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. 2026 Sep 14;11(38):57541–57547. doi: 10.1021/acsomega.6c06111

Rotational Motion Amplifies Oxidative Responses to Polystyrene Nanoplastics: An In Vitro Proof-of-Concept and Particle-Impact Model

Hanzeng Li 1, Qian Zhou 1, Xiaofeng Chen 1,*, Shunqing Xu 1,*
PMCID: PMC13629356  PMID: 42824737

Abstract

Micro- and nanoplastics have been detected in human blood and arterial plaques, but how particle motion modifies cellular responses remains unclear. Here, we used a simple rotational exposure system as an in vitro proof-of-concept to test 80 nm polystyrene (PS) nanoplastics in A549 cells. Exposure to 100 μg/L PS nanoplastics or rotation at 6.7 rpm alone produced no detectable change in tetramethylrhodamine ethyl ester (TMRE) fluorescence or intracellular reactive oxygen species (ROS) under the conditions tested. In contrast, the combined treatment increased TMRE fluorescence and induced a pronounced ROS response without altering the mitochondrial-to-nuclear DNA ratio. Separately, an idealized particle-impact model predicted that single-particle impact force increases strongly with particle size and flow velocity at arterial bifurcations. Because the rotation speed was not calibrated to physiological wall shear stress, and A549 cells are not endothelial cells, these findings do not establish vascular toxicity. Rather, they show that particle motion can modify cellular responses to PS nanoplastics in vitro and support further testing in calibrated endothelial flow systems.

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Introduction

Particle pollutants, including PM2.5, black carbon, and microplastics, have been studied extensively for their capacity to generate reactive oxygen species (ROS) and carry co-occurring contaminants. − Once solid particles enter the bloodstream, they are transported dynamically before cellular uptake. , Particle size, shape, surface features, and local flow conditions can influence particle–wall interactions and cellular responses. , However, the contribution of particle motion to toxicity has rarely been isolated from that of conventional static exposure.

Compared with transition-metal- and organic-rich PM2.5, unmodified polymers such as polystyrene and polyethylene may have lower intrinsic chemical reactivity. − This does not make micro- and nanoplastics biologically inert. Their effects depend on polymer composition, particle size and shape, surface charge, weathering, additives, and adsorbed contaminants, all of which can influence cellular contact, uptake, oxidative stress, and inflammatory responses. ,− ,− After cellular uptake, persistent plastic particles may also disrupt lysosomal function and compromise cell viability. − By comparison, the biological effect of particle motion before or during cell contact remains less well-defined. −

Microplastics have been detected in human arterial tissue and carotid plaques, , and epidemiological and experimental evidence has raised concerns about possible cardiovascular effects. − These observations establish exposure and association but do not demonstrate a causal vascular mechanism. Because endothelial responses are sensitive to local flow conditions, particularly at arterial bifurcations, − particle motion may be an overlooked modifier of microplastic–cell interactions. This possibility requires the experimental separation of motion-related effects from responses observed under static exposure.

Here, we used 80 nm PS nanoplastics and A549 cells as a proof-of-concept epithelial model to test whether low-speed rotational motion modifies the mitochondrial membrane potential and intracellular ROS responses. The rotational condition was not intended as a quantitative reproduction of the arterial wall shear stress. In parallel, we used an idealized collision model to examine how the single-particle impact force scales with particle size and blood-flow velocity at arterial bifurcations. This combined approach identifies a testable physical exposure modifier while defining the experimental and physiological limits of the present study.

