Degradation of Biodegradable Mulch-Derived Microplastics and Their Effects on Bacterial Communities and Radish Growth in Three Vegetable-Cultivated Purple Soils

Abstract

Biodegradable mulch films (BDMs) are considered a promising solution for mitigating plastic residue pollution in agroecosystems. However, the degradation behavior and ecological impacts of their residues on soil–plant systems remain unclear. Here, a pot experiment was conducted using an acidic purple soil (AS), a neutral purple soil (NS), and a calcareous purple soil (CS) to investigate the degradation of 1% (w/w) microplastics derived from polyethylene mulch film (PE-MPs) and polybutylene adipate terephthalate/polylactic acid (PBAT/PLA) mulch film (Bio-MPs), as well as their effects on soil properties, bacterial communities, and radish growth. PE-MPs degraded slightly, while the degradation of Bio-MPs followed the order of NS > CS > AS. PE-MPs and Bio-MPs enhanced the nitrification and radish growth in AS but had no significant effects on soil properties and radish growth in CS. Bio-MPs notably increased the relative abundance of PBAT/PLA degradation-related bacteria, such as Ramlibacter, Bradyrhizobium, and Microbacterium, across the three soils. In NS, Bio-MPs raised soil pH and enriched nitrogen-fixing and denitrifying bacteria, leading to a decrease in NO₃⁻-N content and radish biomass. Overall, the effects of Bio-MPs on soil–plant systems varied with soil properties, which are closely related to their degradation rates. These findings highlight the need to assess the ecological risks of BDM residues before their large-scale use in agriculture.

1. Introduction

The widespread adoption of mulch films in agriculture is attributed to their advantages in controlling soil temperature and retaining moisture, thereby enhancing crop yield and quality [1]. However, concerns are increasing about the impact of plastic residues on the soil ecological environment, especially regarding microplastic (MP) contamination [2]. MPs, defined as plastic fragments with a diameter less than 5 mm [3], exhibit compositions and properties distinct from those of soil particles. MP accumulation can alter the properties of soils such as bulk density, aggregate water stability, pH, and electrical conductivity [4,5], disrupting microbial community structures, enzyme activities, and the biogeochemical cycles of elements such as carbon(C) and nitrogen (N), thus undermining soil health and function [6]. To mitigate concerns associated with the residues from the conventional polyethylene (PE) mulch films, biodegradable mulch films (BDMs) were designed and used worldwide [7]. However, due to their incomplete but easier degradation, BDMs may release more fragments and/or MPs than conventional mulch films do under the actual environmental conditions within a short time [8]. Furthermore, in the conventional agricultural practice, the PE mulch films are retrieved from the soil surface, whereas BDMs are commonly plowed into the soil after crop harvest. Consequently, a higher level of biodegradable MPs (Bio-MPs) may accumulate in soils where the BDMs are repeatedly utilized [9]. With the growing adoption of BDMs, there are increasing concerns regarding the possible environmental hazards posed by the remnants of Bio-MPs in agricultural soils [10].

Previous studies have demonstrated that the effects of Bio-MPs on soil–plant systems are highly variable. Some studies suggested that Bio-MPs exert minimal effects on soil properties and plant growth [11,12], while others highlighted their negative impacts, such as disrupted nutrient availability, altered microbial community composition and functionality [13], and inhibited plant growth [14]. These divergent findings underscore the complexity and uncertainty of the effects of Bio-MPs on soil–plant systems. Except for the influences of MP properties and rates, soil properties were key factors affecting the degradation of biodegradable polyesters, likely playing a major role in these effects. For example, metagenomic analysis showed that the microbial community in lou soil was more responsive to polybutylene adipate terephthalate (PBAT) mulch films than that in fluvo-aquic soil, black soil, and red soil [15]. Research indicates that Bio-MPs application results in a greater rise in CO₂ emissions in sandier soils than in loam soils [16,17]. Similarly, the bacterial communities in the loam sandy loess soils exhibited greater sensitivity to PE-MPs and polylactic acid (PLA) MPs than those present in clay black soils [18]. Moreover, the loam soil type is more water repellent at lower MP concentrations than silty clay–loam–soil [19]. The types of soil are diverse; however, most of the available studies focus on a few soil types with different textures. Furthermore, few studies have compared the differences in plant growth affected by MPs contamination among different soils. It is necessary to explore the degradability of Bio-MPs and their effects on soil properties and plant growth across soils with varying characteristics.

