Bacterial isolation was carried out in the summer of 2020 from the rhizosphere of Epipactis atrorubens (Hoffm). Besser, belongs to the Orchidaceae family, and thrives in a natural forest community on serpentine soil. Serpentine substrates are known for their unfavorable physico-chemical properties for plant growth such as stoniness, poor water-holding capacity, high insolation, significantly high pH value, and large temperature differences. In addition, they differ by a large amount of certain HMs and low concentration of macronutrients such as nitrogen and phosphorus and high Mg/Ca ratio [25, 26, 27].

The rhizosphere soil was collected in a UV-sterilized zip-lock bag and brought to the laboratory. Ten grams of soil sample (on a dry weight basis) were mixed with 90 mL of sterilized distilled water and homogenized by shaking at 180 rpm for 1 hour in an Erlenmeyer conical flask. After serial dilution in sterilized distilled water, a 100 µL aliquot of required suspension was plated on Luria–Bertani (LB) agar (HiMedia, Mumbai, Maharashtra, India) plate and incubated for 3 days at 28 $\pm$ 2 °C in an incubator (Biosan ES-20/60, Riga, Latvia). After incubation, the morphologically different bacterial colonies were picked carefully and further purified by culturing on LB agar media using quadrant streaking. The pure isolates were preliminarily identified by their morphology, physiology, biochemical test, and cross-verified by Bergey’s manual [28].

Biochar (BC), produced by a domestic manufacturer from high-density birch wood by oxygen-free pyrolysis, was crushed and passed through a 0.2 mm sieve, followed by drying for 2 days at a temperature of 70 °C and double sterilized at a temperature of 110 °C for 20 min in an autoclave. An assessment of its physicochemical parameters showed circumneutral pH (6.4 $\pm$ 0.1), electrical conductivity of 273.8 $\pm$ 1.2 µS/cm, moisture content of 2.5 $\pm$ 0.2% and water holding capacity of 265.4 $\pm$ 4.3%. The content of carbon, hydrogen, and oxygen was 76.9 $\pm$ 2.9%, 4.1 $\pm$ 0.3%, and 18.0 $\pm$ 2.7%, respectively. The Cu content in pure BC was 6.05 $\pm$ 0.50 mg/kg of dry weight (DW).

Metal-tolerant PGPR strain Buttiauxella sp. EA20 was used to prepare the biofertilizer (BF). Strain EA20 was grown overnight at 28 °C in 250 mL LB broth at 150 rpm in an orbital shaker incubator (Biosan ES-20/60, Latvia). The bacterial culture (10⁸ cfu/mL), prepared as previously described [17], was mixed with double sterilzed biochar (130 °C for 1 hour) and left overnight in a laminar flow chamber until the level of moisture reaches between 25–35%. The BF was stored in perforated disinfected bags at 27 °C for its use in pot-scale experiment [17].

In April and June of 2021, two separate pot-scale experiments were conducted with spring rapeseed plants, and the averaged outcomes are reported in this study. The B. napus seeds underwent surface sterilization by exposure to 70% ethanol for 30 seconds, followed by a 2-minute wash with 4% sodium hypochlorite, and ultimately, thorough rinsing with sterile double-distilled Millipore water.

The experiment comprises two distinct lines: one without copper (0Cu) and another involving the introduction of 200 mg of Cu (in sulfate form) per kilogram of soil (200Cu) on the 21st day of the plant growth period. Six treatment conditions were applied, namely: peat-containing control substrate (CS), CS + 5% BC (v/v); CS + 5% BF (v/v); CS + 200Cu (200 mg Cu kg^-1 soil); CS + 200Cu + 5% BC (v/v); CS + 200Cu + 5% BF (v/v).

