1 Introduction

The sequestration of atmospheric CO2 into soil organic carbon (SOC) has been identified as a major strategy to combat climate change (Field and Mach 2017; Minasny et al. 2017; IPCC 2022). Therefore, it is imperative to identify soil management practices that not only increase soil C storage (and its associated benefits to agriculture) but also counteract the depletion of SOC (Kopittke et al. 2022). Biochar amendment to agricultural soils has been well-recognized for its potential in soil C sequestration and mitigating climate change (Woolf et al. 2010; Gross et al. 2021; Lehmann et al. 2021). In addition, production and soil amendment with biochar can provide  bioenergy and agronomic benefits, and reduce non-CO2 greenhouse gas emissions (Lehmann 2007; Woolf et al. 2016; Joseph et al. 2021; Kalu et al. 2022).

Biochar soil application leads to an immediate increase in SOC but how biochar interacts with native and exogenous organic substrates in the long term is not clear. In agricultural soils, exogenous organic substrates such as plant litter, root exudates, and organic amendments are continuously added. Biochar can further increase soil C storage by preserving such exogenous organic substrates and their microbial derivatives from decomposition by promoting strong organo-mineral binding on its surface (Pan et al. 2021; Weng et al. 2022). On the other hand, the addition of exogenous organic substrates such as root exudates can accelerate the mineralization of SOC (priming effect) even when it is strongly bound on organo-mineral interfaces (Keiluweit et al. 2015), stimulating the loss of both exogenous and native SOC (Fu et al. 2022). Therefore, the potential of biochar for additional soil C sequestration via preservation of new C inputs or reduced priming effects of native SOC depends on complex interactions between biochar, SOC and exogenous organic substrates. How the fresh root exudates (common exogenous organic substrates) in soil-biochar systems simultaneously affect the fate of old-native SOC and the underlying mechanisms remain elusive. The complex interactions and our limited understanding of the associated mechanisms increase the uncertainty to predict long-term C sequestering potential of biochar. One of the important mechanisms controlling the preservation and stabilization of SOC is occlusion within the soil aggregates and sorption onto mineral surfaces (Lehmann and Kleber 2015). Physical fractionation of SOC can provide insights into the mechanistic understanding of the stabilization of fresh and native SOC after biochar amendment.

Priming effect (PE) is a synergistic change in mineralization rates of native SOC following the addition of fresh organic substrate inputs (e.g., fresh plant litter, root exudates or organic amendments) (Kuzyakov 2002). Positive PE occurs when exogenous organic substrate inputs stimulate soil microbial activities in a way that accelerates the loss of native SOC (Zhou et al. 2022; Yan et al. 2023). Occasionally, soil microbes can shift their preference from native SOC to easily available fresh organic substrates (substrate switching), leading to the negative PE or retardation of SOC decomposition (Guenet et al. 2010). The change in physico-chemical and microbial properties of soil after biochar application can alter the magnitude and direction of PE. A meta-analysis by Wang et al. (2016) concluded that biochar slightly retarded SOC decomposition by 3.8% compared to soil without biochar. This indicates that biochar can contribute to additional C storage in soil, which if continued for a long time, can in some conditions lead to impressive increases in SOC stocks as demonstrated by the famous Terra Preta soils in Amazonia, which contain 250 Mg ha−1 total organic C (TOC) compared to only 50 Mg ha−1 in adjacent Ferralsols, while only 20% of TOC is covered by biochar (Glaser et al. 2001). However, the magnitude and direction of PE following biochar application in soils can vary depending on the characteristics of biochar, soil and environmental variables (Zimmerman et al. 2011; Fang et al. 2015), making it difficult to predict.

Microbial carbon use efficiency (CUE), defined as the ratio between the amount of C allocated to biosynthesis (new biomass and biological products, including exudates) and the amount of C taken up by microbes (Manzoni et al. 2018), indicates the efficiency with which microbes convert available organic substrates into stable biosynthesized products (Geyer et al. 2016). High CUE favors the accumulation of SOC storage through increased microbial biomass and by-products or microbial residues, which can be stabilized in soils in the long term (Sinsabaugh et al. 2013; Liang et al. 2017; Tao et al. 2023). High CUE is expected in environments with better nutrient balance and water dynamics (Manzoni et al. 2012). Since biochar reduces soil environmental stress by improving water and nutrient retention, modulating soil pH and providing suitable habitats for the microbes, biochar has the potential to increase microbial CUE (Liu et al. 2020; Pei et al. 2021). But still, the number of studies and our understanding of the linkage between stabilization or depletion of C in biochar-amended soils and CUE are limited (Giagnoni and Renella 2022).

