MSc Biological Sciences
Limnology & Oceanography
Research Project
Sulfite disproportionation by the extremophilic soda lake bacterium
Desulfurivibrio alkaliphilus.
by
Marijn van Doorn
10001121
36 ECTS Credits
February 2015 – March 2019
Assessor:
Examiner:
Prof. dr. Gerard Muijzer
Prof. dr. Lucas J. Stal
Abstract
Extremophilic bacteria originating from soda lakes are known to play an important role in the element cycling of these environments. The sulfur cycle is one of the most important cycles in these lakes. Even though microbial-regulated soda lake element cycling gained increased attention over the past decades, many aspects of how these systems function still remain unknown. This study focussed on the disproportionation of sulfite by the delta-proteobacterium Desufurivibrio alkaliphilus strain AHT2. After determining what the prefered substrate was for D. alkaliphilus, the bacterium was grown either with or without an electron donor. The latter forcing the bacteria to perform disproportionation. Under both conditions the bacteria showed growth, meaning that they have the ability to perform disproportionation of sulfite. Candidate genes for sulfite oxidation were selected. Homologous genes were selected and aligned with the full genome of D. alkaliphilus. The target gene with the highest query cover and identity, AprAB, was selected for PCR. After designing and testing primers, a PCR-analysis was performed. This confirmed that AprAB is present in the genome of D.
alkaliphilus. RT-qPCR was subsequently performed on AprAB and DsrAB (a gene known to play
a role in the reduction of sulfite). Results from the RT-qPCR show no significant relative increase in the transcription of both genes when the bacteria are forced to perform disproportionation.
Keywords: Sulfite, Disproportionation, Desulfurivibrio alkaliphilus, Soda lakes, Extremophiles, Deltaproteobacteria, Sulfur metabolism.
Introduction
Soda lakes are found all around the world in arid regions with high evaporation rates and are characterized as extreme environments due to their high alkalinity and salinity (Sorokin et al., 2014). Extreme environments like this often harbor microbes able the cope with these circumstances (Van den Burg, 2003). The challenges extreme environments offer, force organisms living there to adapt in more radical ways than organisms living in milder habitats. With current changes in the global climate and the subsequently observed shifts in many ecosystems (Walther et al., 2002), it might be these organisms, holding an important key to adaptation, leading to increased survival (Pikuta, Hoover & Tang, 2007). Even though soda lakes can be found in many different areas around the world, not much is known about the functioning of these ecosystems. The role organisms play and the effects they have on this environment, like bacteria and their contribution to the biochemical cycles, are currently gaining increasing attention.
Soda lakes contain different microbial communities, ranging from alpha- and beta-proteobacteria to clostridia (Tourova, Grechnikova, Kuznetsov & Sorokin, 2014). Microbes are known to fulfill an important role in soda lake element cycling (Foti et al., 2007). The research on isolated microorganisms can provide insights in the mechanisms of element cycling (Falkowski, Fenchel & Delong, 2008). One of the key elements in these soda lakes is sulfur (Sorokin et al., 2014; Sorokin, Kuenen & Muyzer, 2011). Sulfur is an important element because of its function in proteins and its function as an electron donor/acceptor in the sulfur cycle. This broad applicability is mainly due to the large range of valencies of the sulfur atom, with molecules having valencies ranging from +6 (sulfate) to -2 (sulfide). Most of the redox reactions in the sulfur cycle do not happen spontaneously under ambient conditions but are mediated by prokaryotes (Sorokin et al., 2011). These microorganisms possess the ability to use these inorganic sulfur compounds in their energy management, either as electron acceptor or as electron donor, providing energy for their metabolic pathways (Barton, 2013). In the research presented here, the main focus lies on sulfite. Sulfite is present in aqueous environments as the sulfite anion (SO32-). The sulfite anion tends to get oxidized and is
therefore considered unstable in the presence of oxygen (Lindgren, Cedergren, & Lindberg, 1982). Sulfite is well known as a food preservative (Li & Zhao, 2006), the high affinity of sulfite for oxygen prevents the food itself from oxidizing (Wedzicha, 1992). Furthermore, sulfite is used to control bacterial growth (Lester, 1995). There are however a number of bacteria that can use sulfite for their metabolic needs (Hansen, 1994). These bacteria either oxidize sulfite (to sulfate) or reduce sulfite (to hydrogen sulfide) and can therefore be classified as either sulfur oxidizing bacteria (SOB) or sulfur reducing bacteria (SRB).
The present research mainly focuses on sulfur reducing bacteria. Well known bacteria using the reduction of sulfite for their metabolic needs are bacteria from the genera Desulfovibrio, Desulfitobacterium, Desulfotomaculum & Thermodesulfobacterium (Karkhoff-Schweizer, Huber & Voordouw, 1995; Klein et al., 2001). Genes known to play a role in the reduction of sulfite are Dsr (Dissimilatory Sulfite Reductase) (Klein et al., 2001; Wagner, Roger, Flax, Brusseau & Stahl, 1998) and Dsv (desulfoviridin) (Karkhoff-Schweizer et al., 1995a). The reduction of sulfite can be characterized by the following formula.
Recent research shows that some sulfite reducing bacteria, living in the anaerobic sediments of soda lakes, adopted a different method for using sulfite to meet their metabolic needs, this process is known as disproportionation (Finster, 2008). During disproportionation sulfite is reduced and oxidized simultaneously. Disproportionation in sulfite reducing bacteria is comparable to fermentation of organic compounds and is sometimes referred to as ‘inorganic fermentation’ (Finster, 2008). After disproportionation, sulfite is left in an overall reduced state. Even though the energy yield from this process is lower than sulfite reduction, it enables these bacteria to conduct their metabolic processes in the absence of an electron donor (Amend, Edwards & Lyons, 2004). The formula that describes sulfite disproportionation is defined as follows.
In this study, we set out to contribute to the understanding of which of the processes in the biochemical sulfur cycle, with a specific focus on the disproportionation of sulfite, are regulated by bacteria. Furthermore, we want to determine which genes are expressed in order to perform this metabolic process.
The chosen test organism is the deltaproteobacteria Desulfurivibrio alkaliphilus, strain AHT2, a bacterium living in anaerobic conditions, sampled from sediments from soda lakes in Egypt (Sorokin, Tourova, Mußmann & Muyzer, 2008), of which the full genome is sequenced (Melton
et al., 2016). Understanding the role these bacteria play in the biochemical sulfur cycle, under
these extreme conditions, will provide insights in how these processes are regulated. Since the biochemical sulfur cycle is connected to most other element cycles, like the nitrogen- and carbon cycle, a better understanding of the biochemical sulfur cycle might lead to a better understanding of biochemical element cycling as a whole. Different element cycles are connected through processes like mineralization and immobilization (Stevenson & Cole, 1999), in which either organic matter gets decomposed or oxidized, or inorganic matter is converted to organic matter by plants or microbes.
