# Biodegradable mulch films exhibit slower-than-expected degradation with negligible effects on soil microbial communities

Martine Graf, Estelle Choiselat, Michaela K Reay, Rafael Bargiela, Athanasios Dimitriou, Qiuyun Liu, Robert M Elias, Peter N Golyshin, Robert Griffiths, David R Chadwick, et al.

► To cite this version:

Martine Graf, Estelle Choiselat, Michaela K Reay, Rafael Bargiela, Athanasios Dimitriou, et al. "Biodegradable mulch films exhibit slower-than-expected degradation with negligible effects on soil microbial communities." Journal of Hazardous Materials, 2025, 498, 10.1016/j.jhazmat.2025.139871. hal-05420397

HAL Id: hal-05420397

https://hal.science/hal-05420397v1

Submitted on 17 Dec 2025

HAL is a multi-disciplinary open access
archive for the deposit and dissemination of sci-
entific research documents, whether they are pub-
lished or not. The documents may come from
teaching and research institutions in France or
abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est
destinée au dépôt et à la diffusion de documents
scientifiques de niveau recherche, publiés ou non,
émanant des établissements d'enseignement et de
recherche français ou étrangers, des laboratoires
publics ou privés.

<figure><img src="image_1.png" /></figure>

Distributed under a Creative Commons Attribution 4.0 International License
<figure><img src="image_2.png" /></figure>

# Biodegradable mulch films exhibit slower-than-expected degradation with negligible effects on soil microbial communities

Martine Graf<sup>a, *, ID</sup>, Estelle Choiselat<sup>b</sup>, Michaela K. Reay<sup>c</sup>, Rafael Bargiela<sup>d, ID</sup>, Athanasios Dimitriou<sup>e</sup>, Qiuyun Liu<sup>e, ID</sup>, Robert M. Elias<sup>e</sup>, Peter N. Golyshin<sup>d, ID</sup>, Robert Griffiths<sup>a, d</sup>, David R. Chadwick<sup>a</sup>, Davey L. Jones<sup>a, ID</sup>

<figure><img src="image_3.png" /></figure>

## HIGHLIGHTS

* 12-month field degradation of 8 PMF types above and below ground.

* Negligible PMF effect on bacteria and fungi soil communities.

* PMF degradation rate under field conditions much slower than anticipated.

* Fragmentation does not occur without external mechanical stress.

* Negligible PMF effect on soil properties.

## GRAPHICAL ABSTRACT

<figure><img src="image_4.png" /></figure>

## ARTICLE INFO

### Keywords:

Biodegradation testing

Plastic pollution

16S rRNA and ITS2 sequencing

ATR-FTIR

Polymer degradation

## ABSTRACT

The adoption of biodegradable plastic mulch films (PMFs) has increased as a sustainable alternative to conventional PMFs. However, evidence suggests that current biodegradability standards may not reflect in-field conditions, leading to potential accumulation of biodegradable PMFs in soil with yet undetermined effects on soil microbial communities. We therefore conducted a field experiment to investigate the above and below ground degradation of one conventional and seven biodegradable PMF blends in a temperate climate (UK) over 12 months. We found that PMFs had no effect on soil properties and a negligible effect on microbial community structure. The main factors driving change in relative abundance of bacteria and fungi were exposure time, followed by the potential of meso- and macrofauna interactions. PMFs buried in soil showed no sign of fragmentation, regardless of presence of meso- and macrofauna, which was likely the cause for no overall difference in microplastic concentrations. The slow degradation of all PMFs combined with the lack of physical fragmentation under natural conditions, indicates that degradation is slower than expected according to current

<footnote><sup>a</sup> School of Environmental & Natural Sciences, Bangor University, Bangor LL57 2UW, UK</footnote>

<footnote><sup>b</sup> Institut Polytechnique UniLaSalle, Beauvais 60000, France</footnote>

<footnote><sup>c</sup> Organic Geochemistry Unit, School of Chemistry, University of Bristol, Bristol BS8 1TS, UK</footnote>

<footnote><sup>d</sup> Centre for Environmental Biotechnology, Bangor University, Bangor LL57 2UW, UK</footnote>

<footnote><sup>e</sup> The BioComposites Centre, Bangor University, Bangor LL57 2UW, UK</footnote>

<footnote>* Correspondence to: School of Environmental & Natural Sciences, Bangor University, Gwynedd, Bangor LL57 2UW, UK.</footnote>

<footnote>E-mail address: m.graf@bangor.ac.uk (M. Graf).</footnote>
biodegradability standards. This highlights the importance of reframing standards by treating degradation as a system property, influenced by a variety of environmental and intrinsic factors.

# 1. Introduction

The use of agricultural plastic mulch films (PMFs) has increased in recent years, reaching an estimated 2.5 Mt y⁻¹ globally, with production predicted to steadily increase for the next few decades [1]. PMFs have become an integral part of agricultural systems due to their positive effects on soil water retention, weed suppression, crop yield, and reduction of soil erosion [1]. Conventional petroleum-based PMFs are mainly made from low-density polyethylene (LDPE) and are mostly used for a short duration of up to 10 months [1,2], depending on crop type, PMF type, and environmental conditions. After harvest, these PMFs are often ploughed back into the soil or incinerated at field margins due to a lack of recycling infrastructure, leading to an accumulation of legacy plastic in agricultural soils [1]. This legacy plastic degrades and fragments only slowly with time, eventually turning these macroplastic fragments (> 5 mm) into microplastic (MP, ≤ 5 mm) and nanoplastic (NP, ≤ 1 µm) particles, which are more mobile and have the potential to enter watercourses, be taken up by plants, and enter the food chain [1, 3].

As an alternative, the agricultural industry has recently moved towards seemingly more sustainable options, which has led to the gradual adoption of biodegradable PMFs in many countries. Although biodegradable PMFs can be fully bio-based or fossil-fuel based (or a combination), they have the potential to address the problem of legacy plastic accumulation by breaking down relatively rapidly (i.e., within a few years) and being fully degraded by microbes in the soil, resulting in only biomass, CO₂, and H₂O in the process. Current EU standards for biodegradability require PMFs to degrade ≥ 90% within 24 months (EN 17033). However, previous field studies have already shown that degradation rate of biodegradable PMFs is often much slower in realistic field settings [4,5], calling for a revision of biodegradation standards [6]. Treating biodegradation of PMFs as a system property, i.e. taking environmental factors (e.g. climate and soil type) and intrinsic properties of PMFs into account [6], would greatly improve the translation of current standards into real world conditions.

As farmers are actively encouraged to plough biodegradable PMFs into the soil after harvest (in contrast to LDPE PMFs which are supposed to be removed), this could lead to an accumulation of biodegradable PMF macroplastic in the soil. Previous studies have shown that the degradation rate is inversely correlated to plastic concentration in soil, due to nutrient limitation [7]. Based on only one growing season per year, and an average degradation time exceeding 12 months, this poses a high potential for a continued increase in biodegradable PMFs in agricultural fields. This could result in a subsequent decrease of degradation rates, alongside increasing accumulation and leaching potential of additives and other non-intentionally added substances, which have been shown to have a higher leaching potential compared to conventional PMFs [8].

It is well established that degradation of PMFs is influenced by environmental conditions, soil types, and microbial community composition [4,9–12]. However, to date most field-based research involving PMFs focuses less on their degradability and more on their overall impact on soil and crop health [13,14], resulting in limited knowledge about PMF degradation rates, particularly in a temperate climate. The uncertainty regarding degradation rates of different biodegradable PMF blends, coupled with the questions regarding applicability of current standards in real-world field conditions, has formed the basis of this field study.

Here, we investigated the above and below ground degradation rates of one conventional PMF (LDPE), and seven biodegradable PMF blends, consisting of poly(lactic acid) (PLA), poly(butylene adipate-co-

terephthalate) (PBAT), poly(3-hydroxybutyrate-co-3-hydroxyvalerate) (PHBV), and poly(aromatic aliphatic urethane) (PAAU) over a period of 12 months in a field-based study in a temperate climate. The aims were to i) evaluate chemical degradation and physical fragmentation rates of PMF blends after above and below ground degradation; ii) identify the effect of meso- and macrofauna interactions on below ground degradation rates by allowing either only microfauna, or micro-, meso-, and macrofauna interactions; iii) determine the MP production rate after 12 months below ground degradation; and iv) assess the effect of PMF types on physio-chemical soil properties and microbial community composition (archaea, bacteria, and fungi). We hypothesised that i) the degradation and fragmentation rate of biodegradable PMFs would be faster compared to LDPE, subsequently resulting in an increased MP concentration in the surrounding soil; ii) the interaction of meso- and macrofauna would accelerate physical fragmentation, and subsequently chemical degradation compared to only microfauna interactions; iii) the PMF type would have effects on soil nutrients, specifically the soil dissolved organic carbon (DOC) pool (increasing with biodegradable PMF compared to LDPE); and iv) the PMF type would influence microbial community composition.

# 2. Materials and methods

## 2.1. Plastic mulch film material

This study used two commercially available PMFs (one conventional and one biodegradable), and six bespoke biodegradable PMF blends in commercial development. The conventional PMF used was LDPE and the biodegradable PMF was a PLA/PBAT blend with a polybutylene sebacate (PBSe) adhesive [15], from hereon referred to as LDPE and PLA/PBAT, respectively (Table 1). Organic co-contaminants of these two films were identified using a sequential solvent extraction followed by gas chromatography (GC) and GC/MS (mass spectrometry) analyses, and fillers and metals were identified by analysing the ash content of the PMFs using total reflection X-rays fluorescence [8]. See supplementary material for information on organic co-contaminants.

The six bespoke biodegradable PMF blends were formulated and provided by Wells Performance Materials Ltd. (UK) and were split into two separate groups based on the main polymer composition (Group A PBAT only, Group B PBAT with an equal ratio of PHBV and PAAU) which contained different additives. All films contained lubricants which were added during the production process but differed in polymer composition and included additives (Table 1). Information on polymer and substance ratios can be found in Table S1, however as these blends were formulated in collaboration with Wells Performance Materials Ltd. as part of their product innovation, we are unable to provide details on exact blend ratios and type of substances added. Bio A and Bio B served as baseline for the other four blends, to assess if and how the addition of certain stabiliser and catalysts affects the degradation process. All six PMF blends were developed with the aim to further improve biodegradability of already existing commercialised blends.

