Folia Microbiol (2011) 56:477–481 DOI 10.1007/s12223-011-0071-8
Terminal restriction fragment length polymorphism analysis of soil microbial communities reveals interaction of fungi and chlorine bound in organic matter Milan Gryndler & Hana Hršelová & Zora Lachmanová & Nicolas Clarke & Miroslav Matucha
Received: 21 April 2011 / Accepted: 30 August 2011 / Published online: 16 September 2011 # Institute of Microbiology, Academy of Sciences of the Czech Republic, v.v.i. 2011
Abbreviations AOX Organically bound halogen(s) bp Base pair(s) BSA Bovine serum albumin dbRDA Distance-based redundancy analysis FH Fermentation organic soil horizon HEX 6-Carboxy-4,7,2′,4′,5′,7′-hexachlorofluorescein ITS Internal transcribed spacer PCR Polymerase chain reaction TRF Terminal restriction fragment T-RFLP Terminal restriction fragment length polymorphism
Introduction Chlorine belongs to the 20 most abundant elements on the Earth's surface. The main source of chloride ions in the M. Gryndler (*) : H. Hršelová Institute of Microbiology, Academy of Sciences of the Czech Republic, CZ-14220, Prague, Czech Republic e-mail:
[email protected] Z. Lachmanová Forestry and Game Management Research Institute, CZ-15604, Prague, Czech Republic N. Clarke Norwegian Forest and Landscape Institute, N-1431, Aas, Norway M. Matucha Institute of Experimental Botany, Academy of Sciences of the Czech Republic, CZ-14220, Prague, Czech Republic
forest ecosystem is the oceans, in spite of the fact that more chlorine is contained in the Earth's crust and mantle (Graedel and Keene 1996; Winterton 2000). Chlorine bound to organic matter may originate from various sources. In forest ecosystems, it is mainly present as organically bound chlorine or as chloride ion in the soil and in the biomass. It contributes to the degradation of soil organic matter, which represents the major sink of carbon in the forest ecosystem. Forest soil organic matter is formed mostly from litter, which is decomposed largely by basidiomycetes (DeJong and Field 1997). The content of chloride in the environment depends on the geographic situation, i.e., on distance from the sea, while organically bound chlorine is connected with the biogeochemical cycling (Öberg 1998, 2002, 2003; Johansson et al. 2003). The organically bound halogen (AOX) concentrations in the soil of a coniferous forest ecosystem in Scandinavia always exceeded those of chloride (Öberg 2003). Chlorinated organic compounds are thus not only of anthropogenic origin (e.g., chlorinated solvents, PVC, PCBs, or insecticides) as though in the past. They are present in forest soils, are relatively stable, and, as some of them are soluble, they can penetrate into groundwater. They may thus represent a potential menace in regions with drinking water reservoirs. Many hundredths of chlorinated organic compound were found in nature (Winterton 2000; Gribble 2003; Clarke et al. 2009), which are often produced from deposited sea salt in the processes mediated by halogenating enzymes of peroxidase nature and also abiotically (Shaw and Hager 1959; Clarke et al. 2009). Forest soil microbiota interact with the natural chlorine cycle. This is reflected in changes in composition of the soil microbial community when the soil was incubated at high chloride concentration (Gryndler et al. 2008). Further,
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elimination of living microbiota from soil results in a substantial decrease in chlorination rates (Rohlenová et al. 2009). Influenced by the above facts, we were interested in general patterns of association of different groups of soil microorganisms with the concentration of soil organically bound chlorine (estimated as AOX) in forest soils.
