UV irradiation of natural organic matter (NOM): impact

UV irradiation of natural organic matter (NOM): impact on organic carbon and bacteria Andrea Paul, Claudia Dziallas, Elke Zwirnmann, Egil T. Gjessing & HansPeter Grossart Aquatic Sciences Research Across Boundaries ISSN 1015-1621 Volume 74 Number 3 Aquat Sci (2012) 74:443-454 DOI 10.1007/s00027-011-0239-y

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Author's personal copy Aquat Sci (2012) 74:443–454 DOI 10.1007/s00027-011-0239-y

Aquatic Sciences

RESEARCH ARTICLE

UV irradiation of natural organic matter (NOM): impact on organic carbon and bacteria Andrea Paul • Claudia Dziallas • Elke Zwirnmann Egil T. Gjessing • Hans-Peter Grossart

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Received: 10 January 2011 / Accepted: 10 November 2011 / Published online: 8 December 2011 Ó Springer Basel AG 2011

Abstract UV-induced transformation of dissolved organic matter (DOC) is often accompanied by reduction of molecular weight and aromaticity and an increase of low-molecular weight (LMW) matter that can be utilized as a substrate by heterotrophic bacteria. Moreover, the generation of reactive transients and mineralization of DOC occurs. For a better understanding of the modification that starts after irradiation and to distinguish between possible chemical and biological modifications, we selected different natural organic matter (NOM) from Norway and Germany. The aqueous solutions were treated by UV irradiation and divided into two aliquot samples. NaN3 anti-bacterial treatment was applied to one sample, and high-pressure size-exclusion chromatography (HPSEC)

Electronic supplementary material The online version of this article (doi:10.1007/s00027-011-0239-y) contains supplementary material, which is available to authorized users. A. Paul (&) E. Zwirnmann BAM Federal Institute for Materials Research and Testing, Richard-Willsta¨tter-Straße 11, 12489 Berlin, Germany e-mail: [email protected] A. Paul E. Zwirnmann Department of Shallow Lakes and Lowland Rivers, Leibniz Institute of Freshwater Ecology and Inland Fisheries, Mu¨ggelseedamm 301, 12561 Berlin, Germany C. Dziallas H.-P. Grossart (&) Department of Limnology of Stratified Lakes, Leibniz Institute of Freshwater Ecology and Inland Fisheries, Alte Fischerhuette 2, 16775 Stechlin, Germany e-mail: [email protected] E. T. Gjessing Department of Chemistry, University of Oslo, Po Box 1033, 0135 Oslo, Norway

analysis was used for both. In all samples, we found typical modifications of NOM after UV irradiation. Incubation ([7 days) of UV-irradiated NOM samples resulted in lower levels of LMW matter and increased aromaticity. Parallel to these changes of carbon fractions, an increase in bacterial cell numbers was observed. Addition of NaN3 to NOM, however, inhibited the reduction of LMW matter, indicating that microbial activity accounted for the observed changes in NOM. Analysis of the bacterial community composition by denaturing gradient gel electrophoresis (DGGE) of the amplified 16S rRNA genes revealed that bacterial communities of non-irradiated and UV-irradiated NOM were different and that UV selected for specific members of a-proteobacteria, b-proteobacteria, and Bacteriodetes. Our results imply that after UV-irradiation of NOM, specific bacterial members are well adapted to low pH, high LMW DOC concentrations, and oxidative stress, and therefore thrive well on UV-irradiated humic matter. Keywords Humic matter Bacteria High-pressure size-exclusion chromatography (HPSEC) UVC Denaturing gradient gel electrophoresis (DGGE) Sequencing

Introduction Solar radiation is a fundamental ecosystem modulator (Wetzel and Tuchmann 2005). In particular, UV-radiation accelerates the degradation of organic matter either by photolysis or by oxidation of organic compounds to CO2, often followed by enhancing the bio-availability of complex organic substrates to microbes. UV light is conveniently split into three spectral bands: UVC (200–280 nm), UVB