## 2.2. Experimental setup

### 2.2.1. Study site and environmental data

The study was undertaken in an arable field located at the Henfaes Research Centre, Abergwyngregyn, North Wales, UK (53°14′19.4″N, 4°00′54″W). The experimental area has a temperate oceanic climate, no known history of agricultural plastic application, and was previously under a rotation of grass (*Lolium perenne* L.) and wheat (*Triticum aestivum*). The soil is classed as a freely draining sandy clay loam textured
Eutric Cambisol [16]; initial soil properties can be found in Table S2. Environmental data was recorded in 30-minute intervals by a meteorological station on site. Soil surface temperature was recorded hourly using iButtons (DS1922L Thermochron Data Logger, Measurement Systems Ltd., UK) placed on bare soil (n = 3) and under the LDPE PMF surface (n = 3) to determine the average soil surface temperature. Soil moisture and temperature at 5 cm depth were recorded in 30-minute intervals using TDT sensors (n = 12) connected to a data logger (SDI-12, Acclima, USA). The experiment started on 29th October 2021 and ended on 26th October 2022.

### 2.2.2. Above ground degradation and sampling of PMFs

To assess the above ground degradation of PMFs, films were cut into 0.25 × 4 m strips and stretched taut over rotavated soil seed beds in a randomised block design (n = 4 per film type) (Fig. S1), secured by burying 5 cm of the edges in the surrounding soil, and weighed down with soil across the film surface in 1 m intervals to reduce mechanical stress from wind. Films were laid out slightly dome-shaped to allow water run-off and reduce splashback from precipitation.

All films were sampled monthly over the course of 12 months, by removing a 10 × 10 cm piece of exposed film from the end of each row and burying the newly created end with soil to reduce the likelihood of mechanical damage introduced by the sampling, subsequently resulting in shortened rows of PMF throughout the experiment. Samples were cleaned with ultrapure water and stored in paper bags.

### 2.2.3. Below ground degradation and sampling of PMFs

To assess the below ground degradation of PMFs in soil and their potential impact on soil biological and chemical properties, PMFs were cut into 5 × 5 cm pieces (n = 4 per film type) and horizontally placed into 10 × 10 cm nylon mesh bags, surrounded by two layers of soil (ca. 5 mm on each side of the PMF piece). The soil was previously collected from the study site, air dried, then brought to a moisture content of 44%, and weighed out to 20 ± 0.1 g per mesh bag. Control treatments contained only soil. Mesh bags were sealed using a heat gun and stored in aluminium foil at 4 °C overnight before burying them in the field, adjacent to the area designated for the above ground degradation.

Two different nylon mesh sizes were used to assess the impact of micro-, meso-, and macrofauna on the degradation process. The fine mesh (100 µm mesh size, 32% opening) allowed for only microfauna to enter the bags, whereas the coarse mesh (5000 µm mesh size, 72% opening) allowed micro-, meso-, and macrofauna to interact with the contents of the bags [17,18], from hereon referred to as fine and coarse mesh, respectively. Mesh was sourced from Normesh Ltd. (UK), and nylon was selected as an inert material with low likelihood of affecting surrounding soil properties, which would be a risk if using metal. None of the PMF materials used in this study contained nylon, and the control

mesh bags allowed for a creation of a baseline.

Mesh bags were buried horizontally in the soil at 5 cm depth in a randomised block design for fine and coarse mesh bags respectively (Fig. S2). Mesh bags were sampled in 4-month intervals over a 12-month period by excavating them from the soil. The PMF pieces were removed and cleaned using ultrapure water, subsequently stored in paper bags.

### 2.3. Soil property analysis

For each sampling point, the soil from each mesh bag was analysed for pH and EC (electrical conductivity) in a 1:2.5 soil:distilled water solution using standard electrodes. Soil available ammonium ($NH_4^+$) and nitrate ($NO_3^-$) were measured in a 0.5 M $K_2SO_4$ soil extract according to the salicylic acid and vanadium chloride procedure [19,20]. DOC was measured using the same extract on a Multi-N/C Series TOC/TN analyser (Analytik-Jena, Germany). Available phosphate ($PO_4^3-$) was measured in a 0.5 M acetic acid extract according to the molybdate blue method [21].

### 2.4. Microplastic analysis

Microplastic generation of PMFs within the mesh bags and baseline MP contamination of control soil were assessed after 12 months burial in the soil, by oven-drying soil at 40 °C for 24 h and utilising two optimised extraction methods applicable for conventional [22] and biodegradable plastic [23]. Briefly, for the conventional extraction method, 5 ± 0.05 g of dry soil was homogenised with 230 ml of saturated NaCl and left to stand overnight for density separation. The supernatant was decanted into a new beaker and 4 M NaOH was added to a fixed volume of 500 ml, homogenised, and left to stand overnight for further density separation and digestion of organic material. The supernatant was then filtered through a 0.2 µm nylon membrane (Cytiva, USA) using a glass vacuum filtration system. Plastic material was replaced with metal or glass where possible, and plastic was only used in form of pipette tips, membranes, and vial caps.

For the biodegradable extraction method, 5 ± 0.05 g of dry soil was homogenised with 45 ml of distilled water in a new 50 ml polypropylene (PP) plastic centrifuge tube. The solution was mixed for 30 s using a Vortex Genie 2 (Scientific Industries, USA) and further homogenised using a shaker at 225 rpm for 10 min. Samples were then centrifuged at 2800 rcf for 10 min and supernatant discarded. The precipitate was then mixed with 45 ml of 6.24 M $ZnCl_2$ solution and the aforementioned mixing and centrifugation steps were repeated three times, using new $ZnCl_2$ solution each time. The $ZnCl_2$ supernatant was combined in a glass beaker, left standing overnight for further separation, and then filtered through a 0.2 µm nylon membrane using a glass vacuum filtration system.

<table><caption>**Table 1**

Overview of PMF types and properties.</caption><thead><tr><th>PMF type</th><th>Polymer(s)</th><th>Additives</th><th>Colour</th><th>Thickness [µm]</th><th>Supplier</th></tr></thead><tbody><tr><td>LDPE</td><td>LDPE</td><td>See supplementary material</td><td>Black</td><td>23</td><td>Gro-Clean, GroMax, UK</td></tr><tr><td>PLA/ PBAT</td><td>PLA 15%, PBAT 85% with PBSe adhesive</td><td>See supplementary material</td><td>Black</td><td>10.5</td><td>Gro-Clean Bio-Mulch, GroMax UK</td></tr><tr><td>Bio A</td><td>PBAT</td><td>Lubricant</td><td>Clear</td><td>17</td><td>Wells Performance Materials Ltd., UK</td></tr><tr><td>Bio A1</td><td>PBAT</td><td>Lubricant, thermal stabiliser, antioxidant, two catalysts</td><td>Clear</td><td>23</td><td>Wells Performance Materials Ltd., UK</td></tr><tr><td>Bio A2</td><td>PBAT</td><td>Lubricant, thermal stabiliser, one catalyst</td><td>Clear</td><td>23</td><td>Wells Performance Materials Ltd., UK</td></tr><tr><td>Bio B</td><td>PBAT, PAAU, PHBV</td><td>Lubricant</td><td>Clear</td><td>22</td><td>Wells Performance Materials Ltd., UK</td></tr><tr><td>Bio B1</td><td>PBAT, PAAU, PHBV</td><td>Lubricant, thermal stabiliser, antioxidant, one catalyst</td><td>Clear</td><td>29</td><td>Wells Performance Materials Ltd., UK</td></tr><tr><td>Bio B2</td><td>PBAT, PAAU, PHBV</td><td>Lubricant, thermal stabiliser, two catalysts</td><td>Clear</td><td>27</td><td>Wells Performance Materials Ltd., UK</td></tr></tbody></table>
Particle recovery tests for both methods matched ranges in the original referenced methods. For both methods, particles were resuspended from the membrane in 20 ml ethanol via sonication, which was further evaporated to a fixed volume of 5 ml. The solution containing the particles was mixed well and 10 µl of the sample were analysed in triplicates using laser direct infrared chemical imaging (8700 LDIR, Agilent Technologies Inc., USA). Particle spectra were matched against a modified version of the 'Microplastics Starter 2.0' library in the Clarity software (v1.5.58, Agilent Technologies Inc., USA). Particle size range was limited to 20 – 500 µm and the threshold for acceptance of library matches was set to 80 % match quality. Blanks for each extraction method (n = 3), which contained only reagents and underwent the same procedures, were analysed alongside samples. Polyamide was excluded from analysis for both extraction methods due to the use of nylon mesh bags and nylon filter membranes. For samples extracted with the conventional method, identified biodegradable particles were excluded from analysis, and vice versa for samples extracted with the biodegradable method. Control soils were extracted with both methods.

## 2.5. Soil microbial community analysis

Soil from the mesh bags of each timepoint (n = 3 timepoints; n = 4 replicates) was frozen at -80 °C immediately after sampling, and DNA was extracted using the Quick DNA Fecal/Soil Microbe Miniprep Kit (Zymo Research, USA). Quality and concentration of extracted DNA samples were assessed by Qubit 4.0 Fluorometer (Thermo Fisher Scientific, USA) using a dsDNA HS Assay Kit (Life Technologies, USA). DNA extracts were then further used for 16S rRNA and ITS2 amplicon sequencing. The NCBI BioProject accession number is PRJNA1229197.

### 2.5.1. 16S rRNA amplicon sequencing

Libraries of 16S rRNA gene amplicons were prepared by duplicate PCR with double-indexed fusion primers [24]. The V4 16S rRNA gene fragment was amplified using modified forward F515 (5'-GTGBCAGCMGCCGCGGTAA-3') and reverse R806 (5'-GGACTACHVGGGGTWCTAAT-3') prokaryotic primer. Primers were designed to contain: the Illumina adapters and sequencing primers, a 12 bp barcode sequence, a heterogeneity spacer, and 16S rRNA gene universal primers [24]. PCRs were performed using OneTaq DNA Polymerase (New England Biolabs, USA). All reactions were run with no-template negative controls. Thermocycling conditions were as follows: initial denaturation at 95 °C for 2 min, followed by 30 cycles at 95 °C for 30 s, 50 °C for 50 s, and 68 °C for 90 s with a final elongation at 68 °C for 5 min. Amplicons were visualised in a 1.2 % tris-acetate agarose gel and DNA bands of approximately 440 bp were gel-purified using the QIAEX II Gel Extraction Kit (Qiagen, Germany). The purified amplicons were then quantified using the Qubit 4.0 Fluorometer, pooled in equimolar amounts and the final pool was run on the Illumina MiSeq platform (Illumina Inc., USA) using 500-cycle v2 chemistry kits.

Raw sequencing reads were processed according to previously described protocols [24,25]. Briefly, the data was pre-processed to extract the barcodes from sequences, and then cleaned of primer sequences using TagCleaner v0.16 [26]. The barcodes and the sequences were re-matched using in-house Python scripts. The resulting filtered reads were analysed using QIIME v2022.2 [27]. First, the libraries were demultiplexed based on the different barcodes, then sequences were classified on amplicon sequence variants (ASVs) combining de novo and reference-based methods using the SILVA version 138.1 reference database. Analyses of most abundant taxonomic groups were performed using in-house R-scripts, selecting those groups with a relative abundance ≥ 2 % in any of the samples. Selection started at species level and groups were added to the immediate upper taxonomic level if none of the samples of that group passed the 2 % threshold.