Purple soils, originating from the reddish or purple mudstone or sandstone of the Triassic to Cretaceous system, constitute a major soil type in the warm and humid hilly areas of Southwestern China [20]. Based on their pH values, the purple soils are classified into three types: acidic purple soil (pH < 6.5), neutral purple soil (pH 6.5–7.5), and calcareous purple soil (pH > 7.5) [21]. The extensive use of plastic mulch films may lead to the accumulation of mulch-derived MPs. This issue is particularly relevant in vegetable production systems such as those for radish (Raphanus sativus L.), pepper (Capsicum annuum L.), pakchoi (Brassica rapa subsp. chinensis), and tomato (Solanum lycopersicum L.), which are commonly grown with mulch films in southwestern China. Therefore, the three typical vegetable-cultivated purple soils were employed as tested soils, and the MPs from the PBAT/PLA mulch film, which has been widely used in China, were selected as experimental Bio-MPs. This study aims to investigate the degradation of the Bio-MPs and their effects on soil properties, bacterial community, and plant growth in these nutrient-rich vegetable-cultivated purple soils. Furthermore, the effect of MPs from the PE mulch film (PE-MPs) was also assessed. We hypothesized that (i) the degradation of Bio-MPs varied with soil properties, especially soil pH; and (ii) the responses of microbial community and plant growth to Bio-MPs were related to their degradation in soils. The findings of this study help clarify soil–plant–MPs interactions and provide a theoretical basis for evaluating the ecological risks associated with the use of BDMs in agriculture.

2. Materials and Methods

2.1. Preparation of Soils and MPs

Three purple soils, including an acidic purple soil (AS), a neutral purple soil (NS), and a calcareous purple soil (CS), were collected from the topsoil (0–20 cm) of the vegetable fields in the Bishan and Tongnan Districts of Chongqing, China. According to the World Reference Base for Soil Resources (WRB, 2015), all these soils are classified as Cambisols. Post-elimination of plant debris, stones, and other contaminants, the soil samples were air-dried, then ground and passed through a 5 mm mesh. The properties of the tested purple soils are presented in Table S1.

The Bio-MPs were prepared from a PBAT/PLA mulch film supplied by Xifeng Plastic Co., Ltd. (Baishan, China). The PE-MPs were derived from a PE mulch film purchased from Lansheng Plastic Products Co., Ltd. (Linyi, China). Both mulch films exhibited a thickness of 0.01 mm, with a black color for the PBAT/PLA mulch film and a silver/black color for the PE mulch film. The films were manually cut into pieces smaller than 5 mm in size, washed with deionized water, air-dried, and stored at 4 °C for subsequent experiments. The size distribution of these MPs is presented in Table S2.

2.2. Experimental Design

The experiment was performed in a greenhouse of Southwest University from September to December 2022 under natural light, with temperatures ranging from 18 to 28 °C and a relative humidity of between 50% and 70% during the experimental period. Each soil type had three treatments with three replications: CK (control without MPs addition), PE-MPs, and Bio-MPs. A total of 1000 g of soil samples was amended with either PE-MPs or Bio-MPs at a rate of 1% (w/w) relative to the soils, then thoroughly mixed with basal fertilizers at rates of 0.15 g N·kg⁻¹, 0.10 g P₂O₅·kg⁻¹, and 0.10 g K₂O·kg⁻¹, supplied by the chemical agents of urea (CH₄N₂O), potassium dihydrogen phosphate (KH₂PO₄), and potassium chloride (KCl), respectively. The treated soils were subsequently transferred into ceramic pots (14 cm in diameter and 15 cm in height) and incubated for 60 days. The MPs rate applied was set up according to the guidelines of the European Standard Ecotoxicity Test (EN 17033:2018) [22]. After incubation, 15 seeds of radish, a widely cultivated plant in southwestern China, were sown at a depth of 0.8–1 cm. After 10 days, the seedlings were thinned to 10 uniform plants per pot and allowed to grow for another 30 days. Throughout the experiment, the pots were randomly repositioned every 3 days, and soil moisture was maintained at 60% of the maximum water holding capacity by weighing and irrigating with deionized water.

2.3. Plant Harvest and Determination

During the first 10 days after sowing, the germination rates were recorded daily. Prior to harvest, the plant heights were measured using a ruler. The seedlings were harvested, separated into roots and shoots, washed clean, and dried at 70 °C till constant weight for biomass assessment. The contents of nitrogen (N), phosphorus (P), and potassium (K) in the plant were quantified using the Kjeldahl method, the molybdenum blue colorimetric method, and the flame photometric method, respectively, after digestion with an H₂SO₄–H₂O₂ mixture [23].

2.4. Soil Properties and Bacterial Community Determination

After harvesting, the soil samples were divided into three portions: one portion was preserved at −80 °C for bacterial community analysis; another was kept at 4 °C for ammonium nitrogen (NH₄⁺-N), nitrate nitrogen (NO₃⁻-N) and enzyme activities determination; and the third portion was air-dried for further assessment of other soil properties. Soil pH, soil organic matter (SOM), NH₄⁺-N, NO₃⁻-N, available phosphorus (AP), and available potassium (AK) were analyzed following the procedures described by Lu (2000) [23]. The activities of urease, sucrase, and catalase were determined by employing the colorimetric method of phenol-sodium hypochlorite, 3,5-dinitrosalicylic acid, and the titrimetric method of potassium permanganate, respectively [24].