Here, we conducted a soil incubation study for six months with 13C labelling technique with the objectives: (1) to explore the effects of biochar on the mineralization of native SOC and added glucose (as a representative of root exudates) in a fine-textured soil and (2) to shed light on mechanistic understanding of how biochar affects the process of stabilization or depletion of SOC and 13C-glucose. We addressed the following hypotheses: (1) biochar amendment influences mineralization of SOC (priming effect) and fresh labile organic substrate; (2) the mineralization of SOC and fresh labile organic substrate depends on the amounts of biochar added; (3) biochar facilitates microbial uptake of fresh labile organic substrate and decreases their energy expenditure (lowers respiration), leading to higher CUE; (4) biochar preserves and stabilizes the microbial necromass derived from fresh labile organic substrate after they die.

2 Materials and methods

2.1 Biochar and soil

Biochar was produced from hardwood branches and split. The composition of hardwood species was 80–90% willow (Salix spp.), 5–10% birch (Betula spp.), and 5–10% of other hardwood species such as alder (Alnus spp.), bird cherry (Prunus padus L.), and Norway maple (Acer platanoides L.). The hardwood feedstock was slowly pyrolyzed using a 0.3 m3 Kon-Tiki kiln. After the pyrolysis, the biochar was soaked with a mixture of tap water and cattle slurry at a ratio of 7:3. The detailed properties of the biochar were  presented in Kiani et al. (2021). In brief, the biochar had a pH of 9.9, a total C content of 86%, a total N content of 0.3%, and a  specific  surface area of 199 m2 g−1.

The soil used in this study was fine-textured ( silty clay loam with 3% sand, 78% silt, and 19% clay) Endogleyic-Stagnosol (IUSS Working Group WRB 2007) collected from 0–30 cm depth from an agricultural field located in Hyvinkää (60°35′38.5ʺN, 24°56′26.3ʺE) in southern Finland. The soil had  a pH of 6.8 and electrical conductivity of 46 µS cm−1. The total C and N contents of the soil were 3.7% and 0.3%, respectively.

2.2 Soil incubation

The experiment consisted of six treatments (i) Soil-only, (ii) Soil + B15, (iii) Soil + B30, (iv) Soil + G, (v) Soil + B15 + G, and (vi) Soil + B30 + G with four replicates. The “B15” and “B30” refer to two rates of biochar applications corresponding to 15 and 30 Mg ha−1 in field  conditions, respectively and the “G” refers to the addition of 13C-glucose.

After passing the soil through a 4 mm sieve to remove large plant particles and gravel, 41 g fresh soil (= 35 g in dry weight basis) was filled in a 120 mL borosilicate glass bottle. Then 0.29 g and 0.58 g of biochar (= 0.24 g and 0.48 g dry weight basis) were mixed with soil for the respective B15 and B30 treatments. The glass bottles were closed with a rubber septum and further enclosed with an aluminum crimp cap to ensure airtight condition. Altogether, 168 glass bottles (6 treatments × 4 replicates × 7 sets) were first pre-incubated at 15 °C for 78 days at 45% water holding capacity (WHC) to balance out the mineralization and/or priming occurred due to biochar addition. During the pre-incubation, soil respiration was measured continuously, more frequently in the beginning, and sparsely at the end. The purpose of including 7 sets of samples was to take soil samples destructively 7 times during the experiment:  on the day of 1, 4, 7, 14, 28, 97, and 180 after 13C-glucose addition.

After the pre-incubation, 15 mg of uniformly labeled 13C-glucose (20 at%) was added per g soil C (the same rate as in Hartley et al. (2010)) into all glucose addition treatments (indicated by “G” in the treatment name). Glucose was added as a solution (3 ml per soil sample), resulting in the final WHC of 60%. In non-labelled treatments, a corresponding amount of Milli-Q water was added to maintain the same WHC. This soil moisture content was maintained throughout the study by periodically checking the weight of the bottle and adding the necessary amount of Milli-Q water.

2.3 Sampling and measurement

2.3.1 CO2 and 13CO2 production

The CO2 and 13CO2 production were measured throughout the experiment only from the last set of experimental units, intended for destructive soil sampling 180 days after 13C-glucose addition. During pre-incubation, when 13C-glucose was not yet added, CO2 production was measured only from 3 treatments—“Soil-only”, “Soil + B15”, and “Soil + B30” on the day of 1, 3, 5, 7, 14, 21, 28, 42 and 78  after biochar addition. For measuring CO2 flux at this phase, the bottles were first flushed with ambient air for 1 min followed by enclosing the bottles with airtight caps. Thereafter, using a micro-syringe (Hamilton Co), 100 µL of gas samples were taken from the bottles and inserted into a Gas Chromatograph (GC; Hewlett-Packard—HP 6890 GC System by Agilent Technologies Inc) equipped with a flame ionization detector (FID) for measuring CO2 concentration at 0, 6 and 24 h after closing the bottles. The CO2 flux was calculated using linear regression between the CO2 concentrations  and measurement time. On other days when no CO2 flux was measured, the bottles were closed with ventilation caps for air circulation.

