Growth and membrane stress responses in E. coli and Acinetobacter sp. upon exposure to functionalized polystyrene microplastics
Article information
Abstract
Microplastic pollution is increasingly recognized as a potential environmental stressor for microorganisms. This study aimed to explore how surface-functionalized polystyrene (PS) microplastics influence selected cellular-level responses in two Gram-negative bacteria, Escherichia coli and Acinetobacter sp., focusing on growth, viability, biofilm formation, and membrane-associated stress. Bacterial cultures were exposed to PS microplastics with three surface chemistries: non-functionalized PS, aminated PS (PS-NH2), and carboxylated PS (PS-COOH). Exposure to PS microplastic induced species- and surface chemistry-dependent alterations in bacterial responses. Compared to the control, non-functionalized PS reduced E. coli growth and viability to 74.8% and 61.3%, respectively, while Acinetobacter sp. showed reductions to 72.1% and 69.3% following PS exposure. Biofilm formation increased significantly to 143.2% in E. coli with PS, and to 207.2% and 190.7% in Acinetobacter sp. with PS and PS–COOH, respectively. Cytotoxicity assays revealed distinct stress patterns: in E. coli, PS exposure elevated MDA and LDH levels to 155.3% and 120.5% of control levels, respectively, while ROS levels remained near baseline (100.2%), indicating predominant membrane rupture and lipid peroxidation. In contrast, Acinetobacter sp. exhibited markedly elevated ROS (118.5% and 123.5%) and MDA (190.7% and 212.8%) levels upon exposure to PS and PS–COOH, while LDH remained comparable to the control, suggesting sublethal oxidative stress and membrane perturbation. These findings demonstrate that even chemically inert PS microplastics can trigger biologically significant responses in bacteria through surface-mediated mechanisms. The observed interspecies and inter-surface variability underscores the complexity of microplastic–microbe interactions and highlights the need for microbial-level assessments in evaluating the ecological risks of microplastic pollution.
Introduction
Polystyrene (PS) is primarily used in disposable products and packaging materials, and it readily enters both aquatic and terrestrial ecosystems through improper waste disposal and environmental fragmentation [1-4]. The widespread presence of PS debris has been documented across marine, freshwater, and soil environments [1-3]. As this waste undergoes fragmentation through physicochemical and biological processes, the resulting PS microplastics have emerged as a growing ecological concern [5].
Multicellular organisms, including animals and plants, can ingest PS microplastics, which may subsequently transfer through the food chain to higher trophic levels [6-8]. Exposure to these microplastics has been associated with a variety of toxic effects, such as reproductive cell damage, genetic mutations, inflammatory responses, and impaired immune or respiratory functions [9-12].
Toxicological studies have shown that microbial responses to PS microplastics depend largely on particle size [13-15]. Nanoplastics smaller than 100 nm are capable of entering bacterial cells and inducing DNA damage, membrane disruption, and oxidative stress [14, 16-18], while larger microplastics have generally been considered less impactful. However, some studies, including our previous work, indicate that certain micron-sized PS particles can also exert significant toxicity toward bacterial cells [13, 19].
In particular, we previously reported that bacterial growth was most strongly inhibited upon exposure to 1040 nm PS microplastics, compared to exposure to particles of 60, 220, 430, 700, 1700, and 2260 nm [13]. This exposure also led to increased biofilm formation and aggregation of PS particles within bacterial biofilms. These results suggest that physical interactions–particularly between the bacterial cell surface and PS microplastics–may play a key role in toxicity, potentially through membrane disruption [19].
These observations collectively suggest that the surface properties of PS microplastics, including their functional groups, could further influence their interactions with bacteria. In this context, we aimed to evaluate whether surface modifications–specifically, amine (PS-NH2) and carboxyl (PS-COOH) groups–alter bacterial responses compared to non-functionalized PS. Rather than assuming a unidirectional effect of surface chemistry, this study sought to assess how different surface functionalities affect bacterial growth, viability, biofilm formation, and membrane-associated stress in a species-specific manner.
