1 Introduction

As an efficient C-neutral tool for sustainable development, biochar is used in a variety of applications, including carbon sequestration, greenhouse mitigation, soil amendments and agricultural practices (Wang et al. 2021). The high porosity, abundant functional groups, and large and hydrophobic external surface of biochar give it great potential value in the remediation of contaminated soils as a superior sorbent (Ahmad et al. 2014). Natural and anthropogenic conversion of solid wastes into biochar has increased its occurrence in the environment, especially when released to soil, which serves as a dominant sink for many persistent organic pollutants (POPs), such as polycyclic aromatic hydrocarbons (PAHs) (Chen et al. 2017). Microbial degradation is a primary method for the complete attenuation and removal of these PAHs from soil (Ren et al. 2018a). The presence of biochar in soil can strongly alter the bioavailability of PAHs to microbes and hence their biodegradation by indigenous soil microorganisms due to the excellent sorption and immobilization of biochar for such PAHs (Ren et al. 2018b). Therefore, a better understanding of the impact of biochar on the biodegradation and mineralization of PAHs in soil is imperative for elucidating the fate and risk of PAHs in soil ecosystems as well as assessing the effectiveness of biochar applications in soil bioremediation.

The effects of biochars on the microbial availability and degradation of PAHs in soils and sediments have been widely studied (Marchal et al. 2013a; Jia and Gan 2014; Kuśmierz et al. 2016; Zhu et al. 2016). In particular, soil amended with 1% sewage sludge-derived biochar caused a decrease in the freely dissolved concentration (Cfree) of ∑16 PAHs in the soil up to 22%, leading to their reduced bioavailability to soil microbes and increased persistence (Tomczyk et al. 2020). However, Shan et al. (2015) reported that biochar amendment in soil at various doses (0.2, 20 and 2000 mg kg−1) did not result in a significant effect on 14C-catechol mineralization even though the addition of these biochars considerably increased 14C-catechol sorption to the soil. Furthermore, Ni et al. (2018) revealed that amendment of soil with 2% corn straw-derived biochar produced at 300 °C facilitated PAH release from the soil matrix, thus increasing the bioavailability of immobilized PAHs and enhancing their biodegradation. Similarly, Bao et al. (2020) found that the addition of 5% biochar to agricultural soil significantly increased the biodegradation of PAHs, especially 4–6 ring high molecular weight PAHs in soil. These divergent observations for PAH biodegradation due to biochar amendment could result from the dissimilar soil properties, exposure dosage and physicochemical properties of biochars, which vary dramatically with raw materials, charring temperature and pyrolysis conditions (Anyika et al. 2015; García-Delgado et al. 2015; Sigmund et al. 2018). This also reflects that the exact mechanisms regarding the effects of the interactions between biochars and PAHs in soil on the microbial availability of PAHs are still poorly understood.

Although the bioavailability of PAHs to microbes plays a crucial role in their microbial degradation and remediation, the abundance and structure of microbial communities providing basic information on degradation capability/potential might also greatly affect biodegradation. Alteration of soil microbial abundance, extracellular enzyme activities, bacterial and fungal community structure and composition as well as intraspecies and interspecies communications between bacteria due to biochar amendment in soil has been documented (Lehmann et al. 2011; Masiello et al. 2013). This effect would further influence microbial-related processes such as carbon sequestration, nutrient biogeochemical cycling and degradation of organic compounds (Liu et al. 2015; Sheng et al. 2016; Zhang et al. 2018b). For instance, Xu et al. (2014) demonstrated that biochar application increased the relative abundance of Flammeovirgaceae, Chitinophagaceae and Hyphomicrobiaceae, which are closely related to carbon and nitrogen cycling, and increased that of Sphingomonadaceae, which is capable of degrading refractory pollutants and aromatic compounds, while it decreased that of certain bacterial families, namely, Acidobacteriaceae, Koribacteraceae, Solibacteraceae, Bradyrhizobiaceae, Gaiellaceae and Rhodospirillaceae. Ahmad et al. (2021) found that the abundance and expression of the nidA gene, which acts as a universal biomarker for PAH-degrading bacteria and is often positively correlated with PAH degradation, increased with increasing biochar dosage in soil. However, the shifts in microbial communities in the complex and real soil environment as affected by biochars coexposed to PAHs, especially for the specific contaminant degraders and functional genes (e.g., nidA gene), were not identified. It is hypothesized that coexisting PAHs and biochars have synergistic or antagonistic effects on soil microbial communities. To date, knowledge regarding the impact of microbial community responses to biochar alone and biochar-contaminant coexposure on the biodegradation of PAHs is very limited, and the associated physiological mechanisms are not fully understood.

In summary, biochar may greatly affect PAH fate in soil via two main aspects: (1) microbial availability of PAHs affected by immobilization of biochar and (2) microbial activity affected by interference of biochar with soil microbial communities. Therefore, the key objectives of this study were to gain mechanistic insight into how biochar affects the biodegradation and mineralization of phenanthrene (as a representative of PAHs) by indigenous soil microorganisms based on these two critical issues. Most previous studies regarding the biochar effects on either biodegradation or soil microbes are constrained to single-time-point measurements at the end of different incubation periods (Mitchell et al. 2016; Zielińska and Oleszczuk 2016; Li et al. 2019). In this work, we examined the impact of wheat straw-derived biochars produced at two charring temperatures (300 and 500 °C) in low and high amendment levels (0.03% and 0.3%) on the microbial mineralization kinetics of phenanthrene with different initial concentrations (2 and 20 mg kg−1) in soil. The bioavailable fraction of phenanthrene, the biomass of bacterial 16S rRNA and PAH-degradative nidA genes as well as the bacterial community structure and diversity in soil with biochar/phenanthrene exposure alone or together were periodically monitored to explore the possible mechanisms involved in the dynamic changes in phenanthrene mineralization. Findings from this study are important for supervising the manipulation of biochar technology in remediation of PAH-contaminated soil.

2 Materials and methods

2.1 Soil, biochar and chemicals

A pristine soil with a composition of 52.6% sand, 24.1% silt and 23.3% clay was collected from the top 20 cm in forestlands located in Mohe, Heilongjiang Province, China. The soil was air-dried and sieved to less than 2 mm to remove stones and roots prior to use. The soil sample had a pH of 7.2 and a cation exchange capacity of 20.94 cmol kg−1. Its organic matter and total organic carbon (TOC) contents were 184.06 g kg−1 and 7.59%, respectively. The content of Σ16 PAHs on the US EPA priority pollutant list in the soil was 1494.2 ± 16.5 μg kg−1, and the soil background concentration of phenanthrene was 179.3 ± 5.6 μg kg−1.

