bs_bs_banner
Environmental Microbiology Reports (2014) 6(6), 786–791
doi:10.1111/1758-2229.12211
Spatially tracking 13C-labelled substrate (bicarbonate) accumulation in microbial communities using laser ablation isotope ratio mass spectrometry
James J. Moran,1* Charles G. Doll,1 Hans C. Bernstein,1 Ryan S. Renslow,2 Alexandra B. Cory,3† Janine R. Hutchison,4 Stephen R. Lindemann4 and James K. Fredrickson4 1 Signatures Science and Technology Division, National Security Directorate, Pacific Northwest National Laboratory, Richland, WA 99352, USA. 2 Scientific Resources Division, William R. Wiley Environmental Molecular Sciences Laboratory, Pacific Northwest National Laboratory, Richland, WA 99352, USA. 3 Geology Department, Lawrence University, Appleton, WI 54911, USA. 4 Biological Sciences Division, Fundamental and Computational Sciences Directorate, Pacific Northwest National Laboratory, Richland, WA USA. Summary Microbial mats are characterized by extensive metabolic interactions, rapidly changing internal geochemical gradients, and prevalent microenvironments within tightly constrained physical structures. We present laser ablation isotope ratio mass spectrometry (LA-IRMS) as a culture-independent, spatially specific technology for tracking the accumulation of 13 C-labelled substrate into heterogeneous microbial mat communities. This study demonstrates the novel LA-IRMS approach by tracking labeled bicarbonate incorporation into a cyanobacteria-dominated microbial mat system. The spatial resolution of 50 μm was sufficient for distinguishing different mat strata and the approach effectively identified regions of greatest label incorporation. Sample preparation for LA-IRMS is straightforward and the spatial selectivity of LA-IRMS minimizes the volume of mat consumed, leaving material for complimentary analyses. We present analysis of DNA extracted from a sample post-
Received 10 August, 2013; accepted 5 August, 2014. *For correspondence. E-mail [email protected]; Tel. 509 371 6798; Fax 509 375 2227. †Present address: Signatures Science and Technology Division, National Security Directorate, Pacific Northwest National Laboratory, Richland, WA 99352, USA.
© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd
ablation and suggest pigments, lipids or other biomarkers could similarly be extracted following ablation. LA-IRMS is well positioned to spatially resolve the accumulation of any 13C-labelled substrate provided to a mat, making this a versatile tool for studying carbon transfer and interspecies exchanges within the limited spatial confines of such systems. Introduction Microbial mats are characterized by tightly spaced, metabolically interactive microbial populations that frequently show functional stratification along geochemical microgradients existing through the mat structure (Visscher and Stolz, 2005; Dupraz et al., 2009; Franks and Stolz, 2009; Bernstein et al., 2013; Klatt et al., 2013). The microgradients themselves can result from microbial metabolic activity and are commonly transient in nature, displaying large diel or seasonal shifts resulting from changing dominant metabolisms (Canfield and Des Marais, 1993; Des Marais, 2003; van der Meer et al., 2005; Villanueva et al., 2007). The combination of extensive interspecies carbon exchange, generally high metabolic activities, and dynamic biogeochemical processes within the spatial confines of mat strata make understanding microbial interactions at the sub-mat level challenging. Isotope analysis can elucidate nutrient transfers and interactions within these complex systems and some techniques provide varying degrees of spatial resolution. Microradiography can be used to spatially track the location of radioisotope-labelled substrate accumulation and dissemination into a microbial mat (Ginige et al., 2004). Secondary ion mass spectrometry (SIMS) and fluorescence in situ hybridization SIMS provide spatially specific isotope measurements coupled to phylogenetic markers (Orphan et al., 2001; Behrens et al., 2008). Physical subsampling of microbial mat sections followed by traditional stable isotope analysis can enable some degree of spatial resolution (Kelley et al., 2006). Yet, innate challenges such as requiring a radioactive label (microradiography), low sample throughput/high analysis time (SIMS), low instrument availability/access (SIMS) or extensive sample preparation (physical subsampling)
787 J. J. Moran et al. limit application of many of the above methods. In contrast, laser ablation isotope ratio mass spectrometry (LAIRMS) provides spatially resolved isotope analysis with minimal sample preparation and high measurement throughput. Here we provide the first documented demonstration of laser ablation sampling coupled to stable isotope analysis via LA-IRMS for spatially specific carbon (13C) isotope analysis of a microbial mat. The spatial resolution demonstrated here is approximately 50 μm, significantly less than visually observed lamina in the microbial mats. We used isotopically labelled (13C) bicarbonate to track accumulation in a cyanobacteria-dominated mat under ambient light conditions. Coupling LA-IRMS analysis with detailed transport and reaction kinetics for a particular system will enable spatially resolved interpretations of phenomena such as microbially induced carbon assimilation rates. While this represents a future application, the primary goal of the current study is to demonstrate the spatial resolution and accuracy of the LA-IRMS method. This technique consumes only a small portion of a microbial mat sample, is highly amenable to examination of other isotopically, carbon-labelled substrates and is therefore a useful and novel technique for spatially resolving metabolic interactions within microbial systems.
In the laboratory, we used a razor blade to manually cross-section thin (1–2 mm) portions of the frozen mat for LA-IRMS analysis. We paid careful attention to avoid compression of the sample and maintain the mat’s native stratified structure, and determined that cross-sectioning of frozen samples showed improved structural preservation as compared with subsectioning thawed samples. Therefore, LA-IRMS was limited to samples sectioned while frozen. Cross-sections were lyophilized prior to isotope analysis as significant water would interfere with both the ablation process and the isotope analysis. LA-IRMS followed the methods of Moran and colleagues (2011). The ablation system (Cetac LSX 500, Omaha, NE, USA) allowed for visual imaging, magnification and targeting with high spatial resolution (50 μm). This analysis was destructive in the sense that material correlating to the ablation target was removed (Fig. 1). However, the ablation pits were minimal in size (< 3.3 × 10−7 ml based on visual estimation) and the remainder of the sample was physically intact and suitable for any additional downstream analysis compatible with a lyophilized sample. As an example, we used a post-ablated sample for DNA extraction and sequencing. Previous demonstration of a similar laser technique on organic polymers (Moran et al., 2011) demonstrated only negligible production of volatile
Results and discussion We sampled a cyanobacteria-dominated microbial mat from Hot Lake, an epsomitic, hypersaline lake in northcentral Washington state (Anderson, 1958; Lindemann et al., 2013). When sampled on 25 July 2012, the seasonal mat had developed visually distinct strata over its approximately 5 mm thickness. The mat overlays a gypsum-dominated sediment that also contains minor amounts (< 10%) of carbonate (dominated by aragonite, data not shown). Ex vivo incubations of replicate mat samples excised from 55 cm water depth were incubated in lake water sampled concurrently with the mat, but modified by the addition of 2.25 mmoles of 50% 13C bicarbonate in a 740 ml glass incubation container. A clear acrylic sheet (1/8 inch thickness) allowed ambient light penetration. Control containers were wrapped in aluminium foil to exclude light. The experimental incubations quickly showed signs of phototrophic activity, producing visual bubbles (presumably O2) after ∼ 15 min of incubation under bright light conditions (near solar noon). We subsampled the mat at 0.5, 1.0 and 2.5 h after starting the incubation. Immediately following subsectioning, samples were rinsed in lake water devoid of labelled bicarbonate for 30 min to ensure the removal of residual label. We performed these rinses in the dark, and immediately after rinsing, samples were stored on dry ice for storage and transportation back to the laboratory.
