MRSA, and *C. albicans* were grown overnight in respective culture media, described in Section 4.5.1. PU and CyanoCoating samples were disinfected, as described in Section 4.5.2., being then dried with argon stream in a flow hood and transferred to a 24-well tissue culture polystyrene plates (TCPS, Sarstedt, Nümbrecht, Germany). Then, 100 μL of inoculum (1.0 <sup>×</sup> 107 CFUs/mL) were added to each well containing samples pre-hydrated in 900 <sup>μ</sup>L of TSB for 30 min. After a 2 h incubation period at 37 ◦C, surfaces were rinsed three times with sterile PBS and re-incubated with 1000 μL of TSB during 24 h. After incubation, samples were rinsed five times with PBS to remove planktonic and loosely bound bacteria. Then, surfaces were transferred to 5 mL SARSTEDT tubes containing 1 mL of 0.5% Tween 80 in PBS and placed on ice, then sonicated using BactoSonicR (BANDELIN, Heinrichstraße, Berlin, Germany) at 160 W for 15 min, placed on ice for 5 min, sonicated again for 15 min and put on ice. As a control, the adjusted inoculum was submitted to the same sonication protocol to verify if the sonication applied interferes with microorganism viability. After, serial dilutions were done and plated for CFU counting. Results are the average of three replicates of three independent assays.

To ensure that after sonication all bacteria were removed from the surfaces, PU and CyanoCoating samples were transferred to a 24-well plate and fixed with paraformaldehyde 4% (*v*/*v*) in PBS, for 30 min at RT. After rinsing with PBS three times, samples were stained with 4 ,6-diamidino-2-phenylindole (DAPI) (0.1 μg/mL) for 30 min at RT, protected from light. Afterwards, samples were rinsed with PBS and transferred to an uncoated 24-well μ-plate (#82406, IBIDI) with the surface facing the bottom. The image acquisition and analysis were performed, described in Section 4.5.2. The adhesion reduction percentages and respective standard deviations were calculated, as described in Section 4.5.2.

#### *4.6. Encrustation Assay*

The evaluation of the deposition of crystals on the surface of samples was performed using supplemented artificial urine medium, prepared as described by Cox and collaborators [26] (composition: Supplementary Table S3). Samples were immersed in 2 mL of AUS and incubated at 37 ◦C, 60 rpm for 7 days. These experiments were executed in triplicate. After 7 days, the samples were washed gently using distilled water to remove any salts that may be loosely deposited on the surface of the materials. Then, samples were dried in vacuum oven (Trade Raypa, Barcelona, Spain) overnight. The samples conductivity was enhanced by sputtering with Au/Pd for 60 s and 15 mA current using the SPI Module Sputter Coater equipment (Structure Probe, Inc., West Chester, PA, USA). The SEM / EDS analysis was performed using a High resolution (Schottky) Environmental Scanning Electron Microscope with X-Ray Microanalysis and Electron Backscattered Diffraction analysis (JEOL JSM 6301F / Oxford INCA Energy 350, Jeol, Peabody, MA, USA). Micrographs of the surfaces were taken using an electron beam intensity of 5 kV (accelerating voltage) and a magnification of 30×, at CEMUP (University of Porto, Porto, Portugal).

#### *4.7. Assessment of CyanoCoating Stability after Ethylene Oxide Sterilization*

#### 4.7.1. Water Contact Angle (WCA)

To assess the performance of CyanoCoating after clinically relevant sterilization procedure, samples were submitted to ethylene oxide (EO) sterilization (kindly performed at sterilization service of Hospital de São João, Porto, Portugal) and compared to samples disinfected with the protocol described in Section 4.5.2. (control samples). The ethylene oxide sterilization was performed using a sterilizer cabinet EOGas series 3 plus with ampoules system (Andersen Products, Essex, UK) during 16 h (4 h of sterilization plus 12 h of aeration) at 50 ◦C.

Water contact angle measurements were performed using captive bubble method with a goniometer model OCA 15, equipped with a video CCD-camera and SCA 20 software (Data Physics, Filderstadt, Germany). Samples were tape glued to a microscope slide and placed with the surface facing the bottom in a quartz chamber filled with type I water. Subsequently, 10 μL bubbles of room air were introduced using a J-shaped syringe at a dose rate of 2 μL/s. Bubble profiles were fitted using tangent formula, to obtain the contact angle. Results are the average of two measurements of three replicates of three independent assays.

#### 4.7.2. Microbial Assays

In order to understand if EO sterilization process compromises bacterial adhesion in CyanoCoating surface, anti-adhesive assays performance was also evaluated, as described in Section 4.5 using *P. mirabilis* and *E. coli*.

#### *4.8. Statistical Analysis*

Statistical analysis was performed using Mann–Whitney test (*t*-test) and non-parametric Kruskal–Wallis test using the GraphPad Prism program version 6 (GraphPad Software, San Diego, CA, USA). Data is expressed as the mean ± standard deviation (SD) and p values of < 0.05 were considered significant.

#### **5. Conclusions**

Cyanobacteria are a prolific source of extracellular polymeric substances with particular characteristics that represent an untapped source of natural polymers for industrial applications, namely biomedicine. The evaluation of the cyanobacterial polymer-based CyanoCoating demonstrated that this coating is highly efficient in preventing the adhesion of most relevant uropathogens tested here, both in the presence of culture medium or artificial urine, when compared to medical grade PU. Moreover, a significant biofilm formation reduction was observed for three of these uropathogens, namely *E. coli*, *P. mirabilis,* and *C. albicans*. In addition, CyanoCoating is also promising on encrustation mitigation, and is rather stable after being subjected to an industrial sterilization technique (ethylene oxide). In the post-antibiotic era, strategies similar to the one reported here will play an important role as effective and non-cytotoxic solutions in the battle against CAUTIs.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/1660-3397/18/6/279/s1, Table S1: Metal contaminants assessment by Inductively Coupled Plasma–Atomic Emission Spectroscopy (ICP–AES). Table S2: Composition of the artificial urine medium according to Brooks et al., 1997 [32]. Table S3: Composition of the supplemented artificial urine medium according to Cox et al., 1987 [26]. Figure S1: Micrographs of *Proteus mirabilis*, *Klebsiella pneumoniae*, *Staphylococcus aureus* MRSA and *Candida albicans* cells adhered to polyurethane (PU) and CyanoCoating after 24 h incubation at 37 ◦C and stained with Draq5 and propidium iodide (PI) (scale bars—60 μm). Figure S2: Micrographs of *Escherichia coli*, *Proteus mirabilis*, *Klebsiella pneumoniae*, *Staphylococcus aureus* MRSA and *Candida albicans* cells adhered to polyurethane (PU) and CyanoCoating after 24 h incubation at 37 ◦C, in the presence of artificial urine medium (AUM), and stained with Draq5 and propidium iodide (PI) (scale bars—60 μm).