DNA was extracted from 0.5 g of soil using the E.Z.N.A.^® Soil DNA Kit (Omega Bio-tek, Norcross, GA, USA). The quantity and quality of DNA were assessed by NanoDrop 2000 (Thermo Fisher Scientific, Waltham, MA, USA) and 1% agarose gel electrophoresis. The V3–V4 regions of the bacterial 16S rRNA gene were amplified utilizing the primer pairs 338F (5′-ACTCCTACGGGAGGCAGCA-3′) and 806R (5′-GGACTACHVGGGTWTCTAAT-3′). PCR products were separated by 2% agarose gel and purified using the AxyPrep DNA Gel Extraction Kit (Axygen Biosciences, Union City, CA, USA). Sequencing was performed on the Illumina MiSeq PE300 platform (Majorbio, Shanghai, China). Quality-filtered sequences were processed using QIIME (version 1.9.1) and clustered into OTUs at 97% similarity. Sequencing data were deposited in NCBI under accession number SUB15325676.

2.5. MPs Extraction and Characterization

MPs were extracted from the soils through density separation following a modified protocol of Li et al. (2022) [25]. Specifically, 50 g of air-dried soil was mixed with 150 mL of ZnCl₂ (1.6 g·cm⁻³) in a 250 mL conical flask. The mixture was sonicated for 2 min, stirred thoroughly for 30 min, and then settled for 24 h. The supernatant was collected via vacuum filtration using a 0.45 μm membrane filter. MPs retained on the filter were transferred to a 250 mL beaker containing 50 mL of 30% H₂O₂ solution and heated at 60 °C to digest organic matter. Subsequently, the solution underwent a second vacuum filtration through the 0.45 μm filter membrane, and the MPs on the membrane were collected in a Petri dish (60 mm in diameter). These extraction steps were repeated 3 times to fully extract analogous MPs from soils. The MPs were dried at 25 °C, weighed, and kept at 4 °C in the dark for characterization. All the samples were extracted in triplicate. The extraction method has a recovery rate of 90% for MPs. A control sample without soil was included, and no MPs were detected, indicating minimal risk of external pollution during extraction. Additionally, the pristine Bio-MPs were subjected to the same procedure and showed negligible weight loss (<1%).

According to the EN ISO 472:2013 [26] (Plastics–Vocabulary), “degradation” is defined as an irreversible process leading to a significant change in the structure of a material, typically characterized by a change in properties (e.g., integrity, molecular mass or structure, and mechanical strength) and/or by fragmentation, affected by environmental conditions, proceeding over a period of time and comprising one or more steps. In this study, the degradation of MPs was evaluated through the changes in multiple indicators before and after experiments. Briefly, the degradation rate of MPs was assessed according to their weight loss rate after the experiment. The changes in the surface morphology of the MPs were observed by scanning electron microscopy (SEM, Sigma 500, Zeiss, Oberkochen, Germany) at an acceleration voltage of 3–10 kV after being sprayed with gold. Three individual MPs from each treatment were randomly selected, and the representative SEM images are presented in Figure 1. The infrared absorption spectrum of the MPs was gained by an attenuated total reflection-Fourier transform infrared spectrometer (ATR-FTIR, Spectrum Two, PerkinElmer, Waltham, MA, USA). The infrared spectra were recorded with a scan range of 4000–500 cm⁻¹, for 32 scans, and at a resolution of 4 cm⁻¹. To ensure consistent contact between the MPs and the ATR crystal, uniform pressure was applied using the pressure arm of the ATR accessory. In addition, the carbonyl index, based on the ratio of the carbonyl peak area (1790–1600 cm⁻¹) to the reference peak area (1560–1330 cm⁻¹), was estimated to indicate the degradation degree of MPs [27].

2.6. Statistical Analysis

The data are presented as the mean ± standard deviation of three replicates per treatment. Two-way ANOVA followed by Duncan’s test was used to evaluate the differences in soil properties, enzyme activities, microbial diversity, plant biomass, and nutrient uptake among treatments with SPSS Statistics software (v 26). A one-way ANOVA was applied to evaluate differences in the relative abundance of bacterial communities at the phylum and genus levels among MP types within the same soil type. The alpha diversity indices were calculated using Mothur (version 1.30.2). Principal Coordinate Analysis (PCoA) and correlation heatmap analysis were performed using the R “vegan”, “ape”, and “pheatmap” packages (version 4.0.3), respectively.