Fig. 1. Ablated mat cross-section. In preparation for analysis, mat was cross-sectioned with a razor blade then lyophilized prior to mounting in the laser ablation chamber. As seen here, this method retained the internal mat stratification and, going from the surface down, the stratified, orange, green, white, brown and pink layers are visibly distinguished. Laser ablation was performed along three tracks perpendicular to the mat surface, each track acted as replicate profile measurements of the mat sample. Individual spots in each profile represent a 50 μm sample from the mat and contained ample material for isotope ratio determination. Digital microscopy and optics within the laser ablation unit (Cetac LSX-500, Omaha, NE, USA) permitted precise selection and targeting of mat for ablation.
© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 6, 786–791
Spatial tracking of substrate into microbial mat
788
Fig. 2. Label incorporation into the mat as a function of time. We used an elemental analyser (ECS 4010 CHNSO Analyser, Costech Analytical, Valencia, CA, USA) coupled to IRMS (Delta V plus, Thermo Scientific, Bremen, Germany) to measure bulk 13C content of homogenized mat sections. These bulk results showed increasing label incorporation over time in both the total (A) and acid-treated (organic) fractions (B) of the mat. In both cases, we homogenized a ∼ 4 cm2 section of mat taken from immediately adjacent to the laser ablation sample. We compared experimental samples (square) to the dark control (diamond, samples incubated with label in the dark) and error bars are the standard deviation of analytical replicates (n ≥ 3). LA-IRMS results of a mat cross-section (C) likewise show increasing label incorporation over time but more specifically also show a clustering of label towards the top of the mat. We show (C) all LA-IRMS measurement resulting from each of the three tracks through the mat. The mat displayed variable overall thickness within a single sample due to its pinnacled upper surface. In order to compare adjoining sections of mat with different thicknesses, we normalized cross-sectional depth based on proportional thickness into the mat (with 0.0 being the upper and 1.0 being the lower mat surface). The dark control was incubated with the 13C label but kept dark for 2.5 h then harvested. We used the trapezoidal numerical method to integrate the area between an experimental isotope profile for each timepoint and the dark control (with a representative area for the 1.0 h incubation shaded in blue). While mat cross-sections varied between samples, we denoted representative boundaries between the visually distinguished mat strata (orange, green, white, brown and pink).
organic carbon during the ablation (on the order of 1.8 ppb in the carrier gas during ablation). This suggests that thermal decomposition surrounding the ablation pit is minimal and that the sample immediately adjacent to the ablation is preserved for other, complimentary analyses which may include DNA (as demonstrated here), pigment, lipid or other biomarker analyses. We observed increasing 13C accumulation over time in both the total bulk mat (Fig. 2A) and in samples that had been acid washed to remove carbonate material, leaving only organic carbon behind (Fig. 2B). The dark control samples, which we incubated in the presence of the 13C label, did not show significant label accumulation. These data suggest both that the dark conditions prohibited carbon accumulation on this timescale and that a 30 min post-incubation rinse in lake water was effective at removing unincorporated bicarbonate. The combination of advective flow (through microchannels) and diffusive transport control the rate at which the applied labelled bicarbonate moves into the mat and is available for accumulation across the depth profile. Relatively slow transport rates into the mat would favour label accumulation at the mat surfaces while disfavouring accumulation in the mat interior. Our dark control showed that 30 min of rinsing in label-free Hot Lake water enabled sufficient
porewater exchange to remove all label from the sample even after a 2.5 h incubation. The laser ablation data (Fig. 2C) reflect both the increasing label accumulation with time and lack of label uptake in the dark control observed in the bulk analysis data (Fig. 2A and B). The LA-IRMS data presented in Fig. 2C are a compilation of data from each of the three parallel profiles sampled through the mat. Subsequent data analysis considers each of these three profiles independently and is discussed below. In addition to the elemental analysis isotope ratio mass spectrometry (EAIRMS) interpretations, however, the LA-IRMS produces a quantitative profile of label incorporation through the mat cross-section. Incorporation was greatest near the mat surface, with comparatively small amounts at the bottom of the mat. Total label incorporation was estimated by first integrating each profile across relative mat depth (trapezoidal numerical method) and then subtracting the average control (dark) incubation area from each of the respected illuminated areas (schematically depicted for the 1.0 h incubation by the shaded areas in Fig. 2C). Each of the three replicate laser ablation profiles corresponding to an incubation treatment were individually analysed. The total label accumulation calculated from integrating the LA-IRMS profiles through the entire mat thickness was in
© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 6, 786–791
789 J. J. Moran et al.
Fig. 3. Bulk and spatially specific label accumulation and quantification of total label as a function of depth. Comparison between bulk (EA-IRMS) and spatially resolved (LA-IRMS) data (A) shows reasonable agreement in the amount of label associated with each sample at the whole mat scale. Values reported for the bulk mat reflect the increase in isotopic content of the experimental versus control samples with error reported as the sum of squares for the experimental and associated control analytical replicate standard deviations. For the spatially resolved approach, we integrated the area between experimental and control data points from the mat’s upper to lower surface. The spatially resolved data are average difference between the integration of each of the replicate tracks through the 2.5 h incubation mat and the average control data with error reported following sum of squares error propagation of the standard deviation of the experimental and control replicates. Integration of LA-IRMS depth profile restricted to specific depths in the mat (B) enables estimation of where, physically, in the mat, the label accumulation occurs. We report both the relative amount of label accumulation observed at different depths within the mat (outlined squares) and the relative summed accumulation as integrated from the surface to different depths in the mat (black diamonds). Error bars depict either the standard deviation of replicate tracks through the mat (summed accumulation) or sum of squares of the standard deviation of the accumulation through different depths (strata accumulation). Transitions between visually distinguished mat strata (orange, yellow, white, brown and pink) are provided at representative depths in the mat.