Furthermore, this research could provide possible clues for the use of these kind of bacteria for bioremediation, for instance desulfurization of wastewater and gas (Melton et al., 2016). Future research might reveal the production of the amino acids L-serine and C-cysteine by these bacteria, via cysteine synthase A, as shown in the schematic overview of the pathways for sulfite oxidation and reduction and L-Serine and L-Cysteïne production in D. alkaliphilus AHT2 (Figure 1). Combined with the ongoing search for enzymes that are stable under extreme conditions (Van den Burg, 2003), this might even yield possibilities for implication of these bacteria for biotechnology.
SO32- + electron donor <--> H2S + oxidized donor + 3 H2O
* Δ G0 = -222 (KJ/mol SO32-) at 25˚C
4 SO32- + H+ à 3 SO42- + HS- *
To get a better understanding of how these bacteria contribute to the element cycling, by influencing the sulfur cycle, the processes they perform must be identified. Before identifiing these processes, the substrate preferences of the bacteria were investigated by growing them on different carbon sources and with or without yeast extract. Subsequently, the bacteria were grown with or without an electron donor. Growing the bacteria without an electron donor will force them to perform disproportionation.
Expectations are that bacteria growing on the preferred substrate and without an electron donor will start the process of disproportionation. When disproportionation takes place, this means that as well as possessing a genetic pathway encoding for the reduction of sulfite, a pathway responsible for sulfite oxidation needs to be present. Aside from investigating wether the bacterium D. alkaliphilus is capable of disproportionation, this study therefore subsequently aims to make an attempt to start mapping the genetic profile that is responsible for the execution of this process.
Figure 1. Schematic overview of sulfite oxidation and reduction and L-Serine and L-Cysteïne pathways
Material and Methods
Bacteria
The studied bacterium was Desulfurivibrio alkaliphilus strain AHT2, a haloalkaliphilic, Gram-negative deltaproteobacterium. The cells are shaped like curved rods, are non-motile and do not form spores. D. alkaliphilus grows in pH range from 8.5 to 10.3 with an optimum at pH 9.5 and is capable of tolerating sodium carbonate concentrations varying from 0.2 to 2.5 M total Na+ (Sorokin et al., 2008). D. alkaliphilus, from the order of the Desulfovibrionales, belongs to
the family of the Desulfobulbaceae. The genome of this species has been sequenced and characterized, containing a 3.10 Mbp chromosome (Melton et al., 2016).
The samples were obtained from mixed sediments, that originated from 8 different alkaline and hypersaline soda lakes in the Wadi El Natrun (Arabic for natron valley) region in the Libyan desert in Egypt, indicated in Figure 2 (Sorokin et al., 2008).
Cultivation conditions
All cultures were grown in 60 mL serum bottles in 0.6 M Na+ medium (see protocol in Appendix
A). All serum bottles were first cleaned with a 1 M HCl solution and rinsed three times with demi-water and three times with Milli-Q water. After cleaning they were capped with aluminum foil and autoclaved at 120 °C. Each serum bottle was filled under sterile conditions, in the laminar flow cabinet, with 50 mL sterile culture medium, the respective carbon sources and the electron donor. The bottles were closed with butyl rubber stoppers, capped with aluminum crimp caps and made anoxic (see protocol in Appendix B). Two mL of bacterial suspension was added through the stopper, under sterile conditions in the laminar flow cabinet using a 2 mL syringe (BD Plastipak) and needle (Terumo: Neolus 0.55 x 25 mm). All bottles were incubated at 30 °C while shaking.
Figure 2-A. Location of origin of the samples D. alkaliphilus B. Example of one of the soda lakes
To test what the preferred substrate was for the bacteria, they were grown on different carbon sources, thiosulfate or sulfite respectively, and with or without the addition of yeast extract, leading to four different compositions (Table 1). Bacteria were known to grow on elemental sulfur and thiosulfate and it was proposed that sulfite is a key intermediate in the disproportionation of elemental sulfur (Poser et al., 2013). Since a number of sulfur reducing bacteria are able to grow on, and perform disproportionation of thiosulfate (Finster, Liesack & Thamdrup, 1998; Habicht, Canfield & Rethmeier, 1998), this sulfur compound was incorporated in the substrate preference test as a check to see if the bacteria would grow on thiosulfate if they would not grow on sulfite.
To test if the bacteria were able to perform disproportionation, they were grown either with or without an electron donor. To determine if sulfite was disproportionated and growth could not be explained by another cause, a number of controls were added, including a chemical control to verify sulfide would remain at the start concentration and no decline due to either degradation or adherence to any of the materials would take place. An overview of the different cultivation conditions can be found in Table 2.
Table 1. Overview of carbon and sulfur source sources used in the experiment
Treatments Content Concentration Replicates
1 Thiosulfate 10 mM 2 Acetate 5 mM Pyruvate 10 mM Yeast extract * 2 Thiosulfate 10mM 2 Acetate 5 mM Pyruvate 10 mM 3 Sulfite 10 mM 2 Acetate 5 mM Pyruvate 10 mM Yeast extract * 4 Sulfite 10 mM 2 Acetate 5 mM Pyruvate 10 mM
Comparative sequence analysis
In order to identify the genes responsible for reducing and oxidizing sulfite, the KEGG (Kyoto Encyclopedia of Genes and Genomes) pathway specific for sulfur metabolism (Appendix C), was consulted. Possible candidates were selected based on their position in the KEGG pathway, between sulfite and the product of reduction (sulfide) or the product of oxidation of sulfite (sulfate). Genes that were not annotated in D. alkaliphilus, but which were possible candidates for sulfite oxidation were also selected.
The National Center for Biotechnology Information (NCBI) GenBank was used to acquire the sequences from the candidate genes. This search was conducted by entering the name of the gene in the NCBI-search tool and, after selecting for genes, downloading the sequences of these genes from the online library.
The sequences were checked for possible alignments with the complete sequence of the genome of D. alkaliphilus AHT2 using NCBI’s Basic Local Alignment Search Tool (BLAST). Best matches on query cover (> 60%) and identity (> 40%) were selected. The resulting genes were entered in Joint Genome Institute Integrated Microbial Genomes (JGI-IMG) and a search was conducted for the top 200 homologous genes. These 200 genes were selected to create a phylogenetic tree (bootstrap 500) using Molecular Evolution Genetic Analysis (MEGA). Based on their position in the homology tree, the nearest branches having the highest homology, the most closely related organisms were selected. An alignment of the homologous genes was made using MEGA. In this alignment a search for conserved regions was conducted (more than 12 consecutive matching base pairs). Genes that met these criteria were included.
