- Email: [email protected]
Comparison of methods for preservation of activated sludge samples for high-throughput nucleic acid sequencing and bacterial diversity analysis
International Biodeterioration & Biodegradation 157 (2021) 105139
Contents lists available at ScienceDirect
International Biodeterioration & Biodegradation journal homepage: www.elsevier.com/locate/ibiod
Comparison of methods for preservation of activated sludge samples for high-throughput nucleic acid sequencing and bacterial diversity analysis Lívia Carneiro Fid´elis Silva a, 1, D´eborah Romaskevis Gomes Lopes a, 1, Helena Santiago Lima a, Larissa Quartaroli a, Maíra Paula de Sousa b, Vinicius de Abreu Waldow b, Rubens Nobumoto Akamine b, S´ergio Oliveira de Paula a, Cynthia Canˆedo da Silva b, * a b
Department of Microbiology, Universidade Federal de Viçosa, Viçosa, Minas Gerais, Brazil Department of Biotechnology, Leopoldo Am´erico Miguez de Mello Research and Development Center (Cenpes), Petrobras, Rio de Janeiro, RJ, Brazil
A R T I C L E I N F O
A B S T R A C T
Keywords: Microbial community structure Sample preservation methods Environmental sample Amplicon sequencing
Preservation of environmental samples is an important step in maintaining the original microbial community until the nucleic acid extraction in the laboratory. Here, we collected activated sludge samples to both imme diately extract nucleic acids (control) and submit to different storage/preservation methods for 48 h before nucleic acids extraction: room temperature storage, storage on ice, direct storage at − 20 ◦ C, rapid freezing in liquid nitrogen followed by storage at − 20 ◦ C, and preservation with TRIzol®, RNAlater®, and Nucleic Acid Preservation (NAP) buffers. Bacterial community was compared by high-throughput 16S rRNA gene sequencing from total DNA and RNA. Among the evaluated methods, TRIzol® and rapid freezing with liquid nitrogen were the least indicated, since they led to significant changes in the microbial community structure and abundance of functional groups involved in ammonia metabolism. Compared to control, storage on ice and NAP preserved more than 80% and 75% of genera at the DNA and RNA level respectively, indicating that these methods are the most suitable for storage/preservation of activated sludge samples intended for nucleic acids sequencing.
1. Introduction Advances in high-throughput sequencing of nucleic acids have resulted in better coverage and characterization of the microbial di versity from environmental samples (Althani et al., 2016; O’Doherty et al., 2016; Sales et al., 2019). However, a successful employment of these methods requires efficient upstream protocols for sample collec tion, storage and preservation, aiming at conserving the fidelity, integ rity and quantity of the genetic material present in the collected samples (Osborn, 2004; Pontes et al., 2007). Protocols for sample collection and nucleic acids extraction must be appropriate to the individual characteristics of each sample, such as its origin, physical state and extracellular matrix, among others. Thus, specific protocols for DNA and RNA preservation and extraction are constantly being revised and optimized to obtain nucleic acids with high yield and quality (Gabor et al., 2003; Henderson et al., 2013; Dias et al.,
2014; Taberlet et al., 2018). However, information about the effect of preservatives and storage conditions aimed at preserving the integrity of genetic material from environmental samples is still limited (Mitchell and Takacs-Vesbach, 2008; Sales et al., 2019). Ideally, the genetic ma terial is extracted immediately after collection, but the establishment of preservation protocols ensures that samples collected in remote areas can be transported to a laboratory with adequate infrastructure for nucleic acid extraction without greatly compromising the integrity of the metagenome and metatranscriptome as originally present in the sample. Inappropriate methods of sample preservation and storage may result in DNA and RNA degradation, compromising inferences about the microbial community structure and making it difficult to identify rare species (Pilliod et al., 2014; Barnes and Turner, 2016; Hinlo et al., 2017). Sample storage at low temperatures, including freezing, is widely applied for sample preservation, as overall microbial metabolism and enzyme activity are reduced in this condition (Pilliod et al., 2014;
* Corresponding author. Av. Peter Henry Rolfs, s/n, Campus Universit´ ario, Laborat´ orio de Microbiologia Ambiental Aplicada, Viçosa, MG, CEP 36570-900, Brazil. E-mail addresses: [email protected] (L.C.F. Silva), [email protected] (D.R.G. Lopes), [email protected] (H.S. Lima), larissaquartaroli@ gmail.com (L. Quartaroli), [email protected] (M.P. de Sousa), [email protected] (V.A. Waldow), [email protected] (R.N. Akamine), [email protected] (S.O. de Paula), [email protected] (C.C. Silva). 1 These authors contributed equally to this work. https://doi.org/10.1016/j.ibiod.2020.105139 Received 12 August 2020; Received in revised form 21 October 2020; Accepted 12 November 2020 Available online 27 November 2020 0964-8305/© 2020 Elsevier Ltd. All rights reserved.
L.C.F. Silva et al.
International Biodeterioration & Biodegradation 157 (2021) 105139
Eichmiller et al., 2016). Other strategies include mixing the sample with saline buffers containing chelating agents (Camacho-Sanchez et al., 2013; Williams et al., 2016; Ladell et al., 2019) or with commercial organic solutions that generally act by protein denaturation (Menke et al., 2017; Yamanaka et al., 2017). The oil and gas industry is a major consumer of water, with a high ratio of volume of water used per volume of oil processed. Oil extraction and refining processes generate large quantities of contaminated efflu ents, whose composition is complex, containing organic compounds as well as ions with varying concentrations, such as chloride (Cl− ), sodium (Na+), calcium (Ca2+), magnesium (Mg2+), sulfide (S2− ) and ammonium (NH+ 4 ). Wastewater discharges containing high concentrations of nitro gen can be toxic to aquatic life, causing eutrophication and oxygen depletion in the aquatic environment, negatively affecting water qual ity. Thus, according to environmental requirements, industrial effluents must be treated before being reused or discharged into a receiving water body (Fortis et al., 2007; Munirasu et al., 2016). Among biological treatments of effluents, activated sludge system is the most commonly used due to its high efficiency (Da Motta et al., 2003). The biomass of activated sludge systems is composed of a complex extracellular matrix and a high microbial diversity that develops global processes related not only to the nitrogen cycle, but also to carbon and phosphorus cycles (Shchegolkova et al., 2016; Quartaroli et al., 2017; Wang et al., 2018). Characterization of the microbial community in activated sludge is important for monitoring populations and metabolic activity inside re actors of effluent treatment plants. However, these treatment plants are commonly located close to the runoff and/or oil refining stations and distant to laboratory facilities, making it necessary to transport these samples to a laboratory for more complex microbiological analysis. Given the importance of microbial characterization of activated sludges and the absence of a specific preservation/storage protocol for this kind of sample, we collected samples of activated sludge from a pilot reactor, extracted nucleic acids immediately after collection (control) and after being submitted for 48 h to seven different storage/preservation methods: storage at room temperature, storage on ice, direct freezing at − 20 ◦ C, rapid freezing in liquid nitrogen followed by storage at − 20 ◦ C, and mixing with a homemade (Nucleic Acid Preservation Solution – NAP) and two commercial (TRIzol® and RNAlater®) preservation buffer. The comparison of the bacterial community structure in terms of genera composition and functional groups with the control treatment indicated the best storage/preservation method to nucleic acid analysis in activated sludge samples.
2.2. Nucleic acid extraction Total DNA and total RNA were extracted from the replicates of each treatment according to the protocol described by Silva et al. (2010), with modifications. The total volume of each treated sample was centrifuged at 12,000 × g for 5 min, and the pellet was washed three times with SET buffer (20 mM Tris, 75 mM NaCl, 25 mM EDTA, pH 7.5). After washing, the pellet was resuspended in 1.0 mL of SET buffer plus 50 μL of a 100 mg/mL lysozyme solution (GE Healthcare, Chicago, USA), and the so lution was incubated inside a 37 ◦ C water bath and stirred every 10 min. Subsequently, 50 μL of a 10 mg/mL proteinase K solution (Sigma-Al drich, St Louis, USA) and 200 μL of SDS 10% w/v (for a final concen tration of 2% w/v) were added to the solution, and it was then incubated at 60 ◦ C for 30 min and stirred every 10 min. The samples were then subjected to three freeze-thaw cycles of 2 min immersed in liquid ni trogen and 2 min at 65 ◦ C. One milliliter of TRIzol® (Qiagen, Hilden, Germany) was added to the solution and homogenized for 2 min. After centrifugation at 10,000 × g for 5 min, the organic phase was used for DNA extraction and the aqueous phase for RNA extraction. In both phases, an equal volume of chloroform/isoamyl alcohol 24:1 (v/v) so lution was added and centrifuged at 10,000 × g for 5 min. The super natant was collected and a 5 M NaCl solution (10% of the total volume) was added to it, followed by the addition of 2 vol of frozen absolute ethanol and centrifugation for 20 min at 10,000 × g. The pellet was washed with 70% ethanol (v/v) and, after drying, it was eluted in 50 μL of ultrapure water. DNA and RNA samples were respectively treated with RNAse (Sigma-Aldrich, St Louis, USA) and DNAse (Promega Cor poration, Madison, USA), following the manufacturer’s recommenda tions. Nucleic acids were quantified using a NanoDrop™ 2000 Spectrophotometer (Thermo Fisher Scientific, Waltham, USA) and visualized in 1% w/v agarose gel. RNA was used for cDNA synthesis with the GoScript™ Reverse Transcriptase kit (Promega Corporation, Wis consin, USA), according to the manufacturer’s recommendations. DNA and cDNA from replicates of each treatment were lyophilized (Benchtop K, VirTis, New York, EUA) and sent for sequencing at the Molecular Research DNA Laboratory (http://www.mrdnalab.com, Shallowater, USA) using the Illumina MiSeq platform. 2.3. Sequencing and analysis of data The sequences obtained were processed using the “MR DNA analysis pipeline” (MR DNA, Shallowater, USA). The barcodes were removed, chimeras and assembled contigs and sequences smaller than 150 bp were also removed. The operational taxonomic units (OTUs) were defined by grouping under 3% divergence (97% similarity). A repre sentative sequence of each OTU was taxonomically classified using BLASTn against a database with curatorship derived from GreenGenes (http://greengenes.lbl.gov), RDPII (http://rdp.cme.msu.edu) and NCBI (http://www.ncbi.nlm.nih.gov). Multivariate analyses and calculation of diversity indices were performed using the PAST software (“Paleon tological Statistics”) (Hammer et al., 2001). All original nucleotide se quences are available at the NCBI Sequence Read Archive (SRA) under Accession Number PRJNA597347.