In this study, we investigated the interactions between ~1060 nm-sized PS microplastics with different surface functional groups and two model bacterial species: Escherichia coli and Acinetobacter sp. Bacterial growth and viability were evaluated following exposure to PS, PS-NH2, and PS-COOH microplastics. To elucidate potential mechanisms of toxicity, we measured intracellular reactive oxygen species (ROS), lactate dehydrogenase (LDH) release, and malondialdehyde (MDA) accumulation. In addition, changes in membrane integrity were visualized to assess the physical effects of PS microplastic exposure.
Materials and Methods
Bacterial growth
The bacterial strains used in this study, E. coli (KCTC 2791) and Acinetobacter sp. (OQ402585), were selected due to their frequent occurrence on plastic waste surfaces in environmental settings and their well-documented ability to adhere to plastic materials and form biofilms [20-22]. These traits make them suitable model organisms for investigating interactions with functionalized microplastics. E. coli and Acinetobacter sp. were initially cultivated in nutrient broth medium (composed of 5 g of peptone and 3 g of beef extract per liter, MBcell, KisanBio Co., Seoul, Korea) at 37°C and 30°C, respectively, under agitation at 150 rpm for 16 h. Following this pre-cultivation, the optical density (OD₆₀₀) of each culture was adjusted to 1.0, and the corresponding CFU/mL–determined by serial dilution and plate counting–was approximately 8.5 ± 1.8 × 108 for E. coli and 8.1 ± 0.2 × 108 for Acinetobacter sp. Then, 500 μL aliquots were transferred into 125-mL Erlenmeyer flasks containing 49.4 mL of freshly prepared nutrient broth (2.5 g of peptone and 1.5 g of beef extract per liter, MBcell). Subsequently, 100 μL of three different commercially available PS microplastic suspensions were added to the flasks to achieve a final concentration of 50 mg/L: non-functionalized PS (diameter: 1060 nm, Spherotech Inc., Lake Forest, IL, USA), PS-NH2 (diameter: 1020 nm, Spherotech Inc.), and PS-COOH (diameter: 1040 nm, Spherotech Inc.). These microplastics were supplied as aqueous suspensions and were already well-dispersed in water; thus, no sonication was required. Prior to use, the suspensions were vortexed to ensure homogeneity. A set of flasks without microplastic addition served as the negative control group. Cultures were incubated at 37°C for E. coli and 30°C for Acinetobacter sp. with continuous shaking at 150 rpm for 9 h. The optical density at 600 nm (OD600) was using a UV-1800 spectrophotometer (Shimadzu Co., Kyoto, Japan). OD600 was measured at 1-h intervals throughout the incubation period for all experimental conditions, except for Acinetobacter sp. exposed to PS and PS–COOH, where measurements were taken at 0, 1, 4, 5, 6, 7, 8, and 9 h to better capture its extended lag phase and altered growth dynamics under microplastic exposure. The relative OD600 value (%) was calculated using the following equation:
where OD600 Sample and OD600 NC denote the optical density of the test and negative control samples at 6 h (E. coli) and 7 h (Acinetobacter sp.) respectively. All the experimental conditions were conducted in triplicate.
Bacterial cell viability
The bacterial cultures of E. coli and Acinetobacter sp. were harvested by centrifugation at 13200 × g for 10 min, followed by two washing steps using phosphate-buffered saline (PBS, pH 7.4, Sigma-Aldrich, St. Louis, MO, USA). The resulting cell pellets were resuspended in fresh PBS to adjust the final concentrations to approximately 8.5 ± 1.8 × 108 CFU/mL for E. coli and 8.1 ± 0.2 × 108 CFU/mL for Acinetobacter sp. A portion of each suspension was transferred into 125-mL Erlenmeyer flasks, to which PS, PS-NH2, and PS-COOH microplastics were individually added to a final concentration of 50 mg/L. Flasks containing only bacterial suspensions without microplastics served as the negative control group. The cultures were incubated at 37°C (E. coli) and 30°C (Acinetobacter sp.) for 24 h with shaking. These experimental conditions were used for ROS, LDH, and MDA assays unless otherwise specified.
