DOI: 10.5061/dryad.wwpzgmsqn
title: Data from: Photodegradation modifies microplastic effects on soil properties and plant performance

description: Microplastics in soil affect plant-soil systems depending on their shape
 and polymer type. However, previous research has not yet considered the
 effects of degraded plastics, which are the plastic materials actually
 present in the environment. We selected 8 microplastics representing
 different shapes (fibers, films and foams) and polymer types, and exposed
 them to UV-C degradation. Each microplastic was mixed with soil at a
 concentration of 0.4% (w/w). The phytometer Daucus carota grew in each
 pot. At harvest, soil properties and plant biomass were measured.
 Photodegradation altered microplastics physical and chemical properties,
 impacting plant-soil systems. Microplastics degradation effects on plant
 and soil were observed with fibers and foams, but there were negligible
 effects with films. The latter could be explained by the polymer structure
 of films and manufacturer’s additives, potentially delaying their
 degradation. Degraded fibers increased soil respiration more than their
 non-degraded counterparts, as photodegradation increased the positive
 effects of fibers on soil water retention. The emergence of oxygenated
 groups during degradation may have increased the hydrophilicity of fibers,
 enhancing their ability to retain water. Degraded foams increased soil
 respiration, which could be related to the possible leaching of organic
 substances with lower partition coefficients, which may promote soil
 microbial activity. By contrast, degraded foams decreased soil
 aggregation, likely as degradation produced larger holes increasing their
 permeability. Also, the increase of hydrophilic molecules could have
 decreased soil particle cohesiveness. Degraded fibers and foams increased
 shoot and root mass as a result of microplastic effects on soil
 properties. Photodegraded microplastics affected root traits, which could
 be linked to microplastic effects on soil water status and plant coping
 strategies.
description: Plant species and microplastic
 selection We selected
 <em>Daucus carota</em> as a phytometer, which is a herbaceous
 plant typical of grassland ecosystems (Federal
 Agency for Nature Conservation, 2019) that shows clear responses
 to microplastics in soil (Lozano et
 al., 2021b, 2022). Also, we selected 8 secondary microplastics
 widely used in daily life, which represent three microplastic shapes:
 fibers, films and foams and 6 polymer types: polyamide (PA), polyester
 fibers made to at least 85% of polyethylene terephthalate (PET; Council
 Directive, 2011), polypropylene (PP), low density polyethylene (PE),
 polyethylene terephthalate (PET), polystyrene (PS) and polyurethane (PU)
 (Figure 1). Microplastic
 degradation Plastics were
 exposed to UV-C degradation (254 nm irradiation) by using a
 photodegradation chamber with three 36W UV-C lamps (Figure S1). The
 average incident energy in the chamber was of 20.98
 Wm<sup>-2</sup> (photometer; item number HD 2302.0, DeltaOHM),
 which simulated UV-C wavelengths. UV-A (present in solar light) and UV-C
 (used for shorter photodegradation experiments due to its higher energy)
 are ranges of the ultraviolet wavelength spectra that produce similar
 final outputs (e.g., breaking of polymer chains, volatile organic
 compounds production, reactive oxygen species or surface cracking) (Wu et
 al., 2023; de Freitas et al., 2022), and therefore could be compared. Each
 plastic type was photodegraded during two weeks; except PET film, which
 was photodegraded during four weeks as these films showed an extremely
 slow rate of degradation after FTIR analyses. Microplastics were randomly
 distributed in the chamber and their position was shifted twice during the
 photodegradation time. Then, plastic was manually cut with scissors and an
 upper size of 5.0 mm length for fibers and 5.0 mm<sup>2</sup>
 area for films and foams was established. Foams were cut by using a
 Philips HR3655/00 Standmixer (1400 W, ProBlend 6 3D Technologie,
 Netherlands) and sieved through a 4 mm mesh. Soil preparation We collected dry sandy loam soil (Albic
 Luvisol; 0.07% N, 0.77% C, pH 6.66) from a dry grassland plant community
 located in Dedelow, Brandenburg, Germany (53 37’ N, 13 77’ W). The soil
 was sieved (4 mm mesh size), homogenized, and mixed with each microplastic
 type at a concentration of 0.4% (w/w). Thus, 0.76 g of each microplastic
 type were mixed into 190 g of soil for each pot (4 cm diameter, 21 cm