The air-dried wheat straws were cut to approximately 5 mm long and packed into a ceramic crucible with a lid. They were then charred under limited oxygen access in a carbonization furnace at 300 or 500 °C for 3 h. These two temperatures are the most frequently administrated temperatures to produce biochar in both scientific research and practical application (Ok et al. 2018; Zhang et al. 2018b). Furthermore, the formation of large quantities of graphite structures and stable C is initiated at low temperatures of about 300 °C, and a high temperature of 500 °C will continue to cause evident changes in biochar properties so as to compare (Chen and Chen 2009; Lehmann et al. 2011; Zhang et al. 2014, 2015; Chagas et al. 2022). The resulting biochars were mechanically ground to pass through a 60 mesh sieve and stored for use. The detailed characterization methods of the physicochemical properties of the biochars are depicted in the Additional Information, and the results are summarized in Additional file 1: Table S1. The biochars obtained at 300 and 500 °C had similar phenanthrene contents of 169.3 ± 6.8 and 188.1 ± 8.9 μg kg−1, respectively.

A batch equilibration technique with ten initial sorbate concentrations was used to obtain the sorption isotherms of phenanthrene by the tested soil and biochars. The detailed procedures are described in the Additional file 1. The Freundlich isotherms are presented in Additional file 1: Fig. S1, and the fitting parameters are listed in Additional file 1: Table S2.

The 9-14C-labeled phenanthrene with ≥ 98% radiochemical purity and a radioactivity of 50–60 mCi per mmol was provided by Moravek Biochemicals Inc., Brea, CA, USA. Nonlabeled phenanthrene (98% purity) was purchased from Sigma-Aldrich Ltd., Shanghai, China, and its internal standard (d10) was obtained from Accustandard Inc., New Haven, Connecticut, USA. Hydroxypropyl-β-cyclodextrin (β-HPCD, ≥ 98% purity) was supplied by J&K Scientific Ltd. (Beijing, China), and all organic solvents used in this study, including methanol, acetone, n-hexane, dichloromethane and acetonitrile, were of HPLC grade. All other chemicals, such as HCl, NaOH and CaCl2, were of analytical grade.

2.2 Incubation experiments

Briefly, 40 or 400 μL aliquots of a 5000 mg L−1 phenanthrene solution dissolved in acetone were added to a 250 mL glass bottle that contained 100 g of preweighed soil (dry weight) and mixed thoroughly after evaporating the acetone off in a fume hood to yield an initial phenanthrene concentration of 2 or 20 mg kg−1 in the soil. The low pollution level (2 mg kg−1) of phenanthrene occurs in some “hot spots” where this compound is enriched (Tang et al. 2005; Li et al. 2010; Peng et al. 2011). The high phenanthrene pollution level of 20 mg kg−1 was comparable to the concentrations commonly used in the literature (10–50 mg kg−1) and was employed as for the comparison with other relevant studies (Semple et al. 2006; Rhodes et al. 2010; Towell et al. 2011; Oyelami et al. 2015; Zhang et al. 2018a). The soils treated with phenanthrene at low and high concentrations were labeled PLS and PHS, respectively. Then, a specific amount of biochar in dry powder form was added to the bottles and vigorously mixed with the phenanthrene-spiked soil to obtain an application level of 0.03% or 0.3%, which were selected based on our preliminary experiment (data not shown) with the details summarized in Additional file 1. The biochar-amended soil was transferred to a 250 mL glass beaker and rehydrated with 40 mL sterilized deionized water to achieve a soil moisture content of 35%, followed by equilibrating for 12 h and thorough stirring with a stainless-steel spatula for homogenization. The blank control (CK) without phenanthrene and biochar received only sterilized deionized water to reach a similar soil moisture content. The beakers were then sealed with small holes in aluminum foil, allowing ventilation and maintaining soil moisture, and incubated in a climate chamber under darkness at 25 °C with 60% relative humidity. All biochar treatments along with controls were run in triplicate. At 1, 4, 8, 14, 21, 28, 35, 42, 60, 90 and 120 days, 2 g of the soil was sampled for the extraction and determination of total and bioavailable phenanthrene fractions. To evaluate the influence of biochar alone on the biomass of soil microorganisms and soil bacterial community composition and structure, the preparation and incubation of soil amended with biochar but without phenanthrene spiking were the same as described above.

2.3 Total and bioavailable phenanthrene in soil

The total phenanthrene in the soil was extracted with the accelerated solvent extraction (ASE) method. Approximately 1 g of each freeze-dried soil sample was mixed with the ASE preparation diatomaceous earth and extracted with hexane/acetone (v/v = 1:1) using a Dionex ASE 350 Accelerated Solvent Extractor (ThermoFisher Scientific, USA). Detailed extraction conditions were as follows: carrier gas pressure of 0.8 MPa, extractor pressure of 1500 psi (10 MPa), extracting temperature of 100 °C, heating time of 5 min, static extraction time of 5 min, solvent volume of 60% of the extraction cell, nitrogen purging time of 60 s and 2 cycles of extraction. The extracts were concentrated to approximately 1 mL using a rotary vacuum evaporator (RE-52, Qingdao Mingbolm, China) and then purified on a silica gel solid phase extraction column (CNWBOND, Shanghai Anpel, China). The column was subjected to elution with 12 mL of hexane/dichloromethane (v/v = 2:1). The latter eluate was spiked with 200 ng of phenanthrene-d10, condensed to near dryness under slight nitrogen blowing, and then redissolved in 1 mL of methanol as a final volume for further high-performance liquid chromatography (HPLC) analysis.

β-HPCD was used to mildly extract the bioavailable fraction of phenanthrene from the soil (Towell et al. 2011). Briefly, 0.2 g of fresh soil was homogenized with 4 mL of β-HPCD water solution at 50 mM by shaking on an orbital shaker at 200 rpm for 24 h. After mixing, the soil suspension was centrifuged at 4500 rpm for 30 min. The supernatant was transferred to a 15 mL screw cap vial, and 4 mL of n-hexane was added. The vials were vigorously shaken at 150 rpm for 10 min using the same orbital shaker, followed by adding 1 mL of acetone to avoid emulsification. Then, the clear supernatant was condensed to near dryness under a nitrogen stream and reconstituted to 1 mL of methanol for phenanthrene detection by HPLC analysis.