Following the 24-h incubation, culture samples were serially diluted in 0.9% NaCl solution. A 100-μL aliquot of each diluted sample was spread onto nutrient agar plates (containing 5 g/L peptone, 3 g/L beef extract, and 15 g/L agar, Duchefa Biochemie, Haarlem, The Netherlands) and incubated at 37°C for E. coli and 30°C for Acinetobacter sp. for 16 h.
The number of viable cells was determined in CFU/mL by counting the colonies formed on the plates. Cell viability (%) was calculated according to the following formula:
where Viable cell number NC and Viable cell number Sample represent the CFU/mL of the negative control and microplastic-treated samples, respectively. All experiments were independently performed in triplicate.
ROS assay
The intracellular ROS levels were assessed after 24 h of exposure using 2’,7’-dichlorofluorescein diacetate (DCF-DA, Sigma-Aldrich) as previously described [13, 19]. Cultures treated with 0.05% hydrogen peroxide served as the positive control. After staining and incubation in the dark for 30 min, fluorescence was measured (excitation: 488 nm; emission: 535 nm) using a microplate spectrofluorometer (SpectraMax M2, Molecular Devices, San Jose, CA, USA).
The relative ROS level (%) was calculated by comparing the fluorescence intensity (F535) of each treated sample to that of the negative control using the following equation:
where F535 Sample and F535 NC represent the fluorescence intensities of the microplastic-treated sample and the untreated control, respectively.
LDH assay
Membrane damage was evaluated by quantifying LDH release using the CyQUANTTM LDH Cytotoxicity Assay Kit (Thermo Fisher Scientific, Waltham, MA, USA), following the same culture and incubation procedure as described in the Cell viability section and our previous report [19]. To determine maximum LDH release, cells (108 CFU/mL) were lysed on ice using a tip sonicator (VCX 750, Sonics & Materials, Inc., USA) in pulse mode (10 s on, 50 s off) for a total of 20 min. The experiment was conducted according to the manufacturer’s instructions. After the reaction, absorbance was measured at 490 nm and 680 nm.
The relative LDH release was calculated by comparing LDH activity in treated samples (LDH Sample) to that in negative controls (LDH NC), as shown below:
MDA assay
Lipid peroxidation was measured using an MDA assay kit (Sigma-Aldrich), following the method outlined in our previous study [19]. Following lysis and centrifugation, samples were reacted with trichloroacetic acid (Sigma-Aldrich) in glacial acetic acid (Duksan Co., Gyeonggi-do, Korea), incubated at 95°C, and cooled on ice. Absorbance was recorded at 532 nm, and MDA concentrations were determined using a calibration curve.
Relative MDA content (%) was calculated by comparing MDA levels in samples treated with microplastics (MDA Sample) to those of the negative control (MDA NC) using the following equation:
All assays were performed in quintuplicate.
Biofilm formation
A 500-μL aliquot of pre-cultured E. coli and Acinetobacter sp. cells was inoculated into 125-mL Erlenmeyer flasks containing fresh nutrient broth, followed by the addition of PS, PS-NH2, and PS-COOH microplastics to achieve a final concentration of 50 mg/L. Cultures containing bacterial cells without microplastics served as the negative control. Each flask contained a total volume of 25 mL and was incubated at 37°C (E. coli) or 30°C (Acinetobacter sp.) with shaking at 150 rpm for 24 h.
After incubation, 1 mL of each culture was centrifuged at 3381 × g for 3 min. The resulting cell pellets were stained with 0.1% crystal violet solution (Samchun Chemical Co. Ltd., Gyeonggi-do, Korea) for 15 min. Excess, unbound dye was removed by rinsing the cells twice with deionized (DI) water, followed by air-drying at room temperature for 24 h. To quantify biofilm formation, the bound crystal violet was solubilized by incubating the dried samples in 200 μL of 30% acetic acid (Duksan Co.) for 15 min. A 150-μL aliquot of the resulting solution was transferred into a 96-well microplate, and absorbance was measured at 550 nm using a spectrophotometer. Notably, control samples containing only PS microplastics (without bacterial inoculation) exhibited negligible absorbance at 550 nm, indicating that the measured signal primarily originated from biofilm-associated dye bound to bacterial cells.
