Mechanisms of Biochar in Modulating Soil Organic Selenium Transformation and Enhancing Soil Selenium Availability

Abstract

Selenium deficiency poses a significant threat to human health. The low bioavailability of selenium in soil largely limits the improvement of selenium content in crops. Selenium in soil mainly exists in an organically bound form. Biochar has the ability to regulate the organic matter content of soil; however, the impact of biochar on the transformation of organically bound selenium in soil remains poorly understood. Therefore, this study investigates the effect of biochar on organically bound selenium in typical medium–to–high selenium soils from Yimen County, Yuxi City, Yunnan Province. Reed straw (RS), apple wood (AW), and walnut shells (WS) were used as biomass materials for biochar preparation. The study utilized organically bound selenium transformation incubation and pot experiments to explore the role of biochar in transforming organically bound selenium in soil. The results showed that organically bound selenium was the dominant selenium form in the soil, accounting for 66.31% of the total selenium content. Both pot experiments and incubation trials indicated that the addition of biochar significantly increased the levels of water–soluble and exchangeable selenium in the soil. The addition of biochar mainly promotes the conversion of fulvic acid–bound selenium into water–soluble and exchangeable selenium. In the absence of carbon sources, humic acid–bound selenium can also be converted to water–soluble and exchangeable selenium. Correlation analysis revealed that soil water–soluble selenium was significantly negatively correlated with soil total selenium (r = −0.792 **, p < 0.01), soil phosphatase activity (r = −0.645 *, p < 0.05), abundance taxa of Chloroflexi (r = −0.751 *, p < 0.05), Chytridiomycota (r = −0.674 *, p < 0.05), and Basidiomycota (r = 0.722 **, p < 0.05), while it was significantly positively correlated with soil urease activity (r = 0.809 **, p < 0.01), and significantly negatively correlated with abundance taxa of Myxococcota (r = −0.800 **, p < 0.01). Compared with the initial soil, the WS treatment (initial soil water–soluble selenium 0.31 μg·kg⁻¹, exchangeable selenium 0.11 μg·kg⁻¹) significantly increased the soil water–soluble selenium by 34.9 times and exchangeable selenium by 100.2 times. Additionally, the selenium content in garlic increased by 170% compared to the control group.

1. Introduction

Selenium (Se) is a vital nutrient essential to human biology [1,2], primarily absorbed through food [3,4]. Among dietary sources, plant–derived selenium is crucial due to its high absorption efficiency [5]. The selenium content in plants is influenced by soil selenium levels and the soil’s physicochemical properties [6]. Selenium exists in various forms in the soil, and these forms significantly affect selenium’s bioavailability [7,8]. Thus, studying selenium transformation in soil and regulating its bioavailability is essential.

Soil selenium bioavailability varies greatly with soil properties and composition. Processes like adsorption, desorption, precipitation, dissolution, biological oxidation, and reduction continuously affect soil selenium forms [9]. These processes are influenced by soil pH, redox potential (Eh), clay content, soil organic matter (OM), and microbial activity, with soil pH and organic matter being primary factors [10]. Besides a small portion of free water–soluble selenium, most selenium compounds bind to various soil components. Selenium forms are generally categorized into soluble selenium (SOL–Se), exchangeable selenium (EX–Se), iron–manganese oxide bound selenium (FMO–Se), organic bound selenium (OM–Se), and residual selenium (RES–Se) based on water solubility and binding strength [11]. Soluble and exchangeable selenium, considered bioavailable, can be absorbed by plants but generally constitutes less than 5% of total soil selenium, while organic bound, iron–manganese oxide bound, and residual selenium are more prevalent [12].

Selenium mainly exists in residual and organic bound forms [13]. The residual form is mineral–locked and hard to release, while organic–bound selenium is a significant available source. Soil organic matter is a crucial factor in selenium distribution in selenium–rich soils [14]. Biochar can enhance soil properties, improve water retention, reduce nutrient leaching, boost microbial activity, and increase organic carbon content, promoting plant growth [15]. Biochar can have both positive and negative priming effects on soil organic matter mineralization. It can increase soil pH, enhance microbial growth, and promote organic selenium mineralization, thus increasing water–soluble or exchangeable selenium [16]. Conversely, biochar can adsorb humic acids, reduce microbial contact, and lower the mineralization of organic bound selenium, negatively affecting selenium availability [17,18]. Understanding how biochar alters the balance between different soil selenium forms to regulate selenium bioavailability remains complex.

