Bacterial diversity assessment of different soil types from eastern states of India using 16S r DNA sequencing approach.

*Spandan Chaudhary¹, Ravi Shaliwal¹, Pooja Chaudhary¹, Hiren Gajjar¹, Shiv Patel¹, Richa Misra¹, Pushprajsinh Chauhan¹, Jayesh Jalondhara¹, Harita Pandit¹ and Prashanth Bagali¹.

¹Xcelris Labs Limited, 2nd Floor, Heritage Profile, 22-23, Shrimali Society, Opp. Navrangpura Police Station, Navrangpura, Ahmedabad - 380009, Gujarat, INDIA

Yanshan Chen(chenyanshan@nju.edu.cn)

Paola Arfaioli(paola.arfaioli@unifi.it)

Qingye Sun(sunqingye@ahu.edu.cn)

Zimin Wei(weizimin@neau.edu.cn)

Spandan Chaudhary, Bacterial diversity assessment of different soil types from eastern states of India using 16S r DNA sequencing approach.(2017)SDRP Journal of Earth Sciences & Environmental Studies 2(4)

Soil is one of the reservoirs of the diverse group of microorganisms including bacteria, archeae, fungi etc on the earth. Unculturable bacteria are of prime focus in scientific community because of their vital role in production of many important enzymes and degradation capabilities. Mostly, researcher through out the world using NGS technologies to decode their genome which is costly when multiple isolates to be tested at initial level. In order to answer this question, we have tried gold standard Sanger sequencing techniques to sequence metagenomic DNA extracted from soil. We used 16s rDNA approach and found microbes in the samples which are involved in processes such as Nitrogen fixation, oil-bioremadiation, reduction of sulphate compounds, decomposition of aromatic compounds and hydrocarbons, removal of toxic metals etc., to maintain and increase sustainability of environment. This is one of very few studies that have focused on metagenome diversity using sanger technology. This proves use of first generation technology as initial screening of metagenomic samples with desired microbes before going for comparatively expensive NGS sequencing.

Soil is one of the reservoirs of the diverse group of microorganisms including bacteria, archeae, fungi etc on the earth [1,2]. These microorganisms perform the key processes that create and maintain the environment in both terrestrial and aquatic systems. They also play an essential role in survival of other organisms on earth such as nitrogen fixing rhizobacteria in rhizospheric soil fix atmospheric nitrogen to make available it to the plants which utilize nitrogen as macro-nutrient, microorganisms which are found in the soil on polluted sites like sewage dumping site, wastelands, oil-contaminated sites etc. have the potential for conversion of complex organic and inorganic compounds  in their simpler forms, thus, making their degradation easier and  render them less harmful to environment as well as for live beings on earth [3-6]. But this diversity of microorganisms in soil is still in its infancy in terms of accessibility because most of the soil microorganisms can’t be cultured and isolated on standard solid or liquid media and by the current laboratory techniques because we lack critical information on their biology, and this presents both challenges and opportunities [7]. So, the hidden treasure of valuable biological information contained in genome of microorganisms is yet to be revealed because microbial diversity found in soil environment is of immense importance regarding in the molecular aspects like identification and exploitation of specific microorganism for their genome which contains genes coding for products/processes characteristic to that microorganism. These products includes enzymes, antibiotics as well as other organic substances, bioremediation of polluted lands, removal of toxic metals(such as Chromium, Arsenic, Mercury etc.) and compounds, inhibition of pathogenic as well as harmful organism, development of treatment measures for diseases in plants as well as animals to increase productivity and yield. Microbial secondary metabolites are used in organ transplantation, cancer treatment and cholesterol control, as well as serving as insecticides, fungicides etc. Almost every aspect of human health would benefit from a greater diversity and availability of microbial natural products [8-12].