Double sterilized (130 °C for 20 min) peat-containing substrate with pH 6.7 $\pm$ 0.1, EC 714.5 $\pm$ 1.3 µS/cm and Cu content 21.5 $\pm$ 0.7 mg/kg DW was used as a control. Fifteen B. napus seedlings were germinated in 350 mL plastic pots (five replicates for each treatment) in growth chambers with photoperiod, 14:10 (day:night), illumination, 180 $\pm$ 30 µmol/m² sec, temperature, 23 $\pm$ 2 °C and checked for seed germination after 7 days. On the 21st day, all treatments were undergone with the addition of 200 mg of Cu, except control. The extra plants were uprooted and only ten healthy seedlings in each pot were allowed to grow for 36 days in total. At harvest (on the 36th day), the plants were washed with tap water, followed by ultrasonication and finally twice with deionized Millipore water and divided into root and shoot. Half of the plants were harvested to access the biometric growth parameters whereas the rest were used to study the Cu content, biochemical and physiological characteristics of rapeseed.

Pre-dried and homogenized samples, weighing 0.2 g underwent wet digestion with 70% nitric acid (AR, analytical grade) in a MaRS 5 Digestion Microwave System (CEM, Matthews, NC, USA). The atomic absorption spectrometer (AAS) AA240FS (Varian Australia Pvt Ltd., Mulgrave, Victoria, Australia) was employed to determine the total Cu and potassium (K) content in both aboveground and underground rapeseed biomasses. JSC Ural Plant of Chemical Reagents (Verkhnyaya Pyshma, Russia) was used as Standard Reference Materials; GSS 7998-93 for Cu(II) and GSS 8092-94 for K(I) were utilized for the preparation, calibration and analysis of each analytical batch. Calibration coefficients were maintained at $\ge$0.99. The total nitrogen (N) and phosphorus (P) content in B. napus organs were measured at 440 nm and 640 nm, respectively, using a spectrophotometric method [27], following wet digestion with a 1:10 (v/v) mixture of perchloric and sulfuric acids. Total N was measured after reacting with Nessler’s reagent, while total P was determined using ammonium molybdate in an acidic medium.

The biometric parameters of plants such as shoot and root fresh and dry biomass, length, and leaf area were studied [17]. The water content in the organs was calculated by weighing the samples before and after drying at 75 °C for 24 hours. The germination rate of seedlings was also computed following the method reported by Wijewardana et al. [36].

The CO₂ assimilation (µmol CO₂/m² sec) on the 4th leaf (excluding cotyledons) of rapeseed plant was measured in a LeD Light Source chamber (3 $\times$ 4 cm) using a LI-COR (portable infrared gas analyzer LI-6400XT, Lincoln, NE, USA) at a saturating LED light intensity of 1200 µmol/m² sec, with 23 °C chamber temperature, and 50% humidity. The content of chlorophyll a (Chl a), chlorophyll b (Chl b) and carotenoids (Car) were calculated and determed using PD-303 UV spectrophotometer (Apel, Saitama, Japan) at wavelengths of 470, 647, and 663 nm, respectively, and calculated according to Lichtenthaler [37]. Water saturation deficit was measured using the floating discs’s method as described by Maleva et al. [38].

Malondialdehyde (MDA), a marker of lipid peroxidation, was quantified in 0.3 g of fresh rapeseed leaves. The leaves were homogenized with 4 mL of a reaction medium comprising 0.25% thiobarbituric acid in 10% trichloroacetic acid (v/v) as per Heath and Packer [39]. After boiling the extract for 30 min, cooling it on ice, and centrifuging it at 12,000 $\times$g for 15 min, the absorbance of the supernatant was measured at 532 nm and 600 nm. The concentration of TBA-reactive products was determined using the MDA extinction coefficient (155 mmol/cm) and expressed as nmol MDA per gram of dry weight (nmol MDA/g DW). The free proline content was assessed after boiling 0.4 g of freshly crushed rapeseed leaves in water for 15 min [38]. The reaction medium was prepared by adding prepared filtered extract with an amalgamation of glacial acetic acid with ninhydrin reagent in equal proportions (1:1:1; v/v), was placed in a boiling water bath for 30 min for staining and then cooled rapidly on ice. The proline content was determined spectrophotometrically by PD-303 UV (Apel, Saitama, Japan) at a wavelength of 520 nm and calculated as mg proline/g DW. The proline was used for preparing the standard curve (Sigma-Aldrich Chemie GmbH, Taufkirchen, Germany).

