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Article

Extraction and Characterization of Microplastics in Soil: A Case Study from the Hetao Irrigation District

by
Chia Min Ho
1,
Weiying Feng
1,*,
Yuxin Deng
1,
Xiaofeng Li
2 and
Su Kong Ngien
3
1
School of Materials Science and Engineering, Beihang University, Beijing 100191, China
2
China Hebei Construction and Geotechnical Investigation Group Ltd., Shijiazhuang 050227, China
3
Faculty of Civil Engineering Technology, Universiti Malaysia Pahang Al-Sultan Abdullah, Lebuhraya Persiaran Tun Khalil Yaakob, Gambang 26300, Pahang, Malaysia
*
Author to whom correspondence should be addressed.
Water 2025, 17(18), 2700; https://doi.org/10.3390/w17182700
Submission received: 16 August 2025 / Revised: 8 September 2025 / Accepted: 11 September 2025 / Published: 12 September 2025
(This article belongs to the Special Issue Water Environment Pollution and Control, 4th Edition)
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Versions Notes

Abstract

Microplastics (MPs) pollution has become a global environmental issue. Soil, as a key environmental medium, serves as an important sink and carrier of MPs. Accurate and efficient extraction of MPs from soil matrices is essential for understanding their distribution, composition, and environmental behavior. This study presents a refined extraction method that combines two-step density separation with sodium chloride (NaCl, 1.20 g/cm3), hydrogen peroxide (H2O2) digestion for organic matter removal and a Fractionated Filtration Method (FFM) to capture MPs across multiple particle size ranges. Polymer identification and size characterization were performed using the high-throughput Agilent 8700 Laser Direct Infrared (LDIR) imaging system. Method validation demonstrated a recovery rate of 85% based on 100 μm MPs standards spiked into soil and minimal background contamination of 5–8 particles in blank controls, confirming the reliability of the workflow. Applying this method to agricultural soils from the Hetao Irrigation District revealed widespread MP contamination, with concentrations ranging from 5778 to 31,489 particles/kg and an average of 16,461 ± 8097 particles/kg. More than 99% of MPs were smaller than 500 μm, with the 10–30 μm fraction dominating the distribution. Polypropylene (PP), polyamide (PA), and polyethylene (PE) accounted for over 90% of detected MPs. This refined method enables reproducible extraction and accurate characterization of fine MPs in complex soil environments and provides a practical foundation for advancing standardized soil MP monitoring protocols.