Table 2. Different experimental cultivation conditions
Experimental group Contents Concentration Replicates
Treatment Sulfite 10 mM 3 Acetate 5 mM Pyruvate 10 mM Bacteria * Control Sulfite 10 mM 3 Acetate 5 mM Pyruvate 10 mM
Sulfite control Sulfite 10 mM 3 Acetate 5 mM
Pyruvate 10 mM Sulphide 0.5 mM
Disproportionation Sulfite 10 mM 3 Bacteria *
Sterile control Sulfite 10 mM 3 Chemical control Sulfite 10 mM 3
Sulfide 0.5 mM
Primer design
Primers were designed using the web-based software from Primer3Plus (https://primer3plus.com/), an online primer design tool. The sequence of the targeted gene was entered in the tool and parameters were set to meet the following conditions:
- The 3’ prime is most specific site; avoid a T at the 3’ prime.
- The amplification region is not bigger than 200 (250 for qPCR) nucleotides (optimum lies between 60 and 180 nucleotides).
- The CG-content is around 50/60 percent. - The melting temperature is between 60-63 ˚C.
The primer should be specific to one site; no secondary structure should be able to form.
- No more than 3 of the same nucleotides next to each other. For each targeted gene two sets of primers were designed.
Experimental analysis
Sampling
The sampling of the experimental setups was conducted under sterile conditions in the laminar flow cabinet. Two mL samples were taken from the bottles using a syringe and needle and stored for further analysis.
Determination of growth by optical density measurements and sulfide analysis
In order to measure growth and determine the onset of the exponential growth phase, the optical density at 600 nm (OD600) was measured. In addition, the amount of sulfide was
determined using a photometric method. To measure sulfide, 0.5 mL sample was added to 2.0 mL of a 10% (w/v) zinc acetate solution to fix the, highly unstable, sulfide. Using two different reagents the fixed sample was colored blue. Photometric measurements were performed at 675 nm (the full protocol for OD measurements can be found in Appendix D). A calibration curve was made using known concentrations of sulfide (in the form of sodium sulfide, Na2S),
a curve was fitted true the data points (y = 577,7x – 37,031) leading to R2 = 0,9922, the
calibration curve can be found in Appendix E. The exponential formula was used to fit the data from further hydrogen sulfide measurements. For both OD600 and OD765 a Versamax tunable
microplate reader was used, which was connected to a PC using VersaMax Pro 4.8 software. This data was used to determine when the culture would reach the exponential growth phase.
Harvesting
Bacteria were harvested when they were in the exponential growth phase. Using a 10 mL stripette, samples were transferred to two sterile 50 mL Greiner tubes, containing 30 and 20 mL of sample per tube respectively, and were spun down twice, using a centrifuge, at 10.000 rpm for 15 minutes at 4 ˚C. The supernatant was removed, the pellet resuspended and transferred to a 2 mL Eppendorf tube. These were spun down for 10 minutes at 10.000 rpm at 4 ˚C. Supernatant was removed and the pellet, in the Eppendorf tube, was flash frozen in liquid nitrogen and stored at -80 ˚C for DNA/RNA extraction.
DNA extraction
DNA was extracted using the MoBio Powersoil DNA extraction kit (protocol in Appendix F). Samples were run on a 1% (w/v) agarose gel for 40 minutes at 100V (4 μL sample & 1 μL loading dye, a 1 kB Generuler DNA Ladder mix (ThermoFisher) was used as a marker).
RNA extraction
RNA was extracted using the MoBio Powersoil RNA extraction kit (protocol in Appendix G). Samples were run on a 1% (w/v) agarose gel for 40 minutes at a 100 Volt. The extracted RNA was cleaned using the Life Technologies RNA-cleaning kit (see protocol in Appendix H).
Qubit
The concentration of the extracted DNA and RNA was measured using a Qubit fluorometer 2.0, which uses fluorescent dyes and a photospectrometer to measure light absorbance at 260 nm, determining the concentration of either DNA or RNA based on the amount of light that is absorbed.
Polymerase chain reaction
A PCR was conducted in a BioRad T100 Thermal Cycler (protocol in Appendix I), using the GM3F and GM4R primers that amplify the nearly complete 16S rRNA gene. The resulting PCR product was run on a 1% (w/v) agarose gel to see if the RNA-cleanup (Life Technologies) was successful. Cleaned RNA was converted to cDNA using BioRAD iScript™ cDNA synthesis Kit (see protocol in Appendix J).
Primer testing
To test if the primers worked and to determine the optimal annealing temperature, making sure no non-specific bands form, a gradient PCR was performed with temperatures ranging from 40 to 60 ˚C (see Table 4).
RT-qPCR
A real time quantitative PCR was used to determine gene expression. By monitoring the amplification of the targeted DNA during the polymerase chain reaction it shows real time amplification opposed to measuring only at the beginning and the end of the reaction, as is done in a regular PCR.
Expression of the target genes, for both sulfite oxidation and reduction, was measured using CYBR-green fluorescent DNA probes and measuring the fluorescence in real time. An additional plate was used to measure the expression of 16S rRNA genes. The expression of the genes, shown in Table 5, was monitored. The RT-qPCR program for 10 µL samples can be found in Appendix K.
Table 4. Temperatures in gradient PCR
Step 1 2 3 4 5 6 7 8
Temperature 60.0 58.4 55.8 52.2 47.6 43.7 41.3 40.0
Table 5. The genes monitored in the qPCR
Gene Description Function
DsrAB Sulfite reductase, dissimilatory-type alpha subunit sulfite reductor AprAB Adenylylsulfate reductase, alpha subunit sulfite oxidator
The results were used to determine the efficiency of the reaction using the Livak method (Livak & Schmittgen, 2001).
Statistics
De Winter (2013) showed that even when sample sizes are very small the student’s T-test can be used. Therefore, an independent sample T-test was used to determine if there were significant differences between growing the bacteria with or without yeast-extract. To test for normality and equality of variance respectively the Kolmogorov-Smirnov and Levene’s test were used. To test for significant differences in gene expression in the RT-qPCR results also the independent sample T-test was used. To test for normality the Kolmogorov-Smirnov test was carried out, equality of variance was tested with Levene’s test. All statistical analysis were conducted with IBM® SPSS® version 24.
