Open AccessArticle

Microplastics Can Alter Plant Parameters Without Affecting the Soil Enzymatic Activity in White Lupine

Carla Sobarzo-Palma

^1,2

María Dolores López-Belchí

Felipe Andrés Noriega

Raúl Zornoza

⁴

Gonzalo Tortella

^5,*

and

Mauricio Schoebitz

^1,2,*

Department of Soil and Natural Resources, Faculty of Agronomy, Universidad de Concepción, Concepción 4030000, Chile

Biotechnology Center, Renewable Resources Laboratory, Universidad de Concepción, Concepción 4030000, Chile

Department of Vegetal Production Faculty of Agronomy, Universidad de Concepción, Concepción 4070409, Chile

⁴

Sustainable Use, Management and Reclamation of Soil and Water Research Group, Escuela Técnica Superior de Ingenieros Agrónomos (ETSIA), Universidad Politécnica de Cartagena, Paseo Alfonso XIII, 30202 Cartagena, Spain

⁵

Centro de Excelencia en Investigación Biotecnológica Aplicada al Medio Ambiente (CIBAMA), Facultad de Ing. y Ciencias, Universidad de La Frontera, Temuco 4811230, Chile

Authors to whom correspondence should be addressed.

Sustainability 2025, 17(1), 149; https://doi.org/10.3390/su17010149 (registering DOI)

Submission received: 20 November 2024 / Revised: 18 December 2024 / Accepted: 24 December 2024 / Published: 28 December 2024

(This article belongs to the Special Issue Emerging Contaminants in the Environment: Measurement, Transport, Impacts, Bioaccumulation, and Remediation)

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Abstract

The widespread presence of microplastics (MPs) in agricultural soils raises concerns regarding their impact on crop health and productivity, particularly in legumes, which are known to have soil-enhancing properties. This study investigated the effects of low-density polyethylene (LDPE), polypropylene (PP), and polyamide (PA) MPs on white lupine (Lupinus albus L.). Plants were cultivated for 110 days in glass pots containing 700 g of volcanic soil mixed with 2% w/w MPs, with four treatments (control, LDPE, PP, and PA) and five replicates each. The results indicated that PP increased soil ammonium and available nitrogen by 71% and 60%, respectively, compared to the control. LDPE increased root length by 3% and decreased chlorophyll content by 2.7%, whereas PA increased chlorophyll levels by 3.5%. Oxidative stress markers were significantly elevated in the LDPE and PA treatments, with 12% and 5.4% increases, respectively, compared with the control. However, no significant differences were observed in enzyme activity or basal soil respiration. These findings contribute to the understanding of how short-term exposure to MPs affects agricultural soils and emphasize the necessity for long-term studies to elucidate their potential effects.

Keywords:

Andisol; enzyme; roots; agroecosystem; microbial activity; abiotic stress

1. Introduction

The Fabaceae family is the basis for sustainable agriculture because it enhances soil properties and supports N cycling. These benefits are primarily attributed to the symbiotic relationship between legume plants and N₂-fixing bacteria, which convert atmospheric N₂ into bioavailable forms for plants [1]. White lupine (Lupinus albus L.) is widely recognized and cultivated for its dual role in improving soil structure and influencing agroecosystem resilience. As a cover crop or in crop rotations, white lupine enhances soil organic matter, provides carbon inputs, and interrupts pest and disease cycles, making it an invaluable tool for sustainable farming systems [2,3,4]. Microplastics (MPs) have emerged as pervasive contaminants in agricultural soils, introduced mainly through the degradation of farming tools such as nets, irrigation systems, and plastic mulch [5,6]. Low-density polyethylene (LDPE), polypropylene (PP), and polyamide (PA) are among the most commonly identified MPs in soil [7]. These materials are not inert; they interact with the soil−plant system in complex ways [8,9,10]. MPs can alter soil properties by affecting microbial communities, enzymatic activity, and nutrient cycling, often by modifying substrate availability or by directly competing with microorganisms for resources [11,12,13,14,15].

Furthermore, MPs can affect plant physiology by inducing oxidative stress, damaging root structures, and interfering with nutrient uptake [9,10]. Previous studies have reported mixed results regarding the impact of MPs on soil enzymatic activity and microbial dynamics. For instance, LDPE and PP-MPs have been shown to stimulate or inhibit certain enzyme activities depending on the soil type and microbial community composition [9,10,11]. Regarding plant responses, MPs may cause positive and negative effects, such as altering root growth, nodulation, or alteration in root exudates, such as malate, citrate, and succinate, which are critical indicators of plant metabolic activity and responses to environmental stress. These compounds play a key role in nutrient acquisition and signaling, making them relevant for understanding plant-soil interactions under microplastic exposure [16,17,18]. However, the extent to which these changes are driven by MP composition, size, and exposure duration is not fully understood. Moreover, while some studies have focused on individual MP types, comparative analyses across different polymers are limited. In this study, we hypothesized that exposure to LDPE, PP, and particularly PA-MPs, due to their N-containing structure, would significantly influence plant physiology and soil enzymatic activities within the soil-plant system.