3. Results

3.1. MP Degradation in Three Purple Soils

The SEM images showed that the original PE-MPs had an intact and smooth surface (Figure 1a), their ATR-FTIR spectrum had the typical peaks of polyethylene, such as C-H stretching vibrations at 2915 cm⁻¹ and 2848 cm⁻¹, respectively, and the stretching vibration of C=C at 1472 cm⁻¹, as well as the bending vibration of C-H at 719 cm⁻¹ (Figure 1d) [28]. After the experiment, the morphology and the intensity of the above peaks did not change obviously, and the weight loss was less than 1% of the total (Figure 1c), indicating that PE-MPs degraded insignificantly across the three soils.

In the case of Bio-MPs, they had an intact surface with small particles and white striations on them (Figure 1b). After the experiment, their surfaces exhibited uneven protuberances and increased roughness without white striations being observed. Notably, deep cracks and holes appeared on the surface of Bio-MPs from NS. The FTIR spectra (Figure 1e) presented several prominent bands belonging to PBAT, such as those at 2957 cm⁻¹ (-CH₂ stretch), 1710 and 1267 cm⁻¹ (aromatic C=O and C-O absorption, respectively), 1410 cm⁻¹ (benzene ring in the main chain), 1100 cm⁻¹ (C-O-C stretch), and 726 cm⁻¹ (C-H bending vibrations) [7,28,29], as well as a PLA-specific peak at 873 cm⁻¹ (–O–CH–CH₃ bending vibration) [30]. The lower intensity of the PLA-specific peaks than that of the PBAT-related peaks suggested that PBAT has higher amounts than PLA in the Bio-MPs. After the experiments, a decrease in the intensities of those bands was detected, implying the hydrolytic cleavage of ester bonds and the degradation of aromatic structures [7,15]. Furthermore, a new band at 1450 cm⁻¹ (to O-CH₂ vibrational region) was observed for Bio-MPs from the NS, indicating that oxidation reactions occurred during degradation, and a new oxygen-containing functional group formed. The carbonyl index of Bio-MPs changed slightly in AS and CS (p > 0.05), indicating limited oxidative degradation in the two soils. In contrast, a significant increase was observed in NS (p < 0.05), further demonstrating the most pronounced degradation of Bio-MPs in NS.

3.2. Effects of MPs on Soil Properties and Enzyme Activities

The types of soils and MPs, and their interaction, had significant effects on soil pH, NO₃⁻-N, and AP content. In contrast, AK was significantly affected by the types of soils and MPs, while SOM and NH₄⁺-N were significantly influenced by the types of soils (p < 0.01) (

Table 1. The properties of three purple soils amended with PE-MPs and Bio-MPs.

Soils	MPs	pH	SOM (g⋅kg−1)	NH4+-N (mg⋅kg−1)	NO3−-N (mg⋅kg−1)	AP (mg⋅kg−1)	AK (mg⋅kg−1)
AS	CK	5.38 ± 0.05 d	16.25 ± 0.07 c	57.46 ± 4.16 a	5.17 ± 0.66 f	288.47 ± 10.55 b	474.25 ± 5.22 c
	PE-MPs	5.32 ± 0.05 d	16.26 ± 0.05 c	57.01 ± 2.29 a	8.28 ± 1.42 e	296.87 ± 2.42 b	463.88 ± 13.15 c
	Bio-MPs	5.43 ± 0.05 d	16.77 ± 0.14 c	52.40 ± 2.29 a	11.29 ± 0.22 d	331.46 ± 3.96 a	468.24 ± 13.04 c
NS	CK	6.41 ± 0.03 c	20.07 ± 0.17 b	1.00 ± 0.09 b	80.50 ± 3.85 b	289.76 ± 2.38 b	666.60 ± 1.31 a
	PE-MPs	6.50 ± 0.02 c	19.40 ± 0.27 b	1.12 ± 0.15 b	75.25 ± 1.75 b	278.77 ± 5.56 b	643.65 ± 20.65 ab
	Bio-MPs	7.30 ± 0.13 b	20.87 ± 0.40 b	0.94 ± 0.13 b	32.58 ± 4.07 c	322.73 ± 7.90 a	617.67 ± 12.01 b
CS	CK	8.00 ± 0.04 a	27.16 ± 0.05 a	0.32 ± 0.05 c	122.73 ± 12.51 a	105.83 ± 1.83 c	369.37 ± 4.06 d
	PE-MPs	8.00 ± 0.04 a	27.06 ± 0.07 a	0.32 ± 0.02 c	113.41 ± 8.99 a	93.87 ± 9.13 c	353.47 ± 4.87 d
	Bio-MPs	7.96 ± 0.01 a	27.27 ± 0.14 a	0.24 ± 0.02 c	117.93 ± 9.35 a	93.22 ± 6.00 c	349.68 ± 11.92 d
Soils		**	**	**	**	**	**
MPs		**	ns	ns	**	**	**
Soils × MPs		**	ns	ns	**	**	ns

Note: AS, NS, and CS represent acidic, neutral, and calcareous purple soils, respectively. Values represent mean ± standard error (n = 3). Different letters in the same column indicate significant differences among treatments (p < 0.05). ‘Soils’ represents the soil types, ‘MPs’ represents the MP types, and ‘Soils × MPs’ represents the interaction between soil types and MP types. ** means a significance level of p < 0.01; ns indicates no significance.