Approximately 10 μL of sample was injected to analyze the phenanthrene concentration in the extract with an HPLC (UltiMate 3000, ThermoFisher Scientific, USA) equipped with an Agilent ZORBAX Eclipse Plus C18 analytical column (5 μm, 4.6 mm × 250 mm) and a UV detector. The mobile phase was composed of acetonitrile and deionized water (80:20, v/v) with a flow rate of 1 mL min−1. The UV wavelength employed for phenanthrene detection was 275 nm. The phenanthrene concentration was quantified by internal standard calibration. The average recovery ratio for phenanthrene in the soil was 93.91% (relative standard deviation (RSD) < 6.98%, n = 7).

2.4 Microbial mineralization kinetics

The mineralization kinetics experiment was carried out in a respirometer following a method developed in our previous study (Zhang et al. 2016). Briefly, an aliquot (5 g) of clean soil was added to a 40 mL amber glass vial and spiked with 20 μL of 14C-labeled phenanthrene solution prepared in methanol at 90 mg L−1, giving a chemical concentration of 0.36 mg kg−1 and a specific activity of 0.111 μCi g−1. Ten or one hundred microliters of 1000 mg L−1 nonlabeled phenanthrene dissolved in acetone was then added to the amber vial, followed by evaporating the solvent off in a fume hood. The vials were placed on a rotary shaker to homogenize the samples at 50 rpm for 72 h, resulting in a spiking concentration of phenanthrene comparable to that in Sect. 2.2. Soil amendments with biochars were conducted in the same way as described in Sect. 2.2. After homogenization, the soil water content was adjusted to 35%. All respirometers were performed in triplicate and incubated under the same conditions as described in Sect. 2.2. During the incubation period, the evolved 14CO2 from the mineralization of 14C-phenanthrene by indigenous microorganisms in soil was captured by 1.5 mL of 1 M NaOH solution in a 2 mL gas chromatography (GC) vial, which was attached to the Teflon-lined screw cap of the 40 mL glass vial. At regular time points (1, 2, 4, 8, 14, 21, 28, 35, 42, 60, 90 and 120 days), respirometers were sampled by replacing the NaOH solution in the GC vial and adding it to 4 mL scintillation cocktail (Ultima Gold, PerkinElmer, USA) to detect the 14C radioactivity using  a Beckman LS6500 liquid scintillation counter (LSC, USA). The initial total 14C-activity in the soil was measured by combusting the soil at 900 °C with an OX-500 biological oxidizer (R.J. Harvey Instrument Corp., USA). Fifteen milliliters of alkaline scintillation cocktail (Oxysolve C-400, Zinsser Analytic, Germany) was applied to absorb 14CO2 released from combustion, and the radioactivity was then quantified by LSC. The remaining 14C-activity in the soil at the end of incubation was also determined via the same procedure, and a mass balance ≥ 98% was detected in all soil treatments. A very low 14C trap by NaOH (less than 30 dpm) was observed during the incubation of sterilized soil amended with the same amounts of 14C-phenanthrene and biochar, which excluded the influence of possible release (e.g., volatilization) of 14C-phenanthrene into the GC vials on the measurement of 14CO2 over the 120 days incubation period.

2.5 Soil microbial biomass C

The chloroform fumigation extraction method was adopted to measure the soil microbial biomass C at the end of 120 days of incubation (Joergensen and Mueller 1996). Ten grams of the nonfumigated soil and fumigated soil by ethanol-free chloroform for 24 h from each sample were subjected to extraction with 40 mL of 0.5 M K2SO4 by vigorous shaking at room temperature for 1 h. The supernatant was filtered through 0.45 μm syringe filters prior to measuring the organic carbon content in the filtrates using a Multi N/C 3100 TOC analyzer (Jena, Germany). The microbial biomass C was calculated by the equation below:

$${\text{Biomass}}\,C = \frac{{E_{C} }}{{0.45}},$$

where Ec (mg C kg−1 soil) is the mass difference in the extracted TOC between fumigated and nonfumigated soils.

2.6 Soil bacterial diversity analysis using high-throughput sequencing

Approximately 1.5 g of moist soil from each replicate sample of various treatments was collected on Days 14, 28 and 60 in the incubation experiment. The extraction of DNA from 0.5 g of soil sample was performed using a FastDNA® Spin Kit for Soil (MP BioMedicals, USA) according to the instructions provided by the manufacturer. The DNA extract was assessed with 1% agarose gel electrophoresis, and its purity was measured using a UV‒vis spectrophotometer (NanoDrop 2000, ThermoFisher Scientific, USA). To analyze the bacterial diversity and community composition, the universal primer pair 515F and 907R was used to amplify the V4–V5 regions of the bacterial 16S rRNA gene. Details for the PCR amplification reaction and its purification following an established method are shown in Additional file 1. 16S rRNA gene-targeted sequencing was performed on an Illumina MiSeq PE300 platform (San Diego, USA) provided by Majorbio Bio-Pharm Technology Co., Ltd. (Shanghai, China). Raw data obtained from high-throughput sequencing were demultiplexed and filtered using fastp (version 0.19.6) to discard low-quality reads (Chen et al. 2018), and then paired-end reads were merged by FLASH (version 1.2.11) (Magoč and Salzberg 2011), with the detailed procedure summarized in Additional file 1. All bacterial sequences were clustered into operational taxonomic units (OTUs) with a 97% similarity cutoff using UPARSE software (version 7.0.1090) (Edgar 2013). Taxonomic annotations were assigned to the representative sequences of OTUs using RDP Classifier (version 2.11) to align against the latest SILVA database of 16S rRNA (version 138) with a threshold of 70% confidence (Wang et al. 2007; Quast et al. 2012).

Alpha diversity expressed as the Shannon index was analyzed by Mothur (version 1.30.2) to measure the evenness and richness of species in the soil bacterial communities. We calculated the Bray‒Curtis distance of various treatment groups relative to the CK with QIIME (version 1.9.1) to describe the dissimilarity in the bacterial community structure among treatments (beta diversity). Principal coordinate analysis (PCoA) based on the weighted UniFrac distance was conducted to display the similarity or differences in the bacterial community structure.