Results & Discussion
Substrate preference
In order to ensure that the bacterial cultures would grow and to optimize growth conditions, a substrate preference test was performed. Knowing from previous literature (Bak & Pfennig, 1987; Friedhelm, Bak & Cypionka, 1987; Finster et al., 1998; Finster, 2008 & Janssen, Schuhmann, Bak & Liesack, 1996) that this type of bacteria could grow and disproportionate on thiosulfate, this sulfur compound was incorporated in the test next to sulfite. Thereby creating a control to rule out that the bacterial strain was not growing whatsoever. To determine which substrate is best to grow batch cultures, four different mixtures were selected. All batches were grown in duplo, Figure 3 shows the optical density values (OD600),
for the estimation of the bacterial concentration of D. alkaliphilus AHT2 over time with the different substrates. The results from these OD600 measurements show that D. alkaliphilus
strain AHT2 grows on both thiosulfate and sulfite. This matches the literature (Finster, 2008; Pikuta et al., 2003; Sorokin et al., 2011; Sorokin et al., 2008) which shows that this type of bacteria can use sulfite for their metabolic processes.
For the cultures with thiosulfate and sulfite, there were no significant differences between substrate preference with and without addition of yeast extract (t(2)=0.261 p=0.833 and t(2)=-2.056 p=0.177 respectively). 0,01 0,1 0 10 20 30 OD 600 nm Time (days)
A
0,01 0,1 0 10 20 30 OD 600 nm Time (days)D
0,01 0,1 0 10 20 30 OD 600 nm Time (days)B
0,01 0,1 0 10 20 30 OD 600 nm Time (days)C
Figure 3. Growth of D. alkaliphilus on different substrates, the blue and orange lines represent the different replicates, Y-axes are on a logarithmic scale (base 10). A. Thiosulfate, Acetate, Pyruvate & Yeast B. Thiosulfate, Acetate & Pyruvate C. Sulfite, Acetate, Pyruvate & Yeast and D. Sulfite, Acetate & Pyruvate.
Growth
Desufurivibrio alkaliphilus AHT2 was grown in batch culture in order to determine when they
would reach their exponential growth phase. Hydrogen sulfide was measured from the beginning of the inoculation by color-based spectrophotometric analysis. Figure 4 shows the hydrogen sulfide concentration for the cultures grown with different substrates over time.
Additionally, yeast extract was added to half of the cultures in order to see if it would benefit growth, since it is suggested that yeast extract could function as a nitrogen source (Hakobyan, Gabrielyan & Trchounian, 2012). From Figure 4 we can see that there was no difference in sulfide production for cultures with thiosulfate and sulfite, and with or without addition of yeast extract (t(2)=1.005 p=0.427 for thiosulfate and t(2)=-0.210 p=0.857 for sulfite). Therefore, we can conclude that no beneficial growth effects are visible when yeast extract was added. Consequently, the hypothesis that yeast extract would be beneficial for growth was rejected. This observation, combined with the lack of an accurate description of the composition of yeast extract, led to the decision not to use yeast extract in the culture medium. This matches with cultivation conditions described in other literature, using D.
alkaliphilus as their test organism, where there is also no mentioning of using yeast extract in
the culture medium to supply extra nitrogen (Melton et al., 2016; Poser et al., 2013; Sorokin
et al., 2008; Thorup, Schramm, Findlay, Finster & Schreiber, 2017).
0 300 600 900 1200 1500 1800 0 10 20 30 [H S -] (m M ) Time (days)
A
0 300 600 900 1200 1500 1800 0 10 20 30 [H S -] (n M ) Time (days)B
0 300 600 900 1200 1500 1800 0 10 20 30 [H S -] (m M ) Time (days)C
0 300 600 900 1200 1500 1800 0 10 20 30 [H S -] (m M ) Time (days)D
Figure 4. Sulfide concentration in mM for the different substrates, the blue and orange lines represent the different replicates. A. Thiosulfate, Acetate, Pyruvate & Yeast B. Thiosulfate, Acetate & Pyruvate C. Sulfite, Acetate, Pyruvate & Yeast and D. Sulfite, Acetate & Pyruvate.
From the data shown in Figure 4 it can be concluded that the exponential growth phase is reached between day 12 and 21. However, the time points at which growth was measured were too far apart to give an exact time indication. Other studies incorporating growth of D.
alkaliphilus measure growth every seven to eight hours (Thorup et al., 2017) where in this
study it was measured approximately every three days. However, since the exponential growth phase lasts several days, the estimation of the maximum of the exponential growth phase was deemed adequate.
Both the control and sterile control groups showed no production of hydrogen sulfide. In the sulfide control and chemical control groups there was a very slight decrease from the initially added amount of sulfide to reduce the culture medium.
To test if the AHT2 strain of the bacteria D. alkaliphilus was also capable of growing on sulfite without an electron donor present, a substrate containing only growth medium, sulfite and bacterial solution was tested. A series of controls (a chemical-, sterile-, sulfide and a no-bacteria control) was added to the setup to make sure that disproportionation of sulfite was indeed taking place and could not be explained by a different cause. The AHT2 strain indeed seemed capable of disproportionating sulfite, since in the cultivation condition without an electron donor it still managed to grow and produce sulfide. This matches earlier studies conducted with Desulfurivibrio sp. strain AMeS2 (the only other known isolated representative from the Desulfurivibrio genus), were it showed the ability to perform disproportionation of sulfite in the absence of an electron donor (Poser et al., 2013).
The proof that disproportionation of sulfite was possible in the bacteria D. alkaliphilus, led to the assumption that besides having genes coding for sulfite reduction also genes responsible for the oxidation of sulfite should be present.
Comparative sequence analysis
To identify the genes responsible for or involved in the disproportionation of sulfite, possible candidate genes were located in the KEGG pathway specific for D. alkaliphilus. The genes that were picked, based on their position in the D. alkaliphilus specific KEGG pathway (Appendix L), were a dissimilatory-type alpha and beta subunit (DsrAB), which is a sulfite reductase and adenylyl sulfate reductase, alpha and beta subunit (AprAB), a sulfite oxidizer. Sequences were obtained from the NCBI’s Genbank (Appendix M).
Figure 5 and Figure 6 show phylogenetic trees made of sequences of DrsAB and AprAB indicating the position to other species.
The method of using the KEGG pathway to identify the possible candidate genes responsible for or involved in the disproportionation of sulfite in D. alkaliphilus, genes and using these candidate gene sequences to find homologous genes often yields success. However, this method limits the possibility of finding genes involved in sulfite disproportionation that are not homologous to the candidate genes or even fall outside the scope of the candidate genes, when they code for a so far unknown pathway. To gain a more complete overview of genes with possible involvement of sulfite disproportionation, other methods in functional genomics might therefore yield more results. Yet, techniques like genetic interaction mapping or RNA sequencing, are usually highly time consuming and/or very costly. Given the exploratory nature of this study, the used approach was considered sufficient at this stage of the research.