2. Materials and methods 2.1. Sampling and preservation A sample of activated sludge was collected from a pilot reactor ´rio de Microbiologia Ambiental Aplicada of the operating at the Laborato Universidade Federal de Viçosa, situated in the state of Minas Gerais, Brazil. This pilot reactor was inoculated with a sample of the activated sludge from a petroleum terminal effluent treatment plant and fed with effluent with a saline concentration of 5.5 g.L− 1. Sludge aliquots of 2.0 mL were collected in triplicate for DNA and RNA extraction under different conditions: (1) extraction immediately after collection (con trol); and extraction after 48 h under (2) storage at room temperature; (3) storage on ice using a cooling box; (4) direct freezing at − 20 ◦ C; (5) rapid freezing in liquid nitrogen followed by storage at − 20 ◦ C; (6) preservation in TRIzol® (Invitrogen, Carlsbad, USA) by mixing in a 3:1 vol ratio (preservative-to-sample); (7) preservation in RNAlater® solu tion (Qiagen, Hilden, Germany) by mixing in a 5:1 vol ratio; and (8) preservation in Nucleic Acid Preservation Solution (NAP) by mixing in a 3:1 vol ratio (Camacho-Sanchez et al., 2013). TRIzol® and RNAlater® were added in the ratios recommended by their respective manufac turers. All treatments were performed in triplicate.
2.4. Statistical analysis Comparisons of the mean concentrations of extracted DNA and RNA, diversity indices and functional categories were performed using Mini Tab® 17.1.0 software (Minitab, Inc., State College, USA) by ANOVA, followed by Tukey’s test. P-values less than 0.05 were considered significant. 3. Results Both the DNA and the RNA extracted from the samples showed high purity, with an average absorbance ratio of 260/280 nm equal to 1.83 2
L.C.F. Silva et al.
International Biodeterioration & Biodegradation 157 (2021) 105139
and 2.05, respectively. The different preservation/storage methods evaluated did not affect the amount of total DNA recovered when compared to the control (extraction immediately after extraction) (p > 0.05). In relation to RNA, all treatments evaluated resulted in increased concentration recovered compared to control (mean 1832.87 ng/μL), with the exception of RNAlater® and liquid nitrogen (mean 1048.11 ng/ μL) (p < 0.05) (Fig. 1). Alpha diversity analyses performed from the partial sequencing of the 16S rRNA gene demonstrated that the conditions tested had OTU richness and Shannon and Simpson (1-D) index values similar to the control (p > 0.05), with the exception of freezing with liquid nitrogen, in which these values were 349, 2.59 and 0.82, respectively (p < 0.05) (Table 1). When microbial composition was accessed by cDNA sequencing obtained from transcripts of the 16S rRNA gene, only direct storage at − 20 ◦ C and rapid freezing with liquid nitrogen followed by storage at − 20 ◦ C showed greater richness of OTUs than the control (p < 0.05). For the Shannon index, preservation in TRIzol® was the only method that presented a higher value than the control (p < 0.05) (Table 1). Beta diversity analyses using the Bray-Curtis similarity index showed that the different preservation/storage methods altered the bacterial composition of sludge samples. For both DNA and cDNA, preservation in TRIzol® and direct freezing in liquid nitrogen followed by storage at − 20 ◦ C resulted in the greater dissimilarity of the bacterial community relative to the control. For DNA, preservation in RNAlater® was the condition with the most homogeneous replicates and closest to the control; while for cDNA, the condition with the most homogeneous replicates and closest to the control was direct freezing at − 20 ◦ C (Fig. 2). Obtained sequences resulted in 1008 unique OTUs which were classified into 21 phyla, 38 classes, 79 orders, 164 families and 366 genera. The four most abundant genera in all treatments were Hal omonas, Tindallia, Spirochaeta and Halanaerobium, representing 83.64 ± 2.43% (mean ± SD) of the obtained DNA sequences, and 93.72 ± 1.83% (mean ± SD) of the obtained cDNA sequences. TRIzol® was the
preservative that most disturbed the taxonomic profile of sludge samples in relation to the control: the genera Spirochaeta, Rickettsia and Achole plasma were enriched for DNA, while Spirochaeta was enriched and Halanaerobium was diminished for cDNA, presenting less abundance in relation to the control (Fig. 3). Considering the genera involved in ni trogen metabolism, both for DNA and cDNA, the abundance of Nitra tireductor was different among the treatments (p < 0.05). When compared to the control, no treatment presented differences in relation to Nitratireductor abundance when evaluating DNA, while preservation in NAP, TRIzol® and liquid nitrogen freezing provided enrichment of Nitratireductor when evaluating cDNA (p < 0.05). When cDNA was evaluated, an increase in the abundance of Nitrospira was observed with preservation in TRIzol® and liquid nitrogen freezing (p < 0.05). For DNA, there was no difference in this genus between treatments (p > 0.05). The abundance of the genus Nitrosomonas was not altered (Fig. 4). Of 96 genera identified with DNA sequencing, approximately 56% were shared among all treatments, and of 124 genera identified with cDNA sequencing, 32% were shared among all treatments. Storage on ice was the storage/preservation method with the highest number of genera shared with the control (84% for DNA and 83% for cDNA), and directly freezing at − 20 ◦ C was the condition with the lowest number of genera shared with the control (77% for DNA and 47% for cDNA) (Fig. 5). 4. Discussion Considering logistical and practical difficulties of extracting nucleic acids immediately after sample collection, it is necessary to investigate safe and efficient ways of preserving samples without compromising the microbial community structure and nucleic acids integrity. Molecular studies demonstrated that physical methods of storage and preservation, especially those involving low temperatures, are more efficient in maintaining the microbial community structure than chemical methods (Rubin et al., 2013; Cui et al., 2014; Tatangelo et al., 2014). Chemical preservatives interact directly with cellular and matrix components and
Fig. 1. Concentration of DNA and RNA extracted from activated sludge samples subjected to different storage/preservation methods. Control represents the im mediate extraction after sample collection, while storage/preservation methods are represented as: RT (room temperature); NAP (Nucleic Acid Preservation solu tion); Ice (storage on ice); TZ (TRIzol®); RL (RNAlater®); NIT (rapid freezing with liquid nitrogen followed by storage at − 20 ◦ C) and Freeze (direct freezing at − 20 ◦ C). The letters above the bars represent the statistical differences between the average DNA or RNA concentrations. Equal letters represent that there was no statistical difference at 95% of confidence obtained by ANOVA followed by Tukey’s test. 3
L.C.F. Silva et al.
International Biodeterioration & Biodegradation 157 (2021) 105139
Table 1 Diversity indices of activated sludge samples submitted to different storage/preservation methods. DNA
cDNA
OTU Richness Simpson (1-D) Shannon Evenness Chao OTU Richness Simpson (1-D) Shannon Evenness Chao
Control
RT
NAP
Ice
TZ
RL
NIT
Freeze
426.33a 0.85a 2.83a 0.04 702.70 313.67b 0.70abc 1.93bc 0.02abc 511.20c
385.67ab 0.83ab 2.73ab 0.04 589.37 290.67b 0.74ab 2.15ab 0.03a 428.00c
413.67a 0.85a 2.82a 0.04 665.13 335.00b 0.72abc 2.07abc 0.02 ab 485.70c
413.00a 0.83ab 2.73ab 0.04 660.30 329.33b 0.74ab 2.17ab 0.03ab 499.17c
382.00ab 0.85a 2.69ab 0.04 564.73 372.67b 0.75a 2.28a 0.03ab 565.60bc
387.00ab 0.83ab 2.66ab 0.04 623.60 336.33b 0.68bc 1.90bc 0.02bc 616.47bc
349.67b 0.82b 2.59b 0.04 554.90 585.33a 0.66c 1.84c 0.01d 1035.73a
415.33a 0.83ab 2.68ab 0.04 654.47 510.33a 0.68c 1.98abc 0.01cd 855.63ab
Control, extraction of genetic material immediately after collection; RT, storage at room temperature; NAP, Nucleic Acid Preservation solution; Ice, storage on ice; TZ, TRIzol®; RL, RNAlater®; NIT, freezing in liquid nitrogen followed by storage at − 20 ◦ C; and Freeze, direct freezing at − 20 ◦ C. Averages of the indices followed by the same letter have no statistical difference at 95% confidence obtained by ANOVA followed by Tukey’s test. Fig. 2. Principal coordinate analysis (PCoA) based on the Bray-Curtis similarity index of activated sludge samples submitted to different storage/preservation methods. Bacterial diversity was accessed by partial sequencing of the 16S rRNA gene from total DNA (a), samples represented by circles, and cDNA synthesized from total RNA (b), samples represented by triangles. Control represents the extraction of genetic material immediately after collection, while storage/preser vation methods are represented as: RT, storage at room temperature; NAP, Nucleic Acid Preservation solution; Ice, storage on ice; TZ, TRIzol®; RL, RNA later®; NIT, rapid freezing with liquid nitrogen fol lowed by storage at − 20 ◦ C; and Freeze, direct freezing at − 20 ◦ C.
alter the pH of the solution, resulting in discrepancies in bacterial composition between the samples (Rissanen et al., 2010; Tripathi et al., 2012; Tatangelo et al., 2014). Preservation solutions rich in salts (which favor the precipitation of nucleic acids) and chelators of enzymatic co factors (e.g., EDTA) can be purchased commercially, such as RNAlater®, TRIzol®, DNAzol® and LifeGuard™, or produced in the laboratory (“homemade”), such as NAP solution and Longmire’s buffer (Cama cho-Sanchez et al., 2013; Longmire et al., 1997). Additionally, the use of chemical preservatives usually requires large volumes of the chosen preservative, since the preservative-to-sample ratio normally used is 3:1. In this work, relevant information was provided on the effect of different preservation/storage conditions of activated sludge samples on the recovered nucleic acid concentration and on the taxonomic and meta bolic function diversity of the bacterial community. When compared to the control (extraction immediately after
collection), the evaluated preservation/storage methods did not affect the amount of total DNA recovered. In contrast, two methods resulted in a reduced RNA concentration when compared to the control (RNAlater® and rapid freezing with liquid nitrogen followed by storage at − 20 ◦ C), and the other five methods resulted in increased RNA concentration in comparison to the control. These results may be associated with (1) the fact that rates of biosynthesis and biodegradation of RNA transcripts are higher when compared to genomic DNA due to regulation of gene expression in response to environmental conditions, and (2) RNA is more susceptible to hydrolysis than DNA, making it more sensitive to preservation/storage methods (Xia et al., 1998; Deutscher, 2006). For both DNA and RNA, the genus Halomonas was the most abundant in all treatments, as the samples of sludge were all inoculated in a bioreactor fed with saline effluent (5.5 g.L− 1 of salt). This genus com prises halophilic bacteria isolated from saline environments around the 4
L.C.F. Silva et al.