We investigated the effects of MPs on plant growth, stress responses, and soil microbial activity. We focused on PA-MPs because of the potential for incorporated N in their structure to interact with and alter plant-soil dynamics. By comparing the effects of different MP types on a model legume crop, we aimed to elucidate how the chemical composition of MPs differentially affects plant health and soil properties. In this study, we hypothesized that LDPE and PP-MPs, as inert polymers, might have minimal or indirect effects on enzymatic activities but could influence plant physiological parameters such as oxidative stress and chlorophyll content through physical interactions or surface adsorption. In contrast, PA-MPs, with their nitrogen-containing structure, were expected to have a more pronounced impact on enzymatic activities due to their potential interactions with soil microbial processes and nutrient cycling, as well as differential effects on plant physiological parameters, including nodulation and root exudate profiles. We studied the effects of MPs on plant growth, stress responses, and soil microbial activity. By comparing the effects of different MP types on a model legume crop, we aimed to elucidate how the chemical composition of MPs differentially affects plant health and soil properties.

2. Materials and Methods

2.1. Sample Site and Experimental Design

Soil samples were collected from an experimental station in Santa Rosa, INIA Experimental Station, located in the Mediterranean region of south-central Chile. (Lat. 36°31′59.4′′ S; Long 71°55′35.7′′ W). A 15 kg quantity of soil from a depth of 30 cm was acquired using a metal shovel. The soil belongs to the Diguillín series, is derived from modern volcanic ash of the Andisol order, and is classified as Typic Haploxerands [1]. Andisol soil was selected for its high organic matter content, water retention capacity, and prevalence in the study region, making it representative of agricultural soils in volcanic areas and ideal for investigating soil-microplastic interactions. The texture is silt loam, and the soil chemical properties were organic matter, 11.3%; pH 5.2, available N, 48.6 mg kg⁻¹; Olsen-P, 7.0 mg kg⁻¹; and available K, 137 mg kg⁻¹. The soil was not sterilized before experimentation to preserve its natural enzymatic activity and microbial community structure, which are essential for accurately assessing the effects of MPs under more realistic soil conditions.

White lupine (var. INIA-Alboroto) seeds were subjected to a preliminary treatment involving immersion in a 5 g L⁻¹ sodium hypochlorite solution under aeration for 2 h, followed by immersion in distilled water under aeration for 24 h. Subsequently, the treated seeds were sown in MP-contaminated soil from the germination phase onward.

After germination, the seedlings were transplanted into glass pots (1 L) with holes placed at the bottom to ensure drainage and aeration capabilities, and two plants were allocated to each pot (20 glass pots in total). Each pot contained 700 g of soil contaminated with 2% (w/w) MPs (14 g; LDPE, PP, or PA). The 2% concentration of MPs was chosen based on previous studies investigating microplastic contamination in agricultural soils. This concentration represents a high but plausible level of microplastic accumulation in soils subjected to prolonged plastic farming practices, ensuring the ecological relevance of the study by simulating a worst-case scenario. A completely randomized design featured four distinct treatment groups: control, LDPE, PP, and PA. Each treatment group included five replicates. The pots were then placed in a controlled environment chamber and watered three times a week to maintain the soil moisture content at 60% of the water-holding capacity. The air temperature was maintained at 20 ± 2 °C, featuring a 16-h light cycle and an 8-h dark cycle. The chamber conditions included a consistent relative humidity of 60% and a photosynthetically active radiation (PAR) level of 350 μmol m² s⁻¹.

2.2. Microplastics Preparation

LDPE films, PA ropes, and PP yarn were purchased from agricultural supply stores (Polimaq Ltd. Concepción, Chile; PA and PP: Sodimac S.A, Concepción, Chile). LDPE and PP were mechanically ground using an Ultra-Turrax^® homogenizer (Ultra-TurraxT25, IKA, Wilmington, NC, USA) to obtain particles within the size range of the MPs. Particules of PA-MPs were obtained by manually cutting the PA ropes using scissors, resulting in 2–5 mm particles. The LDPE and PP particles were sieved through a 2 mm sieve to ensure the desired size range adherence. Before use, each MP type was subjected to UV sterilization (254 nm) for one hour to minimize microbial contamination. LDPE, PP, and PA microplastics were chosen because they are commonly used in agricultural activities, such as plastic mulch, irrigation systems, and nets, making them representative contaminants in soils.

2.3. Determination of Plant Parameters

Plant height was measured from the stem to the top leaf using a millimeter ruler. The total aerial and root biomass (g) was evaluated after drying at 70 °C for 48 h. Root diameter (cm), length (cm), volume (cm³), nodules, and cluster roots were analyzed using the WinRHIZO Reg software (V5.0, Regent Instrument Inc., Quebec, QC, Canada).

The relative chlorophyll content was measured using a SPAD chlorophyll meter (SPAD-502 Plus, Konica Minolta, Tokio, Japan). Specifically, the SPAD values of the plants were determined using five measurements taken from different leaves. The average SPAD values of all leaves were considered as the relative chlorophyll content.

