bs_bs_banner
Environmental Microbiology Reports (2015) 7(3), 480–488
doi:10.1111/1758-2229.12276
Sphingomonas wittichii RW1 gene reporters interrogating the dibenzofuran metabolic network highlight conditions for early successful development in contaminated microcosms
Edith Coronado, Annabelle Valtat and Jan R. van der Meer* Department of Fundamental Microbiology, University of Lausanne, Lausanne 1015, Switzerland. Summary In order to better understand the fate and activity of bacteria introduced into contaminated material for the purpose of enhancing biodegradation rates, we constructed Sphingomonas wittichii RW1 variants with gene reporters interrogating dibenzofuran metabolic activity. Three potential promoters from the dibenzofuran metabolic network were selected and fused to the gene for enhanced green fluorescent protein (EGFP). The stability of the resulting genetic constructions in RW1 was examined, with plasmids based on the broad-host range vector pME6012 being the most reliable. One of the selected promoters, upstream of the gene Swit_4925 for a putative 2-hydroxy-2,4-pentadienoate hydratase, was inducible by growth on dibenzofuran. Sphingomonas wittichii RW1 equipped with the Swit_4925 promoter egfp fusion grew in a variety of non-sterile sandy microcosms contaminated with dibenzofuran and material from a former gasification site. The strain also grew in microcosms without added dibenzofuran but to a very limited extent, and EGFP expression indicated the formation of consistent small subpopulations of cells with an active inferred dibenzofuran metabolic network. Evidence was obtained for competition for dibenzofuran metabolites scavenged by resident bacteria in the gasification site material, which resulted in a more rapid decline of the RW1 population. Our results show the importance of low inoculation densities in order to observe the population development of the introduced bacteria and further illustrate that the limited
Received 8 October, 2014; accepted 31 January, 2015. *For correspondence. E-mail [email protected]; Tel. +41 (21) 692 5630; Fax +41 (21) 692 5605.
availability of unique carbon substrate may be the most important factor impinging growth. Introduction Inoculation of pure bacterial cultures or enrichments into complex microbial communities with the purpose of introducing or enhancing specific functionalities has enjoyed a long-standing interest (Pritchard, 1992; van Veen et al., 1997; Verschuere et al., 2000; El Fantroussi and Agathos, 2005; Thompson et al., 2005). In the area of environmental biotechnology, the inoculation into contaminated material of specific pre-enriched bacteria, which efficiently degrade one or more of the contaminants under laboratory conditions, has been frequently proposed as a means to accelerate biodegradation rates (El Fantroussi and Agathos, 2005; Thompson et al., 2005). However, even more than 20 years since the first strategic review (Pritchard, 1992), the process of bioaugmentation is still largely unpredictable because this is mostly trial and error-based (Tyagi et al., 2011). In order to improve our general understanding of the process, recent studies have attempted to more systematically measure and interpret global reactions of introduced bacteria for bioremediation into complex systems (Fida et al., 2012; 2013; Iino et al., 2012; Maphosa et al., 2012; Gunasekera et al., 2013; Moreno-Forero and van der Meer, 2015; Scheublin et al., 2014). This has been complemented with global transposon mutagenesis approaches in order to identify gene functions with fitness benefit for growth and survival in soil (Roggo et al., 2013; Fida et al., 2014). Whereas ‘big data’ approaches are powerful to characterize global cellular reactions and identify candidate genes essential for fitness or survival, they need complementation by methods that can measure the variability of individual cellular responses over time under local conditions. Gene reporter technology has been proposed for this (Leveau and Lindow, 2002; Tecon and van der Meer, 2006), and has been successfully applied to study, for example, local sugar availability for plant pathogens on plant leaves (Leveau and Lindow, 2001), or water (Herron
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd. This is an open access article under the terms of the Creative Commons Attribution-NonCommercial-NoDerivs License, which permits use and distribution in any medium, provided the original work is properly cited, the use is non-commercial and no modifications or adaptations are made.
Dibenzofuran detection by S. wittichii RW1 et al., 2010) and iron availability to bacteria in the phyllosphere (Joyner and Lindow, 2000). The goal of the underlying study was to obtain a more complete picture of the factors determining the success or failure of Sphingomonas wittichii RW1 to grow in non-sterile microcosms with contaminated material. Sphingomonas wittichii RW1 metabolizes dibenzofuran (DBF) and dibenzodioxins, which are frequently found in areas contaminated with polycyclic aromatic hydrocarbons (PAH) and after incineration processes (Safe, 1990). Strain RW1 has been studied in detail for its DBF and dioxin metabolic pathways (Wittich et al., 1992; Wilkes et al., 1996; Armengaud et al., 1998; Hartmann and Armengaud, 2014), and has also been applied to some success in bioaugmentation studies (Megharaj et al., 1997; Halden et al., 1999; Nam et al., 2005). Our hypothesis was that the growth of RW1 in nonsterile material would be mostly controlled by the presence of its unique carbon source (DBF) irrespective of the presence of other PAH-contaminated material. In order to study DBF availability for RW1 at the single-cell level, we generated RW1 reporter strains where egfp expression for enhanced green fluorescent protein (EGFP) would be under control of a promoter from the DBF metabolic network (Appendix S1, Fig. S1). As genetic engineering in strains like RW1 is non-trivial, we examined the genetic stability of the constructs and EGFP expression driven from the three regions in liquid cultures under a variety of conditions. The best performing RW1 bioreporter was then inoculated in sandy microcosms spiked or not with DBF or PAH-contaminated material to follow population growth and reporter induction among single cells. Our results indicate that RW1 can indeed grow and induce its DBF metabolic network in contaminated material, depending on the substrate availability. However, our results further suggest that resident bacteria from the contaminated material may compete with RW1 at the level of metabolites formed by RW1 from DBF.
Results and discussion Development of a S. wittichii RW1 bioreporter for DBF degradation Three putative RW1 promoter regions (Tables S1 and S2, Fig. S1) were fused to a promoterless egfp gene, introduced into strain RW1 and evaluated for their expression during growth on DBF, salicylate (SAL) or phenylalanine (PHE). The first is a region upstream of the gene Swit_4925, putatively involved in the transformation of 2-hydroxy-2,4-pentadienoate and 12-fold induced during growth on DBF (Coronado et al., 2012). The second was a putative promoter region upstream of Swit_5102, predicted to code for a gentisate dioxygenase and up to
481
17-fold induced (Coronado et al., 2012). As third promoter, we selected the region upstream of the gene Swit_4897 (dxnA1), which codes for dibenzofuran-4,4adioxygenase, catalysing the initial step in DBF degradation (Bunz and Cook, 1993; Armengaud et al., 1998). Promoter-egfp constructs based on vector pME6012 (Heeb et al., 2000) were stable in RW1 for eight generations in the absence of antibiotic selection compared with pPROBE- or mini-Tn5-based constructions (Fig. S2, Appendix S2). Strain RW1 (pME6012-P4925-egfp) displayed fivefold higher background-corrected fluorescence when growing on DBF than on SAL or PHE (Fig. 1A and D). In contrast, RW1 (pME6012-PdxnA1-egfp) produced a slightly higher EGFP fluorescence when grown with SAL compared with PHE or DBF (Fig. 1B and D). The EGFP expression from RW1 (pME6012-P4925-egfp) but less so from RW1 (pME6012-PdxnA1-egfp) was insensitive to changes in water potential, which might influence its behaviour in unsaturated soil (Fig. S3, Appendix S2). Contrary to our expectations from microarray data (17-fold higher expression of Swit_5102 in DBF-grown compared with PHE-grown cells), strain RW1 (pME6012-P5102-egfp) hardly produced any EGFP fluorescence above background under any growth condition, DBF, PHE or SAL (Fig. 1C and D). Because the exact location of this promoter is not known, it may be that the chosen fragment upstream of Swit_5102 was too short or wrongly placed. We concluded that strain RW1 (pME6012-P4925-egfp) was the best indicator for RW1 growth and DBF metabolism in the sand microcosms. RW1 population growth in contaminated sandy microcosms RW1 (pME6012-P4925-egfp) cells were inoculated at relatively low density in non-sterile sandy microcosms without antibiotic selection pressure [105 colony-forming units (cfu) per g material] and followed during 8 days for population growth and single cell EGFP fluorescence. The bacterial community was washed from the sand at individual time points, appropriately diluted and plated on both selective and non-selective media to count the number of cfu per gram of sand. RW1 population growth was assessed from colonies growing on minimal medium (MM) agar plates with 5 mM SAL and tetracycline (Tc, to select for the pME6012-derivative, Appendix S1). All colonies developing on MM + SAL + Tc agar plates showed green fluorescence, indicating no plasmid loss. No colonies developed on MM + SAL + Tc plates with extracts from non-inoculated controls. The RW1 population increased in microcosms consisting of sand with artificially added DBF (S + DBF, dosage 2.5 μmol g−1) from 6 × 105 to between 6 and 8 × 107 cfu g−1 5 days after inoculation (two independent inoculation series), which decreased to between 1 and
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 7, 480–488
482 E. Coronado, A. Valtat and J. R. van der Meer
Fig. 1. EGFP expression from pME6012-based reporter constructs in S. wittichii RW1 as a function of growth substrate in liquid culture. (A, B, C) Phase contrast (PhC) and corresponding EGFP fluorescence images (eGFP) of RW1 cells carrying the constructs pME6012-P4925-egfp, pME6012-PdxnA1-egfp or pME6012-P5102-egfp, respectively, grown with phenylalanine (Phe), salicylate (Sal) or dibenzofuran (DBF) as carbon sources. Images are scaled to the same intensity levels. (D) Average EGFP intensity of RW1 cells in early stationary phase cultures, ± calculated standard deviations from independent biological triplicates. The dotted line shows the background intensity level of the EGFP microscope images.