After 13C-glucose addition, CO2 production was periodically measured from all six treatments from the last set of the experimental units on  the day of 0, 1, 2, 3, 4, 5, 6, 7, 11, 14, 19, 50, 68 and 106, while 13CO2 production used to calculate the PE was measured until day 19 only. First, the bottles were enclosed with a rubber septum and aluminum crimp cap followed by flushing with CO2-free air at the rate of 1 L min−1 for 1 min to remove all the CO2 present inside the bottle. Then, the bottles were over-pressurized by inserting 7 mL of CO2-free air. Consequently, the CO2 concentrations were measured 6 and 24 h afterward, by introducing 100 µL of gas samples from the bottles to the GC. Right after 13C-glucose addition, higher 13CO2 emissions were expected, therefore, CO2 concentration was measured only after 6 h. After that, 7 mL of gas sample  was taken from the bottle using a syringe and injected into 6 mL evacuated exetainer for measuring 13CO2 production. The gas samples were analyzed for 13CO2 signature using an Isotope Ratio Mass Spectrometer (IRMS; Delta plus XP, Thermo Fisher Scientific, Bremen, Germany) coupled with a Gas Chromatograph (GC-Box, Thermo Fisher Scientific, Bremen, Germany and Poraplot Q, Combi-PAL autosampler, Zwingen, Switzerland) via a Conflo III Interface (Thermo Fisher Scientific, Bremen, Germany). The amount of CO2 produced from the mineralization of either added 13C-glucose or native SOC was calculated using the two-pool mixing model (Eqs. 1 and 2), and the PE was calculated using Eq. 3. The cumulative CO2 or 13CO2 production was calculated through linear interpolation or trapezoidal method.

$${{\text{R}}}_{{\text{Glucose}}}= \frac{{}^{13}{{\text{CO}}}_{2}-{}^{13}{\text{SOC}}}{{}^{13}{{\text{C}}}_{{\text{Glucose}}}-{}^{13}{\text{SOC}}}\times {{\text{R}}}_{{\text{Total}}}$$
(1)
$${{\text{R}}}_{{\text{SOC}}}= {{\text{R}}}_{{\text{Total}}}-{{\text{R}}}_{{\text{Glucose}}}$$
(2)
$$\mathrm{Priming\, effect}= {{\text{R}}}_{{\text{Trt}}}-{{\text{R}}}_{{\text{Ctrl}}}$$
(3)

where RGlucose is CO2 derived from the added 13C-glucose; 13CO2 is 13C-at% of sample; 13SOC is 13C-at% of soil, the 13C-at% of soils in the treatments without 13C-glucose (Soil-only, Soil + B15 and Soil + B30) was the same = 1.08; 13CGlucose is 13C-at% of added 13C-glucose; RTotal is the total CO2 production; RSOC  represents CO2 derived from SOC; RTrt is the CO2 derived from SOC in a sample treatment and RCtrl is the CO2 derived from SOC in the “Soil-only” control treatment.

2.3.2 Soil microbial biomass carbon (MBC) and 13C in microbial biomass

Soil microbial biomass carbon (MBC) was measured on   the day of 1, 4, 7, 14, 28, 97 and 180  after 13C-glucose addition using the chloroform fumigation extraction (CFE) method (Vance et al. 1987). For that, 8 g of moist soil samples were fumigated with chloroform inside a desiccator for 24 h in the dark followed by their extraction with 40 mL 0.05 M K2SO4 by shaking on an orbital shaker (200 rpm, 30 min). The extracts were filtered using Whatman Grade 42 ashless filter paper and subsequently filtered through a 0.45 μm syringe filter (Sartorius, Minisart High Flow, PES). At the same time, another soil sample without fumigation was also extracted similarly. The extracts were then analyzed for dissolved organic C (DOC) using a TOC analyzer (Analytik Jena multi N/C 3100, Jena, Germany). The MBC was calculated by subtracting the DOC of the non-fumigated extract (NF) from the fumigated extract (F).

The K2SO4 extracts of samples obtained on the day of  1, 14, and 180 were freeze-dried followed by 13C analysis using an IRMS (Delta C, Thermo Electron, Bremen, Germany) coupled with an Elemental Analyzer (Costech ECS 4010, Costech Analytical Technologies Inc., Valencia, USA) via a Conflo III Interface (Thermo Electron, Bremen, Germany). The 13C in the microbial biomass (13MBC) and microbial carbon use efficiency (CUE) of 13C-glucose were calculated using Eqs. 4–7 as in Mganga et al. (2022). The CUE was calculated for the day of 1 and 14 after the 13C-glucose addition.

