The Effect of Heavy Metals on Microbial Communities in Industrial Soil in the Area of Piekary Śląskie and Bukowno (Poland)

Abstract

The aim of this study was to determine the activity and structure of microbial communities in soils contaminated with heavy metals (HMs). To achieve this goal, soil samples were taken from two contaminated sites (i.e., Piekary Śląskie and Bukowno) in Poland. A wide range of methods were applied, including: total and metal-tolerant culturable bacteria enumeration; microbial community structure analysis using the phospholipid fatty acid method (PLFA); denaturing gradient gel electrophoresis (PCR-DGGE); and metabolic activity using BIOLOG and EcoPlate^TM. Our studies showed that HMs negatively affected microbial community structure and activity in polluted soils. Apart from the contamination with HMs, other soil parameters like soil pH and water also impacted microbial community structure and growth. Metal-tolerant bacterial strains were isolated, identified and tested for presence of genes encoding HM tolerance using the polymerase chain reaction (PCR) methodology. Contamination with HMs in the tested areas was found to lead to development of metallotolerant bacteria with multiple tolerances toward Zn, Ni, Cd and Cu. Different genes (e.g., czcA, cadA and nccA) encoding HM efflux pumps were detected within isolated bacteria. Culturable bacteria isolated belonged to Proteobacteria, Actinobacteria and Bacteroidetes genera. Among non-culturable bacteria in soil samples, a significant fraction of the total bacteria and phyla, such as Gemmatimonadetes and Acidobacteria, were found to be present in all studied soils. In addition, bacteria of the Chloroflexi genus was present in soil samples from Piekary Śląskie, while bacteria of the Firmicutes genus were found in soil samples from Bukowno.

1. Introduction

The environmental effects of heavy metals (HMs) have been studied for several years [1,2]. Nevertheless, despite various attempts to lower HM emissions, related environmental contamination remains a serious concern [3] because of their non-biodegradable nature and ability to accumulate in the food chain generating negative effects on human health [4]. As an example, cadmium accumulation in soil can be caused by natural processes, but is mainly the result of anthropogenic activities, including wastewater irrigation, fertilizers, agricultural usage of sewage sludge, smelting and mining [5]. Even microelements (e.g., copper, zinc and nickel) can become toxic to cells if their concentration exceeds cellular requirements [6].

Due to their size, ubiquity and key role in metabolic cycles, microorganisms are considered to be the best indicators of the effects of toxic compounds on soil [7]. The negative impacts of HMs on microbial communities in soil have been reported in several studies [8,9,10,11]. Heavy metals are well-known to lower/inhibit activity of soil enzymes as well as disturb carbon, nitrogen and organic matter transformation and reduce microorganism biodiversity and biomass [6], leading to dominance of specific HM-tolerating microorganisms in soils [12,13,14,15].

Bioremediation using microorganisms and their natural mechanisms responsible for tolerance/resistance to toxic metallic ions like intra and extracellular sequestration, active efflux from cells and reduction of HM ions is considered to be a very promising tool for HM removal/recovery in comparison with the conventional chemical and physical techniques, since it is an environmentally friendly (for example, retention of soil structure and no secondary pollution) and low-cost technique [6]. In particular, microbial biosorbents are very effective in removing contaminants, although other options such as bioleaching or phytoremediation are also available [6,16].

Many researchers are concerned about the environmental impact of untreated polluted areas, and various studies have been conducted to improve this situation and understanding on how microbial communities in areas contaminated with HMs are shaped and function [17,18,19] in order to further develop the optimal and most effective strategy for bioremediation [18,20,21,22].

The area selected for this study is described as a territory of ecological disaster and is located in the industrial region of Southern Poland. Both Mining and Metallurgical Plants ‘Waryński’ in Piekary Śląskie and ‘Bolesław’ in Bukowno were operating for over a century, leading to extensive contamination of soil with various HMs [23,24].

The aim of this study was to (1) determine the effects of HMs and other physicochemical properties of soils on microbial communities in industrial soils with a history of long-term contamination with various HMs; (2) evaluate the actual state of microbial diversity, metabolic activity and structure by culturable and non-culturable methods; (3) isolation and identification of strains potentially useful in bioremediation in this area, including tests on the presence of genes typically involved in HM efflux among those isolated.

The results obtained with this research can contribute to the development of eco-friendly bioremediation technology and monitoring of the environmental effect of soil pollution in this area.

2. Material and Methods

2.1. Soil Sample Collection

Soil samples were collected in the proximity of the former Mining and Metallurgical Plants ‘Waryński’ in Piekary Śląskie (Upper Silesia) and ‘Bolesław’ in Bukowno (Lesser Poland), Poland. Selected soil sampling sites were designed as P1 (50°21′54.4″ N, 18°58′02.5″ E), P2 (50°21′56.2″ N, 18°58′10.0″ E) and P3 (50°22′03.7″ N, 18°57′44.9″ E) in Piekary Śląskie and B1 (50°16′27″ N, 19°29′7″ E), B2 (50°16′4″ N, 19°29′13″ E) and B3 (50°15′51″ N, 19°29′22″ E) in Bukowno (Figure 1).

Five soil samples were taken from the top soil layer (0–0.2 m) from a 1 m × 1 m sampling area and pooled together to make a composite soil sample. The composite samples were transported to the laboratory, where the soil was air dried, sieved through a 2-mm mesh to remove the stones and plant debris and then stored at –20 °C until further use. All analyses were carried out in triplicate for every single soil sample.

2.2. Physicochemical Analysis

Different physicochemical soil parameters were analysed including granulometry, soil moisture, pH, soil organic matter (SOM), organic carbon, total nitrogen, total phosphorus, HM (Zn, Cd, Cu, and Ni) concentration and bioavailable forms, as well as K, P, Na, Mg and Ca.

Soil granulometry was determined using laser diffraction (Mastersizer 2000, Malvern Pananalytical, Malvern, UK) for particles ranging 0–1 mm in size and sieve analysis for particles above 1 mm. The soil samples were classified into a particular granulometric group according to PN-88/B-02480. The soil moisture content was determined in agreement with the ISO method 16586:2003 and soil pH was measured according to PN-ISO 10390:1997. The SOM and organic carbon were estimated using the Turin test [25].

Total nitrogen was quantified using colorimetry by the Slandi 300 (Slandi, Michałowice, Poland) photometer. The soil (0.2 g) was mixed with 2 mL of 30% H₂O₂ and left for 24 h. The sample was then boiled until complete evaporation of the solution. A volume of 20 mL of H₂O was added to the residual precipitate and brought to the boil. After cooling and filtering through a qualitative filter in the solution, the concentration of ammonium, nitrite and nitrate were measured in the solution and the results were added to obtain the total nitrogen value in the dry mass of the soil sample.