close agreement with that measured by bulk EA-IRMS analysis for each of the three timepoints (Fig. 3A). In addition to quantifying bulk label accumulation, the LA-IRMS data reveal the extent of label incorporation along the depth profile of the mat. For example, in the 2.5 h incubation, 75% of the total label accumulation
occurred in the upper half of the mat (Fig. 3B) and the upper 20% of the mat accounted for 48% of the label accumulation. The integrated vertical distribution of label incorporation within the mat was nearly identical through each of the three timepoints (data not shown). Previous work on mat from this location (Lindemann et al., 2013) showed photosynthetically active radiation being attenuated to below detection by ∼ 1.5 mm or within the top 30% of the mat during peak daylight. In our experiment, we demonstrate that this region showed the highest rate of label accumulation (> 60% of total accumulation in the upper ∼ 30% of the mat) but that there was label accumulation beneath. Presumably, this deeper label accumulation results from either transportation of fixed carbon to great depth in the mat or autotrophic processes other than photosynthesis. To characterize the community composition and likely metabolisms within the mat, we sectioned post-ablated samples into five pieces corresponding to the coloured zones observed in Fig. 1: orange, green, white, brown and pink (see Fig. 1). We extracted genomic DNA from these subsections and performed pyrosequencing of the 16S rRNA gene on a 454 GSFLX platform, amplified using the V4 primers 515F and 806R (Caporaso et al., 2011). Sequences were analysed in MOTHER v. 1.31 using the 454 SOP [Schloss et al., 2009) (http://www.mothur.org/ wiki/454_SOP (accessed 24 July 2013)] and clustered using an average neighbour algorithm at 3% distance. These clusters were classified using a Wang approach with the Ribosome Database Project training set v. 9 (updated 20 March 2012 and formatted for MOTHER). Areas of highest label incorporation exhibited significant cyanobacterial relative abundance, although we also measured relatively high cyanobacterial amplicon abundance in strata (including the white, green and brown layers) that incorporated only small amounts of label (Fig. 4). This observation highlights the ability of the LA-IRMS approach to localize actual label accumulation as opposed to metabolic inferences derived solely from community composition. While not measured in this study, previous work on the same microbial mat systems (Lindemann et al., 2013) describe irradiance in the mat with very similar profiles to the accumulation trends we observed. This coupled with the minimal label accumulation in the dark controls suggest that light availability exerts strong influence on label accumulation in this system. The results presented here highlight the ability of LA-IRMS to spatially pinpoint locations within a microbial mat where labelled substrate accumulates. While this specific LA-IRMS system was previously demonstrated on organic polymers (Moran et al., 2011) and a similar system was demonstrated on plant material (Wieser and Brand, 1999; Schulze et al., 2004; Skomarkova et al.,
© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 6, 786–791
Spatial tracking of substrate into microbial mat
790
Fig. 4. Community composition of differently colored mat layers. We sectioned the mat along color stratifications (Fig. 1) and extracted genomic DNA. Mat samples were washed (500 mM EDTA, 10 min), homogenized (in 700 μl of 50 mM Tris pH 8.0 + 25 mM EDTA), then heated (5 min at 85°C) prior to cell lysis (lysozyme 30 mg ml−1 for 1 h at 37°C followed by an overnight incubation at 56°C with 10% SDS and 0.2 mg ml−1 Proteinase K). We used a phenol chloroform extraction followed by ethanol (70%) precipitation. We removed PCR inhibitors using a Zymo Research OneStep PCR Inhibitor Removal Kit following the manufacturer’s instructions. Pyrosequencing was performed on a 454 GSFLX platform by Research and Testing Laboratories (Lubbock, TX, USA) and we processed sequences and taxonomy as previously described (Schloss et al., 2011). The relative abundances, based upon a standardized number of sequences (n = 3452 per layer), of major taxonomic orders within the mat are displayed above. Classes of phylum Proteobacteria are marked with brackets around their composite orders. ‘Unclassified’ refers to sequences from kingdom Bacteria that could not be classified at the phylum level, while ‘Other’ refers to reads classified within orders not otherwise depicted. Orders containing sequences from genera known to be autotrophic are in green.
2006), this is, to the best of our knowledge, the first demonstration of the LA-IRMS technique on a microbial mat system. Our results demonstrate that, under high light intensity, the highest rates of label accumulation occur near the mat surface. The utility of LA-IRMS, however, goes beyond this specific example. Tracking the accumulation of specific labelled compounds using LA-IRMS can not only demonstrate where substrates are consumed within a mat but also how the location of these activities changes over diel or seasonal cycles. Sample preparation for analysis is straightforward and can be performed in most laboratories, requiring only sectioning and lyophilization of a suitable mat sample. Adaptation of these preparation methods to enable drying and mounting a sample for analysis could also enable LA-IRMS probing of any microbial system where spatial resolution is important. Potential examples include microbial communities associated with the plant rhizosphere, soil aggregates, surface biofilms and others. The spatial resolution afforded by LA-IRMS allows separate isotope measurement of microbial biomass and associated growth matrices, whether that is organic (plant roots) or inorganic (soil minerals or some biofilm substrates). Finally, the LA-IRMS system has a high sample throughput for carbon-rich samples such as microbial mats. The δ13C measurement of a spot in our mat sample was completed in less than 10 min. Overall, we believe that due to the ease of sample preparation, microbially relevant spatial resolution, and high sample throughput, the LA-IRMS technique fills an analytical niche for culture-independent assessment of metabolic activity in microbial community.