To obtain above advantage, the very first step is to isolate the microorganisms from soil environment and their further screening for information, we need. For this, One can use traditional biochemical tests [13] but the recent molecular biology techniques such as pulsed field gel electrophoresis(PFGE) of whole chromosomal DNA, RAPD & AFLP assays, 16S rDNA analysis, Real Time PCR and microarray based bacterial identification methods have been proved more promising [14-20]. All these methods has revolutionized the environmental microbiology [21]. Dominant species of microorganisms that best adapts to the ecosystem can be more prominently detected using these techniques as metabolically active microorganisms contain more DNA and RNA. In other words ecologically important microorganisms are assessed with molecular techniques and not the inactive ones which do not contribute to ecosystem functions. Another major advantage of these techniques is that microbial communities can be studied without actually cultivating the microorganisms thereby preserving the in situ metabolic status and composition.  The new analytical approaches using DNA extracted directly from the environment enable us to access the genome, which is called metagenome, of all microorganisms inhabiting in the environment [22]. It made capable the researchers to develop effective and efficient culture-independent techniques for direct isolation and further screening and analysis of microbial diversity by using 16S rDNA methods  provide the faster way to identify a specific group as well as diversity of microorganisms in soil and to correlate them to nearest species [23]. In addition to this, many researchers have been used and/or are using 16S rDNA sequences as a tool for taxonomic classification, usually where phenetic methods have proved lengthy and inconclusive [24].

In the present work, a modified protocol of CTAB-method for microbial genomic DNA isolation from different types of soils including agricultural, oil-contaminated, sewage, polluted water, desert soil etc., has been standardized and it is validated by analysis of microbial diversity in each type of extracted genomic DNA sample. The meta-genomic DNA was amplified by using the eubacterial primers designed from variable region of 16S rDNA sequences, the amplified PCR products were sequenced on ABI 3730XL.

2.1 Site and sample description

Different types of soil samples were collected from different sites as mentioned in Table 1.

2.2 Extraction of soil gDNA

Soil gDNA was extracted using modified CTAB protocol (TES-CTAB method) which consists of two phases (a) sample processing and (b) DNA isolation. Sample processing consist of washing of 0.5g to 5g weighed soil (depending upon soil type capable to form pellet after centrifuge) in 50ml1x Phosphate buffer, filtration of soil suspension with filter paper, centrifugation of filtrate at 4000 rpm for 20 minutes at 4°C pellet out the cells attached to soil particles, subsequently dissolving the pellet in to 4ml pre-warmed TES Buffer (100mM Tris, 10mM EDTA, 2% [w/v] sodium dodecyl sulphate, pH 8.0) and distributing it in four tubes. For DNA isolation one of the above tubes was taken and 10µl proteinase K (20mg/ml) is added and tube was incubated for 1 hour at 60°C. Further, 250µl 2% CTAB and 140µl 5M NaCl was added and incubated at 65° C for 20 minutes. 20µl RNase A (20mg/ml) was added in tube and incubated at 37° C for 45 minutes to remove RNA which usually precipitate with DNA. The lysed cells containing DNA in solution was divided into two tubes and mixed with equal volume of chloroform-isoamyl alcohol (24:1, v/v). The aqueous phase containing DNA was recovered by centrifugation at 14,000rpm at room temperature and precipitated with 0.1 volumes 3 M Sodium Acetate (pH-5.5) and 0.6 volume of isopropanol at -20°C for 1-2 hour or at -80°C for half an hour. Pellet of crude nucleic acid was obtained by centrifugation at 14,000rpm for 10 minutes, washed with cold 70 % ethanol and resuspended in sterile nuclease free water to a final volume of 30-50µl. These genomic DNA were quantified by Nanodrop 8000 (Thermoscientific) spectrophotometer. The purity of the extracted DNA was confirmed by running 1 to 2.5μg DNA separately from each sample on agarose gel electrophoresis set at 110V for 30-40 minutes at 25°C. The resultant DNA bands were visualized using Gel-Doc (Bio-Rad).