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(280–315 nm) and UVA (315–400 nm). UVC is absorbed by stratospheric gases and usually does not reach the earth, while UVB is an important photo-activating agent in waters. UVA is absorbed less readily and penetrates more deeply into water. Dissolved humic substances (HS) are the major chromophoric compounds of natural organic matter (NOM) present in all freshwater systems. After absorption of UV/ VIS light by HS, complex photochemical processes, including energy and electron transfer processes, proceed. The photo-degradation of HS is caused both by direct photolysis and by reactions with photo-induced transient oxidants, such as singlet oxygen, hydrogen peroxide (H2O2), hydroxyl radicals, super-oxide, and organo-peroxy radicals, which are classified as reactive oxygen species (ROS). Although quantum yields of ROS formation by humic matter are in general low (Kieber et al. 2003), the strongly UV-absorbing humic matter fraction (HM) of NOM is efficiently degraded by UV-irradiation, whereas the non-absorbing low-molecular weight (LMW) fractions, such as hydrophilic charged and neutral compounds, become more concentrated (Kulovaara et al. 1996; Schmitt-Kopplin et al. 1998; Buchanan et al. 2004). However, UV-irradiation does not increase the bioavailability of NOM in all cases. Kaiser and Sulzberger (2004) found that photochemical transformations may reduce the bioavailability of initially bioreactive LMW and hydrophilic compounds. This was particularly the case for initially bioavailable LMW and hydrophilic compounds. They also found that the bioavailability of initially biorecalcitrant hydrophobic compounds increased upon irradiation, whereas that of HMW compounds did not have any effect. An overview of this topic is given by Sulzberger and Durisch-Kaiser (2009). Even though the impacts that UV-irradiated NOM may have on biota have received considerable attention during the last decade (e.g., Wetzel et al. 1995; Bertilsson and Tranvik 2000), there is apparently some disagreement whether this treatment causes any adverse effects on organisms, in particular on microbial communities in natural waters. A number of laboratory studies suggests that irradiated NOM causes toxic effects in a wide range of aquatic organisms, including bacteria (Lund and Hongve 1994; Paul et al. 2006), algae (Gjessing and Ka¨llqvist 1991; Hessen and van Donk 1994; Sun et al. 2006), and crustaceans (Frimmel 1998; Parkinson et al. 2001). In some cases, long-lasting adverse effects with time spans over several days, hereby eliminating ROS as potential toxic agents, were observed (Gjessing and Ka¨llqvist 1991; Lund and Hongve 1994; Paul et al. 2006). This will eliminate ROS as potential toxic agents. Even H2O2, which may last up to 4 days in clear water at low temperatures (Kieber et al. 2003), could not account for these observed long-lasting

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adverse effects. In one of these reported studies, NOMs contained untypically high amounts of copper and the liberated cupric ions caused the adverse biological effect, as reported by Parkinson et al. (2003). So far, the abovementioned long-lasting adverse effects on various aquatic organisms do not fit any known pattern. Most of these experiments were performed by using UVC irradiation. Oxidative chemical degradation of humic matter by UVC irradiation generates fatty acids, aliphatic and aromatic carboxylic, as well as phenolic and benzenecarboxylic acids (Schmitt-Kopplin et al. 1998). Except for ROS and NOMs with elevated levels of potentially toxic metal species, it seems worthwhile to study changes in the organic matrix following UVC-irradiation. In previous studies, the effects of UV-irradiation on organic carbon were evaluated by using different chromatographic techniques (Frimmel 1998; Parkinson et al. 2003; Buchanan et al. 2004). However, in most cases analytics and chromatographic runs were performed immediately after irradiation and merely yielded a snapshot on the current state of organic carbon without providing information about long-term effects. Here, we focus on studying the long-term effects after UV-irradiation. The present studies are based on solutions of well-characterized NOM isolates from the Norwegian NOM-Typing Project (Gjessing et al. 1999); that particular samples have also been used in a former NOM-UV-irradiation study where Escherichia coli (E. coli) was used as a test organism (Paul et al. 2006). In this study, we used an irradiation protocol that is similar to that used in these former studies. Several different methods, such as high-pressure sizeexclusion chromatography (HPSEC), have been used to characterize NOM during the period of storage.

Materials and methods Sampling of natural organic matter (NOM) NOMs from Humex (HUM), Birkenes (BIR), Trehørningen (TRE), Hellerudmyra (HEO) and Gjerstad (GJU) in Norway were sampled in 1996 using the reverse osmosis procedure (Gjessing et al. 1999). HUM The catchment of this sample is representative for ‘‘coastal surface water’’ and typical for Western Norway, with high annual precipitation (2,000–3,000 mm). Humex B is the reference lake in half of the Humic Lake Acidification Experiment (Gjessing et al. 1998). The character and nature of the catchment and water quality of Lake Skjervatjern

Author's personal copy UV impact on NOM and bacteria

(‘‘Humex lake’’) is well described in more than two dozen publications. BIR This is a sampling station that was established in 1972 in relation to the Norwegian Acid Rain Program [SNSFProject (‘‘Acid Rain, Effects on Forestry and Fish’’ 1972–1980)]. It is located approximately 20 km from the south coast of Norway. The main feature with this sampling station is NOM from an acidified area. TRE This it the upper lake of the water course of Trehørningen, consisting of four lakes. Theoretical retention time for the water in the lake is 9.4 months. The size of the catchment is estimated to be 6 km2. HEO This is a small catchment (0.08 km2) within the water course of Trehørningen. The same sampling site was used for the IHSS Nordic Fulvic and Humic Reference Material (June 1986). The NOM is considered to be ‘‘young’’ in the water phase. GJU That sample is from a control catchment, which at the same time is representative for acidified areas, and should be comparable with the sample from Birkenes (BIR). The catchment is small (0.40 km2). These NOMs have also been extensively characterized in the ‘‘Norwegian NOM Typing Project’’ and reported in a Special Issue of Environment International (Egeberg et al. 1999; Gjessing et al. 1999). Fuchskuhle (FUKU) NOM was sampled in July 2003 from the humic-rich southwest basin of Lake Große Fuchskuhle, an artificially divided lake in the Northeast of Germany (Sachse et al. 2001). FUKU has been well characterized with regard to its chemical and photosensitizing properties (Paul et al. 2004), and tested for effects on the fish mould Saprolegnia parasitica (Meinelt et al. 2007) and for bacterial communities (Burkert et al. 2003; Allgaier and Grossart 2006; Hutalle-Schmelzer et al. 2010).

