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

# Chemical imaging reveals phosphorus mobilization patterns in earthworm-engineered drilosphere

Zhen-Yu Qiang<sup>a</sup>, Dong-Xing Guan<sup>a,∗</sup> <sup>●</sup>, Jia-Lu Gao<sup>a</sup>, Gang Li<sup>b</sup>, Daniel Menezes-Blackburn<sup>c,†</sup>,
Anna Gunina<sup>d,e</sup>, Yvan Capowiez<sup>f,●</sup>, Lena Q. Ma<sup>a</sup>

## ARTICLE INFO

Handling Editor: Dr. Andrew Margenot

### Keywords:

Spatial heterogeneity

Diffusive gradients in thin-films (DGT)

Soil zymography

Epidermal mucus

Burrow

## ABSTRACT

Earthworms are soil ecosystem engineers who play a crucial role in phosphorus (P) cycling, encompassing inorganic P desorption and organic P mineralization. Although the role of earthworms in P mobilization is well documented, the spatial distribution of P mobilization processes and critical microbial species within the earthworm-engineered soil microhabitat, known as the drilosphere, remains to be determined. This study investigated P mobilization and redistribution in the drilosphere by *ex situ* chemical analyses, *in situ* soil zymography and diffusive gradients in thin-films imaging, and assessment of the microorganism communities. Endogeic earthworm species *Metaphire guillelmi* was incubated for 30 days in pots filled with soils Fluvisol or Acrisol with different total P contents (0.6 and 1.2 mg kg⁻¹, respectively). Chemical analyses revealed that total P content in earthworm casts increased by 120 % in Fluvisol and 7.7 % in Acrisol. Available P content increased by 8.5 times in Fluvisol and 4.4 times in Acrisol in the drilosphere compared to bulk soil due to elevated acid/ alkaline phosphatase activities and intensified Fe/Al-bound P desorption amongst others. Imaging identified co- existing and distinct hotspots for available P and acid phosphatase activity in soils surrounding the burrow walls, with hotspot proportions of 0.1 %–3.1 % and 5.4 %–7.5 % of the imaged areas. Earthworm activity increased the abundance of specific bacterial (*Aeromonas* and *Flavobacterium*) and fungal (*Scedosporium* and *Podospora*) taxa potentially contributing to P mobilization. The correlation between the distribution of metal-bound adsorption sites, phosphatase activity, and diffusion mechanisms contributed to the available P redistribution in the soil. These findings provide insights for developing soil management strategies that harness earthworm-mediated P cycling to optimize nutrient use efficiency and reduce external P inputs in sustainable agricultural systems.

## 1. Introduction

Phosphorus (P) is a critical element in soil and often a limiting nutrient for crop yield, terrestrial ecosystem functioning, and planetary health (Liu et al., 2024; Zou et al., 2022). Despite the vast global soil P reservoir, available P is often insufficient to support optimal crop growth. It is estimated that the global soil total P in the 0–50 cm depth is 40.6 Pg, of which only 3.6 Pg is labile inorganic P and 8.6 Pg is organic P, collectively representing just 30 % of the total P content (Yang et al.,

2013). Soil P availability is influenced by multiple factors, including pH, soil organic matter (SOM), metal oxides, and microbial and enzyme activities. Soil pH affects P availability by altering the solubility of metal oxides, with P typically bound to iron/aluminum (Fe/Al) oxides in acidic soils and calcium carbonate in alkaline soils (Chu and Chang, 1966). SOM can compete for binding sites on metal oxides, leading to P desorption (Weng et al., 2011). Additionally, microbial activity, particularly that of P-solubilizing microorganisms, facilitates inorganic P desorption and organic P mineralization through the release of protons

<footnote><sup>a</sup> State Key Laboratory of Soil Pollution Control and Safety, Zhejiang Provincial Key Laboratory of Agricultural Resources and Environment, Institute of Soil and Water Resources and Environmental Science, College of Environmental and Resource Sciences, Zhejiang University, Hangzhou 310058, China</footnote>

<footnote><sup>b</sup> State Key Laboratory of Regional and Urban Ecology, Ningbo Observation and Research Station, Institute of Urban Environment, Chinese Academy of Sciences, Xiamen 361021, China</footnote>

<footnote><sup>c</sup> Department of Soils, Water and Agricultural Engineering, CAMS, Sultan Qaboos University, PO Box 34, Al-khod 123, Sultanate of Oman</footnote>

<footnote><sup>d</sup> Department of Soil Biogeochemistry, OWL University of Applied Sciences and Arts, Lippe 37671, Germany</footnote>

<footnote><sup>e</sup> Institute of Applied Ecology, Chinese Academy of Sciences, Shenyang 110016, China</footnote>

<footnote><sup>f</sup> INRAE, UMR1114, EMMAH, INRAE/Université d'Avignon, Site Agroparc, 84914 Avignon cedex 09, France</footnote>

<footnote>* Corresponding author.
E-mail address: dxguan@zju.edu.cn (D.-X. Guan).</footnote>
and phosphatases (Rawat et al., 2021). In this context, soil fauna, especially earthworms, play a crucial role in modifying these factors and influencing P cycling in terrestrial ecosystems.

Earthworms are “ecosystem engineers” that improve soil fertility and health through processes such as organic carbon (C) stabilization, intensification of nitrogen (N) and P cycling, and increasing nutrient availability for plants (Jouquet et al., 2006; Lavelle et al., 1997). A meta-analysis investigating the impact of earthworm casts on P mobilization has confirmed the positive influence of earthworms on nutrient cycling (van Groenigen et al., 2019). The mechanisms of P solubilization mediated by earthworms have been a focus of research and debate. Key aspects include 1) earthworms stimulating soil microbial activities to increase phosphatase activity and organic P cycling (Chiba et al., 2024); 2) microbial activities leading to the release of dissolved organic matter with negatively charged functional groups that bind to the same positively charged sites on clay and metal oxides where phosphate would otherwise adsorb, thereby displacing inorganic P and increasing its desorption and solubilization (Vos et al., 2022b); and 3) earthworm activity altering soil pH, consequently influencing P speciation and solubility (Vos et al., 2019).

While extensive research has been conducted on earthworm-mediated P mobilization in soils using bulk chemical analysis, the role of earthworms as drivers of stimulating microbial activities in soils warrants further exploration. Similar to roots in the rhizosphere and litter in the detritusphere, earthworms create microbial hotspots known as the “drilosphere” (Andriuzzi et al., 2013). The drilosphere is defined as the zone influenced by earthworm activities, typically extending a few millimeters around the burrow walls (Brown et al., 2000). Some theories also include the casts produced at the soil surface or within the soil as part of the drilosphere (Dallinger and Horn, 2014). However, for clarity in this study, we refer to the drilosphere as only the few millimeters around the burrows (Rogasik et al., 2014), distinguishing it from casts. Understanding the mechanisms and dynamics of P cycling within the drilosphere is crucial for developing sustainable agricultural practices that optimize nutrient availability while minimizing environmental impacts (Luo et al., 2024; McDowell et al., 2024).