1. Introduction

Since 1907, plastic has been widely used due to its durability, lightweight nature, and low cost. In recent years, biodegradable plastics have also been increasingly utilized as alternatives to conventional polymers [1,2]. While plastic has significantly contributed to societal advancement and improved human living standards, the disposal of plastic waste has become a critical global environmental concern. Currently, over 450 million tons of plastics are produced annually, with approximately 12.5 million tons utilized specifically in agricultural practices [3]. Agricultural plastics primarily include mulch films, greenhouse films, and irrigation pipes, which enhance crop yields, conserve water, and protect plants from pests and adverse weather conditions [4]. Once released into the environment, these plastics undergo mechanical abrasion, ultraviolet degradation, and biological breakdown processes, fragmenting into plastic particles smaller than 5 mm, known as microplastics (MPs) [5,6].
MPs have been detected in diverse environments, from the peaks of the Himalayas to the depths of the Mariana Trench, and from coastal regions to remote oceanic waters, spanning the globe from equatorial zones to the polar regions [7,8,9]. Compared to marine environments, soils are increasingly recognized as one of the largest sinks for MPs, yet remain methodologically challenging to study. MPs enter soil through multiple anthropogenic pathways, including the application of sewage sludge and compost, wastewater irrigation, the use of plastic mulch, tire wear, litter deposition, and atmospheric fallout [10]. Study indicates that MP concentrations in soils can exceed those found in marine environments by more than fourfold [11]. The accumulation of MPs in soils negatively impacts soil health, microbial diversity, and the integrity of entire soil ecosystems [12]. MPs can modify soil physical and chemical properties, impair the growth and reproductive capacities of soil organisms, alter microbial community structures, and impede nutrient cycling. Additionally, MPs interact with pollutants such as heavy metals and organic contaminants [13,14], intensifying their detrimental effects on plant health, including impaired root growth, and altering soil parameters like pH and nutrient availability [14].
Despite increasing attention to soil MPs, reliable quantification remains difficult due to the heterogeneity of soil, which consists of complex organo-mineral mixtures with wide particle size distributions [15]. Accurate isolation and identification of MPs in soil are hampered by low recovery rates [16], interference from organic matter [17,18], and variability in particle size detection [19,20]. Density separation methods are commonly employed for isolating MPs from environmental matrices [21]. These methods utilize saturated salt solutions of varying densities, including sodium chloride (NaCl, 1.2 g cm−3), sodium bromide (1.4 g cm−3) and zinc chloride (ZnCl2, 1.7 g cm−3) [22]. While low-density solutions, such as NaCl, are inexpensive and safer to handle, solutions with higher densities (e.g., ZnCl2) are necessary to effectively extract denser plastics like polyvinyl chloride (PVC, density range 1.16–1.58 g cm−3) and polyethylene terephthalate (PET, density range 1.37–1.45 g cm−3) [22]. An alternative method employs low-density oils, leveraging the oleophilic nature of plastics to facilitate MPs separation into an oil layer above an aqueous solution [23]. Additionally, ultrasonic treatment and centrifugation can enhance MPs extraction efficiency [24,25].
Organic matter is another major barrier, as its density can overlap with that of MPs, leading to false positives or analytical interference. Organic digestion is therefore essential, but harsh acid treatments can damage polymers [26]. Thus, milder reagents such as potassium hydroxide (KOH), hydrogen peroxide (H2O2), and enzyme treatments are preferred. Enzymatic digestion has been effective in simpler matrices (e.g., water, marine sediments) [27,28,29], but the diverse and complex organic matter in soil often necessitates multiple enzymes, increasing complexity and cost. H2O2, especially when used with iron catalysts (Fenton’s reagent), efficiently removes organic materials and has demonstrated strong performance in sludge and soil analyses [30]. Elevating reaction temperatures to approximately 50 °C can further improve digestion efficiency without exceeding common MPs’ thermal limits [30].
Post-extraction identification traditionally relies on manual visual inspection, which is subjective and operator-dependent. Advanced techniques, such as Micro-Fourier Transform Infrared Spectroscopy (µ-FTIR) and Micro-Raman Spectroscopy (µ-Raman), provide reliable identification for MPs larger than 1 mm and within the 10–20 µm range, respectively [31,32]. Thermal extraction–desorption gas chromatography–mass spectrometry (TED-GC-MS) allows for precise and efficient quantification but is limited to specific polymer types and inherently destructive, preventing subsequent morphological assessments [33]. Although MP detection methodologies have advanced significantly over the past decade [22,34]. the lack of standardized protocols for complex soil matrices has resulted in inconsistent data and underrepresentation of small particles.
In this context, this study developed and applied a refined extraction workflow for soil MPs that combines two-step NaCl density separation, H2O2 digestion, and fractionated filtration, coupled with high-throughput polymer identification using the Agilent 8700 Laser Direct Infrared (LDIR) imaging system. Two-stage density separation combined with fractionated filtration ensures effective removal of mineral particulates and non-target debris while preserving MPs down to 10 μm. LDIR is a high-throughput, highly sensitive analytical tool capable of accurately identifying MPs down to 10 µm with minimal sample preparation. This study evaluates the performance of the refined protocol through blank and recovery tests and demonstrates its application in agricultural soils from the Hetao Irrigation District, providing new insights into the abundance, size distribution, and polymer composition of soil MPs. The results aim to support the development of reliable and reproducible approaches for terrestrial MPs monitoring.

2. Materials and Methods

2.1. Sample Collection

The samples used in this study were collected from Dengkou County, Wuyuan County, and Urad Front Banner in the Hetao Irrigation District which is one of the largest inland irrigation areas in Asia and is situated in the western part of Inner Mongolia, China. Sampling sites were distributed along the twelve main drainage canals, the sample numbers are S1, S2…… S12 as shown in Figure 1. Before sampling, stainless steel soil augers and shovels were thoroughly rinsed with distilled water. At each sampling location, tools were cleaned to prevent cross-contamination. Soil samples were collected to a depth of 100 cm. Three replicate samples were taken at each site, placed into aluminum boxes, and promptly transported to the laboratory under cooled conditions (−4 °C). The physicochemical characteristic of the soil in Hetao Irrigation District is presented in Table 1. Upon arrival, the samples were dried in an oven at 60 °C until a constant weight was reached. The dried soil was gently ground with a mortar to loosen it, then sieved through a 5 mm stainless steel mesh (manufactured by Shangyu Huafeng Hardware Instrument Co., Ltd., Shaoxing, China). Subsamples of 5 g were taken for MPs detection in soil.