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Fuchskuhle and 3–7 times higher than that of HUM and BIR. Solutions were kept in the dark for 24 h and then filtrated through a 0.45 lm cellulose acetate filter (Millipore) to remove traces of non-dissolved NOM. The UVA/B irradiation (5 tubes ARIMET-B, Cosmedico) was performed from the distance of 15 cm. Sterile glass petri dishes were filled with dissolved NOM to a level of 1 cm and covered by quartz plates to prevent evaporation. The spectral irradiance of the tubes was determined to a value of about 3 and 97 W m-2 in the UVB and UVA, respectively, by a spectral radiometer OL754 coupled with a double monochromator and an integrating sphere (Ulbricht Kugel) optic. For the irradiation time intervals of 2, 4, 6, and 15 h, doses of 72, 144, 216, and 541 J cm-2 were calculated. UVC irradiation was performed by a TNN 15/35 lamp that emits a single narrow line at 254 nm. Such lamps are typically used for water disinfection. Here, we applied the spectral irradiance of about 2.6 W cm-2 (estimated from manufacturer information by taking into account the experimental arrangement) for irradiating dissolved NOM in 1 cm quartz cells at a distance of 2 cm. Irradiation periods of 10, 20, 30, 60, 90, and 120 min resulted in doses of 19, 38, 58, 115, 173, and 230 J cm-2 at the surface of the cells. For comparison: typical doses used for water disinfection range between 40 and 100 mJ cm-2 (Buchanan et al. 2006). For the experiments involving characterization of bacterial community structure by denaturing gradient gel electrophoresis (DGGE), larger sample volumes were needed. In this case, NOM samples were irradiated in 200 mL quartz tubes with an inner diameter of 3.5 cm. Here, we used a higher dilution of NOM in order to keep the similar absorbance. Cells were shaken and tubes were turned several times during irradiation in order to perform a homogeneous irradiation of solutions. Absorbance spectra of non-irradiated references and irradiated samples are provided in Fig. S1. For incubation experiments, samples were split according to the scheme shown in Fig. 1, and 0.02% (m/m) NaN3 was added to one batch to stop microbial growth. To the other batch the same volume of water was added. All samples were kept in flasks in an incubator at 22°C in the dark for a period from 7 days to a maximum of 33 days.

Preparation and irradiation of dissolved NOM

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NOM isolates were re-dissolved in sterile Milli(Q) water to a concentration of about 10–20 mg C L-1; i.e., at the concentration close to the natural one of Lake Große

HPSEC was performed by a custom-designed automated high-pressure size-exclusion chromatograph connected to detectors for UV and organic carbon (Huber and Frimmel

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Total bacterial cell numbers

dissolved NOM

Total bacterial cell numbers were determined after filtration on black polycarbonate filters (0.2 lm pore-size, Millipore, Eschborn, Germany) and staining with 40 ,6diamidino-2-phenylindole (DAPI, Fluka) according to Porter and Feig (1980). Cells were counted under a fluorescence microscope at 1,0009 magnification immediately after filtration and staining.

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Fig. 1 Scheme of incubation experiments of dissolved NOM including UVC irradiation by a TNN 15/35 lamp (spectral irradiance 2.6 W cm-2) and addition of 0.02% (w/w) NaN3 to inhibit microbial growth (control treatments)

1994; Sachse et al. 2001). In general, duplicates of all samples were analyzed and diluted to concentrations below 5 mg L-1 DOC prior to chromatographic analysis. The column for size-exclusion chromatography was packed with Toyopearl HW 50 S resin (250 9 20 mm) and eluted with 29 mmol L-1 phosphate buffer (pH 6.5) at a flow rate of 1 mL min-1. The response of the different fractions in the chromatograms were evaluated using custom-designed software. To calculate the apparent molecular weights (MP) for the HM fraction from the retention time of the peak maximum, the system was calibrated with polydextranes (Mp: 830, 4,400, 9,900, 21,400, 43,500) from Polymer Standards Service, saccharides (raffinose, maltose, glucose), glycerin, and methanol (all from Merck). The specific UV absorbance (sUVa), which is the ratio of the specific absorption coefficient at 254 nm (SAC in m-1) and the organic carbon of the humic matter peak (DOC in mg L-1), was calculated. The sUVa, which is also referred to as ‘‘aromaticity’’, can be used as a measure for aromatic and unsaturated structures in NOM as derived from NMR (Huber and Frimmel 1994). The performance of the system and reliability of the evaluation procedure were checked with mixtures of typical compounds, e.g., dextran, acetic acid, formic acid, glucose, and Suwannee River standard humic acid (1S101H, http://www.ihss.gatech.edu/). Experiments with mixtures of different concentrations and combinations of compounds showed that both the highmolecular and low-molecular weight neutral (LMWN) fraction can be determined with 90% recovery, whereas the low-molecular weight acid (LMWA) fraction can be determined with 50–70% recovery, depending on their initial concentration (Fig. S2). In contrast, high-molecular weight substances were, in most cases, not reliably separated from the HM fraction.