DNA and RNA extraction
To test if the genes were present in the DNA of the test organism D. alkaliphilus, first DNA and RNA was extracted. For both the DNA and RNA isolation, MoBio powersoil® kits were used.
Figure 5.Partial phylogenetic tree of DsrAB sequences compiled using MEGA, the scale bar
indicates the percentage sequence difference. The numbers on the branches indicate the bootstrap values out of 500 iterations.
Figure 6.Partial phylogenetic tree of AprAB sequences compiled using MEGA, the scale bar
indicates the percentage sequence difference. The numbers on the branches indicate the bootstrap values out of 500 iterations.
Other studies isolating DNA and/or RNA from soda lake samples (Duckworth, Grant, Jones & Van Steenbergen, 1996; Melton et al., 2016; Rees, Grant, Jones & Heaphy, 2004; Sorokin et
al., 2001) used comparable methods. In order to determine if the isolation of DNA was
successful, the extracted product was run on an agarose gel. Figure 7 clearly shows a band on the gel, for both replicas (duplo) of the batch cultures, grown on a substrate of sulfite, pyruvate and acetate, around the 1 kB mark of the added gene ruler mix.
To check if the RNA extraction was successful the product was submitted to a gel- electrophoresis (Figure A). After the extraction an RNA clean-up was performed (Figure 8-B).
Qubit
Using the Qubit the quantity of both the DNA- and the RNA-samples was determined. Table 6 and Table 7 show the quantity of the DNA- and RNA-samples respectively.
Figure 8-A. Extracted RNA from D. alkaliphilus AHT2 grown on sulfite, acetate & pyruvate. ‘M’ marks the gene ruler. 1-3 represent the RNA-samples.
Figure 8-B. Extracted RNA from D. alkaliphilus AHT2 grown on sulfite, acetate & pyruvate, after RNA clean-up. ‘M’ marks the gene ruler. 1-3 represent the RNA-samples.
Figure 7.Extracted D. alkaliphilus DNA. ‘M’ marks the gene ruler. All bacteria were grown on a substrate of sulfite pyruvate and acetate. 1-2 represent DNA from replica 1 (measured in duplo) 3-4 represent DNA from replica 2.
M 1 2 3 4 M 250 bp 1000 bp 5000 bp 10000 bp 3000 bp M 1 2 3 250 bp 10000 bp 1000 bp 5000 bp 3000 bp M 1 2 3 250 bp 10000 bp 1000 bp 5000 bp 3000 bp
Figure 7 and Figure 8, showing the isolated DNA and RNA, showed clear bands on the gel electrophoresis and both isolates. Analysis for quantity using qubit measurements confirmed that both DNA and RNA isolates had quantities within the acceptable range for successful PCR and RT-qPCR.
Primer design and testing
In order to be able to perform PCR- and RT-qPCR-analysis, primers were designed for both DsrAB and AprAB. The sequences from the candidate genes were entered in the Primer3Plus online primer design software and parameters were set to match the desired outcome (Abd-elsalam, 2003; Rychlik, 1995), resulting in multiple primer-set sequences for both genes. For both DsrAB and AprAB two sets of primers (forward and reverse) were selected, based on the best match with the criteria previously described. The primer sequences were blasted against the genomic DNA of D. alkaliphilus to ensure a single sequence was targeted. Based on the outcome of the BLAST, two primer-sets per target sequence were selected for a gradient-PCR, to see if the primers worked and to test for their optimal annealing temperature (photo of the product on the gel in Figure 9). Annealing temperatures were found to be optimal at 60 ˚C for both DsrAB and AprAB. An overview of the selected primers, including their sequences and properties, can be found in Appendix N.
Limitations in primer design can most often be found in setting parameters either to strict or not strict enough. This results in lower efficiency, in other words, how close to the theoretical optimum a primer pair is able to amplify a product, either because of extra unrelated amplification when parameters are set to tolerant or because of poor annealing when set to strict (Dieffenbach, Lowe & Dveksler, 1993). Nonetheless, with the parameter settings used in designing the primers used in this study, described in the material and methods section, no limitations in primer functionality were found. Both the PCR and RT-qPCR analysis yielded enough product and no byproduct or primer dimers were found. In previous research, conducted on both the DsrAB and AprAB gene, primers were designed as well (Giloteaux, Goñi-Urriza & Duran, 2010; Karkhoff-Schweizer, Huber & Voordouw, 1995b; Meyer & Kuevert, 2007). These studies used comparable parameter settings in the design of the primers and in all cases, primers were designed succesfull for use in a PCR. There was however a difference in the methods of designing the primers, using either software tools or designing the primers manually. Using software tools to design primers offers the ability to work faster and
Table 6. Quantity of the DNA-samples measured with the Qubit
Sample Substrate Quantity (µg*mL-1)
1 acetate, pyruvate & sulfite 1.01 2 acetate, pyruvate & sulfite 66.4 3 acetate, pyruvate & sulfite 47.2 4 acetate, pyruvate & sulfite 39.3
Table 7. Quantity of the RNA-samples measured with the Qubit
Sample Substrate Quantity (mg*mL-1)
1 acetate, pyruvate & sulfite 22.0 2 acetate, pyruvate & sulfite 24.9 3 acetate, pyruvate & sulfite 24.5
incorporate different parameter settings at once. It does, however, not effect the quality and efficiency of the primer itself (Abd-elsalam, 2003; Ye et al., 2012).
Analyzing the photos from the gel electrophoresis, containing the product from the PCRs, is it clear that all the primers used in the PCR reaction yielded product. This proves that D.
alkaliphilus possesses at least part of the genetic code associated with sulfite oxidation,
namely the gene AprAB.
RT-qPCR
Thorup et al. (2017) state that: “Key genes of known sulfide oxidation pathways are absent from the genome of D. alkaliphilus.”. Giving the fact that in the sulfur oxidation pathway sulfide is only one oxidation step away from sulfite, it is surprising that the genome of D.
alkaliphilus contains genes known to code for the oxidation of sulfite. To measure whether
these genes showed an increased level of transcription, a clear indication of involvement of the gene in the process, when D. alkaliphilus was forced to perform disproportionation of sulfite, an RT-qPCR analysis was performed. Since disproportionation involves simultaneous reduction and oxidation of the compound of interest, transcription of both a gene involved in oxidation and a gene involved in the reduction of sulfite, AprAB and DsrAB respectively, were measured. To quantify the expression of the targeted genes an RT-qPCR analysis was performed using 16S RNA as a reference. The melt curve summary shows values between 87 and 88.5, meaning only one product was produced and there was no off-target amplification. Figure 10-A & 10-B show the mean of the starting quantity values, with 16S RNA as reference for the AprAB and the DsrAB gene respectively.