**Author Contributions:** Conceptualization, R.M., and F.C.; Data curation, B.C., R.M., and F.C.; Funding acquisition, R.M., P.T., M.C.L.M., and F.C.; Investigation, B.C., R.M., and F.C.; Methodology, B.C., R.M., and F.C.; Validation, R.M., P.T., M.C.L.M., and F.C.; Writing—original draft, B.C. and F.C.; Writing—review and editing, B.C., R.M., P.T., M.C.L.M., and F.C. All authors have read and agreed to the published version of the manuscript.

**Funding:** This project has received funding from "la Caixa" Foundation (ID 100010434), under the agreement LCF/TR/CI18/50030020". Additionally, this work was financed by FEDER - Fundo Europeu de Desenvolvimento Regional funds through the COMPETE 2020 - Operacional Programme for Competitiveness and Internationalisation (POCI), Portugal 2020, and by Portuguese funds through FCT - Fundação para a Ciência e a Tecnologia/Ministério da Ciência, Tecnologia e Ensino Superior in the framework of the project POCI-01-0145-FEDER-028779 (PTDC/BIA-MIC/28779/2017) and in the framework of the project "Institute for Research and Innovation in Health Sciences" (UID/BIM/04293/2020). Fabíola Costa also thanks FCT/MCTES for her contract under Stimulus of Scientific Employment 2017 (CEECIND/01921/2017/CP1392/CT0002). The authors acknowledge the support of the i3S Scientific Platform BioSciences Screening (BS), member of the national infrastructure PPBI - Portuguese Platform of Bioimaging (PPBI-POCI-01-0145-FEDER- 022122) and i3S Scientific Platform Biointerfaces and Nanotechnology (BN).

**Acknowledgments:** The authors acknowledge the support of the Biosciences Screening (André Maia) & Biointerfaces and Nanotechnology (Manuela Brás) i3S Scientific Platforms. The authors would like to thank to sterilization unit of Centro Hospitalar de São João, Porto, Portugal, to Daniela Silva for technical assistance with SEM/EDS from Centro de Materiais da Universidade do Porto (CEMUP), Porto, Portugal, and to Laboratório de Análises - REQUIMTE, Faculdade de Ciências e Tecnologia da Universidade Nova de Lisboa, Lisboa, Portugal.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


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## *Article* **The Effect of Molecular Weight on the Antimicrobial Activity of Chitosan from** *Loligo opalescens* **for Food Packaging Applications**

**Luciana C. Gomes 1, Sara I. Faria 1, Jesus Valcarcel 2, José A. Vázquez 2, Miguel A. Cerqueira 3, Lorenzo Pastrana 3, Ana I. Bourbon <sup>3</sup> and Filipe J. Mergulhão 1,\***


**Abstract:** The growing requirement for sustainable processes has boosted the development of biodegradable plastic-based materials incorporating bioactive compounds obtained from waste, adding value to these products. Chitosan (Ch) is a biopolymer that can be obtained by deacetylation of chitin (found abundantly in waste from the fishery industry) and has valuable properties such as biocompatibility, biodegradability, antimicrobial activity, and easy film-forming ability. This study aimed to produce and characterize poly(lactic acid) (PLA) surfaces coated with β-chitosan and β-chitooligosaccharides from a *Loligo opalescens* pen with different molecular weights for application in the food industry. The PLA films with native and depolymerized Ch were functionalized through plasma oxygen treatment followed by dip-coating, and their physicochemical properties were assessed by Fourier-transform infrared spectroscopy, X-ray diffraction, water contact angle, and scanning electron microscopy. Their antimicrobial properties were assessed against *Escherichia coli* and *Pseudomonas putida*, where Ch-based surfaces reduced the number of biofilm viable, viable but nonculturable, and culturable cells by up to 73%, 74%, and 87%, respectively, compared to PLA. Biofilm growth inhibition was confirmed by confocal laser scanning microscopy. Results suggest that Ch films of higher molecular weight had higher antibiofilm activity under the food storage conditions mimicked in this work, contributing simultaneously to the reuse of marine waste.

**Keywords:** chitin; chitosan; marine waste; antimicrobial activity; poly(lactic acid); active packaging

#### **1. Introduction**

One of the major growth segments in the food industry is minimally processed, preservative-free, and ready-to-eat meals and food products [1,2]. As a consequence, the waste from traditional plastic packaging is also increasing (around 4.2% per year) [3] and is considered one of the main factors responsible for short- and long-term environmental pollution [3–5]. Another problem facing the food industry is microbial contamination since microorganisms can attach to food and packaging surfaces and form biofilms in a complex and multifaceted process, causing food spoilage, illness, and shelf-life reduction of food products [1,6]. Biofilms are organized communities of microorganisms that attach to a surface and produce extracellular polymeric substances (EPS), which protect them from adverse environmental conditions [6]. These two factors are putting increasing pressure on the food industry to develop new types of antimicrobial packaging materials,

**Citation:** Gomes, L.C.; Faria, S.I.; Valcarcel, J.; Vázquez, J.A.; Cerqueira, M.A.; Pastrana, L.; Bourbon, A.I.; Mergulhão, F.J. The Effect of Molecular Weight on the Antimicrobial Activity of Chitosan from *Loligo opalescens* for Food Packaging Applications. *Mar. Drugs* **2021**, *19*, 384. https://doi.org/ 10.3390/md19070384

Academic Editors: Tom Turk and Joana Reis Almeida

Received: 31 May 2021 Accepted: 28 June 2021 Published: 2 July 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

mainly based on natural and renewable sources, in order to ensure food safety, quality maintenance, and shelf-life enhancement, as well as to reduce environmental issues caused by non-biodegradable packaging materials [7–9].