International Biodeterioration & Biodegradation 157 (2021) 105139
Fig. 3. Taxonomic composition at the genus level of activated sludge samples submitted to different stor age/preservation methods. Bacterial diversity was accessed by partial sequencing of the 16S rRNA gene from total DNA (a) and cDNA synthesized from total RNA (b). Control represents the extraction of genetic material immediately after collection, while storage/ preservation methods are represented as: RT, storage at room temperature; NAP, Nucleic Acid Preservation solution; Ice, storage on ice; TZ, TRIzol®; RL, RNA later®; NIT, rapid freezing with liquid nitrogen fol lowed by storage at − 20 ◦ C; and Freeze, direct freezing at − 20 ◦ C. Others represents genera with relative abundance less than 0.2% in the control.
Fig. 4. Heatmap representing the relative abun dances of genera involved in nitrogen metabolism in sludge activated samples. Genera accessed by partial sequencing of the 16S rRNA gene from total DNA (a) and cDNA synthesized from total RNA (b). Control represents the extraction of genetic material imme diately after collection, while storage/preservation methods are represented as: RT, storage at room temperature; NAP, Nucleic Acid Preservation solu tion; Ice, storage on ice; TZ, TRIzol®; RL, RNAlater®; NIT, rapid freezing with liquid nitrogen followed by storage at − 20 ◦ C; and Freeze, direct freezing at − 20 ◦ C. Identical letters have no statistical difference at 95% confidence obtained by ANOVA followed by Tukey’s test.
world, including solar salt production facilities, intertidal estuaries, the open ocean and hypersaline lakes (Vreeland, 2015). Our results indicate that preservation in TRIzol® and rapid freezing in liquid nitrogen fol lowed by storage at − 20 ◦ C were the worst methods, since the microbial community diverged most from the control. TRIzol® is used for main taining nucleic acids integrity, as it contains high concentrations of guanidine thiocyanate and acid phenol, which are inhibitors of endo nucleases (Simister et al., 2011). Rapid freezing in liquid nitrogen is a widely-used method of cryogenics for cell cultures in general, since it leads to the formation of smaller water crystals inside the cell, making it possible to maintain its properties after thawing (Weißbecker et al., 2017). Nonetheless, both TRIzol® and liquid nitrogen are expensive and difficult to handle in the field, since TRIzol® has high toxicity and liquid
nitrogen presents the risk of explosion during its transport. Storage on ice and preservation in NAP solution were the most efficient methods for maintaining the microbial community structure in samples: these methods covered more than 80% of the genera at the DNA level and more than 75% at the RNA level in comparison to the control. According to Weißbecker et al. (2017), an overlap of 70–80% of OTUs with the control treatment represents a satisfactory value for a storage/preservation method to be considered good. Sample storage at low temperatures reduces overall enzymatic activity, reducing the degradation of nucleic acids. In addition, mild cooling of samples, unlike freezing, ensures that water crystals are not formed, and thus do not cause cell lysis during thawing (Weißbecker et al., 2017). The main objective of biological treatment of liquid effluents by 5
L.C.F. Silva et al.
International Biodeterioration & Biodegradation 157 (2021) 105139
Fig. 5. Venn diagrams representing the unique genera for each storage/preservation method and those shared with the control of activated sludge samples. Bacterial diversity was accessed by partial sequencing of the 16S rRNA gene from total DNA (a) and cDNA synthesized from total RNA (b). Control represents the extraction of genetic material immediately after collection, while storage/preservation methods are represented as: RT, storage at room temperature; NAP, Nucleic Acid Pres ervation solution; Ice, storage on ice; TZ, TRIzol®; RL, RNAlater®; NIT, rapid freezing with liquid nitrogen followed by storage at − 20 ◦ C; and Freeze, direct freezing at − 20 ◦ C. Only the genera observed in all replicates were considered. Below each diagram is the percentage of treatment sharing in relation to the control.
activated sludge is the removal of pollutants, including ammonia (Da Motta et al., 2003). The biological removal of ammonia can be per formed by different processes involving distinct groups of microorgan isms, through either autotrophic or heterotrophic metabolism, and under both aerobic and anaerobic conditions (Robertson and Kuenen, 1983; Vazoller et al., 2001; Limpiyakorn et al., 2013). The cDNA sequencing data demonstrated that the relative abundances of genera involved in autotrophic nitrification, Nitrosomonas and Nitrospira (Wagner et al., 2002), and anaerobic denitrification, Nitratireductor (Labb´e et al., 2004), were altered by the different preservation/storage methods evaluated, indicating that the chosen method may compromise the characterization of functional groups in sludge samples at the RNA level. Thus, studies aiming at the characterization of metabolically active microorganisms and/or gene expression studies should be more cautious regarding the choice of sample preservation/storage method. Despite being a widely used preservative solution, studies have shown that RNAlater® is not efficient for preserving the microbial sample community. Lackey et al. (2017) evaluated the effect of preser vation with commercial solutions and the storage time of human milk samples maintained at 37 ◦ C on the composition of the bacterial com munity (sequencing of the V1-V3 region of the 16S rRNA gene) and observed that over time the preservation with RNAlater® is compro mised at that temperature. According to the manufacturer, this solution is efficient for preservation for up to 1 day at 37 ◦ C and up to 1 week at 23 ◦ C (room temperature). For samples of monkey feces, Hale et al. (2015) observed that the microbial composition and abundance of samples preserved in RNAlater® differ from the fresh sample. Menke
et al. (2017) observed that the NAP solution stabilizes the microbial community better than commercial preservative media (RNAlater® and RNA/DNA Shield™) for samples of sheep feces, making this homemade solution a viable alternative, in terms of both cost and efficiency, for the study of environmental samples from remote areas in the absence of ice or refrigerators. High-throughput sequencing, as well as other molecular applica tions, requires that the extracted genetic material is of high quantity and good quality in order to adequately access the microbial groups and metabolic functions present in the sample. In this work, we observed that the evaluated preservation/storage methods differentially altered the microbial community structure, but all of them enabled the recovery of DNA and RNA in sufficient concentration and quality. In addition, it was suggested that for activated sludge samples, storage in ice or pres ervation in NAP solution are efficient for preserving the DNA and RNA of the microbial community in the sample. However, the choice of a preservation method will depend on several factors, such as the nature and physicochemical characteristics of the sample, logistics, cost, availability, safety and infrastructure available at the time of collection, as well as the focus of the study (taxonomic profile of the microbial community or gene expression, for example). 5. Conclusion This study provided evidence that freezing of activated sludge sam ples in liquid nitrogen followed by storage at − 20 ◦ C and preservation in TRIzol® results in significant changes in the total structure of the 6
L.C.F. Silva et al.
International Biodeterioration & Biodegradation 157 (2021) 105139
microbial community and the relative abundance of functional groups involved in ammonia metabolism, for both DNA and RNA. Moreover, our results indicated that preservation in NAP solution or storage on ice enabled greater coverage of the total and metabolically active microbial community, accessed by both DNA and cDNA (RNA) sequencing, and therefore the application of these methods is recommended for molec ular and functional studies of activated sludge samples, both from DNA and RNA.
Hinlo, R., Gleeson, D., Lintermans, M., Furlan, E., 2017. Methods to maximise recovery of environmental DNA from water samples. PLoS One 12, e0179251. https://doi. org/10.1371/journal.pone.0179251. Labb´e, N., Parent, S., Villemur, R., 2004. Nitratireductor aquibiodomus gen. nov., sp. nov., a novel α-proteobacterium from the marine denitrification system of the Montreal Biodome (Canada). Int. J. Syst. Evol. Microbiol. 54, 269–273. https://doi.org/ 10.1099/ijs.0.02793-0. Lackey, K.A., Williams, J.E., Price, W.J., Carrothers, J.M., Brooker, S.L., Shafii, B., McGuire, M.A., McGuire, M.K., 2017. Comparison of commercially-available preservatives for maintaining the integrity of bacterial DNA in human milk. J. Microbiol. Methods 141, 73–81. https://doi.org/10.1016/j.mimet.2017.08.002. Ladell, B.A., Walleser, L.R., McCalla, S.G., Erickson, R.A., Amberg, J.J., 2019. Ethanol and sodium acetate as a preservation method to delay degradation of environmental DNA. Conserv. Genet. Resour. 11, 83–88. https://doi.org/10.1007/s12686-0170955-2. Limpiyakorn, T., Fürhacker, M., Haberl, R., Chodanon, T., Srithep, P., Sonthiphand, P., 2013. amoA-encoding archaea in wastewater treatment plants: a review. Appl. Microbiol. Biotechnol. 97, 1425–1439. https://doi.org/10.1007/s00253-012-46507. Longmire, J.L., Baker, R.J., Maltbie, M., 1997. Use of “Lysis Buffer” in DNA isolation and its implication for museum collections. Museum Texas Tech. Univ. 1–3. https://doi. org/10.5962/bhl.title.143318. Menke, S., Gillingham, M.A.F., Wilhelm, K., Sommer, S., 2017. Home-made cost effective preservation buffer is a better alternative to commercial preservation methods for microbiome Research. Front. Microbiol. 8, 1–12. https://doi.org/10.3389/ fmicb.2017.00102. Mitchell, K.R., Takacs-Vesbach, C.D., 2008. A comparison of methods for total community DNA preservation and extraction from various thermal environments. J. Ind. Microbiol. Biotechnol. 35, 1139–1147. https://doi.org/10.1007/s10295-0080393-y. Munirasu, S., Haija, M.A., Banat, F., 2016. Use of membrane technology for oil field and refinery produced water treatment - a review. Process Saf. Environ. Protect. 100, 183–202. https://doi.org/10.1016/j.psep.2016.01.010. O’Doherty, K.C., Virani, A., Wilcox, E.S., 2016. The human microbiome and public health: social and ethical considerations. Am. J. Publ. Health 106, 414–420. https:// doi.org/10.2105/AJPH.2015.302989. Osborn, M., 2004. Molecular microbial ecology, first ed. Taylor & Francis, London. https://doi.org/10.4324/9780203503393. Pilliod, D.S., Goldberg, C.S., Arkle, R.S., Waits, L.P., 2014. Factors influencing detection of eDNA from a stream-dwelling amphibian. Mol. Ecol. Resour. 14, 109–116. https://doi.org/10.1111/1755-0998.12159. Pontes, D.S., Lima-Bittencourt, C.I., Chartone-Souza, E., Amaral Nascimento, A.M., 2007. Molecular approaches: advantages and artifacts in assessing bacterial diversity. J. Ind. Microbiol. Biotechnol. 34, 463–473. https://doi.org/10.1007/s10295-0070219-3. Quartaroli, L., Silva, L.C.F., Silva, C.M., Lima, H.S., de Paula, S.O., de Oliveira, V.M., de C´ assia, S., da Silva, M., Kasuya, M.C.M., de Sousa, M.P., Torres, A.P.R., Souza, R.S., Bassin, J.P., da Silva, C.C., 2017. Ammonium removal from high-salinity oilfieldproduced water: assessing the microbial community dynamics at increasing salt concentrations. Appl. Microbiol. Biotechnol. 101, 859–870. https://doi.org/ 10.1007/s00253-016-7902-0. Rissanen, A.J., Kurhela, E., Aho, T., Oittinen, T., Tiirola, M., 2010. Storage of environmental samples for guaranteeing nucleic acid yields for molecular microbiological studies. Appl. Microbiol. Biotechnol. 88, 977–984. https://doi.org/ 10.1007/s00253-010-2838-2. Robertson, L.A., Kuenen, J.G., 1983. Thiosphaera pantotropha gen. Nov. sp. nov., a facultatively anaerobic, facultatively autotrophic sulphur bacterium. Microbiology 129, 2847–2855. https://doi.org/10.1099/00221287-129-9-2847. Rubin, B.E.R., Gibbons, S.M., Kennedy, S., Hampton-Marcell, J., Owens, S., Gilbert, J.A., 2013. Investigating the impact of storage conditions on microbial community composition in soil samples. PLoS One 8, e70460. https://doi.org/10.1371/journal. pone.0070460. Sales, N.G., Wangensteen, O.S., Carvalho, D.C., Mariani, S., 2019. Influence of preservation methods, sample medium and sampling time on eDNA recovery in a neotropical river. Environ. DNA 1, 119–130. https://doi.org/10.1002/edn3.14. Shchegolkova, N.M., Krasnov, G.S., Belova, A.A., Dmitriev, A.A., Kharitonov, S.L., Klimina, K.M., Melnikova, N.V., Kudryavtseva, A.V., 2016. Microbial community structure of activated sludge in treatment plants with different wastewater compositions. Front. Microbiol. 7, 1–15. https://doi.org/10.3389/ fmicb.2016.00090. Silva, C.C., Vieiro, A.F., Dias, A.C.F., Andreote, F.D., Jesus, E.C., Paula, S.O. De, Torres, A.P.R., Santiago, V.M.J., Oliveira, V.M., 2010. Monitoring the bacterial community dynamics in a petroleum refinery wastewater membrane bioreactor fed with a high phenolic load. J. Microbiol. Biotechnol. 20, 21–29. https://doi.org/ 10.4014/jmb.0906.06001. Simister, R.L., Schmitt, S., Taylor, M.W., 2011. Evaluating methods for the preservation and extraction of DNA and RNA for analysis of microbial communities in marine sponges. J. Exp. Mar. Biol. Ecol. 397, 38–43. https://doi.org/10.1016/j. jembe.2010.11.004. Taberlet, P., Bonin, A., Zinger, L., Coissac, E., 2018. Environmental DNA. Oxford University Press, Oxford. https://doi.org/10.1093/oso/9780198767220.001.0001. Tatangelo, V., Franzetti, A., Gandolfi, I., Bestetti, G., Ambrosini, R., 2014. Effect of preservation method on the assessment of bacterial community structure in soil and water samples. FEMS Microbiol. Lett. 356, 32–38. https://doi.org/10.1111/15746968.12475.
