As agriculture strives to feed an ever-increasing number of people, it must also adapt to increasing exposure to minute plastic particles. To learn about the accumulation of nanoplastics by plants, we prepared well-defined block copolymer nanoparticles by aqueous dispersion polymerisation. A fluorophore was incorporated via hydrazone formation and uptake into roots and protoplasts of Arabidopsis thaliana was investigated using confocal microscopy. Here we show that uptake is inversely proportional to nanoparticle size. Positively charged particles accumulate around root surfaces and are not taken up by roots or protoplasts, whereas negatively charged nanoparticles accumulate slowly and become prominent over time in the xylem of intact roots. Neutral nanoparticles penetrate rapidly into intact cells at the surfaces of plant roots and into protoplasts, but xylem loading is lower than for negative nanoparticles. These behaviours differ from those of animal cells and our results show that despite the protection of rigid cell walls, plants are accessible to nanoplastics in soil and water.

Image data:

Methods

Arabidopsis thaliana root preparation

A. thaliana ecotype Columbia-0 was used for all nanoparticle uptake experiments. The seeds were surface-sterilised using successive washes in 10% Bleach or 70% ethanol prior to sowing onto sterile ½ strength Murashige and Skoog (MS) medium (2.2 g/L MS supplemented with B5 vitamins, 1% sucrose, 1% agar, pH 5.8). Seeds were stratified in the dark for two days at 4°C prior to germination for 5 days at 22°C with 12-hour day length.

Protoplast isolation

Protoplast solution (600 mM Mannitol, 2 mM MgCl2, 2 mM CaCl2, 10 mM KCl, 2 mM MES, 0.1% w/v bovine serum albumin) was prepared and adjusted to pH 5.5 with Tris-HCl, 0.2 µm filtered, and stored at -20°C until use. To isolate protoplasts, 5-day-old Arabidopsis thaliana roots were transferred into enzyme solution (1.5% w/v Cellulase RS, 0.1% w/v Pectolyase in protoplast solution) and chopped finely using a sterile blade. The solution with chopped roots was transferred to a 35 mm round Petri dish and incubated in the dark at 25°C with constant agitation for at least 2 hr. The resulting cell suspension was filtered sequentially through 70 μm and 40 μm meshes pre-soaked with enzyme solution. The protoplasts in solution were carefully transferred to polystyrene culture tubes and an equal volume of fresh protoplast solution added, before centrifugation at 300 g for 5 min at 4°C. The supernatant was discarded, the protoplasts resuspended in the same volume of protoplast solution and centrifugation repeated. The protoplasts were resuspended in 500 μL protoplast solution and used directly.

Confocal Microscopy

Intact Arabidopsis roots were incubated with nanoparticle samples for an hour before visualisation under fluorescence confocal microscopy. Seedling roots were dipped into 100 microlitres of the nanoparticle solution (all 1 mg/ml in water) or water in a microfuge tube. For confocal microscopy, root or protoplast samples were mounted on glass slides (1.0-1.2 mm thick) with a long cover glass (22x50 mm, 0.16-0.19 mm thick) before visualisation using a Zeiss LSM 880 instrument under 40x objectives lens with excitation laser at 488 nm and collecting emission at 496 - 577 nm. The images were stacked and reconstructed by ZEN software and analysed using ImageJ software. After incubation with nanoparticles, roots were also stained with propidium iodide for 10 min and propidium fluorescence was visualised with excitation at 561 nm and emission collected from 580 – 718 nm.

The files are .jpg image files of excel data files for qPCR data.

Files:

Folder names begin with a symbol to indicate the charge type on the nanoparticle for that set of images. This is followed by the name of the polymer used to make that nanoparticle and then the tissue from which the image is taken (root tip, root hair zone or protoplast). The time course files are given in folders named according to the time of collection post-treatment. All images are saved as .jpg files.