Exploring biochar’s impact on soil organic matter mineralization offers new research directions for regulating soil organic bound selenium. This study aims to improve the bioavailability of organic bound selenium in Yunnan bauxite by adding biochar. Using typical plateau soil from Yimen County, Yuxi City, Yunnan Province, this research examines the effects of biochar made from reed straw (RS–BC), apple wood (AW–BC), and walnut shell (WS–BC) on soil organic bound selenium. It also investigates different biochar types effects on various selenium forms, contents, and soil properties. Meanwhile, garlic has the ability to accumulate and metabolize selenium and has a strong enrichment effect on selenium. Selenium is transported from the root and accumulated in the leaves, which can be metabolized into selenium amino acids with important biological functions. The absorption, accumulation, distribution, and metabolism of selenium in garlic plants depend on the form of selenium provided by soil. A pot experiment is conducted to explore the effects of different types of biochar on selenium content in garlic. Clarifying the effect of biochar on the transformation behavior of selenium in soil and its mechanism will provide a theoretical basis for the application of biochar in regulating soil selenium availability.

2. Materials and Methods

2.1. Soil Sample Collection and Materials Pretreatment

2.1.1. Soil Sample Collection

The soil sample was collected in Yimen County, Yunnan Province, China (N: 24°71′–24°73′; E: 101°98′–102°05′), according to the geological survey reports [19], and the soil type is Latosol. The 0–20 cm soil was air–dried and sieved with a sieve size of 0.150 mm. The soil was sealed in polyethylene ziplock bags and stored at room temperature for the next step of physical and chemical property determination and cultivation experiments.

2.1.2. Biochar Pretreatment

Biochar was pyrolyzed using apple wood, reed straw, and walnut shell. Specifically, the raw materials were ground with a Chinese herbal medicine grinder and sieved with a sieve size of 0.150 mm. The milled biomass powder was weighed and placed in a stainless–steel distiller in a M10L–1200 °C small laboratory muffle furnace (Sigma, Shanghai, China). The heating rate was set to 25 °C·min⁻¹, and then the muffle furnace preheated to the target maximum heating temperature of 500 °C. The distiller was placed in the furnace and purged with nitrogen at a flow rate of 4 L·min⁻¹, and pyrolyzed at a specified pyrolysis temperature for 4 h. The distiller was removed from the muffle furnace and blown with nitrogen. Additionally, it was cooled to room temperature and taken out for drying and weighing. The quality of biochar was weighed to quantify the yield, and then the sample was ground and sieved with a sieve size of 0.150 mm for later use. The biochar made by apple wood, reed straw, and walnut shell was labeled as AW, RS, and WS.

2.1.3. Soil Fulvic Acid Se and Humic Acid Se Pretreatment

The extraction of soil organic–bound selenium (including humic acid selenium and fulvic acid selenium) was used in Tessier methods [14]. The specific extraction methods are in the Supplementary Materials. According to step (III) in the Supplementary Materials, after heating and digestion, the precipitate was determined to be humic acid selenium (HA–Se), and the supernatant was determined to be fulvic acid selenium (FA–Se). The extract of step (III) in Supplementary Materials. was freeze–dried to obtain fulvic acid Se and humic acid Se powder.