Among different biopolymers, chitosan (Ch) has received substantial attention from academics and industry for food packaging applications due to its particular physicochemical features, biodegradability, non-toxicity, biocompatibility, good film-forming properties, chemical stability, high reactivity, low cost, and availability in nature [10–12]. Chitosan also has intrinsic antioxidant and antimicrobial activities against fungi, molds, yeasts, and Gram-negative and Gram-positive bacteria [13]. As a result of these properties, chitosan was classified as Generally Recognized as Safe by the US FDA in 2001 [14]. However, inherent drawbacks of Ch, including low mechanical and thermal stability and high sensitivity to humidity, have been restricting its industrial application [15]. One strategy to overcome these disadvantages is to combine chitosan with other biopolymers [16] such as the poly(lactic acid) (PLA) used in the present work. PLA is a commonly used polymer for packaging because it has mechanical, thermal, and barrier properties comparable to the most used synthetic plastics, having the advantage of being biodegradable and obtained from renewable sources [17]. Particularly in the last decade, different research groups have focused on the development of PLA/Ch materials [18–22]. Sébastien et al. [18] and Grande and Carvalho [19] obtained composite films by solution mixing and film casting processes. Although the Ch-PLA films produced by Sébastien et al. [18] showed interesting antifungal activity, their heterogeneity and high water sensitivity restrict their usage as food packaging materials [18]. Later, Soares et al. [20] synthesized biodegradable sheets of PLA and coated them with cross-linked chitosan by both spraying and immersion techniques. Nevertheless, the antimicrobial performance of these films was not further evaluated [20]. Bonilla et al. [21] and Chang et al. [22] prepared chitosan–PLA films containing various amounts of chitosan by extrusion and demonstrated their antibacterial activity in refrigerated meat and fish samples, respectively.

The main source of commercial chitosan is chitin, which is the second most abundant polysaccharide on Earth, only preceded by cellulose. Chitin is formed by N-acetylglucosamine units linked by β-(1→4) glycosidic bonds and is commonly sourced from crustacean shells [23], although some molluscs, such as squid, insects [24], and fungi [25], also incorporate this polysaccharide. The disposition of the chitin chains depends on the source; in the case of a squid pen, the source of the chitosan tested in this work, this is parallel (β-chitin), leading to weaker inter- and intra-molecular forces and, as a result, increased solubility and water-absorbing capacity [26]. Weaker forces also carry advantages for the partial deacetylation of N-acetylglucosamine to glucosamine units in chitin to produce chitosan. A sufficient number of glucosamine units enables dissolution in dilute acids due to the protonation of the free amino groups. Furthermore, deacetylation of squid pen β-chitin by alkaline hydrolysis produces more homogeneous chitosan as acetyl groups are more easily accessible than in other sources [27].

In addition to the chemical potential of the chitin/chitosan extracted from the squid pens, their use can help to solve disposal problems for the processing industry and potential environmental impacts. In fact, global squid captures have risen in the last few decades, reaching almost 4 million tonnes in 2015 [28]. Although only representing 5% of all fish, molluscs, and crustaceans captured, squid processing produces a substantial amount of waste. Considering that the yield of edible flesh in squid ranges from 60 to 80% [29], the annual waste generation can be estimated at 0.8 to 1.6 million tonnes and the disposal can be costly in developed countries [30]. Despite the huge potential of chitin and chitosan, currently, they are only employed in a few areas of industrial chemistry, such as cosmetics, textiles, water treatment, and biomedicine [30]. Therefore, a novel concept of shell biorefinery has been suggested on account of the massive potential of chitin valorization. Shell biorefinery consists of the sustainable conversion of chitin into several nitrogen-rich chemicals for pharmaceuticals, cosmetics, textiles, water treatment, household cleansers, soaps, and carbon dioxide sequestration, which benefits both the economy and the environment [30–32].

Variations in chitosan's antibacterial efficacy arise from numerous parameters, including intrinsic factors of chitosan, such as positive charge density, molecular weight (Mw), concentration, and hydrophilic/hydrophobic characteristics [33,34]. In the present work, the effect of Mw on the antimicrobial activity of chitosan is addressed. Although several studies have focused on this parameter, they have generated contradictory results concerning the relation between bactericidal activity and chitosan Mw. Some studies reported that increasing chitosan Mw leads to decreasing chitosan activity against *Escherichia coli*, while others suggested that high Mw chitosan displays greater activity than low Mw chitosan [34].

This study was undertaken to (1) produce β-chitosan and β-chitooligosaccharides from the *Loligo opalescens* pen, (2) develop and characterize PLA surfaces coated with β-chitosan and its derivatives of different Mw, and (3) evaluate the antimicrobial activity of PLA/Ch composite films against *Escherichia coli* and *Pseudomonas putida*, which are bacterial strains present in food processing environments [35,36] and may be responsible for the spoilage of chilled food products [37,38]. This work encompasses the crucial steps for the synthesis and characterization of PLA/Ch films for application in food contact surfaces, from the extraction of chitin and production of chitosan from marine by-products to antimicrobial tests, using innovative combinations of PLA and chitosan and its derivatives.

#### **2. Results**

#### *2.1. Production and Characterization of Chitosan and Derivatives*

Endoskeleton (pen) by-products of the squid species *Loligo opalescens* were initially processed to obtain chitosan by a combination of enzymatic and alkaline treatments, following the optimal conditions defined in a previous work [39]. Alcalase was the enzyme selected for the first step of pen deproteinization to produce chitin, and NaOH was the alkali utilized for the subsequent conversion of chitin into chitosan (Figure 1a). A highly purified β-chitosan (β-Ch) with a 92% of deacetylation degree (Figure 1b) and molecular weights of Mn (number average molecular weight) = 206 kDa/Mw (weight average molecular weight) = 294 kDa was finally recovered (Figure 1c). Based on the protocol of sodium nitrite depolymerization (see Section 4.1), three β-chitooligosaccharides (β-Cho) were produced: (1) a Ch of Mn = 138 kDa/Mw = 186 kDa (β-ChoA, Figure 2a), (2) a Ch of Mn = 84 kDa/Mw = 129 kDa (β-ChoB, Figure 2b), and (3) a Ch of Mn = 37 kDa/Mw = 61 kDa (β-ChoC, Figure 2c).

The rheological behavior of 1% (*w/v*) β-Ch and β-Cho solutions was assessed by evaluating their flow curves at 25 ◦C (Figure 3). All chitosan solutions revealed a Newtonian behavior, i.e., the viscosity was not dependent on the shear rate. However, it was possible to observe that the β-Ch solution was approximately 10 times more viscous than the three β-Cho samples (β-ChoA, β-ChoB, and β-ChoC).