Materials and methods Soil sampling and chemical analyses Soil samples were taken from 12 forest stands located in different parts of the Czech Republic: Benešovice, Březka, Jizerka, Klepačka, Lásenice, Lazy, Luisino údolí, Mísečky, Medlovice, Nová Brtnice, Všeteč, and Želivka. One additional stand near the south coast of Norway in an area with relatively high chloride deposition, Birkenes, was also sampled. These stands belong to the network of intensively monitored plots (level II) of the International Cooperative Programme—Forests (United Nations Economic Commission for Europe). At each locality, three samples from one sampling point were taken and treated separately: (1) organic horizon FH; (2) mineral surface layer, 0 to 100 mm; and (3) mineral subsurface layer, 100 to 200 mm. The sampling depths were measured after the FH horizon was removed. In total, 39 soil samples were analyzed for the content of chloride, total carbon, total nitrogen, and AOX. Chloride content in soil was determined using potentiometric determination by titration with silver ions. The chloride is leached from the oven-dried soil using deionised water and is then titrated with standard AgNO3 solution using the chloride selective electrode (argentochloride electrode) as the end point detection. Carbon and nitrogen were analyzed using a CNS analyzer (Vario Max elementar analyzer). The principle of CNS analysis is based on catalytic tube combustion of soil samples, separation of foreign gasses, and separation of the desired components (N2, CO2, SO2) by TCD detection. The instrument was used for simultaneous determination of C and N in the weighed-in solid samples. Adsorbable organic halogens (predominantly the organically bound chlorine, AOX) were measured as described by Rohlenová et al. (2009). DNA extraction and PCR conditions DNA was extracted from 250 mg representative fresh soil sample (2 mm mesh fraction) using Power Soil DNA Extraction Kit (MoBio). The resulting DNA extract was diluted to a concentration of 2.5 ng/μL (measured by NanoDrop spectrophotometer) and further used as template in PCR. PCR mixture (50 μL) was composed of 25 μL of 2× Combi-PPP mix (Top-Bio Ltd., Prague, Czech Republic;
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contains hot start-Taq DNA polymerase, 5 mmol/L MgCl2, buffer, deoxyribonucleotides, and loader), 1 μL 5′-HEX-labeled forward primer (10 μmol/L), 1 μL reverse primer (10 μmol/L), 4 μL DNA template, and 19 μL water. Communities of soil saprotrophic fungi, Archaea, and Bacteria were studied in all the 39 samples. For the analysis of fungal communities, the primers ITS1F and ITS4 (White et al. 1990) were used in PCR to amplify the ITS region of the rRNA gene cassette. In this particular case, the thermal cycler program was 95°C for 4 min, followed by 34 cycles of 95°C for 45 s, 52°C for 45 s, and 72°C for 90 s. Final extension at 72°C lasted for 5 min. PCR of bacterial 16S rRNA gene was performed using primers 16Seu27f and 783r (Sakai et al. 2004). After initial incubation for 5 min at 95°C, the mixture was subjected to 34 PCR cycles involving 45 s at 94°C, 45 s at 57°C, and 90 s at 72°C. Final extension at 72°C lasted for 5 min. Archaeal 16S rRNA gene was amplified using primers A751F and UA1406R (Baker et al. 2003). The thermal cycler program was 94°C for 4 min, followed by 34 cycles of 94°C for 60 s, 55°C for 60 s, and 72°C for 90 s. Final extension at 72°C lasted for 5 min. T-RFLP analysis Amplified fungal fragments of the ITS region were purified using UltraClean PCR Clean-Up DNA Purification Kit (Mo Bio), evaporated under vacuum to a volume of 15 μL and combined with two units (U) of TaqI restriction endonuclease (NEB), 0.3 μL of 1% bovine serum albumin (BSA) supplied with the enzyme and 3 μL buffer NEB3. The resulting restriction mixture was supplied with water up to the volume of 30 μL and incubated at 65°C for 1 h, then inactivated for 15 min at 80°C, desalted by Post-
Fig. 1 Correlation between soil inorganic chloride and AOX. r2 indicates the coefficient of correlation; the p value denotes its statistical significance. Thin lines delimit the 95% confidence interval of the correlation trend (thick line)
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Table 1 Results of dbRDA and the Monte Carlo test of interaction of soil organically bound chlorine (AOX) with soil communities of fungi, Bacteria, and Archaea Parameter TRF intensity—environment correlation (4 axes) Monte Carlo test: variability explained by AOX (%) F p Number of fragments analyzed
Fungi
Bacteria
Archaea
0.857
0.769
0.637
3.5
2.7
2.2
1.431
1.251
0.995
0.0300 173
0.0560 117
0.2820 67
The italicized value indicates the significant result of the Monte Carlo test at p≤0.05
Reaction Clean-Up Column (Sigma), and analyzed using capillary electrophoresis as indicated below. PCR-amplified bacterial 16S rRNA gene fragments were cleaved by restriction endonuclease AluI (NEB) as follows: purified PCR products were mixed with 5 μL of buffer NEB4 and 20 U of AluI enzyme, incubated 2 h at 37°C,
Fig. 2 Biplot of dbRDA analysis of the interaction between fungal TRF intensities (scores are presented as small triangles) and AOX (the score is presented as a big arrow), as environmental variable. The score numbers indicate the TRF lengths. Effects of chloride, total carbon, total nitrogen, and sampling depth were subtracted as covariables. Long TRF scores sharing the same direction with the AOX score indicate that corresponding microorganisms correlate positively with soil AOX concentration