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Fifty mL of water and dissolved NOM were filtered through 0.2 lm Nucleopore membranes (Sartorius AG, Go¨ttingen, Germany) to analyze the bacterial community composition by denaturing gradient gel electrophoresis (DGGE) of amplified 16S rRNA genes. Extraction of genomic DNA was performed using a protocol with lysozyme, proteinase K, and zirconium beads (Allgaier and Grossart 2006). Fragments of bacterial 16S rRNA gene fragments were amplified using universal primers 341f-GC and 907r (Brinkhoff and Muyzer 1997). Equal amounts of DNA were loaded in each lane (ca. 500 ng) of a 7% polyacrylamide gel with a denaturing gradient from 40 to 70% (urea/formamide). The running time of DGGE gels was 20 h. Afterwards, DGGE gels were stained with SYBRGold (Molecular Probes) for 30 min, destained with Milli(Q) water for 10 min and illuminated on a UV table (Biometra). Cluster analyses of the DGGE banding patterns were performed employing GelCompare II, Version 3.5 (Applied Maths) software using Dice0 s correlation index (presence/absence of bands, no band intensity) and the unweighted pair-group method with arithmetic means (UPGMA). Sequencing of DGGE bands DNA from excised DGGE bands was re-amplified using primers 341f and 907r and the PCR protocol of Grossart et al. (2005). Aliquots (5 ll) of the amplification products were analyzed by electrophoresis on 1.5% (w/v) agarose gels stained with ethidium bromide (1 mg mL-1). DNA was purified using the NucleoSpin Extract II PCR purification kit (Macherey & Nagel, Germany) according to the manufacturer’s protocol. Purified DNA was sequenced by using the BigDye Terminator v3.1 cycle sequencing kit (Applied Biosystems) and an ABI Prism 3100-Avant genetic analyzer (Applied Biosystems) according to the manufacturer’s instructions. Sequences were compared with those of reference organisms by BLAST search (http://www.ncbi.nlm.nih.gov/blast). The taxonomy browser of the NCBI server was used to determine phylogenetic affiliation. Partial sequences of 16S rRNA genes obtained


in this study were deposited in GenBank with the following accession numbers: FJ268749 to FJ268762. Phylogenetic analyses Phylogenetic analyses were performed using the ARB software package (www.arb-home.de). The sequences from 16S rRNA fragments were imported into an ARB database with approx. 52,100 aligned sequences. The new sequences were aligned automatically with the FastAligner tool and corrected by hand. For calculation of phylogenetic trees, only sequences of more than 1,400 nucleotides were used. A 50% base frequency filter was calculated within ARB to exclude highly variable positions within the sequences and to obtain more robust calculations. Phylogenetic analyses were performed by the maximum-likelihood algorithm using the fast DNAml program of ARB. This tree was compared with trees of other construction methods (i.e., neighbor joining and maximum parsimony) to test the stability. Partial sequences (between 450 and 570 nucleotides) were added to the trees according to maximum-parsimony criteria and applying the 50% base frequency filter. This tool neither corrects for evolutionary distances nor allows changes in overall tree topology.

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UV254-detection mode usually does not display the LMWN fraction (Fig. 2b). The UV-irradiated NOM exhibited an even more complex pattern than the non-irradiated sample. Furthermore, the chromatogramms were strongly affected by the irradiation time (Fig. 2a, b). The OC-detection reveals a clear reduction in HM that further divides into several subfractions (Fig. 2a). In contrast, UV254-detection (Fig. 2b) shows the reduction in HM but only a slight split into the sub-fractions, indicating that the newly formed material at least partly comprises non-UV-absorbing compounds. The shift of the HM fraction to longer retention times further indicates that the MP was significantly reduced, in the case of HUM from 4.9 kDa to 3.2 and 3.0 kDa, respectively. A significant increase in the LMWA fraction could be detected by the organic carbon mode, but not with the UV254-detection mode. The lowest irradiation dose (19 J cm-2) resulted in a small increase of the LMWN fraction. However, it appears from Fig. 2a, b that a doubling of the UVC dose (to 38 J cm-2) resulted only in a small reduction in HM size and in an obvious increase in LMWA concentration. On the other hand, this doubling of

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Results Effects of UVA/B and UVC irradiation of NOM on molecular size distribution The effects of both UVA/B and UVC irradiations were assessed by analyzing HPSEC chromatograms of irradiated NOM. In the organic carbon mode (OC, Fig. 2a), four chromatographic fractions can be separated for non-irradiated HUM (reference) with increasing retention time: high-molecular weight (HMW), HM, LMWA, and ultimately LMWN. HMW matter includes mainly hydrophilic biopolymers, such as polysaccharides and proteins, and ion-organic colloids including mainly polyelectrolytes with a negative charge, neutral poly-hydroxides, and oxyhydrates of Fe, Al, and Si (Huber and Frimmel 1994). The dominant carbon fraction in non-irradiated HUM is the HM fraction, including recalcitrant humic and fulvic acids with a typical high absorbance in the UV range. Furthermore, at longer retention times, the low-molecular weight acids (LMWA), which represent the sum of fractions of all mono- and diprotic low-molecular weight organic acids, can be seen as a pronounced spike. Finally, the minor peak at retention times [60 min represents the low-molecular weight neutral and amphiphilic compounds (LMWN), e.g., sugars, alcohols, aldehydes, ketones, and amino acids. The

