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Article

Inducing Rhizosphere Acidification in White Willow with Bacillus sp. ZV6 Enhances Ni Phytoextraction from Soil and Soil Quality

by
Zaheer Abbas Virk
1,
Muhammad Zubair Yasin
2,
Sebam Gill
1,
Muhammad Fraz Ilyas
1,
Agnieszka Dradrach
3,
Saud Alamri
4,
Alanoud T. Alfagham
4,
Mohd Sayeed Akhtar
5 and
Muhammad Iqbal
1,*
1
Department of Environmental Sciences, Government College University, Faisalabad 38000, Pakistan
2
Department of Emergency, Aziz Fatimah Hospital, Faisalabad 38000, Pakistan
3
Institute of Agroecology and Plant Production, Wrocław University of Environmental and Life Sciences, 50-363 Wrocław, Poland
4
Department of Botany and Microbiology, College of Science, King Saud University, Riyadh 11451, Saudi Arabia
5
Department of Botany, Gandhi Faiz-e-Aam College, Shahjahanpur 242001, India
*
Author to whom correspondence should be addressed.
Minerals 2023, 13(9), 1178; https://doi.org/10.3390/min13091178
Submission received: 19 July 2023 / Revised: 23 August 2023 / Accepted: 30 August 2023 / Published: 8 September 2023
Browse Figures
Versions Notes

Abstract

:
Chelating agents may decrease the extent of Ni phytoextraction by reducing plant growth and soil health due to Ni toxicity during enhanced phytoextraction. Contrarily, inducing acidity in the rhizosphere of Ni-accumulating plants with plant growth-promoting rhizobacteria (PGPR) having rhizosphere acidification ability can enhance Ni phytoextraction by increasing Ni bioavailability in the soil, plant growth, and plant stress tolerance. We investigated the efficacy of a PGPR species with rhizosphere acidification potential, named Bacillus sp. ZV6 (ARB), in enhancing Ni phytoextraction by white willow (Salix alba) from a Ni-affected soil. The plants were grown for 120 days in soil with zero, threshold, and moderate Ni pollution levels (0, 50, and 100 mg Ni kg−1 soil, respectively) with and without ARB inoculation. After harvest, the effects of the treatments on rhizosphere acidification and associated Ni bioavailability in this zone, Ni distribution in plants, and Ni removal from the soil were investigated. Moreover, enzyme activity, count of bacteria, biomass of microbes, and organic C in the soil, together with indices of plant growth and antioxidant defense, were evaluated. The ARB inoculation significantly improved the plant parameters and soil health and reduced plant oxidative stress at each Ni level compared to the treatments lacking ARB. Besides lowering the soil pH and increasing Ni bioavailability in the rhizosphere with respect to the bulk zone, ARB inoculation exerted additional effects. Surprisingly, the Ni 100 + ARB treatment induced the highest decrease in soil pH (0.32 unit) and an increase in DPTA-extractable Ni (0.45 mg kg−1 soil) between that measured in the bulk zones and that obtained in the rhizosphere zone. Ni distribution in plant parts and Ni removal (% of total Ni) from the soil were also significantly improved with ARB inoculation, compared to the Ni treatments without ARB. The extent of Ni removal was similar for the Ni 50 + ARB (0.27%) and Ni 100 + ARB (0.25%) treatments. Concluding, ARB-inoculated Salix alba can remove similar amounts of Ni from the soil, irrespective of the Ni pollution level.