supplied with another 10 U of AluI enzyme, and incubated for another 2 h at 37°C. Finally, the restriction enzyme was inactivated at 65°C for 20 min, and the sample was desalted by Post-Reaction Clean-Up Column (Sigma) and analyzed using capillary electrophoresis. Archaeal DNA restriction cleavage was performed in a mixture of 50 μL purified PCR product, 5 μL buffer NEB3, 0.5 μL 1% BSA, and 20 U TaqI enzyme for 2 h at 65°C. Another 10 U of the enzyme was then added, and the mixture was then incubated for the next 2 h, finally inactivated for 20 min at 80°C, and finally desalted by Post-Reaction Clean-Up Column (Sigma). After restriction cleavage and desalting, 0.5 μL of GeneScan 400HD ROX size standard (Applied Biosystems) per 14-μL sample aliquot was added, and the samples were transferred to the sample plate, denatured at 96°C for 5 min, instantly cooled down to 4°C in a thermocycler (Biometra), and analyzed using capillary electrophoresis (ABI Prism 3,130 Genetic analyzer, Applied Biosystems) at run voltage 15 kV and oven temperature 60°C. POP7 polymer (Applied Biosystems) was used for fragment
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analysis. Samples were loaded electrokinetically (15 s at 1.6 kV). Data analysis The data were tabulated using GeneMarker 1.85 software (Soft Genetics LLC, State College, USA) and further analyzed using distance-based redundancy analysis (dbRDA) (Canoco 4.5 software package, Biometris, Wageningen, The Netherlands). Only signals stronger than 50 fluorescence arbitrary units of fragments fitting into the length range 60 to 400 bp and occurring in at least ten samples were taken into account. Soil AOX concentration was used as environmental variable, while soil chloride, total carbon, total nitrogen, and soil sampling depths were used as covariables. Sampling depths were expressed as three additional nominal parameters defining the blocks of randomization. The Bray–Curtis dissimilarity measure of square root-transformed terminal restriction fragment (TRF) intensity generated by PrCoord Canoco tool was analyzed (Grant and Ogilvie 2003), and negative eigenvalues were corrected. The Monte Carlo test (499 permutations) was exploited to evaluate the significance of the results.
Results and discussion As seen from Fig. 1, a correlation between inorganic soil chloride and organically bound chlorine was found. This means that, being interested in interaction of the soil biota with organically bound chlorine, we cannot simply correlate concentration of AOX with microbial signals. This might lead to incorrect conclusions since soil microbiota are significantly affected by chloride as well (Gryndler et al. 2008). Thus, we had to subtract the effects of other measured soil parameters including inorganic chloride from the analysis output. This may be performed by using these parameters as covariables in dbRDA. The results of the analysis are presented in Table 1. The variability of TRF signals is best explained by the AOX concentration in the case of fungi, which is the only statistically significant result of the Monte Carlo permutation test. The proportion of the variability explained in the case of bacterial TRF signals is smaller and is statistically insignificant with a marginal p value (0.056). A still smaller proportion of the variability is explained in the case of archaeal TRF signals. A biplot of dbRDA analysis of fungal TRFs is shown in Fig. 2. The same orientation of some fungal TRF scores and AOX score indicates a positive correlation of a part of soil
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fungal microbiota with increased concentrations of organically bound chlorine. The results are visualized after subtraction of the possible interference of inorganic soil chloride and other soil parameters as covariables. The biplots describing scores of other than fungal TRFs are not given since the corresponding results of the Monte Carlo test are not significant. A relatively small proportion of the variability in fungal TRF intensities explained by AOX (3.5%, Table 1) can be hypothetically attributed to effects of other varying factors affecting the studied localities, such as plant community composition and age, soil humidity, temperature regime, etc., which introduce an additional variability into the statistical model used in the analysis. We can conclude that the only microbial group significantly correlated with organically bound chlorine in forest soil are fungi. This may be connected with the crucial role of fungi in the humification of organic matter, a process closely connected with the accumulation of organically bound chlorine (Keppler and Biester 2003). Fungi possess a complex range of oxidative enzymes which produce a high number of halogenated (chlorinated) compounds and are the most important source of these compounds in forest litter (DeJong and Field 1997). It has been demonstrated in the past (Matucha et al. 2007) that chlorinated organic compounds are degradation intermediates of organic matter in forest soil. The association of a portion of fungal community with higher concentration of AOX in soil thus seems so be consistent with role of the fungi in the biodegradation of a significant portion of soil organic matter. Acknowledgments This research was supported by the Norwegian Financial Mechanism (project CZ 0135), by the Ministry of Education, Youth and Sports (project no. 7F09026), by the Institute of Microbiology, the Research Concept AV0Z50200510, and the Institute of Experimental Botany, AS CR, the Research Concept AV0Z50380511. We thank the staff of the testing laboratories of the Forestry and Game Management Research Institute and T.G.M. Water Research Institute for the laboratory analyses.