The total content of soluble phenolic compounds and flavonoids was determined in an ethanol extract from freshly crushed rapeseed leaves (0.3 g) pre-filled with 80% ethanol for 24 h. The resulting extract was filtered through the suitable size filter paper and analyzed. The total phenolic content was determined by spectrophotometer (Infinite 200 PRO, Tecan Austria GmbH, Grödig, Austria) at 760 nm after incubating the reaction mixture for 1 h (0.1 mL of extract interacting with 0.5 mL of 0.2 M Folin–Ciocalteu reagent for 5 min, followed by 0.4 mL 7.5% Na₂CO₃ (w/v) solution) at room temperature [40]. Gallic acid sourced from Sigma-Aldrich Chemie GmbH in Germany served as the reference standard in the analysis, with the outcomes reported as milligrams of gallic acid/g DW.

The content of flavonoids was measured by reacting the extracted sample with an equivalent amount of 10% aluminum chloride ethanolic solution followed by a 15 min incubation period at room temperature [38]. The absorbance of the reaction mixture was measured by spectrophometer (Infinite 200 PRO, Tecan Austria GmbH, Grödig, Austria) at 415 nm. The 10% aluminum chloride in the blank was replaced with an equal volume of 80% ethanol. Likewise, a standard solution of rutin from Sigma-Aldrich Chemie GmbH (Taufkirchen, Germany) was treated with aluminum chloride to establish a calibration curve, and the findings were presented as milligrams of rutin/g DW.

The data presented in the Table 1 represent the mean values (Means) of 3 replicates for measurements of PGP-attributes (IAA, siderophore production and phosphate solubilisation). Fig. 1 shows the bootstrap value from 1000 replicates. Fig. 2A demonstrates the appearance of 36-day-old rapeseed plants grown under various treatments. Fig. 2B and Table 2 represent the mean values (Means) from 5 replicates, composed of composite samples from each pot. Fig. 3 represent the Means obtained from 20 replicates, each consisting of four randomly selected plants from every pot, for biometric characteristics. Figs. 4,5,6 represent the Means of 5 composite replicates. Standard errors (SE) are included. Statistical analysis was conducted using STATISTICA 13.0 (StatSoft, Palo Alto, CA, USA) and Excel 16.0 software (Microsoft Office 2016, Microsoft, Redmond, WA, USA), following verification of normality using the Shapiro–Wilk test and homogeneity of variance using Levene’s test. Differences between treatment groups were assessed using the non-parametric Kruskal–Wallis H-test at a significance level of p 0.05. Significant differences between treatments are denoted by distinct lowercase and uppercase alphabetical letters.

Fig. 2.

Effect of 5% biochar (BC) and biofertilizer (BF) based on PGPR Buttiauxella sp. strain EA20 and BC on plant appearance and copper accumulation in shoot and root of B. napus after 36 days of vegetation without Cu (0Cu) and with the addition of 200 mg Cu/kg of soil (200Cu). (A) Plant appearance. (B) Copper content. CS, control substrate. Data are presented as Means $\pm$ SE (n = 5). Significant differences between treatments are denoted by distinct lowercase (a–d) and uppercase (A–C) alphabetical letters at p 0.05 according to the Kruskal–Wallis H-test. DW, dry weight.

Fig. 3.

Effect of 5% biochar (BC) and biofertilizer (BF) based on PGPR Buttiauxella sp. strain EA20 and BC on biometric growth parameters of B. napus after 36 days of vegetation without Cu (0Cu) and with the addition of 200 mg Cu/kg of soil (200Cu). (A) Shoot and root fresh biomass. (B) Shoot and root dry biomass. (C) Shoot and root length. (D) Leaf area. CS, control substrate. Data are presented as Means $\pm$ SE (n = 20). Significant differences between treatments are denoted by distinct lowercase (a–d) and uppercase (A–C) alphabetical letters at p 0.05 according to the Kruskal–Wallis H-test.

Fig. 4.