To quantify phosphate (PO₄³⁻) and Na, K, Mg and Ca content, 0.5 g of soil sample was added to 3 mL of 60% HNO₃. The suspension was mineralised in a microwave oven (Mars 5, CEM, Matthews, NC, USA). The resulting solution was diluted to 50 mL. The concentration of phosphate (PO₄³⁻) was measured using the colorimetric method on Slandi 300 (Slandi, Michałowice, Poland). The results were converted into phosphorus content in dry matter.

The concentration of Na, K, Mg and Ca were estimated using flame atomic absorption spectrometry (FAAS) (SavantAA Σ, GBC, Braeside, Melbourne, Australia). The results were given in mg/kg dry soil mass.

The total content of selected HMs (Cd, Zn, Ni and Cu) in soil was determined using FAAS (iCE 3500 FAAS, Thermo Fisher Scientific, Waltham, MA, USA). A quantity of 300 mg of soil was dried at 105 °C and then mineralised in a mixture of 8 mL 65% HNO₃ and 2 mL H₂O₂ at 190 °C for 50 min using an Ethos Up microwave oven (Milestone, Sorisole, Italy). After mineralisation, the solutions were filtered on 0.45 µm filter paper and diluted in water and analysed by FAAS. Results were given in mg of metal per kg of dry soil and compared with Polish regulations for the maximum allowed concentration of HMs in soils (Regulation of the Minister of Environment, 9 September 2002 on Soil Quality Standards 2002; further RME 2002) for soil class B: Cd 4 mg/kg dry soil mass, Zn 300 mg/kg dry soil mass, Cu 100 mg/kg dry soil mass and Ni 70 mg/kg dry soil mass.

To quantify water-extractable HMs in soil or their bioavailable fraction (which is the amount of metal extracted from soil using 0.01 M CaCl₂), 5 g of soil sample was dried at 105 °C and resuspended in 50 mL of DI water or 50 mL 0.01 M CaCl₂ solution, respectively, and then shaken for 2 h at 120 rpm. The suspension was filtered through 0.45 µm filter and metal content was measured using FAAS (iCE 3500 FAAS, Thermo Fisher Scientific, Waltham, MA, USA). Results were given in mg of metal per kg of dry soil.

2.3. Culturable Bacteria Enumeration

The total number of heterotrophic (HB) and oligotrophic bacteria (OB), as well as metal-tolerant bacteria were determined by the plate method using tryptic soy agar (TSA) (BTL, Warsaw, Poland) and broth agar (BTL, Warsaw, Poland) for HB and 10% TSA (BTL, Warsaw, Poland) for OB. All plates were supplemented with 200 µg/mL Nystatin to avoid fungal contamination. The metal-tolerant fraction of culturable bacteria was determined on 1/10 TSA (BTL, Warsaw, Poland), TSA (BTL, Warsaw, Poland) and broth agar (BTL, Warsaw, Poland) plates supplemented with 1 mM CdCl₂ (Sigma-Aldrich, Inc. St. Louis, MO, USA), 1 mM ZnCl₂ (Sigma-Aldrich, Inc., St. Louis, MO, USA), 1 mM NiCl₂ (Sigma-Aldrich, Inc., St. Louis, MO USA) or 1 mM CuCl₂ (Sigma-Aldrich, Inc., St. Louis, MO, USA), respectively. The inoculated plates were incubated for 7 days at 22 °C. The number of bacteria was expressed as log of colony forming unit (CFU) per g of dry soil.

2.4. Total Bacteria Enumeration

The total bacterial count in soil was quantified using the 16S rRNA gene in real-time PCR. The template used for amplification was genomic DNA isolated from 2 × 0.5 g of soil using GeneMATRIX Soil DNA Purification Kit (EURx Ltd., Gdańsk, Poland) according to the producer’s protocol and combined as one sample.

A fragment of the 16S rRNA gene was amplified using primers pE (5′-AAA CTC AAA GGA ATT GAC GG-3′) and pF (5′-ACG AGC TGA CGA CAG CCA TG-3′) (Edwards et al., 1989). The reaction mixture (25 µL) consisted of 10 µL of LightCycler^® 480 SYBR Green I Master (Roche Diagnostic GmbH, Mannheim, Germany), 1 µL of pE primer (100 µM dil. 1:10 v/v) (Sigma-Aldrich, Inc., St. Louis, MO, USA), 1 µL of pF primer (100 µM dil. 1:10 v/v) (Sigma-Aldrich, Inc., St. Louis, USA), 2 µL of DNA and 6 µL of H₂O. The amplification conditions were: 95 °C for 10 min, 30 cycles of 10 s at 95 °C followed by annealing for 20 s at 57 °C and extension for 30 s at 72 °C. Fluorescence data were acquired at the end of each extension step at 81 °C to avoid detection of primer dimers. For the melting curve analysis of products, temperature was raised from 65 to 95 °C and the melting temperatures were determined using LightCycler^® (Roche Diagnostic GmbH, Mannheim, Germany) following a modified procedure described elsewhere [26].

Amplification efficiency was estimated by serial dilution of the standard, which was a fragment of 16S rRNA cloned into the pTZ57R/T vector. The standard was prepared using a InsTAclone PCR Cloning Kit (Thermo Fisher Scientific, Waltham, MA, USA), following the manufacturer’s instructions. The 16S rRNA gene copies quantification was calculated as described in [27].

2.5. Total Microbial Activity Measurement

General microbial activity was determined according to a procedure described elsewhere [28] with minor modification: centrifugation of suspension at 2000 rpm for 3 min. The concentration of free fluorescein in the filtered solution was measured at 490 nm using a spectrophotometer (Helios Epsilon, Thermo Fisher Scientific, Waltham, MA, USA). A calibration curve was used to calculate the amount (in μg) of fluorescein per g of dried soil, which corresponds to enzymatic activity in the soil sample.