$${}^{13}{{\text{MBC}}}_{{\text{x}}}=\frac{{}^{13}{{\text{DOC}}}_{{\text{F}}}\times {{\text{DOC}}}_{{\text{F}}}-{}^{13}{{\text{DOC}}}_{{\text{NF}}}\times {{\text{DOC}}}_{{\text{NF}}}}{{{\text{DOC}}}_{{\text{F}}}-{{\text{DOC}}}_{{\text{NF}}}}$$
(4)
$${}^{13}\mathrm{MBC\%}= \frac{{}^{13}{{\text{MBC}}}_{{\text{T}}}-{}^{13}{{\text{MBC}}}_{{\text{C}}}}{{}^{13}{\text{Glucose}}-{}^{13}{{\text{MBC}}}_{{\text{C}}}}\times 100$$
(5)
$${}^{13}{\text{MBC}}=\left({{\text{DOC}}}_{{\text{F}}}-{{\text{DOC}}}_{{\text{NF}}}\right)\times \frac{{}^{13}\mathrm{MBC\% }}{100}$$
(6)
$${\text{CUE}}= \frac{{}^{13}{\text{MBC}}}{{}^{13}{\text{MBC}}+{{\text{R}}}_{{\text{Glucose}}}}$$
(7)

where 13MBCx represents either 13MBCT or 13MBCC in Eq. (5); 13DOCF, DOCF, 13DOCNF, and DOCNF represent the 13C-at% and total C concentrations (mg C kg−1 soil) of fumigated (F) and non-fumigated (NF) K2SO4 extracts, respectively. 13MBCT and 13MBCC (denoted as 13MBCx in Eq. 4) are the 13C-at% of sample treatments and the control (Soil-only treatment), respectively; 13Glucose is the 13C-at% of added 13C-glucose solution (20 at%). 13MBC is total microbial growth derived from added 13C-glucose and RGlucose is the cumulative respiration derived from added 13C-glucose.

2.3.3 Soil density fractionation

The remaining soil samples from the 180  day samples were dried at 60 °C for 72 h and stored in room temperature until processed for the fractionation into free-particulate (f-POM), occluded-particulate (o-POM), and mineral-associated organic matter (MAOM) according to methods mentioned in Weng et al. (2017) with slight modification. For the fractionation, about 10 g of dry soil samples were added with 40 mL of 1.85 g cm−3 sodium polytungstate (SPT) solution in a centrifuge tube. After gently inverting the soil SPT mixtures 10 times, it was centrifuged at 2000 g for 1 h. Then, the floating particles were filtered using 45 µm pore-sized filter paper (GN-6 Metricel® membrane, Pall membrane filters), washed with 100 mL of Milli-Q water to remove the SPT solution and dried at 60 °C until the weight remained constant for 24 h. This fraction is the f-POM.

The remaining soil in the centrifuge tube was washed by adding 40 mL of Milli-Q water, centrifuged at 2000 g for 30 min and discarding the supernatant. This process was repeated 5 times to remove all the SPT. After washing, 40 mL of Milli-Q water was added on top of the decanted sample along with 6 glass beads. The mixture was shaken in an orbital shaker overnight (18 h at 250 rpm) to break all the aggregates. Afterward, the mixture was passed through a 63 µm sieve. The fraction retained on the sieve was the o-POM and the one passing through the sieve was MAOM. These fractions were transferred to the pre-weighed aluminum tray and dried at 60 °C until the weight remained constant for 24 h.

These fractions as well as bulk soil were grinded using a mortar and pestle and then analyzed for C and 13C contents using the IRMS (Delta C, Thermo Electron, Bremen, Germany) coupled with the Elemental Analyzer (Costech ECS 4010, Costech Analytical Technologies Inc., Valencia, USA) via the Conflo III Interface (Thermo Electron, Bremen, Germany). The amount of 13C remained after 6 months (180 days) in the bulk soil, and the different density fractions (f-POM, o-POM, MAOM) were calculated using the two-pool mixing model, similarly as in Eq. (1).

2.3.4 Extraction of amino sugars

The soil amino sugar fractions were extracted from samples destructively sampled at the end of the experiment (180 days) from three treatments “Soil-only”, “Soil + G” and “Soil + B30 + G”. In addition, the biochar particles recovered from “Soil + B30” and “Soil + B30 + G” treatments were extracted for amino sugars (biochar particles had to be pooled from different analytical replicates to extract enough amino sugars for the 13C analysis). For determination of 13C in total amino sugar fraction, grinded biochar particles or soil( + biochar mixtures) were hydrolyzed with 6 M HCl as described by Dippold et al. (2014), and purified through cation-exchange columns (Poly-Prep® Columns, AG® 50W-X8, hydrogen form mesh size 100–200; Biorad, Munich, Germany, Cat# 731–6213), in order to remove hydrolyzable cationic compounds such as carboxylic acids and inorganic cations such as iron and aluminum. The 13C in total amino sugar fraction was measured using a EURO EA Elemental Analyzer (EuroVector, Hekatech, Germany) coupled via a Conflo III Interface to an IRMS (Finnigen Delta V Advantage, Thermo Scientific, Bremen, Germany). Sucrose (ANU, IAEA, Vienna, Austria) and CaCO3 (NBS 19, TS limestone) were used as calibration standards. The precision of 13C measurements was 0.2‰.