Effect of 5% biochar (BC) and biofertilizer (BF) based on PGPR Buttiauxella sp. strain EA20 and BC on photosynthetic characteristics of B. napus after 36 days of vegetation without Cu (0Cu) and with the addition of 200 mg Cu/kg of soil (200Cu). (A) Rate of CO₂ uptake. (B) Photosynthetic pigment content. CS, control substrate; Chl a, chlorophyll a; Chl b, chlorophyll b; Car, carotenoids. Data are presented as Means $\pm$ SE (n = 5). Significant differences between treatments are denoted by distinct lowercase (a–c and a–c) and uppercase (A–D) alphabetical letters at p 0.05 according to the Kruskal–Wallis H-test.

Fig. 5.

Effect of 5% biochar (BC) and biofertilizer (BF) based on PGPR Buttiauxella sp. strain EA20 and BC on the content of malondialdehyde (MDA) and water saturation deficit in the leaves of B. napus after 36 days of vegetation without Cu (0Cu) and with the addition of 200 mg Cu/kg of soil (200Cu). (A) MDA content. (B) Water saturation deficit. CS, control substrate. Data are presented as Means $\pm$ SE (n = 5). Significant differences between treatments are denoted by distinct uppercase (A–C) alphabetical letters at p 0.05 according to the Kruskal–Wallis H-test.

Fig. 6.

Effect of 5% biochar (BC) and biofertilizer (BF) based on PGPR Buttiauxella sp. strain EA20 and BC on the content of low molecular weight antioxidants in the leaves of B. napus after 36 days of vegetation without Cu (0Cu) and with the addition of 200 mg Cu/kg of soil (200Cu). (A) Free proline. (B) Total soluble phenolic compounds and flavonoids. CS, control substrate. Data are presented as Means $\pm$ SE (n = 5). Significant differences between treatments are denoted by distinct lowercase (a–d) and uppercase (A–D) alphabetical letters at p 0.05 according to the Kruskal–Wallis H-test.

From the 30 strains isolated from the rhizosphere of the orchid E. atrorubens, one particular isolate i.e., strain EA20, exhibited the highest resistance to metals such as Cu, Ni, Zn, and Cd. EA20 also demonstrated exceptionally positive results across all tested plant growth-promoting attributes, including N₂-fixation, IAA production, phosphate solubilization, and siderophore production. The preliminary identification was performed by assessing its morphological characteristics. The isolate was identified as Gram-negative and aerobic and the pure colony exhibited a smooth, crateriform elevation, light beige color, and a round shape with a rolled appearance around the edge.

The full-length sequence of the 16S rRNA gene was used for genus identification. Utilizing the cloud-based platform EPI2ME Agent, the strain’s genus was determined to be Buttiauxella with an average accuracy of 91.8%. Additionally, a phylogenetic investigation revealed that strain EA20 was situated in a shared cluster with strains from the Buttiauxella genus, including B. izardii, B. noackiae, and B. warmboldiae. This supports the conclusion that strain EA20 certainly falls within the Buttiauxella genus (Fig. 1).

The bacterial strain Buttiauxella sp. EA20 showed resistance against various metals. The strain EA20 exhibited maximum tolerance for Zn = Cd Cu Ni with a range of 1000 600 400 mg/L, respectively (Table 1). The quantitative analysis of phosphate solubilization and IAA synthesis in a liquid medium revealed that the strain EA20 solubilized 110.7 mg/L of insoluble tri-calcium phosphate and produced 12.7 mg/L of IAA. The qualitative analysis of siderophore synthesis was revealed by a clear halo zone around the colony on a CAS-agar plate. N₂-fixation analysis also revealed a positive result for the strain which upsurges the nitrogen assimilation ability of plants (Table 1).

The application of BC or BF showed no significant effect on the germination of B. napus seeds, which averaged 76%. Fig. 2A shows the appearance of B. napus plants grown under different treatments at the end of the experiment. It can be visually seen that the appearance of rapeseed shoots was much better in the variants with the addition of 5% BC or BF, regardless of copper treatment, compared to the control substrate (CS). The addition of 200 mg/kg of Cu into CS resulted in a 2.4-fold increase in B. napus shoot content and a 9.4-fold increase in the root, corresponding to 18 and 79 mg/kg DW, respectively (Fig. 2B).