### 2.5.2. ITS2 amplicon sequencing

Extracted DNA samples were processed and analysed by Novogene

Company Ltd. (UK). Briefly, PCR amplification of the ITS2 region was performed by using ITS3–2024F and ITS4–2409R primers. Quality was determined via 2 % agarose gel electrophoresis and products from each sample were pooled, end-repaired, A-tailed, and further ligated with Illumina adapters. The purified amplicons were quantified using the Qubit 4.0 Fluorometer, pooled in equimolar amounts and the final pool was run on the Illumina MiSeq platform to generate 250 bp paired-end raw reads.

Raw data was spliced and filtered, then DADA2 [28] was used to obtain ASVs and annotated using QIIME v2022.2 [27]. Analyses of most abundant taxonomic groups were performed using in-house R-based scripts, selecting those groups with a relative abundance ≥ 2 % in any of the samples. Selection started at species level and groups were added to the immediate upper taxonomic level if none of the samples of that group passed the 2 % threshold.

## 2.6. PMF analysis

### 2.6.1. Analysis of chemical spectra changes

Chemical changes to the surface of PMFs (both above and below ground) at each sampling point were assessed using attenuated total reflectance-Fourier transform infrared (ATR-FTIR) spectroscopy. The analysis was carried out using a Cary 630 FTIR bench spectrometer (Agilent Technologies Inc., USA) with a diamond ATR crystal attachment in conjunction with a KBr source with a spectral range of 650 – 4000 cm⁻¹ and a resolution of 4 cm⁻¹. Spectra were recorded as reflectance (32 scans per sampling point & background) using the MicroLab Pharma Software (Agilent Technologies Inc., USA). The crystal was routinely cleaned using acetone, and background measurements were taken every 2 min. Five random points per sample (n = 4) were measured, and sides were randomly switched throughout the sampling process, resulting in a total of 20 spectra per film type per time point. Spectra manipulation and transformation were carried out using the open-source online application Open Specy [29]. Spectra were transformed to absorbance according to the Kubelka-Munk equation, baseline correction was carried out with the setting ‘Polynomial 8’, and the CO₂ region between 2200 – 2400 cm⁻¹ was flattened.

To quantify changes in spectra peak intensity, the carbonyl index (CI) was calculated by dividing the area under the carbonyl (C=O) peak by the area of a reference peak (here: mix of carbon-hydrogen (C-H) bonds) [30]. To capture the entire peak area and account for peak shifts caused by degradation, the C=O wavenumber interval of 1536 – 1928 cm⁻¹ [31,32] and CH wavenumber interval of 1384 – 1472 cm⁻¹ [31,33,34] were selected for LDPE. For PLA/PBAT, the C=O wavenumber interval of 1552 – 1808 cm⁻¹ [35–38] and CH wavenumber interval of 1336 – 1496 cm⁻¹ [39–44] were selected. For the six bespoke biodegradable PMFs, the C=O wavenumber interval of 1650 – 1755 cm⁻¹ [35–38] and CH wavenumber interval of 1350 – 1470 cm⁻¹ [39–44] were used. For LDPE, the C=O wavenumber interval includes C=O stretching vibrations, and bending vibrations of CH bonds, and the CH wavenumber interval includes bending or scissoring vibrations of CH bonds (primarily CH₂) as well as some CH stretching. For PLA/PBAT, the C=O wavenumber interval includes stretching vibrations of the C=O group, C-C stretching or bending vibrations, and bending vibrations of CH bonds, and the CH wavenumber interval includes the bending or scissoring of CH bonds (primarily CH₃ and CH₂). For Bio A, A1, and A2 (main polymer: PBAT), the C=O wavenumber interval includes stretching of ester bonds, and the CH wavenumber interval includes scissoring of CH₂ bonds, asymmetric bending of CH₃ bonds, and CH in-plane bending associated with the benzene ring. For Bio B, B1, and B2 (blend of PBAT, PHBV, PAAU), the C=O wavenumber interval includes stretching vibrations of esters, urethanes, and amides, and the CH wavenumber intervals includes scissoring and bending of CH₂ and CH₃ groups, and potential aromatic C=C and CH bending.

The CI change relative to the control (unexposed) PMFs was calculated for above and below ground exposure after 12 months as
percentage change, i.e. the control CI = 100% and CI change was calculated as follows: ((CI<sub>treatment</sub> - CI<sub>control</sub>) / CI<sub>control</sub>) x 100. A positive percentage represents an increase in CI after exposure compared to the control, whereas a negative percentage represents a decrease in CI after exposure.

### 2.6.2. Analysis of morphological surface changes

Changes to PMF thickness for above and below ground films were measured using a thickness gauge (Mitutoyo, Japan) and changes to surface roughness (Ra) were measured using a surface roughness tester (Surftest SJ-210, Mitutoyo, Japan) following the ISO 4287:1997 standard with speed set to 0.5 mm s⁻¹, total measured length = 12.5 mm, λ = 2.5, and measurements split into 5 equal parts to calculate Ra. For both measurements, each sample (n = 4) was measured 5 times and sides were randomly switched throughout the sampling process, resulting in a total of 20 measurements per time point.

## 2.7. Statistical analysis and visualisation

Data analysis and visualisation were carried out using R (v.4.3.3) [45]. Graphs were created using package 'ggplot2' (v.3.5.0) [46], additionally non-metric multi-dimensional scaling (NMDS) plots based on Bray-Curtis distance were created using package 'vegan' (v.2.6-4) [47]. Differences in MP numbers, film thickness, and roughness after 12 months exposure, were analysed using one-way ANOVA with subsequent Tukey HSD tests (95 % confidence interval; *p* ≤ 0.05). Interactions between PMF type and mesh size for soil properties and microbial communities were analysed with linear mixed-effect models using package 'lmerTest' (v.3.1-3) [48], with PMF type, mesh size, and exposure time as fixed, interacting effects and position of samples in the field as random effect. Significance of fixed effects was ascertained using type III ANOVA (*p* ≤ 0.05). Models were checked for normality using QQ plots and by plotting residuals vs. fitted values. All graphs were modified using Affinity Designer (v.1.10.5.1342) (Serif Ltd., UK).

# 3. Results

## 3.1. Environmental conditions

The soil moisture content at 5 cm depth followed the precipitation pattern, with higher soil moisture during periods of high precipitation.

In contrast, soil and air temperature, as well as solar radiation followed an opposing trend, i.e. these parameters increased whilst precipitation and soil moisture decreased. Wind speed was more variable, but followed a similar trend, with higher wind speeds during colder and wetter months, and lower wind speeds during warmer, drier months (Fig. 1). The average soil surface temperature on bare control soil was slightly lower than temperature under LDPE PMFs during warmer, drier months, however there was no difference during colder, wetter months (Table S3).

## 3.2. Above and below ground PMF degradation

### 3.2.1. Changes to PMF chemical properties

Overall, stress values of all NMDS plots (Fig. 2) indicate a good to excellent fit, meaning the displayed 2-dimensional distance between sample points conforms well with the real multivariate distance between samples. The Bray-Curtis distance was used to quantify similarity of spectra based on their relative intensity values across wavenumbers, i.e. similar samples appear closer together and dissimilar samples farther apart.

For LDPE, above ground degradation altered the chemical spectra differentially to below ground degradation in either coarse or fine mesh bags. Above ground degradation caused the largest increase in the carbonyl index (CI) after 12 months (Table 2) and showed the greatest distance in NMDS plots to the control spectra (Fig. 2a). This difference between degradation methods seems largely driven by peak area changes to the C=O area, with an overall decrease after below ground exposure and slight increase after above ground exposure, as the CH peak area decreased similarly for both exposure methods (Table 3, Fig. S3a).

All biodegradable PMFs showed the largest changes in CI compared to the control after below ground exposure compared to above ground exposure, with CI increasing for PLA/PBAT and decreasing for all Bio A and B films (Table 2). Bio A and A2 presented a larger decrease in fine compared to coarse mesh bags, whereas Bio A1, B1 and B2 were relatively similar for both mesh sizes, and CI decreased more for coarse compared to fine mesh bags for Bio B (Table 2). This is supported by the distance increase in fine and coarse mesh bag data points after 12 months exposure in the corresponding NMDS plots (Fig. 2b-h). After 12 months, PLA/PBAT showed the highest decrease in C=O peak area after above > below ground degradation in fine > coarse mesh bags, and

<figure><img src="image_5.png" /><figcaption>Fig. 1. Environmental data across the 12-month study period. a) Total daily precipitation; b) Average daily soil moisture content at 5 cm depth; c) Average daily soil temperature at 5 cm depth; d) Average daily air temperature; e) Total daily solar radiation; f) Average daily wind speed.</figcaption></figure>
<figure><img src="image_6.png" /><figcaption>**Fig. 2.** Non-metric multi-dimensional scaling (NMDS) plots based on Bray-Curtis distance showing the data distribution of control (unexposed) PMFs, and PMFs after above ground (UV), and below ground degradation in coarse (Coarse) and fine (Fine) mesh bags in soil over 12 months for a) LDPE; b) PLA/PBAT; c) Bio A; d) Bio A1; e) Bio A2; f) Bio B; g) Bio B1; and h) Bio B2. Data points are expressed as mean (n = 20).</figcaption></figure>
<caption>**Table 2**</caption>

Carbonyl index (CI) and CI change relative to the control (unexposed PMF) after 12 months above and below ground degradation in coarse (Coarse) and fine (Fine) mesh bags in soil for a conventional PMF (LDPE) and 7 biodegradable PMFs. Values expressed as mean (n = 4). CI values have been rounded to two decimal places; calculations for percentages have been carried out with raw data.