2.2. Materials and Reagents

All solutions were prepared using ultrapure water (18.2 MΩ·cm at 25 °C) from a NeoLab Pure water system. All glassware was thoroughly rinsed with filtered ultrapure water three times before use to minimize contamination.
i.
Sodium chloride (NaCl) (1.2 g cm−3) for density separation was obtained from Xiangyun Huida Co., Ltd. (Beijing, China) (Purity: ≥99.0%). A saturated solution was prepared by dissolving NaCl in ultrapure water.
ii.
Hydrogen peroxide (H2O2) (30% w/w) was purchased from Xiangyun Huida Co., Ltd. (Beijing, China) and used for organic matter digestion without further dilution.
iii.
Anhydrous ethanol (HPLC grade, ≥99.9%) was obtained from Xiangyun Huida Co., Ltd. (Beijing, China) for the final rinsing and concentration steps.
iv.
Metal filter membranes (10 µm pore size, 47 mm diameter) were custom-made by Jiuding High-Tech Filtration Equipment Co., Ltd. (Beijing, China).
v.
Custom vacuum filtration device was fabricated by Boyuan Hongda Biotechnology Co., Ltd. (Beijing, China).
vi.
Standard reflective slides for Laser Direct Infrared (LDIR) analysis were purchased from Agilent Technologies (Santa Clara, CA, USA).
vii.
Polyethylene (PE MP) with 100 µm was purchased from Si Ye Zi Chemical Co., Ltd. (Dongguan, China) for quality control.

2.3. MPs Extraction Methods

To minimize interference from suspended soil particles that could compromise analytical accuracy, this study adopted a modified density separation method for MPs extraction. An additional density separation step was incorporated to improve purification efficiency, followed by oxidative digestion using H2O2. The complete extraction and analysis workflow is illustrated in Figure 2.
Step 1: Density Separation
The sieved soil sample (5 g) was placed in a beaker with saturated NaCl solution (density 1.2 g cm−3). The mixture was stirred using a glass rotor on a magnetic stirrer for 30 min and then allowed to settle for 4–6 h. Once the supernatant was clear, it was decanted into a clean beaker. This density separation procedure was repeated three times for each sample.
Step 2: Filtration and Organic Matter Digestion
The combined supernatants were filtered through a custom-made 10 μm metal filter membrane. Material retained on the filter was rinsed into a beaker with 30 mL of 30% H2O2, added incrementally until bubbling ceased. The solution was digested at room temperature for 2–3 days to remove organic matter.
Step 3: Fractionated Filtration Method (FFM) and Large Particle Analysis
The digested solution was passed through a sequence of 500 μm and 10 μm metal sieves to separate particles into two size classes: 10–500 μm and 500–5000 μm. Particles larger than 500 μm were observed under a stereomicroscope and manually isolated using tweezers. These were transferred to standard reflective slides and analyzed using the LDIR imaging system. Particles smaller than 500 μm were retained for further purification.
Step 4: Secondary Density Separation for Fine Particles
Fine particles (<500 μm) retained on the 10 μm filter were rinsed into an Erlenmeyer flask with saturated NaCl solution and filled close to the rim. The suspension was allowed to settle for 12 h. A custom vacuum filtration device, featuring a funnel with a 30° angled outlet submerged just below the liquid surface, was used for separation. The use of a custom vacuum filtration device with angled outlet and reduced pressure minimizes turbulence and sample loss during fine particle recovery. Vacuum suction from the top maintained low pressure inside the funnel, enabling the supernatant to flow into a clean beaker. The setup was rinsed with ultrapure water. This process was repeated three times.
Step 5: Ethanol Rinse and Concentration
The final supernatants were filtered using a 10 μm membrane. The filter was immersed in chromatography-grade anhydrous ethanol in a clean glass tube and sonicated for 30 min. The membrane was then removed and rinsed with additional ethanol. The ethanol extract was concentrated on a heating plate.
Step 6: Final Sample Preparation for Analysis
The concentrated extract was transferred into a 1.5 mL chromatography vial, rinsed with ethanol, and further reduced to a final volume of 100 μL on a heating plate. The ethanol’s volatility was used to uniformly distribute the sample on a standard reflective slide. Final identification of MPs was performed using the Agilent 8700 LDIR imaging system.