3 × 107 cfu g−1 at day 8 (Fig. 2A and B). This approximately equals eight generations of growth in soil in the absence of Tc, during which plasmid loss can be neglected (Fig. S1). The population of RW1 cells also developed in sand mixed with 10% (w/w) PAH-contaminated material from the Jonction former gasification site (Geneva, Switzerland) to which DBF was added (S + J + DBF, Fig. 2A and B). The maximum population size was smaller (3–4 × 107 cfu g−1 after 4 days), and the decline was more severe than in the case of S + DBF microcosms. In microcosms of sand with 10% w/w Ter Munck (TM, Belgium) loamy agricultural soil (van Gestel et al., 2012) but no aromatic compound contamination (S + TM), the population increased during the first day after inoculation from 1 × 106 to 8 × 106 cfu g−1, after which it decreased until around 1 × 106 cfu g−1 at day 8. Similarly, in the microcosms with sand and 10% Jonction material (S + J), a small increase from 3 × 104 to 1 × 106 cfu g−1 was observed after 1 day, after which the population remained at around 105 cfu g−1 soil (Fig. 2A). Populations developed similarly but not as pronounced for microcosms inoculated with RW1 (pME6012-PdxnA1-egfp, Fig. S4). The total cultivable community in the inoculated sandy microcosms was largely dominated by RW1 (Fig. 2C). Between 45% (S + J + DBF) and 85% (S + TM) of total counts were RW1, whereas counts in S + DBF micro-
cosms on MM + SAL + Tc were even slightly higher than on the PTYG medium (Balkwill and Ghiorse, 1985) (Fig. 2C). Total counts in microcosms (S + J + DBF) without inoculated RW1 were one third of those inoculated with RW1 (Fig. 2D). Very little growth occurred in microcosms consisting of sand plus Jonction (S + J) only, without RW1 being inoculated. Considerable growth occurred in sandy microcosms mixed with Jonction material that received extra phenanthrene (S + J + PHN, Fig. 2D), suggesting that phenanthrene degraders are more common than DBF degraders in the Jonction material. RW1 bioreporter response indicates DBF availability in contaminated sandy microcosms Average EGFP levels of RW1 (pME6012-P4925-egfp) cells inoculated in S + DBF microcosms increased twofold after inoculation on day 2, after which they slowly decreased until the end of the experiment (Fig. 3A). In contrast, average EGFP levels increased very slightly among cells inoculated in S + J + DBF after day 2, after which they decreased slowly to a level below that of inoculated cells. In all the microcosms supplemented with either pristine (S + TM) or contaminated soil (S + J), but without added DBF, the EGFP signals decreased from the day of
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 7, 480–488
Dibenzofuran detection by S. wittichii RW1
483
Fig. 2. Population size development of inoculated S. wittichii RW1 (pME6012-P4925-egfp) and the resident communities in different microcosms. (A, B) Colony-forming units (cfu g−1) in sand over time on MM + SAL + Tc plates (indicative for RW1). (C) Proportion of cfu g−1 sand in RW1 inoculated microcosms on MM + SAL + Tc plates (SAL) versus general medium (PTYG). (D) cfu g−1 sand in microcosms inoculated or not with RW1 on PTYG general medium. S + DBF, sandy microcosms with DBF added; S + J, sandy microcosms mixed with 10% contaminated material from the Jonction former gasification site; S + TM, sand mixed with 10% Ter Munck agricultural soil; S + J + PHN, sand with Jonction and added phenanthrene. Data points show averages from independent triplicate series ± calculated standard deviations.
inoculation onwards and remained lower than in microcosms with DBF (Fig. 3A). Average EGFP signals in RW1 cells from microcosm S + J were slightly and statistically significantly higher than in RW1 cells from S + TM (P < 0.001, one-way analysis of variance followed by post-Tukey test). Microscope images at different time points illustrate the increase in the overall number of RW1 reporter cells in S + DBF microcosms and their higher reporter signal (Fig. 3B). As expected from the behaviour in liquid culture, the EGFP signal intensity of inoculated RW1 (pME6012-PdxnA-egfp) cells did not vary much between microcosms amended with DBF or not, nor did it vary over time (Fig. S4). The distribution of EGFP signals among RW1 cells in S + DBF microcosms was relatively homogeneous (Fig. 4A), although the standard deviation was proportionally higher than in liquid cultures grown with DBF, SAL or PHE, and a more reactive subpopulation of between 3.7% and 13.9% occurred at day 1 (Table S3). In contrast,
EGFP distributions among RW1 cells were more heterogeneous in microcosms S + J + DBF, S + J and even S + TM (Fig. 4A, Fig. S5). Although the average EGFP levels in those microcosms were well below those of the S + DBF microcosms (Fig. 3A), consistently, small subpopulations of cells (∼ 1–10%) with higher EGFP levels occurred at all time points (Figs 3C and 4B, Table S3). These subpopulations, except for the case of the S + TM microcosms, displayed EGFP levels comparable to the population-average EGFP levels in S + DBF microcosms (Fig. 4B). We interpret this as evidence for a small fraction of cells for which DBF or compound causing a similar reporter activation as DBF was available in the microcosms. Various factors have been suggested to influence the potential successful establishment or activity of inoculated strains for bioaugmentation in contaminated material (Pritchard, 1992; van Veen et al., 1997; Tyagi et al., 2011), such as proper environmental growth
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 7, 480–488
484 E. Coronado, A. Valtat and J. R. van der Meer
Fig. 3. EGFP expression among RW1 (pME6012-P4925-egfp) cells in sandy microcosms. (A) Mean EGFP values (plus or minus the 95% confidence internal) over time from independent triplicate series. (B, C) Illustrative phase contrast (PhC) and corresponding EGFP images in S + DBF microcosms over time, or after 8 days in S + J + DBF, S + J and S + TM microcosms respectively. EGFP images scaled to the same intensity levels.
conditions (e.g. temperature, pH, humidity, oxygen) (Chen et al., 2008), general nutrient and carbon availability, presence of competitors (Shi et al., 2001; McGenity et al., 2012) or predators (Goldstein et al., 1985), toxicity
(Sikkema et al., 1995), or lack of availability of the primary target carbon substrates (Bosma et al., 1997; Harms, 1998; Johnsen and Karlson, 2004; Semple et al., 2007; Coppotelli et al., 2010). Many studies have expressed some frustration about the lack of establishment of the inoculated strain (Megharaj et al., 1997; Halden et al., 1999; Nam et al., 2005) but have used rather high starting cell numbers. Seeing that we can repeatedly grow a population of some 108 RW1 cells per gram of sand artificially contaminated with DBF (Fig. 2) (Moreno-Forero and van der Meer, 2015), it is obvious that not much growth can occur when inoculating with 108 cells per g material. Importantly, therefore, low starting cell densities make it easier to study the first episodes after inoculation and to discern the possible avenues of a strain’s establishment and activity. Our results show good growth of RW1 in non-sterile sand artificially contaminated with DBF to a population size as expected from the added amounts of DBF (eight generations, similar growth rates as in liquid culture; Moreno-Forero and van der Meer, 2015). We acknowledge that we did not specifically measure DBF consumption in these experiments but rely on previous similar experiments that showed specific DBF degradation by RW1 in microcosms with the same material and experimental set-up (Moreno-Forero and van der Meer, 2015). Moreover, RW1 population growth in sand alone (Moreno-Forero and van der Meer, 2015) or even in sand mixed with a regular agricultural soil (TM) is far below what we observed in sand with DBF (Fig. 4A). There can, therefore, be little doubt that RW1 is using the DBF in the sand for its growth, and also the pertinently increased EGFP expression from the Swit_4925 promoter under those conditions is evidence for DBF utilization (Fig. 3). However, as previous gene expression data with RW1 in similar microcosms have shown, despite the fact that the strain is growing in non-sterile sand with DBF, its transcriptome is largely different from growth in pure liquid culture (Moreno-Forero and van der Meer, 2015). The strain can, therefore, clearly adapt to a life in a porous sandy system within the limits of favourable boundary conditions (e.g. temperature, moisture), as long as its unique carbon source (DBF) is at its disposition. We find that RW1 also grows in microcosms without any specifically added DBF, such as S + TM (sand mixed with TM agricultural soil), something that we clearly would not have seen when starting at high cell densities. Apparently, the cells can still find carbon to divide some three times on average, and EGFP expression data suggest that a small proportion of cells remains ‘active’ for several days (Fig. 4, Fig. S5, Table S3), displaying higher than the bulk average EGFP levels from the Swit_4925 promoter. While sandy microcosms with artificially added DBF may
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 7, 480–488
Dibenzofuran detection by S. wittichii RW1
485
Fig. 4. EGFP expression variability among inoculated RW1 (pME6012-P4925-egfp) cells in sandy microcosms. (A) Density plots of the EGFP expression values among all observed cells in independent triplicate series at day 5 after inoculation in the indicated microcosms (abbreviations as before). n = total number of observed cells. (B) Quantile–quantile plots illustrating expected normal versus observed distribution of the EGFP expression values. Estimated subpopulation sizes with aberrant expression values above the dotted lines; estimations according to Reinhard and van der Meer (2013). For a full overview of all density plots, see Fig. S5. Estimated subpopulations across all data are reported in Table S3.