#### *2.2. Characterization of Functionalized Poly(lactic acid) (PLA) Films*

#### 2.2.1. Water Contact Angle

Oxygen plasma was applied on the PLA surface to enhance its wettability, adhesion, and biocompatibility [40]. Figure 4 shows the effect of oxygen plasma treatment on the water contact angle of the film. The value of the water contact angle obtained for the PLA before treatment was 79.2 ± 1.1◦. A surface with a water contact angle between 0◦ and 30◦ can be considered hydrophilic, while a hydrophobic surface is characterized by contact angles over 90◦ [41]. Therefore, the PLA surface was quite hydrophobic as the initial contact angle for the untreated surface was close to 90◦. After the plasma treatment, a significant reduction in the contact angle value was observed. Figure 4 also presents the water contact angles after the deposition of chitosan samples on the PLA surface. PLA films were functionalized with the 1% (*w/v*) β-Ch and β-Cho solutions described above by the dip-coating method. After chitosan deposition, a decrease of approximately 45% in the contact angle of the surfaces was observed compared to the non-functionalized PLA after

plasma treatment. This result indicates that chitosan was successfully deposited on the PLA films. Moreover, the type of immobilized chitosan did not influence the wettability of the surfaces since their water contact angles were very similar (around 38◦).

**Figure 1.** (**a**) Flowchart of chitosan (Ch) production from *Loligo opalescens* squid; (**b**) analysis of Ch by nuclear magnetic resonance (NMR) to calculate the purity and degree of deacetylation; (**c**) eluogram of Ch analyzed by gel permeation chromatography (GPC) for molecular weight determination (RID—refractive index signal, RALS—right angle light scattering signal).

**Figure 2.** Eluograms of (**a**) β-ChoA, (**b**) β-ChoB, and (**c**) β-ChoC analyzed by gel permeation chromatography (GPC) for molecular weight determination (RID—refractive index signal, RALS—right angle light scattering signal).

**Figure 3.** Flow curves of 1% (*w/v*) solutions of chitosan and derivatives used for surface preparation: β-Ch (1), β-ChoA (2), β-ChoB (3), and β-ChoC (4).

**Figure 4.** Water contact angles ± standard deviations (SDs) for PLA films before and after oxygen plasma treatment, and PLA/Ch films created by dip-coating method (PLA/Ch, PLA/ChoA, PLA/ChoB, and PLA/ChoC).

#### 2.2.2. Fourier-Transform Infrared Spectroscopy (FTIR) Analysis

FTIR spectra of different types of chitosan immobilized onto the PLA surface are shown in Figure 5. The different types of chitosan deposited onto PLA did not reveal significant differences in FTIR results (Figure 5a). Characteristic bands of chitosan were covered by the presence of bands from the PLA film. The broad \_OH stretching absorption band between 3680 and 2750 cm−<sup>1</sup> and one between 2980 and 2750 cm−<sup>1</sup> assigned to aliphatic C-H stretching indicated the presence of chitosan on PLA films [42]. Furthermore, a characteristic NH stretching band of chitosan with a maximum at 3350 cm−<sup>1</sup> was identified on the functionalized PLA/Ch films (Figure 5a).

The FTIR spectrum of PLA films is included in Figure 5b. Results show that characteristic bands of PLA with high-intensity peaks were represented at 1750 cm−<sup>1</sup> corresponding to CO, at 1188–1090 cm−<sup>1</sup> corresponding to CO, at 1452–1368 cm−<sup>1</sup> corresponding to COH, and at 3000 cm−<sup>1</sup> corresponding to CH. These results are in accordance with those reported by Stoleru et al. [43]. In general, FTIR spectra revealed that surface immobilization of chitosan onto PLA films was successfully achieved.

**Figure 5.** FTIR spectrum of (**a**) different types of chitosan immobilized onto PLA (β-Ch (1), β-ChoA (2), β-ChoB (3), β-ChoC (4)) and (**b**) PLA films (PLA (5)).

#### 2.2.3. X-ray Diffraction (XRD)

The diffraction patterns of PLA films and the effect of different types of chitosan immobilized on PLA are shown in Figure 6. In the case of PLA film, diffraction peaks at 2θ = 16.5◦, 20◦, and 22◦ were observed, which corresponded to the characteristic peaks of PLA, indicating a crystalline PLA matrix [44]. An accentuated band at 2θ = 28◦ suggested the presence of calcium carbonate on PLA films, as described in the literature [45]. The immobilization of chitosan solutions caused a decrease in the intensity peaks at 2θ = 16.5◦, 20◦, and 22◦ [46], which confirmed the deposition of chitosan. This modification could indicate a decrease in the crystalline part of PLA/Ch films. It is also possible to conclude that different types of chitosan did not affect the XRD pattern.

**Figure 6.** X-ray diffraction (XRD) patterns of different types of chitosan immobilized onto PLA surface (β-ChoC (1), β-ChoB (2), β-ChoA (3), and β-Ch (4)) and of PLA film (5).

#### 2.2.4. Morphological Studies

For determination of the surface morphology of the control PLA and Ch-based films, scanning electron microscopy (SEM) analysis was performed. The SEM image corresponding to the PLA film (Figure 7a) reveals a homogeneous surface with a uniform appearance. After the deposition of chitosan onto PLA surfaces, the presence of small particles, which could be unsoluble materials of chitosan, was detected (Figure 7b–e). Although the func-

tionalized surfaces were heterogeneous (Figure 7b–e), no significant differences in PLA/Ch appearance were observed for the different types of chitosan tested.

**Figure 7.** Scanning electron micrographs of (**a**) PLA surface and (**b**–**e**) Ch-based surfaces: (**b**) β-Ch, (**c**) β-ChoA, (**d**) β-ChoB, and (**e**) β-ChoC.

#### *2.3. Antimicrobial Activity of Functionalized PLA Films*

The antimicrobial properties of PLA and PLA/Ch surfaces were evaluated in conditions mimicking the storage conditions of packaged food products, namely a short incubation period (1 day), refrigeration temperature (5 ◦C), and static conditions. Furthermore, two different biofilm-forming bacteria typically associated with the food environment were used: *Escherichia coli* (a model pathogen) and *Pseudomonas putida* (isolated from a salad processing industry) [35,36].

The cellular composition of *E. coli* and *P. putida* single-species biofilms formed on the control PLA and PLA/Ch films was evaluated by counting viable, viable but nonculturable (VBNC), and culturable cells (Figure 8), whereas the spatial distribution of the biofilms developed by both bacterial strains on the surfaces was analyzed by confocal laser scanning microscopy (CLSM; Figures 9 and 10).

The analysis of *E. coli* biofilm cells (Figure 8a) indicated that there was a significant decrease in the number of all cell types considered (viable, VBNC, and culturable cells) on the chitosan-coated PLA surfaces compared to PLA, except in the case of PLA/ChoB (i.e., the PLA coated by the β-chitooligosaccharide of intermediate molecular weights derived from β-chitosan). Indeed, the biofilms formed on PLA/Ch, PLA/ChoA, and PLA/ChoC surfaces exhibited, on average, 70%, 74%, and 63% fewer *E. coli* viable, VBNC,

and culturable cells, respectively, than PLA (*p* < 0.001, Figure 8a). When comparing the antimicrobial efficacy of immobilized native chitosan (PLA/Ch) with its derivatives (PLA/ChoA, PLA/ChoB, and PLA/ChoC, in descending order of molecular weight), it was observed that PLA films coated with depolymerized chitosan had bactericidal behavior similar to PLA films coated with native chitosan (Figure 8a), except for PLA/ChoB.