1. Introduction

Natural processes and various human activities such as electroplating, alloy manufacturing, production of tannery wastewater, mafic and ultramafic rocks weathering, and improper disposal of Ni-rich industrial wastes pollute the soil with Ni [1,2]. Nickel occurs in the soil in different geochemical fractions such as in an exchangeable form, bound to Fe and Mn (hydro)oxides, carbonates, organic matter (OM), etc. [3]. These forms and geochemical behaviors of Ni in the soil are controlled by several factors like type and content of clay, amount of OM, Fe and Mn (hydro)oxides contents, and soil pH. Nickel bound to carbonate and Ni exchangeable fractions are mobile ones, whereas non-residual fractions become mobile when changes occur in soil pH, redox conditions, and content of dissolved organic carbon (DOC) [4]. The amount of Ni bound to Fe and Mn (hydro)oxides is directly dependent on the soil pH [5]. In conclusion, soil pH, ion exchange, complexation, precipitation, biological transformations, redox conditions, and dissolution are the factors that determine the mobility and phytoavailability of Ni in the soil [4]. In agricultural soils, the threshold level of Ni is 50 mg kg−1 [6]. A soil with 100 mg kg−1 of Ni is considered moderately polluted and can cause ecological and health risks [7]. When up taken by food plants, Ni enters the food chain and may cause several health problems in humans and animals [8].
The remediation of soil through traditional ways, such as soil washing, incineration, replacement, vitrification, etc., has several limitations that restrict the ability to decontaminate Ni-polluted soils. Contrarily, a green remediation approach, phytoremediation, has been regarded as a sustainable, environmentally friendly, cost-effective, and biocompatible process for decontaminating Ni-polluted soils [9]. Phytoextraction, a well-renowned phytoremediation approach, involves the usage of plants to uptake Ni from the soil and the accumulation of Ni in above-ground biomass under natural (continuous) or induced (enhanced or chemically assisted) conditions [8,10].
According to the literature, approximately 500 plant species, mostly belonging to the Brassicaceae, Asteraceae, Cunoniaceae, Euphorbiaceae, and Phyllanthaceae families, have been identified as natural Ni hyperaccumulators which can accumulate high concentrations of Ni. However, these hyperaccumulators produce limited biomass, a primary hindrance to the success of phytoextraction [11,12]. Alternatively, several high-biomass-yielding plants, especially those belonging to the genus Salix, can be used to clean Ni-contaminated soils, though they uptake moderate quantities of Ni from the soil, which can prolong the time for soil cleanup [13,14].
In order to counteract these shortfalls, enhanced or chemically assisted phytoextraction is suggested, which involves the enhancement of Ni bioavailability in the soil through several chemical agents and, later, Ni uptake by plants [1,10]. In this context, ethylenediamine disuccinic acid (EDDS), nitrilotriacetic acid (NTA), hydroxy ethylene diamine triacetic acid (HEDTA), and ethylenediaminetetraacetic acid (EDTA) were extensively used for the enhanced phytoextraction of Ni [1,15]. However, several limitations in the use of these chelating agents, such as their high cost, environmental persistence, and the possibility of Ni leaching into groundwater, have limited their usage [15]. Later, low-molecular-weight organic acids, like citric, lactic, oxalic, tartaric, and malic acids, were widely applied to enhance the phytoextraction of Ni and other heavy metals (HMs) from polluted soils [16,17]. Since they degrade quickly in the soil, their regular application is needed, which increases their cost and hence is a limiting factor in the success of phytoextraction [15,18,19]. In the past few years, elemental sulfur (S) was proven to improve Ni phytoextraction by several plants like Vicia sp., Sorghum sp. and Brassica sp., Pisum sativum, and Zea mays [20,21]. Unfortunately, the leaching of SO4 and HMs into groundwater and the harmful effects of S on the activities of microorganisms and soil enzymes may negatively affect HMs phytoextraction [21,22].
According to recent research, some genera of plant growth-promoting rhizobacteria (PGPRs), such as Enterobacter, Bacillus, Pseudomonas, Klebsiella, and Serratia, may enhance Ni phytoextraction from Ni-polluted soils by several plant species [23]. One of the PGPR genera, Bacillus, is fast-growing and strongly resistant to Ni stress. Despite the harmful effects of Ni on different soil microbes, Bacillus spp. adapt to Ni-polluted soil through various mechanisms. The members of the Bacillus genus are excellent biosorbents of Ni on active chemosorption sites linked with peptidoglycan layers in their cell walls [24]. Furthermore, also chitin, present in the cell wall of Bacillus spp., acts as a biosorbent for Ni. Interestingly, the bioaccumulation of Ni in Bacillus spp. is much higher than its release back into the environment [25]. Apart from it, members of this genus have a solid capability to form spores in Ni-polluted soils, thus ensuring their survival in a polluted environment. Other adaptive strategies include the formation of biosurfactants and the presence of metal-resistant genes, which help these bacteria cope with Ni pollution in the soil [25,26]. Therefore, this genus is a candidate of choice to enhance Ni phytoextraction by improving the growth and biomass of plants via increasing nutrient bioavailability, the production of phytohormones, and the secretion of enzymes and strengthening the communities of other beneficial microorganisms in the soil [27,28].
The members of the Salix genus have been successfully used for the phytoextraction of HMs, including Ni, and vary in their ability to extract them from the soil [13]. Compared to other species, Salix alba offers broad advantages such as a deep root system, a high biomass production, a high metal translocation ability from the roots to the shoots, and rapid growth and development. Moreover, Salix alba produces large amounts of woody biomass, which can be used as fuel and for the production of bioenergy after the completion of the phytoextraction process [13,14]. Until now, extensive research has been carried out using different species of Salix and PGPR alone or in combination for Ni phytoextraction from polluted soils. Combining rhizosphere-acidifying PGPR and Salix alba for the enhanced phytoextraction of Ni could be a novel approach to decontaminate Ni-polluted soils in a short time. Herewith, we hypothesized that the inoculation with a PGPR with rhizosphere acidification potential, named Bacillus sp. ZV6 (ARB), would increase Ni bioavailability in the rhizosphere by reducing the pH of this zone, improving plant tolerance against Ni toxicity, enhancing plant growth, and resultantly improving the phytoextraction of Ni by Salix. In this context, we performed a pot study where Salix alba was grown on soil containing Ni at zero, threshold, and moderate pollution levels (at 0, 50, and 100 mg Ni kg−1 soil, respectively) in the presence or absence of ARB. The aim of this study was to observe the effects of these treatments on (1) the growth and biomass of Salix, (2) the tolerance of the plants against Ni stress, (3) microbial abundance and enzyme activities in the soil, and (4) Ni bioavailability in the rhizosphere, its distribution in the plants after its uptake, and the extent of Ni removal from the soil.

2. Materials and Methods

2.1. Collection of the Soil, Its Analysis, and Preparation of Soil with Various Ni Concentrations

Soil (0–20 cm from the surface) from an agriculture field in northwest Faisalabad, Pakistan, was collected, sieved (2 mm), and analyzed for its physiochemical properties, described in Table 1, using standard methodologies, which were described in our previous paper [9]. Then, three levels of Ni, i.e., zero, threshold, and moderate pollution levels (0, 50, and 100 mg kg−1 soil) were achieved in the soil. For this purpose, the calculated Ni(NO3)2 amounts were dissolved in distilled water and homogenized with two soil portions, while a third portion of soil was left untreated, yielding the zero Ni sample. The three soils were packed in polyethylene zippers and incubated for 60 days at 25 °C in a dark room.

2.2. ARB Isolation, Identification, Physicochemical Characteristics, and Addition to the Soil

The bacterial strain used in this study were isolated in our recent research and was named Bacillus sp. ZV6 (ARB), after BLASTn and phylogenetic analyses [29]. The physicochemical characteristics of ARB were determined according to the methodologies stated in our previous work [30]. The ARB strain was aerobic, rod-shaped, Gram-positive, and motile. The examination of colonies revealed that they were fuzzy white, fairly circular, smooth, shiny, and had a convex elevation. Other characteristics of ARB are as follows: growth at 15–35 °C (+), P solubilization (+), citrate (+), indole acetic acid (IAA) production (+), 1-aminocyclopropane-1-carboxylic acid (ACC) deaminase activity (−), and siderophore production (+). For the inoculum preparation, the strain was grown in Nutrient Broth Medium, shaking the culture for two days at room temperature. Then, the medium was centrifuged (2287× g, 2 °C, 5 min) to obtain the bacterial inoculum. The sediment was re-suspended in sterile tap water to obtain an inoculum with 108 colony-forming units (CFUs) mL−1 [31]. Finally, according to the treatment plan, the ARB inoculum (10 mL pot−1) was sprayed on the soil, and later, the soil was homogenized for the proper distribution of the bacteria (Table 2).