2.6. Metallotolerant Bacteria Isolation and Identification

Metallotolerant bacteria were isolated from soil samples using 1/10 TSA, TSA and BA media supplemented with 1 mM CdCl₂ (Sigma-Aldrich, Inc., St. Louis, MO, USA)—to ensure metal-resistant bacteria isolation—and 200 µg/mL Nystatin (Sigma-Aldrich, Inc., St. Louis, MO, USA). After incubation for 5 days at 22 °C, morphologically different bacteria were taken for further studies. Strains were identified based on the 16S rRNA sequence. Genomic DNA was isolated using Genomic Mini (A&A Biotechnology, Gdańsk, Poland). The 16S rRNA fragment was amplified by PCR using 0.5 µL of starter 8F AGAGTTTGATCCTGGCTCAG (100 µM, dil. 1:10 v/v) (Sigma-Aldrich, Inc., St. Louis, MO, USA), 0.5 µL of starter 1492R GGTTACCTTGTTACGACTT (100 µM, dil. 1:10 v/v) (Sigma-Aldrich, Inc., St. Louis, MO, USA), 0.5 µL of polymerase DreamTaq™ 500 U (Thermo Fisher Scientific, Waltham, MA, USA), 2.5 µL of 10× buffer DreamTaq (Thermo Fisher Scientific, Waltham, MA, USA), 0.5 µL of dNTP Mix (10 mM) (Thermo Fisher Scientific, Waltham, MA, USA) and 3 µL of DNA. The PCR amplification was done according to [27].

The PCR products were purified using Gel Out (A&A Biotechnology, Gdańsk, Poland) following the manufacturer’s protocol. The isolated DNA was sequenced at Genomed S.A. (Warsaw, Poland) and compared with the GenBank using the BLAST [29] database to identify bacteria.

2.7. Minimal Inhibitory Concentration (MIC) of HMs

The MICs were determined by plating method using BA plates (BTL, Warsaw, Poland) supplemented with ZnNO₃ (Sigma-Aldrich, Inc., St. Louis, MO, USA), CdCl₂ (Sigma-Aldrich, Inc., St. Louis, MO, USA), CuCl₂ (Sigma-Aldrich, Inc., St. Louis, MO, USA) or NiCl₂ (Sigma-Aldrich, Inc., St. Louis, MO, USA) at different concentrations (e.g., 1–10 mM, 15 mM and 20 mM) with 200 µg/mL of Nystatin (Sigma-Aldrich, Inc., St. Louis, MO, USA). After inoculation, plates were incubated for 7 days at 22 °C. The MIC for each bacterial isolate was determined as the lowest concentration of HM at which no colonies were observed. Plates without added metals were used as positive control while Cupriavidus metallidurans AE104 was used as a negative control for MIC.

2.8. Microbial Community Structure

The structure of microbial communities in soil was estimated using phospholipid fatty acids (PLFAs) as described by other studies [30] with minor modifications. Lipids of microbial origin were extracted, fractioned, methylated and then analysed using gas chromatography (Hewlett-Packard 6890, Agilent, Santa Clara, CA, USA). Methylated fatty acids were identified and quantified using the MIDI-MIS software (Sherlock TSBA6 library, MIDI Inc., Newark, DE, USA) and nonadecanoic acid (19:0) as internal standard. Specific PLFAs for gram-negative bacteria (16:1ω7c, cy17:0, 18:1ω7, cy19:0) [31], gram-positive bacteria (i15:0, a15:0, i16:0, i17:0, a17:0) [31], actinomycetes (10Me17:0, 10Me18:0) [32] and fungi (18:2ω6,9) [33] were used to calculate the total microbial PLFAs and PLFAs for each group of microorganisms, expressed as nmol PLFA per g of dry weight of soil.

2.9. Microbial Community’s Physiological Profile

Microbial community’s catabolic abilities were determined using BIOLOG EcoPlates™ (Biolog Inc., Hayward, CA, USA). The soil solution was prepared with 10 g of dry soil and 90 mL of 0.85% NaCl. The samples were shaken for 2 h (140 rpm) and left for 30 min to settle. The 96-well BIOLOG microplates were inoculated with 120 μL (10⁻²) dilution aliquots of soil solution supernatant and incubated at 22 °C for 5 d. The readings were performed twice per day using a Bio-Tek ELx808 (BioTek-Agilent, Santa Clara, CA USA) microplate reader at 590 nm. The average well colour development (AWCD) was calculated at 144 h and equalled the optical density value of each well, corrected by subtracting the blank well using Equation (1) [34]:

$$\mathrm{AWCD}={\displaystyle \sum }{\mathrm{OD}}_{\mathrm{i}}/31$$

(1)

where AWCD is the average well colour development (units), ODi is optical density value from each well, corrected by subtracting the blank well (inoculated, but without a carbon source), and 31 is number of variables (C substrates).

The sum of the AWCD scores for individual substrates was used as a measurement of total microbial catabolic activity. All analyses were performed in triplicate.

2.10. Genetic Biodiversity of Microbial Community Structure

Total DNA was extracted from 2 × 0.5 g soil samples using a GeneMATRIX Soil DNA Purification Kit (EurX, Gdańsk, Poland) according to the manufacturer’s recommendations and combined in to one sample. The 16S rRNA gene fragment V3–V5 was amplified using primers MF-341-GC CGC CCG CCG CGC CCC GCG CCC GTC CCG CCG CCC CCG CCCG CCT ACG GGA GGC AGC AG with GC clamp [35,36] and MR-907 CCGTCAATTCMTTTGAGTTT [37]. The reaction mixture (25 µL) contained: 0.25 µL of primer MF-341-GC (100 µM, 1:10) (Sigma-Aldrich, Inc., St. Louis, MO, USA), 0.25 µL of primer MR-907 (100 µM, 1:10) (Sigma-Aldrich, Inc., St. Louis, MO, USA), 0.5 µL of polymerase DreamTaq^TM 500U (Thermo Fisher Scientific, Waltham, MA, USA), 2.5 µL of buffer 10× DreamTaq^TM (Thermo Fisher Scientific, Waltham, MA, USA), 0.5 µL of 10mM Mix dNTP (Thermo Fisher Scientific, Waltham, MA, USA), 4 µL of DNA and 17 µL of H₂O. The amplification conditions were: 94 °C for 5 min, and 36 cycles of 94 °C for 20 s, 58 °C for 20 s, 72 °C for 30 s with final 72 °C 5 min.

The electrophoresis of the PCR products was performed in 6% (w/v) polyacrylamide gel (37.5:1 acrylamide:bis-acrylamide) with a linear denaturing gradient ranging from 40 to 70% using DCode Mutation Detection System (Bio-Rad, Hercules, CA, USA). The band pattern was analysed using Image Lab 6.0.1 (Bio-Rad, Hercules, CA, USA) (see Figures S1 and S2 of the Supplementary Materials section). The selected bands were cut out, reamplified, and sequenced. Reamplification was performed using MF-341 CCTACGGGAGGCAGCAG and MR-907 CCGTCAATTCMTTTGAGTTT AG primers. The reaction mixture was prepared as described before and amplification conditions were: 94 °C for 5 min followed by 34 cycles of 94 °C for 20 s, 56 °C for 20 s, 72 °C for 30 s and finally 72 °C for 5 min. The biodiversity of the soil bacterial community was expressed as the Shannon-Wiener index (H’), richness (Rs), and evenness (E_H), calculated according to methodology reported elsewhere [38].