2.4 Data analysis

The differences in treatments were analyzed using one-way analysis of variance (ANOVA) followed by the least significant difference (LSD) test to test the significant differences between the treatments (P < 0.05). The assumptions of normality and homogeneity of variances were tested using the Shapiro–Wilk test and Levene’s test. The Shapiro–Wilk test was carried out on the residuals of ANOVA for testing the normality. All the results shown represent the average of the four replicates (n = 4) ± standard error (SE). The analyses were performed in R Statistical Software v4.2.2 (R Core Team 2022).

3 Results

3.1 CO2 production

During the pre-incubation, CO2 production rapidly increased immediately after the addition of biochar and was proportional to the biochar application rate (Fig. 1). However, only after a few days, CO2 production rates from biochar treatments sharply decreased and were even lower than the control soil without biochar. This trend continued during the entire pre-incubation period of 78 days, indicating that biochar addition significantly reduced the mineralization of native SOC or negative PE. At the end of the pre-incubation, cumulative CO2 productions from biochar-amended soils were 20% to 36% less than in the Soil-only control.

Fig. 1
figure 1

a CO2 production rate and b cumulative CO2 production after the addition of biochar between the treatments during the pre-incubation. The figure presents the average of the four replicates (n = 4) and the error bars represent SE. Different lowercase letters in b represent the significant differences in total cumulative CO2 production between the treatments (P < 0.05)

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After glucose addition, CO2 production rapidly increased for a short period and peaked within 3 days in all glucose-added treatments (Fig. 2). Among them, the treatments with glucose added to biochar-amended soil had significantly lower CO2 production compared to the treatment with glucose added to soil without biochar. There was also a clear effect of biochar amount, i.e., more biochar addition resulted in less CO2 production. The trend in the total CO2 production in treatments without glucose addition was the same as during the pre-incubation i.e., Soil-only > Soil + B15 > Soil + B30.

Fig. 2
figure 2

a CO2 production rate and b cumulative CO2 production after glucose addition between the treatments. The figure presents the average of the four replicates (n = 4) and the error bars represent SE. Different lowercase letters in b represent the significant differences in total cumulative CO2 production between the treatments (P < 0.05)

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Likewise, the production of CO2 derived from SOC and glucose among the glucose-added treatments followed the same trend as the total CO2 production (Figs. 3 and 4) i.e., Soil + G > Soil + B15 + G > Soil + B30 + G. The addition of glucose led to immediate positive priming in all three treatments amended with glucose. However, after 12 days, the mineralization of SOC decreased in biochar-added treatments leading to negative PE, while the Soil + G treatment continued to exhibit positive PE (Fig. 3c). Compared to the Soil + G control, Soil + B15 + G and Soil + B30 + G treatments had 12% and 15% less cumulative CO2 production derived from SOC, respectively. There was no significant difference between the Soil + B15 + G and Soil + B30 + G treatments on the production of CO2 derived from SOC (Fig. 3b).

Fig. 3
figure 3

a CO2 production rate originating from SOC decomposition, b cumulative CO2 production derived from SOC, and c priming effect between the treatments. The figure presents the average of the four replicates (n = 4) and the error bars represent SE. Different lowercase letters in b represent the significant differences in total cumulative CO2 production derived from SOC between the treatments (P < 0.05)

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Fig. 4
figure 4

a CO2 production rate and b cumulative CO2 production originated from glucose between the treatments. The figure presents the average of the four replicates (n = 4) and the error bars represent SE. Different lowercase letters in b represent the significant differences in total cumulative CO2 production derived from glucose between the treatments (P < 0.05)

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Similarly, the biochar treatments (Soil + B15 + G and Soil + B30 + G) significantly decreased the production of 13CO2 derived from glucose compared to the Soil + G treatment (P < 0.05) (Fig. 4b). The higher the biochar application rate, the lower the mineralization of added glucose. The most significant reduction in glucose mineralization by the biochar occurred immediately after the addition (on Days 1 and 3; Fig. 4a). Compared to the Soil + G control, Soil + B15 + G and Soil + B30 + G treatments had 11% and 17% less cumulative CO2 production derived from added glucose, respectively (Fig. 4b). After 19 days, 32–35% of the added glucose had been mineralized in the biochar treatments, whereas 39% of the added glucose had been mineralized in the control treatment.

