Aronia mitschurinii Encapsulated into a Biocompatible Polymer as an Effective Solution for Antifouling Protection

Abstract

Since the 15th century, scientists have been working to address biofouling, the accumulation of biofilm on submerged surfaces, like ships and buoys. Key stages in this process include bacterial precipitation and radical polymerization of bacteria’s metabolites. Along with corrosion and extended fuel consumption by ships, biofilm also contributes to ecological changes by spreading invasive bacterial species across the world when segments of biofilm are detached from moving ships. Historically, effective antifouling solutions have been toxic to marine ecosystems and are now banned in many regions, including the US. While recently developed highly hydrophobic polymers have promise, they are currently expensive and challenging for technology transfer.As an alternative solution, nontoxic formulations have been studied for antifouling coatings that utilize organic renewable superfruits, medicinal herbs, and nonbiofilm-forming algae rich in antioxidants and/or essential oils and terpenes, combined with biocompatible polymers. It is hypothesized that antibacterial essential oils and terpenes would prevent precipitation and growth of bacteria on the protected surfaces, while antioxidants, which are natural free radical scavengers, would significantly slow down polymerization processes in the biofilms. This study focuses on formulations created with extracts of A. mitshurinii, known for its high antioxidant concentrations. Current short-term results indicate that A. mitschurnii serves as an effective antifouling agent in both freshwater and saltwater, but with a higher performance in freshwater environments. The influence of bacteria and phytochemical concentrations will be discussed, along with the current antifouling methodology and postsurface analysis.

Introduction

Biofouling refers to the adhesion and accumulation of microorganisms, such as bacteria and their polymerized metabolites, on submerged surfaces like ships, buoys, and platforms. Yet, it is not limited to these objects. Biofouling is also a reason for poor drainage in all kinds of water, including agricultural, industrial, and civilian water; water purification systems on submarines, space stations, and crafts; and it is believed to be a reason for dental plaque formation. The economic impact is significant; for instance, the shipping industry alone has faced costs exceeding 1 billion dollars over 15 years due to biofouling. This buildup leads to increased surface friction, which degrades vessel speed and efficiency. ,

Biofilm formation, a key component of biofouling, progresses through five stages (Figure ), of which the most critical are bacterial precipitation and radical polymerization of bacterial metabolites into the initial biofilm layer. The challenge of managing biofouling lies in the vast diversity of fouling species and the variation in local conditions, which makes universal antifouling strategies elusive. , Materials such as waxes, tar, copper, and arsenic-based compounds were historically employed to combat biofouling from the 15th to 16th centuries. By the 20th century, the development of polymer-based coatings marked a significant advancement, but environmental and health concerns led to the banning of toxic compounds like tributyltin (TBT) in 2008. In response, research shifted toward biocide-release coatings, which later showed a negative ecological effect on the natural microbiome in water; and nonbiocide-release coatings utilizing naturally occurring antifouling compounds from marine organisms. , Recent innovations include synthetic hydrophobic polymer brushes and nanoparticle-based coatings derived from environmentally safe materials. , However, the complex multistep synthetic process presents both technological and economic challenges in the technology transfer for such coatings.

1.

Five stages of biofilm formation.

Certain marine organisms, such as sea cucumbers and sea urchins, exhibit natural resistance to biofouling. , In contrast, oysters are highly susceptible to biofilm accumulation, which can block their gills and lead to mortality. ,

Free radicals, such as superoxide anions and hydroxyl radicals, can oxidize surfaces, promoting biofilm development. However, antioxidants like vitamins E and C can inhibit this process by neutralizing free radicals. Inhibiting processes of bacterial precipitation through surface modifications or antibacterial agents can impede biofilm formation. Antifouling coatings are classified into fouling-resistant, fouling-release, and fouling-degrading categories, each with specific strategies to combat different fouling mechanisms. ,

This research focuses on encapsulating plant extracts from plants rich in antioxidants, essential oils, and terpenes into poly­(methyl methacrylate) coatings. While a variety of such plants, herbs, and algae is very wide, and random attempts to use some of them for antifouling have been made in the past, here, for the first time, an intelligent approach was taken to selecting potential crops, which is based on the hypothesis that essential oils and terpenes act as antibacterial agents. In contrast, phenolic antioxidants act as natural, free radical scavengers, preventing the radical processes that lead to biofilm formation. To prove this concept, extracts of Aronia (A. mitschurinii), which are fruits that are native to Maryland, US, and known to be one of the highest in the world by its anthocyanins, flavonoids, and polyphenol content, were prepared, encapsulated into biocompatible PMMA, and used to coat samples.

While Aronia mitschurinii is a superfruit, it is unexpensive and easy to grow and use for a wide variety of applications. In North Dakota, A. mitschurinii has been found to grow at a rate of at least 30 pounds per shrub. Along with providing a high yield of fruit, the crop is also known for having high antioxidant concentrations in the pulp, meaning that the waste part of the crop can still be used for other applications, including antifouling research. This research aims to establish the potential of A. mitschurinii in antifouling applications, with plans to analyze individual components to enhance the concept of industrialization. However, the hardiness of the crop, along with the high concentration of desired phytochemicals that can be derived from waste products, indicates that A. mitschurinii is already a good candidate for an industrialized antifouling product.

Full phytochemical characterization for extracts used in antifouling formulations has been conducted, including the ORAC factor as evidence of the ability to capture free radicals. At the same time, potential antimicrobial properties were predicted based on essential oils and terpenes found in the GC-MS analysis of extracts. Here, the results of short-term antifouling tests in salt and fresh water sampled at the Eastern Shore of Maryland are presented. Post-test surface analysis using phase contrast, fluorescent, and laser microscopy, and profilometric results will be presented for A. mitschurinii.