2.11. CadA, CzcA, and NccA Genes Detection

Soil isolated strains were tested for presence of cadA, czcA, and nccA genes using PCR. The DNA isolated from the bacteria using Genomic Mini (A&A Biotechnology, Gdańsk, Poland) according to the manufacturer’s instructions was the template for this reaction. The 605 bp fragment of cadA gene was amplified using degenerated primers cad1 5′ AAR ACI GGI ACI YTI ACI AAR GGI G 3′ and cad2 5′ GIG CRT CRT TIA CIC CRT CIC CIA 3′ [39]; the 581 bp nccA gene was amplified using 2555nccAF 5′AGCCGSGASAACGGCAAGCG 3′ and 3117nccAR 5′ CCGATCACCACCGTYGCCAG 3′; and the czcA gene with primers 1226czcAF 5′GACTTCGGCATCATCRTCGAYGG3′, 2026czcAR 5′CGTTGAASCGCARCTGGATCGG 3′ [40]. The reaction mixture (25 µL) contained 1.5 µL of MgCl2 (Thermo Fisher Scientific, Waltham, MA, USA), 2.5 µL of 10× buffer KCl (Thermo Fisher Scientific, Waltham, MA, USA), 0.5 µL of dNTP Mix (10 mM) (Thermo Fisher Scientific, Waltham, MA, USA), 0.25 µL of primer cad1 or 2555nccAF or 1226czcAF, respectively (100 µM, 1:10 v/v) (Sigma-Aldrich, USA), 0.25 µL of primer cad2, primer 3117nccAR or 2026czcAR 5′, respectively (100 µM, 1:10 v/v) ((Sigma-Aldrich, Inc., St. Louis, MO, USA), 0.5 µL of polymerase Taq (Thermo Fisher Scientific, Waltham, MA, USA), 3 µL of DNA and 16.5 µL of H₂O.

The amplification conditions were: 94 °C for 2 min, followed by 25 cycles at 94 °C for 1 min denaturation, 60 °C for 1 min annealing, 72 °C for 2 min and extension at 72 °C for 5 min. The pKPY11 plasmid with the cloned cadA gene was used as positive control [39].

The amplification conditions for nccA gene were: 94 °C for 2 min for initial denaturation, 25 cycles of 94 °C for 1 min, 59 °C for 1 min and 72 °C for 1 min followed by extension at 72 °C for 5 min. The positive control for the amplification was genomic DNA from C. metallidurans CH34 isolated with Genomic Mini (A&A Biotechnology, Gdańsk, Poland) according to the manufacturer’s instructions.

The czcA gene fragment was amplified using 94 °C for 2 min initial denaturation, 25 cycles at 94 °C for 1 min, 60 °C for 1 min, 72 °C for 2 min and a last extension of 72 °C for 5 min. The positive control for the amplification was genomic DNA from C. metallidurans CH34. The amplification specificity was checked through sequencing of PCR products. Briefly, PCR products were purified using Gel Out (A&A Biotechnology, Gdańsk, Poland) according to the manufacturer’s instruction. The eluted DNA were sequenced in Genomed S.A. (Warsaw, Poland) and BLAST with the GeneBank database.

2.12. Statistical Analysis

Statistically significant differences between samples were calculated using one-way ANOVA and post-hoc Tukey test (p < 0.05) with Excel and Real Statistics 2020. Kendall Tau correlation was used to investigate the effect of environmental factors on microbial communities.

3. Results

3.1. Physicochemical Analysis of Soil Samples

The results of physicochemical analysis of soil samples are summarised in

Table 1. Physicochemical characterization of soil samples.

	Piekary Śląskie			Bukowno
	P1	P2	P3	B1	B2	B3
Water content (%)	26.21 ± 0.24 b	29.48 ± 0.12 a	8.54 ± 0.25 c	5.01 ± 0.28 d	5.34 ± 0.36 d	1.56 ± 0.21 e
pH (H20)	7.71 ± 0.03 b	8.14 ± 0.02 a	7.62 ± 0.01 c	6.79 ± 0.01 e	6.96 ± 0.01 d	5.49 ± 0.01 f
pH (KCl)	8 ± 0.03 b	8.29 ± 0.01 a	7.15 ± 0.01 c	6.77 ± 0.01 e	6.81 ± 0.01 d	4.69 ± 0.01 f
pH (CaCl2)	7.84 ± 0.01 b	8.15 ± 0.01 a	7.04 ± 0.01 c	6.78 ± 0.01 e	6.82 ± 0.01 d	5.07 ± 0.01 f
Total N (mg/kg)	829.9 ± 35.21 a	780.8 ± 15.41 a	997.8 ± 19.28 a	747.97 ± 950.41 a	215.63 ± 6.15 a	151.2 ± 7.81 a
Organic matter (%)	7.76 ± 0.02 b	7.89 ± 0.06 b	10.95 ± 0.55 a	2.62 ± 0.04 c	2.23 ± 0.07 c	0.54 ± 0.04 d
Organic carbon (%)	4.5 ± 0.01 b	4.58 ± 0.03 b	6.35 ± 0.32 a	1.52 ± 0.02 c	1.3 ± 0.04 c	0.32 ± 0.02 d
P (%)	0.83 ± 0.03 a	1.14 ± 0.04 b	0.38 ± 0.01 c	0.2 ± 0.02 d	0.29 ± 0.01 e	0.24 ± 0.01de
Na (mg/kg)	547.77 ± 23.24 a	397.67 ± 63.28 b	131.1 ± 5.11 c	112.3 ± 10.87 cd	38.6 ± 6.35 de	15.17 ± 0.42 e
Mg (g/kg)	37.486 ± 0.877 a	14.965 ± 0.567 b	5.865 ± 0.636 c	19.257 ± 0.024 d	3.420 ± 0.441 e	0.277 ± 0.024 f
K (g/kg)	0.708 ± 0.152 c	1.093 ± 0.055 ab	1.219 ± 0.059 a	0.962 ± 0.111 b	0.333 ± 0.076 d	0.114 ± 0.056 d
Ca (g/kg)	109.124 ± 2.746 a	36.105 ± 3.384 b	9.396 ± 0.425 d	24.361 ± 1.402 c	3.230 ± 0.285 e	0.274 ± 0.049 e
Zn total (g/kg)	1.208 ± 0.106 b	1.282 ± 0.205 b	1.035 ± 0.083 b	1.8143 ± 0.00 a	1.777 ± 0.00 b	14.362 ± 0.00 b
Zn (H20) (mg/kg)	0.49 ± 0.3 f	1.14 ± 0.69 def	3.47 ± 1.54 c	19.38 ± 0.00 a	1.98 ± 0.00 cd	6.25 ± 0.00 b
Zn (CaCl2) (mg/kg)	0.34 ± 0.12 e	0.35 ± 0.11 e	7.92 ± 0.88 d	56.21 ± 0.00 a	14.34 ± 0.00 b	21.43 ± 0.00 c
Cd total (g/kg)	0.293 ± 0.016 c	1.215 ± 0.119 b	0.0459 ± 0.006 d	1.991 ± 0.00 a	0.100 ± 0.00 d	0.017 ± 0.00 d
Cd (H20) (mg/kg)	0.06 ± 0.03 c	0.14 ± 0.07 b	0.03 ± 0.00 c	0.51 ± 0.00 a	0.02 ± 0.00 c	0.01 ± 0.00 c
Cd (CaCl2) (mg/kg)	0.05 ± 0.01 e	0.11 ± 0.03 d	0.24 ± 0.01 c	3.32 ± 0.00 a	0.49 ± 0.00 c	0.21 ± 0.00 b
Ni total (mg/kg)	3.94 ± 0.08 d	3 ± 0.22 d	0.62 ± 0.08 e	54.16 ± 0.00 a	44.63 ± 0.00 b	15.25 ± 0.00 c
Ni (H20) (mg/kg)	0.001 ± 0.001 e	0.002 ± 0.001 de	0.003 ± 0.001 d	0.02 ± 0.00 c	0.04 ± 0.00 a	0.03 ± 0.00 b
Ni (CaCl2) (mg/kg)	0.001 ± 0.002 a	0.00	0.00	0.01 ± 0.00 c	0.02 ± 0.00 b	0.02 ± 0.00 b
Cu total (mg/kg)	580.68 ± 59.12 b	175.65 ± 13.7 c	14.59 ± 0.69 d	1116.87 ± 0.0 a	75.81 ± 0.00 d	0.00 ± 0.00
Cu (H20) (mg/kg)	0.04 ± 0.02 a	0.05 ± 0.01 a	0.03 ± 0.02 a	0.00 ± 0.00 b	0.02 ± 0.00 a	0.01 ± 0.00 a
Cu (CaCl2) (mg/kg)	0.02 ± 0.01 b	0.05 ± 0.00 a	0.01 ± 0.01b	0.01 ± 0.00 b	0.00 ± 0.00 b	0.01 ± 0.00 b