3.2 Microbial biomass carbon and carbon use efficiency

Biochar treatments with glucose addition (Soil + B15 + G and Soil + B30 + G) had significantly higher MBC than other treatments throughout the 180 days of incubation (Fig. 5a). However, there was no difference in MBC between the Soil + B15 + G and Soil + B30 + G treatments. Also, no statistically significant effect of biochar was observed on 13MBC although the average values were slightly higher in the biochar treatments than in the Soil + G treatment throughout the experiment (Fig. 5b).

Fig. 5
figure 5

a Soil microbial biomass carbon (MBC) and b 13C in microbial biomass (13MBC) between the treatments over 180 days of the incubation. The figure presents the average of the four replicates (n = 4) and the error bars represent SE. Different lowercase letters represent the significant differences between the overall average of the treatments (P < 0.05)

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Biochar addition increased microbial CUE. Greater CUE was found in the higher biochar application rate treatment (Fig. 6). On day 1, the greater variance in 13MBC (Fig. 5b) resulted in greater variance in CUE, constraining the statistical significance (P > 0.05). However, on day 14, the Soil + B30 + G had significantly higher CUE than the Soil-only control (P < 0.05).

Fig. 6
figure 6

Microbial carbon use efficiency (CUE) of glucose in 13C-glucose-amended treatments on (a) Day 1 and (b) Day 14 after 13C-glucose addition. The figure presents the average of the four replicates (n = 4) and the error bars represent SE; no error bars are shown for a because of the large SE. Different lowercase letters represent the significant differences between the treatments (P < 0.05)

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3.3 Partitioning of soil C in different fractions

After 6 months, biochar-amended soils had significantly higher soil C content than the Soil-only control (P < 0.05). The 15 Mg ha−1 biochar treatments (Soil + B15 and Soil + B15 + G) had 20% and 30 Mg ha−1 biochar treatments (Soil + B30 and Soil + B30 + G) had 32% higher total soil C content compared to the Soil-only control (Fig. 7a). After 6 months, about 29–30% of the 13C label remained in glucose-amended soils. The percentage of 13C remaining in Soil + B30 + G treatment was slightly higher but not statistically significant (P > 0.05) than Soil-only and Soil + B15 + G treatments (Fig. 7b).

Fig. 7
figure 7

a Soil C contents and b remaining 13C in soil after 6 months of the experiment. The percentage in (b) represents the percentage of added 13C-glucose. The figure presents the average of the four replicates (n = 4) and error bars represent SE. Different lowercase letters represent the significant differences between the treatments (P < 0.05)

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Soil fractionation revealed that most of the soil mass (88–90%; Fig. 8) and the remaining 13C (about 22% out of 29–30% of remaining 13C; Fig. 9) were recovered in mineral-associated fraction (MAOM). Most of the biochar particles were recovered in the f-POM fraction because biochar-amended soils had significantly higher (P < 0.05) mass of f-POM fraction than the Soil-only and Soil + G treatments (Fig. 8). Also, the mass of o-POM fraction was significantly higher (P < 0.05) in biochar treatments (2.8% and 2.6% in Soil + B15 and Soil + B30 treatments, respectively) compared to the Soil-only treatment (2.1%), particularly in treatments without glucose addition (Fig. 8), indicating that biochar increased soil occlusion or aggregation in non-glucose amended treatments.

Fig. 8
figure 8

Proportion of different soil organic matter fractions (f-POM, o-POM, and MAOM) after 6 months of incubation. The y-axis is in logarithmic scale. The figure presents the average of the four replicates (n = 4) and the error bar represents SE. Different lowercase letters represent the significant differences between the treatments (P < 0.05)

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Fig. 9
figure 9

Recovery of added 13C in different soil fractions among the 13C-glucose-added treatments after 6 months. The figure presents the average of the four replicates (n = 4) and the error bars represent SE. Different lowercase letters represent the significant differences between the treatments (P < 0.05)

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The 13C contents in the soil fractions showed a significantly higher (P < 0.05) amount of 13C remaining in the f-POM fraction in the biochar treatments (1.3% and 1.8% of added 13C remaining in Soil + B15 + G and Soil + B30 + G treatments) than in the Soil + G treatment (0.7%) after 6 months (Fig. 9). However, no differences in the remaining 13C were found between the treatments in o-POM and MAOM fractions.

3.4 Amino sugars

Soil amended with biochar (Soil + B30 + G) had significantly higher total amino sugar C and 13C amino sugar contents than controls (Soil-only and Soil + G treatments) (P < 0.05) (Table 1). In addition, 13C at% of amino sugar measured from biochar particles collected from the treatment receiving 13C-glucose (Soil + B30 + G) was higher compared to that of soil from Soil + G or Soil + B30 + G treatments indicating that dead microbial residues derived from consuming 13C-glucose were concentered on biochar particles. The percentage of 13C in amino sugars (percentage of the total 13C remaining in the soil at the end of the incubation) was on average 10.8% in the Soil + G treatment and 11.2% in the Soil + B30 + G. Our results indicated that 13C of added glucose was incorporated into soil microorganisms and stabilized in biochar pores as microbial necromass, i.e., in the form of amino sugars.