References Baker GC, Smith JJ, Cowan DA (2003) Review and re-analysis of domain-specific 16S primers. J Microbiol Meth 55:541– 555 Clarke N, Fuksová K, Gryndler M, Lachmanová Z, Liste HH, Rohlenová J, Schroll R, Schröder P, Matucha M (2009) The formation and fate of chlorinated organic substances in temperate and boreal forest soils. Env Sci Pollut Res 16:127–143 DeJong E, Field JA (1997) Sulfur tuft and turkey tail: biosynthesis and biodegradation of organohalogens by basidiomycetes. Annl Rev Microbiol 51:375–414
Folia Microbiol (2011) 56:477–481 Graedel TE, Keene WC (1996) The budget and cycle of Earth's natural chlorine. Pure Appl Chem 68:1689–1697 Grant A, Ogilvie LA (2003) Terminal restriction fragment length polymorphism data analysis. Appl Environ Microbiol 69:6342– 6343 Gribble GW (2003) The diversity of naturally produced organohalogens. Chemosphere 52:289–298 Gryndler M, Rohlenová J, Kopecký J, Matucha M (2008) Chloride concentration affects soil microbial community. Chemosphere 71:1401–1408 Johansson E, Sandén P, Öberg G (2003) Organic chlorine in deciduous and coniferous forest soils in southern Sweden. Soil Sci 168:347–355 Keppler F, Biester H (2003) Peatlands: a major sink of naturally formed organic chlorine. Chemosphere 52:451–453 Matucha M, Gryndler M, Schröder P, Forczek ST, Uhlířová H, Fuksová K, Rohlenová J (2007) Chloroacetic acids—degradation intermediates of organic matter in forest soil. Soil Biol Biochem 39:382–385 Öberg G (1998) Chloride and organic chlorine in soil. Acta Hydrochim Hydrobiol 26:137–144
481 Öberg G (2002) The natural chlorine cycle—fitting the scattered pieces. Appl Microbiol Biotechnol 58:565–581 Öberg G (2003) The biogeochemistry of chlorine in soil. In: Gribble G (ed) The handbook of environmental chemistry, vol. 3, part P. Springer, Berlin, pp 43–62 Rohlenová J, Gryndler M, Forczek ST, Fuksová K, Handová V, Matucha M (2009) Microbial chlorination of organic matter in forest soil: investigation using 36Cl-chloride and its methodology. Env Sci Technol 43:3652–3655 Sakai M, Matsuka A, Komura T, Kanazawa S (2004) Application of a new PCR primer for terminal restriction fragment length polymorphism analysis of the bacterial communities in plant roots. J Microbiol Meth 59:81–89 Shaw PD, Hager LP (1959) An enzymatic chlorination reaction. J Am Chem Soc 81:1011–1012 White TJ, Bruns T, Lee S, Taylor J (1990) Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: Innis MA, Gelfland DH, Sninsky JJ, White TJ (eds) PCR protocols, a guide to methods and applications. Academic, San Diego, pp 315–322 Winterton N (2000) Chlorine: the only green element-towards a wider acceptance of its role in natural cycles. Green Chem 2:173–225