Lipid peroxidation rates were determined by quantifying malondialdehyde (MDA) equivalents. Freshly germinated samples (0.5 g) were homogenized in phosphate-buffered saline (PBS) and centrifuged at 3040× g for 20 min at 4 °C. The collected supernatants were combined, and 0.1 mL of the extract was mixed with 1 mL of a solution called MDA, which consisted of 15% trichloroacetic acid (TCA), 0.25 M hydrochloric acid (HCl), and 0.01% butylated hydroxytoluene (BHT). The mixture was heated at 100 °C for 15 min and then cooled to room temperature on ice. The absorbance of each sample was measured at 532 nm wavelength (BioTek Epoch, Agilent, Santa Clara, CA, USA). All reagents were obtained from Merck (Darmstadt, Germany). Lipid peroxidation rate equivalents were expressed as nm g⁻¹ fresh weight (FW) [2,3].

Carboxylates were obtained from the complete root system, which was carefully collected and thoroughly washed before being incubated in 50 mL of a 0.2 mM CaSO₄ solution (pH 5.5). The roots were agitated in an orbital shaker for 2 h. Following incubation, the solution was filtered using a sterile syringe fitted with a 0.22 μm filter and then frozen at −20 °C. The frozen solution was lyophilized using a freeze-dryer (Model FD8508; Bondiro, Ilshin Lab, Co. Ltd., Yangju, Korea) and resuspended in 200 μL of chromatography-grade water. Quantification was performed using a high-performance liquid chromatograph (Hitachi Primaide) equipped with a UV-VIS detector. Separation was performed on a reversed-phase column (Kromasil C18). The mobile phase was prepared according to [4] Cawthray 2003), consisting of 93% v/v 25 mM KH₂PO₄ (pH 5.5) and 7% v/v methanol at a flow rate of 1 mL min⁻¹. Citrate, malate, oxalate, and succinate were used as standards, and detection was performed at 210 nm. The results were expressed as the rate of carboxylate exudation per gram of fresh weight per hour (μmol g⁻¹ FW h⁻¹) [5,6].

2.4. Soil Properties

To determine acid β-glucosidase activity, p-nitrophenyl-β-D-glucopyranoside (PNG 0.05 M) was used as a substrate. Aliquots of 2 mL MUB buffer (pH 6.5) and substrate (0.5 mL) were added to 0.5 g of soil. The mixture was incubated at 37 °C for 1 h, followed by incubation in an ice bath to stop the reaction. Subsequently, 0.5 mL of 0.5 M CaCl₂ and 2 mL of TRIS buffer (pH 12) were added. The same procedure was followed for the blank samples. The mixtures were then centrifuged at 2196× g for 8 min. Appropriate dilutions were prepared, and absorbance readings were taken at 398 nm using a spectrophotometer (BioTek Epoch, Agilent) [7].

Urease activity was determined by adding 2 mL of 0.1 M phosphate buffer (pH 7) and 0.5 mL of 6.4% urea solution as a substrate to the sample (0.5 g) and incubating at 30 °C for 90 min. Distilled water, (7.5 mL) was then added. The samples were centrifuged at 2196× g for 8 min. One milliliter of supernatant was diluted with 5.8 mL of distilled water. Then, 0.8 mL sodium citrate, 1.6 mL reagent A (containing sodium salicylate and sodium nitroprusside), and 0.8 mL of dichloroisocyanurate with NaOH were added. The mixture was incubated in the dark for 45 min. Urease activity was determined using a standard ammonium chloride curve to determine the amount of NH₄⁺ released during hydrolysis. The absorbance was measured at 660 nm using a spectrophotometer (BioTek Epoch, Agilent) [8].

P-nitrophenyl phosphate disodium (PNPP, 0.115 M) was used as the substrate to determine acid phosphatase activity. Aliquots of 2 mL MUB buffer (pH 6.5) and substrate (0.5 mL) were added to 0.5 g of soil. The mixture was incubated at 37 °C for 1 h, followed by incubation in an ice bath to stop the reaction. Subsequently, 0.5 mL of 0.5 M CaCl₂ and 2 mL of 0.5 M NaOH were added. The same procedure was followed for the blank samples. The mixtures were then centrifuged at 2196× g for 8 min. Appropriate dilutions were prepared, and absorbance readings were recorded at 398 nm using a spectrophotometer (BioTek Epoch, Agilent) [9,10].

Fluorescein diacetate hydrolysis (FDA) was measured using sodium phosphate buffer (99 mL, 60 mM, pH 7.8), and 0.1 mL of the FDA solution (2 mg/mL in acetone) was added to 1 g of soil sample. The blank contained only 10 mL of buffer. The tubes were vortexed and incubated for 1 h at 20 °C in a thermostatic bath. After incubation, the samples were cooled in an ice-water bath, and 10 mL of acetone was added to the samples and blanks. The tubes were shaken, and the contents were filtered through Whatman No. 40 filter paper. The absorbance of the resulting solutions (samples and blanks) was measured at 490 nm using a spectrophotometer (BioTek Epoch, Agilent). The results are expressed as μg fluorescein g⁻¹ dry soil [11].