2.3. Phytoextraction Study

This experiment was set up in the greenhouse of Government College University, Faisalabad, Pakistan. For this purpose, soil (3 kg) was placed in each planter (height = 22.8 cm, width = 17.8 cm) as per the treatment scheme (Table 2). Each treatment was replicated three times, and the pots were arranged in a completely randomized design (CRD). Then, sterile distilled water (SWt) was poured into the pots and set for two days to achieve the proper moisture. The cuttings (length = 10 cm, width = 0.5 cm) of Salix alba were purchased from Flower Spot Plant Nursery Lahore, Pakistan. The cuttings were initially rinsed with simple tap water and later dipped in ethanol (70%) for 1 min, a NaClO2 solution (2.5%) for 3 min, and ethanol (70%) for 1 min. Ultimately, the cuttings were gently washed with SWt thrice [32] and coated with Hormex Rooting Powder # 8 (Maia products, Inc. Westlake Village, CA, USA) at the basal side, and three of them were planted in every planter. Shoots emerged from the buds after six days from planting. Before harvesting the plants after 120 days, the planters were irrigated with SWt regularly according to the weather conditions.

2.4. Pot Experiment Termination and Analysis

Plant height was estimated with a soft tape measure before plant harvest. After collecting the biomass above the soil level, the roots were retrieved out of each pot. Soil up to 1 mm from the roots was gently removed with a brush and is referred to as rhizosphere soil, while the soil surrounding the roots (distance of 2–5 cm) was also retrieved and is referred to as bulk soil [33]. Finally, uniform samples of bulk and rhizosphere soils were air-dried, sieved (2 mm), and stored for analysis. After washing the shoots and roots with distilled water several times, they were oven-dried at 70 °C, and the dry weight (DW) of the shoots (Shoot DW) and roots (Root DW) was measured.

2.4.1. Parameters of the Soil

Dynamics of Ni Bioavailability and pH of Bulk and Rhizosphere Soil Zones

The values of pH of the rhizosphere and bulk soils were measured with a calibrated pH meter (model WTW 7110, Xylem Analytics, Weilheim, Germany) [34]. Likewise, the rhizosphere and bulk soils were extracted with a diethylenetriaminepentaacetic acid (DTPA) solution (5 mM), and their plant-assessable Ni concentrations were measured with inductively coupled plasma mass spectrometry (ICP−MS) (PerkinElmer’s NexION® 2000, PerkinElmer Inc., Shelton, CT, USA).

Soil Enzymology

In order to estimate the activity of the enzyme dehydrogenase, moist soil (1 g) was mixed in the solution of 0.2 mL of 2,3,5−triphenyl tetrazolium chloride, 0.05 mL of solution of glucose (10 g L−1), and 1 mL of TRIS buffer and incubated for about 24 h at 35 °C. Then, 10 mL of methanol was used to extract the soil. Further, to determine the triphenylformazan amount in this extract, UV spectrophotometry (λ 485 nm) was employed [35]. The method reported by Eivazi and Tabatabai [36] was used to analyze β-glucosidase activity. Briefly, the soil (1 g) was mixed with ρ-nitrophenyl-β-D-glucopyranoside as a substrate and a pH buffer (pH = 9). Later, the solution underwent incubation at 37 °C for 1 h. To halt β-glucosidase activity, Tris was poured into the solution. The cleavage product derived from the substrate, ρ-nitrophenol glucoside, was then measured on a spectrophotometer (at 464 nm). Similarly, for measuring urease activity, the methodology recommended by Kandeler and Gerber [37] was used. The moist soil (1 g) was homogenized in a flask with 4 mL of borate buffer (pH = 10) and a urea solution (0.5 mL), and the mixture underwent incubation (2 h). Next, 1 M KCl (6 mL) was added to the flask, mixing for 30 min. Then, distilled water, Na salicylate/NaOH, and sodium dichloroisocyanurate were poured into the filtrate. Prior to measuring the optical density (690 nm), the solution was placed for 30 min at room temperature to measure the NH4+ content. Likewise, a method by Bollag et al. [38] was used to detect peroxidase activity in the soil. Briefly, a soil extract (2.7 mL) was mixed with 50 µL of 0.5% o−dianisidine, 0.3 mL of 0.06% of H2O2, and 50 mM of phosphate buffer (pH = 6) in methanol. Following this, a spectrophotometer (460 nm) was employed to measure the activity of peroxidase in the solution. The Tabatabai and Bremner [39] technique was used for the measurement of acid phosphatase activity. For that, 1 g of soil was mixed with a solution [4 mL of universal buffer (pH = 5) and p-nitrophenyl phosphate substrate (0.025 M)] and underwent incubation for 60 min (37 °C). Later, the p-nitrophenol absorbance was measured on the spectrophotometer (398 nm).

Soil Microbial Parameters

The method recommended by Wu et al. [40] was employed for determining the bacterial abundance in the soil. Aqueous extracts from the soil samples were diluted and spread in beef extract−peptone medium. Following incubation, the bacteria were counted for 2–5 days under suitable conditions. Likewise, the microbial biomass was determined by using the chloroform fumigation method [41]. The wet oxidation method of Walkley and Black [42] was employed to determine the organic C.

2.4.2. Plant Parameters

Allocation of Ni in Plant and Ni Removal Efficacy from the Soil

The dry biomass (shoots/roots) was ground in a milling machine and sieved using a 0.5 mm sieve. The roots and shoots samples (1 g) were digested through the open flask digestion method with a mixture of HClO4 and HNO3 (1:2, v/v) [43]. ICP-MS was used to measure the Ni concentration in the digests of the shoots and roots, which was expressed as mg kg−1 DW for each plant portion. The Ni content in the shoots and roots was also estimated using Formula (1) and was expressed as mg pot−1.
Ni concentration in the plant portion × biomass of the plant portion (in kg) harvested from the pot
Soil Ni removal was measured by adopting Formula (2). In Formula (2), the total Ni in the soil corresponds to the total soil Ni content (in mg in the whole soil of the pot).
Shoots content of Ni + roots content of Ni/total soil Ni content × 100