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Fig. 2 Typical chromatograms of non-irradiated reference NOM (black line) and UVC-irradiated NOM (low dose: grey line, high dose: light-grey line) from the sampling site Humex B (HUM) with indication of carbon fractions HMW, HM, LMWA, and LMWN. Chromatograms: a by organic carbon (OC) detection and b UV254detection (UV)

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the UVC dose yielded a significant reduction in UV254detectable matter. Furthermore, the first irradiation interval led to a small increase in LMWN. However, with increased UVC dose, a non-structured background instead of a shaped peak was observed by OC-detection. HMW is present in low amounts but is lowered by increasing UVC irradiation. This fraction probably comprises both polysacchararides and inorganic colloids in the HUM. The results described above are related to HUM from the Norwegian ‘‘NOM-Typing project’’. Similar results from the four other isolates are shown in Fig. S3. Here, UVA/B is compared with UVC to estimate its effect on the molecular size distribution of NOM. It can be seen from these results (Fig. S3) that: (i) quality of radiation has a remarkable impact on NOM degradation rather than its energy dose; and (ii) different NOM samples respond differently upon radiation. This is apparently the case for UVA/B irradiation. It should be pointed out that the molecular weight distribution of sample TRE differed only slightly upon an increase in UVA/B dose of more than seven times. Since sample TRE originates from a rather big catchment area, where NOM is rather ‘‘old’’ in the water phase (retention time in the upstream lake is more than 9 months), it has been subjected to global UVAB for an extended period of time leading to an increased photodegradation of this NOM type. Effects of incubation-time on molecular size distribution of NOM after UVC irradiation In order to study the effects of incubation time on molecular size distribution of NOM after UVC irradiation, samples were split according to Fig. 1 into non-treated and NaN3-containing aliquots. NaN3 was added to stop potential microbial growth. In Fig. 3, results of the HPSEC fractionation of the HUM sample are summarized. The three graphs of the top row represent non-irradiated samples: (a) NaN3-treated, and (b) non-treated control with OC detection, and (c) nontreated control with UV detection. In samples with NaN3, no detection by UV was possible due to its strong UV absorbance. The three graphs in the middle represent lowdose irradiated samples (19 J cm-2): (d) NaN3-treated and (e) non-treated with OC detection, and (f) non-treated with UV detection. The lower three graphs (g), (h), and (i) are identical with the middle row, except for the irradiation dose that was enhanced to 38 J cm-2. The different lines on the graphs represent equal days of storage after irradiation (0, 4, 10, 13, and 28 days). The following findings emerge from Fig. 3: (1) Considering the three graphs of the left column (i.e., a, d, and g), a clear effect of NaN3 addition on the molecular weight

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distribution is evident. However, rather small changes on the HPSEC pattern during 33 days of storage occur. (2) The middle graphs (i.e., b, e, and h) indicate both a UVCdose effect on the molecular size distribution and an obvious effect of storage time. (3) The three graphs in the right column (i.e., c, f, and i) suggest smaller changes in the UV254-detectable material during storage (c)—even with small irradiation doses (f: 19 J cm-2). However, with higher UVC dose, (I: 38 J cm-2), a noticeable storage effect can be observed. Overall, these results suggest that microorganisms in the sample remain active during storage of UVC-irradiated NOM, meanwhile changing the amount and quality of the organic matter. In the following paragraphs, changes in DOC, HM, LMWA, LMWN, MP, and SUVA of HUM during incubation are discussed. DOC concentrations in non-irradiated control samples were almost constant over the entire observation period (Fig. 4a). However, in UVC-irradiated samples, a pronounced reduction in DOC concentrations during the first 10 days of storage occurred. A similar pattern was observed for the HM-fraction (Fig. 4b). The higher the UVC dose, the higher the organic matter reduction during the first 10 days of storage. Figure 4c illustrates the apparent molecular weight of HM. There are only very small changes in the Mp values of the non-irradiated sample during 33 days of storage. UVC irradiation reduced the apparent Mp, however, no clear storage effect and no apparent dose effect (i.e., a difference between 19 J cm-2 and 38 J cm-2) were found. Figure 4d illustrates changes in the LMWA fraction (%) with storage time. However, the LMWA content in a nonirradiated sample remained low and constant during storage. In contrast, these acids contribute between 7 and 17% just after UVC-irradiation, whereas their concentrations dramatically decreased during storage. The impact of UVC irradiation and storage time after irradiation on the amount of LMWN are illustrated in Fig. 4e. Although there is a clear trend being similar to that of LMWA, LMWN seems to be less affected by storage after irradiation. According to Fig. 4f, UVC-irradiation reduces the sUVa units from 6 to 5 and further to 4 at the highest UVC dose (38 J cm-2). This confirms a decrease in the content of aromatic compounds as a consequence of UVC irradiation. Furthermore, SUVA values increased with time of storage. To determine whether these observations are specific to HUM or a general property of dissolved organic matter, further incubation experiments were performed with UVC irradiated BIR and FUKU. For both NOM types, similar effects of UVC irradiation and subsequent incubation on DOC fractions were observed (Figs. S4 and S5).