The data represent the mean and standard deviation of three replicates. P1, P2 and P3 indicate individual sampling sites from Piekary Śląskie; and B1, B2 and B3 from Bukowno. Data with different lower-case letters indicate statistically significant differences between results of particular parameter (one-way ANOVA, Tukey test, p < 0.05) and the same letter refers to no significant difference.

. As shown, samples significantly differed in water content, with pH values ranging from slightly acidic to slightly alkaline. The characteristic feature of the tested soils was the high content of macro elements (i.e., Ca, Mg, Na, K and total N, see Table 1). The total content of HMs in different soil samples significantly exceeded acceptable standards for Cd and Zn (Table 1). The only elements whose values remained within acceptable limits were Ni and Cu (with the exception of Cu at point B1).

3.2. Enumeration of Metal Tolerant Microorganisms in Soil Samples

From samples collected at Piekary Śląskie, 64–77% out of the total oligotrophic bacteria (OB) was identified as metal tolerant but no oligotrophic Cu-tolerant bacteria were detected. Neither oligotrophic Ni- and Cd-tolerating bacteria were found in P1, nor Cd-tolerating bacteria in soil sample P2. In soil samples from Bukowno, no Cu-tolerant OB were found. As in the case of Piekary Śląskie, the number of oligotrophic bacteria that tolerated Zn and Ni accounted for 64–93% out of the total bacterial counts from Bukowno.

Metal tolerant bacteria in soil from Piekary Śląskie accounted for 71–96% of all bacteria grown on TSA medium (Heterotrophic bacteria, HB) and 67–89% of bacteria grown on agar broth medium. Cd-tolerant bacteria were found in soil samples from Bukowno (B1 and B2) but Zn-tolerant bacteria were absent in the B2 sample. The number of metallotolerant bacteria in Bukowno samples grown on TSA medium ranged from 73% to 99% (Heterotrophic bacteria, HB). In the case of bacteria grown on agar broth, metal-resistant bacteria accounted for 67–89% of all bacteria (

Table 2. Metallotolerant bacteria enumeration in soil samples.

Bacteria Count,		Piekary Śląskie			Bukowno
(log10 CFU/g Dry Soil)		P1	P2	P3	B1	B2	B3
OB	Total	4.44 ± 0.09 ce	4.79 ± 0.14 cd	4.91 ± 0.03 c	3.86 ± 0.04 f	5.73 ± 0.43 a	5.53 ± 0.16 ab
	Cu	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00
	Cd	0.00 ± 0.00	0.00 ± 0.00	3.80 ± 0.14 a	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00
	Ni	0.00 ± 0.00	3.51 ± 0.17 a	3.44 ± 0.35 ab	2.48 ± 0.15 c	0.00 ± 0.00	0.00 ± 0.00
	Zn	3.10 ±0.00 a	3.11 ± 0.00 a	3.67 ± 0.31 a	3.59 ± 0.03 a	4.58 ± 0.49 a	2.83 ± 2.46 a
HB (TSA)	Total	4.35 ± 0.08 cde	4.50 ± 0.04 ac	4.64 ± 0.67 ab	3.71 ± 0.09 f	4.38 ± 0.10 acd	4.73 ± 0.67 a
	Cu	3.98 ± 0.34 ce	4.26 ± 0.06 c	4.38 ± 0.65 ab	3.28 ± 0.24 f	4.56 ± 0.21 a	4.19 ± 0.63 cd
	Cd	3.10 ± 0.00 a	3.59 ± 0.31a	3.53 ± 0.52 a	0.00 ± 0.00	0.00 ± 0.00	3.47 ± 0.54 a
	Ni	3.99 ± 0.14 a	4.33 ± 0.25 a	4.42 ± 0.65 a	3.87 ± 0.89 a	5.12 ± 0.70 a	4.66 ± 0.65 a
	Zn	3.86 ± 0.28 a	3.47 ± 0.39 a	4.28 ± 0.63 a	3.87 ± 2.06 a	4.64 ± 0.83 a	4.63 ± 0.65 a
HB (Broth agar)	Total	4.65 ± 0.10 ade	4.77 ± 0.07 ac	4.80 ± 0.24 ab	4.65 ± 0.10 ef	4.77 ± 0.07 ad	4.80 ± 0.24 a
	Cu	4.14 ± 0.07 a	3.99 ± 0.09 a	4.03 ± 0.61 a	4.14 ± 0.07 a	3.99 ± 0.09 a	4.03 ± 0.61 a
	Cd	3.33 ± 0.40 a	3.21 ± 0.17 a	4.07 ± 0.22 a	3.33 ± 0.40 a	3.21 ± 0.17 a	4.07 ± 0.22 a
	Ni	3.99 ± 0.38 ade	4.23 ± 0.26 ac	4.16 ± 0.30 ad	3.99 ± 0.38 ef	4.23 ± 0.26 a	4.16 ± 0.30 ab
	Zn	3.77 ± 0.06 a	3.65 ± 0.21 a	4.27 ± 0.05 a	3.77 ± 0.06 a	3.65 ± 0.21 a	4.27 ± 0.05 a
RT-PCR	Total	9.04 ± 0.08 bc	9.11 ± 0.02 ab	9.16 ± 0.03 a	9.02 ± 0.03 bd	9.06 ± 0.03 abd	8.99 ± 0.03 cd