<table><caption>**Table 3**</caption><thead><tr><th rowspan="2">Polymer type</th><th colspan="4">CI</th><th colspan="3">CI change relative to control [%]</th></tr><tr><th>Control</th><th>Above ground</th><th>Below ground (Coarse)</th><th>Below ground (Fine)</th><th>Above ground</th><th>Below ground (Coarse)</th><th>Below ground (Fine)</th></tr></thead><tbody><tr><td>LDPE</td><td>0.33</td><td>0.47</td><td>0.32</td><td>0.39</td><td>+ 44.44</td><td>-2.12</td><td>+ 18.62</td></tr><tr><td>PLA/PBAT</td><td>0.91</td><td>1.43</td><td>2.36</td><td>2.09</td><td>+ 56.97</td><td>+ 159.23</td><td>+ 129.56</td></tr><tr><td>Bio A</td><td>22.27</td><td>20.37</td><td>19.59</td><td>16.15</td><td>-8.51</td><td>-12.02</td><td>-27.46</td></tr><tr><td>Bio A1</td><td>22.03</td><td>21.79</td><td>19.11</td><td>19.33</td><td>-1.12</td><td>-13.26</td><td>-12.27</td></tr><tr><td>Bio A2</td><td>23.91</td><td>21.98</td><td>20.06</td><td>16.22</td><td>-8.09</td><td>-16.09</td><td>-32.16</td></tr><tr><td>Bio B</td><td>30.48</td><td>23.33</td><td>16.87</td><td>20.93</td><td>-23.48</td><td>-44.66</td><td>-31.34</td></tr><tr><td>Bio B1</td><td>30.55</td><td>23.09</td><td>16.03</td><td>16.08</td><td>-24.44</td><td>-47.52</td><td>-47.37</td></tr><tr><td>Bio B2</td><td>31.51</td><td>23.46</td><td>16.96</td><td>16.39</td><td>-25.54</td><td>-46.18</td><td>-47.97</td></tr></tbody></table>

CH and C=O peak area for control (unexposed PMF) and PMFs after 12 months above and below ground degradation in coarse (Coarse) and fine (Fine) mesh bags in soil for a conventional PMF (LDPE) and 7 biodegradable PMFs. Values expressed as mean (n = 4).

<table><thead><tr><th rowspan="2">Polymer type</th><th colspan="2">Control</th><th colspan="2">Above ground</th><th colspan="2">Below ground (Coarse)</th><th colspan="2">Below ground (Fine)</th></tr><tr><th>CH peak area</th><th>C=O peak area</th><th>CH peak area</th><th>C=O peak area</th><th>CH peak area</th><th>C=O peak area</th><th>CH peak area</th><th>C=O peak area</th></tr></thead><tbody><tr><td>LDPE</td><td>4.04</td><td>1.33</td><td>2.85</td><td>1.35</td><td>2.62</td><td>0.84</td><td>2.21</td><td>0.86</td></tr><tr><td>PLA/PBAT</td><td>18.95</td><td>17.24</td><td>6.77</td><td>9.68</td><td>6.04</td><td>14.26</td><td>5.03</td><td>10.52</td></tr><tr><td>Bio A</td><td>1.56</td><td>34.78</td><td>0.85</td><td>17.22</td><td>1.52</td><td>29.72</td><td>0.88</td><td>14.17</td></tr><tr><td>Bio A1</td><td>1.75</td><td>38.60</td><td>1.01</td><td>22.01</td><td>1.49</td><td>28.56</td><td>1.11</td><td>21.42</td></tr><tr><td>Bio A2</td><td>1.93</td><td>46.22</td><td>1.11</td><td>24.48</td><td>1.65</td><td>33.01</td><td>1.01</td><td>16.31</td></tr><tr><td>Bio B</td><td>1.47</td><td>44.72</td><td>0.82</td><td>19.05</td><td>0.93</td><td>15.63</td><td>0.79</td><td>16.53</td></tr><tr><td>Bio B1</td><td>1.68</td><td>51.30</td><td>0.83</td><td>19.09</td><td>0.82</td><td>13.16</td><td>0.65</td><td>10.61</td></tr><tr><td>Bio B2</td><td>1.57</td><td>49.57</td><td>0.88</td><td>20.59</td><td>1.18</td><td>20.00</td><td>0.84</td><td>13.69</td></tr></tbody></table>

decrease in CH peak area for fine > coarse mesh bags > above ground degradation (Table 3, Fig. S3b). Bio A, A1, and A2 displayed the highest decrease in C=O peak area after below ground degradation in fine mesh bags > above ground degradation > coarse mesh bags, whereas B, B1, and B2 showed the highest decrease in fine > coarse mesh bags > above ground degradation (Table 3, Fig. S3c-h). Regarding the decrease in CH peak area, Bio A, A1, and A2 showed a similar reduction after above ground degradation and below ground degradation in fine mesh bags, but less pronounced for coarse mesh bags (Table 3, Fig. S3c-e). Bio B, B1, and B2 showed the highest decrease in CH peak area after below ground degradation in fine mesh bags > above ground degradation > coarse mesh bags (Table 3, Fig. S3f-h).

### 3.2.2. Morphological changes, physical fragmentation, and MP generation

Overall, all buried PMFs were recovered physically intact with no visible tearing or material loss, regardless of mesh size. All above ground degraded PMFs, apart from LDPE, started to disintegrate after approx. 8 months exposure. Surface roughness of all PMF types increased significantly for both exposure methods (Table 4). Changes to film thickness occurred only for PLA/PBAT, Bio A, A2, B, and B1 PMFs, with an increase for PLA/PBAT, Bio A, and Bio B, and a decrease for Bio A2 and B1 (Table 4).

The number of MP particles detected in soil samples after extraction with the conventional method was similar between blanks, controls (soil only), and LDPE in coarse mesh bags, and only showed a significant increase for LDPE in fine mesh bags compared to blanks (p = 0.044)

<caption>**Table 4**</caption>

Surface roughness and thickness of plastic mulch films before exposure (control) and after 12 months above and below ground degradation in coarse (Coarse) and fine (Fine) mesh bags in soil for a conventional PMF (LDPE) and 7 biodegradable PMFs. Values expressed as mean ± SE (n = 4). Significant differences between control and exposed films are indicated by superscript letters for each polymer type.

<table><thead><tr><th rowspan="2">Polymer type</th><th colspan="4">Surface roughness [µm]</th><th colspan="4">Thickness [µm]</th></tr><tr><th>Control</th><th>Above ground</th><th>Below ground (Coarse)</th><th>Below ground (Fine)</th><th>Control</th><th>Above ground</th><th>Below ground (Coarse)</th><th>Below ground (Fine)</th></tr></thead><tbody><tr><td>LDPE</td><td>0.81<br>± 0.02<sup>a</sup></td><td>1.31 ± 0.13<sup>b</sup></td><td>1.12 ± 0.07<sup>ab</sup></td><td>1.28 ± 0.07<sup>b</sup></td><td>23.0 ± 0.26<sup>a</sup></td><td>23.55<br>± 0.35<sup>a</sup></td><td>23.55 ± 0.46<sup>a</sup></td><td>24.2 ± 0.19<sup>a</sup></td></tr><tr><td>PLA/PBAT</td><td>1.53<br>± 0.04<sup>a</sup></td><td>2.19 ± 0.12<sup>b</sup></td><td>2.84 ± 0.18<sup>c</sup></td><td>2.61 ± 0.13<sup>bc</sup></td><td>10.45<br>± 0.21<sup>a</sup></td><td>12.1 ± 0.41<sup>b</sup></td><td>11.65 ± 0.40<sup>ab</sup></td><td>12.05 ± 0.33<sup>b</sup></td></tr><tr><td>Bio A</td><td>0.99<br>± 0.04<sup>a</sup></td><td>2.31 ± 0.19<sup>b</sup></td><td>1.99 ± 0.18<sup>b</sup></td><td>2.16 ± 0.14<sup>b</sup></td><td>17.35<br>± 0.36<sup>a</sup></td><td>20.7 ± 0.73<sup>b</sup></td><td>23 ± 0.66<sup>c</sup></td><td>23.55 ± 0.66<sup>c</sup></td></tr><tr><td>Bio A1</td><td>1.07<br>± 0.03<sup>a</sup></td><td>1.78 ± 0.15<sup>b</sup></td><td>2.15 ± 0.19<sup>b</sup></td><td>1.96 ± 0.16<sup>b</sup></td><td>23.2 ± 0.73<sup>a</sup></td><td>22.55<br>± 1.25<sup>a</sup></td><td>21.3 ± 0.57<sup>a</sup></td><td>21.9 ± 0.77<sup>a</sup></td></tr><tr><td>Bio A2</td><td>1.06<br>± 0.04<sup>a</sup></td><td>2.57 ± 0.27<sup>b</sup></td><td>1.71 ± 0.12<sup>ac</sup></td><td>2.29 ± 0.26<sup>bc</sup></td><td>22.95<br>± 0.70<sup>ab</sup></td><td>25.0 ± 0.54<sup>a</sup></td><td>23.1 ± 0.69<sup>ab</sup></td><td>22.2 ± 0.43<sup>b</sup></td></tr><tr><td>Bio B</td><td>1.04<br>± 0.04<sup>a</sup></td><td>2.64 ± 0.19<sup>b</sup></td><td>2.22 ± 0.14<sup>b</sup></td><td>2.69 ± 0.25<sup>b</sup></td><td>21.85<br>± 0.29<sup>a</sup></td><td>20.85<br>± 0.41<sup>a</sup></td><td>24.4 ± 0.48<sup>b</sup></td><td>23.65 ± 0.53<sup>b</sup></td></tr><tr><td>Bio B1</td><td>0.97<br>± 0.05<sup>a</sup></td><td>2.51 ± 0.23<sup>b</sup></td><td>2.91 ± 0.25<sup>b</sup></td><td>3.69 ± 0.23<sup>c</sup></td><td>28.95<br>± 0.84<sup>a</sup></td><td>27.40<br>± 0.63<sup>ab</sup></td><td>25.7 ± 0.54<sup>b</sup></td><td>25.3 ± 0.59<sup>b</sup></td></tr><tr><td>Bio B2</td><td>0.99<br>± 0.06<sup>a</sup></td><td>2.31 ± 0.22<sup>b</sup></td><td>2.04 ± 0.13<sup>b</sup></td><td>3.45 ± 0.27<sup>c</sup></td><td>27.1 ± 0.33<sup>a</sup></td><td>26.6 ± 0.65<sup>a</sup></td><td>25.5 ± 0.58<sup>a</sup></td><td>27.0 ± 0.48<sup>a</sup></td></tr></tbody></table>
<figure><img src="image_7.png" /><figcaption>Fig. 3. Number of MP polymer particles $g^{-1}$ dry soil after extraction with a) the extraction method for conventional MPs; and b) for biodegradable MPs for coarse (C) and fine (F) mesh bags. Blanks contained only reagents; control samples refer to mesh bags containing only soil. Values expressed as mean ± SE (n = 4 for PMFs & control, n = 3 for blanks).</figcaption></figure>

<figure><img src="image_8.png" /><figcaption>Fig. 4. Relative abundance of archaea and bacteria after 16S rRNA metagenomic amplicon sequencing for control (soil only), PLA/PBAT, LDPE, Bio A, Bio A1, Bio A2, Bio B, Bio B1, and Bio2 PMFs after 4 (pink bars), 8 (blue bars), and 12 months (orange bars) burial in coarse mesh bags.</figcaption></figure>
(Fig. 3a). However, there was no significant difference to the control or LDPE in coarse mesh bags. The dominant polymer types across all samples were rubber > polypropylene (PP) > polytetrafluoroethylene (PTFE) > acrylonitrile butadiene styrene (ABS) > polyethylene (PE) > others (Table S4).