Figure 9. Product of the gradient-PCR. ‘M’ marks the gene rulers. Numbers 2-8 mark the DsrAB gene with the first primerset, 9-16 mark the DsrAB gene with the second primerset, 17-24 mark the AprAB gene with the primerset and 25-32 mark the AprAB gene with the second primerset. The consecutive temperatures at which the primersets were tested were: 60.0, 58.4, 55.8, 52.2, 47.6, 43.7, 41.3 and 40.0 ˚C.
M 17 18 19 20 21 22 23 24 M 1 2 3 4 5 6 7 8 9 10 111213 141516 M 25 26 27 28 29 30 31 32 M 10000 bp 10000 bp 10000 bp 10000 bp 5000 bp 5000 bp 5000 bp 5000 bp 1000 bp 1000 bp 1000 bp 1000 bp 250 bp 250 bp 250 bp 250 bp
Both the AprAB and the DsrAB gene do not show a significant increase or decrease in transcription when the bacteria are forced to perform disproportionation (t(2)=-0.988 p=0.427) for AprAB and (t(2)=-0.993 p=0.425) for DsrAB respectively. This can be explained by different causes. First of all it might be that, even though these genes code for the oxidation of sulfite, they are not involved in the process of disproportionation. This then would lead to the assumption that other genes, also coding for the oxidation of sulfite, are present in the
0,00010 0,00100 0,01000 0,10000 1,00000 10,00000
APR_NTC APR_1 APR_2 APR_3 APR_4 APR_5 APR_6
St ar tin g qu an tit y (g en e co pi es re la tiv e to 1 6S )
Target gene and sample number
0,00001 0,00010 0,00100 0,01000 0,10000 1,00000 10,00000 DSR_NTC DSR_1 DSR_2 DSR_3 DSR_4 DSR_5 DSR_6 St ar tin g qu an tit y (g en e co pi es re la tiv e to 1 6S )
Target gene and sample number
Figure 10-A.Mean of the Starting Quantity calculated using 16S RNA as reference, for the AprAB (Adenylylsulfate reductase)gene. APR_NTC is the AprAB gene No Template Control. Numbers 1-3 are three different replicates of D. alkaliphilus culture grown with an electron donor and number 4-6 are three different replicates grown without an electron donor, thereby forcing the latter to perform disproportionation.
Figure 10-B.Mean of the Starting Quantity calculated using 16S RNA as reference, for the DsrAB (Dissimilatory sulfite reductase) gene. DSR_NTC is the DsrAB gene No Template Control. Numbers 1-3 are three different replicates of D. alkaliphilus culture grown with an electron donor and number 4-6 are three different replicates grown without an electron donor, thereby forcing the latter to perform disproportionation.
genome of D. alkaliphilus and are used in this process. Another explanation is that sample sizes were simply too small to detect a significant difference. De Winter (2013) showed that when a T-test is used with small sample sizes and an unequal variance, the effect size must be large in order to detect a significant difference.
Thorup et al. (2017) suggest that a reversal of the DsrAB gene was used by D. alkaliphilus in order to create an oxidation pathway for sulfide, which, when coupled with dissimilatory reduction of nitrate and nitrite to ammonium, can be used for chemo-lithotrophic growth. When this proves correct, it could reveal that homologous genes from closely related bacteria, coding for oxidation, might not actually play a role in the disproportionation of sulfite, but that instead this process is mediated by the reversal of genes involved in reduction. Both Meyer & Kuevert (2007) and Mussmann et al. (2005) indicate a common evolutionary development of AprAB and DsrAB between sulfate oxidizing bacteria and sulfate reducing bacteria through both horizontal and vertical gene transfer. Further investigation in the large homology of both genes between these two groups might reveal clues for the mechanisms behind the previous described reversal of these genes and thereby create more insight in the genetic background of disproportionation. However, since both the AprAB and the DsrAB gene showed no significant increase in expression when D. alkaliphilus was forced to perform sulfite disproportionation, the genetic background for this process yet remains unknown.
Conclusion
This study aimed to gain more insight in the microbial mediated element cycling in soda lakes, with a specific focus on the disproportionation of sulfite by the bacteria D. alkaliphilus. It was hypothesized that, with a suitable substrate, D. alkaliphilus could use disproportionation of sulfite for chemolithotrophic growth.
In this study we found that D. alkaliphilus is indeed capable of performing disproportionation of sulfite and can couple that to chemolithotrophic growth. Its ability to perform inorganic sulfur disproportionation reactions in laboratory cultures indicates that the necessary genetic pathways for both sulfite reduction and oxidation are present in the genome of this organism. These pathways are so far poorly mapped, therefore further investigation of the D. alkaliphilus AHT2 genomic background may reveal insights in which genes are responsible for this metabolic property. In addition, a more in-depth genome sequence analysis might provide a better understanding of autotrophic carbon metabolism in halo alkaline environments. Overviewing the current study there are a number of specific subjects to highlight. The main conclusion is that the bacteria D. alkaliphilus is capable of disporportionation of sulfite and is thus capable of using sulfite to power its metabolic needs and fuel chemolithotrophic growth, in the absence of an electron donor. Even though homologous genes, involved in sulfite oxidation were shown to be present in the genome of D. alkaliphilus, they did not show a significant increase in expression when forcing the bacteria to perform disproportionation. This leads to the hypothesis that a different genetic pathway is responsible for this process. In order to uncover and map these pathways there are a number of recommendations for future research to consider.
Since in the current study only two different genes were measured in the RT-qPCR, the first step would be to scale up the experiment and use a larger number of possible involved genes. Two of those could be SOR (Sulfur oxygenase/ reductase) and Rubredoxin, since non published data from this research showed that they are present in the genome of D. alkaliphilus. Scaling up the number of replicates within the experiment would lead to more statistical power. As shown by de Winter (2013), a large effect size is needed to show significant differences when variances are unequal and the number of replicates is small. Furthermore, the study of Thorup
et al. (2017), showed that a reversal of the Dsr gene was used in a sulfate oxidation pathway.
Reversal of genes, like Dsr in sulfide disproportionation, might be a more common trade in this clade of bacteria and should therefore be investigated further. Other genes thought to only be responsible for either oxidation or reduction might serve a dual function. A better understanding of which genes possess this ability and insight in how they are able to reverse their working mechanism might lead to a broader understanding of genetic regulation of microbial mediated disproportionation of sulfur compounds. To gain a larger understanding of the role microbes play in the element cycling of soda lakes, a first step could be to compare different types of bacteria involved in tantamount processes, in order to check for homologous genetic pathways, coding for shared properties. This could lead back to understanding the ancestral background of certain pathways and help to understand current diversity. A next step would then be to connect different processes to form (part of) the element cycle and map which parts of these processes are mediated by microbes and what the responsible mechanisms are.
