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Cave Drip Water-Related Samples As a Natural Environment for Aromatic Hydrocarbon-Degrading Bacteria

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Cave Drip Water-Related Samples As a Natural Environment for Aromatic Hydrocarbon-Degrading Bacteria microorganisms Article Cave Drip Water-Related Samples as a Natural Environment for Aromatic Hydrocarbon-Degrading Bacteria Eric L. S. Marques 1,* , Gislaine S. Silva 1, João C. T. Dias 1, Eduardo Gross 2 , Moara S. Costa 1 and Rachel P. Rezende 1,* 1 Department of Biological Sciences, State University of Santa Cruz, Rod. Jorge Amado, Km 16, Ilheus CEP 45662-900, Brazil; [email protected] (G.S.S.); [email protected] (J.C.T.D.); [email protected] (M.S.C.) 2 Department of Agricultural and Environmental Science, State University of Santa Cruz, Rod. Jorge Amado, Km 16, Ilheus CEP 45662-900, Brazil; [email protected] * Correspondence: [email protected] (E.L.S.M.); [email protected] (R.P.R.); Tel.: +55-73-3680-5435 (E.L.S.M.); +55-73-3680-5441 (R.P.R.) Received: 30 November 2018; Accepted: 15 January 2019; Published: 25 January 2019 Abstract: Restricted contact with the external environment has allowed the development of microbial communities adapted to the oligotrophy of caves. However, nutrients can be transported to caves by drip water and affect the microbial communities inside the cave. To evaluate the influence of aromatic compounds carried by drip water on the microbial community, two limestone caves were selected in Brazil. Drip-water-saturated and unsaturated sediment, and dripping water itself, were collected from each cave and bacterial 16S rDNA amplicon sequencing and denaturing gradient gel electrophoresis (DGGE) of naphthalene dioxygenase (ndo) genes were performed. Energy-dispersive X-ray spectroscopy (EDX) and atomic absorption spectroscopy (AAS) were performed to evaluate inorganic nutrients, and GC was performed to estimate aromatic compounds in the samples. The high frequency of Sphingomonadaceae in drip water samples indicates the presence of aromatic hydrocarbon-degrading bacteria. This finding was consistent with the detection of naphthalene and acenaphthene and the presence of ndo genes in drip-water-related samples. The aromatic compounds, aromatic hydrocarbon-degrading bacteria and 16S rDNA sequencing indicate that aromatic compounds may be one of the sources of energy and carbon to the system and the drip-water-associated bacterial community contains several potentially aromatic hydrocarbon-degrading bacteria. To the best of our knowledge, this is the first work to present compelling evidence for the presence of aromatic hydrocarbon-degrading bacteria in cave drip water. Keywords: Sphingomonadales; microorganisms; ndo gene; naphthalene; NGS; Illumina; 16S rDNA 1. Introduction Caves are aphotic environments isolated to some degree from the surface and lacking photosynthetic activity [1–3]. Consequently, a cave environment commonly has low levels of organic matter and some essential micronutrients change according to the mineralogical composition associated with carbonate rocks [4]. However, other factors such as runoff, bats, tourism, invertebrates’ feces, and drip water could exert a significant influence on the organic nutrients available in caves [5] and, thus, on the cave microbial communities. Microbial communities in caves have been studied, with a focus on microbial or biofilm associations in the cave environment [4,6,7] and external contamination caused by tourism [8,9]. Despite all the studies in caves, there is a lack of information regarding the influence of dripping Microorganisms 2019, 7, 33; doi:10.3390/microorganisms7020033 www.mdpi.com/journal/microorganisms Microorganisms 2019, 7, 33 2 of 13 water and its effects on cave floor sediment microbial communities. Microorganisms in dripping water are involved in the formation of speleothems, as well as in biofilm on the walls [10–13]. Many bacteria found in caves can induce calcite precipitation [11,14]. However, dripping water could transport organic and inorganic material from the surface [15–17]. This material carried to caves could change the availability of nutrients and modify microbial communities in favor of a chemoheterotrophic-based community. We hypothesize that drip water microbial communities can degrade transported organic matter such as aromatic compounds. Among those compounds, naphthalene is one of the most well studied, and it has been a model compound for aromatic metabolism [18] and is usually found in caves [16], while several other aromatic compounds are found in low concentration in soils due to the partial degradation of lignin [19]. Among the genes that encode enzymes to degrade naphthalene, the nahAc and phnAc cluster III are those with greatest environmental importance due to their presence in common soil bacteria [20] and codify the large subunit of different naphthalene dioxygenase, and those genes were used in the present study. The presence of aromatic compounds and aromatic hydrocarbon-degrading bacteria in caves is partially unknown with several studies having aimed to detect such compounds in karstic aquifer [21,22], but no relation with drip water was analyzed. Besides alluvial transportation, the presence of aromatic compounds in cave air has also been demonstrated as an alternative transport mechanism for such compounds [23]. This paper aims to analyze the occurrence of potential bacterial degradation of aromatic compounds in the bacterial communities of dripping water, and drip-water-saturated and unsaturated sediment samples from two different carbonate caves. 