**Figure 8.** Cellular composition of (**a**) *E. coli* and (**b**) *P. putida* biofilms on PLA and Ch-based surfaces: viable (-), viable but non-culturable (VBNC) (-), and culturable cells (-). Inferential statistics were performed using unpaired *t*-tests or Mann–Whitney tests according to the normality of the variables' distributions. The means ± SDs for three independent experiments are illustrated. Within the same type of cells, the significance levels were \* *p* < 0.05, \*\* *p* < 0.01, and \*\*\* *p* < 0.001 related to PLA (\*) and PLA/Ch (+).

Regarding the effectiveness of PLA-coated films against *P. putida* biofilm growth (Figure 8b), it was also evident that they were effective in reducing the number of viable, VBNC, and culturable cells by, on average, 73%, 52%, and 87%, respectively. These percentages exclude the cell numbers of the PLA/ChoC surface (i.e., the PLA coated by the β-chitooligosaccharide of higher Mw). Despite having a smaller number of culturable cells than PLA (67%, *p* < 0.001), this coating showed equal or higher numbers of the remaining types of cells.

Looking at the results of the two bacterial strains together (Figure 8a,b), they suggest that the surfaces with the greatest antimicrobial activity were those coated with the native β-chitosan (Mn = 206 kDa/Mw = 294 kDa) and the depolymerized β-chitosan of the highest molecular weight (ChoA, chitosan of Mn = 138 kDa/Mw = 186 kDa). On the other hand, no linear relationship was found between the molecular weight of the tested chitosans obtained from the *Loligo opalescens* pen and their antimicrobial performance under the conditions tested in this study.

The effect of immobilized chitosan and its derivatives against *E. coli* and *P. putida* biofilm formation was also analyzed by CLSM. Both bacteria formed dense and thick biofilms, regardless of the tested surface (Figure 9). Nevertheless, microscopic images revealed that, in general, *E. coli* and *P. putida* biofilms grown on uncoated PLA surfaces were thicker than those developed on PLA films coated with chitosan, which may be related to its antimicrobial activity (Figure 8). These differences in biofilm thickness were particularly evident for PLA/Ch and PLA/ChoA surfaces in the case of *E. coli* (Figure 9b,c), and for PLA/Ch, PLA/ChoA, and PLA/ChoB films with *P. putida* (shadow projection on the right of Figure 9g–i). Furthermore, quantitative data showed a decrease of up to 26% in the average thickness of *E. coli* biofilms formed on PLA/Ch and PLA/ChoA surfaces (shadow projection on the right of Figure 9b), whereas *P. putida* biofilms developed on the same surfaces had approximately 44% and 36% less thickness and biovolume, respectively, than those grown on PLA (shadow projection on the right of Figure 9c,d). Thus, besides

bactericidal activity, the native chitosan and its higher Mw derivative also prevented biofilm growth.

**Figure 9.** Representative biofilm structures of (**a**–**e**) *E. coli* and (**f**–**j**) *P. putida* on PLA and Ch-based surfaces. These images were obtained from confocal *z*-stacks using IMARIS software and present an aerial, three-dimensional (3D) view of the biofilms (images on the left). The shadow on the right represents the vertical projection of the biofilm. The white scale bar is 200 μm and the numerical scale indicated in each panel is in μm.

**Figure 10.** Biovolume and thickness of (**a**,**b**) *E. coli* and (**c**,**d**) *P. putida* biofilms formed on PLA and Ch-based surfaces. The values were obtained from the confocal *z*-stacks using the COMSTAT2 tool associated with the ImageJ software. Statistical analysis was performed using one-way analysis of variance (ANOVA) and the significance levels were \* *p* < 0.05, \*\* *p* < 0.01, and \*\*\* *p* < 0.001 related to PLA (\*) and PLA/Ch (+).

#### **3. Discussion**

The demand for bio-based and safer materials is increasing due to the growth of the human population, industrial development, and environmental concerns. Among known biopolymers, chitosan was selected to confer antibacterial properties to poly(lactic acid) surfaces as it presents remarkable characteristics such as non-toxicity, biocompatibility, and biodegradability [10–12]. From the particular perspective of food packaging, easy film formation and antimicrobial activity are considered the most important properties of chitosan, which have been extensively studied [5,7,12,14,47].

Poly(lactic acid) is a polyester formed from 100% renewable raw materials and is highly transparent, hence its intensive use in many disposable packaging solutions. One of the strategies to modify PLA is to apply a surface treatment using a plasma process in order to improve the wettability, adhesion, and biocompatibility of this biopolymer. As expected, the wettability of PLA films used in the present study increased after treatment with oxygen plasma. Identical behavior was reported by Jordá-Vilaplana et al. [40], who used atmospheric plasma treatment to improve the adhesion capacity of PLA films.

In this work, chitosan and its depolymerized derivatives were successfully obtained from a *Loligo opalescens* squid pen by-products via a combination of enzymatic and alkaline treatments [39] and immobilized onto PLA films through plasma oxygen treatment followed by dip-coating. In the last few years, a considerable number of studies have been published on the production of new polymeric systems incorporating chitosan. Some studies have focused on the synthesis of PLA/Ch composites by solution mixing and film casting [19,48,49], but others prepared PLA films by extrusion, coated them with a Ch solution, and used crosslinking agents [20].

Most of the commercial chitosan solutions reported in the literature revealed a viscoelastic behavior, i.e., a high dependence of viscosity with shear rate, except for the lowconcentration solutions (lower than 0.25–0.5% (*w/v*)) [50,51]. On the contrary, the results here obtained at 1% (*w/v*) of Ch and derivatives showed a Newtonian behavior, i.e. null dependence of viscosity with shear rate. Other authors observed the same flow behavior, with a Newtonian plateau at a low chitosan concentration of 1.7%, but no Newtonian behavior could be observed at higher chitosan concentrations [52], which suggests that the viscosity of chitosan solutions may be affected by other factors, such as the degree of deacetylation, molecular weight, temperature, and pH [53–55]. Additionally, the decrease in viscosity with the decrease in molecular weight among β-Ch and β-Cho was in agreement with the data collected by Chattopadhyay and Inamdar [54]. These authors demonstrated that the viscosity of chitosan is influenced by its Mw and that the intrinsic viscosity of different grades of chitosan decreased approximately ten-fold when the viscosity average molecular weight dropped from 285 to 21 [54]. Furthermore, the decrease in viscosity of β-Cho compared to native chitosan can facilitate the formulation of coatings for application in food technology [56,57].