Materials and Methods

General Procedures

The following chemical reagents were used in the experiments: aluminum chloride (99% extra pure, anhydrous granules), ethyl alcohol (99% ACS spectroscopic grade), and quercetin hydrate (95%), all obtained from Thermo Fisher Scientific. Potassium acetate (certified ACS crystalline) and sodium carbonate anhydrous (HPLC-grade powder) were sourced from Fisher Scientific. Folin and Ciocalteu’s phenol reagent and gallic acid monohydrate (ACS reagent grade) were provided by MP Biomedicals (Santa Ana, CA). Sodium acetate (Sigma Ultra, minimum 99.0%) was purchased from Sigma-Aldrich. Additional reagents, including concentrated HCl, concentrated NaOH, pure ethanol, and 95% ethanol, were also acquired from Fisher Scientific. Distilled water was used in all experimental procedures. The ultraviolet/visible (UV/vis) spectrophotometric analysis of anthocyanins, flavonoids, and polyphenols was performed using an Agilent Cary 60 UV–vis spectrophotometer (USA), following the specific procedures outlined below. Microscopic analysis for morphological evaluation was conducted using a Nikon Eclipse E200 LED Binocular Microscope (#846347, Japan) for high-clarity imaging, a Laxco Seba P4BP Digital Phase Contrast Microscope (USA, Product #C1422–8240–0039) to enhance contrast in transparent specimens, and a BZ-X Series All-in-One Fluorescence Microscope (Keyence, Japan) for fluorescence-based visualization. Additionally, a Chemat KW-4A Spin Coater (USA) was utilized to achieve uniform thin film deposition on slides by controlling rotation speeds and processing times.

Sample Source, Preparation, and Storage

Aronia mitschurinii plants were cultivated at the University of Maryland’s Wye Research and Education Center (WyeREC) in Queenstown, MD. The cultivation took place in Mattapex-Butlertown silt loam soil, characterized by a 0–2% slope and a pH of 6.1. Fruit samples for this study were randomly collected from harvests between 2020 and 2024. Before processing, the fruit samples were stored at −25 °C. Before massing the initial fruit samples, they were defrosted for approximately 3 min. The fruits were then ground using a mortar and pestle, followed by rapid filtration using a vacuum filtration apparatus. The processed samples were stored in Eppendorf vials at −25 °C for further analysis, with annual renewal of samples.

Principle of Crop Selection for Antifouling Protection

Biofilm formation and its subsequent fouling are closely linked with oxidative stress. , Initial radical-catalyzed polymerization and bacteria precipitation play a crucial role in the early stages of biofilm development, influencing the later phases of fouling. To combat these effects, it is essential to neutralize the key constituents necessary for biofilm establishment. Our approach to preventing biofilm formation involves incorporating antiseptic and antibacterial components, alongside antioxidants derived from plants, into the biofilm environment. Antiseptics and antibacterials target bacterial settlement, while antioxidants counteract oxidative stress, thereby inhibiting biofilm growth. These active components are to be encapsulated in biocompatible polymer, in our case poly­(methyl methacrylate) (PMMA), which is currently used as a base for most of paints and would be the easiest for the future technology transfer.

The superfruit Aronia mitschurinii has been selected as a model crop for the development of the concept and methodology because of its extremely high content of phytochemicals of interest – phenolic antioxidants, for which aronia is the crop with the highest known content, and essential oils, which Aronia juice has 7% wt. A. mitschurinii, the cultivar of the wild bread A. melanocarpa commonly known as black chokeberry, is a berry native to the Eastern Shore of Maryland, renowned for its high anthocyanins, flavonoids, and polyphenols content. Aronia berries exhibit the highest antioxidant activity among berries, as measured in their ORAC (Oxygen Radical Absorption Capacity), which is attributed to their high anthocyanin concentration. Anthocyanins, responsible for the purple, blue, and red hues of fruits, are sensitive to light, pH, oxygen, and temperature. To maintain their effective flavylium cation form, anthocyanins require storage at temperatures below 50 °C to prevent degradation.

For the antifouling tests, the aronia harvest was grown at the Wye Research and Education Center facility in Queenstown, MD, from 6-year-old bushes using medium irrigation with a precise irrigation system in place. Such growing conditions, along with 3 g/year/bush of organic nitrogen fertilizer and organic pest treatment for Japanese beetles, were found to be the best way to grow the harvest with a high content of antioxidants. Aronia was harvested at the peak of its antioxidant content and stored frozen in a lab freezer at −25 °C after being washed and fully dried of water in vacuum bags sealed and vacuumed. The processing included the preparation of five different concentrations of extract in ethyl acetate from defrosted berries, as described in the Materials and Methods section. The extracts were used fresh for sample preparation.

Antioxidant Extraction

The extract of the processed plant material (masticated plant or freeze-dried powder) was prepared by mixing in a 250 mL round-bottom flask 10g of plant material and 100 mL of solvent (solvent varies – please see the results part). A magnetic stirrer was added, stirring the mixture at 1000 rpm to prevent settling. The joints were greased with fluorinated grease to avoid mixing of grease with plant extract, and the flask was connected to the extraction apparatus equipped with a condenser to prevent solvent evaporation during the extraction. Cold water was circulated through the condensers, and a heated silicone oil bath maintained the temperature at 50 °C using a thermoregulatory probe. The extraction process was carried out for 72 h. After extraction, the solution was cooled and vacuum filtered. The extract volume was then reduced to one-quarter using a rotary evaporator, resulting in the “1x” aliquot. This aliquot was subsequently diluted to 2x (i.e., diluted twice), 4x (i.e., diluted 4 times), 8x (i.e., diluted 8 times), and 16x (i.e., diluted 16 times) concentrations with the respective solvents.

All five aliquots were stored in 1.5 mL microcentrifuge tubes and frozen for subsequent phytochemical characterization. Each aliquot was characterized as a triplicate for total anthocyanins, total polyphenols, total flavonoids, and total tannins, with mean values and standard errors calculated. Results are available in the Supporting Information file as figures S5–S14. The varying concentrations of the extracts are needed later to demonstrate a correlation between the concentration and antifouling capabilities.

Water Samples for Antifouling Tests

Water Samples for Antifouling Tests. Water samples were collected from multiple GPS-controlled locations, including Chincoteague Oceanside for saltwater tanks, and Pocomoke River for the freshwater tanks. Basic water parameters, including pH, temperature, salinity, oxidation–reduction potential, dissolved oxygen, and conductivity, were measured using an AZ Instruments water quality meter (model AZ-86031).