Results

Rotational Exposure Enhances Cellular Responses to PS Nanoplastics

To test whether particle motion modifies cellular responses, we established a rotational exposure assay using commercially manufactured 80 nm PS nanoplastics. A549 cells were selected as a proof-of-concept epithelial model because they efficiently internalize micro- and nanoplastics. Four groups were examined for 24 h: untreated control, 100 μg/L PS nanoplastics, rotation at 6.7 rpm, and PS nanoplastics plus rotation. The rotational speed was used as an operational low-intensity condition and was not calibrated to physiological wall shear stress. Neither PS nanoplastics nor rotation alone produced a detectable change in TMRE fluorescence (Figure A,B). In contrast, the combined treatment significantly increased TMRE fluorescence relative to the control and both single-treatment groups. Because increased TMRE fluorescence can reflect mitochondrial hyperpolarization, we describe this result as an alteration in mitochondrial membrane potential rather than as a loss of membrane potential. Assuming monodisperse spherical particles with a diameter of 80 nm and a PS density of 1050 kg/m3, the nominal exposure concentration of 100 μg/L corresponds to approximately 3.55 × 1011 particles/L, or 3.55 × 1014 particles/m3. This conversion is an idealized estimate and does not account for aggregation or size dispersion. The mitochondrial-to-nuclear DNA ratio did not differ among groups (Figure C), indicating that the TMRE response was not accompanied by a detectable change in mitochondrial DNA copy number.

1.

1

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Rotational exposure enhances changes in mitochondrial membrane potential and intracellular ROS in A549 cells treated with 80 nm PS nanoplastics. (A) Representative TMRE images; scale bar, 50 μm. (B) TMRE fluorescence intensity. (C) Mitochondrial-to-nuclear DNA ratio measured by qPCR. (D) Representative H2DCFDA images; scale bar, 100 μm. (E) H2DCFDA fluorescence intensity. The control group received no treatment; the rotation group was rotated continuously at 6.7 rpm; the PS group received 100 μg/L 80 nm PS nanoplastics; and the combined group received both treatments. Data are mean ± SEM from six independent biological experiments. One-way ANOVA followed by Tukey’s post-hoc test was used for multiple comparisons. *P < 0.05, **P < 0.01, and ***P < 0.001; ns, not significant.

Mitochondrial hyperpolarization can accompany cellular stress and increased electron leakage. We therefore measured intracellular ROS using H2DCFDA. The combined PS nanoplastic and rotation treatment produced a marked increase in H2DCFDA fluorescence, whereas neither treatment alone induced a detectable response (Figure D,E). These observations show that rotational exposure modifies the cellular response to PS nanoplastics under the conditions tested. Because a formal interaction analysis was not performed, we refer to this as a combined effect, rather than a synergistic effect.

Idealized Modeling of Particle Impact Force at Arterial Bifurcations

To examine how particle–wall collision force scales with particle size and flow velocity, we developed an idealized model using a 60° arterial bifurcation as an illustrative geometry. The model does not reproduce a specific vessel or the full complexity of pulsatile blood flow. Instead, it estimates the normal component of particle velocity and an upper-bound average force during a simplified collision.

For a spherical PS particle, the particle mass is

m=43πr3ρ

where r is the particle radius and ρ = 1050 kg/m3 is the density of PS.

The normal component of particle velocity is v n = v sin θ, where v is the local flow velocity and θ is the bifurcation angle.

For an upper-bound estimate, we assumed reversal of the normal momentum component over a collision interval approximated as the time required to traverse one particle diameter (Δt ≈ 2r/vn ). The resulting average impact force is

F=mvn2r=43πr2ρ(v⁡sin⁡θ)2

Under these assumptions, the estimated single-particle impact force scales with r 2 v 2. The result should be interpreted as an idealized scaling estimate rather than a direct measurement of force at the cell surface.

We calculated single-particle impact forces in Python for particle diameters of 80 nm to 5 μm and flow velocities of 0.1–0.5 m/s. Representative upper-bound estimates are shown in Table .

1. Idealized Upper-Bound Estimates of Single-Particle Impact Force at a 60° Bifurcation.

Diameter (nm) Velocity (m/s) Impact Force (N) Impact Force (pN)
80 0.1 5.28 × 10‑14 0.1
80 0.3 4.75 × 10‑13 0.5
80 0.5 1.32 × 10‑12 1.3
500 0.1 2.06 × 10‑12 2.1
500 0.3 1.86 × 10‑11 18.6
500 0.5 5.15 × 10‑11 51.5
5000 0.1 2.06 × 10‑10 206.2
5000 0.3 1.86 × 10–9 1855.5
5000 0.5 5.15 × 10–9 5154.2
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When particle number concentration (C, particles/m3) is included, multiplying the estimated force by the particle flux toward the wall gives a collision-frequency-weighted impact-loading rate

L=FCv⁡sin⁡θ

where L is the impact-force loading rate, F is the estimated force for one particle, C is particle number concentration, v is local flow velocity, and θ is the bifurcation angle. The quantity L has units of Pa s–1 and is not equivalent to physiological wall shear stress in pascals.