Investigation of Proteins, Phenolic, and Carotenoid in Salix Leaves

Bradford method [44] was chosen to measure the protein content. Briefly, a fresh leaf sample (1 g) was homogenized in a buffer solution [2% polyvinylpyrrolidone, 0.5% Triton X-100, 40 mM tris-HCl], 1 mM phenyl methane sulfonyl fluoride, 1 mM EDTA, and 0.07% β-mercaptoethanol] by pestle and mortar and centrifuged (20,000× g) for 10 min. Then the afloat was collected, and the Bradford reagent was mixed in the solution, which caused a change in the color. The color intensity was measured by a spectrophotometer, and the protein content was reported in mg g−1 FW. Similarly, the carotenoid content was measured through the Hiscox and Israelstam [45] procedure. Samples of fresh leaves (1 g) were homogenized in 20 mL of a solution [methanol chloroform water (12 to 5 to 3 ratio)]. Afterwards, using a spectrophotometer (at 452.5 nm), the carotenoid content was determined. Likewise, the phenolic compound content was measured using the Folin−Ciocalteu method [46]. A mixture was prepared by homogenizing methanol (80%) and a leaf extract. Then, this solution was shaken for 30 min in a flask and underwent centrifugation (15 min) at 25 °C. Next, the solution (100 µL) was diluted with the Folin–Ciocalteu reagent (100 µL) and 1 mL of methanol (80%) and left to rest for 2–3 min. Then, 3 mL of Na2CO3 (20%) was added to the solution, which was stored for 30 min at 25 °C. Later, the absorbance was determined on a spectrophotometer (765 nm).

Assessment of the Plant Defense System

Fresh leaves (500 mg) were homogenized in 4 mL of potassium phosphate buffer (pH = 7), 5 mM mercaptoethanol, 1 mM AsA, 100 mM KCl, and 10% (w/v) glycerol to estimate antioxidant enzymes’ activities. For the collection of an afloat, the solution was centrifuged (11,500 × g) for 10 min. Following that, the same samples were used to evaluate the activities of superoxide dismutase (SOD), catalase (CAT), ascorbate peroxidase (APX), and peroxidase (POD) in the leaves, according to the methodologies of El-Shabrawi et al. [47], Aebi [48], Nakano and Asada [49], and Dias and Costa [50], respectively. Similarly, by employing the techniques described by Jambunathan [51], Velikova et al. [52], and Yang et al. [53], the content of reactive oxygen species (ROS), such as MDA and H2O2, as well as the O2•− generation rate in the leaves, respectively, were determined.

2.5. Statistical Analysis

The current study was conducted according to CRD. All treatments were replicated thrice, and the standard error (SE) was calculated using Microsoft Excel 2013. The findings were interpreted using one-way analysis of variance (ANOVA) in Statistix 8.1. (Analytical software, Tallahassee, FL, USA). To find significant differences (at p < 0.05) between the treatment means, the least significant difference (LSD) test was performed.

3. Results and Discussion

3.1. Agronomical and Biochemical Traits

Plant height and shoot and root DW ranged from 75.7 to 103.4 cm, from 4.62 to 7.31 g pot−1, and from 0.98 to 1.81 g pot−1, respectively. The protein, phenolic, and carotenoid contents ranged from 10.8 to 14.9 mg g−1 FW, from 6.47 to 8.98 mg g−1 FW, and from 0.43 to 0.66 mg g−1 FW, respectively (Figure 1). Without the ARB inoculum, the two tested doses of Ni significantly reduced all of these parameters, compared to the control. Relative to the control, significant reductions in plant height and in the contents of protein and phenolic compounds were observed with the Ni 100 + ARB treatment, whereas the carotenoid content with Ni 50 + ARB and Ni 100 + ARB treatments were reduced. Interestingly, no significant differences were observed in plant height and protein and phenolic contents after the Ni 50 + ARB treatment, in shoot DW and root DW after the Ni 50 + ARB and Ni 100 + ARB treatments, and in carotenoid content after the ARB treatment, compared to control. The sole ARB inoculum was the most efficient treatment, which improved plant height, shoot and root DW, and the contents of protein and phenolic compounds by 8, 19, 22, 10, and 9%, respectively, compared to the control.
The pollution of soil with Ni reduced the DW and the length of shoots and roots for zucchini [54] and rice [55]. Moreover, the phenolic and carotenoid contents of wheat [56] were significantly reduced in Ni-stressed soils. Nickel reduces plant growth and yield by reducing the absorption of nutrients by the roots, damaging thylakoid structures, lowering ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCO) activity, and affecting stomatal closure and the biosynthesis of chlorophyll [57,58]. Interestingly, adding PGPR to Ni-polluted soil notably boosted the growth and the DW of Arabidopsis thaliana L. [59]. The contents of phenolic compounds in wheat [56], while that of protein and carotenoid in Zea mays [60] were boosted in the presence of Bacillus sp. and Proteus mirabilis, respectively. In our experiment, Salix alba growth and biomass were positively affected by the ability of ARB to solubilize P and secrete IAA in the soil (Figure S1). IAA induces cell elongation and division in plant roots, thus promoting their growth and nutrient acquisition [61]. Furthermore, the improvements in the growth and biomass of Salix alba were also attributed to the ability of PGPRs to reduce ethylene production and improve Fe2+ availability to plants through the synthesis of ACC deaminase [23] and siderophores [25], respectively. Interestingly, the Bacillus species produce non-enzymatic antioxidants (phenolics and carotenoids) in the plant rhizosphere, which reduce the entry of HMs ions into the roots and thus neutralize their toxic effects [56].