Table 1 Total amino sugar C and 13C amino sugar contents of soil and biochar particles at different treatments
Full size table

4 Discussion

4.1 Biochar reduced SOC mineralization: negative priming

Immediately after the addition, biochar increased CO2 production for one week (Fig. 1a). This immediate short-term increase in CO2 production after biochar addition is common and mainly attributed to the presence of labile C fraction in biochar, which fuels the microbes in the short term (Maestrini et al. 2015). Previous studies also observed increased short-term mineralization of SOC immediately after biochar addition, but this positive priming decreased over time (Luo et al. 2013; Singh and Cowie 2014). However, similar 13C signatures of soil and biochar in this study did not allow us to separate the proportion from which this CO2 production originated (soil or biochar). Nevertheless, it was evident that after one week, biochar decreased the mineralization of native SOC, i.e., negative priming of native SOC occurred for the rest of the pre-incubation period. This result shows that biochar-induced positive priming is short-lived followed by negative priming (Zimmerman et al. 2011; Maestrini et al. 2015; Weng et al. 2015).

The addition of easily mineralizable glucose indeed induced short-term positive PE from the soil-biochar system for 12 days (Fig. 3c). This is linked to the immediate microbial blooming (Fig. 5a). However, in the longer term (after 12 days of glucose addition), added biochar reduced SOC mineralization i.e., negative PE of native SOM occurred. This biochar-induced negative PE could arguably be explained by an increase in soil occlusion or aggregation by the biochar, as indicated by increased o-POM in the biochar treatments (Fig. 8). Our results conform to a previous study, which also observed an increase in soil aggregate stability when the same soil used in this experiment was amended with a wood-based biochar (Soinne et al. 2014). The binding of soil organic and mineral components at the large surface area of biochar increases inter-particular cohesion, which is the primary step in the aggregate formation and stabilization process (Gul et al. 2015). The increase in soil aggregation by biochar protects the native SOC from microbial degradation (Weng et al. 2017; Fang et al. 2018).

Substrate switching and dilution   effects (preference of the microbes to decompose labile biochar C and other easily mineralizable substrates over native SOC because of biochar) are the other possible mechanisms behind biochar-induced negative priming (Joseph et al. 2021). However, such mechanisms usually result in the transient negative priming immediately after the substrate or biochar addition, followed by a positive priming (Whitman et al. 2014), which was not observed in this study. In some cases, biochar might decrease microbial biomass (Dempster et al. 2012) that could lead to negative priming. On the contrary, MBC increased especially in the glucose-added biochar treatments in this study. Hence, physical protection of SOC due to increased soil aggregation and/or sorption of labile native SOC into the biochar porous matrix  seems plausible mechanisms behind the observed negative PE.

4.2 Biochar particles retained added 13C

Most of the biochar particles were recovered in f-POM fraction. The significantly higher recovery of added 13C in f-POM fraction of biochar treatments (Fig. 9a) denotes that the added 13C was recovered in biochar particles. This demonstrates the additional preservation of labile organic substrates (e.g., root exudates) inside or in close proximity to biochar particles (charosphere region). This increased recovery of 13C in biochar particles could be due to the physico-chemical sorption or via microbial pathways. The physico-chemical sorption of added glucose in biochar particles occurs mostly immediately after the addition of glucose (Melas et al. 2017). However, in the longer term, part or most of the added glucose would have been desorbed and consumed by soil microorganisms. Our results agree with the concept that biochar usually benefits microbial growth and activity because of the supply of labile C present in biochar (Luo et al. 2013; Farrell et al. 2013) as well as serving as the preferred microbial habitat in its large and porous surface matrix (Pietikäinen et al. 2000; Lehmann et al. 2011). The increased microbial biomass and CUE of glucose in biochar-amended soil (especially in the higher biochar application rate treatments) suggest that the added glucose might have been effectively and/or preferentially used by soil microorganisms to synthesize new biomass. This is supported by the increased 13MBC in biochar-amended soils at the beginning of the experiment (Fig. 5b; although not statistically significant). Eventually, 13C in microbial biomass was converted to microbial byproducts (microbial necromass and metabolites).

