Introduction

In recent years, due to the continuous industrialization and various other anthropogenic activities, soils have been polluted to the varying degrees with potentially toxic elements, i.e., heavy metals (HMs), and become a repository for the environmentally released pollutants. More than 10 million soil sites are considered contaminated with priority HMs worldwide (Lwin et al. 2018). In Russia, high concentrations of HM were found in soils near nonferrous metals mining and processing companies (Gorovtsov et al. 2019a; Minkina et al. 2018, 2019). HMs are known for their toxicity, persistence in the environment and for bioaccumulation and bio/geo-transformation in natural ecosystems (Rajput et al. 2019). The excess HMs present in soil could affect plant performance, the activity of soil microbial community. Water bodies can be polluted through leaching of the HM to groundwater, causing a threat to aquatic life and human health. Recent studies showed high accumulation of HMs in tissues of native plants grown in the polluted soils of mining areas (Ghazaryan et al. 2018; Minkina et al. 2019; Rajput et al. 2018a). HM accumulation in edible crops is an issue of global concern due to its relation with human health (Onakpa et al. 2018). Previous studies indicate that the HMs also affected plant growth by changing root morphology, root/shoot lengths, ultrastructure of cellular and subcellular structures and changes in physiological state of plants (Rajput et al. 2018a, b; Wang et al. 2017b). Therefore, the issues related to soil pollution must be urgently addressed.

Chlorophyll fluorescence is a noninvasive measurement for the photosynthetic performance of plants. The maximal quantum yield of photosystem II (Fv/Fm) is one of the most important parameters of chlorophyll fluorescence and has been used as a nondestructive and accurate tool to precisely determine the physiological state of plants (Henriques 2009). Chlorophyll fluorescence is useful to monitor the interaction between soil pollutant and plants (Zurek et al. 2014).

Bioremediation is considered a better option for reclaiming the contaminated soils (Ali et al. 2013; Wuana and Okieimen 2011). Another approach is the immobilization of HMs in soil by addition of various sorbents that prevent the translocation of metals from the soil solution to plants and their leaching into the underlying soil horizons. One of the most promising types of sorbents applicable in remediation is carbonaceous sorbents and specifically biochar (Lim et al. 2019).

Biochar is a carbon-rich material that helps to stabilize the mobility and bioavailability of the metal in the soil (Palansooriya et al. 2019; Wang et al. 2017a). The porous structure of biochar contributes to its large surface and sorption efficiency. Furthermore, the application of biochar could change the soil physicochemical properties and enhance the activities of soil microorganisms that influence soil quality and plant growth (Lwin et al. 2018; Palansooriya et al. 2019). The studies of remediation process are important for better understanding the interactions of biochar with soil, as well as with plants. Several microorganisms, especially bacterial species, are successfully used for bioremediation of HMs due to their significant role in biodegradation of pollutants. Also, microbes are involved in many beneficial soil functions, such as nutrient recycling, organic matter decomposition, soil-structure formation, secretion of plant growth promoters and degradation of organic contaminants (Gorovtsov et al. 2018). Thus, the combined use of biochar and HM-tolerant microbes may be an effective tool for soil remediation.

The territory of south Russia, especially Rostov Region, is the largest producer of spring barley (Hordeum vulgare L.), one of the food grain crops highly important for industrial uses. The study indicated that the H. vulgare L. is an efficient HMs accumulator and could accumulate HMs equal to Indian mustard (Brassica juncea), a hyperaccumulator plant (Ebbs and Kochian 1998). Most studies on plant–metal interactions are focused on the processes involved in metal ion uptake and accumulation in the harvestable parts of the crop as well as on metal-induced phytotoxicity. However, there are limited studies available on the influence of biochar–microbes interaction to plant growth. The aim of this work was to study the biochar and metal-tolerant bacteria application efficiency in polluted soil and their influence on morpho-physiological and anatomical parameters of spring barley.