In *situ* chemical imaging tools, including soil zymography for hydrolytic enzyme activities and diffusive gradients in thin-films (DGT) for labile chemicals, have been used to investigate biogeochemical processes in soil hotspots (such as rhizosphere, detritusphere, and drilosphere). These techniques provide spatially explicit information on nutrient dynamics and enzyme activities (Guan et al., 2022; Li et al., 2023). For example, the shift in plant P acquisition strategy through cluster root formation was revealed, with a 20 % increase in phosphatase activities surrounding the cluster roots observed using soil zymography (Ma et al., 2019). A combination of DGT and soil zymography was applied to elucidate P depletion and revealed increased phosphatase activities in the rhizosphere and detritusphere, following root death and microbial P acquisition (Fang et al., 2021). Additionally, soil zymography provided spatial insights into C and N turnover under earthworm bioturbation (Hoang et al., 2020, Hoang et al., 2016). Recent advances in machine learning and artificial intelligence will offer potential for automated analysis of soil imaging data, enabling more efficient identification of biogeochemical hotspots across diverse environmental conditions (Ahkami et al., 2024; Koley, 2023). Nevertheless, these imaging techniques have seldom been applied to elucidate the mechanism of P dynamics in the drilosphere and visualize the P distribution surrounding the burrow walls.

This study explored the influence of endogeic earthworms (*Metaphire guillelmi*) on P mobilization and spatial heterogeneity within their drilosphere. By combining *ex situ* chemical analysis, *in situ* imaging, and microbial analysis, this study aimed to i) capture P spatial distribution in the drilosphere, providing a detailed understanding of the heterogeneity in P availability within this type of hotspots, and ii) explore the mechanisms involved in P mobilization within the drilosphere, including the role of earthworm activities, physicochemical processes, and microbial

interactions. We hypothesized that: i) the drilosphere functions as a distinct P and microbial hotspot with quantifiable spatial patterns, where available P and phosphatase activities would exceed those in bulk soil and be detectable using *in situ* imaging techniques; and ii) P mobilization mechanisms in the drilosphere would differ between soils with contrasting P availability, involving specific earthworm-mediated physicochemical processes (altered phosphatase activities and reactive mineral metal oxide content) and selective enrichment of P-solubilizing microorganisms, with these mechanisms being more pronounced in P-limited soil. This research provides novel insights into P biogeochemical processes in the drilosphere through chemical imaging, advancing our understanding of nutrient turnover in terrestrial ecosystems.

## 2. Materials and methods

### 2.1. Soils and earthworms

Two topsoil samples (0–20 cm) were collected from distinct locations in Zhejiang Province, China, and were used to verify the effects of earthworm-mediated P mobilization. The first sample, classified as a Fluvisol, was obtained from fallow land in Hangzhou (30°18′09″ N, 120°04′35″ E) that had been uncultivated for a decade. The second sample, classified as an Acrisol, was sourced from actively cultivated farmland in Jinhua (29°10′32″ N, 119°31′49″ E) with a history of crop cultivation using mineral fertilizers and manure application (Fig. S1). The physicochemical properties of both soils are presented in Table 1. Variations in total Fe/Al oxides and organic C content were also considered in the two soils, as these factors influence soil P biogeochemical processes in addition to soil total P content (Vos et al., 2023).

The earthworm species *Metaphire guillelmi* was selected for this study due to its widespread presence in agricultural soils and its capacity to create macropores (Scheu, 1991). Assigning an ecological category to Asian earthworms is challenging, as these categories were originally defined for European Lumbricidae. In literature, *M. guillelmi* has been considered both anecic and endogeic (Bottinelli et al., 2020). For the present study, we classify this species as endogeic based on our observations of its presence within the soil and the absence of surface casts at the collection site.

The earthworm specimens were sourced from a local earthworm farm. Prior to the cultivation experiment, the earthworms were acclimatized to the soil with additional fresh cow manure to ensure their vitality. During this acclimatization period of 14 d, environmental conditions were maintained as follows: temperature, 20–25 °C; light–dark cycle, 12 h each; and soil moisture at approximately 60 % of the water holding capacity (WHC). For the pot experiment, adult earthworms (2–3 g, 5–10 cm) were used, while smaller adult earthworms (~1 g, ~5 cm) were selected for the imaging experiment to better suit the container size. All experimental earthworms had distinct clitella, which are considered indicators of adulthood and species maturity (Deng et al., 2022). The use of smaller earthworms for imaging was based on the need for optimal visualization in confined terrarium spaces. The smaller earthworms allowed for clearer imaging and observation of soil modifications in confined spaces, which is crucial for detailed analysis of P dynamics in the drilosphere. The ratio of earthworm biomass to soil volume was maintained consistently between pot and imaging experiments to minimize differences in their impact on soil P cycling (Heinze et al., 2021).

### 2.2. Experimental design

The experimental system comprised an *ex situ* pot experiment, *in situ* chemical imaging, and microbial analysis (Fig. 1), designed to explore the mechanisms of P mobilization in the drilosphere and the distribution patterns of earthworm-mediated hotspots related to available P and acid phosphatase.

The pot experiment was conducted with Fluvisol and Acrisol soils to
<table><caption>**Table 1**
Physico-chemical properties of soils used in the study.</caption><thead><tr><th>Soil</th><th>pH-CaCl<sub>2</sub></th><th>CEC (cmol kg<sup>-1</sup>)</th><th>TP (g kg<sup>-1</sup>)</th><th>AP (mg kg<sup>-1</sup>)</th><th>TFe (g kg<sup>-1</sup>)</th><th>Fe<sub>ox</sub> (g kg<sup>-1</sup>)</th><th>Tal (g kg<sup>-1</sup>)</th><th>Al<sub>ox</sub> (g kg<sup>-1</sup>)</th><th>TC (g kg<sup>-1</sup>)</th><th>TOC (g kg<sup>-1</sup>)</th><th>C/P</th><th>Sand (%)</th><th>Silt (%)</th><th>Clay (%)</th></tr></thead><tbody><tr><td>Fluvisol</td><td>5.7</td><td>24.9</td><td>0.6</td><td>9.8</td><td>31.2</td><td>2.4</td><td>30.4</td><td>0.7</td><td>10.8</td><td>7.3</td><td>18.0</td><td>10.5</td><td>39.7</td><td>49.8</td></tr><tr><td>Acrisol</td><td>7.1</td><td>24.0</td><td>1.2</td><td>30.3</td><td>18.6</td><td>2.7</td><td>18.7</td><td>1.0</td><td>16.8</td><td>15.9</td><td>14.0</td><td>29.2</td><td>13.0</td><td>57.8</td></tr></tbody></table>

Note: CEC = cation exchange capacity; TP = total phosphorus; AP = available phosphorus; TFe = total iron; Fe<sub>ox</sub> = ammonium oxalate extracted iron; TAl = total aluminum; Al<sub>ox</sub> = oxalate-extractable aluminum; TC = total carbon; TOC = total organic carbon; C/P = carbon-to-phosphorus ratio.