2.7 Quantitative PCR analysis

The copy numbers of the bacterial 16S rRNA gene and key PAH degradation gene (nidA) were measured by real-time quantitative PCR (qPCR) to assess the bacterial abundance. qPCR for the two target genes was conducted following similar protocols with differences in the primer set and standard stock solution. The 16S rRNA gene was amplified using the forward primer Eub338F (ACTCCTACGGGAGGCAGCAG) and reverse primer Eub518R (ATTACCGCGGCTGCTGG). The forward and reverse primers nidAF (TTCCCGAGTACGAGGGATAC) and nidAR (TCACGTTGATGAACGACAAA) were used for amplification of the nidA gene. The recombinant plasmid (pMD18-T) containing an amplified fragment from 16S rRNA or nidA genes was tenfold gradient diluted for the construction of a standard curve and further quantification. The qPCR reactant was 20 μL in total volume, which contained 10 μL of 2× ChamQ SYBR Color qPCR Master Mix (Vazyme Biotech Co., Ltd., Nanjing, China), 0.8 μL of each primer (5 μM), 0.4 μL of 50× ROX Reference Dye, 2 μL of soil DNA template and 6 μL dd H2O. The reaction was performed using an ABI7300 Real-Time PCR system (Applied Biosystems, USA) with the following thermal profile: initial hold at 95 °C for 3 min, followed by 40 cycles of melting at 95 °C for 5 s, annealing at 58 °C for 30 s and extension at 72 °C for 1 min.

2.8 Data analysis

The kinetics of phenanthrene mineralization were fitted with the first-order reaction kinetic model:

$$P = P_{{\max }} (1 - e^{{ - kt}} ),$$

where t (d) is the incubation time, P (%) is the mineralization percentage at time t, Pmax (%) is the maximum mineralization percentage, and k (day−1) is the first-order mineralization rate constant. A formula shown as Pmax/100 × k × [P] was adopted to calculate the maximum mineralization rate (ng g−1 day−1) during the incubation, with [P] representing the initial total concentration of phenanthrene in the experimental system (2000 and 20,000 ng g−1, respectively).

Statistical differences among multiple groups were detected using one-way analysis of variance (ANOVA) or Kruskal‒Wallis analysis. The differences between two groups were compared with Student’s t test or Wilcoxon rank-sum test. Pearson correlation analysis was employed to test significant correlations. All statistical analyses were performed using SPSS 19.0 software with a significance level of 0.05.

3 Results and discussion

3.1 Microbial availability and mineralization of phenanthrene in soil

The mineralization kinetics of phenanthrene at different initial concentrations (2 and 20 mg kg−1) in soil amended with and without biochar by indigenous microorganisms fit well with the first-order equation (R2 = 0.980–0.998) (Fig. 1 and Table 1). The mineralization of high concentrations of phenanthrene in biochar-free soil (Control (PH)) was fast at the beginning of 35 days and stabilized until the end of 120 days of incubation. At approximately 60 days, phenanthrene mineralization in the control reached a maximum equilibrium fraction of 36.76 ± 0.93%. Consistently, the overall extents of microbial mineralization of phenanthrene (50 mg kg−1) in agricultural soil were 36.9–43.1% after 120 days of incubation in the study by Oyelami et al. (2015). Okere et al. (2017) also observed comparable mineralization fractions (39.13–41.44%) of 50 mg kg−1 phenanthrene by indigenous microorganisms in woodland soils from Norway after 30 days of incubation. In comparison, Oyelami and Semple (2015) reported a slightly greater mineralization extent (47.8 ± 0.68%) of 50 mg kg−1 phenanthrene by indigenous microbes in pasture agricultural soil after 14 days of incubation. The mineralization of low-concentration phenanthrene in the control soil (Control (PL)) exhibited a rapid increasing stage in the first 28 days of incubation, followed by a transitional phase after that period to 60 days, and then a very slow increasing stage to the end. After 120 days of incubation, the maximum mineralization percentage of phenanthrene in the control was 17.75 ± 0.39%. Similarly, Shan et al. (2015) observed 18.48 ± 0.85% microbial mineralization of 14C-catechol (0.7 mg kg−1) in agricultural soil after 60 days of incubation. As mineralization is the final process of microbial degradation and is involved in the ultimate transformation of phenanthrene to CO2, the biodegradation percentage of this compound could be larger. At the end of 120 days of incubation, the residue percentages of phenanthrene in the control soil were 20.68 ± 1.12% (Control (PH)) and 31.56 ± 2.55% (Control (PL)), respectively (Additional file 1: Fig. S2a). In contrast, a recent study reported an evidently faster and greater removal of phenanthrene (10 mg kg−1) from petroleum-contaminated soil through natural attenuation with only about 19.6% and 6.2% of phenanthrene remaining in the soil on Days 3 and 6 (Liang et al. 2021). This could be ascribed to the more abundant native degraders stimulated by petroleum hydrocarbons, which are able to decompose the aromatic rings in phenanthrene, compared to those in the forest soil used in the present study. Besides, the phenanthrene molecules in the oil-contaminated soil, which had a much lower TOC content (12.98–39.16 g kg−1) than that of 75.9 g kg−1 in this study, might not be strongly immobilized and could be more available to microbes for degradation.