The data represent the mean and standard deviation of three replicates. P1, P2 and P3 indicate individual sampling sites from Piekary Śląskie; and B1, B2 and B3 from Bukowno. T: total number of heterotrophic bacteria; Cu, Cd, Ni, Zn: the number of Cu-, Cd-, Ni- and Zn-tolerant bacteria. Data with different lower-case letters indicate statistically significant differences between results of particular parameter (one-way ANOVA, Tukey test, p < 0.05) and the same letter refers to no significant difference.

). The use of real-time PCR (RT-PCR) allowed the assessment of the total bacterial number including culturable and non-cultured fractions. Using this method, bacterial numbers were estimated to be over twice as large compared to bacteria able to grow on culture media (Table 2).

3.3. Bacteria Isolated from Soil Samples

A significant number of metallotolerant bacteria were isolated from soil samples obtained from Piekary Ślaskie and Bukowno sampling sites (18 and 12, respectively), according to their 16S rRNA sequence (Table S1 in Supplementary Materials). The different isolates were classified into three phyla: Proteobacteria, Actinobacteria and Bacteroidetes. Proteobacteria phylum found in the soil samples was represented by Methylobacterium from the Alpha-Proteobacteria class, Variovorax from the Betaproteobacteria class, Pseudomonas from the Gammaproteobacteria and Ralstonia classes, and Cupriavidus from the Betaproteobacteria class. The Actinobacteria phylum was represented by three species from the class Actinobacteria (i.e., Arthrobacter, Rhodococcus, Agromyces). The Bacteroidetes phylum was represented by Chryseobacterium from the Flavobacterium class.

Most isolated bacteria were tolerant towards all selected HMs. Special attention should be given to Rhodococcus erythropolis strain P3 4 TSA, which showed tolerance up to 15 mM Zn and 6 mM Cd, Arthrobacter nitroguajacolicus P2 1 TSA tolerated 10 mM Zn and 6 mM Cd, and Arthrobacter oryzae P1 2 TSA, which enabled growth on media containing 10 mM Zn and 4 mM Cd—all of which were isolated from the Piekary Śląskie sampling site. High MIC values were also noted for bacteria isolated from Bukowno. For example, Agromyces sp. p2ba, Arthrobacter nitroguajacolicus ba O3 and Chryseobacterium sp. 1/10 la1 were found to be tolerant of a wide variety of conditions (Table S1 in Supplementary Materials).

3.4. Microorganisms Structure and Metabolic Activity

PLFA analysis showed gram negative bacteria as the dominant group in the different samples, constituting 46–50% of microorganisms in Piekary Śląskie and 47–60% in Bukowno. Actinomycetes were the least numerous phylum of microorganisms in all tested soils (

Table 3. Metabolic activity of microorganisms in soil samples.

Method				Sampling Site
		P1	P2	P3	B1	B2	B3
FDA (µg fluorescein/g dry soil)		0.23 ± 0.08 b	0.27 ± 0.06 b	0.58 ± 0.1 a	0.28 ± 0.1 b	0.15 ± 0.04 b	0.56 ± 0.07 a
PLFA	Total, nmol PLFA/g dry soil	2.83 ± 0.76 bc	8.27 ± 0.95 ac	12.64 ± 1.70 a	5.88 ± 5.89 ac	10.38 ± 1.26 ac	77.62 ± 2.78 ac
	Gram + (%)	38.74 ± 3.75 a	41.54 ± 0.46 a	38.84 ± 0.25 a	32.57 ± 19.47 a	38.19 ± 2.36 a	37.93 ± 3.01 a
	Gram − (%)	46.04 ± 4.33 a	50.49 ± 0.57 a	47.14 ± 0.65 a	59.84 ± 18.32 a	47.75 ± 1.73 a	47.33 ± 2.43 a
	Actinomycetes (%)	15.21 ± 1.91 a	4.65 ± 0.07 a	9.23 ± 0.53 a	6.71 ± 7.39 a	8.23 ± 0.81 a	8.43 ± 1.59 a
	Fungi (%)	0.00 ± 0.00	3.31 ± 0.67 a	4.80 ± 0.12 a	0.88 ± 0.76 a	5.83 ± 0.18 a	6.31 ± 1.37 a
BIOLOG	AWCD	0.82 ± 0.05 ecd	0.78 ± 0.04 cd	1.15 ± 0.17 f	0.09 ± 0.02 a	0.82 ± 0.08 d	1.38 ± 0.07 b
	Carbohydrates (%)	32.87 ± 7.14 b	19.18 ± 1.53 a	24.60 ± 0.10 b	61.69 ± 3.17 b	18.41 ± 2.74 b	25.05 ± 1.99 b
	Carboxylic Acids (%)	4.41 ± 3.81 cdf	17.09 ± 0.46 c	15.42 ± 1.13 cd	7.67 ± 5.37 ce	8.13 ± 3.66 b	18.97 ± 2.07 a
	Amino acids (%)	31.42 ± 3.52 cf	35.48 ± 0.34 ce	30.11 ± 1.85 c	12.16 ± 1.82 cd	41.28 ± 4.47 b	26.94 ± 1.13 a
	Polymers (%)	17.98 ± 0.66 c	16.81 ± 3.78 cd	16.45 ± 0.98 ce	12.24 ± 6.99 cf	18.90 ± 1.92 b	13.52 ± 0.58 a
	Amines and amides (%)	5.58 ± 3.97 bcd	4.36 ± 0.76 bde	5.99 ± 0.40 cb	2.58 ± 3.00 bdf	5.83 ± 0.66 b	7.93 ± 3.40 a
	Miscellaneous (%)	7.74 ± 5.31 c	7.07 ± 3.19 ce	7.44 ± 2.69 cd	3.66 ± 2.56 cf	7.45 ± 1.55 b	7.60 ± 2.27 a
DGGE	H’	1.02 ± 0.08 fde	0.70±0.02 b	0.95±0.04 d	1.01±0.05 ed	0.49±0.03 cb	0.04 ± 0.00 a
	Rs	26 ± 0.00 a	21 ±0.00 a	21 ± 0.00 a	18 ± 0.00 a	24 ± 0.00 a	24 ± 0.00 a
	EH	0.31 ± 0.02 ed	0.23 ± 0.01 cb	0.31±0.01 dc	0.35±0.02 fe	0.23 ± 0.01 b	0.01 ± 0.00 a