2.2. Experiment Design and Sample Analysis

2.2.1. Pot Experiment

The test site was the intelligent greenhouse of Kunming University of Science and Technology. The greenhouse conditions were 25 °C·14 h⁻¹ light conditions, relative humidity of 21.6%, and light intensity of 5682 μmol·(m²·s)⁻¹. This experiment was conducted in a greenhouse with a pot experiment from December 2022 to April 2023. Each pot was filled with 0.7 kg of soil (through a 0.150 mm sieve). Different types of biochar and fertilizers were weighed and manually added to the soil and then mixed evenly; the water–soluble amendments were first dissolved in partially deionized water and then added to the soil to achieve a more uniform effect. Three biochar (including AW–BC, WS–BC, and RS–BC), which were 3% biochar of soil, biochar, and soil, were manually added to the pot according to the biochar/soil mass percentage of 3% (that is, 21 g biochar were manually added to each pot of soil). 0.61 g urea per kg soil, 0.37 g potassium dihydrogen phosphate per kg soil, and 0.50 g potassium chloride per kg soil), and a control (including 0.61 g urea per kg soil, 0.37 g potassium dihydrogen phosphate per kg soil, and 0.50 g potassium chloride per kg soil) was designed. Each treatment was set up with 12 replicates. The pH of basic soil was 6.18, total nitrogen (TN) was 1.8 ± 0.2 g·kg⁻¹, total phosphorus (TP) was 0.7 ± 0.1 g·kg⁻¹, total potassium (TK) was 37 ± 2 g·kg⁻¹, available nitrogen (AN) was 106 ± 7 mg·kg⁻¹, available phosphorus (AP) was 53 ± 2 mg·kg⁻¹, and available potassium (AK) was 175 ± 13 mg·kg⁻¹. Additionally, the total selenium of 267 ± 15 μg·kg⁻¹, of which water–soluble selenium (SOL–Se), exchangeable selenium (EX–Se), fulvic acid selenium (FA–Se), humic acid selenium (HA–Se), iron–manganese oxide–bound selenium (FMO–Se), and residual selenium (RES–Se) were 10.3 ± 0.6 μg·kg⁻¹, 11 ± 1 μg·kg⁻¹, 128 ± 11 μg·kg⁻¹, 49.3 ± 0.6 μg·kg⁻¹, 25 ± 1 μg·kg⁻¹, and 45 ± 6 μg·kg⁻¹, respectively.

The soil and plant samples were collected in the bulb expansion stage of garlic growth (April 2023). The soil was divided into two parts: one part was air–dried for determining soil N, P, K, and Se; the rest part was freeze–dried for determining soil enzyme activity and soil microorganisms. The plant was divided into roots, stems, leaves, and bulbs to analyze the content of selenium in each part.

2.2.2. Organic Bound Selenium Transformation Culture Experiment

The 0.5 g of biochar were manually added to a 15 mL centrifuge tube with 0.1 g of fulvic acid Se or humic acid Se and 2 mL of soil microbial suspension. The treatments are shown in

Table 1. The organic bound selenium transformation culture experiment.

Treatments	Biochar	Soil Microbial Suspensions	Fulvic Acid Se	Huminc Acid Se
ControlFA †	–	–	0.1 g	–
FA–b	–	2 mL	0.1 g	–
RS–Fb	0.5 g RS	2 mL	0.1 g	–
AW–Fb	0.5 g AW	2 mL	0.1 g	–
WS–Fb	0.5 gWS	2 mL	0.1 g	–
ControlHA †	–	–	–	0.1 g
HA–b	–	2 mL	–	0.1 g
RS–Hb	0.5 g RS	2 mL	–	0.1 g
AW–Hb	0.5 g AW	2 mL	–	0.1 g
WS–Hb	0.5 gWS	2 mL	–	0.1 g

^† In Control_FA and Control_HA, the soil microbial suspension was replaced with 2 mL sterile UP water.

. Samples were collected on the 1st, 5th, 10th, 15th, 30th, and 45th days of culture. Each treatment had 18 replicates. The soil microbial suspensions were extracted with the following methods. Fresh soil was added into UP water with a ratio of 1:5 (g·mL⁻¹), as well as an appropriate amount of quartz sand. The mixed liquid was filtered with sterile filter paper before shaking at 200 r·min⁻¹ at 28 °C for 30 min. The filtrate was considered as soil microbial suspensions.

2.3. Sample Analysis

2.3.1. Determination of Soil and Plant Samples

The soil pH, total nitrogen (TN), total phosphorus (TP), total potassium (TK), available nitrogen (AN), available phosphorus (AP), available potassium (AK), soil organic matter (OM), and soil enzyme activity were determined according to the “Methods of soil analysis, part 3: Chemical methods” [20]. The extraction of soil selenium (including SOL–Se, EX–Se, FA–Se, HA–Se, FMO–Se, and RES–Se) was used in Tessier methods and determined by atomic fluorescence spectrometry [14]. The Se content of the plant was determined according to Ohki [21].