Similarly, soils extracted for biodegradable microplastics showed no significant difference in MP number, except for control soil in fine mesh bags, which had a higher particle count compared to controls in coarse mesh bags (p = 0.031) and PLA/PBAT in coarse mesh bags (p = 0.04). No soil in direct contact with any biodegradable PMF showed increased polymer particle numbers compared to the controls (Fig. 3b), however there was an observable increase in PLA and PBAT particles which were not found in blanks or controls (Table S5). The dominant biodegradable polymer types detected across the majority of treatments were PHBV > PBAT > blend of PLA/PBAT > PLA > PAAU (Table S5).

### 3.3. PMF type has no effect on soil properties

PMF type had no significant effect on any soil properties across the 12-month period (Fig. S4). Exposure time (4, 8, 12 months) significantly impacted pH, EC, NO₃, PO₄, and DOC (p < 0.001 for all), and had no effect on NH₄ concentrations. Mesh size significantly affected PO₄ (p < 0.001) and DOC (p = 0.005) concentrations, and the interaction between time and mesh size significantly affected pH (p < 0.001) and EC (p = 0.02) values.

## 3.4. Soil microbial community composition is not affected by PMF type

### 3.4.1. Changes to relative abundance of archaea and bacteria are mainly driven by time and mesh size

Overall, exposure time, mesh size, and PMF type, affected the relative abundance of the majority of taxa in the following order: time (63.5 %) > interaction of time and mesh size (36.5 %) > mesh size (28.6 %) > interaction of PMF type and mesh size (4.8 %) > interaction of PMF type and time (3.2 %) = interaction of PMF type, mesh size, and time (3.2 %) > PMF type (1.6 %) (Figs. 4, 5, Table S6). The relative abundance of *Nitrososphaerales*, *Acidobacteria*, *Acidimicrobiia*, Other *Frankiales*, *Marmoricola*, *Nocardioides*, Other *Propionibacterales*, *Rubrobacter*, *Conexibacter*, Other *Gaeillales*, Other *Thermoleophilia*, *Bacteroidia*, *Chloroflexota*, *Gemmatimonas*, Other *Gemmatimonadota*, *Gemmatales*, Other *Rhizobiales*, *Sphingomonas*, Other *α-Proteobacteria*, *Candidatus Udaeobacter*, and *Pedosphaerales* increased from 4 to 12 months, and decreased for *Subgroup 5*, *Vicinamibacterales*, *Vicinamibacteria*, Other *Acidobacteriota*, *Mycobacterium*, Other *Actinobacteriota*, *Bacillus*, Other *Bacillales*, *Latescibacteriota*, *Polyangiales*, *Methyloligellaceae*, *Mesorhizobium*, *Bradyrhizobium*, *Pseudolabrys*, *Candidatus Alysiosphaera*, *Variovorax*, *Ellin 6067*, Other *γ-Proteobacteria*, and Other *Bacteria* (Figs. 4, 5, Table S6). The relative abundance in fine mesh bags compared to coarse mesh bags was higher in *Subgroup 5*, *Vicinamibacteria*, *Chloroflexota*, *Polyangiales*, *Pseudolabrys*, Other *α-Proteobacteria*, Other *Burkholderiales*, Other *Bacillales*, and Other *Bacteria*, and vice versa for Other *Frankiales*,

<figure><img src="image_9.png" /><figcaption>**Fig. 5.** Relative abundance of archaea and bacteria after 16S rRNA metagenomic amplicon sequencing for control (soil only), PLA/PBAT, LDPE, Bio A, Bio A1, Bio A2, Bio B, Bio B1, and Bio2 PMFs after 4 (pink bars), 8 (blue bars), and 12 months (orange bars) burial in fine mesh bags.</figcaption></figure>
Other *Micrococcales*, *Marmoricola*, Other *Propionibacteriales*, *Bacterioidia*, *Bacillus*, *Mesorhizobium*, *Bradyrhizobium*, Other *Rhizobiales*, and *Massilia*. PMF type only affected *Variovorax*, with overall higher abundance in Bio A, A1, B, B1, and B2 compared to the control, and lower abundance in Bio A2, PLA/PBAT and lowest in LDPE. The interaction of PMF type and time had an effect on *Streptomyces*, with a relative abundance increase from 4 to 12 months in Bio A, A1, A2, B, B2, and LDPE, and a decrease in Bio B, control, and PLA/PBAT, and on *Mesorhizobium* with an increase in Bio A1, A2, control, and LDPE, and a decrease in Bio A, B, B1, B2, and PLA/PBAT. Overall, species richness and microbial diversity (Shannon and Simpson index) showed no significant difference between treatments (Table S7). Rarefaction curves indicate sufficient coverage of taxa and sequencing depth (Fig. S5, S6).

### 3.4.2. Changes to relative abundance of fungi are driven by exposure time and mesh size

Similarly, relative abundance of fungi was affected in the following order: time (59.4 %) > interaction of time and mesh size (43.8 %) > mesh size (15.6 %) > interaction of PMF type and mesh size (3.1 %) = interaction of PMF type, mesh size, and time (3.1 %), and no effect for any taxa for PMF type and interactions of PMF type and time (Figs. 6, 7, Table S8). The relative abundance of *Didymellaceae*, *Fusarium oxysporum*, *Ramophialophora humicola*, Other *Sordariales*, *Mortierella*, and Other *Fungi* increased from 4 to 12 months, and decreased for *Paraphaeosphaeriaorulosa*, Other *Pleosporales*, *Tubeufiaceae*, *Exophiala equina*, Other *Eurotiomycetes*, *Gibellulopsis nigrescens*, *Keithomyces carneus*,

*Trichoderma*, Other *Fusarium*, Other *Nectriaceae*, Other *Asomycota*, Other *Mortierellaceae*. The relative abundance in fine mesh bags compared to coarse mesh bags was higher for *Xanthothecium peruvianum* and *Gibellulopsis nigrescens*, and vice versa for *Didymellaceae*, *Keithomyces carneus*, and Other *Mortierellaceae*. Species richness and Shannon diversity index differed significantly, mainly driven by time and mesh size, whilst Simpson diversity index remained similar (Table S7). Rarefaction curves indicate sufficient coverage of taxa and sequencing depth (Fig. S7, S8).

## 4. Discussion

### 4.1. PMF chemical degradation, physical fragmentation, and MP generation

#### 4.1.1. Additives and polymer blends affect PMF chemical degradation rates

Abiotic degradation, particularly affecting above ground PMFs, is mainly facilitated by photo or thermal oxidation and hydrolysis, whereas biotic degradation (above and below ground) is driven by formation of biofilms and the release of enzymes (e.g. hydrolases, lacases, esterases) by the microbial community, breaking the polymer chains and eventually assimilating and mineralising the polymer [12]. For LDPE, the slight increase in the C=O peak area following above ground degradation indicates that oxidation of the polymer has occurred, creating degradation by-products like ketones, aldehydes, and carboxylic acids [10,49]. Abiotic degradation of PLA/PBAT is mainly

<figure><img src="image_10.png" /><figcaption>**Fig. 6.** Relative abundance of fungi after ITS2 metagenomic amplicon sequencing for control (soil only), PLA/PBAT, LDPE, Bio A, Bio A1, Bio A2, Bio B, Bio B1, and Bio2 PMFs after 4 (pink bars), 8 (blue bars), and 12 months (orange bars) burial in coarse mesh bags.</figcaption></figure>
**Asomycota**

<figure><img src="image_11.png" /><figcaption>**Fig. 7.** Relative abundance of fungi after ITS2 metagenomic amplicon sequencing for control (soil only), PLA/PBAT, LDPE, Bio A, Bio A1, Bio A2, Bio B, Bio B1, and Bio2 PMFs after 4 (pink bars), 8 (blue bars), and 12 months (orange bars) burial in fine mesh bags.</figcaption></figure>

facilitated via chain scission and photocleavage of ester bonds, and bi-
otic degradation via enzymatic hydrolysis, breaking PLA and PBAT into
oligomers and monomers. Overall, the above ground degradation of all
biodegradable PMFs tested in this study showed a similar pattern in
terms of decreasing C=O and CH peak areas after 12 months, though to
differing degrees. This is attributed to the similarity in degradation
mechanisms affecting PLA, PBAT, PHBV, and PAAU [50]. UV exposure
can initially lead to amplification of the C=O peak intensity by creating
radicals and unsaturated fragments through chain scission at existing
ester and urethane sites within the polymer [51]. Whilst this does not
directly generate new C=O groups, secondary oxidation of these groups
can create aldehydes, ketones, and carboxylic acids, thereby amplifying
the C=O peak intensity during early stages of degradation [50,52].
However, prolonged exposure to weathering has likely led to the be-
ginnings of surface ablation and leaching of low-molecular-weight oxi-
dised fragments, reducing both C=O and CH peak intensities at later
stages of degradation [10].

The bespoke PMFs containing only one polymer type (PBAT), Bio A,
A1, and A2, showed the highest degradation in fine mesh bags, followed
by above ground degradation, and coarse mesh bags for both C=O and
CH decrease. It is evident that mesh size affected the degradation rate of
these PMFs; one possibility could be that the interaction of meso- and
macrofauna has affected the microbial community composition, as we
have shown mesh size to have the second largest effect on relative
abundance, aside from exposure time. The presence of macrofauna can

lead to a selective microbial population by inhibiting or promoting
certain taxa [53,54], which can in turn affect the degradation rate of the
polymer. Additionally, the interaction of macrofauna with the polymer
itself can affect degradation rates [55]. Bio A degraded more above
ground than Bio A1 and A2, which is likely due to the lack of a thermal
stabiliser and antioxidant. Bio A2 degraded slightly faster than Bio A1,
which may be due to the presence of an antioxidant in Bio A1, slowing
degradation. However, Bio A1 also contained two rather than only one
catalyst, promoting degradation, suggesting that the presence of the
antioxidant outweighs the effect of two catalysts. As Bio A1 contained
slightly less PBAT compared to Bio A2, a higher degradation would have
been expected in Bio A1, which shows that the polymer percentage was
likely not the driver for the degradation rate difference between those
two PMFs, but rather the additives.

The PMFs containing a blend of three polymer types (PBAT, PHBV, PAAU), Bio B, B1, and B2, likely demonstrated a greater decrease in C=O peak area following below ground exposure compared to above ground exposure due to enzymatic hydrolysis of ester bonds and subsequent metabolization of degradation by-products. Similar to Bio A, A1, and A2 films, mesh size impacted the CH peak area decrease and was slowest in the presence of meso- and macrofauna. Interestingly, Bio B2 showed the slowest degradation overall across the three blends despite the two catalysts. Bio B1 degraded faster than Bio B below ground, but they performed similarly above ground, potentially indicating that the thermal stabiliser, antioxidant, and catalyst in Bio B1 provided a balance
under above ground conditions. However, this balance did not translate into below ground conditions, suggesting that the thermal stabiliser and antioxidant (mainly for abiotic degradation resistance) play a lesser role, and that the catalyst was able to increase degradation. It is also possible that the composition of the polymer blends and interaction with stabilisers and catalysts influenced the degradation rate, as it has been previously shown that pure materials undergo more rapid degradation compared to their blends, and that the polymer ratio can impact degradation rates [56].