 height, 200 ml). Microplastics were manually separated and mixed with the
 soil during 1 min in a large container, before placing it into each pot,
 to help provide an equal distribution of microplastics throughout the
 soil. The same handling was applied to the control soils (without
 microplastics) to provide the same disturbance level. Experimental design In September 2020, soil mixed with
 microplastics was pre-incubated without plants for 42 days in a glasshouse
 with a daylight period set at 12 h, 50 klx, and a temperature regime at
 22/18 ॰C day/night with a
 relative humidity of 40%. This incubation time allowed for the interaction
 between soil microbial communities and microplastic particles, and the
 potential leaching of microplastic chemical substances into the soil. Pots
 were watered with 50 ml of tap water, and covered to avoid evaporation but
 allowing aeration. Seeds of <em>Daucus carota</em> were
 surface-sterilized with 4% sodium hypochlorite for 5 min and 75% ethanol
 for 2 min and then thoroughly rinsed with sterile water. Seedlings of
 similar size were transplanted into pots 3 days after germination, with
 one single seedling per pot. Then, plants grew for 43 days and were
 watered every third day with 40 ml during the first 3 weeks, and then
 every second day with 30 ml of tap water, to maintain water holding
 capacity at ~70%. The experimental design consisted of 8 microplastic
 types (3 fibers, 3 films, 2 foams) x 2 degradation levels (non-degraded,
 degraded) x 7 replicates = 112 pots. Twelve additional pots were
 established as control without microplastics. All pots were randomly
 distributed in the glasshouse chamber, and their position shifted twice
 during the experiment to homogenize environmental conditions. All plants
 survived until the end of the experiment. At harvest, plants were
 separated into above and belowground parts; soil was air-dried and stored
 at 25 ॰C for soil
 aggregation analyses, while fresh soil samples were used to measure soil
 respiration. Microplastics
 characterization We measured
 the water absorption capacity of microplastic pieces of 6 x 6
 cm<sup>2</sup> following the UNE EN ISO 62:2008 standard.
 Plastic was immersed in distilled water for 24 hours and the changes in
 weight and thickness were determined. Water absorption was measured on
 films and foams. Due to the flexibility and brittleness of single fibers
 after degradation, it was not possible to assess their water absorption
 capacity. Identity of the plastics (i.e., polymer type) was confirmed by
 using a FTIR Jasco ATR-FTIR-4100 (Jasco International Co. Ltd., Tokyo,
 Japan), reflection mode, from 4000 to 600 cm<sup>-1</sup>, an
 average of 32 scans performed with the resolution of 4
 cm<sup>-1</sup>. For each plastic type, we measured two
 samples three times. Soil
 Measurements Soil
 aggregation: We measured water-stable soil aggregates (WSA) following a
 protocol by Kemper and Rosenau (1986), modified by Lehmann et al (2019). Briefly, we placed 4.0 g of dried soil (&lt;4 mm) on sieves with a mesh size of 250 µm. Soil was rewetted with deionized water by capillarity and inserted into a sieving machine (Agrisearch Equipment, Eijkelkamp, Giesbeek, Netherlands) for 3 min where the agitation and re-wetting caused the treated aggregates to slake. Then, we dried and weighed the water-stable fraction (dry matter) and subsequently, we extracted the coarse matter, which was also dried at 60 °C for 24 h. Soil aggregation was calculated as: WSA (%) = (dry matter - coarse matter)/(4.0 g - coarse matter). Soil respiration: We measured soil respiration via infrared gas analysis. We placed 25 g of fresh soil in individual 50 ml centrifuge tubes (Sarstedt AG & Co. KG, Nümbrecht Germany, item number 62.548.004) whose lids were modified to control gas exchange via a rubber septum (Supelco, Darmstadt, Germany, item number 27235 U). We measured CO<sub>2</sub> concentration (ppm) at two time points: First, we flushed the tubes with CO<sub>2</sub> free air for five minutes to measure CO<sub>2</sub> concentration at time zero. Then, soil samples were incubated at 20°C for 24 h and we measured CO2 concentration for the second time. At both times, we took a 1-mL air sample and injected it to an infrared gas analyzer (LiCOR- 6400XT photosynthesis system, Li-Cor Biosciences, Lincoln, NE). Measurements were obtained every ~2 secs. The difference between the maximum and the minimum value (peak) was converted to ppm using the calibration equation (ppm= -467 +195.18 peak). Plant measurements Plant biomass and root morphological traits At