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Fig. 3 Typical chromatograms of non-irradiated reference NOM and of UVC-irradiated NOM from the sampling site Humex B (HUM) after incubation. Chromatographic fractions during incubation: boldface black line: immediately after irradiation, black lines: 1, 4, 10, 13 days incubated, bold-face grey line: 28 days incubated at 22°C, a control added NaN3 in OC detection, b control in OC detection,

c control in UV detection, d 19 J cm-2 irradiated sample in OC detection added NaN3, e 19 J cm-2 irradiated sample in OC detection, f 19 J cm-2 irradiated sample in UV detection, g 38 J cm-2 irradiated sample in OC detection added NaN3, h 38 J cm-2 irradiated sample in OC detection, and i 38 J cm-2 irradiated sample in UV detection

Effect of NOM on bacteria

samples, after a delay of 3 days, bacteria were detected at which bacterial numbers increased and peaked at 5 days in the irradiated FUKU sample (Fig. 5b). In contrast, in the non-irradiated FUKU sample, bacterial numbers did not show such a pronounced peak but increased to a lower number. Addition of NaN3 inhibited bacterial growth in both irradiated and non-irradiated samples (Fig. 5a, b). DAPI staining shows that not only numbers but also morphology of bacteria differed depending on the sample and the date (data not shown). To determine the bacterial community composition, larger batches of FUKU and HUM were treated with UVC irradiation (19 J cm-2) and were incubated for 16 days. A volume of 100 mL was sampled weekly to amplify 16S rRNA genes that were subjected to denaturing gradient gel electrophoresis (DGGE). Temporal changes in DOC, HM, LMWA, LMWN, sUVa, and MP derived from the HPSEC for FUKU are summarized in Fig. S5. In general, these results

Analyses of chromatograms from (non-irradiated and UVC-irradiated) NOM samples with or without NaN3 indicated that the observed changes might be caused by microbial activity. Therefore, further experiments were performed that included the determination of bacterial cell numbers and the bacterial community composition in NOM. For this experiment, we resort to FUKU, since bacterial communities of Lake Große Fuchskuhle have been well studied (Burkert et al. 2003; Allgaier and Grossart 2006; Grossart et al. 2008; Hutalle-Schmelzer et al. 2010). After 1 day of incubation, a strong decrease in the LMWA fraction occurred in both samples without NaN3. Subsequently, LMWA in the non-irradiated sample remained low throughout the experiment (Fig. 5a). In the irradiated samples, however, a constant decrease in LMWA was observed during the incubation. In both

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Fig. 4 Parameters derived from HPSEC to characterize dissolved organic matter (DOC) of non-irradiated and UVC-irradiated NOM from the sampling site Humex B (HUM) over an incubation period of 33 days. a DOC: concentration of organic carbon; b HM: HM

fraction; c MP: apparent molecular weight of the HM fraction; d LMWA; and e LMWN; f sUVa: aromaticity. Black squares: nonirradiated reference, (open circles) low dose (19 J cm-2) irradiated sample, and (open triangles) high does (38 J cm-2) irradiated sample

resemble those derived from HUM (Fig. 4). More bacteria were counted in irradiated compared to non-irradiated samples demonstrating a stronger effect in HUM incubations (Fig. S6). After 8 days of incubation, when bacterial numbers were low, FUKU samples were split into equal aliquots and one aliquot of both (i.e., irradiated and nonirradiated samples) were subjected to the second 19 J cm-2 dose of UVC irradiation. After a lag period of 4 days, bacterial cell numbers began to increase again (Fig. S6). The second UVC irradiation dose had a much more pronounced effect on DOC quality than the first irradiation event. The LMWN fraction accounted to 40% of DOC, but was reduced within 1 week to its initial value of approx. 15%, resulting in HM being the dominating fraction (75%; Fig. S5). Reduction of sUVa and MP with a concomitant increase in the LMWA agrees with the effects observed for the DOC fractions reported above and indicates a rapid microbial degradation. Cluster analysis (Fig. 6) of DGGE banding patterns revealed that bacterial communities slightly differed

between German and Norwegian NOM without irradiation (Fig. 7). The second UVC irradiation resulted in bacterial communities that clearly differed from those of samples irradiated only once (Fig. 6), suggesting that the quality of NOM greatly changes as a response to the second irradiation event. We re-amplified the DNA of excised DGGE bands and identified three major bacterial classes: a-proteobacteria, b-proteobacteria, and Bacteriodetes (Fig. 8). Members of the Burkholderiaceae of the b-proteobacteria were particularly prominent (Fig. 8; Table S1). These bacteria appeared in the irradiated treatments of FUKU and in all HUM samples.