After collecting, the water samples were heated and aerated to preserve the bacterial microbial communities. The samples then underwent filtration to assess sediment content, which is believed to influence biofilm formation. The sediment content was determined by comparing the weight of the filter papers before and after baking, providing a measure of the precipitation in the various water sample locations. Water samples were stored for no more than 3 weeks to ensure water quality and stability of water parameters.

Water Sample Processing

Upon collecting, water samples were transported to the laboratory where they were processed as follows: a portion of each sample was immediately filtered to remove mechanical particles and sediment using vacuum filtration and paper filters with large pore sizes. This filtration was essential to prevent solid sediment from settling on the surfaces during the antifouling tests, which could interfere with the subsequent surface analysis. The remaining water was stored in polyethylene containers in the lab freezer for later use. Each 100 mL of water was filtered separately, and gravimetric analysis was performed to quantify the amount of sediment per liter of water at each location.

Microbiome Analysis

An Aquabiomics DNA testing kit for aquariums was used to assess the bacteria in both tanks, both before and after antifouling tests, as well as on the slides after the antifouling tests. For the analysis of slides, the slides were rinsed with RODI as described below and allowed to dry in a dust-free environment to prevent outside contamination. After the slides were dry, they were swabbed and sent for analysis at an Aquabiomics facility.

Gas Chromatography–Mass Spectrometry (GC-MS)

A gas chromatography–mass spectrometer (GC-MS) was employed to analyze Aronia samples and identify essential oils and terpenes within each extract. Specifically, it aids in identifying antimicrobial and anticarcinogenic compounds that may influence the inhibition of biofilm formation, particularly in the second and fourth steps, where microorganisms such as bacteria contribute to biofilm maturation.

The analysis used a Shimadzu GC-MS QP2010 with a standard VF-5 fused silica capillary column (30 m × 0.25 mm, i.e., film thickness 0.25 μm). Helium was used as the carrier gas. Chromatographic conditions included a flow rate of 1 mL/min at 8.2317 psi and an injector temperature of 240 °C. Chromatograms were generated to identify compounds with a match similarity of more than 98% based on NIST database comparisons.

The phytochemical screening of the Aronia samples followed the methods described by Kela et al.

Total Anthocyanins Content of Extracts

Anthocyanin pigment concentration was measured following the method of Lee et al. Two sample dilutions were prepared with 0.025 M potassium chloride (pH 1) and 0.4 M sodium acetate (pH 4.5). The potassium chloride solution highlights anthocyanins in their oxonium and flavylium forms, while the sodium acetate solution corrects for the carbinol form, which is colorless.

Absorbance was recorded at 520 and 700 nm using an Agilent Cary 60 UV–vis spectrophotometer. The anthocyanin pigment content (APC), expressed as cyanidin-3-glucoside equivalents (C3GE) in mg/L, was calculated using the formula:

$$\mathrm{APC}=(\mathrm{A}\times \mathrm{MW}\times \mathrm{DF}\times 103)/(\epsilon \times \mathrm{l})$$

MW is the molecular weight of cyanidin-3-glucoside (449.2 g/mol), DF is the dilution factor (200X), l is the path length in cm, and ε is the molar extinction coefficient for cyanidin-3-glucoside (26,900 L/(mol·cm)). The factor 10∧3 converts the result from grams to milligrams.

Total Flavonoids Content of Extracts

Flavonoid content was determined using the method of Chang et al. Quercetin standards were prepared in 95% ethanol at 0, 30, 60, 90, 120, and 150 g/mL concentrations. Experimental samples were prepared by mixing 125 μL of the sample with 375 μL of deionized water. To this mixture, 25 μL of 10% (w/v) AlCl₃ and 25 μL of 0.4 M potassium acetate were added, followed by 950 μL of deionized water. The samples were vortexed and incubated at room temperature for 30 min.

Absorbance was measured at 405 nm using an Agilent Cary 60 UV–vis spectrophotometer. A calibration curve was created from the standards to calculate the flavonoid concentration in the experimental samples. Results were expressed as μg quercetin equivalents (QE) per mL of juice. Each sample was analyzed in triplicate.

Total Polyphenols

Total polyphenol content was measured following the method described by Singleton et al. Gallic acid standards were prepared in 80% ethanol/20% deionized water at 0, 30, 60, 90, 120, and 150 μg/mL concentrations. For the experimental samples, 198 μL of distilled water was mixed with 2 μL of Aronia juice, followed by the addition of 200 μL of each gallic acid standard.

1250 μL of Folin’s Reagent was added to each sample, and the mixture was vortexed. The samples were allowed to sit at room temperature for 5 min, and then 1500 μL of 7% (w/v) Na₂CO₃ solution was added. The samples were incubated at 40 °C for 15 min, then cooled in a refrigerator for 5 min before measuring absorbance.

Absorbance was recorded at 750 nm using an Agilent Cary 60 UV–vis spectrophotometer. A calibration curve from the standards was used to determine the concentration of polyphenols in experimental samples. Results were expressed as mg gallic acid equivalents per liter of juice and mg gallic acid equivalents per gram. Each sample was analyzed in triplicate.

Total Tannins

Total tannin content was measured using the method described by Antoine et al. The following reagents were prepared: 0.2 M sodium hydroxide (NaOH), 0.01 M Tetrazolium Blue Chloride, 1.5 g/L tannic acid, and 80% ethanol in water. For calibration, a 1.5 g/L tannic acid stock solution was prepared by dissolving 0.15 g of tannic acid in 10 mL of ethanol and then diluting it to 100 mL with distilled water. From this solution, standards ranging from 50 to 500 mg/L were prepared.

Analysis was conducted in triplicate. 300 μL of the sample was added to each glass vial, followed by 100 μL of 0.01 M Tetrazolium Blue Chloride, 100 μL of 0.2 M NaOH, and the mixture was briefly vortexed. Subsequently, 1500 μL of 80% ethanol was added. Absorbance was measured at 527 nm using an Agilent Cary 60 UV–vis spectrophotometer.