Soil respiration was determined by measuring the amount of carbon dioxide (CO₂) released. Duplicate samples weighing 25 g of soil were placed in incubation bottles for each treatment. A centrifuge tube containing 7.5 mL of 0.5 M NaOH was placed in each bottle. After incubation, 1 mL of 0.5 M NaOH from the centrifuge tube was mixed with 2 mL of 1 M BaCl₂ solution. Phenolphthalein indicator (3 drops) was added to the solution beforehand. The resulting solution was then titrated with 0.1 M HCl. The obtained data were expressed as μg CO₂ g⁻¹ soil [11].

Microbial biomass carbon was estimated based on the methodology described by Vance et al. (1987). Soil samples (10 g) conditioned to 60% WPFS capacity were collected from each treatment. Half of the samples were non-fumigated, while the remainder were fumigated with 0.5 mL CHCl₃ and incubated for 30 min at room temperature. A vacuum pump was used for two minutes at a time to remove chloroform, and this process was repeated until complete removal. To the fumigated and non-fumigated samples, 40 mL of K₂SO₄ 0.5 M was added, and the samples were shaken on a rotary shaker at 220 rpm for 1 h. After shaking, the samples were centrifuged at 10,000 rpm for 10 min, filtered through a 0.45 µm syringe filter, and then diluted 1:10 (2 mL in 8 mL of HPLC-type water). The total organic carbon (TOC) (mg L⁻¹) was estimated using a total organic analyzer (TOC-Shimadzu^®-H544160, Shimadzu, Tokyo, Japan). The final microbial mass was calculated using the TOC results and a constant K_EC of 0.20 [12,13].

2.5. Microplastics Analysis

2.5.1. Observation of MPs by Scanning Electron Microscopy (SEM)

Pristine MPs from each treatment type (LDPE, PA, and PP) were stored at the beginning of the experiment. After the exposure period, MP samples were carefully collected from the soil of each treatment using sterile tweezers. Each sample was prefixed with 2.5% glutaraldehyde and stored in an Eppendorf tube at 4 °C without further treatment to preserve any existing bacterial biofilm and prevent its removal during preparation. For SEM analysis, the samples were mounted on aluminum stubs using double-sided carbon tape, sputter-coated with gold to enhance conductivity, and examined using a scanning electron microscope (SEM) (JSM-6380LV, Jeol, Tokyo, Japan). Multiple areas of each sample were imaged to ensure representative observations of surface morphological changes and potential biofilm formation.

2.5.2. MPs Characterization by Fourier Transform Infrared Spectrometer (FTIR)

FITR spectroscopy was employed to confirm the chemical identity of the MPs used in this study and to investigate potential changes in their surface chemistry after soil exposure. Both pristine and soil-exposed MPs (LDPE, PA, and PP) were analyzed using a spectrometer (FTIR, IRTracer-100, Shimadzu, Tokyo, Japan) equipped with an attenuated total reflectance (ATR) accessory.

Before analysis, soil-exposed MPs were rinsed with distilled water to remove attached soil particles and air-dried at room temperature. Spectra were collected in the mid-infrared region (4000–400 cm⁻¹) at a resolution of 4 cm⁻¹, with an average of 32 scans per spectrum. Prior to each measurement, the background spectrum was acquired. After acquisition, all spectra underwent data processing to improve quality and allow for an accurate comparison. These steps were performed using LabSolutions software ver. 1.108 and included atmospheric correction to remove spectral interference from atmospheric water vapor and CO₂. Smoothing to reduce noise while maintaining spectral features, baseline correction to eliminate baseline drift, and ensuring that all spectra had a common baseline and normalization, in which spectra were vector-normalized to account for differences in sample thickness or contact area with the ATR crystal. Once processed, the spectra were used for polymer identification and comparative analysis of the pristine and soil-exposed MPs. The absorption bands observed in the spectra were compared with the reference spectra from the literature to identify the key peaks of the characteristic functional groups of LDPE, PA, and PP. A visual comparison of the spectra of pristine and soil-exposed MPs was conducted to assess potential changes that might reflect alterations in chemistry.

2.6. Statistical Analysis

Data were collected and analyzed for each treatment group. All variables were tested for normality and homogeneity of variance. Significant differences among treatments were assessed using one-way Analysis of Variance (ANOVA) and Fisher’s Least Significant Difference (LDS) post hoc test for pairwise comparisons. The non-parametric Kruskal−Wallis test was employed for data that did not meet the assumptions, followed by Dunn’s test for post hoc pairwise comparisons.

Principal component analysis (PCA) was performed to explore the clustering patterns among the treatments. In addition, a Pearson correlation test (r) was performed to identify significant relationships between the chemical and microbiological parameters of the soil in each treatment.