2.4. Quantity Control

To minimize contamination, all procedures were conducted within a controlled laboratory environment. Wherever possible, aluminum, glass, and stainless-steel materials were used for all equipment. All liquids used during the experiments were filtered through 10 μm stainless steel membranes. To further reduce the risk of sample contamination, all glassware was rinsed with ultrapure water prior to use and covered with aluminum foil. Soil samples were similarly protected using aluminum foil. During laboratory operations, nitrile gloves were worn and 100% cotton laboratory coats were used. Three replicate analyses were conducted for each sampling site to minimize variability.
Despite these precautions, background contamination could not be entirely avoided. Between 5 and 8 particles (<500 μm) were detected in the blank control samples. Accordingly, the number of MPs detected in each soil sample was corrected by subtracting the quantity identified in the corresponding control samples. To determine the recovery rate, 50 pcs of PE MPs were mixed into 5 g of pure plastic-free sand. The mixture was then pretreated and analyzed by following the standard extraction protocol and the experiment was conducted in triplicate. The particles were also counted using an Agilent 8700 LDIR imaging system. The recovery rate of MP in artificially spiked soil can be calculated using Equation (1). From the results, the recovery rate was more than 85%.
R = p 1 − p 2 p 1 × 100 %
where R is recovery rate (%) of MP sample; p 1 is the spiked number of MPs; p 2   is the particles of MP extracted from spiked soil sample.

2.5. Rationale for LDIR Analysis

The quantification and characterization of MPs were performed using Laser Direct Infrared (LDIR) imaging (Agilent 8700). While Fourier-Transform Infrared (μFTIR) and Raman spectroscopy are considered gold standards for polymer identification [35,36], LDIR was selected for this study due to its superior analysis speed and automated counting capabilities, which were essential for processing the large number of samples required to assess spatial heterogeneity across the irrigation district.
Unlike traditional techniques that can be time-intensive for mapping entire filters, LDIR’s quantum cascade laser enables rapid chemical imaging, allowing for a more comprehensive analysis of the sample deposit without subjective operator bias [37]. This is particularly advantageous for detecting small particles (>10 µm) which are easily missed during manual counting. A comparison of key analytical techniques relevant to the small MP size range is provided in Table 2.
A known limitation of the LDIR spectral range (975–1800 cm−1) is that it may not identify polymers with characteristic peaks outside this window. To mitigate this, the custom polymer library is included validated spectra for polymers most commonly used in agricultural applications.

2.6. MP Quantification and Abundance Calculation

To quantify MP abundance in soil, the final MP extract from each sample (originally derived from 5 g of dry soil) was deposited onto a reflective slide and uniformly dried for LDIR analysis. The raw particle count from the scanned area was extrapolated to estimate the total number of particles on the entire filter. The MP abundance was then calculated on a dry mass basis (particles per kilogram of soil) using the following equation:
A b u n d a n c e   p a r t i c l e s k g = N c o u n t e d M s o i l   ×   1000   g
where N c o u n t e d is raw particle count on the scanned are; M s o i l   is dry mass (in gram) of the soil sample processed.

3. Results

3.1. Concentration of MPs

MPs were detected at all 12 sampling sites with abundance values ranging from 5778 to 31,489 particles/kg and an average concentration of 16,461 ± 8097 particles/kg. As shown in Figure 3, the abundance of MPs exhibited significant spatial variability across the Hetao Irrigation District. Statistical differences among sites are denoted by different lowercase letters (a–f) based on p < 0.05. From the result, Site S8, located near the Sixth Drainage Canal in the central-northern part of the irrigation system, exhibited the highest MP concentration of 31,489 particles/kg (a). Sites S2 and S3 also showed high MP abundances, both exceeding 20,000 particles/kg and classified as statistically similar (b), suggesting consistent MP sources in that region. In contrast, the lowest abundances were recorded at sites S4, S5, and S6, with values below 10,000 particles/kg. Notably, S6 had the lowest MP concentration overall and was significantly different from most other sites (f). Sites S1, S7, S9, S10, S11, and S12 formed an intermediate group with abundances ranging from approximately 12,000 to 18,000 particles/kg. These sites were statistically categorized into groups Figure 2, c–e indicating moderate but variable levels of contamination.
Doorgha et al. [39] used attenuated total reflectance Fourier-transform infrared (ATR-FTIR) spectroscopy to detect MPs in agricultural soils in Mauritius, reporting concentrations ranging from 320.0 ± 112.2 to 420.0 ± 244.0 particles/kg. Mercy et al. [40] investigated irrigated farmland soils in Arusha, Tanzania, using dissection microscopy combined with FTIR analysis, and reported MP concentrations between 0.21 and 1.5 items/g. Corradini et al. [41], applying visual inspection with a stereomicroscope, examined urban soils in Melipilla, Chile, and found MP concentrations ranging from 0.6 to 10.4 items/g. The MP abundances observed in this study were higher than those reported in the above studies. This discrepancy may be attributed to elevated MP retention and accumulation levels in the Hetao Irrigation District. This further supports the view that the Hetao Irrigation District, as one of Asia’s largest inland irrigation regions with over 2000 years of agricultural history, is particularly vulnerable to plastic accumulation due to long-term and widespread use of plastic-based agricultural materials.