Biochar facilitated stable microbial necromass formation from the added labile C substrate, as indicated by the higher recovery of 13C in amino sugar by biochar (Table 1). In agreement with our result, some previous studies also suggested that biochar can enhance microbial necromass (Glaser and Birk 2012; Weng et al. 2022; Zhang et al. 2022). Higher accumulation of 13C into amino sugars (microbial necromass) also indicates a higher capacity to store SOC in rather stable forms in biochar-amended soils (Zhu et al. 2020). Microbial necromass is sticky in nature (Dufrêne 2015; Buckeridge et al. 2020) and could strongly adsorb on biochar, hence increasing the long-term CUE and thus 13C retention in biochar particles. The CUE measured after 14 days in this study can be considered ecosystem level CUE, which measures not only the effective uptake of 13C-glucose to synthesize microbial biomass but also includes the efficiency of the substrate (microbial necromass and exudates) recycling, and stabilization in soil (Geyer et al. 2016). Microbial CUE indicates the microbial C sequestration potential in soils because it compares microbial decomposition of organic substrate and stabilization of microbially assimilated C (Manzoni et al. 2018). In our study, biochar enhanced CUE because biochar reduced the mineralization of 13C glucose, while increasing the proportion of microbially assimilated C, and its stabilization as microbial necromass. Greater CUE also indicates that biochar reduced environmental stress to microbes by regulating soil pH (Pei et al. 2021) and providing nutrient supply (Liu et al. 2020).

Previous studies suggested that biochar increases organo-mineral interaction and thus  enhances the retention of (labile) organic matter in MAOM (Weng et al. 2017; Akpinar et al. 2023; Gianetta et al. 2023; Zhang et al. 2023). Biochar did not increase the amount of SOC stabilized in the MAOM fraction in this study (Fig. 9c). A probable explanation for this observation is that the tightly held organo-mineral layer, formed on the biochar surface after  its prolonged exposure to soil (Hagemann et al. 2017), was probably not easily disintegrated into the solution when dispersed in SPT during density fractionation. Yet, the density of biochar with the organo-mineral layer was still lower than 1.85 g cm−3 SPT, hence, it was recovered in the f-POM fraction. Because more clay minerals attached to biochar particles could have been recovered in f-POM fraction in the treatments with high biochar application rate, the mass of clay mineral-dominated MAOM fraction in the higher biochar application rate treatment (Soil + B30 + G) was significantly lower than in the Soil + G treatment (Fig. 8). Interestingly, this significant difference was only observed in glucose-amended treatments. This suggests that glucose or glucose-derived microbial metabolites might have increased the interaction of biochar and clay minerals. In this organo-mineral-biochar interface, multivalent cations from soil minerals can bridge negatively charged functional groups of organic molecules originating either from SOC or newly added organic substrate or their microbial derivatives on one side, and negatively charged biochar surface on the other (Weng et al. 2022).

The retention of added labile organic substrate or its microbial derivates by the biochar was relatively small. For instance, the  retention of 13C in f-POM in Soil + G control was 0.7% whereas that in the biochar treatments: Soil + B15 + G, and Soil + B30 + G treatments were 1.3% and 1.8% of the added 13C, respectively, after 6 months (Fig. 9a). Similarly, the retentions of 13C amino sugars in the Soil + G control and Soil + B30 + G treatments were 10.8% and 11.2% of the total 13C retained, respectively. Nevertheless, the ability of the biochar to retain the added labile organic substrate and its microbial derivates could be expected to be additive over time under field conditions, where plants continuously release root exudates in the soil (Joseph et al. 2021; Weng et al. 2022). Even though this study was limited to only one soil and biochar type, it does provide evidence for (1)  the reduction of SOC mineralization and (2) efficient microbial utilization of added labile substrate in the presence of biochar and (3) eventual protection or stabilization of added labile substrate inside biochar particles as microbial necromass, which helps in promoting soil C sequestration. However, extrapolation of our results with a simple soil-biochar and glucose system from a controlled environment at constant temperature and moisture content to wider fields with complex organic inputs (e.g. other soil amendments, plant litter) should be considered cautiously. Further experiments under field conditions with different organic substrate inputs, soil and biochar types will improve our understanding of the key factors affecting mineralization of fresh and added SOC dynamics after soil biochar application.

5 Conclusions

Biochar reduced the mineralization of SOC by 12–15% and added glucose by 11–17%. Our results demonstrated the additional soil C storage ability of wood-based biochar by preserving SOC from mineralization (negative priming) and stabilizing fresh labile substrate. The higher application rate of biochar (30 Mg ha−1) has higher soil C sequestration potential aided with increased negative priming and increased retention of added labile organic substrate. The inaccessibility of SOC to the microbes due to increased soil occlusion in the biochar-amended soil preserved SOC from mineralization. Similarly, biochar increased microbial CUE, suggesting that biochar helped in the effective conversion of labile organic substrate to stable microbial products. Furthermore, 13C amino sugar analysis of biochar particles confirmed that microbial necromass derived from the labile organic substrate was stabilized inside biochar’s porous matrix. Over the long run, conversion  and stabilization of fresh labile organic substrate into microbial necromass by biochar could be a way to form a greater fraction of stable SOC.