2.3.2. DNA Extraction and Illumina MiSeq Sequencing

Total soil genomic DNA was extracted from the soil samples following the manufacturer’s protocol using the EZNA Soil DNA Kit (Omega, Norcross, GA, USA). The concentration and quality of the genomic DNA samples were tested using a Nanodrop 2000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). Primer pairs, ITS1F(5′–CTTG GTCATTTAGAGGAAGTAA–3′) and ITS2R(5′–GCTGCGTTCTTCATCGATGC–3′), for the fungal ITS1 region and 338F(5′–ACTCCTACGGGAGGCAGCAG–3′) and 806 R(5′–GGACTACHVGGGTWTCTAAT–3′) for the bacterial V3–V4 region, were used to amplify soil fungal and bacterial DNA samples, respectively. PCR was performed according to the methods of a previous study [22]. The prepared samples were detected with MiSeq by Majorbio BioTech Co., Ltd. (Shanghai, China).

Raw sequence data were obtained using Mothur v.1.48.2, applying the standard protocol. Briefly, reads were paired and screened as follows: for fungi, bacteria, reads longer than 280/500 bp or shorter than 230/200 bp were discarded. Sequences with ambiguous bases or homopolymers longer than 13 bp were removed. Fungal sequences were clustered by abundance–based greedy clustering (method = agc). The obtained sequences were compared with the sequence of the Silva v.138 reference file database (https://www.mothur.org/wiki/Silva_reference_files, accessed on 10 December 2024) to obtain taxonomic information [23].

2.4. Statistical Analysis

Venn diagrams were constructed in the apps of origin 2025 to demonstrate the common and unique OTUs of soil bacteria and fungi in response to the different biochar. The Pearson’s correlation analyses between relative abundance of microbial taxa and environmental parameters were conducted with bivariate correlation analysis using SPSS 22.0. Redundancy analysis (RDA) between Se and selected environmental factors or bacteria/fungi with Canoco 5.0 to reveal the relative importance of microorganisms that govern Se form. Mean differences among treatments were determined using one–way ANOVA with Duncan’s test at p < 0.05 in SPSS 22.0.

3. Results and Discussion

3.1. Effects of Biochar on Soil Physical and Chemical Properties

The total nitrogen (N) in soil amended with biochar was higher than that in the control (Figure 1a). This was consistent with other research results [24,25]. On one hand, biochar’s porous structure could adsorb and retain nitrogen, reducing its loss. Studies have shown that the porosity of biochar is significantly positively correlated with soil–available nitrogen, and long–term use leads to a decrease in total soil nitrogen [24,25]. In this study, the BET surface of AW–BC and RS–BC treatments was much lower than those of WS–BC (Table S2), resulting in significantly higher soil total nitrogen contents in AW–BC and RS–BC than those in WS–BC and treatments (p < 0.05); on the other hand, nitrogen–rich biochar (derived from nitrogen–rich feedstocks) can statistically significantly enhance soil nitrogen content (p < 0.05), thereby improving soil fertility. This was supported by the statistically significantly higher nitrogen content in RS–BC and AW–BC treated soils than those in control and WS–BC, due to the high N content of RS–BC and AW–BC (p < 0.05, SI: Table S1).