3.2. pH Alteration and Ni Bioavailability in the Soil

The pH of the soil varied from 6.89 to 7.21 and from 6.62 to 6.99 in the bulk and rhizosphere, respectively (Figure 2). No significant changes in the pH values of the bulk and rhizosphere soils were observed after the Ni 50 and Ni 100 treatments, compared to their matching controls. However, the sole inoculum of ARB and the Ni 50 + ARB and Ni 100 + ARB treatments significantly reduced the pH values by 0.34, 0.37, and 0.34 units in bulk soil and by 0.30, 0.31, 0.30 units in rhizosphere soil, in comparison to their matching controls. Overall, decreases in soil pH up to 0.25, 0.21, 0.25, 0.28, 0.32, and 0.29 units in the control, Ni 50, Ni 100, ARB, Ni 50 + ARB, and Ni 100 + ARB samples, respectively, were noted when comparing the bulk and the rhizosphere zones.
The concentrations of DPTA-extractable Ni varied from 0.041 to 2.74 mg kg−1 and from 0.034 to 2.11 mg kg−1 for rhizosphere and bulk soils, respectively (Figure 2). Excluding ARB, the other treatments notably amplified the DPTA-extractable Ni in both bulk and rhizosphere zones in comparison to their matching controls. The topmost solubilization of Ni was noted with the Ni 100 + ARB treatment. It resulted in the elevation of the bioavailable Ni concentrations by 67.5 times in the rhizosphere zone and by 51.8 times in the bulk zone compared to the control. Overall, increased levels of DPTA-extractable Ni up to 0.006, 0.23, 0.33, 0.012, 0.32, and 0.45 mg kg−1 soil in the control, Ni 50, Ni 100, ARB, Ni 50 + ARB, and Ni 100 + ARB samples, respectively, were noted between the bulk and the rhizosphere zones.
Former research reported lower soil pH values of the rhizosphere zones of mung bean [33] and white willow [14] compared to the bulk soil zones. Depending on the metal species, pH exhibits vital part in the solubility and bioavailability of HMs in the soil. The secretion of H+ ions, exudates, and organic acids by the roots in the rhizospheric zone of several plants, including Salix alba, leads to acidity in this zone, compared to the bulk one. Such rhizosphere acidification results in enhanced solubility of HMs in this vicinity and hence, in their accumulation in the plants [33]. Moreover, a significant reduction in soil pH and, resultantly, enhanced Zn bioavailability were observed in the rhizosphere of Brassica juncea inoculated with PGPR (Rhizobium leguminosarum) [62]. Likewise, Serratia sp. (CTZ4) application reduced soil pH, improved Cd bioavailability, and enhanced Cd phytoextraction by Amaranthus hypochondriacus from a Cd-polluted soil [63].
In this study, we used ARB, a rhizosphere bacterium, to induce acidity in the rhizosphere of Salix. The Bacillus spp. produce cellulolytic enzymes, which break lignocellulose (part of soil OM) into sugars through hydrolysis. Later, Bacillus spp. uptake these sugars, ferment them, and release organic acids in their vicinity [64]. Moreover, plants secrete root exudates in the rhizosphere, which comprise several organic compounds, such as organic acids, vitamins, sugars, volatile organic compounds (VOCs), plant growth-promoting hormones, carbohydrates, flavonoids, and nucleotides [65]. Several vital metabolic pathways, such as fermentation, direct oxidation of these organic compounds, and respiration are key pathways for the secretion of different organic acids by Bacillus spp., which induce acidity in the rhizosphere [66]. Apart from the secretion of organic acids, Bacillus spp. are known to excrete chelates to enhance HMs solubility in the rhizosphere [67]. Furthermore, biosurfactants and siderophores, produced by Bacillus spp., are metal-complexing compounds that improve the mobility and bioavailability of HMs in the soil [25]. In our experiment, apart from the root-induced acidity in the rhizosphere, ARB enhanced Ni bioavailability in this zone by secreting organic acids, chelates, biosurfactants, and siderophores and thus promoted Ni uptake by Salix alba.

3.3. Phytoextraction of Ni by Salix and Removal of Ni from the Soil

The Ni concentrations in shoots and roots varied between 0.13 to 112.50 mg kg−1 DW and 0.07 to 93.37 mg kg−1 DW. Likewise, the Ni content in the roots and shoots increased from 0.0002 to 0.12 mg pot−1 and 0.0004 to 0.63 mg pot−1. The extent of Ni removal from the soil ranged from 0.14 to 0.27% (Figure 3). All treatments, except ARB, significantly increased the Ni concentration in the roots and shoots, the Ni content in these parts, and the Ni removal from the soil compared to the control. Interestingly, the topmost improvements in Ni concentration and in Ni content in the roots and shoots were by 736, 1533, 641, and 1402 times, respectively, and were determined for the Ni 100 + ARB treatment. Likewise, the highest significant values of Ni removal from the soil were found for the Ni 50 + ARB and Ni 100 + ARB treatments with respect to the control.
A study reported that the Ni concentration in the shoots and roots of buttonwood increased with increasing concentrations of Ni in the soil [9]. According to our data, ARB-inoculated plants showed higher concentrations of Ni in both shoots and roots than non-inoculated plants (Figure 3). Herewith, a reduction in soil pH by ARB enhanced Ni bioavailability in the rhizosphere and improved Ni uptake by Salix alba, as discussed in detail (Section 3.2 of this manuscript). The phytoremediation of Ni is strongly dependent on the presence of sufficient plant biomass and on the plant ability to uptake high amounts of Ni in its harvestable parts [9,68]. The inoculation of plants with ARB enhanced the Ni content in the roots and shoots by increasing their Ni concentration and biomasses (Figure 1 and Figure 3). Moreover, the Ni content was higher in the shoots than in the roots (Figure 3), showing the capability of Salix alba to transport greater concentrations of Ni to its harvestable aerial portions and its significance for the (phyto)remediation of Ni-polluted soils. Similar findings were recently reported regarding the accumulation of various HMs, including Ni, by different clones of Salix growing in HMs-polluted soils [13,14]. Significantly, the highest percentages of Ni removal were found for the Ni 50 + ARB and Ni 100 + ARB treatments, compared to the control (Figure 3). In the past, higher percentages of Cd and Zn were efficiently removed by ryegrass in the presence of PGPR (Burkholderia sp. D54), compared to the control [69]. According to our findings, the significantly similar percentages of Ni removal achieved with the Ni 50 + ARB and Ni 100 + ARB treatments showed that Salix alba in the presence of ARB maintains the same potential to remediate soils in the presence of different levels of Ni pollution.