Funding This study was funded by CAPES (grant number 001), CNPq (grant number 001), FAPEMIG (grant number 001) and Petrobras (grant numbers 5850.0107013.18.9 and 5850.0108208.18.9). Declaration of competing interest The authors declare the following financial interests/personal re lationships which may be considered as potential competing interests: The authors declare that there is no conflict of interest. Prof. Cynthia Canedo da Silva Department of Microbiology Universidade Federal de Viçosa – UFV, MG - Brazil Acknowledgments ´leo Brasileiro S.A.) for The authors are grateful to Petrobras (Petro ˜o de Amparo a ` Pesquisa do financial support and FAPEMIG (Fundaça Estado de Minas Gerais), CNPq (Conselho Nacional de Desenvolvimento ´gico) and CAPES (Coordenaça ˜o de Aperfeiçoamento Científico e Tecnolo de Pessoal de Nível Superior) for granting scholarships. References Althani, A.A., Marei, H.E., Hamdi, W.S., Nasrallah, G.K., El Zowalaty, M.E., Al Khodor, S., Al-Asmakh, M., Abdel-Aziz, H., Cenciarelli, C., 2016. Human microbiome and its association with health and diseases. J. Cell. Physiol. 231, 1688–1694. https://doi.org/10.1002/jcp.25284. Barnes, M.A., Turner, C.R., 2016. The ecology of environmental DNA and implications for conservation genetics. Conserv. Genet. 17, 1–17. https://doi.org/10.1007/ s10592-015-0775-4. Camacho-Sanchez, M., Burraco, P., Gomez-Mestre, I., Leonard, J.A., 2013. Preservation of RNA and DNA from mammal samples under field conditions. Mol. Ecol. Resour. 13, 663–673. https://doi.org/10.1111/1755-0998.12108. Cui, H., Wang, C., Gu, Z., Zhu, H., Fu, S., Yao, Q., 2014. Evaluation of soil storage methods for soil microbial community using genetic and metabolic fingerprintings. Eur. J. Soil Biol. 63, 55–63. https://doi.org/10.1016/j.ejsobi.2014.05.006. Da Motta, M., Pons, M.N., Roche, N., Vivier, H., Amaral, A.L., Ferreira, E.C., Mota, M., 2003. Estudo do funcionamento de estaç˜ oes de tratamento de esgotos por an´ alise de imagem: validaç˜ oes e estudo de caso. Rev. Eng. Sanit´ aria e Ambient. 8, 170–181. Deutscher, M.P., 2006. Degradation of RNA in bacteria: comparison of mRNA and stable RNA. Nucleic Acids Res. 34, 659–666. https://doi.org/10.1093/nar/gkj472. Dias, R.S., Silva, L.C.F., Eller, M.R., Oliveira, V.M., De Paula, S.O., Silva, C.C., 2014. Metagenomics: library construction and screening methods. In: Metagenomics: Methods, Applications and Perspectives, pp. 45–65. Eichmiller, J.J., Best, S.E., Sorensen, P.W., 2016. Effects of temperature and trophic state on degradation of environmental DNA in lake water. Environ. Sci. Technol. 50, 1859–1867. https://doi.org/10.1021/acs.est.5b05672. Fortis, R. de M., Ortiz, J.P., Lamparelli, C.C., Nieto, R., 2007. An´ alise Computacional Comparativa da Dispers˜ ao da Pluma do Efluente dos Emiss´ arios Submarinos do Tebar. Petrobras. Rev. Bras. Recur. Hídricos 12, 117–132. https://doi.org/ 10.21168/rbrh.v12n1.p117-132. Gabor, E.M., Vries, E.J., Janssen, D.B., 2003. Efficient recovery of environmental DNA for expression cloning by indirect extraction methods. FEMS Microbiol. Ecol. 44, 153–163. https://doi.org/10.1016/S0168-6496(02)00462-2. Hale, V.L., Tan, C.L., Knight, R., Amato, K.R., 2015. Effect of preservation method on spider monkey (Ateles geoffroyi) fecal microbiota over 8 weeks. J. Microbiol. Methods 113, 16–26. https://doi.org/10.1016/j.mimet.2015.03.021. Hammer, O., Harper, D.A.T., Ryan, P.D., 2001. PAST: paleontological statistics software package for education and data analysis. Palaeontol. Electron. 4, 1–9. Henderson, G., Cox, F., Kittelmann, S., Miri, V.H., Zethof, M., Noel, S.J., Waghorn, G.C., Janssen, P.H., 2013. Effect of DNA extraction methods and sampling techniques on the apparent structure of cow and sheep rumen microbial communities. PLoS One 8, e74787. https://doi.org/10.1371/journal.pone.0074787.
7
L.C.F. Silva et al.
International Biodeterioration & Biodegradation 157 (2021) 105139 activated sludge: the impacts of polymerization degree of proteinaceous substrates. Water Res. 129, 133–142. https://doi.org/10.1016/j.watres.2017.11.008. Weißbecker, C., Buscot, F., Wubet, T., 2017. Preservation of nucleic acids by freezedrying for next generation sequencing analyses of soil microbial communities. J. Plant Ecol. 10, 81–90. https://doi.org/10.1093/jpe/rtw042. Williams, K.E., Huyvaert, K.P., Piaggio, A.J., 2016. No filters, no fridges: a method for preservation of water samples for eDNA analysis. BMC Res. Notes 9, 298. https:// doi.org/10.1186/s13104-016-2104-5. Xia, T., SantaLucia, J., Burkard, M.E., Kierzek, R., Schroeder, S.J., Jiao, X., Cox, C., Turner, D.H., 1998. Thermodynamic parameters for an expanded nearest-neighbor model for formation of RNA duplexes with Watson-Crick base pairs †. Biochemistry 37, 14719–14735. https://doi.org/10.1021/bi9809425. Yamanaka, H., Minamoto, T., Matsuura, J., Sakurai, S., Tsuji, S., Motozawa, H., Hongo, M., Sogo, Y., Kakimi, N., Teramura, I., Sugita, M., Baba, M., Kondo, A., 2017. A simple method for preserving environmental DNA in water samples at ambient temperature by addition of cationic surfactant. Limnology 18, 233–241. https://doi. org/10.1007/s10201-016-0508-5.