The total phosphorus (P) in soil amended with RS–BC was significantly higher than the other biochar treatments and control (Figure 1b). Studies have shown that soil P availability is strongly influenced by soil pH through its regulation of phosphate interactions with metal ions [26]. In acidic soils, low pH promotes the dominance of Al³⁺ and Fe³⁺/Fe²⁺, which immobilize P via precipitation as insoluble Al/Fe phosphates (e.g., AlPO₄, FePO₄). Conversely, in alkaline soils, high pH enhances Ca²⁺ activity, leading to P fixation as Ca–phosphates (e.g., Ca₃(PO₄)₂). Biochar regulates soil pH, thereby altering the strength of these interactions. For instance, RS–BC (pH = 6.78) applied to acidic soils may elevate soil pH toward neutrality, reducing Al³⁺/Fe³⁺ solubility and releasing fixed P, while simultaneously introducing Ca²⁺ from its alkali metal oxides (e.g., CaO), which could form sparingly soluble Ca–P compounds. However, the higher total P content in RS–BC–treated soil compared to AW (pH = 9.20) and WS (pH = 7.23) suggests that pH–driven metal–P interactions alone cannot fully explain the observed trends. Biochar contains alkali metal oxides (e.g., Ca²⁺, Mg²⁺, K⁺) that hydrolyze in water to release soluble salts and OH⁻, particularly in acidic soils. AW–BC, with the highest pH (9.20), significantly increased soil pH compared to WS and RS at the same application rate (Figure 1d). Paradoxically, despite its stronger alkalinity, AW–BC did not enhance total P retention as effectively as RS–BC. This implies that while pH modulation influences P dynamics (e.g., shifting Al/Fe–P to Ca–P equilibria), the intrinsic P content and composition of biochar play a dominant role. For example, if RS–BC inherently contains higher levels of organic P or stable Ca/Mg–P complexes resistant to fixation, it could directly enrich soil total P regardless of pH effects. Therefore, although soil pH regulates P–metal interactions, the total P accumulation in biochar–amended soils likely results from a combination of pH–mediated solubilization and the biochar’s native P content. This dual mechanism warrants further investigation, particularly focusing on biochar–specific P speciation and its long–term stability in contrasting soil pH environments. With respect to soil total potassium content, all three biochar treatments caused a reduction, but no significant difference was observed among those treatments (Figure 1c). With the exception of WS–BC, the soil EC in RS–BC and AW–BC was higher than in control. This might be relative to the ash of biochar. The ash of AW–BC (9.45%) was higher than RS–BC (7.21%) and WS–BC (5.32%). Overall, RS–BC demonstrated the highest efficacy in enhancing soil nutrients and preventing nutrient loss.

3.2. Effect of Biochar on Soil Selenium Form

The content of various forms of Se in soil is shown in Figure 2. Before the study began, the initial soil total Se content was 267.1 μg·kg⁻¹, of which organic matter bound Se, including fulvic acid bound Se (47.96% of total soil Se) and humic acid bound Se (18.35% of total soil Se), accounted for 66.31% of total soil Se. This was consistent with the other studies [27,28,29]. The plant–available Se, including water–soluble Se and exchangeable Se content, accounted for 3.86% and 3.78% of total Se in the soil, respectively.

Compared with the initial soil, the total Se content in the AW–BC, RS–BC, WS–BC, and control soils was significantly reduced. This was because the Se in the soil was absorbed by the garlic. With the addition of biochar, significant changes were observed in the Se composition in the soil. Specifically, fulvic acid–bound Se of initial soil decreased significantly compared to control, AW–BC, RS–BC, and WS–BC treated soil, with control decreasing by 59.5%, AW–BC by 83.4%, RS–BC by 75.5%, and WS–BC by 64.0%. However, the SOL–Se levels in the soil for the control, AW–BC, RS–BC, and WS–BC treatments increased by factors of 4.2, 36.7, 33.9, and 34.9, respectively, compared to the initial soil. Similarly, the EX–Se content in the soil increased by factors of 12.1, 104.3, 110.5, and 100.2 for the control, AW–BC, RS–BC, and WS–BC treatments, respectively, relative to the initial soil. Furthermore, the HA–Se content in the soil increased by factors of 1.6, 1.7, 1.2, and 1.2 in the control, AW–BC, RS–BC, and WS–BC treatments, respectively, compared to the initial soil. This suggests that biochar addition facilitated the transformation of these selenium forms, highlighting its potential to influence selenium speciation and bioavailability in the soil positively [27].

3.3. Effect of Biochar on the Transformation of Organic Bound Selenium

In order to further understand the effect of biochar on the effectiveness of organically bound selenium, we conducted a biochar cultivation experiment. Figure 3 illustrates that in the Control_FA and Control_HA treatments, water–soluble selenium and exchangeable selenium showed no significant changes during the incubation period. However, in the presence of microorganisms, both soluble selenium and exchangeable selenium increased with time. Exchangeable selenium peaked at 15 days, while water–soluble selenium increased rapidly in the first 15 days and then slowly from 15 to 45 days. Fulvic acid–bound selenium and humic acid–bound selenium significantly decreased, indicating that soil microorganisms are the main factor for organic selenium transformation.