Fig. 1
figure 1

Microbial mineralization kinetics of phenanthrene in the PLS (a) and PHS (b) by indigenous microorganisms under various biochar treatments during 120 days of incubation. Control (PL) and Control (PH) indicate soil initially spiked with phenanthrene alone at low (2 mg kg−1) and high (20 mg kg−1) concentrations, respectively. BnL and BnH indicate phenanthrene-spiked soil amended with low (0.03%) and high (0.3%) doses of biochar, respectively. “n” indicates charring temperatures of 300 and 500 °C

Full size image
Table 1 Mineralization rates and extents of phenanthrene by soil indigenous microorganisms under various biochar treatments
Full size table

Generally, the wheat straw-derived biochars used in this study inhibited the mineralization/biodegradation of phenanthrene in soil (Fig. 1), except 0.3% biochars amended into the PHS, with the maximum mineralization rate declining by 19.3–78.3%, 120 days mineralized percentage reduced by 2.89–53.6% (Table 1), and the 120-day  residual fraction increased by 9.75–44.5% (Additional file 1: Fig. S2a). Such an evident suppression of biochars on the mineralization/biodegradation of PAHs in soil could be explained by their inhibitory effects on PAH bioavailability, which has widely been reported in previous studies (Qin et al. 2013; Sigmund et al. 2018). In this study, biochar amendment in soil posed a dosage- and sorption-dependent suppression on phenanthrene bioavailability assessed via β-HPCD extraction (Additional file 1: Fig. S2b). The high amendment level (0.3%) of biochar in soil, providing more effective sorption sites and thus resulting in a greater sorption capacity for phenanthrene, more significantly reduced the HPCD extraction efficiency of this compound than the low amendment level (0.03%) (p < 0.05). For a given amendment level, biochar produced at high temperature (500 °C) exhibited a stronger sorption affinity and capacity for phenanthrene due to its higher aromaticity, surface area and porosity as well as bulk and surface hydrophobicity (Additional file 1: Fig. S1 and Tables S1, S2) and resulted in a lower HPCD extraction efficiency of phenanthrene from soil than that at low temperature (300 °C). Overall, phenanthrene bioavailability in the soil significantly decreased after biochar addition in the following order (p < 0.05): Control (PL/PH) > B3L > B5L > B3H > B5H (BnL and BnH indicate phenanthrene-spiked soil with low and high doses of biochar added, respectively, and n indicates charring temperatures of 300 and 500 °C). Hence, biochar amendments generally suppressed phenanthrene mineralization/biodegradation in soil, and the high-temperature biochar exhibited a stronger inhibition than the low-temperature biochar (p < 0.05). In addition, the mineralization rates and extents of phenanthrene in the PLS amended with and without biochar were obviously lower than those in the PHS treatment group, which could also be attributed to the much lower bioavailable fractions of phenanthrene in the PLS group than those in the PHS group (Additional file 1: Fig. S2b and Table 1).

However, a significant increase in the mineralization/biodegradation of phenanthrene after amendment of PHS with a high level (0.3%) of biochar (B3H and B5H) was observed in this study (Fig. 1b and Additional file 1: Fig. S2a). The results were contrary to the study by Omoni et al. (2020) showing that the addition of low doses (0.01%, 0.1% and 0.2%) of biochar increased the mineralization of phenanthrene with a very high initial concentration (240 mg kg−1) in agricultural soil, whereas that of high doses (0.5%) reduced the mineralization. The maximum mineralization rates of phenanthrene in PHS with B3H and B5H amendments were enhanced by 86.7% and 32.4%, respectively. The 120-day mineralization percentages also increased by 44.7% and 32.0%, respectively, and the 120-day residual fractions decreased by 32.8% and 11.7%, respectively. This illustrated that mineralization/biodegradation of phenanthrene in biochar-amended soil was not solely regulated by its sorption-related bioavailability, especially under high concentration contamination, as further supported by the fact that the maximum mineralization rate and total mineralization fraction at 120 days of phenanthrene in the PHS amended with and without biochar were not correlated with its bioavailable fraction (Pearson correlation coefficient r = − 0.779 and − 0.648, significance level p = 0.120 and 0.237, respectively).

3.2 Mineralization association with soil microbial abundance

In addition to the vital role of bioavailability in microbial mineralization/degradation, the activity and function of microbial communities are also important for this process. Biochar exposure alone and/or in combination with phenanthrene may exert positive or negative effects on soil microbial communities. After 120 days of incubation, phenanthrene exposure alone had no significant effect on the soil microbial biomass C (Additional file 1: Fig. S3), indicating that it has little effect on the growth of total microbes in soil. It is also possible that the microbial biomass C was altered during the incubation period but recovered to the initial state within 120 days. Biochar application caused a significantly positive effect on the microbial biomass C in the absence or presence of phenanthrene (p < 0.05, Additional file 1: Fig. S3). Increments in the biomass C of the total microbial community were 22.0–66.8%, 13.3–47.8% and 9.72–57.7% for biochar amendment in the clean soil, PLS and PHS, respectively. Furthermore, the stimulatory effect was more significant at a high level of a certain biochar. The microbial biomass C enhancement by biochar at a given amendment level decreased with the increase of biochar pyrolysis temperature and the associated C content (Additional file 1: Fig. S3 and Table S1), which was consistent with previous studies (Liu et al. 2016). In comparison to high-temperature biochar, biochar pyrolyzed at a low temperature was more conducive to microbial growth, which was primarily because low-temperature biochar provided more nutrients, such as available N and P, for microbes (Additional file 1: Table S3) (Song et al. 2017; Zhang et al. 2018b).

The bacterial 16S rRNA gene copy numbers of the CK without any amendment remained largely stable over the 60 days of incubation period, and it showed a significant decrease over time when exposed to biochar alone (p < 0.05) (Additional file 1: Fig. S4). The abundance of the bacterial gene increased before 28 days of incubation in each treatment in both the PLS and PHS and then decreased significantly from 28 to 60 days of incubation in all treatments (p < 0.05) (Additional file 1: Fig. S4). Exposure to phenanthrene increased the bacterial gene abundance, and such an increase lasted for the whole period (Fig. 2a and Additional file 1: Fig. S4). The effects of biochars on the total bacterial abundance in soil either in the absence or presence of phenanthrene varied with the exposure duration. Compared with the CK, the addition of biochar significantly increased the abundance of the 16S rRNA gene in soil without phenanthrene on Day 14 (p < 0.05) and resulted in similar abundance on  Day 28, while its abundance decreased on Day 60 (Fig. 2a and Additional file 1: Fig. S4). In the phenanthrene-spiked soil, biochar at both low and high amendment levels led to a significant increase in the copy number of the bacterial 16S rRNA gene relative to the control (PL/PH) on Days 14 and 28 (p < 0.05), and this effect was more significant for the low-temperature biochar (Fig. 2a and Additional file 1: Fig. S4), in agreement with the observed results for microbial biomass C. However, the two biochars obtained at 300 and 500 °C at both amendment levels exerted an inhibitory effect on this value on Day 60.