All statistical analyses were performed using R software (version 4.2.2, R, 2024). The results were considered statistically significant at p < 0.05. Data visualization and graphs were created using GraphPad Prism (version 8) and R software.

3. Results

3.1. Effects of LDPE, PP, and PA on Soil Parameters

After the exposure period, microbial biomass carbon was reduced in the three MP-exposed treatments compared to that in the control. The addition of LDPE-MPs reduced microbial biomass carbon by 50.5%, whereas PA and PP-MPS reduced it by 41.6% and 62.4%, respectively (p < 0.05) (Figure 1d). In contrast, FDA hydrolytic activity, which reflects total microbial metabolic activity, demonstrated a significant difference upon exposure to PA-MPs, exhibiting a 47.7% increase in activity compared with the control (Figure 1c). Soil basal respiration and total organic carbon showed no significant variation among the groups (Figure 1a,b).

Soil analysis indicated no significant differences in nitrate or Olsen P levels between the control and MP-exposed groups. However, exposure to PP-MPs resulted in an increase in ammonium (70.97%) and available nitrogen (59.87%) concentrations compared with the control (Table 1).

Statistical analysis showed no significant changes in β-glucosidase, acid phosphatase, and urease activity after 110 days of MP exposure (Figure 2a–c).

3.2. Microplastics

The SEM images presented in Figure 3 show no microbial biofilm or surface degradation resulting from bacterial activity. In the (A) and (C) pristine images, minor abrasions are visible, likely caused by mechanical abrasion during the formation of MPs. In images (B), (D), and (F), organic matter (soil) is attached to the surface of the MPs.

FTIR spectroscopy confirmed the chemical composition of the MPs used in this study. Pristine LDPE displayed characteristic peaks at 721, 1375, 1468, 2849, and 2913 cm⁻¹, corresponding to CH₃ and CH₂ [14]. PP exhibited absorption bands at 1458, 2833, and 2954 cm⁻¹ [15], whereas PA exhibited peaks at 688, 949, 1195, 1260, 1418, and 1476 cm⁻¹, which are amide bands [16] (Figure 4).

A comparison between the pristine and soil-exposed spectra showed substantial similarity, with no significant shifts in the peak positions. This suggests that the chemical structure of MPs remained largely unaltered throughout the experimental period.

3.3. Effects of LDPE, PP, and PA on Plant Parameters

Scanner analysis showed no significant differences between the control and MPs treatment groups in root dry biomass, volume, number of cluster roots, plant height, or leaf dry biomass (Figure 5b–g). However, root length was 3% higher in the LDPE group and 1.6% lower in the PP group than in the control group (Figure 5a).

The number of root nodules was 34% lower in the LDPE group than in the control group (Figure 5d). The addition of MPs to the soil influenced the relative chlorophyll content of lupines. Specifically, the chlorophyll content significantly decreased by 2.7% in the LDPE group compared to that in the control group. In contrast, in the PA group, it increased by 3.5% relative to that in the control group (Figure 5h). The change in the chlorophyll content between the PA and LDPE groups was 3.85 SPAD.

Oxidative stress showed significant differences (p < 0.05) between LDPE (12% higher than the control) and PA (5.4% higher than the control) compared with PP (24.6% lower than the control) (Figure 5i). Exudate analysis showed that there was no difference in the root exudates among the treatments (Figure 6).

3.4. Relationship Between Microbiological and Chemical Soil Properties

Two-dimensional principal component analysis (PCA) was performed, integrating all variables evaluated in the microcosm experiments. PCA explained 39.77% of the total variability, with PC1 explaining 23.1% and PC2 explaining 16.67% of the variance (Figure 7). Furthermore, PCA revealed a change in soil properties owing to MP exposure. The LDPE and PA treatments clustered similarly, suggesting common effects on soil properties, while the PP treatment separated, indicating less drastic changes compared to LDPE and PA. Samples treated with LDPE-MPs had a distinct effect on variables such as chlorophyll, nodules, MBC, and MDA. Similarly, FDA, MDA, MBC, and chlorophyll contents were measured (Figure 7).

Pearson’s correlation analysis (Figure 8) revealed different patterns between the microbiological and chemical properties of the control soil and soils exposed to different MPs. Soils exposed to MPs tended to have stronger positive correlations than the control. For instance, the correlation between ammonium and available nitrogen was higher in soils with MPs than in the control (r = 1 in LDPE- and PA-MPs exposed soil, r = 0.96 in PP-MPs exposed soil, and r = 0.92 in control soil). The enzyme activities showed varied correlations: AP was positively correlated with MBC (r = 0.88 in the control and 0.95 in PP-MPs), while urease activity showed a strong negative correlation with P. Olsen in PA-MPs exposed soil (r = −0.92). Olsen P also showed a strong positive correlation with FDA in LDPE-MPs exposure (r = 0.91) but a strong negative correlation with MBC in PA-MPs exposed soil (r = −0.90).