The data represent the mean and standard deviation of three replicates. P1, P2 and P3 indicate individual sampling sites from PiekaryŚląskie; and B1, B2 and B3 from Bukowno. Data with different lower-case letters indicate statistically significant differences between results of particular parameter (one-way ANOVA, Tukey test, p < 0.05) and the same letter refers to no significant difference. AWCD (Average Well Colour Development).

). Despite HM contamination, all soil samples showed microbial enzymatic activity (FDA) and microorganisms were found able to break down carbohydrates, carboxylic acids, amino acids, polymers, amines and amides. Their potential to degrade individual carbon substrates, however, varied significantly among tested microbial communities (BIOLOG) (Table 3). Carbohydrates and amino acids were identified as the most important carbon source for microorganisms in Piekary Śląskie and Bukowno. Amines and amides were the least metabolised and constituted 3–7% of the used carbon sources (Table 3).

Genetic structure analysis suggested that Proteobacteria, Gemmatimonadetes and Acidobacteria were present in all studied soils. Representative types Chloroflexi and Actinobacteria were detected in soil from Piekary Śląskie, while in Bukowno sampling sites, Bacteroidetes and Firmicutes were found (Table S2 of Supplementary Materials).

3.5. Genes Encoding Mechanisms of HM Tolerance

Detection of CzcA, NccA and CadA genes, known as part of HM tolerance mechanisms [41], showed that the czcA (822 kb) gene encoding for Co-, Zn- and Cd-tolerance was present in P1 3 1/10 Variovorax paradoxus, P3 3 TSA Variovorax paradoxus, P2 4 ba Arthrobacter sp., P3 1 ba Variovorax sp. and P3 2 ba Arthrobacter sp. isolated from Piekary Śląskie samples, as well as in ba 03 Arthrobacter nitroguajacolicus, 1/10 p2ba Variovorax boronicumulans, TSA 07 Pseudomonas ficuserectae and 1/10 O1 Methylobacterium zatmanii isolated from soil samples from Bukowno. The presence of the nccA (581 kb) gene encoding tolerance for Ni, Co and Cd was found in P1 2 TSA Arthrobacter oryzae (Piekary Śląskie), and ba 03 Arthrobacter nitroguajacolicus, TSA O6 Arthrobacter oxydans, TSA 05 Pseudomonas sp., and1/10 O1 Variovorax boronicumulans (Bukowno). The cadA (1058 kb) gene, determining tolerance to Cd, Zn and Co, was only identified in TSA la3 Pseudomonas sp. (Bukowno). The bands were sequenced and compared with the sequences deposited in the GenBank database (Figure S3 of the Supplementary Materials).

3.6. Kendall Tau Correlation

Interaction between physicochemical and biological soil parameters was tested using the Kendall Tau correlation, Figures S4 and S5 (Supplementary Materials). As shown, negative correlation was observed between soil water content, pH and bacteria cultured on TSA medium, as well as Cd-tolerant and Zn-tolerant cultured bacteria on broth agar, and Zn-tolerant media on 1/10 TSA medium in samples from Piekary Śląskie (Figure S4, Supplementary Materials). The Zn and Cu content negatively affected the number of microorganisms as shown by PLFA and qPCR analysis, as well as culturable fraction of bacteria (TSA medium and 1/10 TSA). These elements also had a negative effect on FDA, BIOLOG index (AUC).

A positive correlation was found between the bioavailable heavy metal fraction (i.e., Zn, Cd) and PLFA (i.e., gram-positive, gram-negative, fungi, actinomycetes) as well as FDA, AUC and Rs determined by the BIOLOG method. Bioavailable Ni and Cd also affected qPCR values in the same way (Figure S4, Supplementary Materials).

In soils collected from Bukowno, soil moisture and pH negatively affected the number of microorganisms (TSA total), soil microorganism activity (FDA), and their ability to break down substrates (BIOLOG). Bioavailable Ni was found positively correlated with the number of cultured microorganisms, PLFA (gram-positive bacteria, actinomycetes, fungi) and microorganism biodiversity determined by DGGE, FDA, as well as parameters from the BIOLOG method (i.e., AUC, H and Rs) (Figure S5, Supplementary Materials).

4. Discussion

4.1. Physicochemical Characterization of Soils

Total metal content in soil is a useful indicator of the overall degree of contamination. It does not, however, provide information about metal mobility, bioavailability and toxicity [42]. Some studies have shown that both total and bioavailable HMs fraction shape microbial communities in contaminated sites [43]. The bioavailable HM fraction found in soil samples from Piekary Śląskie and Bukowno was relatively small compared to the HM total content, probably linked to soil pH, and other key factors determining HM availability in soil. Low pH values decrease HM absorption onto clay minerals, hydrated oxides and organic surfaces [44]. Nevertheless, other physicochemical soil parameters may also contribute to a negative HM effect on soil microbial communities. Results of this research agree with other studies [17,36,45] suggesting that pH and water content have a significant impact on structure and growth of microbial communities (PLFA and culturable bacterial fraction) and are negatively correlated to several biological processes, including enzymatic (FDA) and metabolic activity (BIOLOG), as well as HM availability. The SOM was found not only to be a carbon source but also to influence HM mobility as suggested in the past [42]. The SOM content in both studied soils was low, suggesting that other forms of metal precipitation or immobilisation may occur in the study area. The high amount of nitrogen in soil suggested low enzymatic activity involved in denitrification. Negative effect of Cd on nitrogen transformation has been reported in the past [46] and other studies have suggested that cadmium salts at a concentration of 200 mg/kg soil inhibited denitrification by almost 80% [47] in line with our results.