Developing the Sample Preparation Technology for Antifouling Tests

Polymer Encapsulation

Encapsulation involves entrapping active compounds within a polymeric matrix. The primary goal is to create a controlled-release system that gradually releases the antimicrobial and antioxidant agents over time, thereby preventing microbial precipitation and oxidation of surfaces. The encapsulation process typically begins with selecting suitable antimicrobial and antioxidant extracts, which may be derived from natural sources such as plants, fruits, or marine organisms. These extracts are then incorporated into synthetic polymers through various encapsulation techniques, such as microencapsulation or nanoparticle encapsulation.

During encapsulation, the active compounds are entrapped within the polymer matrix, forming a barrier that protects them from degradation and premature release. This allows for the sustained release of antimicrobial and antioxidant agents, ensuring prolonged protection against microbial growth and oxidative damage.

Blending with Polymers: Determination of the Optimal Ratio of Polymer/Solvent and Coating Techniques

The correct polymer concentration is first obtained before the A. mitschurinii extract is added to the polymer. The correct polymer concentration is obtained by adding increasing amounts of poly­(methyl methacrylate) (PMMA) to the solvent, which is either 10 mL of 50% Ethyl acetate and 50% acetone or 100% chloroform. Solvent is added to 10 mL beakers and heated at 50 °C. PMMA is added and allowed to dissolve. The various amounts of PMMA added into the solvent created a range of viable concentrations from 50 mg/mL to 200 mg/mL. During determination, the addition of PMMA ceased when solubility issues arose. PMMA was fully dissolved at 100 mg/mL for ethyl acetate/acetone and 200 mg/mL for chloroform. The solvent mixture is pipetted onto coverslips fixed onto microscopic slides, ensuring to coat the coverslip completely. Different concentrations resulted in volumes dispensed onto slides that varied between 300 and 500 μL. The solvent is replaced due to evaporation and to maintain concentration after coating the slides. Once dried, the slides are observed visually before using phase contrast microscopy to analyze surface characteristics.

Prepared slides are analyzed through the phase contrast microscope to visualize microscopic cracks and bubbles, potential areas for biofouling to occur, and to ease future comparison of different samples after the antifouling tests. Observations were completed at 4x, 20x, and 40× magnifications. After imaging using phase contrast, slides showing poor characteristics (visually) are omitted from subsequent analysis via microscopy. Triplicates were performed with each slide at three different magnifications.

Spin Coating Samples

Spin coating was used to evenly spread the antifouling formulation over the surface of the slides. We used microscope slides to ease the analysis of samples after the tests. The Chemat KW-4A Spin Coater was used to produce these thin films. This device rotates samples with a liquid blend at high speeds, spreading the blend evenly over the substrate before drying. The spin coater offers two-speed settings in 1000 rpm increments. It allows for adjustments in initial and secondary spinning times, with a maximum of 18 s for the initial cycle and 60 s for the secondary cycle. After experimenting, it was found that 2000 rpm produced the most uniform films. The optimal procedure involved depositing the solution onto the substrate once the spin coater reached 2000 rpm, which takes about 3 s. The remaining time allows the solvent to evaporate completely. Using two different speeds did not improve the film characteristics, so it was decided that a single speed of 2000 rpm for 10 s would be utilized, with polymer deposition occurring between 3 and 5 s.

Poly­(methyl methacrylate) (PMMA) Preparation for Samples

PolyScience-branded PMMA (mW 75,000), stored at room temperature in the antifouling lab and obtained from Sigma-Aldrich, creates polymeric blends. The polymer is added to a solvent/extract mixture and heated at 50 °C until fully dissolved. After dissolution, 1000 μL of the viscous polymer blend is dispensed onto a labeled coverslip adhered to a glass microscope slide using cyanoacrylate. The slide is then spun at 2300 rpm using the spin coater to ensure even spreading and drying of the coating.

Addressing Delamination

Water absorption can lead to delamination of polymeric layers, as shown in Figure . The interaction of water with the polymer causes volume changes and weakens intermolecular forces, leading to delamination.

2.

Delaminated coverslips.

To address this, various methods were tested, including oxygen plasma exposure and sulfuric acid immersion, but these did not effectively enhance adhesion. The most successful approach involved using Cyanoacrylate superglue to affix Superslip No. 1 coverslips onto Fisherbrand Superfrost Disposable Microscope Slides. Applying glue to the corners of the coverslip and pressing it onto the slide improved adhesion and minimized delamination. The vulnerability of polymeric slides in water represents a major limitation, as the intended product would be used on submerged objects. However, for the purpose of observing bacterial growth in a laboratory, glass slides were used. Future testing using metal slides is recommended, as this better models what would happen at the bottom of a ship.

Determination of Thickness and Flatness Using Poly­(methyl methacrylate) (PMMA)

The thickness of PMMA films is influenced by polymer concentration, spinning rate, and solvent type. Higher concentrations typically result in less uniform films, while solvents with low vapor pressure produce more uniform surfaces and thinner films. Lower concentrations are recommended for better uniformity, as per Mohajerani et al. Film thickness and flatness were measured using the Bruker DektakXT (Germany) stylus profiler, where measurements are made by placing and adjusting the R (red reference) and M (green measurement) cursors.

Determination of Optimal Polymer Concentration Using Adhesion Test

An adhesion test was performed to find the optimal polymer concentration for coatings. Glass slides were cleaned with acetone, coated with various polymer concentrations after adding Superslip No. 1, and allowed to dry at room temperature for 2 h. The slides were then cured at 40 °C for 24 h. Coated slides were submerged in deionized water at room temperature for 24 h for the adhesion test. After immersion, slides were inspected for delamination; those without delamination passed the test, indicating good adhesion.

Short-Term Antifouling Test Procedure

The antifouling test involves a custom-built 3D-printed holder to secure glass slides coated with poly­(methyl methacrylate) (PMMA). This holder ensures precise alignment and stability during testing. The slides are immersed in a controlled environment, and the exposure times are carefully timed to ensure consistency across tests. Imaging is conducted using phase-contrast microscopy and laser microscopy to evaluate the extent of biofouling.

Phase-contrast microscopy is used to enhance the visibility of transparent biological organisms, using differences in refractive index for contrast. A Laxco Seba P4BP Digital Microscope System (C1422–8240–0039) is employed for imaging at 10× magnification, capturing images from five distinct points on each slide. These images are analyzed for particle count, area, and size to provide both qualitative and quantitative data. Additionally, laser microscopy (Keyence VK-X3000 Series 3D Surface Profiler) measures surface profiles, film thickness, and roughness with high precision. The laser’s confocal method scans the sample surface, providing 3D topographical maps that highlight surface changes due to fouling.