Acknowledgements This research was supported by the Genomic Science Program (GSP), Office of Biological and Environmental Research (OBER), U.S. Department of Energy (DOE) and is a contribution of the Pacific Northwest National Laboratory (PNNL) Foundational Scientific Focus Area. Alexandra B. Cory was supported by the Lawrence University (Appleton, WI) LUR1 program. A portion of the research described in this paper was conducted under the Laboratory Directed Research and Development Program at Pacific Northwest National Laboratory, a multiprogramme national laboratory operated by Battelle for the U.S. Department of Energy. Hans C. Bernstein and Ryan S. Renslow are grateful for the support of the Linus Pauling Distinguished Postdoctoral Fellowship program. We thank William Chrisler for his assistance in microscope sample photography and Dr. Helen Kreuzer for insightful discussions regarding data interpretation. Conflict of interest: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
References Anderson, G.C. (1958) Some limnological features of a shallow saline meromictic lake. Limnol Oceanogr 3: 259– 270. Behrens, S., Losekann, T., Pett-Ridge, J., Weber, P.K., Ng, W.-O., Stevenson, B.S., et al. (2008) Linking microbial phylogeny to metabolic activity at the single-cell level by using enhanced element labeling-catalyzed reporter deposition fluorescence in situ hybridization (EL-FISH) and NanoSIMS. Appl Environ Microbiol 74: 3143–3150.
© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 6, 786–791
791 J. J. Moran et al. Bernstein, H.C., Beam, J.P., Kozubal, M.A., Carlson, R.P., and Inskeep, W.P. (2013) In situ analysis of oxygen consumption and diffusive transport in high-temperature acidic iron-oxide microbial mats: oxygen consumption in geothermal iron-oxide microbial mats. Environ Microbiol 15: 2360– 2370. Canfield, D.E., and Des Marais, D.J. (1993) Biogeochemical cycles of carbon, sulfur, and free oxygen in a microbial mat. Geochim Cosmochim Acta 57: 3971–3984. Caporaso, J.G., Lauber, C.L., Walters, W.A., Berg-Lyons, D., Lozupone, C.A., Turnbaugh, P.J., et al. (2011) Global patterns of 16S rRNA diversity at a depth of millions of sequences per sample. Proc Natl Acad Sci USA 108: 4516–4522. Des Marais, D.J. (2003) Biogeochemistry of hypersaline microbial mats illustrates the dynamics of modern microbial ecosystems and the early evolution of the biosphere. Biol Bull 204: 160–167. Dupraz, C., Reid, R.P., Braissant, O., Decho, A.W., Norman, R.S., and Visscher, P.T. (2009) Processes of carbonate precipitation in modern microbial mats. Earth Sci Rev 96: 141–162. Franks, J., and Stolz, J.F. (2009) Flat laminated microbial mat communities. Earth Sci Rev 96: 163–172. Ginige, M.P., Hugenholtz, P., Daims, H., Wagner, M., Keller, J., and Blackall, L.L. (2004) Use of stable-isotope probing, full-cycle rRNA analysis, and fluorescence in situ hybridization-microautoradiography to study a methanolfed denitrifying microbial community. Appl Environ Microbiol 70: 588–596. Kelley, C.A., Prufert-Bebout, L.E., and Bebout, B.M. (2006) Changes in carbon cycling ascertained by stable isotopic analyses in a hypersaline microbial mat. J Geophys Res 111: G04012. Klatt, C.G., Inskeep, W.P., Herrgard, M.J., Jay, Z.J., Rusch, D.B., Tringe, S.G., et al. (2013) Community structure and function of high-temperature chlorophototrophic microbial mats inhabiting diverse geothermal environments. Front Microbiol 4: 106. Lindemann, S.R., Moran, J.J., Renslow, R.S., Hutchison, J.R., Cole, J.K., Dohnalkova, A.C., et al. (2013) The epsomitic phototrophic microbial mat of Hot Lake, Washington: community structural responses to seasonal cycling. Front Microbiol 4: 323. doi:10.3389/fmicb.2013. 00323.
van der Meer, M.T.J., Schouten, S., Bateson, M.M., Nubel, U., Wieland, A., Kuhl, M., et al. (2005) Diel variations in carbon metabolism by green nonsulfur-like bacteria in alkaline siliceous hot spring microbial mats from Yellowstone National Park. Appl Environ Microbiol 71: 3978–3986. Moran, J.J., Newburn, M.K., Alexander, M.L., Sams, R.L., Kelly, J.F., and Kreuzer, H.W. (2011) Laser ablation isotope ratio mass spectrometry for enhanced sensitivity and spatial resolution in stable isotope analysis. Rapid Commun Mass Spectrom 25: 1282–1290. Orphan, V.J., House, C.H., Hinrichs, K.U., McKeegan, K.D., and DeLong, E.F. (2001) Methane-consuming archaea revealed by directly coupled isotopic and phylogenetic analysis. Science 293: 484–487. Schloss, P.D., Westcott, S.L., Ryabin, T., Hall, J.R., Hartmann, M., Hollister, E.B., et al. (2009) Introducing mothur: open-source, platform-independent, communitysupported software for describing and comparing microbial communities. Appl Environ Microbiol 75: 7537–7541. Schloss, P.D., Gevers, D., and Westcott, S.L. (2011) Reducing the effects of PCR amplification and sequencing artifacts on 16S rRNA-based studies. PLoS ONE 6: e27310. Schulze, B., Wirth, C., Linke, P., Brand, W.A., Kuhlmann, I., Horna, V., and Schulze, E.-D. (2004) Laser ablationcombustion-GC-IRMS–a new method for online analysis of intra-annual variation of delta13C in tree rings. Tree Physiol 24: 1193–1201. Skomarkova, M.V., Vaganov, E.A., Mund, M., Knohl, A., Linke, P., Boerner, A., and Schulze, E.-D. (2006) Interannual and seasonal variability of radial growth, wood density and carbon isotope ratios in tree rings of beech (Fagus sylvatica) growing in Germany and Italy. Trees 20: 571–586. Villanueva, L., Navarrete, A., Urmeneta, J., White, D.C., and Guerrero, R. (2007) Analysis of diurnal and vertical microbial diversity of a hypersaline microbial mat. Arch Microbiol 188: 137–146. Visscher, P.T., and Stolz, J.F. (2005) Microbial mats as bioreactors: populations, processes, and products. Palaeogeogr Palaeoclimatol Palaeoecol 219: 87–100. Wieser, M.E., and Brand, W.A. (1999) A laser extraction/ combustion technique for in situ delta(13)C analysis of organic and inorganic materials. Rapid Commun Mass Spectrom 13: 1218–1225.
© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 6, 786–791
Copyright of Environmental Microbiology Reports is the property of Wiley-Blackwell and its content may not be copied or emailed to multiple sites or posted to a listserv without the copyright holder's express written permission. However, users may print, download, or email articles for individual use.