3.2. Types of MPs

The types of MPs detected at different sampling sites. Figure 4 shows that polypropylene (PP) and polyamide (PA) were the most frequently detected MP types, both exhibiting high abundance across all samples. PP and PA were present at every site, with PP accounting for 47.83% and 59.69% of the total MPs at sites S1 and S2, respectively, while PA comprised 64.94% and 66.15% of MPs at S4 and S5. Polyethylene (PE) and high-density polyethylene (HDPE) were also widely detected across the sampling points. Totally, polypropylene (PP), polyamide (PA), and polyethylene (PE) were the most prevalent polymer types (>90%). Previous studies on soil MPs have commonly reported PE, PP, PVC, PA, and PS as the dominant types [42,43,44]. PE is widely used in greenhouses and plastic mulching due to its thermal retention and flexibility. PP is typically used in ropes and packaging materials. PVC is often employed in agricultural drip irrigation systems. PA serves in textiles and food packaging, while PS, known for its clarity and gloss, is also commonly used for packaging [16].
In this study, the LDIR imaging system was used for analysis, offering high-throughput automated identification with minimal manual handling, thereby reducing contamination risks. The results differ from those obtained using traditional analytical techniques such as Raman spectroscopy or FTIR spectroscopy, which primarily identify PP and PE. In contrast, focal plane array (FPA)-based FTIR imaging tends to detect a broader range of plastic types [45]. These discrepancies underscore the need to standardize MP analytical methods to improve the comparability of findings across different studies.

3.3. Particles Size of MPs

Figure 5 illustrates the size distribution of MPs at different sampling points. Most detected MPs were smaller than 500 μm, accounting for 99.76% of the total. The average particle size across all sites was 47.09 μm. The minimum particle sizes were relatively consistent across all sampling sites, with a mean value of 20.20 μm. Liu et al. [46] investigated five agricultural soil sites in the Hetao Irrigation District using microscopy combined with laser confocal microscopy, reporting MP abundances ranging from 247.40 to 278.68 particles/kg. Bai et al. [47] conducted a survey across nine counties in the same region using Nile red-assisted visual inspection and Fourier transform infrared spectroscopy, detecting MP abundances ranging from 1810 to 86,331 items/kg of dry soil. In their study, the most frequently observed particle size range was 50–100 μm. In contrast, most MPs detected in the present study were in the 10–30 μm range.
The Sankey diagram in Figure 6 reveals distinct patterns in the size distribution of various MPs polymer types detected in soil samples. Polytetrafluoroethylene (PTFE) exhibited a strong concentration in the 10–30 μm size class. For polymers such as polycarbonate (PC), polylactic acid (PLA), polymethyl methacrylate (PMMA), polyurethane (PU), and polyvinyl chloride (PVC), the number of MPs detected in the 10–30 μm range exceeded that in the 30–100 μm range. In contrast, HDPE and PE exhibited a relatively even distribution between the 10–30 μm and 30–50 μm ranges, each accounting for about 30% of the total particles detected for those types. This indicates that PE-based MPs, which are widely used in mulching films and irrigation pipes, persist in soil across a broader size spectrum, likely due to their high production volume and variable degradation resistance. Polyethylene terephthalate (PET) exhibited the most uniform distribution among all detected polymers, with its abundance spread relatively evenly across the entire size range. Specifically, PET particles were distributed at 23%, 14%, 22%, 13%, 10%, and 15% across the 10–30 μm, 30–50 μm, 50–100 μm, 100–150 μm, 150–200 μm, and >200 μm size intervals, respectively.