Tripathi, B.M., Kim, M., Singh, D., Lee-Cruz, L., Lai-Hoe, A., Ainuddin, A.N., Go, R., Rahim, R.A., Husni, M.H.A., Chun, J., Adams, J.M., 2012. Tropical soil bacterial communities in Malaysia: pH dominates in the equatorial tropics too. Microb. Ecol. 64, 474–484. https://doi.org/10.1007/s00248-012-0028-8. Vazoller, R.F., Gomes, L.P., Baldochi, V.M.Z., Villas-Bˆ oas, D.M.F., Badra, R.J., Povinelli, J., 2001. Biodegradability potential of two experimental landfills in Brazil. Braz. J. Microbiol. 32, 87–92. https://doi.org/10.1590/S151783822001000200003. Vreeland, R.H., 2015. Halomonas. In: Bergey’s Manual of Systematics of Archaea and Bacteria. Wiley, pp. 1–19. https://doi.org/10.1002/9781118960608.gbm01190. Wagner, M., Loy, A., Nogueira, R., Purkhold, U., Lee, N., Daims, H., 2002. Microbial community composition and function in wastewater treatment plants. Antonie van Leeuwenhoek, Int. J. Gen. Mol. Microbiol. 81, 665–680. https://doi.org/10.1023/A: 1020586312170. Wang, B.-B., Liu, X.-T., Chen, J.-M., Peng, D.-C., He, F., 2018. Composition and functional group characterization of extracellular polymeric substances (EPS) in
8
Contents lists available at ScienceDirect
International Biodeterioration & Biodegradation journal homepage: www.elsevier.com/locate/ibiod
Comparison of methods for preservation of activated sludge samples for high-throughput nucleic acid sequencing and bacterial diversity analysis Lívia Carneiro Fid´elis Silva a, 1, D´eborah Romaskevis Gomes Lopes a, 1, Helena Santiago Lima a, Larissa Quartaroli a, Maíra Paula de Sousa b, Vinicius de Abreu Waldow b, Rubens Nobumoto Akamine b, S´ergio Oliveira de Paula a, Cynthia Canˆedo da Silva b, * a b
Department of Microbiology, Universidade Federal de Viçosa, Viçosa, Minas Gerais, Brazil Department of Biotechnology, Leopoldo Am´erico Miguez de Mello Research and Development Center (Cenpes), Petrobras, Rio de Janeiro, RJ, Brazil
A R T I C L E I N F O
A B S T R A C T
Keywords: Microbial community structure Sample preservation methods Environmental sample Amplicon sequencing
Preservation of environmental samples is an important step in maintaining the original microbial community until the nucleic acid extraction in the laboratory. Here, we collected activated sludge samples to both imme diately extract nucleic acids (control) and submit to different storage/preservation methods for 48 h before nucleic acids extraction: room temperature storage, storage on ice, direct storage at − 20 ◦ C, rapid freezing in liquid nitrogen followed by storage at − 20 ◦ C, and preservation with TRIzol®, RNAlater®, and Nucleic Acid Preservation (NAP) buffers. Bacterial community was compared by high-throughput 16S rRNA gene sequencing from total DNA and RNA. Among the evaluated methods, TRIzol® and rapid freezing with liquid nitrogen were the least indicated, since they led to significant changes in the microbial community structure and abundance of functional groups involved in ammonia metabolism. Compared to control, storage on ice and NAP preserved more than 80% and 75% of genera at the DNA and RNA level respectively, indicating that these methods are the most suitable for storage/preservation of activated sludge samples intended for nucleic acids sequencing.
1. Introduction Advances in high-throughput sequencing of nucleic acids have resulted in better coverage and characterization of the microbial di versity from environmental samples (Althani et al., 2016; O’Doherty et al., 2016; Sales et al., 2019). However, a successful employment of these methods requires efficient upstream protocols for sample collec tion, storage and preservation, aiming at conserving the fidelity, integ rity and quantity of the genetic material present in the collected samples (Osborn, 2004; Pontes et al., 2007). Protocols for sample collection and nucleic acids extraction must be appropriate to the individual characteristics of each sample, such as its origin, physical state and extracellular matrix, among others. Thus, specific protocols for DNA and RNA preservation and extraction are constantly being revised and optimized to obtain nucleic acids with high yield and quality (Gabor et al., 2003; Henderson et al., 2013; Dias et al.,
2014; Taberlet et al., 2018). However, information about the effect of preservatives and storage conditions aimed at preserving the integrity of genetic material from environmental samples is still limited (Mitchell and Takacs-Vesbach, 2008; Sales et al., 2019). Ideally, the genetic ma terial is extracted immediately after collection, but the establishment of preservation protocols ensures that samples collected in remote areas can be transported to a laboratory with adequate infrastructure for nucleic acid extraction without greatly compromising the integrity of the metagenome and metatranscriptome as originally present in the sample. Inappropriate methods of sample preservation and storage may result in DNA and RNA degradation, compromising inferences about the microbial community structure and making it difficult to identify rare species (Pilliod et al., 2014; Barnes and Turner, 2016; Hinlo et al., 2017). Sample storage at low temperatures, including freezing, is widely applied for sample preservation, as overall microbial metabolism and enzyme activity are reduced in this condition (Pilliod et al., 2014;
* Corresponding author. Av. Peter Henry Rolfs, s/n, Campus Universit´ ario, Laborat´ orio de Microbiologia Ambiental Aplicada, Viçosa, MG, CEP 36570-900, Brazil. E-mail addresses: [email protected] (L.C.F. Silva), [email protected] (D.R.G. Lopes), [email protected] (H.S. Lima), larissaquartaroli@ gmail.com (L. Quartaroli), [email protected] (M.P. de Sousa), [email protected] (V.A. Waldow), [email protected] (R.N. Akamine), [email protected] (S.O. de Paula), [email protected] (C.C. Silva). 1 These authors contributed equally to this work. https://doi.org/10.1016/j.ibiod.2020.105139 Received 12 August 2020; Received in revised form 21 October 2020; Accepted 12 November 2020 Available online 27 November 2020 0964-8305/© 2020 Elsevier Ltd. All rights reserved.
L.C.F. Silva et al.
International Biodeterioration & Biodegradation 157 (2021) 105139
Eichmiller et al., 2016). Other strategies include mixing the sample with saline buffers containing chelating agents (Camacho-Sanchez et al., 2013; Williams et al., 2016; Ladell et al., 2019) or with commercial organic solutions that generally act by protein denaturation (Menke et al., 2017; Yamanaka et al., 2017). The oil and gas industry is a major consumer of water, with a high ratio of volume of water used per volume of oil processed. Oil extraction and refining processes generate large quantities of contaminated efflu ents, whose composition is complex, containing organic compounds as well as ions with varying concentrations, such as chloride (Cl− ), sodium (Na+), calcium (Ca2+), magnesium (Mg2+), sulfide (S2− ) and ammonium (NH+ 4 ). Wastewater discharges containing high concentrations of nitro gen can be toxic to aquatic life, causing eutrophication and oxygen depletion in the aquatic environment, negatively affecting water qual ity. Thus, according to environmental requirements, industrial effluents must be treated before being reused or discharged into a receiving water body (Fortis et al., 2007; Munirasu et al., 2016). Among biological treatments of effluents, activated sludge system is the most commonly used due to its high efficiency (Da Motta et al., 2003). The biomass of activated sludge systems is composed of a complex extracellular matrix and a high microbial diversity that develops global processes related not only to the nitrogen cycle, but also to carbon and phosphorus cycles (Shchegolkova et al., 2016; Quartaroli et al., 2017; Wang et al., 2018). Characterization of the microbial community in activated sludge is important for monitoring populations and metabolic activity inside re actors of effluent treatment plants. However, these treatment plants are commonly located close to the runoff and/or oil refining stations and distant to laboratory facilities, making it necessary to transport these samples to a laboratory for more complex microbiological analysis. Given the importance of microbial characterization of activated sludges and the absence of a specific preservation/storage protocol for this kind of sample, we collected samples of activated sludge from a pilot reactor, extracted nucleic acids immediately after collection (control) and after being submitted for 48 h to seven different storage/preservation methods: storage at room temperature, storage on ice, direct freezing at − 20 ◦ C, rapid freezing in liquid nitrogen followed by storage at − 20 ◦ C, and mixing with a homemade (Nucleic Acid Preservation Solution – NAP) and two commercial (TRIzol® and RNAlater®) preservation buffer. The comparison of the bacterial community structure in terms of genera composition and functional groups with the control treatment indicated the best storage/preservation method to nucleic acid analysis in activated sludge samples.
2.2. Nucleic acid extraction Total DNA and total RNA were extracted from the replicates of each treatment according to the protocol described by Silva et al. (2010), with modifications. The total volume of each treated sample was centrifuged at 12,000 × g for 5 min, and the pellet was washed three times with SET buffer (20 mM Tris, 75 mM NaCl, 25 mM EDTA, pH 7.5). After washing, the pellet was resuspended in 1.0 mL of SET buffer plus 50 μL of a 100 mg/mL lysozyme solution (GE Healthcare, Chicago, USA), and the so lution was incubated inside a 37 ◦ C water bath and stirred every 10 min. Subsequently, 50 μL of a 10 mg/mL proteinase K solution (Sigma-Al drich, St Louis, USA) and 200 μL of SDS 10% w/v (for a final concen tration of 2% w/v) were added to the solution, and it was then incubated at 60 ◦ C for 30 min and stirred every 10 min. The samples were then subjected to three freeze-thaw cycles of 2 min immersed in liquid ni trogen and 2 min at 65 ◦ C. One milliliter of TRIzol® (Qiagen, Hilden, Germany) was added to the solution and homogenized for 2 min. After centrifugation at 10,000 × g for 5 min, the organic phase was used for DNA extraction and the aqueous phase for RNA extraction. In both phases, an equal volume of chloroform/isoamyl alcohol 24:1 (v/v) so lution was added and centrifuged at 10,000 × g for 5 min. The super natant was collected and a 5 M NaCl solution (10% of the total volume) was added to it, followed by the addition of 2 vol of frozen absolute ethanol and centrifugation for 20 min at 10,000 × g. The pellet was washed with 70% ethanol (v/v) and, after drying, it was eluted in 50 μL of ultrapure water. DNA and RNA samples were respectively treated with RNAse (Sigma-Aldrich, St Louis, USA) and DNAse (Promega Cor poration, Madison, USA), following the manufacturer’s recommenda tions. Nucleic acids were quantified using a NanoDrop™ 2000 Spectrophotometer (Thermo Fisher Scientific, Waltham, USA) and visualized in 1% w/v agarose gel. RNA was used for cDNA synthesis with the GoScript™ Reverse Transcriptase kit (Promega Corporation, Wis consin, USA), according to the manufacturer’s recommendations. DNA and cDNA from replicates of each treatment were lyophilized (Benchtop K, VirTis, New York, EUA) and sent for sequencing at the Molecular Research DNA Laboratory (http://www.mrdnalab.com, Shallowater, USA) using the Illumina MiSeq platform. 2.3. Sequencing and analysis of data The sequences obtained were processed using the “MR DNA analysis pipeline” (MR DNA, Shallowater, USA). The barcodes were removed, chimeras and assembled contigs and sequences smaller than 150 bp were also removed. The operational taxonomic units (OTUs) were defined by grouping under 3% divergence (97% similarity). A repre sentative sequence of each OTU was taxonomically classified using BLASTn against a database with curatorship derived from GreenGenes (http://greengenes.lbl.gov), RDPII (http://rdp.cme.msu.edu) and NCBI (http://www.ncbi.nlm.nih.gov). Multivariate analyses and calculation of diversity indices were performed using the PAST software (“Paleon tological Statistics”) (Hammer et al., 2001). All original nucleotide se quences are available at the NCBI Sequence Read Archive (SRA) under Accession Number PRJNA597347.