3.4. Defense System of Salix

The activities of antioxidants enzymes in the leaves were from 43.0 to 181.9 U min−1 mg−1 protein for CAT, from 0.33 to 0.71 µmol min−1 mg−1 protein for APX, from 12.4 to 61.2 U min−1 mg−1 protein for POD, and from 41.4 to 87.1 U min−1 mg−1 protein for SOD (Table 3). In parallel, the content of H2O2 and MDA and the O2•− generation rate ranged from 14.7 to 48.5 nmol g−1 FW, from 0.25 to 0.84 µmol g−1 FW, and from 31.8 to 77.3 µg g−1 FW, respectively (Table 3). Compared to the control, all treatments significantly affected the activities of APX, POD, and SOD, except for the Ni 50 + ARB treatment in the case of CAT. Similarly, except for the ARB treatment, the rest of the treatments showed significant effects on the content of H2O2 and on the O2•− generation rate in comparison to the control, whereas each treatment had a considerable impact on the content of MDA compared to the control. The greatest improvements in the activities of CAT (by 7%), APX (by 10%), POD (by 12%), and SOD (by 8%), relative to control, were noted in the plants treated with ARB. Furthermore, the treatment Ni 100 resulted in maximum increases of 209%, 174%, and 123% in the contents of H2O2 and MDA and in the O2•− generation rate, respectively, compared to the control.
Reduced activity of APX, SOD, and CAT and an elevated H2O2 content was observed in Arabidopsis thaliana in Ni-polluted soil [59]. In another study, Valivand et al. [54] reported that Ni-polluted soil reduced antioxidant enzyme activities (CAT and POD) and increased the contents of H2O2 and MDA in the zucchini plant. Nickel is known to promote ROS production in plants. Plants having an efficient antioxidant system can withstand Ni toxicity. However, a high bioavailability of Ni in the soil jeopardizes this defense system, leading to oxidative stress and poor plant growth [9]. Cations such as Ca2+ and Zn2+ are essential for membrane stability, while several cofactors vital in antioxidant enzymes contain Fe2+. Unfortunately, due to the competition for absorption by the roots, a higher concentration of Ni in the soil leads to a reduced uptake of Ca2+, Zn2+, and Fe2+, causing lipid peroxidation and membrane instability [8]. In contrast, PGPR inoculation in Ni-polluted soils improved SOD, APX, and POD activities while reducing oxidative stress in sunflower [27]. In our study, at each Ni concentration, the ARB-inoculated plants exhibited lower oxidative stress and higher antioxidant activities than the non-inoculated ones (Table 3). Modulating the expression of several genes controlling ROS accumulation and activating ROS-scavenging enzymes are effective mechanisms of PGPRs to tackle Ni-persuaded oxidative injury in plants [70,71]. Moreover, while colonizing the rhizosphere, PGPRs reduce metal-induced oxidative stress in plants by (1) improving plant growth and nutrition [61] and (2) neutralizing the toxic effects of metals via chelation [25].

3.5. Soil Enzymology and Microbial Traits

Activities of β-glucosidase, dehydrogenase, urease, acid phosphatase, and peroxidase varied between 0.18 to 0.30 μmol PNF g−1 h−1, 1.12 to 2.29 μmol INTF g−1 h−1, 1.87 to 3.41 μmol N-NH4+ g−1 h−1, 17.9 to 35.1 mg PNP kg−1 h−1, and 5.83 to 11.7 mol g−1 h−1, respectively (Figure 4). Both doses of Ni, without ARB inoculum, had a significant negative impact on the activity of these enzymes, compared to the control. Furthermore, significantly lower activities of β-glucosidase, dehydrogenase, and urease, relative to the control, were noted after the Ni 100 + ARB treatment. Interestingly, no significant differences in acid phosphatase and peroxidase activities were found after the Ni 50 + ARB and the Ni 100 + ARB treatments, as well as in β-glucosidase, dehydrogenase, and urease activities after the Ni 50 + ARB treatment, in comparison with the control. The highest significant improvements in β-glucosidase, dehydrogenase, urease, acid phosphatase, and peroxidase activities, by 15, 12, 13, 22, and 36%, respectively, compared to the control, were noted in the presence of the sole ARB inoculum.
The count of bacteria, microbial biomass, and organic C values ranged from 215.6 to 1001.3 × 103 CFU g−1 soil, 144.3 to 288.6 mg kg−1 soil, and 0.43 to 0.79 g C kg−1 dry soil, respectively (Figure 4). All treatments affected the count of bacteria, microbial biomass, and organic C, except for the Ni 50 treatment in regard of bacterial count and the Ni 50 + ARB treatment in regard to organic carbon, with respect to the control. Significant improvements in bacterial count and microbial biomass were observed with the ARB, Ni 50 + ARB, and Ni 100 + ARB treatments and only with the ARB treatment in the case of organic C, compared to the control. However, compared to the control, the greatest improvements of 241% in bacterial count and 26% in organic C were noted with the ARB treatment, and improvements of 48% and 40% in microbial biomass were observed with the ARB and Ni 50 + ARB treatments, respectively. Furthermore, the highest declines in bacterial count, corresponding to 18% and 27%, and in microbial biomass corresponding to 17 and 26%, were found after the Ni 50 and Ni 100 treatments, respectively, while a decline of only 31% in organic C was observed after the Ni 100 treatment, compared to the control.
Microbes secrete enzymes related to nutrient cycling, which improve plant growth and, thus, their capacity for HMs phytoextraction [61,72]. The soil enzymatic activities have a negative correlation (nonlinear) with Ni and other HMs concentrations in the soil and a positive correlation with microbial biomass and microbial biomass C [63,72]. Soil pollution with Ni significantly reduced β-glucosidase, dehydrogenase, urease, and acid phosphatase activities [2,73], as well as the contents of organic C and microbial biomass [74]. Nickel toxicity negatively affects soil enzymatic activities by reducing number of soil microbes through the disintegration of their cell membrane, DNA damage, the enhancement of lipid peroxidation, the impairment of functional proteins, and the inhibition of several enzymes [1]. In previous studies, enhanced activities of dehydrogenase, acid phosphatase, and urease in Ni- [2] and Cd- [63] polluted soils, with PGPR and metal-resistant bacteria (Serratia sp. CTZ4), respectively, were recorded. Adding a PGPR (Serratia sp. strain H3) to Ni-polluted soil significantly improved the bacterial count [75]. In our experiment, the treatments with the ARB inoculum led to higher enzymatic activities, bacterial count, biomass of microbes, and organic C than the matching treatments without the inoculum (Figure 4). This enhanced concentration of organic C in the presence of the ARB inoculum is due to the ability of the Bacillus spp. to secrete several C-rich compounds in the soil, such as phytohormones, ACC deaminase, polyamines, lipopeptides, and acyl homoserine lactone [76]. Moreover, the Bacillus spp. also secrete several VOCs (aldehydes, ketones, alkyls, alcohols, alkenes, esters, acids, ethers, heterocyclic, and phenolic compounds) in the soil. These VOCs act as signaling molecules and are secreted via the chemical interaction of Bacillus spp. with plant roots and other soil microbes. These compounds are responsible for increasing the concentration of organic C in the soil, facilitating plant growth and increasing microbial abundance in the soil [76,77]. Besides secreting soil enzymes [2], PGPRs also support the abundance of indigenous microbes and their capacity to secrete enzymes by (1) providing them nutrition (via IAA, siderophores, and P solubilization) [63] and (2) reducing Ni toxicity via Ni chelation, adsorption, precipitation, and bioaccumulation [25].