not be very realistic for a bioaugmentation process, the behaviour of RW1 was also studied in S + J microcosms, a 10% mixture of buried material from a former gasification work in Jonction with the sand. Inclusion of 10% Jonction material did not cause any apparent toxicity on RW1, given the results in S + J + DBF microcosms (Fig. 2), but it appeared that the inoculated cells found very little available carbon, sufficient for on average six cell doublings (from 3 × 104 to 1 × 106 cfu g−1 after day 1). Also here, EGFP expression data on recovered single cells showed a constant subpopulation of RW1 almost throughout the whole period of 8 days (Table S3), with expression levels approaching those of the bulk population average in S + DBF microcosms (Fig. 4). This indicates extreme population heterogeneity with the majority of cells inactive for DBF metabolism but some 5% of cells finding pockets of available carbon to induce the DBF metabolic network (Table S3). Metabolite competition in non-sterile mixed microcosms In confirmation of our initial working hypothesis, the RW1 population rapidly developed in sand supplemented with DBF. In contrast, its population density in microcosms containing Jonction material (S + J + DBF) fell drastically below that in S + DBF microcosms (Fig. 2D). Also, EGFP expression values from the Swit_4925 promoter in
S + J + DBF microcosms were on average far below those among RW1 cells developing in S + DBF microcosms (Figs 3 and 4). Although the size of the resident bacterial community in the sand is in the order of 104 culturable bacteria per g only (not shown), by mixing in material from Jonction a larger number of bacteria was introduced, which given its high content of PAHs (Appendix S1) likely contains strains capable of degrading aromatic compounds. Indeed, the number of culturable bacteria in uninoculated microcosms with Jonction material increased drastically upon addition of phenanthrene, although much less when DBF was added (Fig. 2D). We, therefore, speculate that poorer growth of RW1 (pME6012-P4925-egfp) in sand microcosms with Jonction and DBF (S + J + DBF) is due to carbon competition, but rather at the level of DBF metabolites produced by RW1 and not by competition for DBF itself. It has been frequently observed in liquid cultures that aromaticcompound degrading bacteria cannot maintain certain key metabolites inside the cell, which leak out and are subsequently taken up again (Schraa et al., 1986; Leveau et al., 1999; Jaspers et al., 2001). Metabolite competition has also been nicely demonstrated in laboratory studies using different combinations of aromatic compound degraders (Christensen et al., 2002). Utilization of DBF metabolites by resident bacteria from Jonction may thus result in earlier collapse of population development of
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 7, 480–488
486 E. Coronado, A. Valtat and J. R. van der Meer inoculated RW1. In order to test this, we inoculated RW1 (pME6012-P4925-egfp) in liquid medium with DBF, in the presence or absence of a mix of meta-toluate degrading strains isolated from Jonction material. Indeed, RW1 populations in co-cultures yielded only about half the population size attained in pure culture, whereas the mix of m-toluate degraders alone did not grow on DBF (Fig. S6). When assuming that the Swit_4925 promoter is induced by a DBF metabolite rather than DBF itself (Fig. S1), it would also make sense that EGFP expression in RW1 cells in S + J + DBF microcosms is strongly diminished compared with S + DBF alone. Value of bioreporter studies The use of a S. wittichii RW1 bioreporter was instrumental for the observations in this study. By being able to observe the first episodes of the inoculation process, we could conclude that RW1 cells can even grow to some extent in heavily contaminated material, but here the lack of available substrate is the key reason for the rapid cessation of its growth. In microcosms with contaminated material and amended with DBF, we could see that RW1 cells have an initial advantage, but this seems to lead to scavenging of DBF metabolites by others, leading to a decline in the RW1 population. Reporter strains faithfully showing local substrate availability and competition have not been used very frequently in bioremediation studies (Liu et al., 2010; Farhan Ul Haque et al., 2013), but have been more successfully applied in phyllosphere and rhizosphere studies, demonstrating the occurrence of extreme local substrate heterogeneity (Joyner and Lindow, 2000; Leveau and Lindow, 2001; Herron et al., 2010; Farhan Ul Haque et al., 2013; Parangan-Smith and Lindow, 2013). Despite frequently being more difficult to genetically engineer than typical laboratory model bacteria, specific bioreporter variants of strains intended for bioaugmentation will thus be a major asset to better assess the particular conditions for suitable growth and activity of the cells in contaminated sites. Acknowledgements This work was supported by Grant KBBE-211684 from European Commission within the FP7 Framework Programme. The authors thank Mr. Enga Luye for samples of the Jonction material and Dirk Springael for samples of Ter Munck agricultural soil. Vladimir Sentchilo is thanked for his help in the competition experiments.
References Armengaud, J., Happe, B., and Timmis, K.N. (1998) Genetic analysis of dioxin dioxygenase of Sphingomonas sp. strain
RW1: catabolic genes dispersed on the genome. J Bacteriol 180: 3954–3966. Balkwill, D.L., and Ghiorse, W.C. (1985) Characterization of subsurface bacteria associated with two shallow aquifers in Oklahoma. Appl Environ Microbiol 50: 580–588. Bosma, T.N.P., Middeldorp, P.J.M., Schraa, G., and Zehnder, A.J.B. (1997) Mass transfer limitation of biotransformation: quantifying bioavailability. Environ Sci Technol 31: 248– 252. Bunz, P.V., and Cook, A.M. (1993) Dibenzofuran 4,4adioxygenase from Sphingomonas sp. strain RW1: angular dioxygenation by a three-component enzyme system. J Bacteriol 175: 6467–6475. Chen, J., Wong, M.H., Wong, Y.S., and Tam, N.F. (2008) Multi-factors on biodegradation kinetics of polycyclic aromatic hydrocarbons (PAHs) by Sphingomonas sp. a bacterial strain isolated from mangrove sediment. Mar Poll Bull 57: 695–702. Christensen, B.B., Haagensen, J.A., Heydorn, A., and Molin, S. (2002) Metabolic commensalism and competition in a two-species microbial consortium. Appl Environ Microbiol 68: 2495–2502. Coppotelli, B.M., Ibarrolaza, A., Dias, R.L., Del Panno, M.T., Berthe-Corti, L., and Morelli, I.S. (2010) Study of the degradation activity and the strategies to promote the bioavailability of phenanthrene by Sphingomonas paucimobilis strain 20006FA. Microb Ecol 59: 266– 276. Coronado, E., Roggo, C., Johnson, D.R., and van der Meer, J.R. (2012) Genome-wide analysis of salicylate and dibenzofuran metabolism in Sphingomonas wittichii RW1. Front Microbiol 3: 300. El Fantroussi, S., and Agathos, S.N. (2005) Is bioaugmentation a feasible strategy for pollutant removal and site remediation? Curr Opin Microbiol 8: 268–275. Farhan Ul Haque, M., Nadalig, T., Bringel, F., Schaller, H., and Vuilleumier, S. (2013) Fluorescence-based bacterial bioreporter for specific detection of methyl halide emissions in the environment. Appl Environ Microbiol 79: 6561– 6567. Fida, T.T., Breugelmans, P., Lavigne, R., Coronado, E., Johnson, D.R., van der Meer, J.R., et al. (2012) Exposure to solute stress affects genome-wide expression but not the polycyclic aromatic hydrocarbon-degrading activity of Sphingomonas sp. strain LH128 in biofilms. Appl Environ Microbiol 78: 8311–8320. Fida, T.T., Moreno-Forero, S.K., Heipieper, H.J., and Springael, D. (2013) Physiology and transcriptome of the polycyclic aromatic hydrocarbon-degrading Sphingomonas sp. LH128 after long-term starvation. Microbiology 159: 1807–1817. Fida, T.T., Breugelmans, P., Lavigne, R., van der Meer, J.R., De Mot, R., Vaysse, P.J., and Springael, D. (2014) Identification of opsA, a gene involved in solute stress mitigation and survival in soil, in the polycyclic aromatic hydrocarbondegrading bacterium Novosphingobium sp. strain LH128. Appl Environ Microbiol 80: 3350–3361. van Gestel, C.A., McGrath, S.P., Smolders, E., Ortiz, M.D., Borgman, E., Verweij, R.A., et al. (2012) Effect of long-term equilibration on the toxicity of molybdenum to soil organisms. Environ Pollut 162: 1–7.
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 7, 480–488
Dibenzofuran detection by S. wittichii RW1 Goldstein, R.M., Mallory, L.M., and Alexander, M. (1985) Reasons for possible failure of inoculation to enhance biodegradation. Appl Environ Microbiol 50: 977–983. Gunasekera, T.S., Striebich, R.C., Mueller, S.S., Strobel, E.M., and Ruiz, O.N. (2013) Transcriptional profiling suggests that multiple metabolic adaptations are required for effective proliferation of Pseudomonas aeruginosa in jet fuel. Environ Sci Technol 47: 13449–13458. Halden, R.U., Halden, B.G., and Dwyer, D.F. (1999) Removal of dibenzofuran, dibenzo-p-dioxin, and 2-chlorodibenzo-pdioxin from soils inoculated with Sphingomonas sp. strain RW1. Appl Environ Microbiol 65: 2246–2249. Harms, H. (1998) Bioavailability of dioxin-like compounds for microbial degradation. In Biodegradation of dioxins and furans. Wittich, R. (ed.). Austin, TX, USA: Springer-Verlag and R.G. Landes Company, pp. 135–163. Hartmann, E.M., and Armengaud, J. (2014) Shotgun proteomics suggests involvement of additional enzymes in dioxin degradation by Sphingomonas wittichii RW1. Environ Microbiol 16: 162–176. Heeb, S., Itoh, Y., Nishijyo, T., Schnider, U., Keel, C., Wade, J., and Haas, D. (2000) Small, stable shuttle vectors based on the minimal pVS1 replicon for use in gram-negative, plant-associated bacteria. Mol Plant Microbe Interact 13: 232–237. Herron, P.M., Gage, D.J., and Cardon, Z.G. (2010) Microscale water potential gradients visualized in soil around plant root tips using microbiosensors. Plant Cell Environ 33: 199–210. Iino, T., Wang, Y., Miyauchi, K., Kasai, D., Masai, E., Fujii, T., et al. (2012) Specific gene responses of Rhodococcus jostii RHA1 during growth in soil. Appl Environ Microbiol 78: 6954–6962. Jaspers, M.C., Schmid, A., Sturme, M.H., Goslings, D.A., Kohler, H.P., and Van der Meer, J.R. (2001) Transcriptional organization and dynamic expression of the hbpCAD genes, which encode the first three enzymes for 2hydroxybiphenyl degradation in Pseudomonas azelaica HBP1. J Bacteriol 183: 270–279. Johnsen, A.R., and Karlson, U. (2004) Evaluation of bacterial strategies to promote the bioavailability of polycyclic aromatic hydrocarbons. Appl Microbiol Biotechnol 63: 452– 459. Joyner, D.C., and Lindow, S.E. (2000) Heterogeneity of iron bioavailability on plants assessed with a whole- cell GFPbased bacterial biosensor. Microbiology 146: 2435–2445. Leveau, J.H.J., and Lindow, S.E. (2001) Appetite of an epiphyte: quantitative monitoring of bacterial sugar consumption in the phyllosphere. Proc Natl Acad Sci USA 98: 3446– 3453. Leveau, J.H.J., and Lindow, S.E. (2002) Bioreporters in microbial ecology. Curr Opin Microbiol 5: 259–265. Leveau, J.H.J., König, F., Füchslin, H.-P., Werlen, C., and van der Meer, J.R. (1999) Dynamics of multigene expression during catabolic adaptation of Ralstonia eutropha JMP134 (pJP4) to the herbicide 2, 4-dichlorophenoxyacetate. Mol Microbiol 33: 396–406. Liu, X., Germaine, K.J., Ryan, D., and Dowling, D.N. (2010) Whole-cell fluorescent biosensors for bioavailability and biodegradation of polychlorinated biphenyls. Sensors (Basel) 10: 1377–1398.