2.3 PCR Amplification of Extracted Soil DNA

 DNA samples of each type of soil were amplified using eubacterial variable region 16S rRNA gene primes V₅F: 5'AAACTYAAARRATTGACGGG3' as forward primer and   V₆R:5'CGACRRCCATGCANCACCT3' as reverse primer specific for Bacteria. The PCR amplification was carried out in Eppendorf Thermal cycler with 20 μl of final reaction volume containing 16.0μl DNase-RNase free water, 4μl 5XPCR  reaction buffer (Roche),1.0μl DMSO, 1.0μl BSA, 0.02μl Taq DNA Polymerase mix(Roche), 1.0 μl forward primer V₅F, 1.0 μl reverse primer and 0.8 to 1.5μl diluted DNA(30ng/μl). The PCR was initiated with initial denaturation of DNA at 95°C for 5min and subsequently the number of cycles (94°C for 30s, 47°C for 30s and 72°C for 1min) were set to 35, and the final extension was performed at 72°C for 10min. 5 μl from the resulting PCR amplicons were mixed separately with 1μl of 6X gel loading dye and analyzed on 1.5% agarose gel containing ethidium bromide (0.1 μg/ml) at constant electric field of 5V/cm for 30min in 1X TAE buffer. The amplified PCR products 16S rDNA variable region sequnces of bacteria were confirmed as 200bp compact single band DNA visualized separately under UV-light using gel documentation system (Bio-Rad).

2.4 Sequencing and analysis of 16S rRNA gene sequences

The amplicons were purified with ExoSAP (USB) and subjected to automated DNA sequencing on ABI 3730xl Genetic Analyzer (Applied Biosystems, USA). Sequencing was carried out using Big Dye Terminator v3.1 Cycle sequencing kit following the manufacturer’s protocol, where sequencing cycle was set with the thermal ramp rate of 1°C per second for 30 cycles (96°C for 5s; 47°C for 5s and 50°C for 4min). The resulted forward and reverse sequences of 16S rRNA genes of each type of soil sample were aligned with Codon Code aligner software and the consensus 16S rRNA gene sequences were obtained. These consensus gene sequences were used to identify the bacterial isolates with BLASTN analysis using NCBI GenBank Nr database. Based on maximum identity score twelve best 16S rRNA gene sequences were selected and aligned using multiple alignment software program ClustalW. The phylogentic tree was generated by neighbor-joining method using Mega v.4 software tool.

3.1 DNA Extraction

In present study, it has been observed that modified CTAB protocol (TES-CTAB method) is effective to efficiently extract reasonably high molecular weight DNA from different types of soils with good yield which is also dependent upon soil type, pH, organic matter, clay and silt content as these factors can influence either the growth of certain microbial taxa, or the formation of aggregates which host microorganisms [21, 24, 25]. However, DNA yield is not only indicator of DNA extraction efficacy. Indeed, greater amounts of DNA do not necessarily mean that a greater number of texa can be detected. It is likely that extracted DNA mainly comes from easily lysed cells and aggregates [21, 26- 28], and therefore, differences in microbial cell wall structure and micro habitats will affect the extraction of DNA and thus analyses of diversity. Quantification of Extracted DNA was carried out using Nanodrop spectrophotometer (Table 2). The quality and purity of these DNA samples were further confirmed by with agarose gel (0.8%) electrophoresis resulting in the single band of high molecular weight DNA under UV illumination (Figure 1).

3.2 PCR Amplification and Sequencing Analysis

The PCR was performed using Eubacterial primers for Variable 16S rRNA region V5 and V6 and resulting PCR amplicons were visualized as a single intact band of expected size 160-180bp DNA using 1.5% agarose gel electrophoresis (Figure 2). Bacterial diversity was detected in phylogentic tree for 16S rRNA sequence of each type of soil sample and found that most of them are uncultured bacteria. Bacterial Community was generally represented by Protobacteria, Acidobacteria, Fermicutes, Bacteriodetes.