Additionally, throughout the incubation period, the content of dissolved selenium in biochar treatments was significantly higher than in FA–b or HA–b treatments, suggesting that biochar has a “positive priming effect” on the decomposition of fulvic and humic acids when they are the sole carbon sources. Studies show that biochar promotes the mineralization of soil organic carbon and biochar itself, accelerating the loss of soil organic carbon and causing a decrease in soil organic carbon content. The strength of the priming effect produced by biochar is related to the type of biochar material. For example, Zimmerman et al. found that biochar made from herbaceous plants exhibited a more significant positive priming effect on soil organic carbon mineralization compared to oak wood biochar. In this study, WS biochar exhibited a greater priming effect on fulvic acid–bound and humic acid–bound selenium than AW and RS biochar. The C/N ratio of the WS treatment was 102.6, which was much higher than that of the AW and RS treatments. Under the same conditions, soil microorganisms preferentially used organic matter that was easily degradable and highly available. Therefore, the addition of biochar, which was difficult to degrade, promoted the degradation of fulvic acid selenium and humic acid selenium in the system [30]. The order of the increase of SOL–Se content under the three biochar treatments was WS > AW > RS, among which the effect of adding walnut shell biochar was the best. After 45 days of culture, the content of SOL–Se converted from FA–Se in the samples treated with WS biochar was 1.63 times that of FA–b and 4.15 times that of Control_FA.

The content of SOL–Se transformed by HA–Se increased by 107.7% compared with the control. It shows that the addition of biochar is beneficial to the activity of microorganisms, thus promoting the conversion of organic bound selenium to water–soluble selenium with high plant availability and improving the effectiveness of selenium. In the process of conversion of organic bound selenium to EX–Se. with the extension of culture time, the exchangeable selenium content in the samples treated with three biochar increased first and then decreased, while the control group without biochar showed an upward trend, which may be due to the partial conversion of EX–Se to SOL–Se in the later stage of culture. With the transformation of FA–Se, the content of EX–Se increased gradually in the first 10 days and began to decrease slowly after 15 days of culture. Compared with the control, in general, the order of the increase of EX–Se content in the three biochar treatments was WS–Fb > AW–Fb > RS–Fb. Under the action of microorganisms, the addition of biochar treatment increased the conversion of HA–Se to EX–Se to a certain extent compared with CK.

3.4. Effect of Biochar on Soil Enzyme Activity

Soil enzyme activity is a crucial indicator for evaluating soil quality, as it reflects the intensity of various biochemical reactions occurring in the soil [31]. Soil invertase activity is indicative of the accumulation and transformation of organic matter, and it serves as an important measure of soil fertility and microbial activity [32]. Figure 4 shows biochar addition significantly increased soil invertase activity and soil urease activity (Figure 4a,b). The RS––BC treatment (3.027 mg·(g·24 h)⁻¹) exhibited the highest invertase activity, followed by WS––BC (2.010 mg·(g·24 h)⁻¹) and AW––BC (1.652 mg·(g·24 h)⁻¹). Invertase activity in the RS––BC treatment was 2.1, 1.8, and 1.5 times higher than that in the control, AW–BC, and WS–BC treatments, respectively. Urease was involved in the hydrolysis of urea into ammonium nitrogen, a form that is readily absorbed by plants, thus promoting nitrogen transformation in the soil [32]. The variation in urease activity between treatments may result from differences in biochar types and application rates, which influence soil physical and chemical properties and thus the activity of urease [31]. In this study, the C/N ratio of RS biochar (41:1) was much lower than that of AW–BC (57:1) and WS–BC (218:1) (Table S1). This resulted in higher microbial abundance and activity in the RS–treated soil and significantly higher soil sucrase activity compared to the other treatments.