Materials and methods

Description of used soils and their chemical and physical properties

Soil samples from the top layer (0–20 cm depth) were collected in the vicinity of the Atamanskoe Lake and immediately transported to the laboratory. The high levels of pollution are due to wastes that were released into the lake by industrial enterprise in 1940–1980s. At present, the lake is dry, and active soil-forming processes continue on the surface of bottom sediments. Soil at the territory adjacent to the lake classified as Stagnic Phaeozem was taken as unpolluted soil. The polluted soil collected at the bottom of the dried lake was classified as Spolic Technosol. The physicochemical properties of these soils had the following physical and chemical properties: content of C org 1.9–2.6%, pH 7.4–7.9, physical clay 52–53%; CaCO3 1.3–5.3% exchangeable bases (cmol kg–1): Ca2+ 31.0–35.2%; Mg2+ 4.5–5.0%. The collected soil samples were analyzed by X-ray fluorescence using a MAKC-GV spectroscan. The unpolluted soil contents (mg/kg) Zn: 90.0, Mn: 230.4, Cr: 97.9, Cu: 43.7, Pb: 26.1, Ni: 40.9, Cd: 0.2, while polluted soil contents Zn: 62,032.1, Mn: 426.4, Cr: 154.1, Cu: 146.9, Pb: 1591.2, Ni: 80.5, Cd: 10.6 (Bauer et al. 2018).

Experimental setup and plant growth conditions

Twenty healthy seeds of barley without any further treatments were sown in each pot containing 2 kg soil. Unpolluted soils were considered as a control. Three HMs-tolerant spore-forming bacteria identified as Bacillus cereus, B. pumilus and B. atrophaeus (Gorovtsov et al. 2019a) were applied into polluted soil layer-wise at a dose of approximately 1010 CFU/kg soil to achieve the most uniform distribution of bacterial cells throughout the soil. Four variants were used: (1) control (unpolluted soil), (2) polluted soil; (2) polluted soil + biochar, (3) polluted soil + bacteria and (4) polluted soil + biochar + bacteria. The soil was added layer by layer into the pot, pouring each layer with a suspension of bacterial strains, evenly consuming soil and water. Pots were incubated (approximately for 1 week), before H. vulgare L. was sown. Industrially obtained wood biochar (2.5%) was used in the experiment with the surface area: 500–800 m2/g, pore width: 8–12 nm and sorption capacity: 200–500 g/kg.

All the pots were placed in the laboratory, with the temperature ranging from 25 ± 5 °C (day) to 18 ± 2 °C (night) at normal lighting mode in aerobic conditions. The moisture was maintained by adding distilled water through vertical pipes from the bottle to the top in the center of pot to avoid leaching of metal. After 2 weeks of growth, plants were thinned to 15. No additional fertilizers and pesticides were applied during the experiment. Each treatment was prepared in triplicate.

Morphometric observations and heavy metals analysis in plant tissues

The morphometric parameters, i.e., shoot, leaf and root lengths, were measured after 60 days of growth. Plant height was estimated by measuring the length from the bottom of the main stem to the end of the emerging third leaf. Primary root length was determined by measuring the mean length of the longest roots. Differences in these measurements between the control and the contaminated plants were used to evaluate metal toxicity.

Roots, stems and leaves were separated, washed thoroughly with distilled water, and oven-dried at 65 °C ground and passed through a 0.25-mm sieve for HMs analysis. The preparation of plant tissues for heavy metals determination was performed by burning the 1 g air-dried plant sample at 450 °C for 6 h. Then, the ash was dissolved in 5 ml of 20% HCl and filtered through 0.45 µm Whatman filter paper (Minkina et al. 2019). The analysis was performed by atomic absorption spectrophotometer (AAS) (KVANT 2-AT, Kortec Ltd, Russia) at room temperature with a wavelength diapason: for Zn—213.9 nm, Mn—279.5 nm, Cr—357.9 nm, Cu—327.4 nm, Pb—283.3 nm, Ni—352.1 nm, Cd—228.8 nm.