Post-test Sample Treatment

Post-test cleaning of the slides is essential for accurate results. The following steps were applied:

• 1. RODI Washes: water is ultrapure water produced through reverse osmosis filtration followed by deionization to remove residual salts, minerals, and impurities. In antifouling tests, slides are washed three times with RODI water to eliminate nonadhered particles and contaminants that could interfere with polymer adhesion, biofouling analysis, or surface characterization. This ensures consistency, prevents unwanted chemical interactions, and maintains the accuracy of experimental results.
• 2. Drying: After washing, slides were dried in a dust-free environment to prevent contamination from airborne dust particles.

These procedures are crucial for maintaining the integrity and reliability of the antifouling test results.

Antifouling Post-test Analysis Methodology

Phase-Contrast Microscopy

Phase-contrast microscopy was used to enhance the visibility of transparent biological specimens by exploiting differences in refractive index. This method, which improves contrast and detail, was performed using a Laxco Seba P4BP Digital Microscope System (#C1422–8240–0039), Bothell, Washington, USA. The microscope allowed detailed examination at five distinct points on polymer-coated slides, capturing images at 10× magnification. Height, Width, and Area measurements were included to ensure accurate and reproducible findings. This technique facilitated both qualitative and quantitative analysis of the slides.

Profilometry

The Bruker (Germany) DektakXT stylus profiler system measures surface texture by moving a diamond-tipped stylus over the sample. The stylus is connected to a Linear Variable Differential Transformer (LVDT), which converts surface variations into electrical signals. These signals are processed into a digital format for display and analysis. The Vision64 software calculates and displays results for various surface parameters, allowing detailed characterization of the sample profile.

Methylene Blue Solution

A final concentration of 0.01% of methylene blue was used to stain slides.

Laser Microscopy

The Keyence laser profiler (3D laser surface analyzer) was used to measure various film thickness parameters, including average thickness, maximum and minimum thickness, standard deviation, three times the standard deviation (3 sigma), start and end film positions, horizontal distance, and correction coefficient. The 3-sigma value represents a statistical range encompassing 99.73% of data points in a normal distribution, helping to assess variability in film thickness measurements. To determine film thickness, the average thickness of control samples (containing only polymer and solvent) was subtracted from the average thickness of treated samples.

The scheme of profilometry experiments with a laser microscope is presented in Figure at 10× magnification. The image to its right shows the three cross-sectional profile lines captured using the 2-point profile tool. The typical cross-sectional measurement data is captured for these three points and averaged. Each diluted sample is measured in this triplicate configuration. Each measurement also produces a 3D profile imager in color (left).

3.

Example of the scheme for film thickness cross-sectional profilometry measurement scan of a blank sample. A 3D profile in color (left) and a cross-sectional profile map (right) are shown.

Image Analysis Using ImageJ Software

ImageJ, an image processing software, was employed to analyze phase-contrast images, including particle counting and area measurements. The process involved converting images to grayscale, applying thresholding for segmentation, and using the “Analyze Particles” tool for quantification. Images were calibrated and preprocessed using ’Image > Scale > Set Scale,’ ’Process > Subtract Background,’ and ’Process > Enhance Contrast.’ Thresholding (’Image > Adjust > Threshold’) isolated key features, and particle analysis was conducted with ’Analyze > Analyze Particles’. The ROI (Region of Interest)­Manager (’Analyze > Tools > ROI Manager’) was used to focus on specific areas. ROI refers to a selected area within an image that is analyzed separately to extract relevant data. In this study, the ROI Manager in ImageJ/Fiji (’Analyze > Tools > ROI Manager’) was used to define and focus on specific regions of interest for analysis. The extracted data was exported using ’Results > Show Results,’ and Batch Processing (’Process > Batch > Macro···’) was applied to automate the handling of multiple images. To ensure accuracy, measurements were averaged across five observation points. This methodology provided a comprehensive analysis of the antifouling properties of the treated glass slides.

Statistical Analysis

Each experiment was repeated three times, and each sample for individual measurement was performed three times. The data was then analyzed statistically for Mean ± Standard Error, t test, and one-way analysis of variance (ANOVA) within the 95% confidence limit. The values of p < 0.05 were considered significant.

Results and Discussion

Locations and Analysis of Water Samples for Antifouling Tests

Water samples were collected from six distinct locations, encompassing a variety of environments: ocean, bay, river, and standing ponds, including fresh and saltwater. The details regarding these sampling locations and the basic water parameters measured with a portable water probe are summarized in Table .

1. Water Sampling Locations and Water Parameters for Antifouling Tests.

Location	GPS Location	pH	Temperature (°C)	Salinity (ppt)	Dissolved Oxygen(mg/L)
Chincoteague Oceanside	37.0238° N, 74.9989° W	8.12	17.9	26.8	117.6
Pocomoke River	38.0765° N, 75.5706° W	7.62	15.5	0.08	119.4

In order to keep any mechanical debris from effecting the slides, all water samples were filtered as described in the materials and methods. During this procedure, the weight of sediment was calculated in order to compare the water types. The results of this analysis are detailed in Table .

2. Determination of Sediment Amount in Water Samples.

Sample Location	Replicates	Water Volume, mL	Sediment Weight, g	Sediment conc,g/mL	Average Conc.,g/mL	Standard Deviation
Chincoteague Oceanside	1	100	0.024	0.00024	0.02733	±2.03 × 10–05
	2	100	0.027	0.00027
	3	100	0.031	0.00031
Pocomoke River	1	100	0.023	0.00023	0.02300	±1.92 × 10–20
	2	100	0.023	0.00023
	3	100	0.023	0.00023

Microbiome Community Analysis

DNA analysis was performed on both tanks and their slides to understand the impact of different microbes on the antifouling extract. Figure shows the abundance of bacteria in the Chincoteague water samples. The samples were found to have 90 different types of microbes and 36 different families. The most prevalent species were Cellvibionaceae, Flavobacteriaceae, Hyphomonadaceae, Hyphomonadaceae, Magnetospiraceae, Methyloligellaceae, Micavibrionaceae, NRL2, OM190, Pedosphaeraceae, Porticoccaceae, Rhondobacteraceae, Terasakiellaceae, and uncultured.