<figure><img src="image_2.png" /><figcaption>**Fig. 1.** Schematic representation of the experimental procedure used to investigate earthworm-driven phosphorus cycling in the soil, focusing on the drilosphere. The experimental system combined *ex situ* chemical analysis (a), *in situ* chemical imaging (b) and microbial analysis (c). *Ex situ* chemical analysis composed of available P content and acid / alkaline phosphatase activity. *In situ* imaging comprised soil zymography for enzyme activity and diffusive gradients in thin-films with colorimetric imaging densitometry (DGT-CID) for labile P distribution. Microbial analysis used amplicon sequencing of the bacterial 16S rRNA gene (V4) and the fungal ITS2 region to assess community composition, diversity, and relative abundance.</figcaption></figure>

provide contrasting available P content environments under the controlled cultivation conditions described in Section 2.1. Three distinct sample types were collected from each earthworm treatment pot: bulk soil, drilosphere soil, and earthworm casts. Each treatment was conducted in triplicate for physico-chemical properties and in quintuplicate for microbial indicators. Each pot (inner size: 13.5 cm in diameter; 14 cm in height) was filled with 980 g of air-dried and sieved (2 mm) soil mixed with 20 g of cow manure. The soil-manure mixture was incubated for 14 d at 60 % WHC. Subsequently, five earthworms were introduced into each pot to ensure sufficient drilosphere soil formation for subsequent analyses. These earthworms had been individually rinsed with deionized water and depurated for 24 h in darkness to clear their gastrointestinal tracts (Deng et al., 2021). After 30 d of incubation, earthworms were manually separated and placed in dark containers for 24 h to collect fresh gastrointestinal contents as casts. The remaining soil was carefully sampled to collect approximately 200 g of bulk soil (unaffected by earthworm activity, primarily from upper soil layers) and 20 g of drilosphere soil (from the 2 mm thick layer surrounding visible burrows, excluding any areas with visible cast material). The 2 mm thick layer of drilosphere soil was collected by carefully scraping the inner walls of visible earthworm burrows using a small spatula. The sampling process involved: (1) identifying clearly visible burrow openings and channels within the soil matrix; (2) using a thin spatula to systematically

scrape soil material from the burrow wall surface to a consistent depth of 2 mm; (3) collecting the scraped material in clean containers while avoiding contamination with bulk soil or cast material; and (4) pooling material from multiple burrows within each pot to obtain the required sample mass of approximately 20 g. This careful separation ensured that drilosphere samples represented soil primarily influenced by earthworm epidermal mucus secretion and physical bioturbation rather than gut-processed material (Ni et al., 2025). Bulk and drilosphere soil samples were air-dried and sieved through a 2 mm mesh for physico-chemical analysis, while casts were freeze-dried and stored at -80 °C. For microbial analysis, 3 g soil samples were collected from each treatment and immediately stored at -80 °C to preserve DNA integrity until further analysis.

Chemical imaging was conducted in earthworm boxes or 2D terraria (Evans, 1947) with the two soils, one terrarium for each soil. Soil pre-process was identical to pot experiment. Custom terraria (15 cm × 20 cm × 3 cm) were designed as Fig. S2a and the front panel can be open to attach imaging membrane on the soil profile. In the pre-cultivation period, the terraria were filled with 300 g of soil at 60 % WHC, which were shielded with aluminum foil to block light simulating soil environment. Following 7 days, two earthworms (~1 g each, approximately 5 cm long) were introduced into each terrarium for a 30-d incubation at 25 °C in a dark environment. Throughout cultivation, the terraria were
shielded with aluminum foil that had several holes pre-punched at the top to block light, prevent the earthworms from escaping, and provide sufficient air for survival. At the end of the incubation period, terraria were carefully opened and a soil profile with conserved burrows was chosen for further imaging. The spatial distributions of acid phosphatase activity and labile P flux, which reflect acidic/neutral soil available P indicators (Soltangheisi et al., 2019), were assessed using soil zymography and DGT tools, respectively. Soil zymography was conducted following Ma et al. (2019), using a phosphatase substrate containing 4-Methylumbelliferyl phosphate (4-MUF-P) at pH 6.5. A nylon filter (5 × 5 cm², 0.45 µm pore size, Taoyuan, China) was immersed in 10 mM 4-MUF-P for 15 min, then attached to the soil profile containing one conserved earthworm burrow for 1 h in darkness. The membrane was then removed, gently cleaned to remove any attached soil particles, and imaged using a camera (EOS 200DII, Canon, Japan) under 355 nm UV light, offering a spatial resolution of approximately 100 µm (Fig. S2b). A calibration curve was established using various MUF concentrations (0.05–10 mM) to standardize the grayscale values obtained from the zymograms in relation to the enzyme activities (Fig. S3a). The DGT with colorimetric imaging densitometry (CID) analysis followed the procedure described by Li et al. (2024). A precipitated zirconia binding gel (5 × 5 cm²) was attached to the same sampling zone as the nylon filter for available P imaging after 24 h incubation in the dark. After rinsing, the binding gel was immersed in a molybdenum blue developing solution for 10 min. After color development, any excess solution was removed using deionized water. Image analysis was conducted using a flat-bed scanner (Laserjet Pro MFP M429dw, Hewlett-Packard, USA) at a resolution of 1200 dpi, corresponding to a vertical and lateral image resolution of ~21.6 µm (Fig. S2c). A calibration curve was established with various KH₂PO₄ concentrations (0–2000 µg L⁻¹) to standardize the identified grayscale values corresponding to the mass of absorbed P in the binding gel (Fig. S3b). The images of P were built using labile P flux values (µg cm⁻² h⁻¹), representing P availability.