Preparation of samples for anatomical and ultrastructural observation

Preparation of plant tissue samples for transmission electron microscope (TEM) was carried out according to the method described by Fedorenko et al. (2018). Two millimeter-sized samples were collected from the middle part of the plant leaf and fixed using 2.5% glutaraldehyde/0.1 M phosphate buffer solution (PBS) at room temperature for 2 h. After washing with PBS, samples were incubated for 1 h in 1% OsO4/0.2 M PBS solution and then dehydrated by increasing the concentration of ethanol and acetone (separately 50, 70, 96 and 100%). After dehydration, samples were embedded in Epon-812 embedding medium. After polymerization of the obtained blocks, ultrathin sections were prepared by microtome (Leica EM UC6, Germany), which were further contrasted in lead citrate and examined under the light-optical microscope (Mikmed-6 St. Petersburg, Russia) and TEM (Tecnai G2 Spirit Bio TWIN, the Netherlands). For the light-optical microscope, semi-thin sections (about 1 μm thick) were prepared and additionally stained with 1% toluidine blue solution (O’Brien et al. 1964), and examined with Micmed-6 microscope (LOMO, Russia).

Measurements of chlorophyll fluorescence and transpiration rate

Plants were dark-adapted in the laboratory room with dark curtains and no artificial light source for 20 min before chlorophyll fluorescence measurements. During the measurement, the room temperature was the same as the growth environment of plants. The maximal quantum yield of photosystem II (Fv/Fm) was measured using pulse amplitude-modulated (PAM) fluorometer (Diving PAM, Waltz, Germany). Photosynthetic photon flux density (PPFD) of red measuring light was 0.15 mmol photons m−2 s−1, and PPFD of white saturating flash was 1800 mmol photons m−2 s−1. Fifteen readings were recorded by selecting fresh leaves, and all the leaves were measured under the same conditions.

Transpiration rate was evaluated gravimetrically measuring leaf mass loss using balances (Adventurer™ Pro, Ohaus) of 10 µG sensitivity connected to the computer and calculated as:

$$\frac{\Delta F}{\Delta t}$$

where F denotes the difference between initial and final weight and t indicates time of weighing which was in min.

Statistical analyses

The multiple comparisons between the groups were performed with the use of one-way ANOVA. Tukey’s honestly significant difference (HSD) was used as a post hoc test. All the calculations were made with the Statistica 12 software.

Results

Accumulation of heavy metals in plant tissues

The growth of H. vulgare L. in polluted soil caused the accumulation of the HMs in the range of 2.79–53.95 and 1.96–30.87 mg/kg in above- and belowground parts of the plant, respectively (Figs. 1, 2). The application of biochar and HM-tolerant bacteria decreased the HMs content by 1.78–9.23- and 1.79–5.30-folds than the control, respectively, while the addition of both biochar and bacteria has led to a decrease in HMs content by 3.11–17-folds and was the most effective to decrease the translocation of all the studied metals, especially Pb.

Fig. 1
figure 1

Heavy metals accumulation in roots of barley

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Fig. 2
figure 2

Heavy metals accumulation in aboveground tissues of barley

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Morpho-physiological parameters of spring barley

The poor growth of H. vulgare L. was observed in polluted soil, and it was recorded that the root length decreased by 46.74% and shoot length decreased more than twice (Table 1). The biochar alone was unable to improve the plants’ morphometric parameters, except for the shoot length that was increased by 18.4% compared to polluted variant. However, the addition of HM-tolerant bacteria influenced the H. vulgare L. growth to a large extent. The addition of both biochar and bacteria had even more pronounced effects. Use of biochar and bacteria had no additional benefit for the root and stem lengths when compared to the use of bacteria alone. The contrary was observed for leaves length and dry weight of plants: The parameters were significantly higher and reached those of unpolluted control.