Acknowledgements
First and foremost my gratitude goes out to dr. Emily Melton and prof. dr. Gerard Muijzer (whom both are funded by the ERC), for all their work and support during the lab work and during the process of writing my thesis, and dr. Michiel Kraak who fulfills the role of mentor during my research master. Furthermore, I would like to thank Cherel Balkema for her help in the lab and with the RT-qPCR analysis, the MSE (Microbial Systems Ecology) research group, part of IBED (Institute for Biodiversity and Ecosystem Dynamics), for their valuable contribution during presentations and team meetings and my examiner prof. dr. Lucas J. Stal for taking the time to review this thesis.
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Appendices
Appendix A
Protocol 0.6 M Na+ medium
Substance Amount Unit
Na2CO3 17.5 g/L NaHCO3 13.9 g/L NaCl 6.1 g/L K2HPO4 1.0 g/L Vitamin solution 1.0 mL/L Trace minerals* 1.0 mL/L SeW 1.0 mL/L NH4Cl 4.0 mL/L MgCl2• 6H20 1.0 mL/L
* (Pfennig & Lippert, 1966)
EDTA (Trilon B) 5.0 g/L FeSO4• 7H2O 2.2 g/L ZnSO4• 7H2O 0.10 g/L MnCl2 • 4H2O 0.03 g/L H3BO3 0.03 g/L CoCl2• 6H2O 0.20 g/L CuCl2• 2H2O 0.03 g/L NiCl2• 6H2O 0.03 g/L Na2MoO4• 2H2O 0.03 g/L
Appendix B
Anoxic environment serum bottles.
All steps take place in the laminar flow cabinet.
Attach 0.2-micrometer filters (VWR international, sterile syringe filter: w/0.2 µm Polyethersulfone membrane. Art. no. 514-0073) to tubes of both nitrogen-tap and vacuum pump (Millipore model no. WP6122050; 230V, 50Hz, 1.7A). Attach a long needle (B Braun; Sterican® 0.80 x 120mm) at the filter-end of the N2 tube, and a shorter needle (Terumo; Neolus 0.55 x 25mm) for the vacuum pump. First open green nitrogen-tap, then open the main nitrogen tap until the pressure is 15 psi.
Execute per bottle:
1. Put alcohol on cap of serum bottle
2. Pierce cap with nitrogen needle, into the fluid 3. Wait until bubbling stops
4. Pierce cap with vacuum needle (bubbling should recommence) 5. Pull the long needle up to gas phase and wait 30 seconds
6. Close the cramp on the nitrogen tube, close the green nitrogen tap 7. Turn vacuum pump on
8. Beat serum bottle until gas escapes from aqueous phase, wait for thirty seconds 9. Turn green nitrogen tap on, loosen cramp on the tube
10. Turn vacuum pump off
11. Let nitrogen flow in for thirty seconds
12. Turn vacuum pump on and close cramp on nitrogen tube, close the green nitrogen tap 13. Wait thirty seconds
14. Turn green nitrogen tap on, loosen the cramp on the tube 15. Turn vacuum pump off
16. Let nitrogen flow in for thirty seconds
17. Turn vacuum pump on and close cramp on nitrogen tube, close green nitrogen tap 18. Wait thirty seconds
19. Turn green nitrogen tap on, loosen cramp on the tube
20. Pull (short) vacuum needle up into cap, wait a moment and completely pull out 21. Turn vacuum pump off, let nitrogen flow in for thirty seconds
22. Pull (long) nitrogen needle up into cap. Wait a moment and slowly pull out 23. Label bottle with ‘N2’ sticker
Appendix C
The sulfur KEGG pathway
Appendix D
Liquid microanalysis of sulfide (HS-)
1. Prepare a number of 15 mL Falcon tubes with 2.0 mL 10% ZnAc solution. 2. Prepare a dilution series to calibrate the assay.
3. Prepare the following solutions (under the fume hood!):
10% Zn acetate – after dissolving, add 0.1 mL (100%) acetic acid/L (prepare 1 L)
Sulfide reagent A: dissolve 2 g dimethylparafenyldiamine (oxalate salt) in 200 mL of DI
water, add 200 mL of 96% H2SO4, adjust volume to 1 L (with DI water) and store in a dark
bottle (fume hood, gloves, glasses).
Sulfide reagent B: Dissolve 10 g Fe(NH4)(SO4)2 • 12 H2O in 50 mL DI water, add 2 mL of
96% H2SO4 and adjust the volume to 100 mL with DI water.
All suspended matter must be removed from the sample since it is absorbing the formed methylene blue. Samples can be filtered or centrifuged, but filtration is better since centrifuge takes longer and sulfide is very unstable.
For every measurement you need 0.5 mL sample. If you measure in triplicate (recommended), then you need 1.5 mL samples in total.
1. Pipet 0.5 mL sample into 2.0 mL 10% Zn acetate solution
(Preloaded into a 15 mL Falcon tubes so that the final volume is 2.5 mL)
2. Agitate (shake) the tube immediately to allow the sulfide to react with the Zn so that it forms insoluble ZnS. (Now the sample is fixed and stable.)
3. Add 6.4 mL DI-water to the fixed sample (in the Falcon tube).
4. Add 1.0 mL of sulfide reagent A and 0.1 mL sulfide reagent B. Immediately close the tube and mix well.
5. Incubate for 15 minutes to allow the blue color to develop
Blue color indicates the presence of sulfide. The full color development depends on the presence of other sulfur compounds. In the presence of thiosulfate color development is retarded. However, in contrast to what is written in the literature, dr. D. Sorokin discovered that the results in presence and absence of thiosulfate are the same. Therefore, it is not necessary to separate thiosulfate from the sample beforehand. In order to remove thiosulfate, the ZnS must be centrifuged. The supernatant with thiosulfate can than be removed and the volume replaced with demi water.
6. Measure optical density 675 nm with whatever control you deem necessary as a blank. 7. To calculate the sulfide concentration, use a factor of 33 x OD675 x dilution factor μM;
i.e. if undiluted sample gives OD675 1 unit, it contains 33 μM sulfide
If the pH is <7 and there is a substantial gas phase, then a certain fraction of sulfide is present in the gas phase as H2S which also needs to be taken into account. It can be calculated from
the distribution formula after analyzing HS- in the liquid phase.
Preparing the stock solutions for the calibration curve.
In the fume hood!
Prepare anoxic water (or whatever other matrix you are using) by bringing it to boiling point and then purging it with N2 gas whilst cooling to room temperature.