The relative level of biofilm formation (%) was calculated by comparing the absorbance values of treated samples (OD550 Sample) to that of the negative control (OD550 NC) using the following equation:
Cell membrane integrity assay
A 500-μL aliquot of pre-cultured E. coli and Acinetobacter sp. was inoculated into nutrient broth, and microplastics–PS, PS-NH2, and PS-COOH–were added to achieve a final concentration of 50 mg/L. Cultures containing bacterial cells alone were prepared as the negative control. A total volume of 10 mL for each bacterial culture was incubated at 37°C (E. coli) and 30°C (Acinetobacter sp.) with agitation at 150 rpm for 24 h.
After incubation, 1 mL of each culture was transferred into 1.7-mL microcentrifuge tubes and centrifuged at 17177 × g for 3 min. The collected cells were washed twice with PBS and resuspended in fresh PBS. Each sample was stained by adding 500 μL of 50 μg/mL propidium iodide (PI, Sigma-Aldrich) and incubated in the dark at room temperature for 15 min.
Following staining, 20 μL of each sample was placed on a microscope slide and covered with a cover slip. The samples were then air-dried for 30 min and visualized under a fluorescence microscope (Axio Observer 3, Zeiss, Oberkochen, Germany) equipped with a rhodamine filter set (excitation: 551–556 nm; emission: 582–595 nm).
Hydrodynamic diameter, zeta potential, and FT-IR measurements
To evaluate the surface properties and colloidal stability of the microplastics, hydrodynamic diameters, zeta potentials, and Fourier-transform infrared (FT-IR) spectra of PS, PS-NH2, and PS-COOH were measured. All samples were loaded into flow cells (Otsuka Electronics, Osaka, Japan), and their surface charge and hydrodynamic diameter were determined using a zeta potential and particle size analyzer (ELS-Z2, Otsuka Electronics, Osaka, Japan) based on laser Doppler velocimetry. For FT-IR analysis, microplastic samples were first freeze-dried and then mixed with potassium bromide (KBr) to form pellets. The FT-IR spectra were obtained using a Nicolet iS50 spectrophotometer (Thermo Fisher Scientific).
T-test analysis
Statistical comparisons were performed using a t-test, to assess significant differences between the means of two groups–specifically, bacterial cells alone (negative control) and bacterial cells exposed to either PS, PS-NH2, and PS-COOH microplastics–on each measured variable. A p-value of less than 0.05 was considered statistically significant. All statistical analyses were conducted using the SigmaPlot software package (Systat Software, Inc., San Jose, CA, USA).
Results
Physicochemical characterization of PS microplastics
The hydrodynamic diameters of PS, PS-NH2, and PS-COOH microplastics were measured to be 1112.0 ± 27.6 nm, 1315.3 ± 58.3 nm, and 1167.1 ± 41.7 nm, respectively (Fig. 1a). Zeta potential analysis revealed surface charges of -30.2 ±0.7 mV for PS, -25.1 ± 0.4 mV for PS-NH2, and -49.7 ± 0.2 mV for PS-COOH (Fig. 1b), indicating differences in surface electrostatic properties depending on functionalization.
(a) Hydrodynamic diameters, (b) zeta potentials, and (c) FT-IR spectra of PS, PS-NH2, and PS-COOH. Dotted circles indicate characteristic peaks corresponding to functional groups in PS-NH2 and PS-COOH.