Environmental Microbiology Reports (2014) 6(6), 786–791
doi:10.1111/1758-2229.12211
Spatially tracking 13C-labelled substrate (bicarbonate) accumulation in microbial communities using laser ablation isotope ratio mass spectrometry
James J. Moran,1* Charles G. Doll,1 Hans C. Bernstein,1 Ryan S. Renslow,2 Alexandra B. Cory,3† Janine R. Hutchison,4 Stephen R. Lindemann4 and James K. Fredrickson4 1 Signatures Science and Technology Division, National Security Directorate, Pacific Northwest National Laboratory, Richland, WA 99352, USA. 2 Scientific Resources Division, William R. Wiley Environmental Molecular Sciences Laboratory, Pacific Northwest National Laboratory, Richland, WA 99352, USA. 3 Geology Department, Lawrence University, Appleton, WI 54911, USA. 4 Biological Sciences Division, Fundamental and Computational Sciences Directorate, Pacific Northwest National Laboratory, Richland, WA USA. Summary Microbial mats are characterized by extensive metabolic interactions, rapidly changing internal geochemical gradients, and prevalent microenvironments within tightly constrained physical structures. We present laser ablation isotope ratio mass spectrometry (LA-IRMS) as a culture-independent, spatially specific technology for tracking the accumulation of 13 C-labelled substrate into heterogeneous microbial mat communities. This study demonstrates the novel LA-IRMS approach by tracking labeled bicarbonate incorporation into a cyanobacteria-dominated microbial mat system. The spatial resolution of 50 μm was sufficient for distinguishing different mat strata and the approach effectively identified regions of greatest label incorporation. Sample preparation for LA-IRMS is straightforward and the spatial selectivity of LA-IRMS minimizes the volume of mat consumed, leaving material for complimentary analyses. We present analysis of DNA extracted from a sample post-
Received 10 August, 2013; accepted 5 August, 2014. *For correspondence. E-mail [email protected]; Tel. 509 371 6798; Fax 509 375 2227. †Present address: Signatures Science and Technology Division, National Security Directorate, Pacific Northwest National Laboratory, Richland, WA 99352, USA.
© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd
ablation and suggest pigments, lipids or other biomarkers could similarly be extracted following ablation. LA-IRMS is well positioned to spatially resolve the accumulation of any 13C-labelled substrate provided to a mat, making this a versatile tool for studying carbon transfer and interspecies exchanges within the limited spatial confines of such systems. Introduction Microbial mats are characterized by tightly spaced, metabolically interactive microbial populations that frequently show functional stratification along geochemical microgradients existing through the mat structure (Visscher and Stolz, 2005; Dupraz et al., 2009; Franks and Stolz, 2009; Bernstein et al., 2013; Klatt et al., 2013). The microgradients themselves can result from microbial metabolic activity and are commonly transient in nature, displaying large diel or seasonal shifts resulting from changing dominant metabolisms (Canfield and Des Marais, 1993; Des Marais, 2003; van der Meer et al., 2005; Villanueva et al., 2007). The combination of extensive interspecies carbon exchange, generally high metabolic activities, and dynamic biogeochemical processes within the spatial confines of mat strata make understanding microbial interactions at the sub-mat level challenging. Isotope analysis can elucidate nutrient transfers and interactions within these complex systems and some techniques provide varying degrees of spatial resolution. Microradiography can be used to spatially track the location of radioisotope-labelled substrate accumulation and dissemination into a microbial mat (Ginige et al., 2004). Secondary ion mass spectrometry (SIMS) and fluorescence in situ hybridization SIMS provide spatially specific isotope measurements coupled to phylogenetic markers (Orphan et al., 2001; Behrens et al., 2008). Physical subsampling of microbial mat sections followed by traditional stable isotope analysis can enable some degree of spatial resolution (Kelley et al., 2006). Yet, innate challenges such as requiring a radioactive label (microradiography), low sample throughput/high analysis time (SIMS), low instrument availability/access (SIMS) or extensive sample preparation (physical subsampling)
787 J. J. Moran et al. limit application of many of the above methods. In contrast, laser ablation isotope ratio mass spectrometry (LAIRMS) provides spatially resolved isotope analysis with minimal sample preparation and high measurement throughput. Here we provide the first documented demonstration of laser ablation sampling coupled to stable isotope analysis via LA-IRMS for spatially specific carbon (13C) isotope analysis of a microbial mat. The spatial resolution demonstrated here is approximately 50 μm, significantly less than visually observed lamina in the microbial mats. We used isotopically labelled (13C) bicarbonate to track accumulation in a cyanobacteria-dominated mat under ambient light conditions. Coupling LA-IRMS analysis with detailed transport and reaction kinetics for a particular system will enable spatially resolved interpretations of phenomena such as microbially induced carbon assimilation rates. While this represents a future application, the primary goal of the current study is to demonstrate the spatial resolution and accuracy of the LA-IRMS method. This technique consumes only a small portion of a microbial mat sample, is highly amenable to examination of other isotopically, carbon-labelled substrates and is therefore a useful and novel technique for spatially resolving metabolic interactions within microbial systems.
In the laboratory, we used a razor blade to manually cross-section thin (1–2 mm) portions of the frozen mat for LA-IRMS analysis. We paid careful attention to avoid compression of the sample and maintain the mat’s native stratified structure, and determined that cross-sectioning of frozen samples showed improved structural preservation as compared with subsectioning thawed samples. Therefore, LA-IRMS was limited to samples sectioned while frozen. Cross-sections were lyophilized prior to isotope analysis as significant water would interfere with both the ablation process and the isotope analysis. LA-IRMS followed the methods of Moran and colleagues (2011). The ablation system (Cetac LSX 500, Omaha, NE, USA) allowed for visual imaging, magnification and targeting with high spatial resolution (50 μm). This analysis was destructive in the sense that material correlating to the ablation target was removed (Fig. 1). However, the ablation pits were minimal in size (< 3.3 × 10−7 ml based on visual estimation) and the remainder of the sample was physically intact and suitable for any additional downstream analysis compatible with a lyophilized sample. As an example, we used a post-ablated sample for DNA extraction and sequencing. Previous demonstration of a similar laser technique on organic polymers (Moran et al., 2011) demonstrated only negligible production of volatile
Results and discussion We sampled a cyanobacteria-dominated microbial mat from Hot Lake, an epsomitic, hypersaline lake in northcentral Washington state (Anderson, 1958; Lindemann et al., 2013). When sampled on 25 July 2012, the seasonal mat had developed visually distinct strata over its approximately 5 mm thickness. The mat overlays a gypsum-dominated sediment that also contains minor amounts (< 10%) of carbonate (dominated by aragonite, data not shown). Ex vivo incubations of replicate mat samples excised from 55 cm water depth were incubated in lake water sampled concurrently with the mat, but modified by the addition of 2.25 mmoles of 50% 13C bicarbonate in a 740 ml glass incubation container. A clear acrylic sheet (1/8 inch thickness) allowed ambient light penetration. Control containers were wrapped in aluminium foil to exclude light. The experimental incubations quickly showed signs of phototrophic activity, producing visual bubbles (presumably O2) after ∼ 15 min of incubation under bright light conditions (near solar noon). We subsampled the mat at 0.5, 1.0 and 2.5 h after starting the incubation. Immediately following subsectioning, samples were rinsed in lake water devoid of labelled bicarbonate for 30 min to ensure the removal of residual label. We performed these rinses in the dark, and immediately after rinsing, samples were stored on dry ice for storage and transportation back to the laboratory.