Take some crystals from the NaS stock and wash them in a petri dish with some (anoxic) DI-water to remove oxidation products and impurities. Blot dry with some paper and weigh the correct amount.
When you prepare the solution, it should be colourless. Over time it will become green/bluish depending on the sulfide concentration. This is due to polysulfides (which form as a function of the sulfide concentration and the pH). Be aware that these polysulfides could act as a catalyst for the oxygenation of sulfide if you are using water as solvent, the catalytic activity is not as strong at alkaline pH values).
Appendix E
Calibration curve HS-
Appendix F
Protocol MoBio PowerSoil® DNA Isolation Kit
EXPERIENCED USER PROTOCOL
PowerSoil® DNA Isolation Kit Catalog No. 12888-50 & 12888-100
Please wear gloves at all times
1. To the PowerBead Tubes provided, add 0.25 grams of soil sample. 2. Gently vortex to mix.
3. Check Solution C1. If Solution C1 is precipitated, heat solution to 60 °C until dissolved before use.
4. Add 60 μL of Solution C1 and invert several times or vortex briefly.
5. Secure PowerBead Tubes horizontally using the MO BIO Vortex Adapter tube holder for the vortex (MO BIO Catalog# 13000-V1-24) or secure tubes horizontally on a flatbed vortex pad with tape. Vortex at maximum speed for 10 minutes.
Note: If you are using the 24-place vortex adapter for more than 12 preps, increase the vortex time by 5-10 min
6. Make sure the PowerBead Tubes rotate freely in your centrifuge without rubbing. Centrifuge tubes at 10,000 x g for 30 seconds at room temperature.
CAUTION: Be sure not to exceed 10,000 x g or tubes may break.
7. Transfer the supernatant to a clean 2 mL Collection Tube (provided).
Note: Expect between 400 to 500 µL of supernatant. Supernatant may still contain some soil particles.
8. Add 250 μL of Solution C2 and vortex for 5 seconds. Incubate at 4 °C for 5 minutes. 9. Centrifuge the tubes at room temperature for 1 minute at 10,000 x g.
10. Avoiding the pellet, transfer up to, but no more than, 600 μL of supernatant to a clean
2 mL Collection Tube (provided).
11. Add 200 μL of Solution C3 and vortex briefly. Incubate at 4 °C for 5 minutes. 12. Centrifuge the tubes at room temperature for 1 minute at 10,000 x g.
13. Avoiding the pellet, transfer up to, but no more than, 750 μL of supernatant into a clean 2 mL Collection Tube (provided).
14. Shake to mix Solution C4 before use. Add 1200 μL of Solution C4 to the supernatant and vortex for 5 seconds.
15. Load approximately 675 μL onto a Spin Filter and centrifuge at 10,000 x g for 1 minute at room temperature. Discard the flow through and add an additional 675 μL of supernatant to the Spin Filter and centrifuge at 10,000 x g for 1 minute at room temperature. Load the remaining supernatant onto the Spin Filter and centrifuge at 10,000 x g for 1 minute at room temperature.
16. Add 500 μL of Solution C5 and centrifuge at room temperature for 30 seconds at 10,000 x g.
17. Discard the flow through.
18. Centrifuge again at room temperature for 1 minute at 10,000 x g.
19. Carefully place spin filter in a clean 2 mL Collection Tube (provided). Avoid splashing any Solution C5 onto the Spin Filter.
20. Add 100 μL of Solution C6 to the center of the white filter membrane. Alternatively, sterile DNA-Free PCR Grade Water may be used for elution from the silica Spin Filter membrane at this step (MO BIO Catalog# 17000-10).
21. Centrifuge at room temperature for 30 seconds at 10,000 x g.
22. Discard the Spin Filter. The DNA in the tube is now ready for any downstream application. No further steps are required.
We recommend storing DNA frozen (-20 ° to -80 °C). Solution C6 contains no EDTA. To concentrate the DNA, see the Hints & Troubleshooting Guide.
Appendix G
Protocol MoBio PowerSoil® RNA Isolation Kit
EXPERIENCED USER PROTOCOL
Wear RNase-Free Gloves (1556) at all times and remove RNase from the work area using Lab Cleaner (12095) for RNase Removal. Both of these products are available from MO
BIO. Please see the “Products recommended for you” section at the end of this manual.
1. Add up to 2 g of soil to the 15 mL Bead Tube (provided).
Note: Please refer to Hints and Troubleshooting Guide for information
regarding the amount of soil to process.
2. Add 2.5 mL of Bead Solution to the Bead Tube followed by 0.25 mL of Solution SR1 and 0.8 mL of Solution SR2.
3. Add 3.5 mL of phenol: chloroform: isoamyl alcohol (pH 6.5 – 8.0, [User supplied]) to the bead tube, cap and vortex to mix until the biphasic layer disappears.
4. Place the Bead Tube on the Vortex Adapter (MO BIO Catalog # 13000-V1-15) and vortex at maximum speed for 15 minutes.
5. Remove the Bead Tube from the Vortex Adapter and centrifuge at 2500 x g for 10 minutes at room temperature.
6. Remove the Bead Tube from the centrifuge and carefully transfer the upper aqueous phase (avoiding the interphase and lower phenol layer) to a clean 15 mL Collection
Tube (provided). The thickness of the interphase will vary depending on the type of
soil used. Discard the phenol: chloroform: isoamyl alcohol in an approved waste receptacle.
Note: The biphasic layer will be thick and firm in soils high in organic matter
and may need to be pierced to remove the bottom phenol layer.
7. Add 1.5 mL of Solution SR3 to the aqueous phase and vortex to mix. Incubate at 4 °C for 10 minutes.
8. Centrifuge at 2500 x g for 10 minutes at room temperature. Transfer the
supernatant, without disturbing the pellet (if there is one), to a new 15 mL Collection
Tube (provided).
9. Add 5 mL of Solution SR4 to the Collection Tube containing the supernatant, invert or vortex to mix, and incubate at room temperature for 30 minutes.
Note: The previous protocol instructions were to incubate at -20 °C for 30
minutes. If you’ve used the -20 °C incubation before and know that your soil type yields good results at that temperature, you may continue to follow that protocol.
10. Centrifuge at 2500 x g for 30 minutes at room temperature.
11. Decant the supernatant and invert the 15 mL Collection Tube on a paper towel for 5 minutes.
Note: Depending on soil type, the pellet may be large and/or dark in color.
12. Shake Solution SR5 to mix. Add 1 mL of Solution SR5 to the 15 mL Collection Tube and resuspend the pellet completely by repeatedly pipetting or vortexing to disperse the pellet.