Table 1 Influence of soil pollution on root, shoot and leaves length and dry weight of barley
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The growth of H. vulgare L. in contaminated soil had adverse effects on both transpiration and photosynthetic performance. The results showed that soil contamination suppressed maximal quantum yield of photosystem II in H. vulgare L. compared to plants grown on unpolluted soil. At the same time, the addition of biochar, bacteria and biochar + bacteria improved the photosynthetic performance compared to the plants grown on polluted soil. The maximal quantum yield of photosystem II decreased 8.32% in plants grown in contaminated soil compared to the control, whereas this decrease was less in biochar (7.17%) and bacteria (5.83%) and in bacteria + biochar applied together (4.72%) (Fig. 3). As for the transpiration rate, the plants grown in polluted soil have shown a decrease in transpiration by 25.78% compared to control. The addition of biochar did not improve the transpiration rate (> 1% difference from polluted variant). Application of bacteria yielded an insignificant 3.28% improvement, compared to polluted variant. At the same time, the simultaneous application of biochar and bacteria has led to a significant 14.28% improvement of the transpiration rate, compared to polluted variant. This variant was still significantly lower than unpolluted control (15.18%).

Fig. 3
figure 3

Performance of photosynthetic system measured by chlorophyll fluorescence

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Anatomical and ultrastructural parameters of barley

Light-optical observations of control plants showed that hypostomatic leaves contain a single-row epidermis, columnar and spongy parenchyma, as well as vascular bundles in the middle part of the leaf blade (Fig. 4a). In the leaves of plants grown on contaminated soil, there was a violation of the integrity of the epidermal layers. The number of cells in them was reduced, while the intercellular space of the parenchyma was increased. An increase in the optical density of chlorenchyma cells and a violation of their ordered arrangement were noted. The thickness of the leaf blade (adaxial to abaxial cells) was significantly decreased (Fig. 4b). These changes (described for control plants) are less pronounced when biochar (Fig. 4c) or bacteria (Fig. 4d) are added to the soil. Even less differences in the structure of the leaves, compared with the control, were observed during their simultaneous application (Fig. 4e). In this variant, the thickness of the leaf blade reached control values; however, the increased optical density of individual chlorenchyma cells remained.

Fig. 4
figure 4

Light-optical observation of barley leaf sections. Ep epidermis, VB vascular bundles, scale bar 100 µm

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The ultrastructure of the control plants leaf mesophyll cells was typical of cereal plants in the control variant. In chloroplasts of the correct lenticular form, the developed thylakoid system, immersed in a fine-grained stroma, was clearly visible. The thylakoid system is represented by large grains containing densely packed lamellae, up to 30 units or more. Between the separate thylakoids of stroma, there are scattered plastoglobules with a diameter of up to 0.1 µ and large starch grains with a diameter of up to 0.5 µ. In rounded mitochondria (with a diameter of 0.6 µ), drop-shaped cristae were noticeable, and small rounded peroxisomes (with a diameter of 1.1 µ) had a uniform and denser structure than mitochondria (Fig. 5, 1a, b). The control plants are characterized by the preferential localization and grouping of these organelles (mitochondria and peroxisomes) directly near chloroplasts. Numerous ribosomes were evenly distributed throughout the cell cytoplasm.

Fig. 5
figure 5

Transmission electron micrographs of barley leaf ultrathin cross section micrographs in sub-fig. (1) a, b—control, (2) a, b—pollution, (3) a, b—pollution + biochar, (4) a, b—pollution + bacteria and (5) a, b—pollution + biochar + bacteria. In figure, subcellular organelles are indicated as: Cl chloroplast, CW cell wall, ER endoplasmic reticulum, GT granal thylakoids, M mitochondria, N nucleus, Pg plastoglobule, Pr peroxisome, SG starch grain, ST stromal thylakoids, V vacuole, LD lipid droplet. The scale bars (in μm) for the sub-figures were 1 (a—1, b—2, insert—0.20), 2 (a, b—1), 3 (a—1, b—0.25, insert—0.25), 4 (a—1, b—0.5); and 5 (a—1, b—0.5, insert—0.25)

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Significant changes in the ultrastructure of cellular organelles of chlorenchyma were noted in plants grown on contaminated soil. In particular, the chloroplasts of mesophyll cells looked more rounded, their matrix density was slightly increased and the thylakoid system was less visible in it (Fig. 5, 2a, b). The grana are reduced in size, and the number of lamellae in them is about 15–20. The size of the plastoglobules almost doubled and reached about 0.2 microns, while their electron density increased significantly. The shape of most mitochondria was elongated (Fig. 5a, b), and mitochondrial cristae looked swollen. Peroxisomes contained a matrix that was heterogeneous in electron density. The cytoplasm of most cells was almost completely devoid of ribosomes, and the elements of the endoplasmic reticulum were vacuolated.