## 2.3. Chemical analysis

The analyzed samples included bulk soil, drilosphere soil, and earthworm casts. Soil available P was extracted using 0.5 M NaHCO₃ (P-Olsen method) with a solid-to-liquid ratio of 1:20 (w/v), shaking for 30 min at 180 rpm, followed by centrifugation at 4000 rpm for 10 min. The supernatant was filtered through 0.45-µm cellulose acetate membrane filters and quantified using the molybdenum-blue colorimetric method at 880 nm wavelength (Olsen et al., 1954). For total P, Fe, and Al measurements, 0.02 g samples were acid-digested with 15 mL of 50 % HNO₃ and 2 mL of 30 % H₂O₂ at 105 °C following USEPA Method 3050B. After cooling, the digestate was diluted to 50 mL with deionized water, filtered through 0.45-µm cellulose acetate membrane filters, and analyzed by inductively coupled plasma optical emission spectrometry (ICP-OES, iCAP Pro X, Thermo Fisher Scientific, USA) in 20 mL solution diluted with deionized water through a 0.45-µm cellulose acetate membrane filter. Sample pH was measured at a soil-to-solution ratio of 1:10 (w/v) using 0.01 M CaCl₂ after 30 min equilibration. Total C (TC) and total organic C (TOC) content was analyzed using a TOC analyzer (MULTI N/C 3100, Germany), while sample texture was assessed using a laser particle size analyzer (ZEN3690/MPT-2, Malvern, UK). Dissolved organic C (DOC) concentration was detected using a TOC analyzer with deionized water extraction at a soil-to-solution ratio of 1:10 (w/v), shaking for 2 h at 180 rpm, followed by centrifugation and filtration (Vos et al., 2019). Cation exchange capacity was measured according to a standardized procedure (HJ889–2017). Oxalate-extractable P, Fe, and Al (P<sub>ox</sub>, Fe<sub>ox</sub>, and Al<sub>ox</sub>) were extracted using 0.2 M acid ammonium oxalate (pH 3.0) at a soil-to-solution ratio of 1:10 (w/v), shaking in darkness for 4 h at 180 rpm, followed by centrifugation at 4000 rpm for 10 min and filtration through 0.45-µm cellulose acetate membrane filters before ICP-OES analysis. To assess the extent of P loading on the Fe/Al oxides, the degree of P saturation (DPS) was calculated using Eq. (1)

(van der Zee and van Riemsdijk, 1988). Extracellular acid phosphatase (ACP) and alkaline phosphatase (ALP) activities were assessed using commercial test kits (BC0140 and BC0280, Solarbio, Beijing, China), respectively, following manufacturer protocols.

$$DPS = \frac{P_{ox}}{0.5 \times (Fe_{ox} + Al_{ox})} \quad (1)$$

## 2.4. Microbial DNA extraction and sequencing

Microbial community analysis was performed on bulk soil and drilosphere soil samples. Total soil DNA was extracted from 0.5 g of soil samples using the SPINeasyTM DNA Pro Kit for Soil following manufacturer's instructions. Extracted DNA quality was evaluated by 1 % agarose gel electrophoresis to verify fragment integrity, and concentration was quantified using a NanoDrop Spectrophotometer (Thermo Scientific, USA). PCR amplification targeted bacterial 16S V4 regions using primers 515F (5'-GTGCCAGCMGCCGCGGTAA-3') and 806R (5'-GGACTACHVGGGTWTCTAAT-3') (Chen et al., 2023), while fungal ITS2 regions were amplified using primers ITS3-F (5'-GCATCGATGAA-GAACGCAGC-3') and ITS4-R (5'-TCCTCCGCTTATTGATATGC-3') (Zuo et al., 2018). PCR products were purified and target DNA fragments recovered for library construction using the NEBNext® Ultra™ II DNA Library Prep Kit for Illumina®, which included end repair, adapter ligation, and library amplification. The final library underwent paired-end 250 bp (PE250) sequencing on the Illumina Nova 6000 platform at Guangdong Magigene Biotechnology Co., Ltd. (Guangzhou, China). This high-throughput sequencing approach enabled comprehensive analysis of microbial community composition and diversity in the soil samples.

Raw paired-end reads underwent sliding-window quality trimming using fastp (v.0.14.1) with parameters -W4 and -M20. Primers were subsequently removed using cutadapt (v.1.14), resulting in quality-controlled clean reads. Paired reads were merged using search-fastq_mergepairs (v.10.0.240), followed by filtering of non-mergable or low-quality reads to generate raw merged sequences (Raw Tags). For bacterial sequences, operational taxonomic units (OTUs) were clustered at 97 % sequence similarity using the UPARSE algorithm. Taxonomic annotation was performed using the RDP classifier against the Silva database (version 138). Fungal sequences underwent identical OTU clustering procedures, with taxonomic annotation performed against the UNITE database (version 8.0). Alpha diversity indices (Shannon) and beta diversity metrics (Bray-Curtis dissimilarity) were calculated to assess community structure and composition differences between bulk soil and drilosphere samples.

## 2.5. Statistical analysis and image processing

Quality control and statistical analysis were performed to ensure the reliability of the experimental results. A soil reference material (GBW(E) 070332 ASA-11) was used to assess the quality of available P measurements, yielding a recovery rate of 96.5 % ± 6.9 %. All data are presented as mean ± standard deviation (SD). Enzyme activities from conventional assays and phosphatase activity from zymography, as well as labile P flux from DGT, were calculated using different approaches, with detailed equations provided in the Supplementary Data (Eqs. (S1)–(S3)), reflecting the difference between mass-based (traditional chemical assays) and area-based (zymography and DGT) analytical methods. Statistical analyses were carried out using SPSS 20.0 software. Before performing analysis of variance (ANOVA), data homoscedasticity was evaluated using a Bartlett test, and the normality of residuals was visually inspected. ANOVA was then conducted, followed by Tukey's test based on the LSD method for multiple comparisons, with significance determined at *p* < 0.05.

Two representative 2D observations with apparent index variation gradients were selected for each imaging technique. Image analysis of
DGT-CID and soil zymography measurements was performed using ImageJ software. The process involved three main steps. I) Analyte quantification: color images were converted to 8-bit grayscale, and pseudo-color was applied using the “Lookup Tables” function to visualize gradients in labile P flux and acid phosphatase activity levels. This color mapping step is crucial as it transforms subtle grayscale intensity differences into a more perceptible color spectrum, making spatial patterns easier to detect visually. Calibration curves based on MUF and KH₂PO₄ gradient concentrations were established using the “calibrate” function. The concentration scale was then incorporated into the image using the “calibration bar” function. II) Spatial distribution analysis: to examine analyte distribution across burrows, a line perpendicular to the burrow direction was drawn using the “line” tool. The “Plot Profile” function was then used for locational quantification. III) Hotspot identification: the “measure” function was used to obtain maxima, minima, and mean values for the entire image. Hotspots were defined as areas with content exceeding the mean plus twice the standard deviation (Mean + 2SD), as described by Bilyera et al. (2020). While other methods have been proposed for analyzing soil physicochemical hotspots (e.g., Top-25 % of values, > 50 % of mean values, > 20 % of mean values), the Mean + 2SD method was found to be effective for hotspot identification in zymography and potentially in DGT-P (Bilyera et al., 2020). The hotspot area proportion was calculated using the “Histogram” function as the ratio of hotspot area to total image area.