The soil exposed to LDPE showed significant positive correlations between nitrate and AN (r = 0.96) and ammonium (r = 0.94). In contrast, soil exposed to PA-MPs showed significant negative correlations between SBR and AN (r = −0.91) and ammonium (r = −0.88).

4. Discussion

4.1. Effects of MPs on Plant Physiology and Growth

Compared with the control group, exposure to LDPE and PP increased and reduced root length, respectively, which is consistent with previous results [17]. The increased root length under LDPE contamination might be attributed to MPs reducing the soil bulk density and facilitating root biomass production [18]. Conversely, the reduction observed with PP could be due to particle adhesion to the root surface, which creates a physical barrier that affects nutrient and chemical absorption [19]. Previous studies have shown that MPs, as abiotic stressors, can cause plant oxidative damage and inhibit root growth [20]. Although not statistically significant, a reduction in the number of cluster roots was observed in the PP treatment. This could be attributed to MPs affecting root function or nutrient uptake, given that cluster roots are specialized structures that enable plants to maximize nutrient absorption from the soil. Additionally, LDPE plant exposure significantly reduced the number of nodules, which could be attributed to potential root damage, as documented in previous studies, where increased reactive oxygen species (ROS) resulting from cell damage disrupted root cell membrane stability, thereby impairing nodulation [21,22]. Moreover, LDPE can perturb the rhizosphere microbial community, indirectly influencing nodulation owing to its association with specific nitrifying bacteria [23].

The lack of statistically significant changes in plant height could be attributed to the tissue-dependent effects of MPs. Although certain MP types may promote root elongation, as observed in this study, their impact on plant height may depend on the specific tissues affected. A previous study demonstrated that PE exposure reduced wheat shoot height at high concentrations but stimulated root elongation, suggesting tissue-specific responses to MP stress [24,25]

A significant difference (p < 0.05) in chlorophyll content was observed between the LDPE and PA exposure treatments. Previous research has indicated that polyethylene (PE) MPs may decrease chlorophyll content and rubisco activity in lettuce leaves [26]. Although the changes in chlorophyll content are statistically significant, their biological relevance to plant health remains uncertain. These variations may reflect initial physiological adjustments to microplastic-induced stress, such as altered nutrient dynamics or oxidative stress. However, the magnitude of these changes suggests that they may have a limited immediate impact on photosynthetic efficiency under short-term exposure. Even minor decreases in chlorophyll content over prolonged periods could potentially influence plant growth and productivity, warranting further investigation in long-term studies.

Although no significant differences (p < 0.05) were observed compared with the control, elevated levels of oxidative stress were found in plants exposed to PA and LDPE. It is hypothesized that the significantly lower level of oxidative stress in the PP exposure group is due to greater soil compaction, potentially decreasing PP’s capacity to be absorbed and mobilized internally in the plant, thus not activating the oxidative stress response.

4.2. Effects of MPs on Soil Properties

In the soil analysis, the FDAse activity was significantly higher after PA exposure. Previous studies have demonstrated a positive correlation between soil microbial activity and porosity, specific surface area, and aggregate structure. Therefore, it has been hypothesized that exposure to PA-MPs may lead to an increase in porosity, enhanced soil aeration, and enriched aerobic microorganisms [27,28,29,30,31]. Additionally, Pearson’s correlation analysis suggested that exposure to PA-MPs might affect nutrient interactions due to physical changes in the soil structure or nutrient retention, which could contribute to the observed FDA activity.

Carbon biomass reflects the total amount of carbon present, and a decrease in biomass could indicate a lower number of microorganisms or a decrease in carbon content per cell [32]. Considering the reduction in microbial biomass carbon content after exposure to MPs, but an increase in FDA activity upon exposure to PA, it is believed that exposure may have favored the growth of specific microbial groups that are more enzymatically active for the hydrolysis of fluorescein; however, overall, there is a lower microbial biomass. Furthermore, PCA results and Pearson’s correlation showed that the relationship between chemical and microbiological properties was weakened in soils with MPs (especially PA and PP) compared to the control. These findings suggest that PA-MPs significantly affected soil interactions by altering the correlation between soil microbiological and chemical properties.

No significant changes were observed in basal soil respiration following MP exposure. The absence of observable effects can be attributed to the temporal nature of the analyses. Although MPs may induce reactions in soil microbial activity, the relatively long exposure period of 110 days compared with the duration of the analysis (3 days) suggests that any effects may have occurred beyond the scope of this study. Similar findings have been reported in the literature, where the effects of MPs on basal soil respiration varied depending on the type of MPs, exposure duration, and soil conditions. For instance, Zhao et al. (2021) observed that MPs can influence soil respiration as a function of their shape, polymer type, and exposure time, with some shapes and polymers increasing respiration under specific conditions [33]. Similarly, Fei et al. (2020) found that MPs can affect soil enzyme activities and bacterial communities, which may indirectly affect soil respiration dynamics [34]. These results highlight the complexity of soil-microplastic interactions and the need for further research to understand their temporal dynamics.