4. Conclusions

Soil pollution with Ni poses severe ecological hazards and public health threats. Enhanced phytoremediation of Ni using chelating agents can decontaminate Ni-polluted soils but could be costly due to the high prices of the chelating agents. Nickel is present in the soil in several fractions, e.g., in its exchangeable form, bound to OM, carbonates, Fe and Mn (hydro)oxides, etc. A reduction in soil pH can enhance the solubility of Ni, releasing it from most of these fractions, and thus increase its phytoavailability to phytoextraction crops. Surprisingly, plant growth-promoting rhizobacteria (PGPR), which can induce acidity in the rhizosphere, can augment Ni phytoextraction by enhancing Ni solubility in soil and plant growth and by reducing Ni-induced oxidative stress in plants. Our current investigation showed that a PGPR with rhizosphere acidification potential, named Bacillus sp. ZV6 (ARB), induced significant acidity in willow rhizosphere and augmented the solubility of Ni in this zone. Moreover, ARB had positive effects on plant growth and Ni distribution in the plant shoots and roots and reduced plant oxidative stress in the presence of all tested Ni concentrations (0, 50, and 100 mg kg−1 soil), compared to inoculum-free treatments. Improvements in enzymatic activities, count of bacteria, biomass of the microbes, and organic C in soil were also observed. Interestingly, the percentage removal of Ni from the soil was similar for both soil Ni levels tested (50 and 100 mg kg−1 soil), showing the ability of ARB-inoculated Salix alba to remove similar amounts of Ni in the presence of different levels of Ni pollution in the soil. Our findings suggest that Ni phytoextraction with Salix alba inoculated with ARB can efficiently decontaminate Ni-polluted soils irrespective of the extent of Ni contamination at a lower cost compared to other commonly used methods.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/min13091178/s1, Figure S1: The potential of ARB to produce IAA (μg mL−1) and solubilize P (μg mL−1).

Author Contributions

Conceptualization, M.I. and S.G.; methodology, Z.A.V., M.S.A. and M.Z.Y.; software, M.F.I., S.A., A.T.A. and Z.A.V.; validation, A.D. and M.I.; formal analysis, Z.A.V.; investigation, Z.A.V. and M.I.; resources, Z.A.V. and M.I.; data curation, S.G., Z.A.V. and A.D.; writing—original draft preparation, Z.A.V., S.A., A.T.A. and S.G.; writing—review and editing, M.I.; visualization, M.F.I.; supervision, M.I. and M.S.A.; project administration, Z.A.V.; funding acquisition, M.I. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the Researchers Supporting Project number (RSP2023R194), King Saud University, Riyadh, Saudi Arabia.

Data Availability Statement

The data of this study are present in the manuscript.

Acknowledgments

Our thanks to Veysel Turan, at Bingöl University, Turkey, for performing many of the analyses related to this study. This study was supported by the Researchers Supporting Project number (RSP2023R194), King Saud University, Riyadh, Saudi Arabia.

Conflicts of Interest

The authors declare no conflict of interest.