Fig. 2
figure 2

Copy numbers of 16S rRNA (a) and nidA (b) genes in soil under single and combined exposures of phenanthrene and biochar on various incubation days. CK indicates soil without any amendment. PL and PH indicate that containing only phenanthrene at 2 and 20 mg kg−1, respectively. BnL and BnH indicate clean soil amended with low (0.03%) and high (0.3%) doses of biochar, respectively. PLBnL and PLBnH indicate low-concentration phenanthrene-spiked soil amended with low and high doses of biochar, respectively. PHBnL and PHBnH indicate high-concentration phenanthrene-spiked soil amended with low and high doses of biochar, respectively. “n” indicates charring temperatures of 300 and 500 °C. Different letters indicate significant differences at p < 0.05

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The nidA gene encoding the α-subunit of PAH-ring hydroxylating dioxygenase (PAH-RHDα), which initiates the aerobic metabolism of PAHs, can be used as an indicator of PAH degradation activity and potential in soils (DeBruyn et al. 2007; Jurelevicius et al. 2012). Phenanthrene exposure at 2 and 20 mg kg−1 induced significant increases in the catabolic gene biomarker nidA in soil by 98–109 and 116–133 times, respectively (Fig. 2b). Biochar on its own did not result in a significant effect on the relative abundance of the nidA gene, but its coexposure with phenanthrene more significantly increased the nidA gene abundance than phenanthrene exposure alone (Fig. 2b, p < 0.05). This illustrated that the stimulation of the communities carrying the nidA gene induced by the complex of biochar and phenanthrene was much stronger than biochar or phenanthrene alone.

These observations suggested a positive effect of biochars on the biomass of total microbial, bacterial and nidA-carrying communities in phenanthrene-contaminated soil, which may have enhanced the mineralization rate and extent of phenanthrene. Nevertheless, the inhibition of phenanthrene mineralization/biodegradation in soil after biochar amendment was observed in most cases in this study (Fig. 1 and Additional file 1: Fig. S2a). This demonstrated that it was the restricted bioavailability of phenanthrene rather than the promoted microbial activity that generally influenced phenanthrene mineralization/biodegradation in biochar-amended soil. Similar findings have been observed in a modeling experiment by Marchal et al. (2013b).

However, it was further found that bioavailability sometimes contributed less than microbial activities to the mineralization/biodegradation of phenanthrene in biochar-amended soil in this study. A high level of biochar amendment in the PLS resulted in lower mineralization/biodegradation and higher residues of phenanthrene in the soil, and the results were opposite for that in the PHS (Fig. 1 and Additional file 1: Fig. S2a). The difference could be attributed to the different roles played by biochar in soil contaminated with different concentrations of phenanthrene. In the PHS, which likely had a high fraction of unbound phenanthrene in comparison to PLS (Additional file 1: Fig. S2b), the tested biochar played a more critical role in the nutrient stimulation and microbial proliferation for potential PAH degraders over the sorption enhancement and bioavailability inhibition in affecting PAH biodegradation (Song et al. 2017). As a result, biochar at a high amendment level may more strongly facilitate the growth of potential degrading microorganisms and thus phenanthrene degradation than biochar at a low amendment level. In contrast, biochar at a high amendment level added to the PLS was more inclined to cause a stronger sorption and immobilization of phenanthrene in soil, leading to a lower microbial availability and degradation of this compound compared to that at a low amendment level.

For the nidA gene, the spiking of phenanthrene significantly increased its abundance in soil, and the addition of biochar further stimulated nidA gene abundance; such a stimulation was more significant at the high amendment level (Fig. 2b). The abundance of nidA-carrying degraders in the PHS amended with high levels of biochar (PHB3H and PHB5H) was considerably higher than that with low amendment levels of biochar (PHB3L and PHB5L), which were all significantly higher than that in the control soil (PH) (p < 0.05), probably making the influence of the activity of PAH-degrading bacteria on phenanthrene degradation much greater than that of phenanthrene bioavailability. Therefore, the mineralization/biodegradation of phenanthrene in high-level biochar-amended PHS (B3H and B5H) was the highest regardless of its lowest bioavailable fraction, exceeding that in low-level biochar-amended PHS (B3L and B5L) and even surpassing that in the control (PH) (Fig. 1b and Additional file 1: Fig. S2). The phenanthrene mineralization/biodegradation in the PHS amended with low levels of biochar (B3L and B5L) was comparable to or slightly lower than that in the control (PH) despite its much lower bioavailable fraction. In addition, the mineralization fraction of phenanthrene in the PHS with biochar amendment was significantly and positively correlated with the nidA gene abundance during 120 days of incubation (p < 0.05, Fig. 3a), but it was not correlated with the bioavailable fraction of this compound (p = 0.384, 0.538 and 0.279, respectively), confirming that the stimulation of potential nidA-carrying degraders in PHS by biochar should greatly contribute to the biodegradation and removal of phenanthrene.

Fig. 3
figure 3

Correlations between the mineralization fraction of phenanthrene and the nidA gene abundance in PHS (a) or its bioavailable fraction in PLS (b) on different incubation days

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In contrast, although the PLS amended with high levels of biochar (PLB3H and PLB5H) contained the highest abundance of nidA-carrying degraders, followed by that with low amendment levels of biochar (PLB3L and PLB5L) and the control soil (PL) (Fig. 2b), phenanthrene mineralization/biodegradation in high-level biochar-amended PLS (B3H and B5H) was lower than that in low-level biochar-amended PLS (B3L and B5L), which was much lower than that in the control (PL) (Fig. 1a and Additional file 1: Fig. S2a). The mineralization/biodegradation of phenanthrene in the PLS with and without biochar addition followed the order Control (PL) > B3L > B5L > B3H > B5H, which was consistent with the declining order of phenanthrene bioavailable fraction induced by biochar (Fig. 1a and Additional file 1: Fig. S2). This finding consistently indicated that phenanthrene bioavailability could still dominantly control its biodegradation rather than bacterial degradation activity when amending the PLS with biochar, given the very low fraction of bioavailable phenanthrene resulting from biochar amendment. This was further supported by the significantly positive correlations between the phenanthrene mineralization fraction in the PLS amended with and without biochar on different incubation days and the corresponding bioavailable fraction of phenanthrene (p < 0.01, Fig. 3b), while its correlations with the nidA gene abundance were poor (p = 0.268, 0.362 and 0.293, respectively). Overall, the abundance of the nidA gene was promoted by biochar in both the PLS and PHS but was only closely correlated with the phenanthrene mineralization fraction in the PHS.