The studied soils were also characterised by a high content of macro- and microelement which, in addition to their biological role, can generate precipitation of nutrients. For example, Ca may generate phosphorus precipitation, necessary for microorganisms to grow [48]. Our results suggest that, regardless of HM contamination, other factors such as essential nutrient precipitation might influence growth and composition of microbial communities in the study area.

4.2. The Influence of HMs on Bacterial Counts

Over 60% of the total heterotrophic bacteria count showed tolerance towards selected HMs, independently of the collection site. Presence of metal-tolerant bacteria in soils exposed to prolonged HM exposure has been repeatedly reported in the past [15,49]. For example, metal-tolerant bacteria have been found accounting for almost 80% of total soil bacterial number in both laboratory and field experiments [50]. Long-term HM soil contamination undoubtedly leads to selection of microorganisms that possess appropriate mechanisms to enable them to survive under these stressful conditions [51]. The number of soil bacteria calculated by real-time PCR was twice as high as those obtained by culture methods, suggesting that non-cultivated bacteria constitute a large group of soil microbiome. Similar differences between total bacteria number measured either by plate count or real-time PCR methods were also noted [52].

4.3. Microbial Community Structure and Activity in HM-Contaminated Soils

As determined by PLFA, gram-negative bacteria dominated in all tested soils. Nevertheless, gram-positive bacteria also constituted a relatively large group. Actinomycetes accounted for the smallest microorganism group. Fungi are considered by some authors to have higher HM resistance and to be even capable of increasing their number under certain conditions [53]. In this work, however, only small amounts of fungal biomarkers were observed compared to bacteria biomarkers.

This type of controversial observation is frequently discussed. For example, the negative impact of Pb on fungi and fungal colony-forming units (CFUs) of PLFA biomarkers has been reported in the past [54]. Similarly, in another study [55], soil fungal population showed higher sensitivity to HM contamination than bacteria. In contrast, a higher fungal tolerance to metals was observed compared to bacteria by another study [56]. These apparently conflicting findings can be explained by the effect of other factors, such as soil pH [57], nutrient content [58], or presence of other contaminants [59] which can mask the harmful effects of HMs.

The test used for microorganism genetic biodiversity (PCR-DGGE) is particularly useful in environments with small biodiversity, and allows accumulation of data on non-culturable fractions known to be dominant in soil [60]. In soil samples from Piekary Śląskie, Proteobacteria, Chloroflexi, Acidobacteria and Gemmatimonadetes outweighed the others. For samples from Bukowno, Bacteroidetes, Firmicutes, Acidobacteria, Proteobacteria and Gemmatimonadetes were the dominant microorganisms. Similar bacteria groups were found using DGGE analysis in other HM contaminated areas. For example, Bacteroidetes, Acidobacteria, Gemmatimonadetes, Actinobacteria and Betaproteobacteria, with predominance of the former three, were observed in HM-polluted soils from Katowice, Poland [61]. Presence of Acidobacteria, Betaproteobacteria, Gammaproteobacteria and Chloroflexi in long-term HM contaminated soils was also reported by other studies [13].

Bacteria belonging to the Gemmatimonadetes are considered to make up 2% of all bacteria in soil. This phylum is often found in HM contaminated areas [62,63] and it is mostly detected by non-culturing methods [64]. Prevalence of bacteria from this phylum in soils is probably related to their ability to adapt to low water content and neutral or slightly acidic soil pH [64].

Moreover, Acidobacteria is a large, diverse phylum found in different soils with most of the bacteria being non-culturable [65,66], the abundance of which is also correlated with soil pH [67,68]. Chloroflexi phylum members have been reported to colonise a variety of environments, including contaminated soils, while being mainly non-culturable in agreement with our results [62,63,69,70]. Firmicutes have been found as the dominant phylum in Cr-contaminated environments [71] while Proteobacteria is a well-known ubiquitous phylum among HM contaminated soils [72,73], in line with our results using the DGGE method. Microorganism metabolic activity analysis using BIOLOG suggested that carbohydrates and amino acids were the main carbon source while amines and amides were metabolised to the least extent, which agrees with similar observations of carbohydrates and amino acids usage as a primary carbon source in HM contaminated areas [74].

4.4. Isolated Bacteria Characteristics

Proteobacteria, Bacteroides and Actinobacteria were detected using culturable methods. Actinobacteria represent one of the most dominant groups of microorganisms and is the most effective in decomposing organic matter, including hemicellulose and lignin [75,76]. Other studies have suggested that, among others, Actinobacteria tolerate much higher HM concentrations than any other soil bacteria for the same conditions [62,63,69,76]. Most bacteria isolated from both studied areas were found tolerant to at least three of the selected HMs tested (i.e., Cd, Cu, Ni, Zn), in addition to reaching high MIC values. We have also been able to show the genes encoding the mechanisms of tolerance to HMs in isolated bacteria. Thus, czcA and nccA were detected in Variovorax sp., Arthrobacter sp., Pseudomonas sp. and Methylobacterium, while cadA was found in Pseudomonas sp. These same genes have been reported in other studies using culturable techniques by other authors [40,77,78]. Their absence in other bacteria inhabiting tested soils suggests presence of alternative mechanisms enabling them to survive under these conditions, including transporting HM out of the cell, metal precipitation outside or inside the cell, or HM binding through exopolysaccharides [79].

Some of the isolated bacteria were well-known for their high metabolic potential and HM tolerance, such as Cupriavidus sp. [80], Arthrobacter sp. [81], Variovorax sp. [82] and Pseudomonas sp. [83]. Because of their characteristics and origin, these microorganisms can be used in HM contaminated area restoration or laboratory-scale tests to identify the mechanisms enabling them to tolerate high HM concentrations. Furthermore, some of these bacteria (e.g., Agromyces or Variovorax sp.) remain poorly understood, therefore their isolation from soil and genome sequencing is expected to allow for better understanding of their role in contaminated environments.

5. Conclusions

It has been demonstrated in this study that the structure and function of microbial communities in both tested contaminated soils were shaped not only by the specific HM and its concentration, but also by the other physicochemical features of the soil.

It seems that pH and water were the key factors affecting not only the biological part of the soil, but also its structural properties such as bioavailability of HMs. Other factors including low amounts of SOM and high concentration of microelements together with HMs contributed in determining the microbial communities in this area.

Undoubtedly long-term contamination of the soils with HMs has led to the development and domination of metal tolerant fractions of bacteria in these areas. In addition, the non-culturable fraction of bacteria was found to be a significant component of microbial communities in the HM-polluted soils, although they are omitted by the traditional methods.

Nevertheless, culture techniques still having high value, as isolated bacteria may be used in bioremediation of contaminated sites and this method, being environmentally friendly, is currently favoured by many researchers.