4. Discussion

This study presents a comprehensive investigation into the extraction, identification, and characterization of MPs in agricultural soils within the Hetao Irrigation District which is the one of Asia’s largest inland irrigation regions with a long-standing history of plastic-intensive agriculture. The observed variability and abundance of MPs across 12 sampling points are not only indicative of regional contamination patterns but also reflect the enhanced detection capability afforded by the optimized extraction protocol developed in this study.
The combination of two-step density separation, hydrogen peroxide digestion, and Agilent 8700 LDIR imaging system represents a significant methodological advancement in MP analysis for complex soil matrices. The repeated density flotation increases the extraction efficiency, while H2O2-based digestion minimizes the polymer degradation and removes interfering organics. LDIR-based analysis enables the high-throughput and particle-level chemical identification across a wide size range. The resulting recovery efficiency exceeding 85%, combined with minimal background contamination (5–8 particles in blanks) shows the reliability and reproducibility of the proposed protocol. More importantly, the methodological refinement enabled the detection of particles as small as 10 μm, which are typically underrepresented in soil studies due to analytical resolution limits.
MPs were identified at all sampling sites, with concentrations ranging from 5778 to 31,489 particles/kg and an average of 16,461 ± 8097 particles/kg (Figure 2). These values far exceed those reported in global studies and reveal that this discrepancy is not solely due to regional accumulation, but also a reflection of improved detection sensitivity offered by the methodological optimization. The detailed polymer profiling revealed that PP and PA were the dominant polymers across all sites, followed by PE and HDPE (Figure 3). PP is most probably dominated by the degradation of plastic twines and ropes for baling and tying, as well as packaging materials such as fertilizer and feed sacks [48]. While the potential local sources for PA are the use of synthetic fishing nets and ropes in aquaculture, which is commonly in water-rich irrigated areas, and agricultural nets used for crop protection [49]. Similar findings are summarized in Table 3. These findings are consistent with their widespread agricultural applications, such as in mulch films, irrigation systems, ropes, and textile packaging.
At the concentrations reported, MPs can directly alter soil structure. The vast number of small, predominantly fibrous PP and PA particles can affect soil porosity, water infiltration, and water-holding capacity [51]. In addition, MPs can be ingested by a wide range of soil fauna, including earthworms, nematodes, and microarthropods, potentially causing physical harm and toxicological effects [52]. Furthermore, MP surfaces serve as novel substrates for distinct microbial communities (the “plastisphere”), which can alter the broader soil microbiome and its essential functions, such as organic matter decomposition, nutrient cycling (N, P), and soil respiration [53,54]. The high surface area of the small MPs we detected vastly increases this interface for microbial colonization, potentially disrupting these critical ecological processes [52]. There is growing evidence that MPs can be taken up by plant roots and translocated to aerial tissues. It results in root uptake, root damage, cracks, or association with root exudates could facilitate entry. Once inside, MPs could induce oxidative stress, impair growth, and ultimately affect crop yield and quality [55].
Notably, biodegradable MPs such as PLA were also detected at several sites, although at lower proportional. This highlights both the feasibility and emerging importance of monitoring degradable polymer residues in agricultural soils. However, due to their hydrophilic and oxidative-sensitive nature, biodegradable MPs are more prone to partial degradation during H2O2 digestion or loss during density separation, especially when using saturated NaCl, which may not float certain biodegradable polymers with higher densities [56,57]. Nevertheless, the successful identification of PLA using LDIR demonstrates the method’s capability in detecting a broader spectrum of MPs, including those from degradable sources, and underlines its applicability for future assessments of biodegradable plastics in terrestrial environments.
In terms of particle size distribution, 99.76% of all detected MPs were <500 μm, with a dominant size range of 10–30 μm (Figure 4). This ultrafine particle detection reflects the methodological strength of LDIR imaging, which not only identifies particles based on chemical composition but also retains their morphological information without destruction. The Sankey diagram (Figure 5) further illustrates the link between polymer type and particle size, showing that polymers like PTFE, PMMA, PLA, and PVC are predominantly present in finer size fractions, while PE, HDPE, and PET span a broader size spectrum. The implementation of Agilent’s 8700 LDIR system facilitated precise polymer identification across a wide particle size spectrum.
In this study, soil samples were collected from a relatively homogeneous agricultural zone, and their key properties are summarized in Table 1. The soils were alkaline (pH 8.65) and moderately saline (510 ± 15 mg/kg), with a total organic matter content of 12.98 ± 0.34 g/kg. The cation exchange capacity was relatively high (Ca: 5400 ± 75 mg/kg; Mg: 1350 ± 90 mg/kg), while available nitrogen, phosphorus, and potassium concentrations were 118.34 ± 35.21, 42.99 ± 15.19, and 92.33 ± 28.36 mg/kg, respectively. The relatively low organic content and alkaline pH likely minimized organic coating and degradation effects, facilitating more effective peroxide digestion and cleaner LDIR imaging. In contrast, soils with higher humic content or acidic pH might lead to greater polymer masking or oxidative damage, reducing recovery and spectral clarity [58]. Similarly, the moderate salinity and ionic composition may have influenced the ionic strength and dispersion behavior of fine MP particles, although no clear inhibitory effects were observed in this case.