4.

Bacterial analysis of Chincoteague water.

Figure shows the bacterial analysis of slides developed in Chincoteague water. There was significantly more diversity, with 116 different microbes and 55 more families. For this sample, families Yersiniaceae (22.529%) and Rhodobacteraceae (5.959%), were the more prevalent species. These results indicate that the antifouling coating may be a better environment for bacterial growth, whether that came from outside contamination during the coating process or these communities developed during the antifouling test.

5.

Bacterial analysis of slides in Chincoteague water.

Figure represents the bacterial analysis of the Pocomoke water during the antifouling tests. This water was more diverse than the saltwater samples, with 152 microbes and 50 different families. The most dominant families are Alteromonadaceae, Burkholderiaceae, Candidatus_Kaiserbacteria, Caulobacteraceae, Cellvibrionaceae, Chloroplast, Clade_III, cvE6, Fimbriimonadaceae, Flavobacteriaceae, Holosporaceae, Hyphomicrobiaceae, Legionellaceae, Moraxellaceae, Parcubacteria, Rhodobacteraceae, Saccharimonadales, Simkaniaceae, and uncultured.

6.

Bacterial analysis of Pocomoke water.

Figure shows the diversity of the swabbed slide developed in the Pocomoke River tank. This sample, unlike the Chincoteague Sample, showed less diversity than the water sample. In the swabbed sample, there were 103 different microbes with 50 families. The most dominant families include Spongiibacteraceae (29.957%) and Solimonadaceae (17.903%). These results may indicate that the bacteria are more likely to thrive in the water than on the substrate.

7.

Bacterial analysis of swabbed slides in Pocomoke River.

Analyses of Aronia Extracts Used for Antifouling Formulations

A. mitschurinii was processed by crushing and extracting with ethyl acetate at 50 °C for 72 h. This solvent was chosen for its ability to dissolve antioxidants while maintaining reasonable poly­(methyl methacrylate) (PMMA) solubility at temperatures not exceeding 50 °C to prevent aronia’s antioxidant degradation. We also tested extraction from freeze-dried Aronia berries, which may offer advantages for industrial use due to better storage stability and preserved antioxidant content. Once prepared, all aronia extracts were diluted to have five concentrations of the extract. This is necessary to prove that the potential antifouling effect is attributed to the phytochemicals of aronia and not to the solvent or polymer in antifouling formulations. All prepared extracts were fully phytochemically characterized, as described below.

Phytochemical screening is a qualitative method only performed for the highest extract concentration. The screening for Aronia berries reveals the presence of flavonoids, glucosides, phytosterols, polyphenols, essential oils, tannins, and carbohydrates.

Table shows the values of polyphenols, tannins, anthocyanins, and flavonoids in aronia mitschurinii samples extracted in ethyl acetate. Since it has been observed that phytochemicals degrade over time when exposed to heat and or mechanical stirring, the phytochemicals were measured per dilution. The average value of these phytochemicals and their standard deviation per dilution can be found in the supplemental file as figures S1–S10, along with the values for acetone and chloroform extracts.

3. Concentration of Phytochemicals in A. Mitschurinii Extract (Ethyl Acetate).

Phytochemical	Concentration
Anthocyanins	64 (mg C3G/mL)
Polyphenols	8.56 (mg GAE/g)
Flavonoids (mg QE/g)	2.11 (mg QE/g)
Tannins	4.98 (mg TAE/g)

GC-MS Analysis of Aronia Extract

Figure presents the chromatograph A. mitschurinii, analyzed using Gas Chromatography–Mass Spectrometry (GC-MS). The analysis revealed several notable antibacterial oils, including Neophytadiene, Ethyl Linoleate, Eicosane (Icosane), Oleamide, Ethyl Oleate, Palmitamide, Tetratetracontane, and Dibutoxyethyl phthalate. These compounds are detailed in Table , which provides their generic names, IUPAC names, chemical formulas, classes, and functions. This data highlights the diverse chemical profile of A. mitschurinii and its potential relevance in antifouling applications.

8.

GC-MS results for A. mitschurinii.

4. Aronia Phytochemicals Detected by GC-MS and Their Properties.

Generic Name	IUPAC Name	Formula	Class	Role/Function
Neophytadiene	7,11,15-trimethyl-3-methylenehexadec-1-ene	C15H24	Hydrocarbon, specifically a diterpene	Neophytadiene is a naturally occurring hydrocarbon found in certain plants. It has been studied for its potential biological activities, including anti-inflammatory and antioxidant properties.
Ethyl Linoleate	ethyl(9Z,12Z)-octadeca-9,12-dienoate	C20H36O2	Fattyacid ester.	Ethyl linoleate is an ester formed from linoleic acid, an essential omega-6 fatty acid.
Eicosane (Icosane)		C20H42	Hydrocarbon, specifically an alkane	It has no known direct role in human metabolism or nutrition. Industrially, it is used as a lubricant, solvent, and in the synthesis of other chemicals.
Oleamide	(Z)-octadec-8-enamide	C18H35NO	Fatty acid primary amide.	Oleamide is an endogenous fatty acid primary amide found in the brain and other tissues
Ethyl Oleate	Ethyl(9Z)-9-octadecenoate	C20H38O2	Fatty acid ester	Ethyl oleate is an ester formed from oleic acid, a monounsaturated omega-9 fatty acid. Oleic acid is found in various oils.
Palmitamide	Hexadecanamide	C16H33NO	Fatty acid primary amide	Palmitamide, also known as hexadecanamide. It has been studied for its potential anti-inflammatory and neuroprotective effects.
Palatinol	Dibutoxyethyl phthalate	C16H22O4	Phthalate ester	In human metabolism and nutrition, it is not naturally occurring and is primarily encountered through environmental exposure. Efforts are made to minimize exposure to this compound due to its potential health risks.
Tetratetracontane		C44H90	Hydrocarbon, specifically an alkane	Tetratetracontane is a straight-chain hydrocarbon belonging to the alkane family. It is a component of certain waxes and mineral oils.