Under the simplified formulation, the impact-loading rate increases linearly with particle number concentration and strongly with particle size and velocity. Calculations were performed over particle diameters of 80 nm–5 μm, velocities of 0.1–0.5 m/s, and an illustrative concentration range of 106–1012 particles/m3. These ranges were used to examine model behavior and should not be interpreted as a calibration of the in vitro rotation system or as a validated distribution of human blood concentrations.

The estimated single-particle force increased markedly with particle size and velocity (Figures and ). At 0.5 m/s, the estimate for a 5 μm particle was approximately 3900-fold greater than that for an 80 nm particle. Figure shows the collision-frequency-weighted loading-rate calculation across an illustrative concentration range. Representative values from this calculation are summarized in Table . These results describe model scaling only and do not demonstrate that particle impacts generate physiological or pathological wall shear stress Figure .

2.

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Idealized single-particle impact force as a function of flow velocity for PS particles of different diameters at a 60° bifurcation. Estimates are shown for particle diameters of 80 nm, 500 nm, 1 μm, and 5 μm and velocities of 0.1–0.5 m/s. The model illustrates the dependence of the upper-bound force estimate on particle size and velocity.

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Heatmap of the idealized single-particle impact force as a function of particle diameter and flow velocity at a 60° bifurcation. Values are shown for four particle diameters (80 nm–5 μm) and three velocities (0.1, 0.3, and 0.5 m/s). The color scale indicates the magnitude of the upper-bound force estimate.

3.

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Collision-frequency-weighted impact-loading rate for 500 nm PS particles across an illustrative range of particle number concentrations and three bifurcation angles. The loading rate was calculated as L = F × C × v sin θ at v = 0.3 m/s.

2. Illustrative Collision-Frequency-Weighted Impact-Loading Rates under the Simplified Particle-Flux Calculation.

Particle diameter Concentration (particles/m3) Impact-force loading rate (Pa s–1)
80 nm 109–1011 0.0001–0.01 Pa s–1
500 nm 109–1011 0.005–0.5 Pa s–1
5 μm 109–1011 0.5–48 Pa s–1
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The model supports a qualitative conclusion that particle–wall interactions depend strongly on the particle size, concentration, velocity, and collision geometry. It does not establish equivalence between the rotational assay and arterial flow, and the calculated loading rate cannot be compared directly to blood pressure or wall shear stress because the quantities have different physical meanings and units. Calibrated fluid dynamic measurements are required before physiological thresholds can be assigned.

Discussion

The present study shows that rotational exposure modifies the response of A549 cells to 80 nm PS nanoplastics. Neither 100 μg/L PS nanoplastics nor rotation at 6.7 rpm alone produced a detectable TMRE or H2DCFDA response, whereas the combined treatment increased both signals. The unchanged mitochondrial-to-nuclear DNA ratio indicates that the TMRE response was not accompanied by a detectable change in mitochondrial DNA copy number. These data support a combined effect of particle exposure and motion in this in vitro system; they do not establish synergy, endothelial injury, or a cardiovascular mechanism.

The idealized collision model provides a separate scaling analysis. It indicates that the estimated single-particle impact force increases with particle size and flow velocity, but it does not calibrate 6.7 rpm to the arterial wall shear stress. Mechanosensitive channels offer one possible route by which membrane deformation could influence cellular signaling. In vascular endothelium, Piezo1 mediates Ca2+ influx in response to shear and membrane deformation , and has been linked to oxidative and inflammatory responses in vascular disease. − Particle motion could therefore influence Ca2+-dependent mitochondrial and inflammatory signaling. This remains a hypothesis: we did not measure Ca2+ flux, inhibit Piezo1, or use endothelial cells. Direct testing with endothelial models, GsMTx4 or genetic perturbation, and calcium imaging is required.