487
McGenity, T.J., Folwell, B.D., McKew, B.A., and Sanni, G.O. (2012) Marine crude-oil biodegradation: a central role for interspecies interactions. Aquat Biosystems 8: 10. Maphosa, F., Lieten, S.H., Dinkla, I., Stams, A.J., Smidt, H., and Fennell, D.E. (2012) Ecogenomics of microbial communities in bioremediation of chlorinated contaminated sites. Front Microbiol 3: 351. Megharaj, M., Wittich, R.-M., Blasco, R., Pieper, D.H., and Timmis, K.N. (1997) Superior survival and degradation of dibenzo-p-dioxin and dibenzofuran in soil by soil-adapted Sphingomonas sp. strain RW1. Appl Microbiol Biotechnol 48: 109–114. Moreno-Forero, S.K., and van der Meer, J.R. (2015) Genome-wide analysis of Sphingomonas wittichii RW1 behaviour during inoculation and growth in contaminated sand. ISME J 9: 150–165. doi:10.1038/ismej.2014.101. Nam, I.H., Hong, H.B., Kim, Y.M., Kim, B.H., Murugesan, K., and Chang, Y.S. (2005) Biological removal of polychlorinated dibenzo-p-dioxins from incinerator fly ash by Sphingomonas wittichii RW1. Water Res 39: 4651–4660. Parangan-Smith, A., and Lindow, S. (2013) Contribution of nitrate assimilation to the fitness of Pseudomonas syringae pv. syringae B728a on plants. Appl Environ Microbiol 79: 678–687. Pritchard, R.P. (1992) Use of inoculation in bioremediation. Curr Opin Biotechnol 3: 232–243. Reinhard, F., and van der Meer, J.R. (2013) Improved statistical analysis of low abundance phenomena in bimodal bacterial populations. PLoS ONE 8: e78288. Roggo, C., Coronado, E., Moreno-Forero, S.K., Harshman, K., Weber, J., and van der Meer, J.R. (2013) Genome-wide transposon insertion scanning of environmental survival functions in the polycyclic aromatic hydrocarbon degrading bacterium Sphingomonas wittichii RW1. Environ Microbiol 15: 2681–2695. Safe, S. (1990) Polychlorinated biphenyls (PCBs), dibenzop-dioxins (PCDDs), dibenzofurans (PCDFs), and related compounds: environmental and mechanistic considerations which support the development of toxic equivalency factors (TEFs). Crit Rev Toxicol 21: 51–88. Scheublin, T.R., Deusch, S., Moreno-Forero, S.K., Müller, J.A., van der Meer, J.R., and Leveau, J.H. (2014) Transcriptional profiling of Gram-positive Arthrobacter in the phyllosphere: induction of pollutant degradation genes by natural plant phenolic compounds. Environ Microbiol 16: 2212–2225. Schraa, G., Boone, M.L., Jetten, M.S., van Neerven, A.R., Colberg, P.J., and Zehnder, A.J. (1986) Degradation of 1,4-dichlorobenzene by Alcaligenes sp. strain A175. Appl Environ Microbiol 52: 1374–1381. Semple, K.T., Doick, K.J., Wick, L.Y., and Harms, H. (2007) Microbial interactions with organic contaminants in soil: definitions, processes and measurement. Environ Poll 150: 166–176. Shi, T., Fredrickson, J.K., and Balkwill, D.L. (2001) Biodegradation of polycyclic aromatic hydrocarbons by Sphingomonas strains isolated from the terrestrial subsurface. J Ind Microbiol Biotechnol 26: 283–289. Sikkema, J., de Bont, J.A., and Poolman, B. (1995) Mechanisms of membrane toxicity of hydrocarbons. Microbiol Rev 59: 201–222.
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 7, 480–488
488 E. Coronado, A. Valtat and J. R. van der Meer Tecon, R., and van der Meer, J.R. (2006) Information from single-cell bacterial biosensors: what is it good for? Curr Opin Biotechnol 17: 4–10. Thompson, I.P., van der Gast, C.J., Ciric, L., and Singer, A.C. (2005) Bioaugmentation for bioremediation: the challenge of strain selection. Environ Microbiol 7: 909–915. Tyagi, M., da Fonseca, M.M., and de Carvalho, C.C. (2011) Bioaugmentation and biostimulation strategies to improve the effectiveness of bioremediation processes. Biodegradation 22: 231–241. van Veen, J.A., van Overbeek, L.S., and van Elsas, J.D. (1997) Fate and activity of microorganisms introduced into soil. Microbiol Mol Biol Rev 61: 121–135. Verschuere, L., Rombaut, G., Sorgeloos, P., and Verstraete, W. (2000) Probiotic bacteria as biological control agents in aquaculture. Microbiol Mol Biol Rev 64: 655–671. Wilkes, H., Wittich, R., Timmis, K.N., Fortnagel, P., and Francke, W. (1996) Degradation of chlorinated dibenzofurans and dibenzo-p-dioxins by Sphingomonas sp. strain RW1. Appl Environ Microbiol 62: 367–371. Wittich, R.-M., Wilkes, H., Sinnwell, V., Francke, W., and Fortnagel, P. (1992) Metabolism of dibenzo-p-dioxin by Sphingomonas sp. strain RW1. Appl Environ Microbiol 58: 1005–1010.
Supporting information Additional Supporting Information may be found in the online version of this article at the publisher’s web-site: Fig. S1. Metabolic network for DBF and aromatic compound metabolism by S. wittichii RW1, displayed using CYTOSKAPE 2.8.3, as inferred for cells growing on DBF (Moreno-Forero and van der Meer, 2014). Nodes represent substrates and metabolic intermediates. Edges represent enzyme reactions converting the linked compounds. Edge line thickness is a representation of normalized expression of the gene coding for the particular enzyme that carries out the reaction between two nodes. Normalization was carried out per gene among six conditions, taking the highest expression value as 100% (=line width 15). Line widths are a linear scale representation of normalized gene expression, notation and the predicted gene function. For compound abbreviations, see Moreno-Forero and van der Meer (2014). Fig. S2. Stability of the pPROBE-PTac-egfp (A), pME6012PTac-egfp (B) and miniTn5::PTac-egfp (C) constructs in Sphingomonas wittichii RW1. Shown is marker stability over
four subsequent culture transfers (∼ 28 generations) in liquid medium with SAL as sole carbon source without (no AB) or with (AB) addition of antibiotic selecting for the plasmid or transposon insertion. Proportions show the average number of antibiotic resistant colonies (Resist) divided by the number of colonies on non-selective media, or the number of fluorescent colonies amidst all resistant colonies (R/Fluo), ± calculated standard deviation from independent triplicates. For pPROBE and mini-Tn5, Km was used as antibiotic, whereas Tc was used for pME6012. Fig. S3. EGFP production rate as a function of growth rate of RW1 carrying pME6012-P4925-egfp, pME6012-PdxnA1egfp or mini-Tn5-PuspA-egfp. Rates were calculated from linear regression of ln-transformed normalized EGFP intensities or culture turbidity values versus time. Decreasing exponential growth rate in liquid culture was achieved by successive addition of NaCl (open symbols) or polyethylene glycol (closed symbols), which lowers the medium water potential (Johnson et al., 2011). Fig. S4. Population development and reporter gene expression of RW1 (pME6012-PPdxnA1-egfp) inoculated in sandy microcosms S + DBF (red symbols) and S + TM (green symbols). (A) Colony-forming units on MM + SAL + Tc plates (cfu g−1 soil) over time, representative for RW1. (B) Mean EGFP expression from triplicate series ± calculated 95% confidence interval. Note that the RW1 population for calculating EGFP values is much smaller in S + TM than in S + DBF. Fig. S5. EGFP expression density plots for bacterial cells extracted from the sandy microcosms at the different time points. All plots scaled to the same EGFP intensity level (x-axis). Fig. S6. Competition for DBF intermediates. (A) Population development of RW1 (pME6012-P4925-egfp) alone (open symbols) or in co-culture (closed symbols) with a mixture of three meta-toluate degrading strains isolated from Jonction in liquid medium with DBF as sole carbon source (arrow indicates the time of their addition). (B) Average EGFP levels of individual RW1 reporter cells under both conditions measured by flow cytometry. Data points are averages from independent triplicates ± calculated standard deviations. Table S1. Strains used in this study. Table S2. Primers used in this study. Table S3. Variability of EGFP expression among individual RW1 (pME6012-P4925-egfp) cells under different growth conditions. Appendix S1. Supplementary experimental methods. Appendix S2. Supplementary data.