Fig. 1. Ablated mat cross-section. In preparation for analysis, mat was cross-sectioned with a razor blade then lyophilized prior to mounting in the laser ablation chamber. As seen here, this method retained the internal mat stratification and, going from the surface down, the stratified, orange, green, white, brown and pink layers are visibly distinguished. Laser ablation was performed along three tracks perpendicular to the mat surface, each track acted as replicate profile measurements of the mat sample. Individual spots in each profile represent a 50 μm sample from the mat and contained ample material for isotope ratio determination. Digital microscopy and optics within the laser ablation unit (Cetac LSX-500, Omaha, NE, USA) permitted precise selection and targeting of mat for ablation.
© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 6, 786–791
Spatial tracking of substrate into microbial mat
788
Fig. 2. Label incorporation into the mat as a function of time. We used an elemental analyser (ECS 4010 CHNSO Analyser, Costech Analytical, Valencia, CA, USA) coupled to IRMS (Delta V plus, Thermo Scientific, Bremen, Germany) to measure bulk 13C content of homogenized mat sections. These bulk results showed increasing label incorporation over time in both the total (A) and acid-treated (organic) fractions (B) of the mat. In both cases, we homogenized a ∼ 4 cm2 section of mat taken from immediately adjacent to the laser ablation sample. We compared experimental samples (square) to the dark control (diamond, samples incubated with label in the dark) and error bars are the standard deviation of analytical replicates (n ≥ 3). LA-IRMS results of a mat cross-section (C) likewise show increasing label incorporation over time but more specifically also show a clustering of label towards the top of the mat. We show (C) all LA-IRMS measurement resulting from each of the three tracks through the mat. The mat displayed variable overall thickness within a single sample due to its pinnacled upper surface. In order to compare adjoining sections of mat with different thicknesses, we normalized cross-sectional depth based on proportional thickness into the mat (with 0.0 being the upper and 1.0 being the lower mat surface). The dark control was incubated with the 13C label but kept dark for 2.5 h then harvested. We used the trapezoidal numerical method to integrate the area between an experimental isotope profile for each timepoint and the dark control (with a representative area for the 1.0 h incubation shaded in blue). While mat cross-sections varied between samples, we denoted representative boundaries between the visually distinguished mat strata (orange, green, white, brown and pink).
organic carbon during the ablation (on the order of 1.8 ppb in the carrier gas during ablation). This suggests that thermal decomposition surrounding the ablation pit is minimal and that the sample immediately adjacent to the ablation is preserved for other, complimentary analyses which may include DNA (as demonstrated here), pigment, lipid or other biomarker analyses. We observed increasing 13C accumulation over time in both the total bulk mat (Fig. 2A) and in samples that had been acid washed to remove carbonate material, leaving only organic carbon behind (Fig. 2B). The dark control samples, which we incubated in the presence of the 13C label, did not show significant label accumulation. These data suggest both that the dark conditions prohibited carbon accumulation on this timescale and that a 30 min post-incubation rinse in lake water was effective at removing unincorporated bicarbonate. The combination of advective flow (through microchannels) and diffusive transport control the rate at which the applied labelled bicarbonate moves into the mat and is available for accumulation across the depth profile. Relatively slow transport rates into the mat would favour label accumulation at the mat surfaces while disfavouring accumulation in the mat interior. Our dark control showed that 30 min of rinsing in label-free Hot Lake water enabled sufficient
porewater exchange to remove all label from the sample even after a 2.5 h incubation. The laser ablation data (Fig. 2C) reflect both the increasing label accumulation with time and lack of label uptake in the dark control observed in the bulk analysis data (Fig. 2A and B). The LA-IRMS data presented in Fig. 2C are a compilation of data from each of the three parallel profiles sampled through the mat. Subsequent data analysis considers each of these three profiles independently and is discussed below. In addition to the elemental analysis isotope ratio mass spectrometry (EAIRMS) interpretations, however, the LA-IRMS produces a quantitative profile of label incorporation through the mat cross-section. Incorporation was greatest near the mat surface, with comparatively small amounts at the bottom of the mat. Total label incorporation was estimated by first integrating each profile across relative mat depth (trapezoidal numerical method) and then subtracting the average control (dark) incubation area from each of the respected illuminated areas (schematically depicted for the 1.0 h incubation by the shaded areas in Fig. 2C). Each of the three replicate laser ablation profiles corresponding to an incubation treatment were individually analysed. The total label accumulation calculated from integrating the LA-IRMS profiles through the entire mat thickness was in
© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 6, 786–791
789 J. J. Moran et al.
Fig. 3. Bulk and spatially specific label accumulation and quantification of total label as a function of depth. Comparison between bulk (EA-IRMS) and spatially resolved (LA-IRMS) data (A) shows reasonable agreement in the amount of label associated with each sample at the whole mat scale. Values reported for the bulk mat reflect the increase in isotopic content of the experimental versus control samples with error reported as the sum of squares for the experimental and associated control analytical replicate standard deviations. For the spatially resolved approach, we integrated the area between experimental and control data points from the mat’s upper to lower surface. The spatially resolved data are average difference between the integration of each of the replicate tracks through the 2.5 h incubation mat and the average control data with error reported following sum of squares error propagation of the standard deviation of the experimental and control replicates. Integration of LA-IRMS depth profile restricted to specific depths in the mat (B) enables estimation of where, physically, in the mat, the label accumulation occurs. We report both the relative amount of label accumulation observed at different depths within the mat (outlined squares) and the relative summed accumulation as integrated from the surface to different depths in the mat (black diamonds). Error bars depict either the standard deviation of replicate tracks through the mat (summed accumulation) or sum of squares of the standard deviation of the accumulation through different depths (strata accumulation). Transitions between visually distinguished mat strata (orange, yellow, white, brown and pink) are provided at representative depths in the mat.