5. Limitations

While recovery tests were successfully conducted for 100 µm particles, the quantitative accuracy for the dominant size fraction observed in the samples (10–30 µm) remains uncertain due to the lack of size-specific validation. It is well-established that extraction efficiency for MPs is strongly size-dependent, with lower recovery rates expected for smaller particle sizes [20,22]. This is primarily due to increased adhesion to glassware and equipment surfaces, higher potential for loss via aspiration or during filtration steps, and challenges in visual identification during transfer. Hence, the absolute abundances reported for the 10–30 µm size fraction are likely underestimates. The reported values should be interpreted as conservative, operational estimates based on the method rather than absolute environmental concentrations. However, the consistent application of the method across all samples allows for robust relative comparisons of MP abundance and distribution between the sampled sites within this study. The potential for particle loss in the smallest size fractions means that the true dominance of MPs in the 10–30 µm range may be even greater than we report. The results highlight the critical need for future studies to employ size-specific validation protocols, especially when targeting nanoplastics and small MPs. Future work should employ validated methods for sub-50 µm particles to better constrain the quantitative accuracy for this environmentally significant size fraction.

6. Conclusions

This study contributes to the broader effort to standardize soil MP extraction and identification protocols. The lack of harmonized methodologies has historically limited the comparability of MP research across geographic regions and environmental matrices. By demonstrating a repeatable, high-efficiency protocol applicable to heterogeneous soil environments, this study addresses a critical gap in the field. The successful coupling of chemical digestion, size-selective separation, and automated spectroscopic analysis provides a methodological blueprint for future soil MP research. This approach proved highly effective for the alkaline, low-organic matter soils of the Hetao Irrigation District, which likely minimized common interferences like organic coating and facilitated cleaner with IR-based analysis. As such, the proposed workflow has the potential to serve as a reference protocol for environmental monitoring agencies and research institutions aiming to quantify MPs in terrestrial systems with both accuracy and efficiency.

Author Contributions

C.M.H.: Writing—original draft; W.F.: Writing, Reviewing, Supervision and Funding; Y.D.: Experiment, Data Collection, Data Analysis; X.L.: Reviewing and editing; S.K.N.: Reviewing and editing. All authors have read and agreed to the published version of the manuscript.

Funding

This research was jointly supported by the Key Science Technology Project of Inner Mongolia Province (2022YFHH0044), the Science and Technology Major Project of Ordos City (ZD20232301) and the National Natural Science Foundation of China (42177400).

Data Availability Statement

The original contributions presented in this study are included in the article. Further inquiries can be directed to the corresponding author.

Conflicts of Interest

Author Xiaofeng Li was employed by the company China Hebei Construction and Geotechnical Investigation Group Ltd. The remaining authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