Overall, these results indicate that the chemical degradation of biodegradable PMF blends used in this study progresses slowly and that polymer ratio, added stabilisers, and catalysts influence the degradation process in soil. Whilst these additives provide the PMFs with certain desired properties, e.g. decreased degradation rate under abiotic conditions through antioxidants and thermal stabilisers, or increased degradation rate through catalysts, optimising them to last for their intended lifespan, we have shown that these additives, especially when unknown, could have unintended effects on PMF degradation once incorporated into the soil. It is also noteworthy that the colour of PMFs can affect degradation rates, with black PMFS showing slightly lower degradation compared to white PMFs [57]. Due to the variety in polymer types and additives in the PMFs used in this study, this was however not investigated.

### 4.1.2. Chemical degradation does not equal physical fragmentation and MP generation

The observed above ground physical fragmentation was mainly attributed to impact of external mechanical forces, and loss of material to wind dispersal rather than chemical degradation. The below ground degraded PMF pieces were recovered without visible signs of fragmentation although changes to surface morphology were detected, which highlights the importance of differentiation between chemical degradation and physical fragmentation when considering the biodegradability of PMFs. During the initial stages of degradation, fragmentation only occurred through external mechanical forces, and the analysis of chemical degradation rate of PMFs suggests that material loss is not due to complete degradation of the polymers but rather environmental impact.

Additionally, our results indicate that there is no detectable formation of MPs for any PMF type after 12 months burial in the soil, regardless of only micro-, or micro-, meso-, and macrofauna interactions, i.e. there was no evidence of physical fragmentation of PMFs. Based on previous studies, we anticipated a higher MP level in soil surrounding biodegradable PMFs compared to LDPE [57]. The lack of increase in PE particles in soil from LDPE mesh bags, compared to controls and blanks, confirms that physical fragmentation of LDPE did not occur to a detectable level, and that degradation is occurring at a slow rate. It is noteworthy, that the thickness of the LDPE PMF used in this study was higher than the average PMF thickness of many LDPE PMFs used in other countries (e.g. China) [8]. This could certainly play a role in the fragmentation and ablation process generating MPs [9]. The higher particle number in LDPE in fine mesh bags was largely driven by rubber particles, which was the dominant conventional polymer, and likely attributed to deposition from regular farm practices. For biodegradable PMFs, the presence of PLA and PBAT particles in PLA/PBAT, Bio A, A1, A2, B, and B1 could indicate some formation of microplastic particles, potentially through early stages of surface ablation [9], although there was no overall difference in particle numbers between controls and PMF treatments. The leading polymer type, PHBV, was detected in similar quantities across all PMF types and controls, although only Bio B, B1, and B2 contained PHBV. This suggests that the PHBV particles likely originated from a different source (e.g. a microplastic experiment utilising PHBV in a neighbouring field) and have been deposited through wind dispersal.

### 4.1.3. Degradation of biodegradable PMFs is much slower than anticipated

Our findings indicate that fragmentation and subsequent generation of MP particles in soil is not linked to interaction with micro-, meso-, or macrofauna, and that microplastic generation requires prior mechanical stress on the PMF material leading to the initial fragmentation. This underpins the importance of differentiating between fragmentation and degradation, as these can be both independent and interactive processes. The below ground degradation and change to chemical spectra after 12 months of burial in soil was not sufficient to fragment the PMFs and generate MP particles. These findings highlight the importance of reframing current biodegradability standards, treating degradation as a system property, which is influenced by environmental factors and material properties. We acknowledge that this study used six biodegradable PMFs, which were developed for improved biodegradability, but not yet commercially available, and did purposefully not implement standardised biodegradability tests. The realistic in-field approach used here however, raises questions regarding the applicability of current standards for the assessment of degradation of biodegradable PMFs in agricultural soils. Farmers are actively encouraged to plough biodegradable PMFs into the soil after use, however we have shown that in the environmental conditions and soil type of this study, macroplastic PMF pieces left in the soil do not degrade and fragment at the proposed standard degradation rate (≥ 90% after 24 months [6]). This could lead to an accumulation of macroplastic in soil over time, further slowing the degradation process [7] and posing the issue of pollution swapping and release and accumulation of additives and other added substances [8].

Whilst we acknowledge that the buried PMFs in this study have not undergone prior above ground exposure, which would usually occur for the duration of several months (i.e. one growing season) and start the degradation process, increasing surface roughness and thereby surface area for microbes to attach, and making C more bioavailable for assimilation [10,58,59], we have also shown that above ground degradation for the biodegradable PMFs used in this study progresses slowly, and that fragmentation of above ground exposed PMFs exceeds the duration of one growing season. We therefore assume that had the same PMFs been degraded above ground for several months prior to soil burial, the observed chemical degradation would have likely increased, and potentially more MP particles would have been released into the soil [10], however as current biodegradation standards are based on new, non-degraded polymers, we believe it vital to reframe current standards for the classification of biodegradable polymers according to climatic conditions, environmental factors, and polymer types.

## 4.2. PMF type has no impact on relative abundance of bacteria and fungi

The overall negligible effect of PMF type on the relative abundance of archaea, bacteria, and fungi indicates that the polymers, regardless of type, were not utilised as a main C source by any taxa. This could be due to the low level of observed degradation and lack of fragmentation of the PMFs in soil, as well as no prior above ground exposure of buried PMFs, making them less bioavailable. Although many studies have reported significant shifts in the microbial community after plastic addition [60–62], our findings align well with other field studies which have observed no significant effect of LDPE, PLA/PBAT, PE, or PHBV at realistic concentrations on soil microbial community structure under the climatic conditions and soil type used in this study [63–65]. We attribute our neutral results mainly to a longer time span, a realistic environmental setting, and realistic macroplastic concentrations in the soil, which is often not possible to replicate in mesocosm settings. Here, time was the main driver for change in relative abundance, followed by mesh size, either independently or in conjunction with time. However, bacterial taxa known to degrade plastic (e.g. *Bacillus* [12]) decreased in abundance after 12 months, indicating that the PMFs were not a significant C source. Although *Streptomyces*, known to degrade PBAT [12, 66], increased in relative abundance over time for some PBAT-containing PMFs, it also decreased for others, resulting in no clear
pattern. Similarly, fungi like *Fusarium oxysporum*, Other *Fusarium*, *Tri-choderma*, and *Cladosporium*, which have been previously linked to plastic degradation [12,67-69], have shown opposing patterns in their relative abundance over time and no link to PMF type. This was also reflected in the lack of effect of PMF type on soil chemical properties and nutrient levels. With microbial degradation of polymers in soil, a change in DOC caused by the breakdown of C chains and release of bioavailable C, particularly for biodegradable PMFs, would be anticipated [70,71], however this was not observed here, further supporting that microbial degradation was neither supressed nor promoted by any particular PMF type. Overall, our findings indicate that PMF type had no impact on relative abundance at a realistic concentration under the climatic conditions and soil type of this study, and that changes in microbial and fungal communities were mainly driven by time. This suggests that below ground degradation in these environmental conditions is very slow and likely does not meet current standards and regulation for biodegradability.

## 5. Conclusion

In this field study, we have assessed the above and below ground (with and without meso- and macrofauna interactions in soil) degradation rate of one conventional LDPE and seven biodegradable PMFs composed of PLA, PBAT, PHBV, and PAAU, and evaluated their effects on soil properties and microbial community structure over 12 months in a temperate climate.

We anticipated the degradation and fragmentation rate of biodegradable PMFs to be faster compared to LDPE, subsequently resulting in an increased MP concentration in the surrounding soil. Additionally, we anticipated the interaction of meso- and macrofauna to accelerate physical fragmentation, and subsequently chemical degradation compared to only microfauna interactions. However, whilst the biodegradable PMFs generally showed a higher level of degradation compared to LDPE, their degradation rate was still slower than expected, and we observed no physical fragmentation of below ground degraded PMFs, regardless of presence of meso- and macrofauna. The lack of physical fragmentation of PMFs buried in soil underpins the importance of differentiating between fragmentation and degradation of polymers, as these terms are often used synonymously. The lack of physical fragmentation is likely the cause for no overall difference in MP numbers, indicating that 12 months of degradation in soil without application of external mechanical forces is not sufficient to physically fragment PMFs, but will result in changes to chemical spectra and surface morphology.

Further, we hypothesised that the PMF type differentially affects soil nutrients, particularly soil DOC. Our results however, showed that PMF type had no effect on the measured soil properties, but that changes were driven by time and mesh size instead.

Lastly, we anticipated PMF type to influence the soil microbial community composition. However, we found PMF type to have a negligible effect on microbial community structure under the climatic conditions and soil type of this study. The main factors driving change in relative abundance of bacteria and fungi were time, followed by the potential of meso- and macrofauna interactions. The presence of meso- and macrofauna led to a decrease in degradation rate potentially due to selective pressure exhibited on the microbial community. However, we conclude that in this study realistic concentrations of macroplastic in this soil type had no adverse or promotional effects on microbial community structure.

Overall, we conclude that the observed degradation rate of biodegradable PMFs in this study was much slower than expected. The current biodegradability standard proposes a degradation rate of ≥ 90% over 24 months under specific conditions. Despite the fact that our study was conducted for 12 months in an open environment and considering that degradation does not occur in a linear fashion, we believe that our results strongly highlight the need to reframe degradation as a system property in an open environment, as this level of degradation would

likely not have been reached by any of the tested PMFs. Biodegradability assessments must account for realistic environmental conditions, soil types, and polymer types and sizes, to improve the sustainability and adoption rate of biodegradable PMFs. A failure to adapt standards to environmental conditions and material properties could lead to the risk of pollution swapping through an accumulation of biodegradable macroplastic in agricultural soil, which subsequently decreases degradation rates and increases risk of accumulation and release of additives and other substances into the soil.

## Environmental Implication

Biodegradable plastic mulch films (PMFs) are gaining popularity as a more sustainable alternative to conventional PMFs. However, the knowledge regarding their impact and degradation under field conditions is still limited. Here, we have shown the importance of treating PMF degradation as a system property and highlighted the importance of reframing standards according to realistic environmental conditions, soil types, and polymer types and sizes, to improve the sustainability and adoption rate of biodegradable PMFs. A failure to adapt standards could lead to the risk of pollution swapping through an accumulation of biodegradable macroplastic in agricultural soil.