close agreement with that measured by bulk EA-IRMS analysis for each of the three timepoints (Fig. 3A). In addition to quantifying bulk label accumulation, the LA-IRMS data reveal the extent of label incorporation along the depth profile of the mat. For example, in the 2.5 h incubation, 75% of the total label accumulation
occurred in the upper half of the mat (Fig. 3B) and the upper 20% of the mat accounted for 48% of the label accumulation. The integrated vertical distribution of label incorporation within the mat was nearly identical through each of the three timepoints (data not shown). Previous work on mat from this location (Lindemann et al., 2013) showed photosynthetically active radiation being attenuated to below detection by ∼ 1.5 mm or within the top 30% of the mat during peak daylight. In our experiment, we demonstrate that this region showed the highest rate of label accumulation (> 60% of total accumulation in the upper ∼ 30% of the mat) but that there was label accumulation beneath. Presumably, this deeper label accumulation results from either transportation of fixed carbon to great depth in the mat or autotrophic processes other than photosynthesis. To characterize the community composition and likely metabolisms within the mat, we sectioned post-ablated samples into five pieces corresponding to the coloured zones observed in Fig. 1: orange, green, white, brown and pink (see Fig. 1). We extracted genomic DNA from these subsections and performed pyrosequencing of the 16S rRNA gene on a 454 GSFLX platform, amplified using the V4 primers 515F and 806R (Caporaso et al., 2011). Sequences were analysed in MOTHER v. 1.31 using the 454 SOP [Schloss et al., 2009) (http://www.mothur.org/ wiki/454_SOP (accessed 24 July 2013)] and clustered using an average neighbour algorithm at 3% distance. These clusters were classified using a Wang approach with the Ribosome Database Project training set v. 9 (updated 20 March 2012 and formatted for MOTHER). Areas of highest label incorporation exhibited significant cyanobacterial relative abundance, although we also measured relatively high cyanobacterial amplicon abundance in strata (including the white, green and brown layers) that incorporated only small amounts of label (Fig. 4). This observation highlights the ability of the LA-IRMS approach to localize actual label accumulation as opposed to metabolic inferences derived solely from community composition. While not measured in this study, previous work on the same microbial mat systems (Lindemann et al., 2013) describe irradiance in the mat with very similar profiles to the accumulation trends we observed. This coupled with the minimal label accumulation in the dark controls suggest that light availability exerts strong influence on label accumulation in this system. The results presented here highlight the ability of LA-IRMS to spatially pinpoint locations within a microbial mat where labelled substrate accumulates. While this specific LA-IRMS system was previously demonstrated on organic polymers (Moran et al., 2011) and a similar system was demonstrated on plant material (Wieser and Brand, 1999; Schulze et al., 2004; Skomarkova et al.,
© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 6, 786–791
Spatial tracking of substrate into microbial mat
790
Fig. 4. Community composition of differently colored mat layers. We sectioned the mat along color stratifications (Fig. 1) and extracted genomic DNA. Mat samples were washed (500 mM EDTA, 10 min), homogenized (in 700 μl of 50 mM Tris pH 8.0 + 25 mM EDTA), then heated (5 min at 85°C) prior to cell lysis (lysozyme 30 mg ml−1 for 1 h at 37°C followed by an overnight incubation at 56°C with 10% SDS and 0.2 mg ml−1 Proteinase K). We used a phenol chloroform extraction followed by ethanol (70%) precipitation. We removed PCR inhibitors using a Zymo Research OneStep PCR Inhibitor Removal Kit following the manufacturer’s instructions. Pyrosequencing was performed on a 454 GSFLX platform by Research and Testing Laboratories (Lubbock, TX, USA) and we processed sequences and taxonomy as previously described (Schloss et al., 2011). The relative abundances, based upon a standardized number of sequences (n = 3452 per layer), of major taxonomic orders within the mat are displayed above. Classes of phylum Proteobacteria are marked with brackets around their composite orders. ‘Unclassified’ refers to sequences from kingdom Bacteria that could not be classified at the phylum level, while ‘Other’ refers to reads classified within orders not otherwise depicted. Orders containing sequences from genera known to be autotrophic are in green.
2006), this is, to the best of our knowledge, the first demonstration of the LA-IRMS technique on a microbial mat system. Our results demonstrate that, under high light intensity, the highest rates of label accumulation occur near the mat surface. The utility of LA-IRMS, however, goes beyond this specific example. Tracking the accumulation of specific labelled compounds using LA-IRMS can not only demonstrate where substrates are consumed within a mat but also how the location of these activities changes over diel or seasonal cycles. Sample preparation for analysis is straightforward and can be performed in most laboratories, requiring only sectioning and lyophilization of a suitable mat sample. Adaptation of these preparation methods to enable drying and mounting a sample for analysis could also enable LA-IRMS probing of any microbial system where spatial resolution is important. Potential examples include microbial communities associated with the plant rhizosphere, soil aggregates, surface biofilms and others. The spatial resolution afforded by LA-IRMS allows separate isotope measurement of microbial biomass and associated growth matrices, whether that is organic (plant roots) or inorganic (soil minerals or some biofilm substrates). Finally, the LA-IRMS system has a high sample throughput for carbon-rich samples such as microbial mats. The δ13C measurement of a spot in our mat sample was completed in less than 10 min. Overall, we believe that due to the ease of sample preparation, microbially relevant spatial resolution, and high sample throughput, the LA-IRMS technique fills an analytical niche for culture-independent assessment of metabolic activity in microbial community.
