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Keywords:

  • phenol hydroxylases;
  • leaf litter;
  • Platanus acerifolia ;
  • LmPH

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References

Microorganisms are responsible for the decomposition of plant litter due to their enhanced enzyme capabilities. Among extracellular enzymes, those involved in lignin decomposition are especially relevant in leaf degradation. However, the knowledge of the bacterial contribution to the decomposition of phenol-derived compounds in submerged leaf litter is limited. We have used the large unit of the multicomponent bacterial phenol hydroxylase (LmpH) as a genetic proxy to describe changes in the phenol-degrading bacterial community during the decomposition of Platanus acerifolia leaves in a forested stream. Significant differences were found in the phenol-degrading community when three decomposition stages, initial (day 7), midterm (day 58), and late (day 112), were compared. Estimated Shannon's diversity values decreased significantly from 1.93 (initial) to 0.98 (late). According to the deduced amino acid sequences and the corresponding theoretical kinetic parameters of phenol hydroxylases, the initial community showed a low degree of specialization, presumably resulting from random colonization of leaves. At the late decomposition stage, the bacterial community became more specialized, and LmpH genes similar to high-affinity phenol hydroxylases of Comamonas sp. and Burkholderia cepacia increased. The observed changes in the bacterial community suggested an active role of bacteria during litter decomposition in aquatic environments.


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References

In forested rivers and streams, the input of leaf litter from riparian vegetation represents a fundamental organic matter source for microbial decomposers (Pascoal et al., 2003; Gulis et al., 2008). Fungi and bacteria decompose and mineralize plant material, which then enters the river food web (Hieber & Gessner, 2002). The most important microbial enzymes for leaf litter decomposition are those that break down plant fibers, such as cellulases, hemicellulases, pectinases, and phenol oxidases (Sinsabaugh et al., 2002). During leaf litter decomposition, different enzymatic activities may arise in function of the available material in the leaf and of the biodegradability and/or recalcitrance of this material. Because lignin is one of the most recalcitrant compounds, its specific degradation might be a relevant limiting step for complete mineralization of plant material. Major enzymes involved in lignin degradation include phenol oxidases, which oxidize phenols at the expense of oxygen. Phenol oxidase activity has been related to an increase in the relative content of lignin and free phenolic compounds (Sinsabaugh, 2010; Artigas et al., 2011).

Although fungi and bacteria both participate in the decomposition and mineralization of plant material (Moorhead & Sinsabaugh, 2000), the major participation of bacteria is restricted to the later stages of leaf litter degradation only after leaf material has been partially broken down by fungi (Newell, 1993; Baldy et al., 1995; Kominkova et al., 2000). Most works reveal fungi, especially aquatic hyphomycetes, as the dominant players, in terms of activity and biomass increase, during early decomposition of leaf litter in aquatic ecosystems (Baldy et al., 1995; Romaní et al., 2006). However, phenol-degrading bacteria may also be involved in decomposition of recalcitrant plant material in aquatic environments, although their potential role is much less investigated.

Phenol-degrading bacteria are highly adaptive, as observed through the analysis of key functional genes in communities growing in biological wastewater treatment plants (Futamata et al., 2003; Basile & Erijman, 2010). Phenol hydroxylases, which convert phenol into catechol derivatives via hydroxylation, are specific phenol oxidases generally involved in the degradation of organic compounds. These enzymes have been extensively studied at the molecular level, and they can now be detected in natural samples by high-throughput analytical methods. Multicomponent phenol hydroxylases (mPHs) are considered to be predominant in nature (Nordlund et al., 1993; Watanabe et al., 2002). The largest subunit of multicomponent phenol hydroxylases (LmPHs) has been used as a molecular marker to assess the functional and genetic diversities of biotechnologically relevant phenol-degrading bacteria (Futamata et al., 2005; Viggor et al., 2008). Moreover, phenol-degrading bacteria have been isolated and characterized from the phyllosphere of trees showing that leaves may contain a significant bacterial diversity with respect to LmPH sequence similarities (Sandhu et al., 2009). However, to the best of our knowledge, no experimental report exists describing the change in the bacterial phenol-degrading community during leaf litter by the use of selected molecular markers targeting to functional genes.