Post-test Analyses of Samples in Short-Term Tests

Phase-contrast images of polymer-coated slides with extracts are presented in Table . Five observation points were captured for each dilution. Parameters include Width, Height, Area, the corresponding label, and scale, with each image captured at 10× magnification. To accurately estimate the amount of precipitation per slide, the following formula was used: Precipitation per Slide (count/mm)=(Total Precipitation Count in Observation Area/Area of Observation Point (μm/1000))*×Total Area of Slide (mm).

5. Phase Contrast Microscopy of Aronia Formulation Slides after Submerging to Water Samples for Three-Week Tests.

The total precipitation count was determined using ImageJ, and the area of the observation point was measured by the phase contrast microscope. The total area of the slide was measured directly. The result is expressed in counts per mm.

The first row of Table displays the ″control″ slides, which were examined before being submerged in their respective water samples. The first column shows the ″Blank″ slides, which underwent the antifouling test but contained no extract, only polymer and solvent. It is clearly seen that the most concentrated samples of protected slides exhibit significant inhibition of biofilm formation. The effect becomes weaker, and the concentration of aronia extract in formulations decreases. This effect can be clearly seen in both salt and freshwater. However, freshwater protection is stronger than saltwater protection. This aligns with the finding of fewer bacteria on the swabbed freshwater slides versus the swabbed saltwater slides. It is hypothesized that the saltwater environment may cause defects on the slides over time, allowing the bacteria to thrive more in the exposed polymer than in the water.

In both cases, Blank slides demonstrate the highest precipitation. All images from Table were then quantified at five observation points using the ImageJ software, as described in the Materials and Methods section. These quantified results for this and all other samples can be found in the Supporting Information available for downloading with no charge as Tables S1–S4.

In both salt and fresh water, the most concentrated 1x sample performed the best and clearly demonstrated effective protection. More diluted samples do not protect as well but are much better than unprotected blank samples. The data suggest that in the future, it might be feasible to work with more concentrated extracts of even our aronia juice to increase the protective effect.

Results of the same series of samples, but in water stained with methylene blue, are presented in Table . The area colored blue indicates bacterial colonies inside the precipitated biofilm. Since such colonies are responsible for metabolites from which the biofilm is polymerized, the overall amount of such colonies indicates higher precipitation and lower antifouling protection. While this is an indirect observation of the number of precipitates, this method does not include occasional observation of mechanical defects on slides and mechanical particle precipitation

6. Phase Contrast Microscopy of Aronia Formulation Slides after Submerging into Water Samples with Methylene Blue for Three-Week Tests.

Table shows qualitative results consistent with our hypothesis. The freshwater and saltwater treated with methylene blue dye show significantly less precipitation with greater concentrations of aronia extract. Qualitatively, the control slides show comparable results to those in pure water samples, absent fluorescent dye. Increased cracks, bubbles, and higher amounts of precipitation were observed in samples with greater dilution.

When compared to other crops and solutions, aronia mitschurinii appears to be a comparable antifouling agent. Research on Curcuma longa demonstrated a 38.9% reduction in bacterial growth over 30 days when combined with tin paint; however, this inhibition dropped to 0% after 60 days. Long-term tests on a. mitschurinii are needed to see if it is truly comparable past the 3 week mark. Additionally, a study on extracts from Nerium oleander showed a barnacle density of 275.22, significantly lower than the control group, which had a mean barnacle density of 1499.

Laser Microscopy

The results for film thickness measurements of Aronia-based antifouling coatings in freshwater are presented with all raw data in the Supporting Information. The blank sample exhibited an average film thickness of 825.234 μm, with a maximum of 1944.339 μm and a minimum of 605.558 μm, serving as the baseline reference without antifouling treatment. In comparison, the 16x diluted Aronia sample showed a lower average thickness of 740.740 μm, with a maximum of 1824.302 μm and a minimum of 539.604 μm, indicating some reduction in film formation. The 1x diluted Aronia sample exhibited the most significant reduction, with an average film thickness of 643.727 μm, a maximum of 1726.995 μm, and a minimum of 602.334 μm. The progressive decrease in film thickness with increasing Aronia extract concentration suggests that higher antioxidant content enhances antifouling performance by limiting excessive film formation. These findings align with previous results in saltwater conditions, further supporting the potential of Aronia-based antifouling coatings.

The data from the surface film thickness measurements for Aronia in freshwater is presented in Table , while the data in saltwater is presented in Table . Table reveals the average precipitation levels across different experimental conditions, shedding light on the relationship between concentration levels and precipitation. The control, a slide with no coating that did not pass any tests, serves as the baseline and is considered to have zero precipitation. The rest of the slides have the same thickness as the control slide, plus precipitation. The Blank slide, which is coated by PMMA in solvent only and measured after the test, exhibits the highest average precipitation level at 155.3569 units. As the dilution of the extract in the antifouling formulation increases from 1x to 16x, there is a corresponding rise in average precipitation levels, confirming that the protective effect is due to the presence of aronia and is not related to PMMA and/or the solvent used.

7. 3D Surface Film Thickness for Aronia Freshwater Samples (μm).

	Slide with no coating/test	1x	2x	4x	8x	16x	Blank
Avg Precipitation	0	36.43343	38.54935	41.88955	134.1522	142.214	155.3569
Mean	609.9839	646.4173	648.5332	651.8734	744.1361	752.1979	765.3408
Standard Error	1.753652	4.409548	0.393797	3.355418	13.84652	7.831278	30.67738
Median	609.4633	643.727	648.715	652.0238	737.9618	748.6793	746.9243
Standard Deviation	3.037414	7.637561	0.682076	5.811754	23.98288	13.56417	53.13477
Sample Variance	9.225883	58.33234	0.465228	33.77649	575.1785	183.9867	2823.304
Skewness	0.748658	1.388447	–1.11396	–0.11634	1.081712	1.088762	1.37233
Range	6.007527	14.54701	1.327332	11.62059	46.75843	26.43494	101.3692
Minimum	607.2404	640.489	647.7787	645.988	723.844	740.7397	723.8644
Maximum	613.248	655.036	649.106	657.6086	770.6024	767.1747	825.2336
Sum	1829.952	1939.252	1945.6	1955.62	2232.408	2256.594	2296.022
Count	3	3	3	3	3	3	3

8. 3D Surface Film Thickness for Aronia Saltwater Samples (μm).