Acknowledgements This research was supported by the Genomic Science Program (GSP), Office of Biological and Environmental Research (OBER), U.S. Department of Energy (DOE) and is a contribution of the Pacific Northwest National Laboratory (PNNL) Foundational Scientific Focus Area. Alexandra B. Cory was supported by the Lawrence University (Appleton, WI) LUR1 program. A portion of the research described in this paper was conducted under the Laboratory Directed Research and Development Program at Pacific Northwest National Laboratory, a multiprogramme national laboratory operated by Battelle for the U.S. Department of Energy. Hans C. Bernstein and Ryan S. Renslow are grateful for the support of the Linus Pauling Distinguished Postdoctoral Fellowship program. We thank William Chrisler for his assistance in microscope sample photography and Dr. Helen Kreuzer for insightful discussions regarding data interpretation. Conflict of interest: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
References Anderson, G.C. (1958) Some limnological features of a shallow saline meromictic lake. Limnol Oceanogr 3: 259– 270. Behrens, S., Losekann, T., Pett-Ridge, J., Weber, P.K., Ng, W.-O., Stevenson, B.S., et al. (2008) Linking microbial phylogeny to metabolic activity at the single-cell level by using enhanced element labeling-catalyzed reporter deposition fluorescence in situ hybridization (EL-FISH) and NanoSIMS. Appl Environ Microbiol 74: 3143–3150.
© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 6, 786–791
791 J. J. Moran et al. Bernstein, H.C., Beam, J.P., Kozubal, M.A., Carlson, R.P., and Inskeep, W.P. (2013) In situ analysis of oxygen consumption and diffusive transport in high-temperature acidic iron-oxide microbial mats: oxygen consumption in geothermal iron-oxide microbial mats. Environ Microbiol 15: 2360– 2370. Canfield, D.E., and Des Marais, D.J. (1993) Biogeochemical cycles of carbon, sulfur, and free oxygen in a microbial mat. Geochim Cosmochim Acta 57: 3971–3984. Caporaso, J.G., Lauber, C.L., Walters, W.A., Berg-Lyons, D., Lozupone, C.A., Turnbaugh, P.J., et al. (2011) Global patterns of 16S rRNA diversity at a depth of millions of sequences per sample. Proc Natl Acad Sci USA 108: 4516–4522. Des Marais, D.J. (2003) Biogeochemistry of hypersaline microbial mats illustrates the dynamics of modern microbial ecosystems and the early evolution of the biosphere. Biol Bull 204: 160–167. Dupraz, C., Reid, R.P., Braissant, O., Decho, A.W., Norman, R.S., and Visscher, P.T. (2009) Processes of carbonate precipitation in modern microbial mats. Earth Sci Rev 96: 141–162. Franks, J., and Stolz, J.F. (2009) Flat laminated microbial mat communities. Earth Sci Rev 96: 163–172. Ginige, M.P., Hugenholtz, P., Daims, H., Wagner, M., Keller, J., and Blackall, L.L. (2004) Use of stable-isotope probing, full-cycle rRNA analysis, and fluorescence in situ hybridization-microautoradiography to study a methanolfed denitrifying microbial community. Appl Environ Microbiol 70: 588–596. Kelley, C.A., Prufert-Bebout, L.E., and Bebout, B.M. (2006) Changes in carbon cycling ascertained by stable isotopic analyses in a hypersaline microbial mat. J Geophys Res 111: G04012. Klatt, C.G., Inskeep, W.P., Herrgard, M.J., Jay, Z.J., Rusch, D.B., Tringe, S.G., et al. (2013) Community structure and function of high-temperature chlorophototrophic microbial mats inhabiting diverse geothermal environments. Front Microbiol 4: 106. Lindemann, S.R., Moran, J.J., Renslow, R.S., Hutchison, J.R., Cole, J.K., Dohnalkova, A.C., et al. (2013) The epsomitic phototrophic microbial mat of Hot Lake, Washington: community structural responses to seasonal cycling. Front Microbiol 4: 323. doi:10.3389/fmicb.2013. 00323.
van der Meer, M.T.J., Schouten, S., Bateson, M.M., Nubel, U., Wieland, A., Kuhl, M., et al. (2005) Diel variations in carbon metabolism by green nonsulfur-like bacteria in alkaline siliceous hot spring microbial mats from Yellowstone National Park. Appl Environ Microbiol 71: 3978–3986. Moran, J.J., Newburn, M.K., Alexander, M.L., Sams, R.L., Kelly, J.F., and Kreuzer, H.W. (2011) Laser ablation isotope ratio mass spectrometry for enhanced sensitivity and spatial resolution in stable isotope analysis. Rapid Commun Mass Spectrom 25: 1282–1290. Orphan, V.J., House, C.H., Hinrichs, K.U., McKeegan, K.D., and DeLong, E.F. (2001) Methane-consuming archaea revealed by directly coupled isotopic and phylogenetic analysis. Science 293: 484–487. Schloss, P.D., Westcott, S.L., Ryabin, T., Hall, J.R., Hartmann, M., Hollister, E.B., et al. (2009) Introducing mothur: open-source, platform-independent, communitysupported software for describing and comparing microbial communities. Appl Environ Microbiol 75: 7537–7541. Schloss, P.D., Gevers, D., and Westcott, S.L. (2011) Reducing the effects of PCR amplification and sequencing artifacts on 16S rRNA-based studies. PLoS ONE 6: e27310. Schulze, B., Wirth, C., Linke, P., Brand, W.A., Kuhlmann, I., Horna, V., and Schulze, E.-D. (2004) Laser ablationcombustion-GC-IRMS–a new method for online analysis of intra-annual variation of delta13C in tree rings. Tree Physiol 24: 1193–1201. Skomarkova, M.V., Vaganov, E.A., Mund, M., Knohl, A., Linke, P., Boerner, A., and Schulze, E.-D. (2006) Interannual and seasonal variability of radial growth, wood density and carbon isotope ratios in tree rings of beech (Fagus sylvatica) growing in Germany and Italy. Trees 20: 571–586. Villanueva, L., Navarrete, A., Urmeneta, J., White, D.C., and Guerrero, R. (2007) Analysis of diurnal and vertical microbial diversity of a hypersaline microbial mat. Arch Microbiol 188: 137–146. Visscher, P.T., and Stolz, J.F. (2005) Microbial mats as bioreactors: populations, processes, and products. Palaeogeogr Palaeoclimatol Palaeoecol 219: 87–100. Wieser, M.E., and Brand, W.A. (1999) A laser extraction/ combustion technique for in situ delta(13)C analysis of organic and inorganic materials. Rapid Commun Mass Spectrom 13: 1218–1225.
© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 6, 786–791
Copyright of Environmental Microbiology Reports is the property of Wiley-Blackwell and its content may not be copied or emailed to multiple sites or posted to a listserv without the copyright holder's express written permission. However, users may print, download, or email articles for individual use.