After the synthesis of the PLA/Ch surfaces, they were tested against two bacterial strains (*E. coli* and *P. putida*) in conditions that simulated the short-term food packaging environment (refrigeration without agitation and 1 day of contact with the cell suspensions). To the best of our knowledge, this is one of the very few studies that evaluates the antibiofilm effect of PLA surfaces coated with native chitosan and derivatives without adding any other compounds, such as essential oils or metallic nanoparticles, to reinforce the antimicrobial properties of coatings and extend the shelf-life of packaged food products [58]. In general, the native chitosan surfaces demonstrated bactericidal activity against the Gram-negative bacteria tested. They decreased the number of biofilm viable, viable but nonculturable, and culturable cells by up to 73%, 74%, and 87%, respectively, compared to PLA. Although the exact mechanism of antibacterial activity of chitosan is still unclear, several mechanisms have been proposed. First, the antimicrobial activity of Ch films is dependent on the degree of deacetylation since the presence of charged amino groups on chitosan molecules can disturb the negatively charged phosphoryl groups on the bacterial cell membrane, leading to its degradation followed by cell death [47]. Chitosan may also form an impermeable layer around the bacterial cell and block the exchange of essential solutes between the intra- and extracellular environment, affecting the physiological state of bacteria and, ultimately, causing cell death [59]. This biopolymer can also diffuse through the cell wall, disrupt the cytoplasmic membrane of bacteria and affect its integrity, as well as suppress the synthesis of RNA and proteins by binding to DNA molecules [58,59].

The antimicrobial activity of chitosan and its derivatives relies on numerous intrinsic and extrinsic factors, including pH, microorganism species, degree of deacetylation of Ch, Mw, concentration, hydrophilic/hydrophobic characteristics, etc. [33,34]. In this work, the surfaces with immobilized chitosan showed different bactericidal performance and inhibited biofilm growth differently. Since the water contact angles and SEM images were identical for all Ch-based surfaces, it is believed that the physicochemical properties and morphology of the films did not affect their antibacterial behavior. Therefore, the main parameter influencing this behavior seems to be the molecular weight of chitosan. Several studies have discussed the relation between bactericidal activity and chitosan Mw, although with contradictory results. Some authors indicated that increasing chitosan Mw leads to decreasing chitosan activity against *Escherichia coli*, while others suggested that high Mw chitosan displays greater activity than low Mw chitosan [34,60]. Hirano et al. [61] showed

that chitosans with a Mw of 1.5–4.5 kDa exhibited better inhibitory activities than those with a higher Mw (6.5–12.0 kDa). On the other hand, Tokura et al. [60] disclosed that a 9.3 kDa chitosan inhibited the growth of *E. coli*, whereas a 2.2 kDa chitosan increased bacterial growth. Moreover, some authors have shown that intermediate molecular weights are more effective [34,62]. For instance, Li et al. [62] demonstrated that chitosan suppressed *E. coli* growth, but its inhibitory activity differed with Mw (from 3 to 1000 kDa), with the Ch of 50 kDa Mw having the strongest effect. Similar to Li et al. [62], no linear relationship was found between the Mw of the tested chitosans obtained from the *Loligo opalescens* pen and their antimicrobial performance under the tested conditions. Moreover, the results indicate that the surfaces with the greatest antibiofilm activity were those coated with the native β-chitosan and the depolymerized β-chitosan of the highest molecular weight (ChoA). It is possible that the chitosan of higher Mw interacted with the bacterial cells adhered to the substrate and altered cell permeability, resulting in cell lysis [34]. At the same time, the positive charge of Ch given by the functional amino groups (NH3 +) of N-acetylglucosamine units is expected to react electrostatically with the negatively charged biofilm components such as EPS, proteins, and DNA [57,63], which may explain the lower biovolume and thickness of biofilms exposed to surfaces coated with higher Mw chitosan. It is also reported that high Mw water-soluble chitosan and solid chitosan may form an impermeable layer around the cell surface, thus blocking the transport of nutrients into the cell and causing cell lysis [64,65].

Another interesting result of this work is the capacity that the chitosan-based films showed to reduce the number of viable but nonculturable cells in the biofilms formed by both microorganisms when compared to uncoated PLA (up to 74%). The VBNC state is a unique survival strategy of many bacteria in the environment in response to adverse conditions. The foodborne pathogens may enter the VBNC state during food processing operations, such as disinfection, preservation, and low-temperature storage, and represent a threat to food safety and public health as cells in this state are not detectable through conventional food and water testing methods [66,67]. Indeed, VBNC bacteria cannot be cultured on routine microbiological media, but they remain viable and retain virulence. Therefore, this study suggests that replacing PLA films with PLA coated with chitosan may be an interesting solution to eliminate most VBNC cells in the food packaging environment, preventing foodborne infections and increasing food safety. These Ch-based surfaces may also be beneficial in improving the shelf-life of food products, which is dependent on microbial contamination since certain microorganisms can modify the odor, flavor, color, and textural properties of food products [68].

#### **4. Materials and Methods**

#### *4.1. Production and Chemical Characterization of Chitosan and Chitooligosaccharides*

Chitosan previously isolated from the pen of *Loligo opalescens* squid [69] was depolymerized by reaction with sodium nitrite [70]. High molecular weight chitosan of 206 kDa (Mn, number average molecular weight) was initially purified by overnight dissolution in 5% (*v/v*) acetic acid at 9 g/L, followed by filtration (FILTER-LAB®ref. 1250, 10–13 μm; Filtros Anoia, S.A., Barcelona, Spain) and precipitation with methanol:25% ammonia in a proportion 1:3 (*v/v*) chitosan solution:methanolic ammonia. The mixture was left at 4 ◦C for 1 h, then centrifuged in 1 L bottles at 13,261× *g* on a Beckman Coulter Avanti J-25I centrifuge (Beckman Coulter, Inc, Indianapolis, IN, USA). The precipitates were washed three times with water followed by a final acetone wash, dried overnight at 50 ◦C on a stove, and freeze-dried. Purified chitosan was milled to a fine powder and dissolved overnight in triplicate in 0.05 M HCl at 8 g/L. Depolymerization reactions were carried out under stirring at room temperature by adding the appropriate amounts of a 1.6 g/L sodium nitrite solution to each chitosan solution, according to the following equation [39]:

$$\frac{1}{M\_f} - \frac{1}{M\_o} = \frac{n}{m} \tag{1}$$

where *Mf* is the molecular weight of chitosan after depolymerization, *M0* the initial molecular weight of chitosan, *n* the moles of sodium nitrite, and *m* the initial mass of chitosan. After 4 h of reaction, chitosan was precipitated with 5 M NaOH, the solids separated by centrifugation as described above for the purification process, and washed with water until neutrality. Finally, depolymerized samples were freeze-dried and milled to a fine powder.