	Slide with no coating	1x	2x	4x	8x	16x	Blank
Avg Precipitation	0	4.499333	6.492333	8.218667	8.562	16.574	31.87567
Mean	609.982	614.4813	616.4743	618.2007	618.544	626.556	641.8577
Standard Error	1.752238854	1.978473	0.986933	2.704008	3.634102	4.937026	9.707703
Median	609.463	614.591	615.614	616.893	621.915	623.044	632.196
Standard Deviation	3.034966721	3.426816	1.709419	4.683479	6.294449	8.55118	16.81423
Sample Variance	9.211023	11.74307	2.922112	21.93497	39.62008	73.12267	282.7185
Skewness	0.747027038	–0.14386	1.691128	1.158489	–1.71876	1.536422	1.731992
Range	6.003	6.851	3.077	9.089	11.153	15.984	29.169
Minimum	607.24	611.001	615.366	614.31	611.282	620.32	632.104
Maximum	613.243	617.852	618.443	623.399	622.435	636.304	661.273
Sum	1829.946	1843.444	1849.423	1854.602	1855.632	1879.668	1925.573
Count	3	3	3	3	3	3	3

The data in saltwater (Table ) shows trends similar to the data in fresh water but with an overall lower level of precipitation; thus, the blank slide shows the highest average measurement at 31.87567 units, as compared to 155.3569 units in saltwater, serving as a reference point for comparison. As the dilution of Aronia extract escalates from 1x to 16x, a consistent upward trend in average measurements of average thickness is observed, indicating a clear correlation between concentration and the protective effect.

As previously reported, scanning electron microscopy (SEM) was also employed to image the surfaces of nonmetallic samples. The SEM images revealed that the blank samples coated by PMMA only displayed a flake-shaped diatom, likely of the genus Cyclotella and possibly the species Cyclotella. In contrast, the samples with 1x extract of Aronia encapsulated into PMMA do not show any evidence of diatom precipitation, indicating that the advanced processes of biofilm growth are suppressed in samples protected with Aronia extracts.

Conclusions

We have demonstrated the efficacy of extracts from Aronia mitschurinii encapsulated in PMMA as an effective formulation for antifouling protection. This proof of concept aims to provide a safer, more effective antifouling solution than previous solutions, such as tributyl tin. Future research on copepods will be conducted to observe the toxicity of PMMA.

The protective effects observed are primarily attributed to the extracts of aronia and not to PMMA or solvent, as evidenced by the concentration-dependent nature of protection, confirmed by phase contrast microscopy, laser 3D profiling, and observation of bacterial colonies stained with methylene blue. This finding highlights the crucial role of plant extracts rich in antioxidants and antibacterial compounds in combating fouling organisms. It opens the path to probing some other crops the lab is working with now.

Post-test analysis performed by different methods has shown that the higher the concentration of Aronia extract in the formulation, the stronger the protective effects. While some of our methods target overall precipitation, others target the bacteria inside the browsing biofilm. These results show that the number of bacteria also decreases with increased concentration of extracts in the formulations.

Our results reveal that the protective effect is more pronounced in freshwater environments than in saltwater, although significant protection is still evident under saltwater conditions. Moreover, the effectiveness of the protection remains consistent over time, showing robust performance after several weeks of testing. Long-term testing is still needed to determine how long this protective nature will protect slides, as this product would need to last in harsher environments over longer periods of time in order to be economically feasible for those purchasing it.

We have further set up experimental designs for longer-term studies, ranging from 2 to 6 months, to validate our formulations’ durability and effectiveness. These experiments are currently in progress and are expected to provide deeper insights into the long-term performance of our antifouling solutions.

Future directions in antifouling research with specialty crops will involve exploring various berries, superfruits, and medicinal herbs to enhance antifouling efficacy. Beyond that, isolated compounds will also be used in order to prepare for the future process of industrialization.

In conclusion, our research provides compelling evidence for the efficacy and practicality of utilizing plant extracts encapsulated into PMMA for antifouling protection.

Glossary

Abbreviations

AF

Antifouling

C3G

Cyanidin-3-Glucoside

DNA

Deoxyribonucleic Acid

DO

Dissolved Oxygen

EIS

Energy Dispersive X-ray Spectroscopy

EPS

Extracellular Polysaccharides

GAE

Gallic Acid Equivalents

KOH

Potassium Hydroxide

ORAC

Oxygen Radical Absorption Capacity

PLL-G-PEG

Poly­(l-lysine)-graft-poly­(ethylene glycol)

PMMA

Poly­(methyl methacrylate)

QE

Quercetin Equivalents

QSI

Quorum Sensing Inhibitor

RODI

Reverse Osmosis Deionized Water

SEM

Scanning Electron Microscopy

TDS

Total Dissolved Solids

UV–vis

Ultraviolet–Visible Spectrophotometry

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsomega.5c05826.

• Bacterial analysis of water and swabbed slides of Chincoteague and Pocomoke; all phytochemical screening results for anthocyanins, polyphenols, flavonoids and tannins per dilution; polymer encapsulation; imagej calculations; film thickness calculations, cross-sectional maps, and 3D profiles of aronia slides (PDF)

This work is supported by the Department of NAVY MSI grant and distinguished faculty fellowship no. N0014–21–1–2756, AFRI-EWD-REEU program, grant no. 2020–69018–30655, from the U.S. Department of Agriculture, National Institute of Food and Agriculture, the USDA-NIFA Evans Allen formula grant at the University of Maryland Eastern Shore, and the United States Environmental Protection Agency under assistance agreement 84058501 to the University of Maryland Eastern Shore. The contents of this document do not necessarily reflect the views and policies of the Environmental Protection Agency, nor does EPA endorse trade names or recommend the use of commercial products mentioned in this document. Any opinions, findings, conclusions, or recommendations expressed in this publication are those of the author(s). They should not be construed to represent any official USDA or U.S. Government determination or policy.

The authors declare no competing financial interest.