The homologous organisms for bacterial community present in crop land soil (Bamboo as well as Chandan) as shown in the Table 3 & 4 where maximum similarity (80%-91%) was found to the genus Pseudomonas that were uncultured. Majority of these uncultured strains belonged to the phyla Delta-Protobacteria. However, most probable nearest neighborhood strain may be considered as the, Coriobacteriaceae bacterium clone Pad-127(JX505374.1) and Uncultured organism clone SBZP_5567 (JN538754.1) for bacterial community present in Bamboo and Chandan Soil, respectively which is also evident from the phylogenetic tree shown in the Figure 3 & 4. Similarly, bacterial strains present in Rhizosheric soil also showed homology with uncultured strains of diverse type viz  forest soil bacterium, Actinobacterium, Rubrobacteridae bacterium, Solirubrobacter sp. Clone, Gemmatimonadetes bacterium in which nearest neighborhood strain is Marinobacter flavimaris strain SDT4S11(JQ068802.1) that is reported as halophilic, hydrocarbonoclastics bacterium with diazotrophic potential (Figure 5) majority of them are found in hypersaline waters and soils. Experimental evidence suggests their nitrogen-fixation potential [29]. Strains of this species can successfully mineralized crude oil in nutrient media as well as in hypersaline soil or water microcosms without the use of any nitrogen fertilizers.

In case of oil rich and oil contaminated soil, Phylogentic tree (Figure 6 & 11) revealed that uncultured strains are predominant but somewhat different strains in both type of soil like uncultured bacterium clone EMIRGE_OTU_s6b4a_7194 (JX224145.1) and uncultured Sphingobacteriales bacterium clone GE7GXPU01A91FX(HM975819.1) which are nearest neighbor of bacterial strains present in oil-rich soil  and also shows the closeness with each other whereas uncultured bacterium clone SM2F31(EU879395.1) and uncultured bacterium clone nbw775c10c1(GQ009344.1) are the nearest neighbor of bacterial strains in oil-contaminated soil with maximum similarity (80-81%). Besides this Sphingobacteriales, Rikenellaceae, Desulfobulbus and Protobacteria, Actinobacteria, Stenotrophomonas, Lysobacter, Pseudomans also showed homology with bacterial strains present in both type of soil (Table 6 & 11).

Phylogenetic tree of bacterial strains present in  Desert Soil (Sand) shows homology with uncultured bacterial strains as well as many known strains such as Rhodospira trueperi strain ATCC 700224 (AJ001276.1), Brevibacillus agri strain ABRII11(JN604902.1),  Brevibacillus borstelensis  strain: AHK190 (AB491169.1), Brevibacillus thermoruber strain T1SS10 (GQ342691.1) with maximum similarity (77%-78%). But uncultured Syntrophaceae bacterium clone F5oHPNU07H3PTE (HQ050952) and Nitrosococcus oceani strain SDT3S16 (JQ068780.1) are nearest neighbor of bacterial strains present in Desert Soil (Sand). Among these, Nitrosococcus oceani is a member of the evolutionary oldest taxonomic group capable of lithotrophic ammonia catabolism. The gammaproteobacterium Nitrosococcus oceani (ATCC 19707) is a gram-negative obligate chemolithoautotroph capable of extracting energy and reducing power from the oxidation of ammonia to nitrite [30].