3.3 Mineralization association with soil bacterial community diversity and structure

The alpha diversity of the soil bacterial community as described by the Shannon index in each treatment generally increased throughout the whole period of incubation (Additional file 1: Fig. S5a), presumably because of the ideal temperature and moisture conditions over the experimental duration. Phenanthrene exposure at both concentrations initially caused a significant decrease in the soil bacterial alpha diversity (p < 0.05), and this decrease was eliminated within 60 days (Additional file 1: Figs. S5a and S6a). It has been generally accepted that contamination by organic pollutants (e.g., PAHs) reduces the soil microbial diversity (Deng et al. 2011). As the biochar was coexposed with phenanthrene, the treatment groups displayed a higher value of diversity indices compared to that with phenanthrene alone (Additional file 1: Figs. S5a and S6a). This illustrated that biochar amendment mediated the reduction in soil bacterial diversity resulting from PAH contamination. Considering no significant change in soil bacterial alpha diversity upon exposure to biochar alone (Additional file 1: Fig. S6a), the positive role of biochar in maintaining the diversity of the bacterial community in contaminated soil could be ascribed to its sorption of toxic compounds, which alleviated the stress caused by contaminants on soil bacteria (Beesley et al. 2011), instead of providing nutrients and habitats for soil bacteria or improving soil physicochemical properties. In addition, more biochar addition (0.3%) further reduced the bioavailability and toxicity of phenanthrene and enhanced the diversity of bacteria in the soil (Additional file 1: Figs. S5a and S6a). Song et al. (2017) also reported that 1% wheat straw-derived biochar amended into PAH-contaminated soil helped to maintain bacterial diversity. However, a greater amount (2%) of biochar did not further enhance the diversity of soil bacteria, which the authors attributed to the changed soil properties (e.g., increased C/N ratio) by the addition of greater amounts of biochar that suppressed the growth of specific bacteria in the soil.

The differences among soil bacterial communities treated with biochar were dependent on the amendment level (Additional file 1: Figs. S5b and S6b). A high level (0.3%) of biochar amendment in the clean soil and PLS resulted in a greater beta diversity (Bray‒Curtis distance) than that at a low amendment level (0.03%), indicating that a greater amount of biochar exerted a stronger effect on the soil bacterial community structure, while this effect for the PLS was time-dependent and diminished within 60 days (Additional file 1: Fig. S5b). Moreover, the difference in the soil bacterial community structure among coexposure to biochar with phenanthrene and phenanthrene exposure alone was greater at the 0.3% amendment level than that at the 0.03% amendment level in a relatively shorter duration, while their effects tended to be similar over time (Additional file 1: Fig. S5b). However, there were no significant differences in the bacterial community structure among the various biochar treatments in the PHS during the entire incubation (Additional file 1: Figs. S5b and S6b).

On the basis of PCoA of all samples on different incubation days (Additional file 1: Fig. S7a), the soil bacterial communities after exposure to phenanthrene clustered separately from those without phenanthrene exposure regardless of the biochar present, indicating that the bacterial community structure showed a dramatic shift upon exposure to phenanthrene, especially at high concentrations of phenanthrene. However, the amendment of biochar into the original soil did not lead to a significant shift in the bacterial community structure, which was reflected by the PCoA results that the bacterial communities after various biochar treatments were not distinctly separated from the CK and were also not distinguishable from each other. Similarly, the bacterial communities under coexposure to biochar-phenanthrene exhibited a close distance from those exposed to phenanthrene alone, suggesting little net effect of biochars on the bacterial community structure. Similar results were also reported in studies conducted with pine wood-derived biochar in clean soil (Tian et al. 2016) and wheat straw-derived biochar in PAH-contaminated soil (Song et al. 2017). Other studies have reported that biochar strongly shifts the soil bacterial community structure (Xu et al. 2014). The discrepant findings may occur, which resulted from the differences in the types and doses of biochar, soil properties, exposure duration and conditions in these studies. The present study revealed that organic pollutants (e.g., phenanthrene) performed more prominently than biochars in affecting the bacterial community structure. Additionally, the incubation time was another main factor contributing to the changes in the soil bacterial community structure, as reflected by the fact that the bacterial communities under a given phenanthrene exposure concentration (0, 2 or 20 mg kg−1) incubated for 14, 28 and 60 days were clustered tightly and separated from each other (Additional file 1: Fig. S7b).

Overall, biochar amendment was helpful for maintaining the alpha diversity and sometimes increasing the beta diversity of the bacterial community in phenanthrene-contaminated soil. A diverse bacterial community structure is beneficial to the degradation of PAHs in soil (Pazos et al. 2003). However, in this study, the similarity and difference in the soil bacterial community structure were largely dependent upon not only the pollutant concentration but also the incubation time. The soil bacterial community structure was not significantly impacted by biochars, which contributed little to phenanthrene mineralization/biodegradation in biochar-amended soil.

3.4 Mineralization association with soil bacterial community composition

To further recognize the specific bacterial taxa related to phenanthrene mineralization/biodegradation at the phylum and genus levels, the relative abundances of the phylotypes in various treatments were recorded (Additional file 1: Fig. S8). The results clearly suggested that the soil bacterial community composition was strongly altered by the addition of phenanthrene and biochar. A total of 31 bacterial phyla were detected in each treated soil after 28 days, and 11 phyla among them had relative abundances greater than 1%, accounting for approximately 96.4–98.0% of the whole bacterial community. Actinobacteria was the most abundant phylum in all treatments, accounting for 36.4–41.7%, followed by Proteobacteria (24.4–29.5%) and Acidobacteria (12.0–16.9%). Many potential PAH degraders, including Rhodococcus, Mycobacterium and Nocardia, belong to the phylum Actinobacteria (Zeinali et al. 2008; Sawulski et al. 2014). Proteobacteria have been found to be the main bacterial phylum in phenanthrene-polluted (Zhang et al. 2018a) and PAH-polluted soils (Xu et al. 2014; Song et al. 2017), reflecting the degradation potential of this bacterial community for PAHs. A significant increase was observed for Actinobacteria and Proteobacteria abundances upon phenanthrene exposure, which was more pronounced for high concentrations of phenanthrene (Additional file 1: Fig. S9). Moreover, they increased with biochar amendment in both the clean soil and PHS but decreased with biochar amendment in the PLS. The opposite results were observed for the phylum Acidobacteria (Additional file 1: Fig. S9), which has also been supposed to play an important role in degrading PAHs (Sawulski et al. 2014; Zhang et al. 2020).