When biochar was added (to the soil), the density of the matrix of plastids and mitochondria increased, while no significant changes in their shape and size were observed (Fig. 5, 3). The number of grana increased, and the number of lamellae in them was up to 25 (15–20). Compared with the control, a decrease in the size of mitochondria and peroxisomes was noted (0.3 and 0.5 microns, respectively). The starch-formation process has intensified (an increase in the size of starch grains). (Figure 5, 3a) The number of plastoglobules increased, although their size somewhat decreased to 0.12 μm. The cytoplasm of some cells contained a few ribosomes, and the appearance of small lipid droplets with a diameter of up to 0.3 (0.2) μm was noted (Fig. 5, 3b).

The state of the plastids ultrastructure in plants under the action of only bacteria is generally similar to the effect of biochar (Fig. 5, 4a, b). With a complex treatment (biochar + bacteria), a further normalization of the ultrastructure was noted (Fig. 5, 5a, b). In some chloroplasts, numerous outgrowths and invaginations of large sections of the cell cytoplasm deep into the stroma appeared (Fig. 5, 5a). The invaginations of cytoplasm into the chloroplast increase its surface, and as a result, the interaction area between the contents of the organelle and the cytoplasm increased significantly. This may be one of the ways to optimize cell metabolism under metal stress conditions. No other significant differences from control samples were found in the fine structure of plant cell organelles.

Discussion

In the present study, accumulation of HMs in the aboveground tissues of H. vulgare decreased in all treatments as compared to the variant with polluted soil. Application of both the biochar and HM-tolerant bacteria and both individual treatments has led a decrease in accumulation of HMs in the aboveground tissues. In this connection, it is interesting that biochar from plant waste adsorbs Pb from the contaminated soil that leads to an increase in the soil microbial biomass and an increase in the enzymatic activity (Igalavithana et al. 2017; Gorovtsov et al. 2019b). The study conducted on wild plant species in Cu-contaminated soils of a mining region indicates high Cu accumulation in roots (dry weight) of Thymus kotschyanus (775 mg/kg) and in shoots of Phleum pratense (243 mg/kg) (Ghazaryan et al. 2018). A higher accumulation of HMs was observed in roots compared to the shoots in a number of plant species grown in a native environment of the iron and copper mining areas (Nouri et al. 2009). In turn, it should be considered that uptake of metal ions is largely influenced by their bioavailability in soil and plant. The mechanisms of metal uptake, translocation, compartmentation, toxicity and detoxification differ greatly with soil and plant type, chemical speciation of metals in soils, soil particle size, cation-exchange capacity, soil pH, organic matter content and microbial activity (Nouri et al. 2009). In addition, HMs distribution in the above- and belowground parts of plants also depends on plant family (Chaplygin et al. 2018).

Application of biochar could decrease HMs accumulation. Thus, it was found that the initial accumulation levels of Cd, Cu, Pb and Zn (79–66, 13–19, 87–86 and 37–36%, respectively) were decreased with application of husk and maple leaves biochar-treated soils (Derakhshan and Jung 2017). Despite the decrease in HMs accumulation capacity of biochar, the simultaneous impacts of biochar and microbes are significant. The application of biochar could provide a suitable habitat for native or applied microorganisms and enhance their activity in soil and reduce the HMs uptake to plant. The application of biochar significantly improved the relative abundance of two soil bacteria (Sphingomonas and Pseudomonas) under cotton cropping system (Han et al. 2017). It was also reported that application of biochar increases the microbial activity and biomass (Lehmann et al. 2011).

Morphometric observations indicated a biochar and bacteria improved the root and shoot growth of H. vulgare L. grown in polluted soil (Gorovtsov et al. 2019a). However, in combination, biochar and bacteria had no additional benefit for the root growth. Another study also showed that the addition of biochar to soil increased shoot biomass, while it did not increase the root biomass Oryza sativa and Solanum lycopersicum (Anyanwu et al. 2018).