The degree of acetylation of chitosan was estimated from nuclear magnetic resonance (NMR) experiments. Chitosan samples (7 g/L) were dissolved in 0.056 M deuterated trifluoroacetic acid (TFA-d in D2O), and the corresponding 1H NMR spectra were recorded at 400 MHz (Bruker Avance II; Brucker, USA). The degree of acetylation was calculated from the relative integrals of acetyl (N-acetyl and AcOH) and combined H2–H6 protons (GlcN and GlcNAc) [71,72]. Chemical shifts were expressed in ppm with the HOD solvent signal acting as a reference. Mestrenova 10.0 software (Mestrelab Research, S.L., Santiago de Compostela, Spain) was used for spectral processing.

The molecular weight of chitosan samples was determined by gel permeation chromatography (GPC) on an Agilent 1260 system equipped with a quaternary pump, injector, column oven, and refractive index and static dual-angle light scattering detectors. Chitosan was separated with a set of four columns: Novema Precolumn (10 mm, 8 × 50 mm), Novema 30 Å (10 mm, 8 × 300 mm), Novema 1000 Å (10 mm, 8 × 300 mm), and Novema 1000 Å (10 mm, 8 × 300 mm) from Polymer Standards Service, Mainz, Germany. Column oven and light scattering detector were kept at 30 ◦C and the refractive index detector was maintained at 40 ◦C. Samples were eluted with 0.15 M ammonium acetate–0.2 M acetic acid (pH 4.5) as mobile phase at 1 mL/min. Chitosan samples were dissolved in the GPC buffer at a concentration of 1 g/L. Detectors were calibrated with a polyethylene oxide standard of 106 kDa and polydispersity index (PDI) of 1.06 (PSS Polymer Standards Service GmbH, Mainz, Germany). Molecular weight of chitosan was estimated using a refractive index increment (dn/dC) value of 0.18 [73]. From this, two types of chitosan size determination were considered: (1) the average molecular weight of the biopolymer (Mw) and (2) the number average molecular weight of the biopolymer (Mn).

#### *4.2. Immobilization of Chitosan and Chitooligosaccharides onto Poly(lactic acid) (PLA) Films*

Solutions of chitosan (Ch) and three chitooligosaccharides (ChoA, ChoB, and ChoC) at 1% (*w/v*) were immobilized onto poly(lactic acid) (PLA) films (Goodfellow, UK). For the immobilization of chitosan and derivatives onto the surfaces of PLA films, a plasma oxygen treatment (Harrick Plasma, PJS-14-0240) was applied using a moderate intensity for 15 min. After this, PLA films with dimensions of 1 cm × 1 cm were dipped in the different solutions (Ch, ChoA, ChoB, and ChoC) for 15 min and dried with nitrogen for 5 min.

#### *4.3. Rheological Measurements of Chitosan Solutions*

Flow curves were obtained using a Discovery Hybrid Rheometer (DHR1) from TA Instruments (New Castle, DE, USA) with Peltier temperature set to 25 ◦C. TRIOS Software (New Castle, DE, USA) was used to control the equipment and to acquire rheological parameters. A stainless-steel cone-plate geometry of 60 mm, with an angle of 2.006◦ and truncation of 64 μm, was used due to its capability of generating a uniform shear rate across the samples. Steady-state flow curves were obtained working in a controlled-stress mode, over the shear rate range of 1–300 s<sup>−</sup>1. All the samples (Ch, ChoA, ChoB, and ChoC at 1% (*w/v*)) were measured in triplicate.

#### *4.4. Surface Characterization*

#### 4.4.1. Water Contact Angle

Films' surface hydrophobicity was evaluated through the measurement of contact angle by sessile drop technique using a DSA 100E drop shape analysis system (Kruss Gmbh, Hamburg, Germany). A water droplet (2 μL) was deposited at different points on the film surface, and afterwards, a digital camera connected to DSA 3 drop shape image analysis

software recorded drop images. The image produced was used to calculate the contact angle via circle fitting method [74]. At least 10 measurements were taken per tested film.

#### 4.4.2. Fourier-Transform Infrared Spectroscopy (FTIR)

FTIR spectra of the films were recorded with VERTEX 80v FTIR spectrometer (Bruker, Germany) in the wavelength range 4000–400 cm−<sup>1</sup> at a resolution of 4 cm<sup>−</sup>1, using Platinum Attenuated Total Reflection mode (ATR) (Bruker, Germany). The absorbance of each FTIR spectrum was normalized between 0 and 1 [75].

#### 4.4.3. X-ray Diffraction (XRD)

XRD was used to investigate the presence and influence of chitosan on the crystalline structure of the polymer matrix. The XRD patterns of PLA films with and without chitosan were determined using a diffractometer (PanAnalytical X Pert PRO MRD system, Malvern, UK). The scanning range varied from 2θ = 10◦ to 50◦ [76].

#### 4.4.4. Scanning Electron Microscopy (SEM)

The morphology of the film's surface was observed using a scanning electron microscope (Quanta 650 FEG, FEI Europe B.V., Eindhoven, The Netherlands) with an accelerating voltage of +5 kV at different magnifications [75]. The samples were cut with a blade and mounted on sample holders with double-sided adhesive and sputtered with a 10 nm layer of gold.

#### *4.5. Antimicrobial Activity of Functionalized PLA Films*

#### 4.5.1. Bacterial Strains and Culture Conditions

A model food pathogen—*Escherichia coli* SS2 expressing the green fluorescent protein (GFP) (*E. coli* SS2 GFP)—and an industrial isolate from a salad processing plant— *Pseudomonas putida*—were the bacteria chosen for this study [35,36,77]. Stock cultures were maintained at −80 ◦C in Tryptone Soy Broth (TSB; BioMérieux, Marcy-l'Étoile, France) containing 20% (*v/v*) glycerol. Before each experiment, frozen cells were subcultured twice in TSB at 30 ◦C with a constant orbital agitation of 120 rpm [36].