At the genus level, 21 of the 464 bacterial genera had relative abundances higher than 1% in each treatment, which accounted for 49.5–60.1% of the total bacteria. Several genera, such as Sphingomonas, Rhodococcus, Lysobacter, Mycobacterium, Vicinamibacter and Pseudomonas, among the top 21 abundant bacterial genera were identified to have a strong association with the mineralization of phenanthrene. Phenanthrene exposure alone induced dramatic responses of these genera in soil bacterial communities, with the relative abundances increasing by 3.07–36.9% (Fig. 4a). Moreover, the abundances of these six genera all belonging to the phyla Actinobacteria, Proteobacteria and Acidobacteria after incubation for 28 days were significantly and positively correlated with the mineralization fractions of phenanthrene on Day 28 in soil amended with and without biochar (p < 0.05, Additional file 1: Table S4).

Fig. 4
figure 4

Percentage change in the relative abundance of 6 genera associated with mineralization under phenanthrene exposure (a) or biochar amendment in the clean soil (b), PLS (c) and PHS (d) for 28 days. PL and PH indicate soil spiked with low (2 mg kg−1) and high (20 mg kg−1) concentrations of phenanthrene alone, respectively. BnL and BnH indicate soil amended with low (0.03%) and high (0.3%) doses of biochar, respectively. “n” indicates charring temperatures of 300 and 500 °C

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More interestingly, although the bacterial community structure was similar in soil exposed to phenanthrene alone and coexposed to phenanthrene-biochar in the present study (Additional file 1: Fig. S7), the relative abundances of the six genera were higher in biochar-amended PHS than in unamended PHS (positive data in Panel d of Fig. 4), whereas they were lower in biochar-amended PLS than in unamended PLS (negative data in Panel c of Fig. 4). The difference could also be ascribed to the different roles that biochar played in soil polluted with different concentrations of phenanthrene. As PLS had a much lower fraction of mobile phenanthrene than PHS, amending biochar into the PLS would place greater limits on the phenanthrene fraction available for degrading species to utilize relative to that into PHS (Additional file 1: Fig. S2b), which was detrimental for the growth of the six bacterial communities at the genus level. Therefore, their abundances significantly decreased after biochar amendment (Fig. 4c), which contributed to the inhibition of the phenanthrene mineralization fractions in the PLS (Fig. 1a). Similarly, Xia et al. (2015) proposed that the abundances of PAH-degrading bacteria depended on PAH bioavailability as supported by the significantly positive correlations between the copy numbers of the nahAc gene, which typically serves as a biomarker for naphthalene-degrading bacteria, in the deposited sediment and the freely dissolved concentrations of naphthalene in the pore water. Omoni et al. (2020) found that phenanthrene-degrading microbial populations declined as the soil was amended with increasing amounts of biochar, which was also due to the reduction in bioavailable fraction of phenanthrene. In contrast, biochar on its own amended into the clean soil had a positive effect on the abundances of most of the six genera with the exception of Vicinamibacter (Fig. 4b). The relative abundances of the six degrading genera significantly increased after amendment of biochar into the PHS (Fig. 4d), which could be because biochar played a more critical role in providing available nutrients and living conditions for bacteria and modifying the physicochemical properties of soil (Additional file 1: Table S3) rather than reducing the substrate bioavailability to bacteria, thus stimulating the growth of specific degrading bacteria. The effective mineralization of phenanthrene in the biochar-amended PHS and the further mineralization enhancement from biochar amendment at a high level (0.3%) may stem from their impact on these genera (Figs. 1b and 4d).

Of the six bacterial genera, Rhodococcus and Mycobacterium within the phylum Actinobacteria as well as Sphingomonas and Pseudomonas within the phylum Proteobacteria have been frequently reported to have a close relationship with PAH biodegradation/mineralization in biochar-amended soils and sediments (Li et al. 2020; Zhao et al. 2022). For instance, Kong et al. (2018) observed that the abundance of some soil bacterial taxa involved in the degradation of hydrocarbons (i.e., Sphingomonas and Alcanivorax) was promoted by adding 5% sawdust- and wheat straw-derived biochars prepared at 300 and 500 °C to petroleum-polluted soil; therefore, PAH biodegradation in soil was accelerated with an enhanced removal efficiency of at least 20.3%. Ahmad et al. (2021) reported that Pearl River sediment amended with 1% maize straw-derived biochar mainly enriched several genera, including Flavobacterium, Hydrogenophaga, Mycobacterium, Rhodococcus, Flavihumibacter, Pseudomonas and Novosphingobium, responsible for PAH metabolism, contributing to increased pyrene degradation in biochar-amended treatments.

4 Conclusions

In this study, biochar played multiple roles in phenanthrene-polluted soil: reduction of phenanthrene bioavailability, maintenance of soil bacterial diversity, enrichment of the overall microbial/bacterial communities and specific PAH degradation gene (i.e., nidA) as well as alteration in bacterial community composition by increasing or decreasing the abundances of potential PAH degraders, thus greatly affecting the removal of phenanthrene from soil. Whether biochar enhanced or inhibited phenanthrene biodegradation was dependent on not only the amendment level and pyrolysis temperature of the biochar but also the phenanthrene pollution level in the soil. The inhibition of phenanthrene mineralization in the PLS with biochar amendments could be ascribed to their suppression effects on bioavailable phenanthrene for degradation as well as that to specific degrading bacterial genera, which hindered the growth of these degraders and reduced their abundances. In contrast, the effective mineralization of phenanthrene in the PHS after biochar amendments and its further enhancement at a high amendment level could be ascribed to the enhancement of the growth of overall microbial/bacterial communities and the selective stimulation of specific degrading genera and PAH-degradative nidA gene by the nutrient supply from biochar. The findings imply that in the cases of low and high PAH pollution levels in soil, biochar should be designed to immobilize PAHs as an engineered sorbent and facilitate PAH degradation, respectively. Therefore, the concentrations of PAHs are necessary to consider when amending PAH-contaminated soil with biochar.