Uncultured Desulfobulbus sp. clone GE7GXPU01CGYJL (HM506760.1) and uncultured Desulfobulbus sp. clone GE7GXPU01B52YQ (HM501684.1) were found to be the nearest neighbor of the bacterial community present in marshy soil with maximum similarity of  83%. The genus Desulfobulbus (which is placed under class Deltaprotobacteria) have been studied earlier as sulphate reducing bacteria (SRB) found in the anaerobic sediments at eutrophicated sites polluted with heavy metals, particularly with mercury [31]. It has been also reported that SRB can destroy organic pollutants and can bind heavy metal ions from solutions to non-soluble sulfides. Many toxic metals like cadmium, mercury, tin, zinc, nickel, cobalt, gold, silver and uranium were found in reservoirs are known to have a toxic effects. Conversely, Acidobacteria, Unidentified soil bacteria and uncultured bacterial strains have shown homology to bacterial community present in sewage soil with maximum similarity of 85%-89%. But uncultured Syntrophorhabdus sp. clone F5OHPNU07HX3UX (HQ060377.1) and uncultured bacterium clone EMIRGE_OTU_s8b4e_473 (JX225464.1) were showing the  nearest neighborhood with  bacterial community present in sewage soil as evident from  phylogenetic tree (Figure 9). Peripheral 16S rRNA gene sequences in the databases indicated that the proposed new family Syntrophorhabdaceae is largely represented by abundant bacteria within anaerobic ecosystems mainly decomposing aromatic compounds [32]. The polluted water soil have diverse bacterial community which were showed homology with various types of bacteria such as Geobacter sp., Luteimonas sp., Xanthomonas sp., Desulfuromonas sp., secondary symbiont of Stomaphis quercus etc but the nearest neighbor was found to be Actinobacterium01QJ5 (EU810872.1), uncultured Syntrophobacterales bacterium clone Agri_anode1_191(JN540148.1). It has been reported that Actino bacteria include some of the most common soil life soil life, freshwater life, and marine life, playing an important role in the decomposition of organic materials, such as cellulose and chitin, and thereby playing a vital part in organic matter turnover and the carbon cylcle. In the soil, this replenishes the supply of nutrients and is an important part of humus formation whereas genus Syntrophobacter consists of rod-shaped bacteria growing in syntrophic association with hydrogen- and formate-scavenging microorganisms. Many of the Syntrophobacter spp. is able to use sulfate as the electron acceptor for propionate oxidation and some other organic compounds and hydrogen. The other nearest could be Deinococcus roseus strain TDMA-uv51 (NR_041481.1) and Deinococcus cellulosilyticus strain 5516J-15 (NR_043994.1). These bacteria have thick cell walls that give them gram-positive stains but they include a second membrane and so are closer in structure to those of gram-negative bacteria. They are also characterized by the presence of the carotenoid pigment Deinoxhantin that give them their pink color, and a high resistance to gamma and UV radiation and are usually isolated according to these two criteria. The first one is gamma- and UV-radiation resistant, Gram-positive, red- or pink-pigmented, rod-shaped, strictly aerobic, oxidase- and catalase-positive bacterial strain, was isolated from fresh water collected at Misasa, a radioactive site in Japan [33]. Phylogenetic analysis based on 16S rRNA gene sequences placed it in a distinct lineage in the family Deinococcaceae, along with another similar strain TDMA-25T.The strains degraded gelatin, casein, starch and Tween 80. Unique physiological characteristics, differences in their fatty acid profiles, and genotypic and phylogenetic features, differentiated strains TDMA-25T and TDMA-uv51T from closely related Deinococcus species. Hence, the two strains are described as novel species of the genus Deinococcus. The names Deinococcus misasensis sp. nov. (type strain TDMA-25T=JCM 14369=NBRC 102116=CCUG 53610) and Deinococcus roseus sp. nov. (type strain TDMA-uv51T=JCM 14370=NBRC 102117=CCUG 53611) are proposed [33].

The results presented here demonstrate that the Sanger sequencing method can be used for initial screening of diversity samples before going for high throughput data generation. Standard practice for diversity studies includes DNA isolation from natural samples and subsequently their sequencing by advanced sequencing platforms using 16S rDNA approaches but present study has provided the evidences that Sanger sequencing can also be used for small scale diversity studies.

Authors like to thank management of Xcelris Labs Ltd. for providing infrastructure and facility to conduct the study.

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