A similar trend was observed in the plants treated with 5% BC. The application of BF without Cu did not affect its content in the shoot, rather a 2-fold enhanced accumulation was observed in the root (compared to CS and 5% BC). When copper was applied (BF + Cu), it mainly accumulated in rapeseed root (4.6 times higher than in shoot). The addition of BF also contributed to an increase in the accumulation of Cu in shoot and root by 3.3 and 13.3 times, respectively, as compared to CS or BC without Cu, and by 1.4 and 1.5 times as compared to CS or BC with 200Cu addition (Fig. 2B).

The use of BC without copper did not significantly affect the phosphorus content, however slightly increased the nitrogen and potassium content in the shoot (15%), while there were no significant differences observed for NPK concentration between CS and BC in root (Table 2). Simultaneously, plants treated with BF demonstrated 1.4-fold higher NPK content in their shoot and root as compared to CS. The combined application of 200Cu and BC increased NPK content in the shoot by 20% compared with single copper. Wherein, the nitrogen content in the root increased by 15%, but the phosphorus and potassium content remained unchanged. The application of Cu and BF resulted in a significant increase of 1.5 times on average in NPK content in both aboveground and underground organs (Table 2).

Both in control and BC treated plants, the addition of Cu showed a negative effect on the growth parameters of rapeseed (Fig. 3). An average decrease of 18% was observed for fresh and dry biomass of shoot while root biomass was reduced by 35% as compared to CS and CS + 5% BC treatments (Fig. 3A,B). Moreover, a significant decrease of 28% (on average) in the shoot and root length as well as leaf area was also observed (Fig. 3C,D). In the absence of Cu, BF enhanced the aboveground and underground fresh biomass by an average of 1.9 times as compared to control and only BC treated plants (Fig. 3A,B). Simultaneously, shoot and root length did not change (Fig. 3C), while the leaf surface area increased significantly (by 47%) compared to CS and BC (Fig. 3D). When Cu was applied together with BF, the fresh shoot biomass was increased almost by 2-fold (Fig. 3A), while the dry shoot biomass upsurge only by 30% (Fig. 3B), compared to CS and CS + Cu. At the same time, an almost quadruple increase in underground biomass for both fresh and dry was observed (Fig. 3A,B). The addition of BF along with Cu did not have a significant effect on the shoot and root length compared to the CS, however, it was 1.4 times higher than CS + Cu (Fig. 3C). Simultaneously, the leaf area increased by 1.4 and 1.7 times compared to CS and CS + Cu, respectively (Fig. 3D).

The rate of CO₂ assimilation in the control plants decreased by 40% in the presence of copper (Fig. 4A), while the content of photosynthetic pigments remained more stable (Fig. 4B). Similar trend was observed for 5% BC treated plants. The addition of BF (both with and without copper) had a positive effect on photosynthetic activity and pigment content in rapeseed leaves which increased them by 1.4- and 1.8-fold, respectively (Fig. 4).

The application of copper contributed to a significant (37%) increase in the MDA content of control rapeseed plants (Fig. 5A). In BC + Cu and BF + Cu treated plants, the MDA content increased to a lesser extent (by 17% from CS on average). In the presence of copper, a significant increase of 25% in water saturation deficit was observed for control plants (Fig. 5B). BC and especially BF caused a significant decrease in water deficit after its application.

The content of free proline in rapeseed leaves significantly increased (by 30% on average) when BF was applied, especially in the presence of copper (Fig. 6A). The Cu addition led to a significant increase in the content of soluble phenolics in the CS-plants, and especially (almost 1.5 times) flavonoids (Fig. 6B). The application of BF also activated the synthesis of soluble phenolics, while the proportion of flavonoids increased by almost 2-fold compared to the control. The copper in the presence of BF stimulated the synthesis of phenolics even more, however the proportion of flavonoids decreased.