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 7, 480–488
Environmental Microbiology Reports (2015) 7(3), 480–488
doi:10.1111/1758-2229.12276
Sphingomonas wittichii RW1 gene reporters interrogating the dibenzofuran metabolic network highlight conditions for early successful development in contaminated microcosms
Edith Coronado, Annabelle Valtat and Jan R. van der Meer* Department of Fundamental Microbiology, University of Lausanne, Lausanne 1015, Switzerland. Summary In order to better understand the fate and activity of bacteria introduced into contaminated material for the purpose of enhancing biodegradation rates, we constructed Sphingomonas wittichii RW1 variants with gene reporters interrogating dibenzofuran metabolic activity. Three potential promoters from the dibenzofuran metabolic network were selected and fused to the gene for enhanced green fluorescent protein (EGFP). The stability of the resulting genetic constructions in RW1 was examined, with plasmids based on the broad-host range vector pME6012 being the most reliable. One of the selected promoters, upstream of the gene Swit_4925 for a putative 2-hydroxy-2,4-pentadienoate hydratase, was inducible by growth on dibenzofuran. Sphingomonas wittichii RW1 equipped with the Swit_4925 promoter egfp fusion grew in a variety of non-sterile sandy microcosms contaminated with dibenzofuran and material from a former gasification site. The strain also grew in microcosms without added dibenzofuran but to a very limited extent, and EGFP expression indicated the formation of consistent small subpopulations of cells with an active inferred dibenzofuran metabolic network. Evidence was obtained for competition for dibenzofuran metabolites scavenged by resident bacteria in the gasification site material, which resulted in a more rapid decline of the RW1 population. Our results show the importance of low inoculation densities in order to observe the population development of the introduced bacteria and further illustrate that the limited
Received 8 October, 2014; accepted 31 January, 2015. *For correspondence. E-mail [email protected]; Tel. +41 (21) 692 5630; Fax +41 (21) 692 5605.
availability of unique carbon substrate may be the most important factor impinging growth. Introduction Inoculation of pure bacterial cultures or enrichments into complex microbial communities with the purpose of introducing or enhancing specific functionalities has enjoyed a long-standing interest (Pritchard, 1992; van Veen et al., 1997; Verschuere et al., 2000; El Fantroussi and Agathos, 2005; Thompson et al., 2005). In the area of environmental biotechnology, the inoculation into contaminated material of specific pre-enriched bacteria, which efficiently degrade one or more of the contaminants under laboratory conditions, has been frequently proposed as a means to accelerate biodegradation rates (El Fantroussi and Agathos, 2005; Thompson et al., 2005). However, even more than 20 years since the first strategic review (Pritchard, 1992), the process of bioaugmentation is still largely unpredictable because this is mostly trial and error-based (Tyagi et al., 2011). In order to improve our general understanding of the process, recent studies have attempted to more systematically measure and interpret global reactions of introduced bacteria for bioremediation into complex systems (Fida et al., 2012; 2013; Iino et al., 2012; Maphosa et al., 2012; Gunasekera et al., 2013; Moreno-Forero and van der Meer, 2015; Scheublin et al., 2014). This has been complemented with global transposon mutagenesis approaches in order to identify gene functions with fitness benefit for growth and survival in soil (Roggo et al., 2013; Fida et al., 2014). Whereas ‘big data’ approaches are powerful to characterize global cellular reactions and identify candidate genes essential for fitness or survival, they need complementation by methods that can measure the variability of individual cellular responses over time under local conditions. Gene reporter technology has been proposed for this (Leveau and Lindow, 2002; Tecon and van der Meer, 2006), and has been successfully applied to study, for example, local sugar availability for plant pathogens on plant leaves (Leveau and Lindow, 2001), or water (Herron
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd. This is an open access article under the terms of the Creative Commons Attribution-NonCommercial-NoDerivs License, which permits use and distribution in any medium, provided the original work is properly cited, the use is non-commercial and no modifications or adaptations are made.
Dibenzofuran detection by S. wittichii RW1 et al., 2010) and iron availability to bacteria in the phyllosphere (Joyner and Lindow, 2000). The goal of the underlying study was to obtain a more complete picture of the factors determining the success or failure of Sphingomonas wittichii RW1 to grow in non-sterile microcosms with contaminated material. Sphingomonas wittichii RW1 metabolizes dibenzofuran (DBF) and dibenzodioxins, which are frequently found in areas contaminated with polycyclic aromatic hydrocarbons (PAH) and after incineration processes (Safe, 1990). Strain RW1 has been studied in detail for its DBF and dioxin metabolic pathways (Wittich et al., 1992; Wilkes et al., 1996; Armengaud et al., 1998; Hartmann and Armengaud, 2014), and has also been applied to some success in bioaugmentation studies (Megharaj et al., 1997; Halden et al., 1999; Nam et al., 2005). Our hypothesis was that the growth of RW1 in nonsterile material would be mostly controlled by the presence of its unique carbon source (DBF) irrespective of the presence of other PAH-contaminated material. In order to study DBF availability for RW1 at the single-cell level, we generated RW1 reporter strains where egfp expression for enhanced green fluorescent protein (EGFP) would be under control of a promoter from the DBF metabolic network (Appendix S1, Fig. S1). As genetic engineering in strains like RW1 is non-trivial, we examined the genetic stability of the constructs and EGFP expression driven from the three regions in liquid cultures under a variety of conditions. The best performing RW1 bioreporter was then inoculated in sandy microcosms spiked or not with DBF or PAH-contaminated material to follow population growth and reporter induction among single cells. Our results indicate that RW1 can indeed grow and induce its DBF metabolic network in contaminated material, depending on the substrate availability. However, our results further suggest that resident bacteria from the contaminated material may compete with RW1 at the level of metabolites formed by RW1 from DBF.
Results and discussion Development of a S. wittichii RW1 bioreporter for DBF degradation Three putative RW1 promoter regions (Tables S1 and S2, Fig. S1) were fused to a promoterless egfp gene, introduced into strain RW1 and evaluated for their expression during growth on DBF, salicylate (SAL) or phenylalanine (PHE). The first is a region upstream of the gene Swit_4925, putatively involved in the transformation of 2-hydroxy-2,4-pentadienoate and 12-fold induced during growth on DBF (Coronado et al., 2012). The second was a putative promoter region upstream of Swit_5102, predicted to code for a gentisate dioxygenase and up to
481
17-fold induced (Coronado et al., 2012). As third promoter, we selected the region upstream of the gene Swit_4897 (dxnA1), which codes for dibenzofuran-4,4adioxygenase, catalysing the initial step in DBF degradation (Bunz and Cook, 1993; Armengaud et al., 1998). Promoter-egfp constructs based on vector pME6012 (Heeb et al., 2000) were stable in RW1 for eight generations in the absence of antibiotic selection compared with pPROBE- or mini-Tn5-based constructions (Fig. S2, Appendix S2). Strain RW1 (pME6012-P4925-egfp) displayed fivefold higher background-corrected fluorescence when growing on DBF than on SAL or PHE (Fig. 1A and D). In contrast, RW1 (pME6012-PdxnA1-egfp) produced a slightly higher EGFP fluorescence when grown with SAL compared with PHE or DBF (Fig. 1B and D). The EGFP expression from RW1 (pME6012-P4925-egfp) but less so from RW1 (pME6012-PdxnA1-egfp) was insensitive to changes in water potential, which might influence its behaviour in unsaturated soil (Fig. S3, Appendix S2). Contrary to our expectations from microarray data (17-fold higher expression of Swit_5102 in DBF-grown compared with PHE-grown cells), strain RW1 (pME6012-P5102-egfp) hardly produced any EGFP fluorescence above background under any growth condition, DBF, PHE or SAL (Fig. 1C and D). Because the exact location of this promoter is not known, it may be that the chosen fragment upstream of Swit_5102 was too short or wrongly placed. We concluded that strain RW1 (pME6012-P4925-egfp) was the best indicator for RW1 growth and DBF metabolism in the sand microcosms. RW1 population growth in contaminated sandy microcosms RW1 (pME6012-P4925-egfp) cells were inoculated at relatively low density in non-sterile sandy microcosms without antibiotic selection pressure [105 colony-forming units (cfu) per g material] and followed during 8 days for population growth and single cell EGFP fluorescence. The bacterial community was washed from the sand at individual time points, appropriately diluted and plated on both selective and non-selective media to count the number of cfu per gram of sand. RW1 population growth was assessed from colonies growing on minimal medium (MM) agar plates with 5 mM SAL and tetracycline (Tc, to select for the pME6012-derivative, Appendix S1). All colonies developing on MM + SAL + Tc agar plates showed green fluorescence, indicating no plasmid loss. No colonies developed on MM + SAL + Tc plates with extracts from non-inoculated controls. The RW1 population increased in microcosms consisting of sand with artificially added DBF (S + DBF, dosage 2.5 μmol g−1) from 6 × 105 to between 6 and 8 × 107 cfu g−1 5 days after inoculation (two independent inoculation series), which decreased to between 1 and
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 7, 480–488
482 E. Coronado, A. Valtat and J. R. van der Meer
Fig. 1. EGFP expression from pME6012-based reporter constructs in S. wittichii RW1 as a function of growth substrate in liquid culture. (A, B, C) Phase contrast (PhC) and corresponding EGFP fluorescence images (eGFP) of RW1 cells carrying the constructs pME6012-P4925-egfp, pME6012-PdxnA1-egfp or pME6012-P5102-egfp, respectively, grown with phenylalanine (Phe), salicylate (Sal) or dibenzofuran (DBF) as carbon sources. Images are scaled to the same intensity levels. (D) Average EGFP intensity of RW1 cells in early stationary phase cultures, ± calculated standard deviations from independent biological triplicates. The dotted line shows the background intensity level of the EGFP microscope images.