The maximal quantum yield of photosystem II is an indication of dissociation of light-harvesting pigment systems of PSII. The maximal quantum yield of photosystem II was recorded in H. vulgare L. grown in polluted soil. However, biochar amendment with bacteria has improved photosynthetic activity compared to individual application of bacteria and biochar. The results of one-way ANOVA show that the combined treatment with biochar and bacteria led to a significant improvement of the maximal quantum yield, while in their separate application the increase was not significant. However, none of the treatments was able to reduce the toxic effect to the control level. The reduction in H. vulgare L. growth and biomass could be a reason for the inhibition of photosynthesis (Dong et al. 2005; Shanmugaraj et al. 2019). The improvement in H. vulgare L. growth may be due to increases in water retention by application of the biochar (Abdalla 2011; Günal et al. 2018). Similarly, other study showed that the application of 5% rice straw biochar decreased the bioavailability of HMs and improved microbial counts and soil enzyme activities (Li et al. 2019).

Significant changes in cellular organelles caused by soil contamination and the subsequent normalization of their ultrastructure when biochar and bacteria were added to the soil were found in this study. These structural changes in barley leaves (in the case of contamination) are generally similar to the data obtained in previous light-optical and electron microscopic studies of plants grown on contaminated soils (Minkina et al. 2019; Koushi et al. 2016; Wang et al. 2017b). At the same time, the authors noted mitochondrial cristal swelling, structureless cytoplasm, few and fragmented peroxisomes, impaired tonoplast integrity, granoplast formation disruption in chloroplasts. The revealed destructive changes in these organelles during soil contamination are obviously associated with a decrease in the level of metabolic processes that ensure plant growth. The subsequent normalization of the ultrastructure when biochar and bacteria are added to contaminated soil, on the contrary, activates cellular metabolism and leads to an improvement in the morphometric characteristics of plants.

Changes in the chloroplast level in leaf mesophyll cells during contamination were noted by other authors in the leaves of O. vulgare (Panou-Filotheou et al. 2001; Taylor and Foy 1985). The same observations, as well as the complete disorganization of these organelles, were characteristic for Pistia stratiotes (Upadhyay and Panda 2009). Ouzounidou et al. (1992) working with Thlaspi ochroleucum, and Panou-Filotheou et al. (2001) with O. vulgare, confirmed the disappearance of starch grains. According to these authors, with a decrease in the number and volume of chloroplasts, a decrease in the chlorophyll content and a subsequent decrease in photosynthesis occurred due to excess copper. In p experiments, an increase in the size of starch grains was observed in the variants after adding biochar and bacteria to contaminated soil. In the variants with soil contamination, “swelling” of plastids (manifested in their rounding) was observed along with an increase in their size. Some authors view this effect as a marker of external influence (Kaplan and Guy 2004; Trunova 2007; Vella et al. 2012). As a rule, changes of this kind are accompanied by an increase in the density of the stroma and indicate rapid rearrangements in its chemical composition, including enhanced synthesis of proteins and enzymes. These changes can act as an indirect indicator of the intensification of enzymatic processes (Kratsch and Wise 2000).

At the same time, a decrease in the level of the process of grana formation is shown; a reduction in the number of lamellae in grana; the expansion of inter-thylakoid space; and a decrease in the total number of grana themselves in organelles. At present, the significance of such a structural transformation of chloroplasts remains not entirely clear. Nevertheless, it is the ability of fast membrane rearrangements, according to some authors, that is an important element of plant resistance to adverse factors (Hudak and Salaj 1999; Rajput et al. 2018b). The ability to “trigger” such changes in chloroplasts is considered one of the mechanisms that reduce the likelihood of photo-inhibition and optimize the process of photosynthesis under stressful conditions (Pribil et al. 2014).