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Water 17 02700 g001
Figure 1. Sampling sites.
Figure 1. Sampling sites.
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Figure 2. Optimized extraction method of MPs.
Figure 2. Optimized extraction method of MPs.
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Figure 3. MPs abundance in 12 sampling sites. a, b, c, d, e, and f indicate levels of statistical significance; different letters represent significant differences between groups.
Figure 3. MPs abundance in 12 sampling sites. a, b, c, d, e, and f indicate levels of statistical significance; different letters represent significant differences between groups.
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Figure 4. Composition and relative abundance of MPs polymer types at different sampling sites.
Figure 4. Composition and relative abundance of MPs polymer types at different sampling sites.
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Figure 5. Size Distribution of MPs Detected at Different Sampling Sites.
Figure 5. Size Distribution of MPs Detected at Different Sampling Sites.
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Figure 6. Relationship between MPs types and their corresponding size characteristics. The left side shows the identified polymer types, while the right side presents the particle size categories (10–30 μm, 30–50 μm, 50–100 μm, 100–150 μm, 150–200 μm, and >200 μm). The width of each flow line represents the relative abundance of each polymer within the corresponding size range.
Figure 6. Relationship between MPs types and their corresponding size characteristics. The left side shows the identified polymer types, while the right side presents the particle size categories (10–30 μm, 30–50 μm, 50–100 μm, 100–150 μm, 150–200 μm, and >200 μm). The width of each flow line represents the relative abundance of each polymer within the corresponding size range.
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Table 1. Physicochemical characteristics of soil from the Hetao Irrigation District.
Table 1. Physicochemical characteristics of soil from the Hetao Irrigation District.
Physicochemical CharacteristicValue
pH8.65
Salinity (mg/kg)510 ± 15
Organic matter (g/mg)12.98 ± 0.34
Exchangeable calcium (mg/kg)5400 ± 75
Exchangeable magnesium (mg/kg)1350 ± 90
Available nitrogen (mg/kg)118.34 ± 35.21
Available phosphorus (mg/kg)42.99 ± 15.19
Available potassium (mg/kg)92.33 ± 28.36
Table 2. Gold Standard Methods for MP Detection.
Table 2. Gold Standard Methods for MP Detection.
MethodSize RangeKey AdvantagesMajor Limitations
FTIR [36]10–20 μm to 5 mmExcellent polymer identification, Quantitative capabilities, relatively standardizedLimited spatial resolution, Time-consuming mapping
Raman [35]1 μm to 5 mmSuperior spatial resolution, no water interference, Detailed molecular informationFluorescence interference, longer analysis times
Py-GC-MS [38]Not size-specificExcellent sensitivity, Unambiguous identification, Provides mass concentrationDestructive, No particle information, Complex sample preparation
LDIR10 μm to 5 mmRapid analysis, Automated counting, good sensitivityLimited spectral range, Polymer library dependence
Table 3. Comparison of Dominant Polymer Profiles in Selected Agricultural Soil Studies.
Table 3. Comparison of Dominant Polymer Profiles in Selected Agricultural Soil Studies.
Study LocationDominant Polymer(s)Postulated Main SourcesReference
Hetao Irrigation District, ChinaPolypropylene (PP)
Polyamide (PA)		Agricultural nets, agricultural irrigation, aquaculture activitiesThis study
Vegetable farmlands of suburb Wuhan, ChinaPolyamide (PA)
Polypropylene (PP)		Plastic mulch films, sewage and wastewater[48]
Cotton fields in Xinjiang Uygur Autonomous Region, ChinaPolyethylene (PE)Plastic mulch films[50]
Jiangshe Modern Agricultural Demonstration Park, ChinaPolypropylene (PP)
Polyethylene (PE)
Polyamide (PA)		Agricultural film, domestic wastewater[49]
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Ho, C.M.; Feng, W.; Deng, Y.; Li, X.; Ngien, S.K. Extraction and Characterization of Microplastics in Soil: A Case Study from the Hetao Irrigation District. Water 2025, 17, 2700. https://doi.org/10.3390/w17182700

AMA Style

Ho CM, Feng W, Deng Y, Li X, Ngien SK. Extraction and Characterization of Microplastics in Soil: A Case Study from the Hetao Irrigation District. Water. 2025; 17(18):2700. https://doi.org/10.3390/w17182700

Chicago/Turabian Style

Ho, Chia Min, Weiying Feng, Yuxin Deng, Xiaofeng Li, and Su Kong Ngien. 2025. "Extraction and Characterization of Microplastics in Soil: A Case Study from the Hetao Irrigation District" Water 17, no. 18: 2700. https://doi.org/10.3390/w17182700

APA Style

Ho, C. M., Feng, W., Deng, Y., Li, X., & Ngien, S. K. (2025). Extraction and Characterization of Microplastics in Soil: A Case Study from the Hetao Irrigation District. Water, 17(18), 2700. https://doi.org/10.3390/w17182700

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