2. Materials and Methods 2.1. Sampling and DNA Extraction Two caves were selected for this study: Gruta do Bom Pastor (cave identified as GBP in the text and G associated with a sample, 10◦39005.99” S, 37◦55026.87” W) and Furna do Fim do Morro do Parafuso (cave identified as FFMP or F, 10◦38025.89” S, 37◦52004.13” W). Sampling was authorized by the Ministério do Meio Ambiente (MMA/ICMBio/SISBIO – No. 26304). These caves are located 8 kilometers from each other in Paripiranga, Bahia State, Brazil (Figures S1 and S2), a hot semi-arid location. The municipal area is based on Neoproterozoic rocks, with meta-limestone (metamorphosed carbonate rock), meta-dolomite and metapelite prevailing in the caves. Both caves are in limestone/meta-limestone hydrogeological domains that represent an aquifer system dominated by limestone, magnesian limestone, and dolomite rocks that can undergo the karstic dissolution that led to caves, sinkholes, dolines, and other erosive features related to this type of rock. The linear development of the studied caves put GBP and FFMP at 236 m and 234 m respectively. However, GBP has a considerable vertical projection for a small cave, at approximately 75 m. The first of the 3 chambers in GBP was also used as a pilgrimage destination; however, the sample sites appear preserved, especially the last chamber, which was not used in this way. No bats nor guano were visible in the sampling sites, but there are bats and guano in other chambers of both caves. Runoff occurs in GBP, and a variety of invertebrates were observed in FFMP. GBP and FFMP are located respectively 12 and 5 km from the closest city (Paripiranga), but small villages and family houses are located nearby. Above GBP is a corn plantation, while FFMP is located beneath native vegetation. Composite sediment samples were collected from 5 subsamples in a 3 m2 area to obtain approximately 200 g. Dripping water was collected in sterile containers from several stalactites to reach at least 50 mL (identified as F1 and G1). The sediment samples collected were drip-water-saturated sediment (F2 and G2) and drip-water-unsaturated sediment (F3 and G3). All sediment samples were placed in sterile plastic bags and transported immediately on ice to the laboratory, where part of them was analyzed and processed for their physicochemical characteristics. For each sediment sample, Microorganisms 2019, 7, 33 3 of 13 the temperature (Incoterm, São Paulo, Brazil) was measured in situ, after sampling, with a thermometer in contact with the sediment. DNA from non-sieved sediment samples was extracted using the MoBio PowerSoilTM DNA kit (MoBio Laboratories Inc., Carlsbad, CA, USA) according to the manufacturer’s instructions. The kit was also used for DNA extraction from drip water cells collected on a 0.22 µm membrane. 2.2. Gas Chromatography Analysis Aromatic compounds were extracted in triplicate from 10 g of soil or 10 mL of distilled water. To extract those compounds, 10 mL of hexane:acetone (1:1, v/v; FMaia LTDA, Cotia, Brazil) were added and sonicated (Ultrasonic cleaner, Maxiclean 1600, Indaiatuba, Brazil) for 30 min. After that, an additional 10 mL hexane:acetone was added, vortexed for 1 min, and sonicated again for 30 min. The extracted solution was stored in a glass vial containing Na2SO4 before analysis. Chromatographic analyses were performed on a Shimadzu gas chromatograph using an RTX-5 column (Restek, Bellefonte, PA, USA; 30 m, 0.25 mm; 0.30-µm film) with hydrogen as the carrier gas. One microliter of each sample was injected at 280 ◦C for 1 min. The temperature column was followed: 40 ◦C for 5 min followed by a heating speed of 10 ◦C/min up to 250 ◦C, hold for 20 min. A polynuclear aromatic hydrocarbons mix (Supelco, Sigma-Aldrich, St. Louis, MO, USA) was used as a standard for identification and quantification of corresponding peaks. The quantification was made in a serial dilution of standard. 2.3. 16S rDNA Gene Sequencing The amplification of the 16S rRNA gene V3/V4 region was carried out using the 341F (50-CCT ACG GGR SGC AGC AG-30) and 806R (50-GGA CTA CHV GGG TWT CTA AT-30) primers [24,25]. The 16S rRNA gene libraries were constructed following a PCR-based protocol [26] and sequenced using the MiSeq Sequencing System (Illumina Inc., San Diego, CA, USA) with the V2 kit, 300 cycles. Reads were separated by barcoding and trimmed using Geneious 11.0.4. Chimeras were detected using DECIPHER [27]. Operational taxonomic units (OTUs) assigned by clustering the sequence against the Ribossomal Database Project (RDP Tools version 2.12) [28] and EZBioCloud database [29]. A cluster analysis based on the abundance of each OTU was made on PAST 3.1 software (Natural History Museum - University of Oslo, Oslo, Norway, https://folk.uio.no/ohammer/past/).
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