). Compared with CK, PE-MPs showed negligible effects on pH, SOM, NH₄⁺-N, AP, and AK across the three purple soils, but significantly increased NO₃⁻-N content in the AS (p < 0.05). Bio-MPs markedly enhanced NO₃⁻-N and AP contents (p < 0.05), exhibiting insignificant alterations in the other properties of AS. In the NS, Bio-MPs also did not markedly affect NH₄⁺-N content but significantly elevated soil pH and AP by 13.88% and 11.38%, respectively, as well as notably decreasing NO₃⁻-N and AK contents by 59.53% and 7.34%, respectively. In contrast, the application of Bio-MPs in CS did not result in significant changes in any of the measured parameters.

The activities of the soil enzymes catalase, urease, and sucrase were measured as indicators of microbial functional responses to MP addition. Catalase reflects oxidative stress and aerobic microbial activity [31,32], urease is involved in nitrogen cycling [33], and sucrase contributes to carbon metabolism, with prior studies reporting its sensitivity to MP exposure [31,34]. The types of soils and MPs, and their interaction, also significantly affected catalase activity. Conversely, urease activity was notably affected only by soil type, and sucrase activity exhibited a strong dependence on both soil type and its interaction with MP type (p < 0.01) (Figure 2). Compared with CK, PE-MPs had an insignificant effect on the activities of these enzymes across the soils, but Bio-MPs significantly boosted catalase activity by 52.38%, 48.79%, and 8.35% in the AS, NS, and CS, respectively. Furthermore, Bio-MPs also enhanced the activities of urease and sucrase by 44.44% and 22.58% in NS, respectively, but displayed insignificant influences regarding the activities of the two enzymes in AS and CS.

3.3. Effects of MPs on Soil Bacterial Community

3.3.1. Alpha and Beta Diversity

A total of 539,031, 577,548, and 590,624 sequences were clustered into 2943, 5463, and 7828 operational taxonomic units (OTUs) in AS, NS, and CS, respectively. Based on the OTUs, the alpha diversity was estimated (Figure 3a). The types of soils and MPs, and their interaction, significantly affected the alpha diversity indices except for the Ace index, which was affected only by soil types (p < 0.05). Among the soils, CS exhibited the highest Chao, Ace, and Shannon indices, followed by the NS and AS. Compared with CK, PE-MPs did not significantly alter these indices in any of the three soils, except for a significant increase in the Simpson index in AS. Bio-MPs also had an insignificant impact on these indices in AS and CS, but markedly decreased the Chao, Ace, and Shannon indices, while elevating the Simpson index (p < 0.05) relative to CK in NS.

Beta diversity was assessed using PCoA to compare the bacterial community variations among treatments. The PC1 and PC2 axes explained 63.62% and 19.67% of the total variations, respectively (Figure 3b). The bacterial communities in AS, NS, and CS were distinctly distributed in the fourth, second, and third quadrants, respectively. However, no significant separation was observed between PE-MPs and CK across the three soils. Moreover, the Bio-MPs treatments overlapped with PE-MPs and CK in AS and CS, but they showed a pronounced divergence from PE-MPs and CK in NS, indicating that the bacterial community in NS was more sensitive to Bio-MPs than those in AS and CS.

3.3.2. Soil Bacterial Community Composition

Figure 4a illustrates the relative abundance of the top 10 bacterial phyla under different treatments. The dominant phyla varied depending on soil type. For instance, Firmicutes, Proteobacteria, Actinobacteriota, and Chloroflexi dominated the bacterial community in AS and NS, accounting for 71.71–74.99% and 75.79–83.22% of the total bacterial phyla, respectively. In contrast, Proteobacteria, Actinobacteriota, Acidobacteriota, and Chloroflexi accounted for 72.41–73.91% of the phyla in CS. Compared with CK, PE-MPs significantly increased the abundance of Firmicutes in AS, enhanced the abundance of Firmicutes and Patescibacteria, and markedly decreased Actinobacteriota abundance in NS, while causing no significant changes in the bacterial phylum composition in CS. For Bio-MPs, minimal changes in phylum composition were observed in AS and CS, whereas significant shifts occurred in NS. Specifically, Bio-MPs significantly increased the abundances of Proteobacteria and Actinobacteriota while causing notable decreases in the abundances of Firmicutes, Chloroflexi, Acidobacteriota, Gemmatimonadota, Patescibacteria, and Planctomycetota in NS.