3 × 107 cfu g−1 at day 8 (Fig. 2A and B). This approximately equals eight generations of growth in soil in the absence of Tc, during which plasmid loss can be neglected (Fig. S1). The population of RW1 cells also developed in sand mixed with 10% (w/w) PAH-contaminated material from the Jonction former gasification site (Geneva, Switzerland) to which DBF was added (S + J + DBF, Fig. 2A and B). The maximum population size was smaller (3–4 × 107 cfu g−1 after 4 days), and the decline was more severe than in the case of S + DBF microcosms. In microcosms of sand with 10% w/w Ter Munck (TM, Belgium) loamy agricultural soil (van Gestel et al., 2012) but no aromatic compound contamination (S + TM), the population increased during the first day after inoculation from 1 × 106 to 8 × 106 cfu g−1, after which it decreased until around 1 × 106 cfu g−1 at day 8. Similarly, in the microcosms with sand and 10% Jonction material (S + J), a small increase from 3 × 104 to 1 × 106 cfu g−1 was observed after 1 day, after which the population remained at around 105 cfu g−1 soil (Fig. 2A). Populations developed similarly but not as pronounced for microcosms inoculated with RW1 (pME6012-PdxnA1-egfp, Fig. S4). The total cultivable community in the inoculated sandy microcosms was largely dominated by RW1 (Fig. 2C). Between 45% (S + J + DBF) and 85% (S + TM) of total counts were RW1, whereas counts in S + DBF micro-
cosms on MM + SAL + Tc were even slightly higher than on the PTYG medium (Balkwill and Ghiorse, 1985) (Fig. 2C). Total counts in microcosms (S + J + DBF) without inoculated RW1 were one third of those inoculated with RW1 (Fig. 2D). Very little growth occurred in microcosms consisting of sand plus Jonction (S + J) only, without RW1 being inoculated. Considerable growth occurred in sandy microcosms mixed with Jonction material that received extra phenanthrene (S + J + PHN, Fig. 2D), suggesting that phenanthrene degraders are more common than DBF degraders in the Jonction material. RW1 bioreporter response indicates DBF availability in contaminated sandy microcosms Average EGFP levels of RW1 (pME6012-P4925-egfp) cells inoculated in S + DBF microcosms increased twofold after inoculation on day 2, after which they slowly decreased until the end of the experiment (Fig. 3A). In contrast, average EGFP levels increased very slightly among cells inoculated in S + J + DBF after day 2, after which they decreased slowly to a level below that of inoculated cells. In all the microcosms supplemented with either pristine (S + TM) or contaminated soil (S + J), but without added DBF, the EGFP signals decreased from the day of
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 7, 480–488
Dibenzofuran detection by S. wittichii RW1
483
Fig. 2. Population size development of inoculated S. wittichii RW1 (pME6012-P4925-egfp) and the resident communities in different microcosms. (A, B) Colony-forming units (cfu g−1) in sand over time on MM + SAL + Tc plates (indicative for RW1). (C) Proportion of cfu g−1 sand in RW1 inoculated microcosms on MM + SAL + Tc plates (SAL) versus general medium (PTYG). (D) cfu g−1 sand in microcosms inoculated or not with RW1 on PTYG general medium. S + DBF, sandy microcosms with DBF added; S + J, sandy microcosms mixed with 10% contaminated material from the Jonction former gasification site; S + TM, sand mixed with 10% Ter Munck agricultural soil; S + J + PHN, sand with Jonction and added phenanthrene. Data points show averages from independent triplicate series ± calculated standard deviations.
inoculation onwards and remained lower than in microcosms with DBF (Fig. 3A). Average EGFP signals in RW1 cells from microcosm S + J were slightly and statistically significantly higher than in RW1 cells from S + TM (P < 0.001, one-way analysis of variance followed by post-Tukey test). Microscope images at different time points illustrate the increase in the overall number of RW1 reporter cells in S + DBF microcosms and their higher reporter signal (Fig. 3B). As expected from the behaviour in liquid culture, the EGFP signal intensity of inoculated RW1 (pME6012-PdxnA-egfp) cells did not vary much between microcosms amended with DBF or not, nor did it vary over time (Fig. S4). The distribution of EGFP signals among RW1 cells in S + DBF microcosms was relatively homogeneous (Fig. 4A), although the standard deviation was proportionally higher than in liquid cultures grown with DBF, SAL or PHE, and a more reactive subpopulation of between 3.7% and 13.9% occurred at day 1 (Table S3). In contrast,
EGFP distributions among RW1 cells were more heterogeneous in microcosms S + J + DBF, S + J and even S + TM (Fig. 4A, Fig. S5). Although the average EGFP levels in those microcosms were well below those of the S + DBF microcosms (Fig. 3A), consistently, small subpopulations of cells (∼ 1–10%) with higher EGFP levels occurred at all time points (Figs 3C and 4B, Table S3). These subpopulations, except for the case of the S + TM microcosms, displayed EGFP levels comparable to the population-average EGFP levels in S + DBF microcosms (Fig. 4B). We interpret this as evidence for a small fraction of cells for which DBF or compound causing a similar reporter activation as DBF was available in the microcosms. Various factors have been suggested to influence the potential successful establishment or activity of inoculated strains for bioaugmentation in contaminated material (Pritchard, 1992; van Veen et al., 1997; Tyagi et al., 2011), such as proper environmental growth
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 7, 480–488
484 E. Coronado, A. Valtat and J. R. van der Meer
Fig. 3. EGFP expression among RW1 (pME6012-P4925-egfp) cells in sandy microcosms. (A) Mean EGFP values (plus or minus the 95% confidence internal) over time from independent triplicate series. (B, C) Illustrative phase contrast (PhC) and corresponding EGFP images in S + DBF microcosms over time, or after 8 days in S + J + DBF, S + J and S + TM microcosms respectively. EGFP images scaled to the same intensity levels.
conditions (e.g. temperature, pH, humidity, oxygen) (Chen et al., 2008), general nutrient and carbon availability, presence of competitors (Shi et al., 2001; McGenity et al., 2012) or predators (Goldstein et al., 1985), toxicity
(Sikkema et al., 1995), or lack of availability of the primary target carbon substrates (Bosma et al., 1997; Harms, 1998; Johnsen and Karlson, 2004; Semple et al., 2007; Coppotelli et al., 2010). Many studies have expressed some frustration about the lack of establishment of the inoculated strain (Megharaj et al., 1997; Halden et al., 1999; Nam et al., 2005) but have used rather high starting cell numbers. Seeing that we can repeatedly grow a population of some 108 RW1 cells per gram of sand artificially contaminated with DBF (Fig. 2) (Moreno-Forero and van der Meer, 2015), it is obvious that not much growth can occur when inoculating with 108 cells per g material. Importantly, therefore, low starting cell densities make it easier to study the first episodes after inoculation and to discern the possible avenues of a strain’s establishment and activity. Our results show good growth of RW1 in non-sterile sand artificially contaminated with DBF to a population size as expected from the added amounts of DBF (eight generations, similar growth rates as in liquid culture; Moreno-Forero and van der Meer, 2015). We acknowledge that we did not specifically measure DBF consumption in these experiments but rely on previous similar experiments that showed specific DBF degradation by RW1 in microcosms with the same material and experimental set-up (Moreno-Forero and van der Meer, 2015). Moreover, RW1 population growth in sand alone (Moreno-Forero and van der Meer, 2015) or even in sand mixed with a regular agricultural soil (TM) is far below what we observed in sand with DBF (Fig. 4A). There can, therefore, be little doubt that RW1 is using the DBF in the sand for its growth, and also the pertinently increased EGFP expression from the Swit_4925 promoter under those conditions is evidence for DBF utilization (Fig. 3). However, as previous gene expression data with RW1 in similar microcosms have shown, despite the fact that the strain is growing in non-sterile sand with DBF, its transcriptome is largely different from growth in pure liquid culture (Moreno-Forero and van der Meer, 2015). The strain can, therefore, clearly adapt to a life in a porous sandy system within the limits of favourable boundary conditions (e.g. temperature, moisture), as long as its unique carbon source (DBF) is at its disposition. We find that RW1 also grows in microcosms without any specifically added DBF, such as S + TM (sand mixed with TM agricultural soil), something that we clearly would not have seen when starting at high cell densities. Apparently, the cells can still find carbon to divide some three times on average, and EGFP expression data suggest that a small proportion of cells remains ‘active’ for several days (Fig. 4, Fig. S5, Table S3), displaying higher than the bulk average EGFP levels from the Swit_4925 promoter. While sandy microcosms with artificially added DBF may
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 7, 480–488
Dibenzofuran detection by S. wittichii RW1
485
Fig. 4. EGFP expression variability among inoculated RW1 (pME6012-P4925-egfp) cells in sandy microcosms. (A) Density plots of the EGFP expression values among all observed cells in independent triplicate series at day 5 after inoculation in the indicated microcosms (abbreviations as before). n = total number of observed cells. (B) Quantile–quantile plots illustrating expected normal versus observed distribution of the EGFP expression values. Estimated subpopulation sizes with aberrant expression values above the dotted lines; estimations according to Reinhard and van der Meer (2013). For a full overview of all density plots, see Fig. S5. Estimated subpopulations across all data are reported in Table S3.