In this study, we have used the LmpH gene as a molecular proxy to analyze the changes in the phenol-degrading bacterial community during the decomposition of submersed Platanus acerifolia [Aiton] Willd. leaves in a forested stream. We hypothesize that phenol-degrading bacteria might contribute to leaf litter breakdown and that their community structure might change throughout the decomposition process as higher amounts of free phenolic compounds are available. To test this hypothesis, three discrete sampling dates were chosen according to mass weight and enzymatic activity data from a previous experiment of leaf litter decomposition. Selected samples covered the main observed changes in microbial activity and biomass. The observed changes of the bacterial community indicate that a specialization of potential phenol-degrading bacteria exists during the decomposition of leaves.

Materials and methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References

Sample preparation and analysis

Platanus acerifolia leaf samples were obtained from a litter decomposition experiment performed in an oligotrophic forested stream (the Fuirosos, NE Spain, latitude 41°42′N, longitude 2°34′E; Artigas et al., 2011). Briefly, recently fallen leaves were placed in leaf litter bags and immersed in the stream; samples were collected intensively for bacterial biomass and enzymatic activity until day 112 after immersion. Leaf samples were collected, rinsed with filtered stream water (0.2 μm), and cut to disks (1.1 cm diameter) with a metal borer. For phenol oxidase activity assays, disk samples were kept at 4 °C until analyzed in the laboratory (within 20 h). Samples for the determination of bacterial density were fixed with formaldehyde (2%). Finally, samples for molecular analyses were stored frozen (−20 °C).

Bacterial densities were estimated according to the protocol of Porter & Feig (1980). Leaf disks were sonicated (2 + 2 min) in an ultrasonic bath (40 W power, 40 kHz frequency; Selecta, Spain), diluted (1 : 4), and stained for 5 min with 4, 6-diamidino-2-phenylindole (DAPI) at a final concentration of 2 μg mL−1. Bacterial suspensions were, then, filtered through 0.2 μm irgalan black–stained polycarbonate filters (Nuclepore; Whatman International Ltd., Maidstone, UK) and counted using a fluorescence microscope (Nikon Eclipse 600W, Tokyo, Japan) under ×1250 magnification. Bacterial densities were transformed into biomass units based on 2.2 × 10−13 g C μm3 conversion factors (Bratbak & Dundas, 1984) and using a mean bacterial biovolume of 0.163 μm3 (J. Artigas, unpublished data).

Phenoloxidase enzyme activity (EC 1.10.3.2 and 1.14.18.1) was determined using L-3,4-dihidroxyphenylalanine (L-DOPA) substrate and following the methodology described by Sinsabaugh et al. (1994).

Nucleic acids extraction and PCR conditions

Triplicate leaf samples from each sampling date were pooled for the DNA extraction. The DNA was extracted from 100 to 200 mg of lyophilized leaf material. Nucleic acids were extracted with the FastDNA® SPIN for Soil Kit (MP Biomedicals) following the instructions provided by the manufacturer, with the following modifications. The homogenizing step was repeated three times in a FastPrep Instrument (MP Biomedicals) using cycles of 30 s at a speed setting of 5.5. Samples were placed on ice for 5 min between every homogenizing step.

The LmPH gene was amplified in a GeneAmp PCR system 2700 with the primer pair PheUf/PheUr (Futamata et al., 2001). PCR mixtures contained 1× PCR buffer, 1.5 mM MgCl2, 200 μM total dNTPs, 0.5 μM of each primer, 10 ng of the DNA extracts, and 0.5 units of Taq polymerase (Go Taq; Promega, Madison, WI) in a total volume of 30 μL. Amplification reactions were carried out exactly as previously described (Futamata et al., 2001). PCR products were analyzed by electrophoresis on 1.5% agarose gels and visualized after staining with ethidium bromide (0.2 mg L−1).