FT-IR spectroscopy further confirmed the presence of surface functional groups. In the spectrum of PS-NH2, a characteristic peak was observed in the range of 300–3500 cm-1, corresponding to N–H stretching vibrations (Fig. 1c) [14, 23]. For PS-COOH, a distinct peak at 1708 cm-1 was detected, attributed to C=O stretching vibrations [24, 25]. These findings confirm the successful surface modification of PS microplastics with amine and carboxyl functional groups.
Effects of PS microplastics with different functional groups on the bacterial growth and cell viability
The growth dynamics of bacterial cells were distinctly influenced by the addition of PS microplastics, with the degree of impact varying depending on the surface functional groups of the microplastics (Fig. 2).
(a), (c) Growth curves and (b), (d) relative OD600 value at 6 h and 7 h of incubation for E. coli and Acinetobacter sp. cells, respectively, after exposure to PS, PS-NH2, and PS-COOH microplastics.
For E. coli, a 2-h lag phase was consistently observed under all conditions, regardless of PS microplastic exposure, indicating that the initial adaptation period of the cells was not affected by the presence of PS particles (Fig. 2a). However, the maximum cell density, as measured by OD600 after 6 h of incubation, was notably reduced in the presence of PS microplastics. Specifically, the relative OD600 values decreased to 74.8% with non-functionalized PS, 86.6% with PS-NH2, and 88.5% with PS-COOH, compared to the control group without microplastics (Fig. 2b). These findings indicate that non-functionalized PS microplastics have the strongest inhibitory effect on E. coli growth, while surface-functionalized microplastics, particularly PS-NH2 and PS-COOH, exhibited a less pronounced impact.
In contrast, Acinetobacter sp. exhibited a more distinct growth suppression pattern upon exposure to PS microplastics (Fig. 2c). Under control conditions, Acinetobacter sp. displayed a 2-h lag phase followed by a steady increase in OD600 that plateaued at around 7 h of incubation. However, in the presence of PS microplastics, both the lag phase duration and final cell density were adversely affected. The addition of PS-NH2 microplastics extended the lag phase to 3 h, while PS and PS-COOH microplastics prolonged it further to 4 h.
Moreover, the maximum OD600 values were significantly reduced by microplastic treatment. The relative OD600 values dropped to 72.1% with PS and 74.5% with PS-COOH, while a milder reduction was observed with PS-NH2 (86.0%). These findings indicate that PS and PS-COOH microplastics exerted a greater inhibitory effect on bacterial proliferation than PS-NH2. Taken together, the prolonged lag phases and reduced growth yield demonstrate that exposure to PS microplastics–particularly those lacking or carrying carboxyl groups–can significantly impair the growth kinetics of Acinetobacter sp.
The cell viability results of E. coli and Acinetobacter sp. showed consistent with the patterns observed in the growth curve analysis (Fig. 3), further supporting the inhibitory effects of microplastic exposure. In both species, the addition of PS microplastics–regardless of surface functionalization–led to a measurable decrease in viable cell numbers compared to the control.
(a), (c) Viable cell counts and (b), (d) cell viability of E. coli and Acinetobacter sp. (hatched bars) after exposure to PS, PS-NH2, and PS-COOH microplastics. Asterisks (*) indicate statistically significant differences compared to the negative control (p< 0.05).
Among the tested particles, PS microplastics caused the greatest reduction in cell viability, decreasing to 61.3 ± 6.5% for E. coli and 69.3 ± 7.2% for Acinetobacter sp. (Figs. 3b and 3d). This reflects a strong cytotoxic effect induced by non-functionalized PS.
In comparison, exposure to surface-functionalized microplastics (PS-NH2 and PS-COOH) also resulted in reduced viability, but to a lesser extent. For E. coli, viability decreased to 77.4 ± 9.9% and 75.5 ± 8.2% in the presence of PS-NH2 and PS-COOH, respectively. Similarly, Acinetobacter sp. exhibited a reduction to 83.6 ± 5.2% (PS-NH2) and 72.8 ± 12.6% (PS-COOH). These results suggest that PS-COOH microplastics may have slightly more adverse effects on Acinetobacter sp. cells than PS-NH2, while in E. coli, both functionalized variants induced comparable reductions in viability.