2. Materials and methods 2.1. Sampling and preservation A sample of activated sludge was collected from a pilot reactor ´rio de Microbiologia Ambiental Aplicada of the operating at the Laborato Universidade Federal de Viçosa, situated in the state of Minas Gerais, Brazil. This pilot reactor was inoculated with a sample of the activated sludge from a petroleum terminal effluent treatment plant and fed with effluent with a saline concentration of 5.5 g.L− 1. Sludge aliquots of 2.0 mL were collected in triplicate for DNA and RNA extraction under different conditions: (1) extraction immediately after collection (con trol); and extraction after 48 h under (2) storage at room temperature; (3) storage on ice using a cooling box; (4) direct freezing at − 20 ◦ C; (5) rapid freezing in liquid nitrogen followed by storage at − 20 ◦ C; (6) preservation in TRIzol® (Invitrogen, Carlsbad, USA) by mixing in a 3:1 vol ratio (preservative-to-sample); (7) preservation in RNAlater® solu tion (Qiagen, Hilden, Germany) by mixing in a 5:1 vol ratio; and (8) preservation in Nucleic Acid Preservation Solution (NAP) by mixing in a 3:1 vol ratio (Camacho-Sanchez et al., 2013). TRIzol® and RNAlater® were added in the ratios recommended by their respective manufac turers. All treatments were performed in triplicate.
2.4. Statistical analysis Comparisons of the mean concentrations of extracted DNA and RNA, diversity indices and functional categories were performed using Mini Tab® 17.1.0 software (Minitab, Inc., State College, USA) by ANOVA, followed by Tukey’s test. P-values less than 0.05 were considered significant. 3. Results Both the DNA and the RNA extracted from the samples showed high purity, with an average absorbance ratio of 260/280 nm equal to 1.83 2
L.C.F. Silva et al.
International Biodeterioration & Biodegradation 157 (2021) 105139
and 2.05, respectively. The different preservation/storage methods evaluated did not affect the amount of total DNA recovered when compared to the control (extraction immediately after extraction) (p > 0.05). In relation to RNA, all treatments evaluated resulted in increased concentration recovered compared to control (mean 1832.87 ng/μL), with the exception of RNAlater® and liquid nitrogen (mean 1048.11 ng/ μL) (p < 0.05) (Fig. 1). Alpha diversity analyses performed from the partial sequencing of the 16S rRNA gene demonstrated that the conditions tested had OTU richness and Shannon and Simpson (1-D) index values similar to the control (p > 0.05), with the exception of freezing with liquid nitrogen, in which these values were 349, 2.59 and 0.82, respectively (p < 0.05) (Table 1). When microbial composition was accessed by cDNA sequencing obtained from transcripts of the 16S rRNA gene, only direct storage at − 20 ◦ C and rapid freezing with liquid nitrogen followed by storage at − 20 ◦ C showed greater richness of OTUs than the control (p < 0.05). For the Shannon index, preservation in TRIzol® was the only method that presented a higher value than the control (p < 0.05) (Table 1). Beta diversity analyses using the Bray-Curtis similarity index showed that the different preservation/storage methods altered the bacterial composition of sludge samples. For both DNA and cDNA, preservation in TRIzol® and direct freezing in liquid nitrogen followed by storage at − 20 ◦ C resulted in the greater dissimilarity of the bacterial community relative to the control. For DNA, preservation in RNAlater® was the condition with the most homogeneous replicates and closest to the control; while for cDNA, the condition with the most homogeneous replicates and closest to the control was direct freezing at − 20 ◦ C (Fig. 2). Obtained sequences resulted in 1008 unique OTUs which were classified into 21 phyla, 38 classes, 79 orders, 164 families and 366 genera. The four most abundant genera in all treatments were Hal omonas, Tindallia, Spirochaeta and Halanaerobium, representing 83.64 ± 2.43% (mean ± SD) of the obtained DNA sequences, and 93.72 ± 1.83% (mean ± SD) of the obtained cDNA sequences. TRIzol® was the
preservative that most disturbed the taxonomic profile of sludge samples in relation to the control: the genera Spirochaeta, Rickettsia and Achole plasma were enriched for DNA, while Spirochaeta was enriched and Halanaerobium was diminished for cDNA, presenting less abundance in relation to the control (Fig. 3). Considering the genera involved in ni trogen metabolism, both for DNA and cDNA, the abundance of Nitra tireductor was different among the treatments (p < 0.05). When compared to the control, no treatment presented differences in relation to Nitratireductor abundance when evaluating DNA, while preservation in NAP, TRIzol® and liquid nitrogen freezing provided enrichment of Nitratireductor when evaluating cDNA (p < 0.05). When cDNA was evaluated, an increase in the abundance of Nitrospira was observed with preservation in TRIzol® and liquid nitrogen freezing (p < 0.05). For DNA, there was no difference in this genus between treatments (p > 0.05). The abundance of the genus Nitrosomonas was not altered (Fig. 4). Of 96 genera identified with DNA sequencing, approximately 56% were shared among all treatments, and of 124 genera identified with cDNA sequencing, 32% were shared among all treatments. Storage on ice was the storage/preservation method with the highest number of genera shared with the control (84% for DNA and 83% for cDNA), and directly freezing at − 20 ◦ C was the condition with the lowest number of genera shared with the control (77% for DNA and 47% for cDNA) (Fig. 5). 4. Discussion Considering logistical and practical difficulties of extracting nucleic acids immediately after sample collection, it is necessary to investigate safe and efficient ways of preserving samples without compromising the microbial community structure and nucleic acids integrity. Molecular studies demonstrated that physical methods of storage and preservation, especially those involving low temperatures, are more efficient in maintaining the microbial community structure than chemical methods (Rubin et al., 2013; Cui et al., 2014; Tatangelo et al., 2014). Chemical preservatives interact directly with cellular and matrix components and
Fig. 1. Concentration of DNA and RNA extracted from activated sludge samples subjected to different storage/preservation methods. Control represents the im mediate extraction after sample collection, while storage/preservation methods are represented as: RT (room temperature); NAP (Nucleic Acid Preservation solu tion); Ice (storage on ice); TZ (TRIzol®); RL (RNAlater®); NIT (rapid freezing with liquid nitrogen followed by storage at − 20 ◦ C) and Freeze (direct freezing at − 20 ◦ C). The letters above the bars represent the statistical differences between the average DNA or RNA concentrations. Equal letters represent that there was no statistical difference at 95% of confidence obtained by ANOVA followed by Tukey’s test. 3
L.C.F. Silva et al.
International Biodeterioration & Biodegradation 157 (2021) 105139
Table 1 Diversity indices of activated sludge samples submitted to different storage/preservation methods. DNA
cDNA
OTU Richness Simpson (1-D) Shannon Evenness Chao OTU Richness Simpson (1-D) Shannon Evenness Chao
Control
RT
NAP
Ice
TZ
RL
NIT
Freeze
426.33a 0.85a 2.83a 0.04 702.70 313.67b 0.70abc 1.93bc 0.02abc 511.20c
385.67ab 0.83ab 2.73ab 0.04 589.37 290.67b 0.74ab 2.15ab 0.03a 428.00c
413.67a 0.85a 2.82a 0.04 665.13 335.00b 0.72abc 2.07abc 0.02 ab 485.70c
413.00a 0.83ab 2.73ab 0.04 660.30 329.33b 0.74ab 2.17ab 0.03ab 499.17c
382.00ab 0.85a 2.69ab 0.04 564.73 372.67b 0.75a 2.28a 0.03ab 565.60bc
387.00ab 0.83ab 2.66ab 0.04 623.60 336.33b 0.68bc 1.90bc 0.02bc 616.47bc
349.67b 0.82b 2.59b 0.04 554.90 585.33a 0.66c 1.84c 0.01d 1035.73a
415.33a 0.83ab 2.68ab 0.04 654.47 510.33a 0.68c 1.98abc 0.01cd 855.63ab
Control, extraction of genetic material immediately after collection; RT, storage at room temperature; NAP, Nucleic Acid Preservation solution; Ice, storage on ice; TZ, TRIzol®; RL, RNAlater®; NIT, freezing in liquid nitrogen followed by storage at − 20 ◦ C; and Freeze, direct freezing at − 20 ◦ C. Averages of the indices followed by the same letter have no statistical difference at 95% confidence obtained by ANOVA followed by Tukey’s test. Fig. 2. Principal coordinate analysis (PCoA) based on the Bray-Curtis similarity index of activated sludge samples submitted to different storage/preservation methods. Bacterial diversity was accessed by partial sequencing of the 16S rRNA gene from total DNA (a), samples represented by circles, and cDNA synthesized from total RNA (b), samples represented by triangles. Control represents the extraction of genetic material immediately after collection, while storage/preser vation methods are represented as: RT, storage at room temperature; NAP, Nucleic Acid Preservation solution; Ice, storage on ice; TZ, TRIzol®; RL, RNA later®; NIT, rapid freezing with liquid nitrogen fol lowed by storage at − 20 ◦ C; and Freeze, direct freezing at − 20 ◦ C.
alter the pH of the solution, resulting in discrepancies in bacterial composition between the samples (Rissanen et al., 2010; Tripathi et al., 2012; Tatangelo et al., 2014). Preservation solutions rich in salts (which favor the precipitation of nucleic acids) and chelators of enzymatic co factors (e.g., EDTA) can be purchased commercially, such as RNAlater®, TRIzol®, DNAzol® and LifeGuard™, or produced in the laboratory (“homemade”), such as NAP solution and Longmire’s buffer (Cama cho-Sanchez et al., 2013; Longmire et al., 1997). Additionally, the use of chemical preservatives usually requires large volumes of the chosen preservative, since the preservative-to-sample ratio normally used is 3:1. In this work, relevant information was provided on the effect of different preservation/storage conditions of activated sludge samples on the recovered nucleic acid concentration and on the taxonomic and meta bolic function diversity of the bacterial community. When compared to the control (extraction immediately after
collection), the evaluated preservation/storage methods did not affect the amount of total DNA recovered. In contrast, two methods resulted in a reduced RNA concentration when compared to the control (RNAlater® and rapid freezing with liquid nitrogen followed by storage at − 20 ◦ C), and the other five methods resulted in increased RNA concentration in comparison to the control. These results may be associated with (1) the fact that rates of biosynthesis and biodegradation of RNA transcripts are higher when compared to genomic DNA due to regulation of gene expression in response to environmental conditions, and (2) RNA is more susceptible to hydrolysis than DNA, making it more sensitive to preservation/storage methods (Xia et al., 1998; Deutscher, 2006). For both DNA and RNA, the genus Halomonas was the most abundant in all treatments, as the samples of sludge were all inoculated in a bioreactor fed with saline effluent (5.5 g.L− 1 of salt). This genus com prises halophilic bacteria isolated from saline environments around the 4
L.C.F. Silva et al.