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Discussion The effect of UV light and especially UVC irradiation on NOM has been studied using HPSEC (Frimmel 1998; Lehtola et al. 2003; Parkinson et al. 2003; Buchanan et al. 2006). Except for Frimmel (1998), all other studies


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Dice (Tol 1.0%-11.0%) (H>0.0% S>0.0%) [0.0%-100.0%] 100

95

90

85

80

75

70

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DGGE Humex 16.05.07 irr Humex 23.05.07 irr Humex 30.05.07 irr Fuku 16.05.07 irr Fuku 23.05.07 irr Fuku 30.05.07 irr Fuku 30.05.07 2xirr Fuku 23.05.07 2xirr Humex 16.05.07 Ref Humex 23.05.07 Ref Humex 30.05.07 Ref Fuku 16.05.07 Ref Fuku 23.05.07 Ref Fuku 30.05.07 Ref

Fig. 6 Cluster diagram of DGGE banding patterns for bacteria associated with NOM from Lake Große Fuchskuhle (FUKU) and from the sampling site Humex B (HUM). The dendrogram was obtained by Dice’s correlation index and the unweighted pair-group method with arithmetic means (UPGMA)

detected organic carbon only by its UV absorbance, and therefore provided only restricted information on nonabsorbing carbon fractions. In contrast, the system used here provides information on both UV absorbance and organic carbon distribution of NOM from the same chromatographic run. However, all authors reported similar fractionation patterns of UV-irradiated dissolved organic matter as presented here, which in detail depend on the applied dose, the geometry of irradiation, the type, and the concentration of HS and DOC. For example, Buchanan

1 2

Fig. 5 Incubation experiment with NOM from Lake Große Fuchskuhle (FUKU). a LMWA in relation to DOC derived from HPSEC. b bacterial cell numbers counted by DAPI. FUKU was irradiated in quartz tubes in air equilibrated solution. (squares) FUKU (nonirradiated reference); (circles) FUKU irradiated with high dose (19 J cm-2); open symbols: samples added 0.02% (w/w) NaN3 to stop microbial growth

1 4

time of storage / d

1 3

time of storage / d

Fig. 7 DGGE banding patterns for bacteria associated with NOM from sampling sites Große Fuchskuhle (FUKU) and from Humex B (HUM). Numbers indicate excised and sequenced bands

et al. (2006) analyzed chromatograms of UVC-irradiated surface water samples after fractionation by a sophisticated XAD resin technique, which yielded four fractions: very hydrophobic acids (VHA), slightly hydrophobic acids (SHA), hydrophilic charged molecules (CHA), and hydrophilic neutral molecules (NEU). According to the chromatograms presented and the properties of fractions provided in the text, it appears that VHA ? SHA may correspond to the HM fraction, as well CHA to the LMWA. However, NEU is not identical to the LMWN fraction obtained here, since in our case no UV absorbance of the LMWN was detected. The irradiation by UVC at comparable doses (23–233 J cm-2) reduced mostly the VHA but had no net effect on the SHA. The latter result was interpreted in terms of two overlaying effects including parallel reduction of SHA and accumulation of newly formed material from degradation of VHA. Such an accumulation of SHA-like material could be the clue to the stronger fractionation among the HM fraction that was observed after irradiation (Fig. 3). No long-term HPSEC study so far has investigated the fate of NOM after irradiation. Our experiments demonstrate that especially during the first days of incubation both the irradiated and the non-irradiated dissolved NOM of different origin undergo significant changes. In NOM, rapid reduction of both the LMWA and LMWN fractions was observed. Moreover, after 1 week of incubation, the resulting chromato-grams resembled those of non-irradiated NOM samples (Fig. 3) except MP which remained almost constant. As no comparable reduction was observed in the NaN3-treated samples, it seems most likely that microbes that had survived the irradiation caused this degradation. Although it seems surprising that bacteria can survive UVC doses of 38 J cm-2, our results are in accordance with studies by Lehtola et al. (2003), Buchanan

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Fig. 8 Phylogenetic tree of bacterial 16S rRNA gene sequences associated with humic matter (HM). Sequences from this study are indicated with bold letters and the other sequences were part of the primary analyses (sequences with more than 1,400 nucleotides—number of nucleotides are given after the name of the sequence). GenBank accession numbers are given in parentheses

HM, band 12 (FJ268760) HM, band 7 (FJ268755) HM, band 3 (FJ268751) HM, band 5 (FJ268753) HM, band 6 (FJ268754) HM, band 9 (FJ268757) HM, band 10 (FJ268758) HM, band 4 (FJ268752) Burkholderia fungorum (AF215705) Burkholderia sp (U37344) Burkholderia sp (AJ505302) Burkholderia plantarii (U96933) Burkholderia multivorans (AY486372) Bradyrhizobium sp. (AY187548) uncultured bacterium (AY425767) soil DNB (AB003459) HM, band 11 (FJ268759) HM, band 2 (FJ268750) Caulobacter vibrioides (AY512823)

Burkh olderi a group

Betaproteobacteria

Alphaproteobacteria

HM, band 14 (FJ268762) HM, band 13 (FJ268761) Sphingomonas sp (AY349411) HM, band 1 (FJ268749) Sep Reservoir clone S10.17 (AY752128) uncultured Bacteroidetes (AJ318121) uncultured nanoarchaeote, 1460 uncultured nanoarchaeote, 1460 Nanoarchaeum equitans, 1500 Methanopyrus kandleri, 1451