Biofilm formation of bacteria in the presence of PS, PS-NH2, and PS-COOH microplastics
To further investigate bacterial responses to microplastic exposure beyond growth inhibition, biofilm formation was assessed as a key adaptive and survival strategy. Biofilm production in E. coli was selectively promoted depending on the surface chemistry of microplastics. When exposed to non-functionalized PS microplastics, E. coli exhibited a significant enhancement in biofilm formation, reaching 143.2 ± 13.8% relative to the negative control group (i.e., E. coli cells alone) (p< 0.05, Fig. 4a). In contrast, the presence of surface-functionalized microplastics, including aminated (PS-NH2, 90.5 ± 4.2%) and carboxylated (PS-COOH, 91.5 ± 5.6%) variants, resulted in only slight changes in biofilm levels, and these changes were not statistically significant compared to the control (p> 0.05). This result suggests that unmodified PS particles can strongly promote biofilm development, possibly due to their hydrophobic surface facilitating bacterial attachment.
Relative biofilm formation (%) of (a) E. coli and (b) Acinetobacter sp. (hatched bars) after exposure to PS, PS-NH2, and PS-COOH microplastics. Asterisks (*) indicate statistically significant differences compared to the negative control (p< 0.05).
A different pattern was observed for Acinetobacter sp. In this species, biofilm formation was significantly enhanced upon exposure to all three types of microplastics tested (p< 0.05, Fig. 4b). The relative increase in biofilm biomass reached 207.2 ± 25.9% with PS, 183.3 ± 4.8% with PS-NH2, and 190.7 ± 15.2% with PS-COOH, respectively. Although the biofilm-promoting effect of PS was slightly greater than those of the functionalized variants, the differences among treatments were not statistically significant (p> 0.05), implying that Acinetobacter sp. may interact broadly with various microplastic surfaces regardless of their surface chemistry.
Cytotoxicity assay of PS microplastics with different functional groups on bacteria
To elucidate the underlying mechanisms of growth and viability inhibition, cytotoxicity assays focusing on oxidative and membrane-related stress were conducted. The cytotoxic effects of PS microplastics with different surface functionalities (PS, PS- NH2, and PS-COOH) were evaluated based on oxidative stress markers, membrane damage indicators, and fluorescence-based membrane integrity imaging.
In E. coli, intracellular ROS levels remained statistically unchanged upon microplastic exposure, with relative values of 100.2 ± 5.2% (PS), 99.9 ± 3.1% (PS- NH2), and 103.5 ± 7.6% (PS-COOH) compared to the control (p> 0.05, Fig. 5a). However, significant increases were observed in both LDH release and MDA content, indicating membrane damage and lipid peroxidation, respectively. LDH levels rose to 120.5 ± 9.2%, 114.2 ± 10.5%, and 112.6 ± 9.0%, while MDA levels reached 155.3 ± 21.8% (PS), 136.9 ± 13.0% (PS- NH2), and 139.0 ± 18.1% (PS-COOH) (Figs. 5c, e). This suggests that membrane damage, rather than oxidative stress, plays a primary role in the growth and viability inhibition observed in E. coli (Figs. 2a–b, 3a–b).
(a), (b) Relative ROS intensities, (c), (d) LDH release levels, and (e), (f) MDA contents of E. coli and Acinetobacter sp. (hatched bars) after treatment with PS, PS-NH2, and PS-COOH microplastics. Asterisks (*) indicate statistically significant differences compared to the negative control (p< 0.05).