## 3. Results

### 3.1. Physicochemical properties of two soils

The physicochemical properties of the two soils are presented in Table 1. Fluvisol contained lower total P (0.6 g kg⁻¹) and available P (9.8 mg kg⁻¹) compared to Acrisol (1.2 g kg⁻¹ and 30.3 mg kg⁻¹, respectively). For the purposes of this study, we refer to Fluvisol as the lower-P soil and Acrisol as the higher-P soil to distinguish their relative P availability levels. Fluvisol had higher cation exchange capacity and higher total Fe and Al contents than Acrisol, but lower reactive Fe and Al contents. TC and TOC contents in Fluvisol (10.8 g kg⁻¹ and 7.3 g kg⁻¹) were lower than those in Acrisol (16.8 g kg⁻¹ and 15.9 g kg⁻¹). Fluvisol was acidic (pH 5.7) with a silt-clay texture, while Acrisol was neutral (pH 7.1) with a clay texture.

### 3.2. Earthworm-induced changes in phosphorus pools and phosphatase activities

Chemical analysis of soil samples and earthworm casts revealed distinct differences in P content and phosphatase activity among bulk soil, drilosphere soil, and earthworm casts (Fig. 2). Statistical analysis confirmed significant differences between these treatments for most measured parameters (Table S1).

Earthworm activities increased P levels in the drilosphere (Fig. 2a and b). Total P content in drilosphere soil increased by 28.8 % compared to bulk soil in Fluvisol, while changes were negligible in Acrisol. Notably, earthworm casts contained the highest total P content,

<figure><img src="image_3.png" /><figcaption>**Fig. 2.** Total (a) and available P contents (b), acid (c) and alkaline phosphatase activities (d) in bulk soil, drilosphere and earthworm casts of a fallow soil (Fluvisol) and a farmland soil (Acrisol). BU, DR, and CA refer to bulk soil, drilosphere soil, and earthworm casts, respectively. Data represent the mean ± standard deviation of the three replicates. Letters denote significant differences at *p* < 0.05.</figcaption></figure>
exceeding levels in bulk soil by factors of 120 % in Fluvisol and 7.7 % in Acrisol (Fig. 2a). Available P followed a similar pattern, with earthworm casts showing the highest available P content, being 8.5 times higher in Fluvisol and 4.4 times higher in Acrisol compared to respective bulk soils. Drilosphere soil contained intermediate available P levels in both soils (Fig. 2b).

Phosphatase activity exhibited soil-specific patterns in the drilosphere (Fig. 2c and d). In Fluvisol, both acid phosphatase and alkaline phosphatase activities increased progressively from bulk soil to drilosphere soil to earthworm casts. In contrast, Acrisol showed different trends: acid phosphatase activity remained relatively stable between bulk soil and drilosphere soil, while alkaline phosphatase activity decreased in drilosphere soil compared to bulk soil (Fig. 2c and d). The highest alkaline phosphatase activity in Acrisol was observed in earthworm casts (Fig. 2d).

Factors influencing inorganic P desorption showed soil-specific patterns between Acrisol and Fluvisol, including variations in reactive Fe/Al oxides content, pH, and DOC content (Fig. S4). Statistical analysis confirmed significant differences between bulk soil and drilosphere soil for most measured parameters (Table S1). Oxalate-extractable P (P<sub>ox</sub>) showed minimal variation between drilosphere soil and bulk soil in both soils (Fig. S4a). However, oxalate-extractable Fe (Fe<sub>ox</sub>) and Al (Al<sub>ox</sub>) contents increased in drilosphere soil of Acrisol (Fig. S4b and c), resulting in a lower DPS in drilosphere soil (Fig. S4d). Soil pH increased in the drilosphere compared to bulk soil in Fluvisol, while it decreased in Acrisol (Fig. S4e). DOC content was highest in earthworm casts but showed significant increases in drilosphere soil compared to bulk soil in both soils, though the magnitude was more modest than in casts (Fig. S4f, Table S1). This pattern reflects the transient nature of DOC elevation in drilosphere soil, where initial epidermal mucus-derived organic matter undergoes rapid microbial processing after earthworm passage. These combined factors suggest that inorganic P desorption

processes were more pronounced in Acrisol than in Fluvisol.

### 3.3. Spatial distribution of labile phosphorus and phosphatase activities in the drilosphere

The drilosphere functioned as a locus for P accumulation and enzyme activity expression, as evidenced by *in situ* chemical imaging (Fig. 3). Chemical imaging analyses of the area surrounding earthworm biopores aligned with the presence of earthworm casts and undisturbed areas (Fig. 3a and f).

Visualization of P distribution using DGT-CID (Fig. 3b and g) revealed elevated labile P content within several millimeters surrounding the burrow walls. In Fluvisol, labile P hotspots were recorded at 5.1 and 6.8 µg cm⁻² h⁻¹ on both sides of the burrow (Fig. 3c). In Acrisol, labile P hotspots were found at 2.3 and 2.2 µg cm⁻² h⁻¹ on both sides of the burrow wall (Fig. 3h). Quantitative analysis of drilosphere hotspots showed that labile P levels (maximum and mean) in Fluvisol were approximately twice those in Acrisol, while the minimum level in Fluvisol was only one-third that of Acrisol (Table S2). Hotspot proportions for labile P flux were 3.1 % and 0.1 % in Fluvisol and Acrisol, respectively.