harvest, roots were carefully removed from the soil and gently washed by hand in order to measure morphological traits of fine roots (i.e., &lt;2 mm in diameter which included mostly first to third order roots). We measured length, surface area, volume and root average diameter on a fresh sample (random portion of the middle part of the root system) using the WinRhizoTM scanner-based system (v.2007; Regent Instruments Inc., Quebec, Canada). We calculated different root morphological traits: specific root surface area (SRSA; cm<sup>2</sup> mg<sup>−1</sup>), specific root length (SRL; cm mg<sup>−1</sup>), root average diameter (RAD; mm) and root tissue density (RTD; root dry weight per volume mg cm<sup>−3</sup>). Shoot and root mass were measured after drying samples at 60 °C for 72 h. Statistical analyses We performed principal component analysis (PCA) on our response variables for each microplastic shape either degraded or non-degraded, using the function “prcomp” and “fviz_pca” from the R package “factoextra” (Kassambara and Mundt, 2017). Ellipses in the graph grouped the treatments with a confidence level of 0.95. To validate differences between treatments we used the function “manova” from the R package “MASS” (Venables and Ripley, 2002). We tested the null hypothesis by means of the Pillai trace statistic, which is robust to violation of multivariate normality and homogeneity of the variance-covariance matrix (Quinn and Keough, 2002). We performed linear models to test the effect of microplastic degradation on plant-soil variables. Microplastic shape, polymer type and degradation were considered as explanatory variables. Polymer was nested within microplastic shape as each shape had different polymer types, and was included as a random factor in the model (Schielzeth and Nakagawa, 2013) as follows: ~ A*B +(1|A/C), with A representing ‘Shape’, B ‘Degradation’, and C ‘Polymer’. Response variables were log-transformed to fulfill linear models assumptions. The function “lmer” from the “lme4” R package was used in the mixed models. Then, we implemented the function “emmeans” from the eponymous R package to define pairwise comparisons. “Tukey” tests were used to compare each degraded microplastic type (shape and polymer) with its non-degraded counterpart, while the “Dunnett” test was used to compare each them with the control. Secondly, polymer and degradation were considered as fixed factors in the linear model. Residuals were checked to validate assumptions of normality and homogeneity and when necessary, we implemented the function “varIdent” from the “nlme” R package (Pinheiro et al., 2021) to account for heterogeneity in the treatment. Then, we implemented the function “glht” and the “Tukey” or “Dunnett” test from the “multcomp” R package to compare among treatments (Bretz et al., 2011). Statistical analyses were done in R 4.2.3 (R Core Team, 2023).
description: # Photodegradation modifies microplastic effects on soil properties and
 plant performance
 [https://doi.org/10.5061/dryad.wwpzgmsqn](https://doi.org/10.5061/dryad.wwpzgmsqn) As microplastics that enter the soil environment are those that have been previously degraded and not pristine plastic (without degradation), we think that we are only telling half of the story in regards to microplastics effects on plant-soil systems as we have not accounted for the effect of degraded microplastics. ## Description of the data and file structure We have worked with different microplastic shapes: Films, fibers and foams. Each shape made of different polymer types (LDPE: Low-density polyethyelene, PET: polyethylene terephthalate, PP: Polypropylene, PA: Polyamide, PU: Polyurethane, PS: Polystyrene). Each kind of microplastics was subjected to degradation or not (degraded, non-degraded). Then, the microplastic was mixed with soil at a concentration of 0.4% w/w and the plant Daucus carota grew there. A control treatment was established with soil without added microplastics. At harvest, we measured: Shoot mass, root mass, soil aggregation (WSA), Soil respiration (CO2\_ppm), and different root traits ( SRL: specific root length, SRSA: specific root surface area, RTD: root tissue density and diameter). Also, we measured water absorption (WaterAbsorb) in degraded and non-degraded plastics. Three replicates of each film and foam were measured, while the other three were not measured ("-"). We could not measure water absorption in fibers due to their small size and flexibility (cells containing "-"). Control did not have microplastics to be measured either (cells containing "-").