In contrast, Acinetobacter sp. showed a distinct cytotoxic response pattern. ROS levels were significantly elevated upon exposure to all three types of microplastics, with relative levels of 118.5 ± 5.8% (PS), 117.0 ± 3.4% (PS- NH2), and 123.5 ± 5.9% (PS-COOH) (Fig. 5b), indicating enhanced oxidative stress. LDH release remained relatively stable, showing values of 93.8% (PS and PS-NH2) and 92.2 ± 4.1% (PS-COOH), suggesting minimal membrane disruption (Fig. 5d). However, MDA content was markedly increased to 190.7 ± 6.3%, 205.8 ± 10.5%, and 212.8 ± 6.6% for PS, PS-NH2, and PS-COOH, respectively (Fig. 5f), indicating severe lipid peroxidation irrespective of surface chemistry. These results imply that Acinetobacter sp. cells are more sensitive to oxidative and lipid-related stress rather than membrane rupture.
To complement the biochemical assays and further confirm membrane integrity changes, PI fluorescence staining was conducted for both E. coli and Acinetobacter sp. (Fig. 6). In control conditions, both strains exhibited weak red fluorescence, indicating minimal membrane damage during incubation. However, upon exposure to PS, PS- NH2, and PS-COOH microplastics, a pronounced increase in red fluorescence intensity was observed in both bacterial species.
Optical and fluorescence images of (a) E. coli and (b) Acinetobacter sp. showing cell membrane integrity. Upper panels show bacterial cells without microplastics, and lower panels show cells in the presence of PS, PS-NH2, and PS-COOH microplastics. Arrows indicate 1020–1060 nm PS microplastics.
In E. coli, this result is in line with the significantly elevated LDH release and MDA content (Figs. 5c and e), suggesting that microplastic exposure leads to clear membrane rupture and lipid peroxidation.
In Acinetobacter sp., however, LDH release remained relatively unchanged across all treatments (Fig. 5d), while red fluorescence increased substantially. This apparent discrepancy can be attributed to the distinct detection sensitivities between the two assays. LDH release reflects severe membrane damage resulting in cytoplasmic leakage, whereas PI staining detects even subtle increases in membrane permeability. Therefore, the increased red fluorescence likely reflects sublethal membrane perturbation, not captured by the LDH assay. This interpretation is further supported by the significant rise in MDA content observed in Acinetobacter sp. cells, indicating that lipid peroxidation and oxidative membrane stress occurred, even in the absence of extensive membrane lysis.
Discussion
The present study demonstrates that exposure to PS microplastics can adversely affect bacterial responses, including growth, viability, biofilm formation, and stress-related markers (Figs. 2–6). These effects were dependent not only on the surface functional groups of the microplastics but also on the bacterial species involved, indicating a complex interplay between microplastic properties and microbial traits.
Previous studies have reported that microplastics can alter bacterial growth and community composition by acting as both physical stressors and attachment surfaces [26, 27]. Our findings align with these observations, showing that non-functionalized PS particles exerted the strongest inhibitory effects, particularly on E. coli (Figs. 2 and 3). This pronounced inhibition is likely due to the high hydrophobicity of non-functionalized PS, which can promote strong hydrophobic interactions with the bacterial membrane. These interactions may compromise membrane integrity, increase permeability, and disrupt ion gradients, ultimately leading to cellular stress and suppressed growth [28]. Interestingly, Acinetobacter sp. exhibited broader sensitivity to all microplastic types, with PS and PS–COOH causing more pronounced growth inhibition than PS–NH2. For PS–COOH, the presence of carboxyl groups–despite their negative charge–may allow hydrogen bonding and interactions with surface components such as membrane proteins or lipopolysaccharides, thereby facilitating close contact and inducing stress responses [29]. Additionally, carboxylation has been reported to enhance oxidative stress and increase membrane permeability in bacterial cells, which may explain its pronounced inhibitory effects [30]. In contrast, PS–NH2, bearing a positive charge, is generally expected to interact strongly with negatively charged bacterial surfaces. However, merely displacing bound water from phosphatidyl lipid head groups exerts minimal disruption to membrane integrity [29]. Overall, these findings are consistent with recent studies suggesting that both surface charge and hydrophobicity of microplastics influence bacterial adhesion, membrane interaction, and stress induction [28, 31, 32].