## CRediT authorship contribution statement

**Martine Graf:** Writing – review & editing, Writing – original draft, Visualization, Validation, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. **Estelle Choiselat:** Writing – review & editing, Investigation. **Michaela K. Reay:** Writing – review & editing. **Rafael Bargiela:** Visualization, Validation, Data curation. **Athanasios Dimitriou:** Writing – review & editing, Methodology. **Qiuyun Liu:** Resources, Conceptualization. **Robert M. Elias:** Writing – review & editing, Resources, Conceptualization. **Peter N. Golyshin:** Writing – review & editing. **Robert Griffiths:** Writing – review & editing. **David R. Chadwick:** Writing – review & editing, Supervision, Funding acquisition, Conceptualization. **Davey L. Jones:** Writing – review & editing, Supervision, Funding acquisition, Conceptualization.

## Declaration of Competing Interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

## Acknowledgements

This study was part of a project funded by the UK Natural Environment Research Council Global Challenges Research Fund programme on Reducing the Impacts of Plastic Waste in Developing Countries (NE/ V005871/1). The authors thank Benjamin I. Collins and Gwion B. Williams for their support in sample processing, and Gary Ogden (Wells Performance Materials Ltd., Staffordshire, UK) for the formulation and provision of bespoke biodegradable PMF blends.

## Appendix A. Supporting information

Supplementary data associated with this article can be found in the online version at doi:10.1016/j.jhazmat.2025.139871.

## Data availability

Data will be made available on request.
## References

[1] FAO, 2021. Assessment of agricultural plastics and their sustainability. A call for action. Rome.

[2] Plastics Europe, 2022. Plastics-the facts 2022.

[3] Huerta Lwanga, E., Mendoza Vega, J., Ku Quej, V., Chi J de los, A., Sanchez del Cid, L., Chi, C., et al., 2017. Field evidence for transfer of plastic debris along a terrestrial food chain. Sci Rep 7, 14071. https://doi.org/10.1038/s41598-017-14588-2.

[4] Griffin-LaHue, D., Ghimire, S., Yu, Y., Scheenstra, E.J., Miles, C.A., Flury, M., 2022. In-field degradation of soil-biodegradable plastic mulch films in a Mediterranean climate. Sci Total Environ 806, 150238. https://doi.org/10.1016/j.scitotenv.2021.150238.

[5] Zhang, H., Flury, M., Miles, C., Liu, H., Devetter, L., 2020. Soil-biodegradable plastic mulches undergo minimal in-soil degradation in a perennial raspberry system after 18 months. Horticulturae 6, 47. https://doi.org/10.3390/horticulturae6030047.

[6] European Commission, 2022. EU Policy Framework on Biobased, Biodegradable and Compostable plastics. COM(2022) 682 final. Brussels.

[7] Reay, M.K., Graf, M., Greenfield, L.M., Bargiela, R., Onyije, C., Lloyd, C.E.M., et al., 2025. Microbial degradation of bioplastic (PHBV) is limited by nutrient availability at high microplastic loadings. Environmental Science Advances 4, 133-146. https://doi.org/10.1039/d4va00311j.

[8] Reay, M.K., Graf, M., Murphy, M., Li, G., Yan, C., Bhattacharya, M., et al., 2025. Higher potential leaching of inorganic and organic additives from biodegradable compared to conventional agricultural plastic mulch film. J Hazard Mater 488, 137147. https://doi.org/10.1016/j.jhazmat.2025.137147.

[9] Zhang, K., Hamidian, A.H., Tubić, A., Zhang, Y., Fang, J.K.H., Wu, C., et al., 2021. Understanding plastic degradation and microplastic formation in the environment: a review. Environ Pollut 274, 116554. https://doi.org/10.1016/j.envpol.2021.116554.

[10] Chamas, A., Moon, H., Zheng, J., Qiu, Y., Tabassum, T., Jang, J.H., et al., 2020. Degradation rates of plastics in the environment. ACS Sustain Chem Eng 8, 3494-3511. https://doi.org/10.1021/acssuschemeng.9b06635.

[11] Beltrán-Sanahuja, A., Benito-Kaesbach, A., Sánchez-García, N., Sanz-Lázaro, C., 2021. Degradation of conventional and biobased plastics in soil under contrasting environmental conditions. Sci Total Environ 787, 147678. https://doi.org/10.1016/j.scitotenv.2021.147678.

[12] Mendoza, J.E., Tineo, D., Chuquibala-Checan, B., Atalaya-Marin, N., Taboada-Mitva, V.H., Tafur-Culqui, J., et al., 2024. Comparative perspectives on the biodegradation of LDPE in agricultural systems. Front Microbiol 15, 1510817.

[13] Shan, X., Zhang, W., Dai, Z., Li, J., Mao, W., Yu, F., et al., 2022. Comparative analysis of the effects of plastic mulch films on soil nutrient, yields and soil microbiome in three vegetable fields. Agronomy 12, 506. https://doi.org/10.3390/agronomy12020506.

[14] Chen, N., Li, X., Simünek, J., Shi, H., Hu, Q., Zhang, Y., 2021. Evaluating the effects of biodegradable and plastic film mulching on soil temperature in a drip-irrigated field. Soil Tillage Res 213, 105116. https://doi.org/10.1016/j.still.2021.105116.

[15] Monkley, C., Reay, M.K., Evershed, R.P., Lloyd, C.E.M., 2024. Mass spectral characterisation of cyclic oligoesters in a biodegradable mulch film. Rapid Commun Mass Spectrom 38, e9726. https://doi.org/10.1002/rcm.9726.

[16] IUSS Working Group WRB, 2015. World reference base for soil resources 2014, update 2015. International soil classification system for naming soils and creating legends for soil maps. World soil resources reports no. 106. FAO, Rome.

[17] Bradford, M.A., Tordoff, G.M., Eggers, T., Jones, T.H., Newington, J.E., 2002. Microbiota, fauna, and mesh size interactions in litter decomposition. Oikos 99, 317-323. https://doi.org/10.1034/j.1600-0706.2002.990212.x.

[18] Bokhorst, S., Wardle, D.A., 2013. Microclimate within litter bags of different mesh size: implications for the "arthropod effect" on litter decomposition. Soil Biol Biochem 58, 147-152. https://doi.org/10.1016/j.soilbio.2012.12.001.

[19] Mulvaney, R.L., 1996. Nitrogen—Inorganic forms. In: Sparks, D.L., Page, A.I., Helmke, P.A., Loeppert, R.H., Soltanpour, P.N., Tabatabai, M.A., Johnston, C.T., Summer, M.E. (Eds.), Methods of Soil Analysis: Part 3 Chemical Methods, 5, pp. 1123-1184.

[20] Miranda, K.M., Espey, M.G., Wink, D.A., 2001. A rapid, simple spectrophotometric method for simultaneous detection of nitrate and nitrite. Nitric Oxide 5, 62-71. https://doi.org/10.1006/niox.2000.0319.

[21] Vaz, M.D.R., Edwards, A.C., Shand, C.A., Cresser, M.S., 1994. Changes in the chemistry of soil solution and acetic-acid extractable p following different types of freeze/thaw episodes. Eur J Soil Sci 45, 353-359. https://doi.org/10.1111/j.1365-2389.1994.tb00519.x.

[22] Ren, S., Graf, M., Wang, K., Zhang, J., Zhang, H., Liu, X., et al., 2025. Separation and identification of conventional microplastics from farmland soils. J Vis Exp 217, e67064. https://doi.org/10.3791/67064.

[23] Li, S., Ding, F., Flury, M., Wang, J., 2023. Dynamics of macroplastics and microplastics formed by biodegradable mulch film in an agricultural field. Sci Total Environ 894, 164674. https://doi.org/10.1016/j.scitotenv.2023.164674.

[24] Fadrosh, D.W., Bing Ma, P.G., Sengamalay, N., Ott, S., Brotman, R.M., Ravel, J., 2014. An improved dual-indexing approach for multiple 16S rRNA gene sequencing on the illumina MiSeq platform. Microbiome 2, 6. https://doi.org/10.1186/2049-2618-2-6.

[25] Korzhenkov, A.A., Toshchakov, S.V., Bargiela, R., Gibbard, H., Ferrer, M., Teplyuk, A.V., et al., 2019. Archaea dominate the microbial community in an ecosystem with low-to-moderate temperature and extreme acidity. Microbiome 7. https://doi.org/10.1186/s40168-019-0623-8.

[26] Schmieder, R., Lim, Y.W., Rohwer, F., Edwards, R., 2010. TagCleaner: identification and removal of tag sequences from genomic and metagenomic datasets. BMC Bioinforma 11, 341. https://doi.org/10.1186/1471-2105-11-341.

[27] Bolyen, E., Rideout, J.R., Dillon, M.R., Bokulich, N.A., Abnet, C.C., Al-Ghalith, G. A., et al., 2019. Reproducible, interactive, scalable and extensible microbiome data science using QIIME 2. Nat Biotechnol 37, 1091. https://doi.org/10.1038/s41587-019-0209-9.

[28] Callahan, B.J., McMurdie, P.J., Rosen, M.J., Han, A.W., Johnson, A.J.A., Holmes, S. P., 2016. DADA2: High-resolution sample inference from illumina amplicon data. Nat Methods 13, 581-583. https://doi.org/10.1038/nmeth.3869.

[29] Cowger, W., Steinmetz, Z., Gray, A., Munno, K., Lynch, J., Hapich, H., et al., 2021. Microplastic spectral classification needs an open source community: open specy to the rescue! Anal Chem 93, 7543-7548. https://doi.org/10.1021/acs.analchem.1c00123.

[30] Almond, J., Sugumaar, P., Wenzel, M.N., Hill, G., Wallis, C., 2020. Determination of the carbonyl index of polyethylene and polypropylene using specified area under band methodology with ATR-FITR spectroscopy. EPolym 20, 369-381. https://doi.org/10.1515/epoly-2020-0041.

[31] Tofa, T.S., Ye, F., Kunjali, K.L., Dutta, J., 2019. Enhanced visible light photodegradation of microplastic fragments with plasmonic Catalysts 9, 819. https://doi.org/10.3390/catal9100819.

[32] Gardette, M., Perthue, A., Gardette, J.L., Janeska, T., Földes, E., Pukánszky, B., et al., 2013. Photo- and thermal-oxidation of polyethylene: comparison of mechanisms and influence of unsaturation content. Polym Degrad Stab 98, 2383-2390. https://doi.org/10.1016/j.polymdegradstab.2013.07.017.

[33] Gulmine, J.V., Janissek, P.R., Heise, H.M., Akcelrud, L., 2002. Polyethylene characterization by FTIR. Polym Test 21, 557-563. https://doi.org/10.1016/S0142-9418(01)00124-6.

[34] Tofa, T.S., Kunjali, K.L., Paul, S., Dutta, J., 2019. Visible light photocatalytic degradation of microplastic residues with zinc oxide nanorods. Environ Chem Lett 17, 1341-1346. https://doi.org/10.1007/s10311-019-00859-z.