#### 4.5.2. Biofilm Formation

Biofilm assays were performed on 12-well plates (VWR International, Carnaxide, Portugal) for 1 day at 5 ◦C under static conditions in order to mimic the conditions typically found in short-term food packaging. Before each experiment, the PLA film and the PLAcoated surfaces produced as described in Section 4.2 were sterilized by ultraviolet (UV) radiation for 30 min. Then, the sterilized surfaces were placed on the wells and inoculated with 3 mL of an overnight culture of *E. coli* SS2 GFP or *P. putida* in TSB adjusted to an optical density (OD) of 0.01 at 610 nm (1:10 dilution from an initial cell suspension at OD610 nm = 0.1). The microplates were then kept at 5 ◦C for 1 h to promote bacterial attachment to the surface materials [36,77]. After this adhesion step, the wells were emptied and refilled with 3 mL of sterile TSB, and the microplates were incubated to allow biofilm development. Furthermore, 3 mL of TSB was added to the wells containing sterilized surfaces to monitor their sterility throughout the experiments.

Biofilm formation experiments were performed in three independent assays, each one with three technical replicates.

#### 4.5.3. Biofilm Cell Quantification

Biofilm cell suspensions were obtained by dipping each surface in 2 mL 0.85% (*v/v*) NaCl and vortexing for 3 min. Biofilm cell culturability was assessed after serial dilutions in Tryptone Soy Agar plates (TSA; BioMérieux, Marcy-l'Étoile, France) in the case of *P. putida*, and TSA plates supplemented with 0.1 g/L of ampicillin for *E. coli* SS2 GFP. In turn, biofilm viability was evaluated by staining the biofilm suspension with the Live/Dead® BacLight™ Bacterial Viability kit (Invitrogen Life Technologies, Alfagene, Portugal) as previously

described [78] and observing it in an epifluorescence microscope (Leica DM LB2; Leica Microsystems, Wetzlar, Germany). A minimum of twenty fields of view was analyzed for each stained sample using the ImageJ software (version 1.52p, U.S. National Institutes of Health, Bethesda, MD, USA) and the number of viable cells was counted. Finally, the number of VBNC cells was determined by subtracting the number of culturable cells from that of viable cells [79]. The number of culturable cells was presented as CFU/cm2, whereas the numbers of viable and VBNC cells were expressed as cells/cm2.

#### 4.5.4. Confocal Scanning Electron Microscopy (CLSM)

Single-species biofilms of *E. coli* and *P. putida* that developed after 1 day on all tested surfaces were observed using a 10× dry objective (Leica HC PLAN APO CS) in an inverted microscope Leica DMI6000-CS (Leica Microsystems, Wetzlar, Germany). *E. coli* cells were pinpointed from the GFP expression, while *P. putida* biofilms were stained in red with 5 μM SYTO® 61 (Invitrogen Life Technologies, Alfagene, Portugal), a cell-permeant fluorescent nucleic acid marker. For this reason, *E. coli* biofilms were observed with a 488 nm argon laser, whereas *P. putida* biofilm samples were scanned at an excitation wavelength of 633 nm (helium-neon laser) [36]. A minimum of six stacks of horizontal plane images (512 × 512 pixels, corresponding to 1550 × 1550 μm) with a *z*-step of 1 μm were acquired for each sample.

Three-dimensional (3D) projections of biofilm structures were reconstructed using the "Easy 3D" tool of IMARIS 9.1 software (Bitplane, Zurich, Switzerland) directly from the CLSM acquisitions. The plug-in COMSTAT2 associated with the ImageJ software was used to determine the biovolume (μm3/μm2) and biofilm thickness (μm) [80].

#### *4.6. Statistical Analysis*

Descriptive statistics were used to calculate the mean and standard deviations (SDs) for the number of viable, VBNC, and culturable cells (Figure 8), and biovolume and biofilm thickness (Figure 10). Differences in the number of cells in relation to PLA and PLA/Ch (Figure 8) were evaluated using unpaired *t*-tests or Mann–Whitney tests according to the normality of the variables' distributions. Quantitative parameters obtained from confocal microscopy (Figure 10) were compared using a one-way analysis of variance (ANOVA). All tests were performed with a confidence level of 95% (*p*-values < 0.05). Data analysis was performed using IBM SPSS Statistics version 24.0 for Windows (IBM SPSS, Inc., Chicago, IL, USA).

#### **5. Conclusions**

The recent increase in sensitivity towards environmental issues arising from plastic packaging has fostered an interest in alternative sustainable packaging materials. Chitosan and its derivatives were efficiently prepared from fishery waste and immobilized onto PLA films. Their antimicrobial effects were demonstrated in the composite films, with a strong reduction in viable, VBNC, and culturable cell counts in *E. coli* and *P. putida* biofilms. Furthermore, the surfaces with the highest antibiofilm activity were those coated with the native Ch and the Cho with the highest Mw. This is the first time that PLA/Ch surfaces have been shown to be able to eliminate most VBNC cells in the food environment, which is a very interesting result given that such cells retain a public health risk. Further research is needed to bring this biopolymer to industrial levels for food packaging applications.

**Author Contributions:** Conceptualization, F.J.M., J.A.V. and L.P.; methodology, L.C.G., S.I.F., J.A.V., J.V., A.I.B. and M.A.C.; formal analysis, L.C.G. and S.I.F.; investigation, L.C.G., S.I.F., J.A.V., J.V., A.I.B. and M.A.C.; resources, F.J.M. and L.P.; data curation, L.C.G. and S.I.F.; writing—original draft preparation, L.C.G.; writing—review and editing, F.J.M., J.A.V., J.V., A.I.B. and M.A.C.; supervision, F.J.M. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by Base Funding—UIDB/00511/2020 of the Laboratory for Process Engineering, Environment, Biotechnology and Energy (LEPABE) funded by national funds through the FCT/MCTES (PIDDAC), and "CVMAR+I—Industrial Innovation and Marine Biotechnology Valorization" project, funded by INTERREG V Espanha Portugal (POCTEP) (0302\_CVMAR\_I\_1\_P). The research was also supported by the SurfSAFE project funded by the European Union's Horizon 2020 research and innovation programme under grant agreement no. 952471. L.C.G. thanks the Portuguese Foundation for Science and Technology (FCT) for the financial support of her work contract through the Scientific Employment Stimulus—Individual Call—(CEECIND/01700/2017). J.A.V. and J.V. also offer their thanks to Xunta de Galicia by Xunta de Galicia (Grupos de Potencial Crecimiento, IN607B 2018/2019) for the financial support.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The data presented in this study are available on request from the corresponding author. The data are not publicly available yet as some data sets are being used for additional publications.

**Conflicts of Interest:** The authors declare no conflict of interest.

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