The enhancement of biofilm formation observed in both bacterial species (Fig. 4), especially in response to non-functionalized PS, supports the idea that microplastics serve as artificial substrates for biofilm development, as noted in several marine and freshwater studies [33-35]. In our study, E. coli showed biofilm enhancement primarily with PS, whereas Acinetobacter sp. responded similarly to all tested microplastics. This may reflect species-specific differences in surface sensing or adhesion mechanisms, which could be influenced by differences in outer membrane structure, motility, or extracellular polymeric substance (EPS) production.
Cytotoxicity assays revealed distinct stress responses between species (Fig. 5). In E. coli, the lack of significant ROS generation alongside elevated MDA and LDH levels implies that lipid peroxidation and membrane rupture were the primary modes of damage, rather than oxidative stress. On the other hand, Acinetobacter sp. exhibited increased ROS and MDA levels, while LDH release remained stable, suggesting sublethal membrane permeability changes coupled with oxidative damage. This divergence may stem from inherent differences in membrane composition, oxidative stress response systems, or tolerance to envelope damage, as previously proposed in stress-response studies involving environmental isolates [36-38].
Furthermore, the increased red fluorescence observed in both species upon PI staining suggests that membrane integrity was compromised in a manner not always detected by LDH release assays (Fig. 6). This aligns with prior work emphasizing that PI uptake can reflect subtle changes in membrane permeability, even in the absence of complete cell lysis [39, 40]. Our findings reinforce the idea that different cytotoxicity assays reflect distinct facets of cell damage, and that combining biochemical and imaging approaches provides a more comprehensive view.
Taken together, these findings suggest a mechanistic pathway in which microplastic–cell contact leads to enhanced surface attachment, biofilm development, and local membrane and oxidative stress, ultimately resulting in impaired growth and viability. The degree of this response is modulated by the surface chemistry of the microplastics, with non-functionalized or negatively charged particles generally causing greater effects. Importantly, the species-specific nature of these responses highlights the need for targeted evaluation of microplastic toxicity across diverse microbial taxa, particularly those with ecological or clinical relevance.
Overall, this study adds to a growing body of evidence that microplastics are not biologically inert and can actively alter microbial functions and survival. As microplastics continue to accumulate in natural and engineered environments, their interactions with microbial communities–both planktonic and biofilm-forming–must be considered in environmental risk assessments. This study has certain limitations, including the use of a single microplastic concentration (50 mg/L) and the focus on one polymer type–PS. These constraints, while enabling controlled experimental conditions, may not fully capture the concentration-dependent and polymer-specific nature of microbial responses observed in complex environmental settings. Future studies should explore a broader range of concentrations, polymer types, long-term adaptations, community-level interactions, and the potential for horizontal gene transfer on microplastic surfaces, which may have broader implications for microbial ecology and public health.
Conclusions
This study demonstrates that PS microplastics, including those with different surface functional groups, can induce species-specific physiological responses in Gram-negative bacteria through surface-mediated interactions. The observed alterations in growth, viability, biofilm formation, and stress responses indicate that even chemically inert microplastics can actively influence microbial behavior.
Our findings reveal a dual role of microplastics–as both attachment substrates and environmental stressors–and underscore the context-dependent nature of microbe–microplastic interactions. This study provides novel, quantitative evidence at the cellular level, contributing to a mechanistic understanding of how microplastics affect microbial physiology.
Given the ecological importance of microbial communities in biogeochemical cycling and environmental stability, these results highlight the need to incorporate microbial-level assessments into environmental risk evaluations of microplastic pollution. Future work should extend to diverse taxa and long-term exposure settings to elucidate broader ecological consequences.
Notes
Acknowledgement
This study was supported by a 2-Year Research Grant of Pusan National University.
Conflict of interest
The authors declare no conflict of interest.
CRediT author statement
SYK: Methodology, Visualization, Data curation, Investigation, Writing-original draft preparation; SWL: Methodology, Data curation, Investigation; EHL: Conceptualization, Methodology, Visualization, Investigation, Data Curation, Supervision, Writing- original draft, Reviewing and Editing.