[35] Singh, R.K., Ruj, B., Sadhukhan, A.K., Gupta, P., 2020. Thermal degradation of waste plastics under non-sweeping atmosphere: part 2: effect of process temperature on product characteristics and their future applications. J Environ Manag 261, 110112. https://doi.org/10.1016/j.jenvman.2020.110112.

[36] Siyamak, S., Ibrahim, N.A., Abdolmohammadi, S., Wan Yunus, W.M.Z., Rahman, M.Z.A.B., 2012. Effect of fiber esterification on fundamental properties of oil palm empty fruit bunch fiber/poly(butylene adipate-co-terephthalate) biocomposites. Int J Mol Sci 13, 1327-1346. https://doi.org/10.3390/ijms13021327.

[37] Pietrosanto, A., Scarfato, P., Di Maio, L., Nobile, M.R., Incarnato, L., 2020. Evaluation of the suitability of poly(lacticide)/poly(butylene adipate-co-terephthalate) blown films for chilled and frozen food packaging applications. Polymers 12, 804. https://doi.org/10.3390/POLYM12040804.

[38] Al-Itry, R., Lamnawar, K., Maazouz, A., 2012. Improvement of thermal stability, rheological and mechanical properties of PLA, PBAT and their blends by reactive extrusion with functionalized epoxy. Polym Degrad Stab 97, 1898-1914. https://doi.org/10.1016/j.polymdegradstab.2012.06.028.

[39] Copinet, A., Bertrand, C., Govindin, S., Coma, V., Couturier, Y., 2004. Effects of ultraviolet light (315 nm), temperature and relative humidity on the degradation of polylactic acid plastic films. Chemosphere 55, 763-773. https://doi.org/10.1016/j.chemosphere.2003.11.038.

[40] Barragán, D.H., Pelacho, A.M., Martin-Closas, L., 2016. Degradation of agricultural biodegradable plastics in the soil under laboratory conditions. Soil Res 54, 216-224. https://doi.org/10.1071/SR15034.

[41] Qi, R., Jones, D.L., Liu, Q., Liu, Q., Li, Z., Yan, C., 2021. Field test on the biodegradation of poly(butylene adipate-co-terephthalate) based mulch films in soil. Polym Test 93, 107009. https://doi.org/10.1016/j.polymertesting.2020.107009.

[42] Porras, M.A., Cubitto, M.A., Villar, M.A., 2016. A new way of quantifying the production of poly(hydroxyalkanoate) using FTIR. J Chem Technol Biotechnol 91, 1240-1249. https://doi.org/10.1002/jctb.4713.

[43] Cai, Y., Lv, J., Feng, J., 2013. Spectral characterization of four kinds of biodegradable plastics: poly (Lactic Acid), poly (Butylenes Adipate-Co-Terephthalate), poly (Hydroxybutyrate-Co-Hydroxyvalerate) and poly (Butylenes Succinate) with FTIR and Raman spectroscopy. J Polym Environ 21, 108-114. https://doi.org/10.1007/s10924-012-0534-2.

[44] Ruggero, F., Carretti, E., Gori, R., Lotti, T., Lubello, C., 2020. Monitoring of degradation of starch-based biopolymer film under different composting conditions, using TGA, FTIR and SEM analysis. Chemosphere 246, 125770. https://doi.org/10.1016/j.chemosphere.2019.125770.

[45] R Core Team, 2024. R: A language and environment for statistical computing.

[46] Wickham H., 2016. ggplot2: Elegant Graphics for Data Analysis.

[47] Oksanen, J., Simpson, G., Blanchet, F., Kindt, R., Legendre, P., Minchin, P., et al., 2022. Vegan Community Ecol Package.

[48] Kuznetsova, A., Brockhoff, P., Christensen, R., 2017. Lmertester package: tests in linear mixed effects models. J Stat Softw 82, 1-26. https://doi.org/10.18637/jss.v082.i13.

[49] Graf, M., Reay, M.K., Dimitriou, A., Chadwick, D.R., Jones, D.L., 2025. Assessment of four artificial methods for aging plastic mulch films according to efficiency, rate, and similarity to natural Field-Aged plastics. J Polym Environ 33, 1542-1560. https://doi.org/10.1007/s10924-024-03481-5.

[50] Hayes, D.G., Wadsworth, L.C., Sintim, H.Y., Flury, M., English, M., Schaeffer, S., et al., 2017. Effect of diverse weathering conditions on the physicochemical properties of biodegradable plastic mulches. Polym Test 62, 454-467. https://doi.org/10.1016/j.polymertesting.2017.07.027.
[51] Zhao, S., Liu, L., Li, C., Zheng, H., Luo, Y., Pang, L., et al., 2022. Photodegradation of biobased polymer blends in seawater: a major source of microplastics in the marine environment. Front Mar Sci 9, 1046179. https://doi.org/10.3389/fmars.2022.1046179.

[52] Xie, F., Zhang, T., Bryant, P., Kurusingal, V., Colwell, J.M., Laycock, B., 2019. Degradation and stabilization of polyurethane elastomers. Prog Polym Sci 90, 211–268. https://doi.org/10.1016/j.progpolymsci.2018.12.003.

[53] Scheu, S., Schlitt, N., Tiunov, A.V., Newington, J.E., Jones, T.H., 2002. Effects of the presence and community composition of earthworms on microbial community functioning. Oecologia 133, 254–260. https://doi.org/10.1007/s00442-002-1023-4.

[54] Medina-Sauza, R.M., Álvarez-Jiménez, M., Delhal, A., Reverchon, F., Blouin, M., Guerrero-Analco, J.A., et al., 2019. Earthworms building up soil microbiota, a review. Front Environ Sci 7, 81. https://doi.org/10.3389/fenvs.2019.00081.

[55] Khaldoon, S., Lalung, J., Maheer, U., Kamaruddin, M.A., Yhaya, M.F., Alsolami, E. S., et al., 2022. A review on the role of earthworms in plastics degradation: issues and challenges. Polymers 14, 4770. https://doi.org/10.3390/polym14214770.

[56] Fu, Y., Wu, G., Bian, X., Zeng, J., Weng, Y., 2020. Biodegradation behavior of Poly (Butylene Adipate-Co-Terephthalate) (PBAT), Poly(Lactic Acid) (PLA), and their blend in freshwater with sediment. Molecules 25, 3946. https://doi.org/10.3390/molecules25173946.

[57] Yang, Y., Li, Z., Yan, C., Chadwick, D., Jones, D.L., Liu, E., et al., 2022. Kinetics of microplastic generation from different types of mulch films in agricultural soil. Sci Total Environ 814, 152572. https://doi.org/10.1016/j.scitotenv.2021.152572.

[58] Shah, A.A., Hasan, F., Hameed, A., Ahmed, S., 2008. Biological degradation of plastics: a comprehensive review. Biotechnol Adv 26, 246–265.

[59] Blesa Marco, Z.E., Sáez, J.A., Andreu-Rodríguez, F.J., Penalver, R., Rodríguez, M., Eissenberger, K., et al., 2024. Effect of abiotic treatments on agricultural plastic waste: efficiency of the degradation processes. Polymers 16, 359. https://doi.org/10.3390/polym16030359.

[60] Brown, R.W., Chadwick, D.R., Zang, H., Graf, M., Liu, X., Wang, K., et al., 2023. Bioplastic (PHBV) addition to soil alters microbial community structure and negatively affects plant-microbial metabolic functioning in maize. J Hazard Mater 441, 129959. https://doi.org/10.1016/j.jhazmat.2022.129959.

[61] Graf, M., Greenfield, L.M., Reay, M., Bargiela, R., Williams, G.B., Onyije, C., et al., 2022. Increasing concentration of pure Micro- and Macro-LDPE and PP plastic

negatively affect crop biomass, nutrient cycling, and microbial biomass. J Hazard Mater 458, 131932. https://doi.org/10.1016/j.jhazmat.2023.131932.

[62] Dong, D., Guo, Z., Wu, F., Yang, X., Li, J., 2024. Plastic residues alter soil microbial community compositions and metabolite profiles under realistic conditions. Sci Total Environ 906, 167352. https://doi.org/10.1016/j.scitotenv.2023.167352.

[63] Graf, M., Greenfield, L.M., Reay, M.K., Bargiela, R., Golyshin, P.N., Evershed, R.P., et al., 2024. Field-based assessment of the effect of conventional and biodegradable plastic mulch film on nitrogen partitioning, soil microbial diversity, and maize biomass. Appl Soil Ecol 202, 105595. https://doi.org/10.1016/j.apsoil.2024.105595.

[64] Greenfield, L.M., Graf, M., Rengaraj, S., Bargiela, R., Williams, G., Golyshin, P.N., et al., 2022. Field response of N2O emissions, microbial communities, soil biochemical processes and winter barley growth to the addition of conventional and biodegradable microplastics. Agric Ecosyst Environ 336, 108023. https://doi.org/10.1016/j.agee.2022.108023.

[65] Brown, R.W., Chadwick, D.R., Thornton, H., Marshall, M.R., Bei, S., Distaso, M.A., et al., 2022. Field application of pure polyethylene microplastic has no significant short-term effect on soil biological quality and function. Soil Biol Biochem 165, 108496. https://doi.org/10.1016/j.soilbio.2021.108496.

[66] Fernandes, M., Salvador, A.F., Vicente, A.A., 2024. Biodegradation of PHB/PBAT films and isolation of novel PBAT biodegraders from soil microbiomes. Chemosphere 362, 142696. https://doi.org/10.1016/j.chemosphere.2024.142696.

[67] Maestri, C., Plancher, L., Duthoit, A., Hébert, R.L., Di Martino, P., 2023. Fungal biodegradation of polyurethanes. J Fungi 9, 760.

[68] Gong, Z., Jin, L., Yu, X., Wang, B., Hu, S., Ruan, H., et al., 2023. Biodegradation of low density polyethylene by the fungus cladosporium sp. Recovered from a landfill site. J Fungi 9, 605. https://doi.org/10.3390/jof9060605.

[69] Wróbel, M., Szymańska, S., Kowalkowski, T., Hrynkiewicz, K., 2023. Selection of microorganisms capable of polyethylene (PE) and polypropylene (PP) degradation. Microbiol Res 267, 127251. https://doi.org/10.1016/j.micres.2022.127251.

[70] Meng, F., Yang, X., Riksen, M., Geissen, V., 2022. Effect of different polymers of microplastics on soil organic carbon and nitrogen – a mesocosm experiment. Environ Res 204, 111938. https://doi.org/10.1016/j.envres.2021.111938.

[71] Meng, K., Teng, Y., Ren, W., Wang, B., Geissen, V., 2023. Degradation of commercial biodegradable plastics and temporal dynamics of associated bacterial communities in soils: a microcosm study. Sci Total Environ 865, 161207. https://doi.org/10.1016/j.scitotenv.2022.161207.