A growing number of plastoglobules in plants due to soil contamination have also been observed in other studies (Ouzounidou et al. 1992; Panou-Filotheou et al. 2001). These structures, found in chloroplasts, function as active centers in the synthesis and recycling of proteins under stress (Ytterberg et al. 2006). Plastoglobules create a functional metabolic relationship between the inner membrane and thylakoid membranes and play a key role in the mechanism of protection against oxidative stress (Ytterberg et al. 2006). Thus, it can be assumed that an increase in the number of plastoglobules in leaf mesophyll cells is a protective mechanism against damage to the photosynthetic apparatus.

An important property of peroxisomes is their high plasticity, which allows changing the shape and localization in the cell under adverse conditions (Del-Rio et al. 2006; Rajput et al. 2018b; Reunov 2014). Peroxisomes are the main generators of reactive oxygen species (ROS), but on the other hand, they also participate in the neutralization of ROS by antioxidant enzymes (Nyathi and Baker 2006; Palma et al. 2009). Therefore, the changes in the shape, density and localization of peroxisomes noted in different experimental variants can be considered as an indicator of changes in the redox balance of cells (Rodriguez-Serrano et al. 2009). Numerous peroxisomes and mitochondria were found near the chloroplasts in all variants of contamination. This may indicate an increase in contacts between these organelles. The outgrowths and invaginations that appear in chloroplasts in this case contribute to an increase in the surface area of plastids and facilitate the exchange of metabolites with cytoplasm and other cell organelles (Buchner et al. 2007; Holzinger et al. 2007; Lütz and Engel 2007).

The above-described ultrastructural changes in plastids were accompanied with a significant decrease in quantum yield of photosystem II (Fv/Fm) observed in our work, which is widely used to evaluate phytotoxicity and environmental impact on plants being correlated with the efficiency of the thylakoid noncyclic electron transport as well as with the efficiency of oxygenic photosynthesis and plant health in whole (Murchie and Lawson 2013; Lazár 2015). The studies indicate the application of soil amendments improves the growth and photosynthetic pigments, helping plants to overcome from various biotic and abiotic stresses, especially metal stress (Abdalla 2011; Bello et al. 2014; Zhao et al. 2019). It is known that chloroplasts, mitochondria and peroxisomes are the main organelles of the glycolate photosynthesis cycle, which serves as a source of intermediate substances of the various syntheses providing plant growth (Kuznetsov and Dmitrieva 2005).

Conclusions

The growth of H. vulgare L. in contaminated soil caused the accumulation of heavy metals in both roots and aboveground tissues of the plants. The metal toxicity was manifested in all levels of plant organization. At the subcellular level, obvious changes in cell organelles ultrastructure were noted. The metal toxicity affected multiple organelles such as mitochondria, peroxisomes and the endoplasmic reticulum. The inner structure of the chloroplasts was also significantly altered. These ultrastructural changes caused changes in physiological state of H. vulgare L. Soil contamination suppressed maximal quantum yield of photosystem II and transpiration rate in H. vulgare L. compared to plants grown on unpolluted soil. At the tissue level, soil contamination damaged the integrity of epidermal layer of the leaves and caused an increase in the intercellular space of the parenchyma and changes in the mesophyll cells arrangement. At the organism level, the overall performance of the plants was reduced, and the main morphometric parameters were lower, compared to the control. The application of biochar, metal-resistant bacteria and especially both treatments together reduced the adverse effects of soil pollution, and these positive changes can be consistently traced at all levels of the organization of living matter, starting with cellular, subcellular structures. This improvement is most likely based on the reduction in metal bioavailability to the plants, which is supported by the reduced metal content in both above and belowground tissues of plants, grown in soil treated with biochar, bacteria or both. The combined use of biochar and bacteria outcompeted each of the treatments taken separately. This implies that there is an active mechanism of interaction between the metal-tolerant bacteria and biochar, leading to an improved immobilization of the HM. The use of biochar in combination with metal-tolerant bacteria is an efficient tool for remediation of soils, contaminated with heavy metals. However, the present level of knowledge is not sufficient to understand the mechanisms that take place in the rhizosphere in the presence of biochar and bacterial inoculum. The processes of interaction between the bacterial cells and biochar particles, taking place in the close vicinity of the plant roots, are yet to be studied.