Figure 4b presents the relative abundances of the top 15 bacterial genera under different treatments. In AS, dominant genera, such as Bacillus, norank_f_JG30-KF-AS9, Sphingomonas, and norank_f__LWQ8, accounted for 37.62–42.56% of the total genera. In NS, the predominant genera included Bacillus, Sphingomonas, Arthrobacter, and Gemmatimonas, representing 12.65–34.05% of the total genera. In contrast, CS was characterized by six genera, including Knoellia, Bacillus, norank_f__norank_o__Vicinamibacterales, norank_f_JG30-KF-CM45, Marmoricola, and norank_f__Vicinamibacteraceae, which together comprise approximately 22.22–22.59% of the total genera. One-way ANOVA revealed that a total of 8, 11, and 9 genera with a relative abundance above 0.1% exhibited significant differences between PE-MPs and CK in AS, NS, and CS, respectively (Tables S3–S5). Specifically, compared with CK, PE-MPs significantly increased the relative abundance of Bacillus in AS, enhanced Bacillus, unclassified_f_Symbiobacteraceae, norank_f_norank_o_Chloroplast in NS, and increased norank_f_TRA3-20 in CS. Conversely, PE-MPs markedly decreased the relative abundances of Arthrobacter in NS, as well as Rhodococcus, Sphingomonas, and Marmoricola in CS. Bio-MPs significantly increased or decreased a total of 9, 56, and 15 genera in AS, NS, and CS, respectively. Specifically, the relative abundances of Ramlibacter, unclassified_f_Comamonadaceae, and Bradyrhizobium were notably boosted by 45.0–183.6%, 93.5–160.2%, and 25.5–199.6% under Bio-MPs treatments, respectively, with the increase in the order of NS > CS > AS. Notably, a dramatic increase was observed with Microbacterium (from 0.02% to 14.05%), unclassified_f_ Microbacteriaceae (from 0.27% to 1.05%), and Herpetosiphon (from 0.19% to 1.05%), alongside the unique Rhizobacter (a nitrogen-fixing bacterium), with a relative abundance of 17.05% in Bio-MPs treated NS. Furthermore, Bio-MPs significantly reduced the relative abundances of 4 and 8 bacterial genera in AS and CS, respectively, but a number of genera were significantly decreased by Bio-MPs in NS, including Bacillus, Arthrobacter, Gemmatimonas, norank_f__Roseiflexaceae, norank_f__JG30-KF-CM45, etc. These findings implied that the bacterial community exhibited a more pronounced response to Bio-MPs than to PE-MPs at the genus level. Notably, the responses were most significant in NS, while being least significant in AS.

3.4. Effects of MPs on Radish Seedling Growth and Nutrient Contents

The germination rate was not significantly affected by the type of MP and its interaction with soil type. In contrast, the plant biomass was significantly impacted by the types of soils and MPs and their interaction (p < 0.05) (Figure 5a–c). PE-MPs increased radish biomass in all three purple soils, with a significant increase in shoot biomass in NS and CS (p < 0.05). Conversely, Bio-MPs slightly increased the radish biomass in AS, had no effect in CS, but significantly reduced shoot and root biomass in NS by 20.77% and 11.11%, respectively. The types of soils and MPs significantly affected radish N content, while radish P and K contents were notably affected only by soil type (p < 0.01) (Figure 5d–f). Compared with the CK, PE-MPs did not significantly alter N, P, or K contents in radish seedlings across the three soils. Similarly, Bio-MPs had no significant effect on radish N content in AS and CS or on radish P and K contents in all soils, but notably reduced N content by 32.08% in NS.