not be very realistic for a bioaugmentation process, the behaviour of RW1 was also studied in S + J microcosms, a 10% mixture of buried material from a former gasification work in Jonction with the sand. Inclusion of 10% Jonction material did not cause any apparent toxicity on RW1, given the results in S + J + DBF microcosms (Fig. 2), but it appeared that the inoculated cells found very little available carbon, sufficient for on average six cell doublings (from 3 × 104 to 1 × 106 cfu g−1 after day 1). Also here, EGFP expression data on recovered single cells showed a constant subpopulation of RW1 almost throughout the whole period of 8 days (Table S3), with expression levels approaching those of the bulk population average in S + DBF microcosms (Fig. 4). This indicates extreme population heterogeneity with the majority of cells inactive for DBF metabolism but some 5% of cells finding pockets of available carbon to induce the DBF metabolic network (Table S3). Metabolite competition in non-sterile mixed microcosms In confirmation of our initial working hypothesis, the RW1 population rapidly developed in sand supplemented with DBF. In contrast, its population density in microcosms containing Jonction material (S + J + DBF) fell drastically below that in S + DBF microcosms (Fig. 2D). Also, EGFP expression values from the Swit_4925 promoter in
S + J + DBF microcosms were on average far below those among RW1 cells developing in S + DBF microcosms (Figs 3 and 4). Although the size of the resident bacterial community in the sand is in the order of 104 culturable bacteria per g only (not shown), by mixing in material from Jonction a larger number of bacteria was introduced, which given its high content of PAHs (Appendix S1) likely contains strains capable of degrading aromatic compounds. Indeed, the number of culturable bacteria in uninoculated microcosms with Jonction material increased drastically upon addition of phenanthrene, although much less when DBF was added (Fig. 2D). We, therefore, speculate that poorer growth of RW1 (pME6012-P4925-egfp) in sand microcosms with Jonction and DBF (S + J + DBF) is due to carbon competition, but rather at the level of DBF metabolites produced by RW1 and not by competition for DBF itself. It has been frequently observed in liquid cultures that aromaticcompound degrading bacteria cannot maintain certain key metabolites inside the cell, which leak out and are subsequently taken up again (Schraa et al., 1986; Leveau et al., 1999; Jaspers et al., 2001). Metabolite competition has also been nicely demonstrated in laboratory studies using different combinations of aromatic compound degraders (Christensen et al., 2002). Utilization of DBF metabolites by resident bacteria from Jonction may thus result in earlier collapse of population development of
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 7, 480–488
486 E. Coronado, A. Valtat and J. R. van der Meer inoculated RW1. In order to test this, we inoculated RW1 (pME6012-P4925-egfp) in liquid medium with DBF, in the presence or absence of a mix of meta-toluate degrading strains isolated from Jonction material. Indeed, RW1 populations in co-cultures yielded only about half the population size attained in pure culture, whereas the mix of m-toluate degraders alone did not grow on DBF (Fig. S6). When assuming that the Swit_4925 promoter is induced by a DBF metabolite rather than DBF itself (Fig. S1), it would also make sense that EGFP expression in RW1 cells in S + J + DBF microcosms is strongly diminished compared with S + DBF alone. Value of bioreporter studies The use of a S. wittichii RW1 bioreporter was instrumental for the observations in this study. By being able to observe the first episodes of the inoculation process, we could conclude that RW1 cells can even grow to some extent in heavily contaminated material, but here the lack of available substrate is the key reason for the rapid cessation of its growth. In microcosms with contaminated material and amended with DBF, we could see that RW1 cells have an initial advantage, but this seems to lead to scavenging of DBF metabolites by others, leading to a decline in the RW1 population. Reporter strains faithfully showing local substrate availability and competition have not been used very frequently in bioremediation studies (Liu et al., 2010; Farhan Ul Haque et al., 2013), but have been more successfully applied in phyllosphere and rhizosphere studies, demonstrating the occurrence of extreme local substrate heterogeneity (Joyner and Lindow, 2000; Leveau and Lindow, 2001; Herron et al., 2010; Farhan Ul Haque et al., 2013; Parangan-Smith and Lindow, 2013). Despite frequently being more difficult to genetically engineer than typical laboratory model bacteria, specific bioreporter variants of strains intended for bioaugmentation will thus be a major asset to better assess the particular conditions for suitable growth and activity of the cells in contaminated sites. Acknowledgements This work was supported by Grant KBBE-211684 from European Commission within the FP7 Framework Programme. The authors thank Mr. Enga Luye for samples of the Jonction material and Dirk Springael for samples of Ter Munck agricultural soil. Vladimir Sentchilo is thanked for his help in the competition experiments.
References Armengaud, J., Happe, B., and Timmis, K.N. (1998) Genetic analysis of dioxin dioxygenase of Sphingomonas sp. strain
RW1: catabolic genes dispersed on the genome. J Bacteriol 180: 3954–3966. Balkwill, D.L., and Ghiorse, W.C. (1985) Characterization of subsurface bacteria associated with two shallow aquifers in Oklahoma. Appl Environ Microbiol 50: 580–588. Bosma, T.N.P., Middeldorp, P.J.M., Schraa, G., and Zehnder, A.J.B. (1997) Mass transfer limitation of biotransformation: quantifying bioavailability. Environ Sci Technol 31: 248– 252. Bunz, P.V., and Cook, A.M. (1993) Dibenzofuran 4,4adioxygenase from Sphingomonas sp. strain RW1: angular dioxygenation by a three-component enzyme system. J Bacteriol 175: 6467–6475. Chen, J., Wong, M.H., Wong, Y.S., and Tam, N.F. (2008) Multi-factors on biodegradation kinetics of polycyclic aromatic hydrocarbons (PAHs) by Sphingomonas sp. a bacterial strain isolated from mangrove sediment. Mar Poll Bull 57: 695–702. Christensen, B.B., Haagensen, J.A., Heydorn, A., and Molin, S. (2002) Metabolic commensalism and competition in a two-species microbial consortium. Appl Environ Microbiol 68: 2495–2502. Coppotelli, B.M., Ibarrolaza, A., Dias, R.L., Del Panno, M.T., Berthe-Corti, L., and Morelli, I.S. (2010) Study of the degradation activity and the strategies to promote the bioavailability of phenanthrene by Sphingomonas paucimobilis strain 20006FA. Microb Ecol 59: 266– 276. Coronado, E., Roggo, C., Johnson, D.R., and van der Meer, J.R. (2012) Genome-wide analysis of salicylate and dibenzofuran metabolism in Sphingomonas wittichii RW1. Front Microbiol 3: 300. El Fantroussi, S., and Agathos, S.N. (2005) Is bioaugmentation a feasible strategy for pollutant removal and site remediation? Curr Opin Microbiol 8: 268–275. Farhan Ul Haque, M., Nadalig, T., Bringel, F., Schaller, H., and Vuilleumier, S. (2013) Fluorescence-based bacterial bioreporter for specific detection of methyl halide emissions in the environment. Appl Environ Microbiol 79: 6561– 6567. Fida, T.T., Breugelmans, P., Lavigne, R., Coronado, E., Johnson, D.R., van der Meer, J.R., et al. (2012) Exposure to solute stress affects genome-wide expression but not the polycyclic aromatic hydrocarbon-degrading activity of Sphingomonas sp. strain LH128 in biofilms. Appl Environ Microbiol 78: 8311–8320. Fida, T.T., Moreno-Forero, S.K., Heipieper, H.J., and Springael, D. (2013) Physiology and transcriptome of the polycyclic aromatic hydrocarbon-degrading Sphingomonas sp. LH128 after long-term starvation. Microbiology 159: 1807–1817. Fida, T.T., Breugelmans, P., Lavigne, R., van der Meer, J.R., De Mot, R., Vaysse, P.J., and Springael, D. (2014) Identification of opsA, a gene involved in solute stress mitigation and survival in soil, in the polycyclic aromatic hydrocarbondegrading bacterium Novosphingobium sp. strain LH128. Appl Environ Microbiol 80: 3350–3361. van Gestel, C.A., McGrath, S.P., Smolders, E., Ortiz, M.D., Borgman, E., Verweij, R.A., et al. (2012) Effect of long-term equilibration on the toxicity of molybdenum to soil organisms. Environ Pollut 162: 1–7.
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 7, 480–488
Dibenzofuran detection by S. wittichii RW1 Goldstein, R.M., Mallory, L.M., and Alexander, M. (1985) Reasons for possible failure of inoculation to enhance biodegradation. Appl Environ Microbiol 50: 977–983. Gunasekera, T.S., Striebich, R.C., Mueller, S.S., Strobel, E.M., and Ruiz, O.N. (2013) Transcriptional profiling suggests that multiple metabolic adaptations are required for effective proliferation of Pseudomonas aeruginosa in jet fuel. Environ Sci Technol 47: 13449–13458. Halden, R.U., Halden, B.G., and Dwyer, D.F. (1999) Removal of dibenzofuran, dibenzo-p-dioxin, and 2-chlorodibenzo-pdioxin from soils inoculated with Sphingomonas sp. strain RW1. Appl Environ Microbiol 65: 2246–2249. Harms, H. (1998) Bioavailability of dioxin-like compounds for microbial degradation. In Biodegradation of dioxins and furans. Wittich, R. (ed.). Austin, TX, USA: Springer-Verlag and R.G. Landes Company, pp. 135–163. Hartmann, E.M., and Armengaud, J. (2014) Shotgun proteomics suggests involvement of additional enzymes in dioxin degradation by Sphingomonas wittichii RW1. Environ Microbiol 16: 162–176. Heeb, S., Itoh, Y., Nishijyo, T., Schnider, U., Keel, C., Wade, J., and Haas, D. (2000) Small, stable shuttle vectors based on the minimal pVS1 replicon for use in gram-negative, plant-associated bacteria. Mol Plant Microbe Interact 13: 232–237. Herron, P.M., Gage, D.J., and Cardon, Z.G. (2010) Microscale water potential gradients visualized in soil around plant root tips using microbiosensors. Plant Cell Environ 33: 199–210. Iino, T., Wang, Y., Miyauchi, K., Kasai, D., Masai, E., Fujii, T., et al. (2012) Specific gene responses of Rhodococcus jostii RHA1 during growth in soil. Appl Environ Microbiol 78: 6954–6962. Jaspers, M.C., Schmid, A., Sturme, M.H., Goslings, D.A., Kohler, H.P., and Van der Meer, J.R. (2001) Transcriptional organization and dynamic expression of the hbpCAD genes, which encode the first three enzymes for 2hydroxybiphenyl degradation in Pseudomonas azelaica HBP1. J Bacteriol 183: 270–279. Johnsen, A.R., and Karlson, U. (2004) Evaluation of bacterial strategies to promote the bioavailability of polycyclic aromatic hydrocarbons. Appl Microbiol Biotechnol 63: 452– 459. Joyner, D.C., and Lindow, S.E. (2000) Heterogeneity of iron bioavailability on plants assessed with a whole- cell GFPbased bacterial biosensor. Microbiology 146: 2435–2445. Leveau, J.H.J., and Lindow, S.E. (2001) Appetite of an epiphyte: quantitative monitoring of bacterial sugar consumption in the phyllosphere. Proc Natl Acad Sci USA 98: 3446– 3453. Leveau, J.H.J., and Lindow, S.E. (2002) Bioreporters in microbial ecology. Curr Opin Microbiol 5: 259–265. Leveau, J.H.J., König, F., Füchslin, H.-P., Werlen, C., and van der Meer, J.R. (1999) Dynamics of multigene expression during catabolic adaptation of Ralstonia eutropha JMP134 (pJP4) to the herbicide 2, 4-dichlorophenoxyacetate. Mol Microbiol 33: 396–406. Liu, X., Germaine, K.J., Ryan, D., and Dowling, D.N. (2010) Whole-cell fluorescent biosensors for bioavailability and biodegradation of polychlorinated biphenyls. Sensors (Basel) 10: 1377–1398.