This work has several limitations. The rotation system was designed as a simple proof of concept and does not reproduce pulsatile flow, spatial shear gradients, blood viscosity, circulating blood cells, vessel geometry, or the endothelial phenotype. The rotation speed was not converted to wall shear stress, and the particle impact model was not calibrated against the in vitro platform. In addition, TMRE and H2DCFDA report mitochondrial polarization and intracellular oxidation, respectively, but they do not identify the initiating mechanosensor or establish cell injury. Future studies should use vascular endothelial cells in calibrated microfluidic devices, quantify local shear and particle trajectories, and validate key findings in ex vivo vessel preparations or other models that preserve vascular architecture. Accordingly, the present results identify particle motion as a potential exposure modifier rather than demonstrating a mechanism of cardiovascular toxicity. −

Methods

Cell Culture and Nanoplastic Exposure

A549 cells were seeded on glass coverslips in 35 mm culture dishes and allowed to reach approximately 50% confluence after overnight incubation. Four groups were exposed for 24 h: (1) untreated control; (2) 100 μg/L 80 nm PS nanoplastics; (3) continuous rotation at 6.7 rpm; and (4) PS nanoplastics plus rotation. During rotation, culture dishes were continuously rotated at 6.7 rpm, while all other incubation conditions and medium volumes were kept identical among groups. Cells were subjected to orbital shaking using a benchtop orbital shaker (Model: NS-OSR-200, NuoSi Scientific Instruments Co., Ltd., Shanghai, China). Culture dishes were placed horizontally on the shaker platform and subjected to orbital motion with an orbital diameter of 20 mm at 6.7 rpm. The 6.7 rpm condition was used as a low-intensity rotational perturbation and was not calibrated to arterial wall shear stress. After exposure, cells were analyzed using TMRE, H2DCFDA, or qPCR as described below.

TMRE Staining

TMRE was used to assess mitochondrial membrane potential (ΔΨm). A 100 μM TMRE stock solution was diluted 1000-fold in culture medium to prepare a 100 nM working solution. After treatment, cells were washed three times with PBS and incubated with 100 nM TMRE at 37 °C in the dark for 30 min. Coverslips were mounted on glass slides and imaged immediately using a Nikon Eclipse Ni upright fluorescence microscope. Identical acquisition settings were used for all groups within an experiment.

ROS Detection

Intracellular ROS-associated fluorescence was measured using H2DCFDA. The probe was dissolved in anhydrous DMSO to prepare a 10 mM stock solution and diluted in PBS to a final concentration of 10 μM. After the 24 h exposure, cells were washed three times with PBS and incubated with 10 μM H2DCFDA at 37 °C in the dark for 30 min. Coverslips were then mounted and imaged using a Nikon Eclipse Ni upright fluorescence microscope with identical acquisition settings across groups within an experiment.

Statistical Analysis

Data are presented as mean ± SEM from at least six independent biological experiments. Group differences were evaluated by one-way ANOVA followed by Tukey’s post-hoc test using GraphPad Prism 9.0. A two-sided P < 0.05 was considered statistically significant.

Supplementary Material

ao6c06111_si_001.pdf (103.5KB, pdf)

Acknowledgments

This study was supported by the National Science Foundation of China (32400724 to HL), the Natural Science Foundation of Hainan Province (823QN230 to HL), the First-Class Discipline Breakthrough Initiative of Hainan University (No. XKTP2025A04 to SX), Hainan Key Research Funding (ZDYF2025SHFZ021 to SX), and the Innovation Fund for Scientific and Technological Personnel of Hainan Province (KJRC2023B02 to SX).

The experimental and computational data that support the findings of this research are available in this article or upon request from the corresponding authors, Hanzeng Li (hanzeng.li@hainanu.edu.cn) or Shunqing Xu (xus@hainanu.edu.cn).

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsomega.6c06111.

  • Codes used in this manuscript (PDF)

†.

H.L. and Q.Z. contributed equally to this work. H.L. and Q.Z.: Methodology, investigation, and writingoriginal draft. X.C.: Data analysis and critical review. H.L. and S.X.: Supervision, writingreview and editing, funding acquisition, and academic guidance.

The authors declare no competing financial interest.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

ao6c06111_si_001.pdf (103.5KB, pdf)

Data Availability Statement

The experimental and computational data that support the findings of this research are available in this article or upon request from the corresponding authors, Hanzeng Li (hanzeng.li@hainanu.edu.cn) or Shunqing Xu (xus@hainanu.edu.cn).

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