4. Discussion

4.1. The Degradation of MPs in Different Purple Soils

The degradation of MPs in soil is largely determined by their polymer composition. In this study, PE-MPs exhibited minimal degradation in all purple soils. This is likely due to the highly stable carbon–carbon backbone of polyethylene, which resists microbial degradation over short periods [18]. In contrast, Bio-MPs contain multiple oxygen-containing functional groups, making them more susceptible to microbial utilization and degradation. As microbial activity is highly sensitive to environmental conditions, the degradation of Bio-MPs varies with soil type. In this study, the most pronounced degradation of Bio-MPs was observed in NS (Figure 1), which is consistent with the previous studies. For example, Zhang et al. (2022) [27] demonstrated that PLA/PBAT film exhibited higher degradation efficiency in soils with a pH of 7.0 compared to those with pH 5.5 and 8.5 after 105 days of soil burial. Han et al. (2021) [15] observed that soils with higher pH values (>7) exhibited greater PBAT hydrolase gene abundance and higher cumulative CO₂-C mineralization compared to soils with lower pH (−4.7). Chen et al. (2025) [35] also reported that Mollisol with a pH of 6.7 exhibited a higher PLA-MPs mineralization rate than Ferralsol and Alfisol, which had pH values of 4.2 and 4.4, respectively. Wu et al. (2023) [36] reported that PBAT-degrading hydrolases exhibit optimal activity at pH 7–8, facilitating effective PBAT degradation. In this study, AS and NS shared similar natural pedogenic conditions and properties, except that AS had a much lower pH. Correspondingly, AS exhibited a significantly lower degradation rate of Bio-MPs than NS, highlighting the crucial role of soil pH in the degradation of Bio-MPs. However, CS with a pH of 8.02 exhibited significantly lower Bio-MPs degradation than NS, which might be attributed to its high contents of SOM and clay. Under high SOM conditions, microorganisms might not need to utilize Bio-MPs [15], while soil clay can protect MPs from microbial degradation by the formation of organo-mineral complexes [37]. Furthermore, Huo et al. (2024) [37] found that Ferralsol with a lower pH value but a higher SOM content exhibited more MPs mineralization than Vertisol (pH 7.8) and Solonetz (pH 5.3) at 20 and 40 °C, but at 30 °C, Vertisol had a higher MPs mineralization rate than the other two soils. Therefore, it can be speculated that the degradation of Bio-MPs is affected by the comprehensive effect of soil properties.

4.2. The Changes in Soil Properties in Different Purple Soils

Soil properties are essential for maintaining soil quality, promoting nutrient cycling, and enhancing crop growth [3]. The minimal changes in the contents of SOM and NH₄⁺-N contents across three soils illustrated that, as substances rich in organic carbon, PE-MPs and Bio-MPs exhibited minimal effects on SOM and the soil ammonification process, which are consistent with those reported by Zhang et al. (2024) [38] and Xiang et al. (2023) [39]. However, the effects of MPs on the other tested properties varied with soils. In AS, which exhibited weak nitrification, both types of MPs significantly increased NO₃⁻-N content. This could be attributed to improved soil aeration following MP addition, which likely promoted soil nitrification. In NS, PE-MPs had no significant effect on NO₃⁻-N content, whereas Bio-MPs significantly decreased it. This reduction may result from nitrogen immobilization by microorganisms, triggered by the release of labile carbon during the degradation of Bio-MPs [10,40]. Furthermore, the lactic acid oligomers released from PBAT/PLA during degradation can also serve as an electron donor in the denitrification process [41], thereby potentially promoting denitrification and reducing NO₃⁻-N content in NS. Neither PE-MPs nor Bio-MPs significantly affected the pH of AS and CS, but Bio-MPs significantly increased the pH of NS (Table 1). This increase may result from the nitrification process (generating H⁺) and the enhancement of the nitrate-reduction process (consuming H⁺) [10], as supported by strong positive correlations between pH and the relative abundances of Rhizobacter (a nitrogen-fixing genus) and Bradyrhizobium (a denitrifier genus) (Figure S1) in the Bio-MPs-treated NS. Furthermore, the release of additives such as CaCO₃ may also cause an increase in pH of NS [42] due to its high Bio-MPs degradation rate in NS.

Owing to their rapid and sensitive response to environmental disturbances, soil enzymes have been widely employed as indicators for evaluating the ecological effects of MPs. In this study, PE-MPs did not notably affect the activities of the tested enzymes across the three soils (Figure 2). In contrast, Bio-MPs markedly increased catalase activity in all the soils, had no significant effect on urease and sucrase activities in the AS and CS, while markedly increasing them in NS. These findings can probably be explained by the more pronounced alteration in soil properties and bacterial community compositions observed in the Bio-MPs-treated NS than those in AS and CS. Catalase serves as an indicator of soil aeration; thus, the change in its activity can be explained by the increased porosity resulting from MP addition [32]. Elevated sucrase and urease activities suggest a stimulation of microbial carbon and nitrogen metabolism in NS.