This study revealed that the addition of MPs did not significantly affect urease, acid phosphatase, or β-glucosidase activity. Previous studies have investigated the impact of MPs on soil enzymatic activities and have reported significant effects that vary depending on MPs type, soil type, and properties [30,34,35,36]. For example, Wang et al. (2024) observed that PE MPs inhibited urease and β-glucosidase activity. In contrast, in cadmium-treated soil, both enzymatic activities were higher [37]. Moreover, Liu et al. (2022) reported a slight reduction in urease activity after exposure to phenantherene-PE, whereas PE exposure significantly increased enzymatic activity [38]

Although our study focused on short-term effects (110 days), the results highlight the potential long-term consequences of MPs in Andisol soils, particularly regarding nutrient cycling and microbial interactions. MPs may alter soil properties over long periods through gradual changes in microbial community composition and soil organic matter dynamics. For example, shifts in microbial populations due to selective pressures exerted by MPs could influence enzymatic activities indirectly, such as reductions in β-glucosidase activity, if cellulose-degrading microbes are outcompeted. Additionally, MPs may adsorb or release organic compounds over time, thereby modifying substrate availability and nutrient cycling processes. These interactions could lead to imbalances in soil functionality, potentially affecting plant productivity and soil health in agricultural systems reliant on Andisols. Therefore, long-term studies are essential to assess these cumulative effects and understand how MPs interact with the unique properties of Andisol soils, such as their high organic matter content and variable charge.

Finally, the particle size of the microplastics used in this study may explain the lack of significant differences observed in some soil and plant parameters. Larger particle sizes (2–5 mm) are less likely to interact closely with soil microorganisms and plant roots than smaller microplastics, which have a greater surface area, higher reactivity, and increased mobility in the soil matrix. Numerous studies [17,18,19,20,21,22,23,24,25,26] have demonstrated that smaller microplastics can exert more substantial ecological effects, such as altering microbial activity, enzyme function, and nutrient cycling. However, it is important to note that when plastics first enter the soil, they often form larger fragments and gradually break down into smaller particles over time. This study reinforces the idea that during the early stages of degradation, when plastic sizes range between 2 and 5 mm, their impact on soil properties and microbial communities may be minimal. Future studies should further explore this size-dependent behavior to better understand the risks associated with microplastics at different stages of fragmentation and degradation.

4.3. MPs

SEM and FTIR analyses revealed no substantial changes in the surface morphology or chemical composition of LDPE, PP, and PA-MPs after exposure compared with the pristine samples. SEM showed no biofilm formation or significant surface alterations, whereas the FTIR spectra remained unchanged, indicating no measurable chemical transformation during the 110-day experimental period. Although no measurable changes in the chemical structure of MPs were observed during the 110-day experimental period, longer exposure times could potentially yield different outcomes. Extended durations may allow for cumulative effects such as oxidative degradation, partial depolymerization, or biofilm formation on MP surfaces. These processes could influence soil microbial dynamics and nutrient cycling, further impacting plant-soil interactions. Future studies should explore these aspects to better understand the long-term implications of MP in agricultural soils.

5. Conclusions

This study evaluated the influence of three types of MPs (LDPE, PA, and PP) on plant growth and Andisol’s chemical and microbiological properties. These results suggest that MPs significantly alter the interactions among soil chemical properties, particularly nitrogen dynamics (ammonium and available nitrogen). These changes could indirectly affect specific microbiological properties, such as FDAse activity and microbial biomass carbon (MBC). However, no significant differences were observed in the enzymatic activities of urease, β-glucosidase, or acid phosphatase. Moreover, the effects of MPs on plant parameters were minor, with changes in root length, nodules, chlorophyll content, and malondialdehyde (MDA) levels. Additionally, no physical changes or microbial film formation were observed on the surface of the MPs, suggesting limited direct interaction with the particles.

Understanding how different types and doses of MPs affect nutrient cycling and microbial activity is critical for better management of cultivable soils. Future research should focus on long-term evaluation of the effects across different soil types to provide insight into mitigating the impact of MP pollution in agricultural systems.

Author Contributions

Conceptualization, M.S. and C.S.-P.; methodology, C.S.-P.; software, C.S.-P.; validation, C.S.-P. and F.A.N.; formal analysis, C.S.-P.; investigation, C.S.-P.; resources, M.S., M.D.L.-B. and G.T.; data curation, C.S.-P.; writing—original draft preparation, C.S.-P.; writing—review and editing, C.S.-P., M.S., R.Z. and G.T.; visualization, C.S.-P.; supervision, M.S.; project administration, M.S.; funding acquisition, M.S. and G.T. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by FONDECYT 1220425, 1230529, 1240947, and ANID/ATE220087.

Institutional Review Board Statement

There were no human subjects in this study, and informed consent was not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The data used to support the findings of this study are included in this article.

Conflicts of Interest

The authors declare no conflicts of interest.

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Figure 1. Soil Parameters. (a) Total Organic Carbon shown as mean ± standard error; (b) soil basal respiration shown as mean ± standard error; (c) FDA activity shown as mean ± standard error; (d) microbial biomass carbon shown as mean ± standard error. Different letters indicate significant differences among the groups (one-way ANOVA with Fisher’s test, p < 0.05).

