Background

Waste plastic from consumer and industrial sources disperses freely into the environment, where it degrades into micro- (10–1,000 μm) and eventually nano (<1 μm) scale materials by natural weathering processes such as solar radiation and physical abrasion. These plastic fragments of micrometer and nanometer sizes are abbreviated in this article as MPs and NPs, respectively. They have become ubiquitous pollutants, with more than 250,000 metric tons of plastic estimated to persist across all waterways worldwide [1–3]. Of particular concern are NPs, because they are not only too small to be remediated by most conventional approaches [4] but are also small enough to infiltrate biological tissues and impact living systems on various cellular levels [5]. Biomimetic lipid bilayers have been extensively shown to be deformed and perforated when interacting with nanoparticles [6–11], where these damaging phenomena are reported to be driven by a combination of nanomaterial composition, size, shape, surface charge, and surface chemistry. Cell culture and toxicological studies have similarly shown various deleterious effects on biological function due to NP exposure [5,12,13].

Despite the vast differences in size, shape, and composition of environmental nanoplastic wastes, over 90% of research literature discussed in a recent review employs pristine or virgin plastic material engineered to have uniform shapes (most commonly spherical beads) and sizes. Furthermore, while polystyrene (PS) is the model plastic used in ~96% of those studies, PS comprises less than 6% of typical environmental samples [14,15]. Indeed, polystyrene (nano-PS) and polymethyl methacrylate (nano-PMMA) are often the only commercially available engineered nanoplastics [4]. The densities of nano-PS and nano-PMMA are greater than that of water/culture media, making them convenient choices for cell toxicity studies, and their particle uniformity in commercial preparations allows for predictable and consistent behavior in vivo [4]. On the other hand, “real-world” nanoplastic pollution is found with a variety of different compositions, shapes, surface chemistries, surface charges, additives, and adulterants due to the chaotic nature of environmental aging and the wide variety of plastic waste source streams [14]. Such adulterant matrices include, but are not limited to, non-plastic organic debris, including leaves and other plant matter, animal protein and remains, humic acid, soil, sediment, motor oil, bacteria, metals, paper, and fibers like cotton and rope from the environment that can be found alongside plastic waste [4]. There is thus a significant gap in understanding the biological impact of weathered waste NPs, as the vast majority of current literature uses NP models that are not environmentally realistic. In other words, those studies can demonstrate an impact on living systems, but because plastic waste is seldom uniformly shaped and pristine, further studies may be needed to determine real-world effects.

In this protocol, we describe a straightforward, customizable, and more environmentally relevant method to experimentally monitor the effects of nanoplastic pollution using, for example, rat alveolar epithelial cell monolayers for assessment of active and passive ion transport properties. NPs are generated by accelerated photoaging/photooxidation of pulverized plastic waste using an ultraviolet (UV) light source coupled with an ozone (O₃) gas generator. It is estimated that this aging process simulates years of solar irradiation within an exposure time of just 72–144 h [16]. NPs generated by this method can be easily concentrated (up to ~120 mg/mL) and mixed with other NP materials as desired. Changes in cell functions (e.g., ion transport and membrane damage) of lung alveolar epithelial cell monolayers exposed apically to these different environmental NP contaminant models can then be monitored using a simple bioelectric assay by tracking both spontaneous potential difference (PD) and electrical resistance (Rt) of cell monolayers over time at different concentrations of various fabricated NPs. An equivalent short-circuit current (Ieq), which represents the active ion transport rate across the monolayer, can be estimated from the ratio PD/Rt as described previously [17,21–23]. Finally, NPs can be tagged with a fluorescent labeling molecule for convenient tracking, if needed. This will allow researchers using confocal microscopy to easily visualize NPs interacting with cells, to quantify NPs with particle counters, to track the flux of NPs across cell monolayers [17,21], and/or to further shed light on mechanisms underlying NP interactions with cells [e.g., NP interaction with lipid bilayers, endocytosis, autophagy, intracellular organelles (mitochondria, endoplasmic reticulum, Golgi, and lysosomes), and exocytosis].

The method for nanoplastic generation described herein is an improved and expanded version of the method described by Sorasan and coworkers [16]. In that original method, only PE, PP, and PS were considered, and the raw plastic materials were sourced from various plastic-strewn beaches in Spain and Portugal. While true environmental samples are the best models for studying NP pollution impact, collecting field samples can be an expensive and arduous task, since alternative methods such as Fourier transform infrared spectroscopy (FTIR) must be used to first identify the plastics found. In addition, the composition of plastics globally found in the environment is mostly PE and PP [14,16] but will vary widely among local sources. The method described herein is much more rapid, high-throughput, controllable, and cost-effective, as the researcher can “mix and match” any consumer plastic sources and have total control over the composition of the NP mixtures [e.g., PE, PP, PS, or polyethylene terephthalate (PET); any plastic amenable to grinding (e.g., PMMA, polycarbonate and nylon) can also work]. Furthermore, much less field travel time and collection and processing efforts are required. Finally, if desired, the researcher could easily adapt this procedure for field-collected samples, with the only difference being the sourcing of the plastic.