In contrast to invertase, urease activity was highest in the AW–BC treatment (1.304 mg·(g·24 h)⁻¹), significantly surpassing the RS–BC (1.117 mg·(g·24 h)⁻¹), WS–BC (0.989 mg·(g·24 h)⁻¹), and control (0.909 mg·(g·24 h)⁻¹) treatments. The response of neutral phosphatase activity to different biochar treatments was less consistent. The AW–BC treatment had the most significant effect on increasing soil neutral phosphatase activity, whereas RS–BC and WS–BC treatments reduced phosphatase activity. This suggested that phosphatase activity in soil was influenced by a range of environmental factors. Biochar increased phosphatase activity on urease and phosphatase by increasing pH [33]. The pH of AW–BC was significantly higher than RS–BC and WS–BC, leading to the significantly higher urease and phosphatase activity. The catalase activity of three biochar did not show a significant difference with each other, as well as with control.

3.5. Effects of Biochar on Bacterial Community Structure in Soil

The Operational Taxonomic Units (OTU) data for rhizosphere bacteria and fungi under different treatments is shown in Figure 5. A total of 4631 bacterial OTUs and 1400 fungal OTUs were detected across all treatments. Compared to the control, the number of bacterial and fungal OTUs significantly decreased under biochar treatment, with the OTU distribution in the three biochar treatments ranking as RS > AW > WS (Figure 5). There were 1844 common bacterial species across all treatments, with the control having 1351 unique bacteria, WS having 830, AW having 993, and RS having 910 unique bacteria (Figure 5a). Similarly, there were 717 common fungal species, with the control having 156 unique fungi, WS having 155, AW having 128, and RS having 175 unique fungi (Figure 5b). The dominant bacterial phyla in all treatments included Proteobacteria, Actinobacteriota, Chloroflexi, Acidobacteriota, Bacteroidota, Gemmatimonadota, and Myxococcota, accounting for over 78.5% of total bacterial OTUs (Figure 6). Ascomycota and Basidiomycota were the dominant fungal phyla, accounting for over 79.4% of total fungal OTUs.

Compared to the control, all biochar treatments significantly reduced the ACE index and Chao1 index, indicating that biochar addition decreased bacterial species richness. Furthermore, the Shannon index for the RS treatment was significantly higher than both the control and other biochar treatments, indicating the greatest species diversity in RS treatment. The WS treatment had the lowest Simpson index, while the AW treatment had the highest. Unlike bacteria, the fungal ACE index, Chao1 index, and Shannon index in the RS treatment did not significantly differ from the control but were significantly higher than those in the AW treatment, suggesting that AW biochar significantly reduces fungal richness and species diversity (

Table 2. Bacteria and fungi diversity index in different biochar treatments †.

	Treatments	ACE Index	Chao 1 Index	Shannon Index	Simpson Index
Bacteria	Control	5435 ± 43 a	5010 ± 53 a	6.69 ± 0.10 b	0.0087 ± 0.0004 b
	RS	4550 ± 23 b	4289 ± 84 b	6.72 ± 0.14 a	0.0086 ± 0.0003 b
	AW	4502 ± 38 b	4220 ± 67 b	6.67 ± 0.17 b	0.0091 ± 0.0003 a
	WS	4310 ± 48 c	4045 ± 64 c	6.69 ± 0.12 b	0.0082 ± 0.0003 b
Fungi	Control	1535 ± 36 a	1522 ± 24 a	4.98 ± 0.16 a	0.0199 ± 0.0003 c
	RS	1537 ± 27 a	1538 ± 32 a	4.92 ± 0.14 a	0.0225 ± 0.0003 b
	AW	1452 ± 25 b	1437 ± 37 c	4.67 ± 0.17 b	0.0326 ± 0.0002 a
	WS	1535 ± 37 a	1499 ± 31 b	4.89 ± 0.18 a	0.0210 ± 0.0002 bc

† The different lowercase letters indicate the significant difference at p < 0.05 using ANOVA with Duncan analysis.

).

3.6. Effects of Biochar on the Selenium Uptake of Garlic

As shown in Figure 7, the selenium concentration in garlic roots, stems, and leaves treated with biochar was higher than that in the control, indicating that biochar could enhance the absorption of selenium in garlic. Among the treatments, RS biochar resulted in the highest selenium content in garlic roots, significantly higher than AW, WS, and the control (Figure 7a). For the stems and leaves, the WS biochar treatment had the highest selenium content, followed by AW and RS, with the control group being the lowest. In terms of overall selenium absorption by garlic, biochar treatments greatly increased selenium absorption compared to the control group, with RS, AW, and WS biochar increasing selenium absorption by 73.7%, 126.0%, and 170.4%, respectively (Figure 7b). This aligns with the role of biochar in decomposing fulvic and humic acid–bound selenium.