Figure 2. Soil enzyme activity. (a) β-Glucosidase activity is shown as mean ± standard error; (b) acid phosphatase activity is shown as mean ± standard error; and (c) urease activity is shown as mean ± standard error. Different letters indicate significant group differences (Kruskal–Wallis test, p < 0.05).

Figure 3. SEM images of pristine MPs and those extracted from each soil treatment. (A) Pristine Polypropylene (500× magnification), (B) Polypropylene extracted from experiment (1000× magnification), (C) Pristine Low-Density Propylene (500× magnification), (D) Low-Density Propylene extracted from experiment (1000× magnification), (E) Pristine Polyamide (500× magnification), (F) Polyamide extracted from experiment (1000× magnification).

Figure 4. FTIR spectra of MPs samples from each treatment. Characteristic peaks corresponding to the (a) LDPE, (b) PP, and (c) PA functional groups were also observed.

Figure 5. Plant Parameters. (a) Root length shown as mean ± standard error; (b) root dry biomass shown as mean ± standard error; (c) root volume shown as mean ± standard error; (d) number of nodules shown as mean ± standard error; (e) number of cluster roots shown as mean ± standard error; (f) plant height shown as mean ± standard error; (g) leaf dry biomass shown as mean ± standard error; (h) chlorophyll content shown as mean ± standard error; and (i) oxidative stress (MDA content) shown as mean ± standard error. Different letters indicate significant differences among the groups (one-way ANOVA with Fisher’s test, p < 0.05).

Figure 6. Root exudates. (a) Oxalate is shown as mean ± standard error; (b) malate is shown as mean ± standard error; (c) citrate is shown as mean ± standard error; and (d) succinate is shown as mean ± standard error. Different letters indicate significant differences among the groups (one-way ANOVA with Fisher’s test, p < 0.05).

Figure 7. Principal component analysis (PCA) of different soil treatments, their chemical and microbiological properties, and plant parameters.

Figure 8. Significant Pearson correlations were observed between the chemical and microbiological properties. (a) Control soil, (b) soil with LDPE-MPs, (c) soil with PA-MPs, and (d) soil with PP-MPs.

Table 1. Chemical properties of control soil and MPs-exposed soil after 110 days of exposure.

Soil Properties	Control	PP	LDPE	PA
Nitrate (mg kg⁻¹)	1.76 ± 0.6 ^a	2.02 ± 0.6 ^a	2.30 ± 1.8 ^a	1.46 ± 0.3 ^a
Ammonium (mg kg⁻¹)	7.44 ± 1.5 ^b	12.72 ± 1.9 ^a	15.28 ± 9.5 ^ab	10.78 ± 1.7 ^ab
Available N (mg kg⁻¹)	9.22 ± 1.5 ^b	14.74 ± 1.6 ^a	17.58 ± 11.2 ^ab	12.22 ± 2.0 ^ab
P Olsen (mg kg⁻¹)	28.44 ± 0.7 ^a	29.12 ± 1.4 ^a	29.1 ± 1.9 ^a	30.4 ± 1.2 ^a

Data are presented as the mean ± standard error (n = 5). Lowercase letters in the row represent statistically significant differences (p < 0.05) among treatments (p < 0.05), as revealed by one-way ANOVA followed by the LDS Fisher’s test. PP: Polypropylene exposure treatment, LDPE: low-density polypropylene exposure treatment, PA: Polyamide exposure treatment.

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Sobarzo-Palma, C.; López-Belchí, M.D.; Noriega, F.A.; Zornoza, R.; Tortella, G.; Schoebitz, M. Microplastics Can Alter Plant Parameters Without Affecting the Soil Enzymatic Activity in White Lupine. Sustainability 2025, 17, 149. https://doi.org/10.3390/su17010149

AMA Style

Sobarzo-Palma C, López-Belchí MD, Noriega FA, Zornoza R, Tortella G, Schoebitz M. Microplastics Can Alter Plant Parameters Without Affecting the Soil Enzymatic Activity in White Lupine. Sustainability. 2025; 17(1):149. https://doi.org/10.3390/su17010149

Chicago/Turabian Style

Sobarzo-Palma, Carla, María Dolores López-Belchí, Felipe Andrés Noriega, Raúl Zornoza, Gonzalo Tortella, and Mauricio Schoebitz. 2025. "Microplastics Can Alter Plant Parameters Without Affecting the Soil Enzymatic Activity in White Lupine" Sustainability 17, no. 1: 149. https://doi.org/10.3390/su17010149

APA Style

Sobarzo-Palma, C., López-Belchí, M. D., Noriega, F. A., Zornoza, R., Tortella, G., & Schoebitz, M. (2025). Microplastics Can Alter Plant Parameters Without Affecting the Soil Enzymatic Activity in White Lupine. Sustainability, 17(1), 149. https://doi.org/10.3390/su17010149

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