International Biodeterioration & Biodegradation 157 (2021) 105139
Fig. 3. Taxonomic composition at the genus level of activated sludge samples submitted to different stor age/preservation methods. Bacterial diversity was accessed by partial sequencing of the 16S rRNA gene from total DNA (a) and cDNA synthesized from total RNA (b). Control represents the extraction of genetic material immediately after collection, while storage/ preservation methods are represented as: RT, storage at room temperature; NAP, Nucleic Acid Preservation solution; Ice, storage on ice; TZ, TRIzol®; RL, RNA later®; NIT, rapid freezing with liquid nitrogen fol lowed by storage at − 20 ◦ C; and Freeze, direct freezing at − 20 ◦ C. Others represents genera with relative abundance less than 0.2% in the control.
Fig. 4. Heatmap representing the relative abun dances of genera involved in nitrogen metabolism in sludge activated samples. Genera accessed by partial sequencing of the 16S rRNA gene from total DNA (a) and cDNA synthesized from total RNA (b). Control represents the extraction of genetic material imme diately after collection, while storage/preservation methods are represented as: RT, storage at room temperature; NAP, Nucleic Acid Preservation solu tion; Ice, storage on ice; TZ, TRIzol®; RL, RNAlater®; NIT, rapid freezing with liquid nitrogen followed by storage at − 20 ◦ C; and Freeze, direct freezing at − 20 ◦ C. Identical letters have no statistical difference at 95% confidence obtained by ANOVA followed by Tukey’s test.
world, including solar salt production facilities, intertidal estuaries, the open ocean and hypersaline lakes (Vreeland, 2015). Our results indicate that preservation in TRIzol® and rapid freezing in liquid nitrogen fol lowed by storage at − 20 ◦ C were the worst methods, since the microbial community diverged most from the control. TRIzol® is used for main taining nucleic acids integrity, as it contains high concentrations of guanidine thiocyanate and acid phenol, which are inhibitors of endo nucleases (Simister et al., 2011). Rapid freezing in liquid nitrogen is a widely-used method of cryogenics for cell cultures in general, since it leads to the formation of smaller water crystals inside the cell, making it possible to maintain its properties after thawing (Weißbecker et al., 2017). Nonetheless, both TRIzol® and liquid nitrogen are expensive and difficult to handle in the field, since TRIzol® has high toxicity and liquid
nitrogen presents the risk of explosion during its transport. Storage on ice and preservation in NAP solution were the most efficient methods for maintaining the microbial community structure in samples: these methods covered more than 80% of the genera at the DNA level and more than 75% at the RNA level in comparison to the control. According to Weißbecker et al. (2017), an overlap of 70–80% of OTUs with the control treatment represents a satisfactory value for a storage/preservation method to be considered good. Sample storage at low temperatures reduces overall enzymatic activity, reducing the degradation of nucleic acids. In addition, mild cooling of samples, unlike freezing, ensures that water crystals are not formed, and thus do not cause cell lysis during thawing (Weißbecker et al., 2017). The main objective of biological treatment of liquid effluents by 5
L.C.F. Silva et al.
International Biodeterioration & Biodegradation 157 (2021) 105139
Fig. 5. Venn diagrams representing the unique genera for each storage/preservation method and those shared with the control of activated sludge samples. Bacterial diversity was accessed by partial sequencing of the 16S rRNA gene from total DNA (a) and cDNA synthesized from total RNA (b). Control represents the extraction of genetic material immediately after collection, while storage/preservation methods are represented as: RT, storage at room temperature; NAP, Nucleic Acid Pres ervation solution; Ice, storage on ice; TZ, TRIzol®; RL, RNAlater®; NIT, rapid freezing with liquid nitrogen followed by storage at − 20 ◦ C; and Freeze, direct freezing at − 20 ◦ C. Only the genera observed in all replicates were considered. Below each diagram is the percentage of treatment sharing in relation to the control.
activated sludge is the removal of pollutants, including ammonia (Da Motta et al., 2003). The biological removal of ammonia can be per formed by different processes involving distinct groups of microorgan isms, through either autotrophic or heterotrophic metabolism, and under both aerobic and anaerobic conditions (Robertson and Kuenen, 1983; Vazoller et al., 2001; Limpiyakorn et al., 2013). The cDNA sequencing data demonstrated that the relative abundances of genera involved in autotrophic nitrification, Nitrosomonas and Nitrospira (Wagner et al., 2002), and anaerobic denitrification, Nitratireductor (Labb´e et al., 2004), were altered by the different preservation/storage methods evaluated, indicating that the chosen method may compromise the characterization of functional groups in sludge samples at the RNA level. Thus, studies aiming at the characterization of metabolically active microorganisms and/or gene expression studies should be more cautious regarding the choice of sample preservation/storage method. Despite being a widely used preservative solution, studies have shown that RNAlater® is not efficient for preserving the microbial sample community. Lackey et al. (2017) evaluated the effect of preser vation with commercial solutions and the storage time of human milk samples maintained at 37 ◦ C on the composition of the bacterial com munity (sequencing of the V1-V3 region of the 16S rRNA gene) and observed that over time the preservation with RNAlater® is compro mised at that temperature. According to the manufacturer, this solution is efficient for preservation for up to 1 day at 37 ◦ C and up to 1 week at 23 ◦ C (room temperature). For samples of monkey feces, Hale et al. (2015) observed that the microbial composition and abundance of samples preserved in RNAlater® differ from the fresh sample. Menke
et al. (2017) observed that the NAP solution stabilizes the microbial community better than commercial preservative media (RNAlater® and RNA/DNA Shield™) for samples of sheep feces, making this homemade solution a viable alternative, in terms of both cost and efficiency, for the study of environmental samples from remote areas in the absence of ice or refrigerators. High-throughput sequencing, as well as other molecular applica tions, requires that the extracted genetic material is of high quantity and good quality in order to adequately access the microbial groups and metabolic functions present in the sample. In this work, we observed that the evaluated preservation/storage methods differentially altered the microbial community structure, but all of them enabled the recovery of DNA and RNA in sufficient concentration and quality. In addition, it was suggested that for activated sludge samples, storage in ice or pres ervation in NAP solution are efficient for preserving the DNA and RNA of the microbial community in the sample. However, the choice of a preservation method will depend on several factors, such as the nature and physicochemical characteristics of the sample, logistics, cost, availability, safety and infrastructure available at the time of collection, as well as the focus of the study (taxonomic profile of the microbial community or gene expression, for example). 5. Conclusion This study provided evidence that freezing of activated sludge sam ples in liquid nitrogen followed by storage at − 20 ◦ C and preservation in TRIzol® results in significant changes in the total structure of the 6
L.C.F. Silva et al.
International Biodeterioration & Biodegradation 157 (2021) 105139
microbial community and the relative abundance of functional groups involved in ammonia metabolism, for both DNA and RNA. Moreover, our results indicated that preservation in NAP solution or storage on ice enabled greater coverage of the total and metabolically active microbial community, accessed by both DNA and cDNA (RNA) sequencing, and therefore the application of these methods is recommended for molec ular and functional studies of activated sludge samples, both from DNA and RNA.
Hinlo, R., Gleeson, D., Lintermans, M., Furlan, E., 2017. Methods to maximise recovery of environmental DNA from water samples. PLoS One 12, e0179251. https://doi. org/10.1371/journal.pone.0179251. Labb´e, N., Parent, S., Villemur, R., 2004. Nitratireductor aquibiodomus gen. nov., sp. nov., a novel α-proteobacterium from the marine denitrification system of the Montreal Biodome (Canada). Int. J. Syst. Evol. Microbiol. 54, 269–273. https://doi.org/ 10.1099/ijs.0.02793-0. Lackey, K.A., Williams, J.E., Price, W.J., Carrothers, J.M., Brooker, S.L., Shafii, B., McGuire, M.A., McGuire, M.K., 2017. Comparison of commercially-available preservatives for maintaining the integrity of bacterial DNA in human milk. J. Microbiol. Methods 141, 73–81. https://doi.org/10.1016/j.mimet.2017.08.002. Ladell, B.A., Walleser, L.R., McCalla, S.G., Erickson, R.A., Amberg, J.J., 2019. Ethanol and sodium acetate as a preservation method to delay degradation of environmental DNA. Conserv. Genet. Resour. 11, 83–88. https://doi.org/10.1007/s12686-0170955-2. Limpiyakorn, T., Fürhacker, M., Haberl, R., Chodanon, T., Srithep, P., Sonthiphand, P., 2013. amoA-encoding archaea in wastewater treatment plants: a review. Appl. Microbiol. Biotechnol. 97, 1425–1439. https://doi.org/10.1007/s00253-012-46507. Longmire, J.L., Baker, R.J., Maltbie, M., 1997. Use of “Lysis Buffer” in DNA isolation and its implication for museum collections. Museum Texas Tech. Univ. 1–3. https://doi. org/10.5962/bhl.title.143318. Menke, S., Gillingham, M.A.F., Wilhelm, K., Sommer, S., 2017. Home-made cost effective preservation buffer is a better alternative to commercial preservation methods for microbiome Research. Front. Microbiol. 8, 1–12. https://doi.org/10.3389/ fmicb.2017.00102. Mitchell, K.R., Takacs-Vesbach, C.D., 2008. A comparison of methods for total community DNA preservation and extraction from various thermal environments. J. Ind. Microbiol. Biotechnol. 35, 1139–1147. https://doi.org/10.1007/s10295-0080393-y. Munirasu, S., Haija, M.A., Banat, F., 2016. Use of membrane technology for oil field and refinery produced water treatment - a review. Process Saf. Environ. Protect. 100, 183–202. https://doi.org/10.1016/j.psep.2016.01.010. O’Doherty, K.C., Virani, A., Wilcox, E.S., 2016. The human microbiome and public health: social and ethical considerations. Am. J. Publ. Health 106, 414–420. https:// doi.org/10.2105/AJPH.2015.302989. Osborn, M., 2004. Molecular microbial ecology, first ed. Taylor & Francis, London. https://doi.org/10.4324/9780203503393. Pilliod, D.S., Goldberg, C.S., Arkle, R.S., Waits, L.P., 2014. Factors influencing detection of eDNA from a stream-dwelling amphibian. Mol. Ecol. Resour. 14, 109–116. https://doi.org/10.1111/1755-0998.12159. Pontes, D.S., Lima-Bittencourt, C.I., Chartone-Souza, E., Amaral Nascimento, A.M., 2007. Molecular approaches: advantages and artifacts in assessing bacterial diversity. J. Ind. Microbiol. Biotechnol. 34, 463–473. https://doi.org/10.1007/s10295-0070219-3. Quartaroli, L., Silva, L.C.F., Silva, C.M., Lima, H.S., de Paula, S.O., de Oliveira, V.M., de C´ assia, S., da Silva, M., Kasuya, M.C.M., de Sousa, M.P., Torres, A.P.R., Souza, R.S., Bassin, J.P., da Silva, C.C., 2017. Ammonium removal from high-salinity oilfieldproduced water: assessing the microbial community dynamics at increasing salt concentrations. Appl. Microbiol. Biotechnol. 101, 859–870. https://doi.org/ 10.1007/s00253-016-7902-0. Rissanen, A.J., Kurhela, E., Aho, T., Oittinen, T., Tiirola, M., 2010. Storage of environmental samples for guaranteeing nucleic acid yields for molecular microbiological studies. Appl. Microbiol. Biotechnol. 88, 977–984. https://doi.org/ 10.1007/s00253-010-2838-2. Robertson, L.A., Kuenen, J.G., 1983. Thiosphaera pantotropha gen. Nov. sp. nov., a facultatively anaerobic, facultatively autotrophic sulphur bacterium. Microbiology 129, 2847–2855. https://doi.org/10.1099/00221287-129-9-2847. Rubin, B.E.R., Gibbons, S.M., Kennedy, S., Hampton-Marcell, J., Owens, S., Gilbert, J.A., 2013. Investigating the impact of storage conditions on microbial community composition in soil samples. PLoS One 8, e70460. https://doi.org/10.1371/journal. pone.0070460. Sales, N.G., Wangensteen, O.S., Carvalho, D.C., Mariani, S., 2019. Influence of preservation methods, sample medium and sampling time on eDNA recovery in a neotropical river. Environ. DNA 1, 119–130. https://doi.org/10.1002/edn3.14. Shchegolkova, N.M., Krasnov, G.S., Belova, A.A., Dmitriev, A.A., Kharitonov, S.L., Klimina, K.M., Melnikova, N.V., Kudryavtseva, A.V., 2016. Microbial community structure of activated sludge in treatment plants with different wastewater compositions. Front. Microbiol. 7, 1–15. https://doi.org/10.3389/ fmicb.2016.00090. Silva, C.C., Vieiro, A.F., Dias, A.C.F., Andreote, F.D., Jesus, E.C., Paula, S.O. De, Torres, A.P.R., Santiago, V.M.J., Oliveira, V.M., 2010. Monitoring the bacterial community dynamics in a petroleum refinery wastewater membrane bioreactor fed with a high phenolic load. J. Microbiol. Biotechnol. 20, 21–29. https://doi.org/ 10.4014/jmb.0906.06001. Simister, R.L., Schmitt, S., Taylor, M.W., 2011. Evaluating methods for the preservation and extraction of DNA and RNA for analysis of microbial communities in marine sponges. J. Exp. Mar. Biol. Ecol. 397, 38–43. https://doi.org/10.1016/j. jembe.2010.11.004. Taberlet, P., Bonin, A., Zinger, L., Coissac, E., 2018. Environmental DNA. Oxford University Press, Oxford. https://doi.org/10.1093/oso/9780198767220.001.0001. Tatangelo, V., Franzetti, A., Gandolfi, I., Bestetti, G., Ambrosini, R., 2014. Effect of preservation method on the assessment of bacterial community structure in soil and water samples. FEMS Microbiol. Lett. 356, 32–38. https://doi.org/10.1111/15746968.12475.