Soil zymography identified localized areas (hotspots) of increased acid phosphatase activity adjacent to the earthworm biopores (Fig. 3d and i), with more pronounced effects in Acrisol. Higher enzyme activity was detected in the soil surrounding the burrow walls compared to the interior of the walls. In Fluvisol, two hotspots were found at 35.6 and 36.2 µmol cm⁻² h⁻¹ on both sides of the burrow, with another at 36.4 µmol cm⁻² h⁻¹ within the burrow (Fig. 3e). In Acrisol, hotspots were measured at 35.0 and 36.0 µmol cm⁻² h⁻¹ on both sides of the burrow, and at 36.6 µmol cm⁻² h⁻¹ within the burrow (Fig. 3j). Based on quantitative analysis, the mean values of acid phosphatase activities were 7.5 times higher in Acrisol than in Fluvisol, primarily due to

<figure><img src="image_4.png" /><figcaption>**Fig. 3.** Identification and quantitation of labile P and acid phosphatase hotspots in the two drilospheres. Mapping of the original soil profile (a, f), distributions (b, d for Fluvisol and g, i for Acrisol) of labile P flux and acid phosphatase activities in the drilosphere, respectively. In particular, the identified areas of labile P (c, h) and acid phosphatase (e, j) are represented as dashed lines (A'A' and C'C' for labile P hotspots; B'B' and D'D' for acid phosphatase hotspots). The blue circles refer to feature points on the dash lines. The labile P flux distribution was captured by DGT-CID, and the acid phosphatase activity distribution was captured by soil zymography. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>
differences in minimum values (19.0 µmol cm⁻² h⁻¹ in Acrisol versus 0.5 µmol cm⁻² h⁻¹ in Fluvisol). Hotspot proportions for acid phosphatase activity were 5.4 % and 7.5 % in Fluvisol and Acrisol, respectively (Table S2). The soil zymography data correlated well with available P obtained by the DGT-CID method, emphasizing the co-localization and potential causal relationship between acid phosphatase activity and labile P flux in the drilosphere (Table S3).

### 3.4. Shifts in microbial community composition associated with earthworm activity

The microbial community in the drilosphere exhibited distinct characteristics compared to bulk soil (Figs. 4 and S5). Earthworm bioturbation decreased microbial abundance and the fungi-to-bacteria ratio (Fig. S5a and b), especially in Fluvisol. Earthworms reduced bacterial community abundance while the fungal community remained relatively stable (Fig. 4a and b). Bacterial β-diversity analysis revealed that the drilosphere harbored a distinct microbiome compared to bulk soil, though differences were less pronounced for the fungal community. Additionally, these microbial community changes were less profound in Acrisol compared to Fluvisol (Fig. 4c and d).

Sphingomonas and Ohtaekwangia dominated the bacterial community in bulk soil; earthworm bioturbation increased the abundance of Aeromonas and Flavobacterium (Fig. 4e). In the fungal community, Scedosporium and Penicillium were dominant in bulk soil of Fluvisol, while Scedosporium and Podospora were abundant in the drilosphere (Fig. 4f). Among the top five bacteria in terms of abundance, only Ohtaekwangia (R² = 0.32, *p* < 0.05) and Aeromonas (R² = 0.53, *p* < 0.05) showed positive correlations with total P. Bacillus generally showed positive correlations with soil physicochemical indicators but a negative correlation with total P (Fig. 4e and g). Among the top five fungi in terms of abundance, Podospora and Humicola correlated positively with most soil physicochemical properties but exhibited negative correlations with total P (Fig. 4f and h).

## 4. Discussion

### 4.1. Soil-specific mechanisms of earthworm-mediated phosphorus mobilization

The effects of earthworm-mediated P mobilization differed between the two soils (Fig. 2 and Table 1). Available P increased in the drilosphere of both soils compared to bulk soil, while total P increased only in Fluvisol (Fig. 2a and b). The elevated available P and total P content in the drilosphere resulted from earthworm feeding selectivity, burrowing behavior, and epidermal mucus composition (Liu et al., 2020). Earthworms preferentially inhabited soil with high organic matter content, which provided more available resources for earthworm assimilation, thereby increasing soil available P content through residual digestion (Vos et al., 2022a). In soil with long-term fertilizer application, nutrients and organic matter were enriched and predominantly existed in forms available to plants and soil organisms, potentially explaining the negligible changes observed under earthworm bioturbation (Fonte et al., 2010). Mineral-associated Fe/Al oxides captured inorganic and organic P through bonding interactions with -OH groups, a process indirectly regulated by dissolved organic matter (Yan et al., 2023). The moderate but significant increase in dissolved organic matter content in the drilosphere (Fig. S4, Table S1) reflects the ephemeral nature of earthworm-induced organic matter inputs, where initial epidermal mucus-derived DOC is rapidly processed by soil microorganisms following earthworm passage, distinguishing it from the more persistent accumulation observed in casts.

The observed differences in phosphatase activities between the two soils reflect distinct mechanisms of P mobilization in the drilosphere. Higher acid phosphatase and alkaline phosphatase activities were exhibited in the drilosphere compared to bulk soil for Fluvisol, while

negligible variation in acid phosphatase activity and lower alkaline phosphatase activity were presented in the drilosphere compared to bulk soil for Acrisol (Fig. 2c and d). These elevated phosphatase activities in Fluvisol accelerated organic P mineralization through enzymes released by microorganisms affected by earthworm bioturbation. Earthworm epidermal mucus secreted during burrowing provided carbon-rich substrate for indigenous microbes, which stimulated P-solubilizing microbes activities to organic P assimilation (Hou et al., 2024). This process involved diverse microbial species interactions regulated by earthworm activity, which contributed to P mobilization in the drilosphere and constructed specific soil microbial food webs that achieved cross-trophic nutrient turnover (Fig. 4) (Wan et al., 2025). The ultimate product of these processes was reflected in earthworm casts, which contained the highest total and available P, potentially attributed to gut-excreted phosphatases and gut microbial interactions with intestinal mucus (Satchell and Martin, 1984) (Fig. 2). However, available P content variations have been reported among different earthworm species (Vos et al., 2019), suggesting species-specific effects on P cycling. While the mechanisms of P mobilization mediated by earthworms remain partly unresolved, novel imaging techniques are needed to identify and quantify P distribution patterns in the drilosphere.

### 4.2. Drilosphere as a functional hotspot for phosphorus cycling and enzyme activities

Labile P and acid phosphatase hotspots were identified using DGT and soil zymography techniques, which were combined for the first time to explore P spatial distribution in the drilosphere and to infer potential mechanisms of P mobilization based on correlative patterns and literature support (Figs. 1, 3, and 5). In soil science, gel-extractable P is commonly used to assess plant-available P, reflecting the desorption capacity of orthophosphate from mineral surfaces (Menezes-Blackburn et al., 2016). We employed a similar methodology utilizing DGT-P, defined as labile P with high correlation to Olsen-P (Six et al., 2014), to characterize the spatial distribution of available P around earthworm burrows. Based on DGT imaging, a layer approximately 0.5 cm thick within the burrow wall contained higher labile P content than outside areas, providing almost twice the value corresponding to the quantification from imaging (Fig. 3c and h).