4.3. The Responses of the Bacterial Community in Different Purple Soils

Soil microorganisms play a vital role in regulating key biological processes within ecosystems, such as energy flows and nutrient cycling within soils. Previous studies have reported conflicting results regarding the impact of PE-MPs on soil bacterial communities, varying from significant changes [43] to minor effects [18]. Our study revealed that PE-MPs exerted no notable influence on bacterial Alpha and Beta diversity (Figure 3), but they altered the bacterial community compositions across the three soils differently (Figure 4). Like PE-MPs, Bio-MPs had negligible effects on bacterial diversity in the AS and CS, yet significantly reduced bacterial diversity in NS (Figure 3). In general, soils with a higher abundance of degradative microorganisms tend to respond more rapidly to Bio-MPs by utilizing them as carbon or energy sources, which promotes the enrichment of specific microbial groups and alters the overall microbial community structure [15]. The more extensive alterations of microbial compositions in NS suggested that this soil possessed a greater abundance of microorganisms with high polyester degradation activity than AS and CS. Proteobacteria and Actinobacteriota have been reported as the key bacterial phyla breaking down MPs [44]. Some genera from these phyla, such as Ramlibacter, Bradyrhizobium, and Microbacterium, have demonstrated an enhanced response to Bio-MPs addition and were recognized as the contributors responsible for degrading PBAT/PLA plastics in soils [15,45,46]. Comamonadaceae is also known as a hydrocarbon decomposer [47]. The significant and positive correlation between the degradation rates and the enhancement of these genera supports their potential involvement in the degradation of Bio-MPs across the three purple soils (Figure S2). The more dramatic enhancement of the four genera may contribute to the highest Bio-MPs degradation rate in NS. Additionally, Rhizobacter and Bradyrhizobium are involved in nitrogen-fixing bacteria [48,49], while Comamonadaceae function as denitrifying bacteria [50]. However, Ramlibacter is crucial in soil N cycling [51]. The dramatic enrichments of these bacteria in Bio-MPs-treated NS suggested that the regulation of bacteria involved in the nitrification and nitrate-reduction process by Bio-MPs affects N cycles, thereby decreasing their availability in soil.

4.4. The Responses of Radish Growth in Different Purple Soils

Our observations indicated that PE-MPs and Bio-MPs had negligible influence on radish germination across the three purple soils (Figure 5a), aligning with the discoveries of a previous study [52]. This occurrence can be explained by the predominant influence of nutrients within the seeds on germination [53]. However, PE-MPs promoted radish biomass across the soils, possibly due to their lower density relative to the soil, which enhances aeration and reduces mechanical resistance to root penetration [18]. In contrast, a significant negative correlation between radish relative biomass and Bio-MPs weight loss (Figure S3) suggests that differences in Bio-MPs degradation may underlie the varying plant responses among soils. Previous studies have indicated that the degradation products (e.g., adipic acid, terephthalic acid, and butanediol) from PBAT MPs (2% w/w) may severely inhibit Arabidopsis growth [45]. Additionally, high levels of volatiles (e.g., dodecanal) from biodegradable plastic residues suppressed wheat (Triticum aestivum) growth [54]. Although the specific degradation products were not analyzed in this study, it is speculated that the release of additives, intermediates, and degradation byproducts from Bio-MPs may directly inhibit radish growth in NS, where the Bio-MPs degradation rate was highest. The slight increase in radish biomass observed in AS might be caused by the indirect effects of physical property improvements of soils [55].

MPs can influence plant growth by disrupting their nutrient uptake. For example, the addition of PBAT MPs significantly reduced N, P, and K contents in lettuce (Lactuca sativa L.) leaves [56]. Liang et al. (2024) [57] observed that Bio-MPs (PBAT/PLA) significantly decreased N and P contents in the root and shoot of cucumbers (Cucumis sativus L.) and adzuki beans (Vigna angularis), while increasing N content in strawberry (Fragaria × ananassa Duch.) shoots and N and K contents in the cabbage (Brassica rapa var. glabra Regel). This result indicates that the impact of Bio-MPs on plant nutrient uptake varies among plant species. In our study, Bio-MPs significantly reduced radish N content in NS but had no significant effect in AS or CS, suggesting that the influence of Bio-MPs on plant N uptake is soil-dependent. This reduction in N content likely contributed to the decreased biomass observed in NS. The insignificant changes in the radish P and K contents across the soils (Table 1) suggested that in soils with rich P and K nutrients, the impacts of MPs on P and K uptake are small.

5. Conclusions

The study indicates that PE-MPs exhibited negligible degradation, whereas Bio-MPs degraded obviously, and their degradation was affected by the combined influences of soil properties. The effects of MPs on soil properties, plant growth, and underlying mechanisms varied with soil type and MP characteristics, closely linked to their degradation behavior. Both PE-MPs and Bio-MPs enhanced nitrification and promoted radish growth in AS but had no significant effects on radish growth in CS. Conversely, in NS with the highest Bio-MPs degradation, Bio-MPs remarkably increased the pH and decreased the contents of NO₃⁻-N and plant N content. The negative effects of Bio-MPs on plant growth in NS may stem from the direct toxicity of their degradation products or indirect effects, such as reduced N availability and plant N uptake, potentially driven by the enrichment of nitrogen-fixing or denitrifying bacteria. These findings indicate that in the environments suitable for biodegradation, BDMs still have adverse effects on the soil ecosystem. Therefore, it is crucial to evaluate the degradation performance and environmental risks of these mulch films before their large-scale application in agriculture. Moreover, as the findings were derived from a short-term greenhouse pot experiment, long-term field studies are needed to comprehensively evaluate their environmental impacts under realistic soil conditions.