Funding This study was funded by CAPES (grant number 001), CNPq (grant number 001), FAPEMIG (grant number 001) and Petrobras (grant numbers 5850.0107013.18.9 and 5850.0108208.18.9). Declaration of competing interest The authors declare the following financial interests/personal re lationships which may be considered as potential competing interests: The authors declare that there is no conflict of interest. Prof. Cynthia Canedo da Silva Department of Microbiology Universidade Federal de Viçosa – UFV, MG - Brazil Acknowledgments ´leo Brasileiro S.A.) for The authors are grateful to Petrobras (Petro ˜o de Amparo a ` Pesquisa do financial support and FAPEMIG (Fundaça Estado de Minas Gerais), CNPq (Conselho Nacional de Desenvolvimento ´gico) and CAPES (Coordenaça ˜o de Aperfeiçoamento Científico e Tecnolo de Pessoal de Nível Superior) for granting scholarships. References Althani, A.A., Marei, H.E., Hamdi, W.S., Nasrallah, G.K., El Zowalaty, M.E., Al Khodor, S., Al-Asmakh, M., Abdel-Aziz, H., Cenciarelli, C., 2016. Human microbiome and its association with health and diseases. J. Cell. Physiol. 231, 1688–1694. https://doi.org/10.1002/jcp.25284. Barnes, M.A., Turner, C.R., 2016. The ecology of environmental DNA and implications for conservation genetics. Conserv. Genet. 17, 1–17. https://doi.org/10.1007/ s10592-015-0775-4. Camacho-Sanchez, M., Burraco, P., Gomez-Mestre, I., Leonard, J.A., 2013. Preservation of RNA and DNA from mammal samples under field conditions. Mol. Ecol. Resour. 13, 663–673. https://doi.org/10.1111/1755-0998.12108. Cui, H., Wang, C., Gu, Z., Zhu, H., Fu, S., Yao, Q., 2014. Evaluation of soil storage methods for soil microbial community using genetic and metabolic fingerprintings. Eur. J. Soil Biol. 63, 55–63. https://doi.org/10.1016/j.ejsobi.2014.05.006. Da Motta, M., Pons, M.N., Roche, N., Vivier, H., Amaral, A.L., Ferreira, E.C., Mota, M., 2003. Estudo do funcionamento de estaç˜ oes de tratamento de esgotos por an´ alise de imagem: validaç˜ oes e estudo de caso. Rev. Eng. Sanit´ aria e Ambient. 8, 170–181. Deutscher, M.P., 2006. Degradation of RNA in bacteria: comparison of mRNA and stable RNA. Nucleic Acids Res. 34, 659–666. https://doi.org/10.1093/nar/gkj472. Dias, R.S., Silva, L.C.F., Eller, M.R., Oliveira, V.M., De Paula, S.O., Silva, C.C., 2014. Metagenomics: library construction and screening methods. In: Metagenomics: Methods, Applications and Perspectives, pp. 45–65. Eichmiller, J.J., Best, S.E., Sorensen, P.W., 2016. Effects of temperature and trophic state on degradation of environmental DNA in lake water. Environ. Sci. Technol. 50, 1859–1867. https://doi.org/10.1021/acs.est.5b05672. Fortis, R. de M., Ortiz, J.P., Lamparelli, C.C., Nieto, R., 2007. An´ alise Computacional Comparativa da Dispers˜ ao da Pluma do Efluente dos Emiss´ arios Submarinos do Tebar. Petrobras. Rev. Bras. Recur. Hídricos 12, 117–132. https://doi.org/ 10.21168/rbrh.v12n1.p117-132. Gabor, E.M., Vries, E.J., Janssen, D.B., 2003. Efficient recovery of environmental DNA for expression cloning by indirect extraction methods. FEMS Microbiol. Ecol. 44, 153–163. https://doi.org/10.1016/S0168-6496(02)00462-2. Hale, V.L., Tan, C.L., Knight, R., Amato, K.R., 2015. Effect of preservation method on spider monkey (Ateles geoffroyi) fecal microbiota over 8 weeks. J. Microbiol. Methods 113, 16–26. https://doi.org/10.1016/j.mimet.2015.03.021. Hammer, O., Harper, D.A.T., Ryan, P.D., 2001. PAST: paleontological statistics software package for education and data analysis. Palaeontol. Electron. 4, 1–9. Henderson, G., Cox, F., Kittelmann, S., Miri, V.H., Zethof, M., Noel, S.J., Waghorn, G.C., Janssen, P.H., 2013. Effect of DNA extraction methods and sampling techniques on the apparent structure of cow and sheep rumen microbial communities. PLoS One 8, e74787. https://doi.org/10.1371/journal.pone.0074787.
7
L.C.F. Silva et al.
International Biodeterioration & Biodegradation 157 (2021) 105139 activated sludge: the impacts of polymerization degree of proteinaceous substrates. Water Res. 129, 133–142. https://doi.org/10.1016/j.watres.2017.11.008. Weißbecker, C., Buscot, F., Wubet, T., 2017. Preservation of nucleic acids by freezedrying for next generation sequencing analyses of soil microbial communities. J. Plant Ecol. 10, 81–90. https://doi.org/10.1093/jpe/rtw042. Williams, K.E., Huyvaert, K.P., Piaggio, A.J., 2016. No filters, no fridges: a method for preservation of water samples for eDNA analysis. BMC Res. Notes 9, 298. https:// doi.org/10.1186/s13104-016-2104-5. Xia, T., SantaLucia, J., Burkard, M.E., Kierzek, R., Schroeder, S.J., Jiao, X., Cox, C., Turner, D.H., 1998. Thermodynamic parameters for an expanded nearest-neighbor model for formation of RNA duplexes with Watson-Crick base pairs †. Biochemistry 37, 14719–14735. https://doi.org/10.1021/bi9809425. Yamanaka, H., Minamoto, T., Matsuura, J., Sakurai, S., Tsuji, S., Motozawa, H., Hongo, M., Sogo, Y., Kakimi, N., Teramura, I., Sugita, M., Baba, M., Kondo, A., 2017. A simple method for preserving environmental DNA in water samples at ambient temperature by addition of cationic surfactant. Limnology 18, 233–241. https://doi. org/10.1007/s10201-016-0508-5.
Tripathi, B.M., Kim, M., Singh, D., Lee-Cruz, L., Lai-Hoe, A., Ainuddin, A.N., Go, R., Rahim, R.A., Husni, M.H.A., Chun, J., Adams, J.M., 2012. Tropical soil bacterial communities in Malaysia: pH dominates in the equatorial tropics too. Microb. Ecol. 64, 474–484. https://doi.org/10.1007/s00248-012-0028-8. Vazoller, R.F., Gomes, L.P., Baldochi, V.M.Z., Villas-Bˆ oas, D.M.F., Badra, R.J., Povinelli, J., 2001. Biodegradability potential of two experimental landfills in Brazil. Braz. J. Microbiol. 32, 87–92. https://doi.org/10.1590/S151783822001000200003. Vreeland, R.H., 2015. Halomonas. In: Bergey’s Manual of Systematics of Archaea and Bacteria. Wiley, pp. 1–19. https://doi.org/10.1002/9781118960608.gbm01190. Wagner, M., Loy, A., Nogueira, R., Purkhold, U., Lee, N., Daims, H., 2002. Microbial community composition and function in wastewater treatment plants. Antonie van Leeuwenhoek, Int. J. Gen. Mol. Microbiol. 81, 665–680. https://doi.org/10.1023/A: 1020586312170. Wang, B.-B., Liu, X.-T., Chen, J.-M., Peng, D.-C., He, F., 2018. Composition and functional group characterization of extracellular polymeric substances (EPS) in
8