The correlation between metal-bound adsorption sites and available P distribution became evident through our spatial analysis. Reactive mineral-associated metals functioned as carriers of phosphate in acidic soil, affecting the adsorption/desorption processes of orthophosphate (Gérard, 2016). Under earthworm burrowing and metabolism, higher reactive Fe/Al oxide content and DOC content were observed in the drilosphere compared to bulk soil (Fig. S4). Higher reactive Fe/Al oxide content provided more adsorption sites, but simultaneously, the competitive binding of dissolved organic matter to these sites facilitated P desorption. This pattern is consistent with DOC competitively binding to adsorption sites on the surface of reactive metal oxides, which may facilitate inorganic P desorption from mineral surfaces (Vos et al., 2022b). This dynamic equilibrium between adsorption and desorption processes, mediated by earthworm-derived organic inputs, created the spatial heterogeneity observed in our DGT images.

Similar to DGT imaging, soil zymography revealed higher acid phosphatase activities surrounding the burrow wall corresponding to labile P content (Fig. 3e and j). The spatial co-localization of phosphatase activity and available P demonstrates the interconnected nature of these processes. Phosphatase enzymes released by enriched microbial communities not only mineralized organic P but also contributed to the local pH and organic acid environment that influenced P desorption from metal oxides, creating a positive feedback loop where enhanced enzyme activity promoted further P mobilization through both direct mineralization and indirect geochemical effects.

The diffusion mechanisms underlying P redistribution in the drilosphere involve both physical and chemical gradients created by
<figure><img src="image_5.png" /><figcaption>**Fig. 4.** Bacterial and fungal community variation in the bulk and drilosphere samples of two soils. (a, b) Alpha-diversity analysis of bacterial and fungal communities. (c, d) Principal component analysis of the bacterial and fungal communities. (e, f) Community composition analysis of bacterial and fungal species. (g, h) Correlation analysis between the dominant bacterial and fungal species and environmental factors. BU, bulk soil; DR, drilosphere soil. TP, total P; DOC, dissolved organic carbon; $Al_{ox}$, oxalate-extractable aluminum; DPS, degree of P saturation; $Fe_{ox}$, oxalate-extractable iron; $P_{ox}$, oxalate-extractable phosphorus; ALP, alkaline phosphatase activity; ACP, acid phosphatase activity; AP, available P.</figcaption></figure>
<figure><img src="image_6.png" /><figcaption>**Fig. 5.** Conceptual framework of earthworm-mediated P mobilization and P hotspot distribution in the drilosphere. The left panel shows the relative changes in available P (AP), acid phosphatase activity (ACP), pH, alkaline phosphatase activity (ALP), bound inorganic P (P<sub>ox</sub>), reactive iron oxides (Fe<sub>ox</sub>), reactive aluminum oxides (Al<sub>ox</sub>), dissolved organic carbon (DOC), and total P (TP) across bulk soil, drilosphere and cast. The center panel depicts the key components of the drilosphere: burrow and epidermal mucus. The lower left panel illustrates the mechanistic processes where microbial pH alteration and DOC competitive adsorption facilitate inorganic P desorption from mineral-associated particle surfaces, while phosphatase excreted by microbes contributes to organic P mineralization. The right panel visualizes P hotspot distribution as detected by DGT (DGT-P) and soil zymography (zymography-ACP), revealing spatial patterns of labile P and phosphatase activity within the few millimeters surrounding the earthworm burrow, which drives P dynamics in the drilosphere microzone under earthworm bioturbation.</figcaption></figure>

earthworm activity. The porous structure of burrow walls facilitates diffusive transport of dissolved P from high-concentration zones near epidermal mucus deposits to surrounding soil matrix. Simultaneously, the steep chemical gradients in pH, DOC, and enzyme activity create driving forces for P mobilization that extend several millimeters from the burrow interface. This combination of enhanced diffusion pathways and chemical driving forces explains the measurable P hotspots observed at distances up to 0.5 cm from burrow walls.

Phosphatase hotspots mediated by earthworm activity were influenced by microbial species and activity (Vos et al., 2014). Earthworm bioturbation increased the abundance of dominant species while decreasing overall species abundance and deviation (Fig. 4). Notably, the abundance of *Aeromonas* and *Flavobacterium* in the bacterial community increased under earthworm bioturbation, with the latter species being critical components of the earthworm gut community (Hu et al., 2018). *Aeromonas* may release orthophosphate through participation in soil organic mineralization, regulated by soil total P content (Tikhonov et al., 2016). *Flavobacterium* has the potential to solubilize P along with other mineral elements (such as potassium and silicon) (Hu et al., 2018; Meena et al., 2015). *Scedosporium* and *Podospora* were abundant in the fungal community of the drilosphere. The effect of earthworms on fungal community composition was minimal, likely due to the high availability of nutrients for fungi (Burke et al., 2011). Since high abundance of *Scedosporium* was found in vermicompost but not in the earthworm gut, this species can be stimulated by earthworm secretions (Martín-Gil et al., 2008). *Podospora*, which may originate from manure, can also be stimulated by earthworms to accelerate soil P turnover (Ding et al., 2017).

Additionally, interactions between microbial communities and soil properties influenced soil P mobilization (Fig. 4g and h). Available P content increased with the abundance of *Bacillus*, a species recognized as critical in microbial fertilizers that boost soil fertility and plant nutrition (Liu et al., 2022) (Fig. 4g). *Bacillus* can i) excrete organic acids to accelerate desorption of bound P from reactive Fe/Al oxides (Hu and Boyer, 1996), and ii) release acid and alkaline phosphatases to increase soil organic P mineralization (Wang et al., 2023). The inorganic soluble P mobilized by *Bacillus* would be further transferred to microbial

biomass (Fig. 4g). DOC increased with the abundance of *Nitrospira*, potentially influencing soil C:N:P stoichiometry and facilitating bound P desorption from reactive metal oxides through competitive binding of DOC at adsorption sites (Fig. 4g) (Aanderud et al., 2018; Vos et al., 2022b). The abundance of *Preussia* increased with total P and decreased with pH, indicating that this genus could be regulated by soil physico-chemical properties (Fig. 4h) and could serve as an indicator of healthy soil. In addition, *Podospora* and *Humicola* may facilitate P mobilization through secretion of organic acids and phosphatase (Kaul et al., 2019). Thus, bacteria appeared to mine soil P primarily through desorption of inorganic labile P rather than through organic P mineralization. In contrast, P mobilization by fungi was mainly affected by soil physico-chemical properties, likely relying on soil food webs comprising diverse species. These integrated processes are conceptualized in Fig. 5, which synthesizes the relationships between microbial communities, soil properties, and P cycling mechanisms.