3.7. The Relationship Between Soil Biological, Physical, and Chemical Properties and Soil Selenium Forms

Correlation analysis (Figure 8) showed that soil water–soluble selenium was significantly negatively correlated with soil total Se (r = 0.792 **, p < 0.01), soil phosphatase activity (r = 0.645 *, p < 0.05), Chloroflexi (r = 0.751 *, p < 0.05), Chytridiomycota (r = 0.674 *, p < 0.05); and Basidiomycota (r = −0.722 **, p < 0.05). Exchangeable Se was significantly positively correlated with soil urease activity (r = 0.809 **, p < 0.01), and significantly negatively correlated with Myxococcota (r = −0.800 **, p < 0.01). To better understand the effect of biochar on Se speciation, we used the redundancy analysis (RDA) between soil physicochemical properties and soil bacterial communities (Figure 9). RDA1 and RDA2 could explain 84.02% of the difference in bacterial structure between all samples affected by detected soil selenium content, soil enzyme activity, and soil physical and chemical properties (Figure 9a), as well as RDA1 and RDA2 could explain 78.4% of the differences in fungal structure in all samples (Figure 9b). The RDA showed that SOL–Se was positively correlated with Chloroflexi, Olpidiomycota, and Chytridiomycota, and negatively correlated with Firmicutes. The EX–Se was positively correlated with Verruomicrobiota, Mortierellomycota, Mucoromycota, Basidiobolomycota, and negatively with Actinobacteriota, Myxococcota, Glomeromycota, Rozellomycota. These patterns suggest a microbial–mediated equilibrium between Se mobilization and immobilization: Chloroflexi and Chytridiomycota likely drive SOL–Se release by decomposing fulvic acid (FA–Se) and humic acid–bound Se (HA–Se) through extracellular oxidases [34] or plant residue degradation [35], respectively. In contrast, Myxococcota and Basidiomycota may promote HA–Se stabilization via humic acid complexation (Myxococcota) [36] or lignin–derived phenolic copolymers [37]. Enzymatic activities further modulate this process—phosphatase enhances SOL–Se liberation by hydrolyzing organic phosphorus–Se bonds [38], while urease facilitates EX–Se adsorption onto humic acid carboxyl/phenolic groups via NH4+–induced cation exchange [39]. Fungal–bacterial synergies also contribute: Mortierellomycota and Mucoromycota physically protect HA–Se within hypha–stabilized microaggregates [40], whereas Actinobacteriota compete for Se via siderophores [41], redirecting it into humic matrices. Collectively, microorganisms regulate FA/HA–Se transformation through decomposition–polymerization dynamics, governed by enzyme activities and functional guild interactions, ultimately determining Se bioavailability and environmental fate in biochar–amended soils [42].

4. Conclusions

The WS–BC treatment can better increase soil pH (increased by 0.65 units), reduce soil electrical conductivity (EC) (reduced by about 19.8%), and increase soil enzyme activity (soil invertase activity increased by 40.8%, soil urease activity increased by 9%). RS–BC and AW–BC treatments reduced the relative abundance of the five dominant bacterial phyla in the rhizosphere soil and increased the relative abundance of the three dominant fungal phyla. The WS–BC treatment significantly increased the conversion of FA–Se and HA–Se to available selenium (SOL–Se and EX–Se) compared to other treatments. In the samples treated with WS–BC, the content of sol se transformed by FA–se increased by 63.3% compared with the control, and the content of sol se transformed by HA–se increased by 107.7% compared with the control. WS–BC treatment significantly increased the selenium accumulation in garlic plants and promoted the transfer of selenium from roots to stems and leaves (compared with the control, it increased by 0.0115 mg·kg⁻¹ and 0.0284 mg·kg⁻¹, respectively), which was beneficial to increase the selenium content of edible parts of plants. In conclusion, the application of biochar has shown positive effects in improving the availability of soil selenium, improving soil properties, regulating microbial communities, and promoting selenium accumulation in plants.