487
McGenity, T.J., Folwell, B.D., McKew, B.A., and Sanni, G.O. (2012) Marine crude-oil biodegradation: a central role for interspecies interactions. Aquat Biosystems 8: 10. Maphosa, F., Lieten, S.H., Dinkla, I., Stams, A.J., Smidt, H., and Fennell, D.E. (2012) Ecogenomics of microbial communities in bioremediation of chlorinated contaminated sites. Front Microbiol 3: 351. Megharaj, M., Wittich, R.-M., Blasco, R., Pieper, D.H., and Timmis, K.N. (1997) Superior survival and degradation of dibenzo-p-dioxin and dibenzofuran in soil by soil-adapted Sphingomonas sp. strain RW1. Appl Microbiol Biotechnol 48: 109–114. Moreno-Forero, S.K., and van der Meer, J.R. (2015) Genome-wide analysis of Sphingomonas wittichii RW1 behaviour during inoculation and growth in contaminated sand. ISME J 9: 150–165. doi:10.1038/ismej.2014.101. Nam, I.H., Hong, H.B., Kim, Y.M., Kim, B.H., Murugesan, K., and Chang, Y.S. (2005) Biological removal of polychlorinated dibenzo-p-dioxins from incinerator fly ash by Sphingomonas wittichii RW1. Water Res 39: 4651–4660. Parangan-Smith, A., and Lindow, S. (2013) Contribution of nitrate assimilation to the fitness of Pseudomonas syringae pv. syringae B728a on plants. Appl Environ Microbiol 79: 678–687. Pritchard, R.P. (1992) Use of inoculation in bioremediation. Curr Opin Biotechnol 3: 232–243. Reinhard, F., and van der Meer, J.R. (2013) Improved statistical analysis of low abundance phenomena in bimodal bacterial populations. PLoS ONE 8: e78288. Roggo, C., Coronado, E., Moreno-Forero, S.K., Harshman, K., Weber, J., and van der Meer, J.R. (2013) Genome-wide transposon insertion scanning of environmental survival functions in the polycyclic aromatic hydrocarbon degrading bacterium Sphingomonas wittichii RW1. Environ Microbiol 15: 2681–2695. Safe, S. (1990) Polychlorinated biphenyls (PCBs), dibenzop-dioxins (PCDDs), dibenzofurans (PCDFs), and related compounds: environmental and mechanistic considerations which support the development of toxic equivalency factors (TEFs). Crit Rev Toxicol 21: 51–88. Scheublin, T.R., Deusch, S., Moreno-Forero, S.K., Müller, J.A., van der Meer, J.R., and Leveau, J.H. (2014) Transcriptional profiling of Gram-positive Arthrobacter in the phyllosphere: induction of pollutant degradation genes by natural plant phenolic compounds. Environ Microbiol 16: 2212–2225. Schraa, G., Boone, M.L., Jetten, M.S., van Neerven, A.R., Colberg, P.J., and Zehnder, A.J. (1986) Degradation of 1,4-dichlorobenzene by Alcaligenes sp. strain A175. Appl Environ Microbiol 52: 1374–1381. Semple, K.T., Doick, K.J., Wick, L.Y., and Harms, H. (2007) Microbial interactions with organic contaminants in soil: definitions, processes and measurement. Environ Poll 150: 166–176. Shi, T., Fredrickson, J.K., and Balkwill, D.L. (2001) Biodegradation of polycyclic aromatic hydrocarbons by Sphingomonas strains isolated from the terrestrial subsurface. J Ind Microbiol Biotechnol 26: 283–289. Sikkema, J., de Bont, J.A., and Poolman, B. (1995) Mechanisms of membrane toxicity of hydrocarbons. Microbiol Rev 59: 201–222.
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 7, 480–488
488 E. Coronado, A. Valtat and J. R. van der Meer Tecon, R., and van der Meer, J.R. (2006) Information from single-cell bacterial biosensors: what is it good for? Curr Opin Biotechnol 17: 4–10. Thompson, I.P., van der Gast, C.J., Ciric, L., and Singer, A.C. (2005) Bioaugmentation for bioremediation: the challenge of strain selection. Environ Microbiol 7: 909–915. Tyagi, M., da Fonseca, M.M., and de Carvalho, C.C. (2011) Bioaugmentation and biostimulation strategies to improve the effectiveness of bioremediation processes. Biodegradation 22: 231–241. van Veen, J.A., van Overbeek, L.S., and van Elsas, J.D. (1997) Fate and activity of microorganisms introduced into soil. Microbiol Mol Biol Rev 61: 121–135. Verschuere, L., Rombaut, G., Sorgeloos, P., and Verstraete, W. (2000) Probiotic bacteria as biological control agents in aquaculture. Microbiol Mol Biol Rev 64: 655–671. Wilkes, H., Wittich, R., Timmis, K.N., Fortnagel, P., and Francke, W. (1996) Degradation of chlorinated dibenzofurans and dibenzo-p-dioxins by Sphingomonas sp. strain RW1. Appl Environ Microbiol 62: 367–371. Wittich, R.-M., Wilkes, H., Sinnwell, V., Francke, W., and Fortnagel, P. (1992) Metabolism of dibenzo-p-dioxin by Sphingomonas sp. strain RW1. Appl Environ Microbiol 58: 1005–1010.
Supporting information Additional Supporting Information may be found in the online version of this article at the publisher’s web-site: Fig. S1. Metabolic network for DBF and aromatic compound metabolism by S. wittichii RW1, displayed using CYTOSKAPE 2.8.3, as inferred for cells growing on DBF (Moreno-Forero and van der Meer, 2014). Nodes represent substrates and metabolic intermediates. Edges represent enzyme reactions converting the linked compounds. Edge line thickness is a representation of normalized expression of the gene coding for the particular enzyme that carries out the reaction between two nodes. Normalization was carried out per gene among six conditions, taking the highest expression value as 100% (=line width 15). Line widths are a linear scale representation of normalized gene expression, notation and the predicted gene function. For compound abbreviations, see Moreno-Forero and van der Meer (2014). Fig. S2. Stability of the pPROBE-PTac-egfp (A), pME6012PTac-egfp (B) and miniTn5::PTac-egfp (C) constructs in Sphingomonas wittichii RW1. Shown is marker stability over
four subsequent culture transfers (∼ 28 generations) in liquid medium with SAL as sole carbon source without (no AB) or with (AB) addition of antibiotic selecting for the plasmid or transposon insertion. Proportions show the average number of antibiotic resistant colonies (Resist) divided by the number of colonies on non-selective media, or the number of fluorescent colonies amidst all resistant colonies (R/Fluo), ± calculated standard deviation from independent triplicates. For pPROBE and mini-Tn5, Km was used as antibiotic, whereas Tc was used for pME6012. Fig. S3. EGFP production rate as a function of growth rate of RW1 carrying pME6012-P4925-egfp, pME6012-PdxnA1egfp or mini-Tn5-PuspA-egfp. Rates were calculated from linear regression of ln-transformed normalized EGFP intensities or culture turbidity values versus time. Decreasing exponential growth rate in liquid culture was achieved by successive addition of NaCl (open symbols) or polyethylene glycol (closed symbols), which lowers the medium water potential (Johnson et al., 2011). Fig. S4. Population development and reporter gene expression of RW1 (pME6012-PPdxnA1-egfp) inoculated in sandy microcosms S + DBF (red symbols) and S + TM (green symbols). (A) Colony-forming units on MM + SAL + Tc plates (cfu g−1 soil) over time, representative for RW1. (B) Mean EGFP expression from triplicate series ± calculated 95% confidence interval. Note that the RW1 population for calculating EGFP values is much smaller in S + TM than in S + DBF. Fig. S5. EGFP expression density plots for bacterial cells extracted from the sandy microcosms at the different time points. All plots scaled to the same EGFP intensity level (x-axis). Fig. S6. Competition for DBF intermediates. (A) Population development of RW1 (pME6012-P4925-egfp) alone (open symbols) or in co-culture (closed symbols) with a mixture of three meta-toluate degrading strains isolated from Jonction in liquid medium with DBF as sole carbon source (arrow indicates the time of their addition). (B) Average EGFP levels of individual RW1 reporter cells under both conditions measured by flow cytometry. Data points are averages from independent triplicates ± calculated standard deviations. Table S1. Strains used in this study. Table S2. Primers used in this study. Table S3. Variability of EGFP expression among individual RW1 (pME6012-P4925-egfp) cells under different growth conditions. Appendix S1. Supplementary experimental methods. Appendix S2. Supplementary data.
© 2015 The Authors. Environmental Microbiology Reports published by Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 7, 480–488