### 4.3. Applications and limitations of imaging techniques for studying biogeochemical processes in soil microhabitats

The mechanisms of earthworm-mediated P mobilization revealed in this study have important implications for global P cycling and sustainable agriculture (Helfenstein et al., 2024; Vidal et al., 2023). With growing concerns about P scarcity and the need to reduce dependence on finite phosphate rock reserves, understanding natural biological processes that boost P availability becomes increasingly critical (Desmidt et al., 2015). Earthworms are present in agricultural soils worldwide, and our findings suggest that management practices that support earthworm populations could contribute to more efficient P cycling and reduce fertilizer requirements. The spatial heterogeneity we observed indicates that earthworm-mediated P hotspots could serve as localized nutrient sources, potentially reducing the need for uniform P fertilization across entire fields. This biological approach to P management aligns with principles of regenerative agriculture and could help address both food security and environmental sustainability challenges (Elser et al., 2024).

Our study demonstrates the value of combining *in situ* imaging
techniques with microbial community analysis to understand P cycling in the drilosphere. The application of DGT and zymography revealed distinct spatial patterns of P availability and phosphatase activity surrounding earthworm burrows, providing insights that conventional bulk soil sampling cannot capture. These findings show that earthworm mucus and casts create microscale hotspots with elevated P concentrations, while specific microbial taxa (*Aeromonas*, *Flavobacterium*, *Scedosporium*, and *Podospora*) become enriched in these zones. The spatial co-localization of P availability and enzyme activity suggests coordinated biogeochemical processes that drive nutrient cycling in this soil micro-habitat (Fig. 5). From an engineering perspective, our methodology provides a framework for monitoring earthworm bioturbation effects on soil properties and could inform sustainable land management practices by identifying biological hotspots of nutrient activity (Ahkami et al., 2024).

The imaging approaches developed in this study have broader applications across soil microhabitats. Similar to our findings in the drilosphere, DGT and zymography have revealed P hotspots in the rhizosphere where root exudates drive P mobilization through pH changes and phosphatase activity (Bilyera et al., 2022; Ma et al., 2021). The methodological framework we established, combining spatial imaging with microbial community analysis, could be applied to understand P cycling in other biologically active soil zones. Our demonstration that earthworm-mediated P mobilization involves both inorganic P desorption and organic P mineralization provides insights into soil organic matter-driven P activation mechanisms that may serve as key bridges linking carbon and P dynamics across different soil microhabitats, as illustrated in the conceptual framework (Fig. 5).

Despite the valuable insights provided by this study, several limitations should be acknowledged. The 2D terraria may not fully represent natural soil conditions, potentially constraining earthworm behavior and P redistribution patterns. The short-term nature of our experiment (30 d) may not capture long-term effects of earthworm-mediated P cycling that occur over seasons or years. Phosphatase activity assays were conducted under standardized pH conditions that may differ from actual *in situ* soil pH, affecting the interpretation of enzymatic activities in natural soil environments. While our microbial community analysis identified taxa potentially associated with P mobilization, direct causal links between specific microorganisms and P transformation processes cannot be definitively established without additional functional analyses. The spatial resolution of our imaging techniques, while advanced, still provides an averaged measurement over small areas and may not detect finer-scale heterogeneity in P distribution and enzyme activities that could be biologically significant.

Looking forward, several research directions could address these limitations and build upon our findings: (1) optimization of imaging protocols through systematic evaluation of terrarium dimensions and soil properties to better represent natural conditions; (2) development of imaging approaches to track transformations of diverse P species, including organic P and sparingly-soluble inorganic P, which would provide more comprehensive understanding of P cycling mechanisms we observed; (3) integration of advanced analytical techniques such as synchrotron-based spectroscopy and nuclear magnetic resonance spectroscopy to provide molecular-level insights into the biochemical processes underlying our spatial patterns; (4) exploring approaches to link laboratory-based hotspot characterization with field-scale applications by identifying mappable proxy indicators, addressing the scale gap between our microscale observations and ecosystem-level P management; and (5) integrating machine learning approaches with our multidimensional spatial datasets to develop automated detection algorithms that could enable real-time monitoring of biogeochemical processes and inform precision management strategies for nutrient cycling across agricultural and natural ecosystems.

## 5. Conclusions

Our comprehensive study, integrating *ex situ* chemical analysis, *in situ* chemical imaging, and microbial analysis, characterized the heterogeneous distribution and activation mechanisms of P mobilization in the drilosphere. Our findings support both of our initial hypotheses. First, we confirmed that the drilosphere functions as a distinct P and microbial hotspot with quantifiable spatial patterns, where available P and phosphatase activities exceeded those in bulk soil and were successfully detected using *in situ* imaging techniques. Second, we demonstrated that P mobilization mechanisms in the drilosphere differed between soils with contrasting P availability, with stronger available P increases in the lower-P soil (Fluvisol) due to elevated phosphatase activities, increased reactive mineral metal oxides content, and greater abundance of P-solubilizing microorganisms (*Aeromonas*, *Flavobacterium*, *Scedosporium*, and *Podospora*).

Specifically, we found that earthworm activity increased soil available P content, with earthworm excretions (epidermal mucus and casts) elevating soil total P levels and facilitating organic P mineralization through boosted phosphatase activity. Earthworm activities also intensified the desorption of inorganic P from Fe/Al oxides through increased dissolved organic matter content and altered pH conditions. *In situ* chemical imaging revealed the crucial role of soil microhabitats in shaping soil P distribution, with the drilosphere emerging as a key zone for P and phosphatase turnover characterized by distinct hotspot patterns. While earthworm activity decreased overall bacterial and fungal community biodiversity, it selectively enriched specific microorganisms involved in P mobilization. These findings advance our understanding of earthworm-driven P mobilization processes and offer valuable insights into the biogeochemical cycling and heterogeneous distribution of P mediated by soil fauna.

### CRediT authorship contribution statement

**Zhen-Yu Qiang:** Writing – original draft, Visualization, Software, Investigation, Formal analysis, Data curation. **Dong-Xing Guan:** Writing – review & editing, Supervision, Methodology, Funding acquisition, Data curation, Conceptualization. **Jia-Lu Gao:** Investigation. **Gang Li:** Writing – review & editing, Resources. **Daniel Menezes-Blackburn:** Writing – review & editing. **Anna Gunina:** Writing – review & editing. **Yvan Capowiez:** Writing – review & editing. **Lena Q. Ma:** Writing – review & editing, Resources, Funding acquisition.

### Declaration of competing interest

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

### Acknowledgments

This work was funded by the National Natural Science Foundation of China (42177194 and 42330703) and Yangtze Delta Region Healthy Agriculture Institute (HAI-YFWW-202403-01). We would like to thank Weizhen Hu from the Agricultural Experiment Station at Zhejiang University for the support in utilizing the greenhouse. We would also like to thank the handling editor and three anonymous reviewers for their detailed, constructive feedback throughout the review process, which substantially improved our manuscript.

## Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.geoderma.2025.117497.
# Data availability

Data will be made available on request.

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