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Minerals 13 01178 g001
Figure 1. Effects of ARB inoculation on the plant height (A), shoot dry weight (Shoot DW) (B), root dry weight (Root DW) (C), protein (D), phenolic (E), and carotenoid (F) of Salix alba under varying Ni concentrations after plant harvest. The results of each treatment are indicated by the mean ± SE of three replicates. Significant variations are denoted by different alphabet letters (LSD test, p < 0.05).
Figure 1. Effects of ARB inoculation on the plant height (A), shoot dry weight (Shoot DW) (B), root dry weight (Root DW) (C), protein (D), phenolic (E), and carotenoid (F) of Salix alba under varying Ni concentrations after plant harvest. The results of each treatment are indicated by the mean ± SE of three replicates. Significant variations are denoted by different alphabet letters (LSD test, p < 0.05).
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Figure 2. ARB inoculation effects on the soil pH values (A) and concentrations of plant-accessible Ni (B) in the rhizosphere and bulk soil portions under varying Ni concentrations after plant harvest. The results of each treatment are indicated by the mean ± SE of three replicates. Significant variations are denoted by different alphabet letters (LSD test, p < 0.05).
Figure 2. ARB inoculation effects on the soil pH values (A) and concentrations of plant-accessible Ni (B) in the rhizosphere and bulk soil portions under varying Ni concentrations after plant harvest. The results of each treatment are indicated by the mean ± SE of three replicates. Significant variations are denoted by different alphabet letters (LSD test, p < 0.05).
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Minerals 13 01178 g003
Figure 3. ARB inoculation effects on Ni concentration in shoots (A) and roots (B), Ni content in shoots (C) and roots (D), and soil Ni removal (E) in the presence of varying Ni concentrations after plant harvest. The results of each treatment are indicated by the mean ± SE of three repeats. Significant variations are denoted by different alphabet letters (LSD test, p < 0.05).
Figure 3. ARB inoculation effects on Ni concentration in shoots (A) and roots (B), Ni content in shoots (C) and roots (D), and soil Ni removal (E) in the presence of varying Ni concentrations after plant harvest. The results of each treatment are indicated by the mean ± SE of three repeats. Significant variations are denoted by different alphabet letters (LSD test, p < 0.05).
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Minerals 13 01178 g004
Figure 4. ARB inoculation effects on the activities of enzymes, i.e., β-glucosidase (A), dehydrogenase (B), urease (C), acid phosphatase (D), and peroxidase (E) as well as on count of bacteria (F), microbial biomass (G), and organic C (H) in the soil under varying Ni concentrations after plant harvest. The results of each treatment are indicated by the mean ± SE of three repeats. Significant variations are denoted by different alphabet letters (LSD test, p < 0.05).
Figure 4. ARB inoculation effects on the activities of enzymes, i.e., β-glucosidase (A), dehydrogenase (B), urease (C), acid phosphatase (D), and peroxidase (E) as well as on count of bacteria (F), microbial biomass (G), and organic C (H) in the soil under varying Ni concentrations after plant harvest. The results of each treatment are indicated by the mean ± SE of three repeats. Significant variations are denoted by different alphabet letters (LSD test, p < 0.05).
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Table 1. Characteristics of the soil used in this study.
Table 1. Characteristics of the soil used in this study.
PropertiesUnitValue(s)
Texture of the soil−Clay loam
Clayg kg−1355
Siltg kg−1265
Sandg kg−1380
pH (H2O)−7.22
Electrical conductivity (EC) dSm−11.83
Cation exchange capacity (CEC)cmolc kg−17.91
Organic matter (OM)g kg−16.8
CaCO3g kg−119.7
DTPA−extractable Nimg kg−10.14
Total Nimg kg−12.07
Exchangeable Kmg kg−171.7
Available Pmg kg−16.48
Table 2. Experimental treatments in this study.
Table 2. Experimental treatments in this study.
TreatmentsAcronymsBacterial Inoculum Volume (OD600 = 1.0)The added Amount of Ni (mg kg−1 Soil)
No additiveControl−0
Ni (threshold level)Ni 50−50
Ni (moderate level)Ni 100−100
Sole bacterial (Bacillus s. ZV6) inoculumARB10 mL pot−10
Threshold Ni level + Bacillus s. ZV6 inoculumNi 50 + ARB10 mL pot−150
Moderate Ni level + Bacillus s. ZV6 inoculumNi 100 + ARB10 mL pot−1100
Table 3. ARB inoculation effects on catalase (CAT), ascorbate peroxidase (APX), peroxidase (POD), and superoxide dismutase (SOD) activities, as well as on the content of reactive oxygen species (ROS), i.e., the content of hydrogen peroxide (H2O2) and malondialdehyde (MDA), and on the generation rate of oxygen radical (O2•−) in the leaves of Salix alba under varying Ni concentrations. The results of each treatment are indicated by the mean ± SE of three replicates. Significant variations are denoted by different alphabet letters (LSD test, p < 0.05).
Table 3. ARB inoculation effects on catalase (CAT), ascorbate peroxidase (APX), peroxidase (POD), and superoxide dismutase (SOD) activities, as well as on the content of reactive oxygen species (ROS), i.e., the content of hydrogen peroxide (H2O2) and malondialdehyde (MDA), and on the generation rate of oxygen radical (O2•−) in the leaves of Salix alba under varying Ni concentrations. The results of each treatment are indicated by the mean ± SE of three replicates. Significant variations are denoted by different alphabet letters (LSD test, p < 0.05).
ROS and Antioxidant EnzymesUnitsTreatments
ControlNi 50Ni 100ARBNi 50 + ARBNi 100 + ARB
CATU min−1 mg−1 protein169.5 ± 4.28 b46.5 ± 1.15 d43.0 ± 1.06 d182 ± 4.60 a160.9 ± 4.07 bc152.3 ± 3.84 c
APXµmol min−1 mg−1 protein0.64 ± 0.01 b0.36 ± 0.01 d0.32 ± 0.01 d0.71 ± 0.02 a0.58 ± 0.02 c0.54 ± 0.01 c
PODU min−1 mg−1 protein54.7 ± 1.38 b13.2 ± 0.31 e12.4 ± 0.28 e61.2 ± 1.53 a46.7 ± 1.18 c40.6 ± 1.01 d
SODU min−1 mg−1 protein80.5 ± 2.02 b43.7 ± 1.09 e41.4 ± 1.03 e87.1 ± 2.19 a73.0 ± 1.84 c66.3 ± 1.67 d
H2O2nmol g−1 FW15.7 ± 0.38 e44.8 ± 1.13 b48.5 ± 1.21 a14.7 ± 0.35 e37.2 ± 0.92 d41.7 ± 1.04 c
MDAµmol g−1 FW0.3 ± 0.01 d0.7 ± 0.02 b0.8 ± 0.03 a0.3 ± 0.01 e0.5 ± 0.02 c0.6 ± 0.02 c
O2•−µg g−1 FW34.7 ± 0.87 d68.2 ± 1.73 b77.3 ± 1.94 a31.8 ± 0.78 d55.7 ± 1.42 c64.0 ± 1.62 b
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MDPI and ACS Style

Virk, Z.A.; Yasin, M.Z.; Gill, S.; Ilyas, M.F.; Dradrach, A.; Alamri, S.; Alfagham, A.T.; Akhtar, M.S.; Iqbal, M. Inducing Rhizosphere Acidification in White Willow with Bacillus sp. ZV6 Enhances Ni Phytoextraction from Soil and Soil Quality. Minerals 2023, 13, 1178. https://doi.org/10.3390/min13091178

AMA Style

Virk ZA, Yasin MZ, Gill S, Ilyas MF, Dradrach A, Alamri S, Alfagham AT, Akhtar MS, Iqbal M. Inducing Rhizosphere Acidification in White Willow with Bacillus sp. ZV6 Enhances Ni Phytoextraction from Soil and Soil Quality. Minerals. 2023; 13(9):1178. https://doi.org/10.3390/min13091178

Chicago/Turabian Style

Virk, Zaheer Abbas, Muhammad Zubair Yasin, Sebam Gill, Muhammad Fraz Ilyas, Agnieszka Dradrach, Saud Alamri, Alanoud T. Alfagham, Mohd Sayeed Akhtar, and Muhammad Iqbal. 2023. "Inducing Rhizosphere Acidification in White Willow with Bacillus sp. ZV6 Enhances Ni Phytoextraction from Soil and Soil Quality" Minerals 13, no. 9: 1178. https://doi.org/10.3390/min13091178

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