Bacteriodetes

root

0.10

et al. (2004, 2006) reporting on biofouling as a common problem in water plants after UVC treatment. Sharpless and Linden (2003) demonstrated that small amounts of UVC-absorbing compounds, even H2O2, may reduce light penetration and can thus hamper photodegradation of dissolved compounds or disinfection. Further evidence for the UVC survival of microbes is presented by Coohill and Sagripanti (2008) who reviewed data on UVC inactivation for 38 different bacteria. The typical survival at a dose of approximately 20 J cm-2 would amount to 10%, and even at higher doses [80 J cm-2, a 10-3 fraction of bacteria could survive. Bacteria possibly survive in organisms and bacterial aggregates that act as shields to those further downstream from the beam or in the core of a large clump (Tang et al. 2010). Moreover, a number of repair mechanisms may help to overcome thymine dimer formation, which is the typical damage by UVC exposure. Since we found higher bacterial numbers in irradiated NOM compared to non-irradiated samples, we conclude that our irradiation procedure selects for such microorganisms that both survive UVC irradiation and efficiently utilize the accumulated LMW compounds as the carbon source. Investigation of bacterial communities yielded small differences between the non-irradiated German and Norwegian NOM (similarity of 86% between the reference samples of FUKU and HUM) and bacterial communities that were more similar to each other (81%) after irradiation than to the original communities (60%). Obviously, the harsh UVC treatment selected for a specific bacterial community mainly composed of a-proteobacteria,

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b-proteobacteria, and Bacteriodetes. Members of the Burkholderiaceae have frequently been found in FUKU (Burkert et al. 2003; Allgaier and Grossart 2006; Grossart et al. 2008; Hutalle-Schmelzer et al. 2010), and their activity and abundance seem to correlate with seasonal changes in DOM photoreactivity in the water column. When investigating the effect of elevated singlet oxygen levels and solar irradiation on bacterial community structure in Lake Fuchskuhle, Glaeser and colleagues (2010) frequently found members of the Burkholderiaceae among their isolates that resisted oxidative stress well. Based on these and our own results (I. Salka, pers. com.), we assume that members of the Burkholderiaceae are highly resistant to UVC irradiation and/or reactive oxygen species. The high resistance of natural bacteria to UV irradiation demonstrates their high potential to rapidly adapt to such harmful conditions. A recently isolated strain of Burkholderia from Lake FUKU was exposed for [20 min to harsh UV radiation of a clean bench, which is commonly used for sterilization of the bench. The bacterial strain survived the UV exposure and rapidly recovered its metabolic activity after UV radiation (I. Salkar, pers. com.). In dystrophic Lake Fuchskuhle, low bacterial activities have been measured upon intensive UV radiation; however, due to their high UV-resistance in parallel to frequent mixing of the upper water layer these bacteria can rapidly respond to LMW generated by photoxidation once the light-induced stress has been reduced. Short periods of intensive irradiation are typical for natural lake systems, which are characterized by frequent mixing of the upper water layer.


In Lake Fuchskuhle, mixing of the upper water layer occurs due to convective water movement, which exposes bacteria for a short period of time to high UV irradiation at the water surface. Hence our approach of high UV-radiation could be of great relevance also for lake ecosystems (see Glaeser et al. 2010). On the other hand, a-proteobacteria of the Sphingomonadaceae are common but not dominant in the bacterioplankton of FUKU (Glo¨ckner et al. 2000; Burkert et al. 2003). Although low in abundance, they seem to play an important role in the breakdown of recalcitrant DOC and aromatic compounds, a distinguished feature found in many Sphingomonadaceae (Basta et al. 2005; HutalleSchmelzer et al. 2010). Almost all bacterial phylotypes of this study, including those of the Bacteroidetes, can be found in bacterial communities of acidic and humic matterrich charcoal mining lakes in East Germany (Kampe et al. 2010), suggesting that these bacterial phylotypes are closely associated with humic matter and its degradation products. In our experiments, these bacteria were solely introduced with the respective humic matter extract. Whereas the non-irradiated NOM is characterized by its own distinctive bacterial community, the UVC irradiation resulted in producing a bacterial community that was able to survive and utilize LMW substrates produced by photolysis (see above). This result is in agreement with Langenheder et al. (2006) who report on changes of both bacterial community composition and function after exposure to different intensities of natural UV radiation. The authors hypothesized that photochemical transformations of DOM in surface water triggered bacterial respiration not only by increasing the size of the available substrate pool, but also by favoring a well-adapted bacterial community with specific functional capabilities. Our results shed new light on the observed long-term adverse effects of UVC irradiated NOM (Gjessing and Ka¨llqvist 1991; Lund and Hongve 1994; Paul et al. 2006). In our opinion, the observed adverse effects on E. coli could be the result of a competition between NOM-introduced phyla and E. coli or the preferential metabolite uptake by NOM-specific bacteria. Acknowledgments We acknowledge grant PA 1655/1-1 given to AP and GR 1540/11-1/2 given to HPG from the German Science Foundation, DFG. AP kindly thanks D. Tuma for many helpful discussions.

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