Cloning and sequencing

The analysis of LmPH gene diversity was determined through cloning experiments. Before cloning, PCR products were cleaned by QIAquick® PCR Purification Kit (Qiagen, Germany) according to the manufacturer instructions. Cloning experiments were conducted using the pGEM-T® Easy vector (Promega). Ligated products were transformed into Escherichia coli TOP10 competent cells, and positive transformants were color-screened on LB plates supplemented with ampicillin (100 μg mL−1), X-Gal (80 μg mL−1), and isopropyl-beta-d-thiogalactopyranoside (IPTG 0.5 mM). Clones were selected using primers M13F-20 and M13R and selected according to the expected size (620 bp) of the amplified LmPH gene fragment. Positive PCR products of the expected size were sequenced using the vector-specific primer M13F-20 at the Macrogen service (Macrogen, Seoul, South Korea). Sequences were manually refined using the BioEdit package. Amino acid-derived sequences were further aligned using clustalw.

Community and phylogenetic analyses

Amino acid-derived sequence alignments of partial LmPH were used to construct a distance matrix using the online package implemented in mothur v1.13 (Schloss et al., 2009). Rarefaction curves were calculated at a cutoff value of 90% similarity and were used to determine the number of operational taxonomic units (OTUs) in each sample. A 90% cutoff value of the LmPH gene approaches a species-level OTU definition according to comparisons between available 16S rRNA and LmPH gene sequences of cultured phenol oxidizers (results not shown). Estimated richness (SChao, and SAce), Shannon diversity index (H′), and evenness (E′) indices were calculated according to the OTUs distribution. Jaccard similarity coefficients were calculated pairwise by using either the presence of shared OTUs between two different communities (OTU based approach) or the relative abundance of individuals that belong to shared OTUs (abundance-based test).

Phylogeny was reconstructed using mega v.4. The Amino Poisson correction and pairwise deletion methods were used. Bootstrap analysis was conducted with 1000 replications. Additionally, to estimate the diversity between different bacterial communities using the phylogenetic information, UniFrac (UniFrac weighted algorithm) and parsimony tests were calculated using the above phylogenetic tree. The outcomes of these analyses reflect the evolutionary distance between the members of the analyzed bacterial communities (Lozupone et al., 2011).

LmPH sequences obtained in this study have been submitted to GenBank under accession numbers JF806548JF806617 and JQ069975JQ070053.

Results and discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References

Sample selection for diversity analyses

During the duration of the whole experiment (112 days), a significant relationship between leaf bacterial biomass and phenol oxidase activity was observed, suggesting a link between bacteria and degradation of phenols in leaves (Fig. 1). To investigate the potential role of phenol-degrading bacteria, three dates were selected for molecular analysis of the largest subunit of multicomponent phenol hydroxylases (LmPHs). The three dates corresponded to three distinct stages throughout the leaf decomposition sequence: a initial stage (day 7), coinciding with the initiation of bacterial colonization; a midterm stage (day 58), coinciding with increasing bacterial biomass and phenol oxidase activity and the maximum fungal biomass (Artigas et al., 2011); and a late stage (day 112), when the maximum bacterial biomass was measured although phenol oxidase slightly decreased (Fig. 1).

image

Figure 1. Linear regression analysis between phenol oxidase activity and biomass of bacteria accumulated in Platanus acerifolia leaves throughout the decomposition process at the Fuirosos stream (data obtained from Artigas et al., 2011). The R square (r2) and probability (P) values after the linear regression analysis are shown. The initial (day 7), midterm (day 58), and late (day 112) stage samples selected for the analysis of LmPH genes are indicated in arrows.

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Phylogenetic analysis of bacterial phenol hydroxylases

The LmPH gene was amplified by PCR from the three leaf litter decomposition stages, and a total number of 148 good quality sequences were obtained from cloning experiments. The estimated rarefaction curves in each sample approached saturation, indicating a good coverage of LmPH gene richness (Fig. 2). All subsequent analyses were performed using an OTU-based approach of the deduced amino acid sequences at a 0.1 cutoff level. The analysis of sequences from the three stages resulted in 16 different OTUs, nine of them being specific for either the initial or the midterm stage. OTU 14 was the most abundant and contained LmPH sequences from the initial (11 sequences), the midterm (22), and late (33) stages. The second most abundant OTU 3 (12 sequences) was exclusively composed of sequences from the initial stage. Other highly represented OTUs, such as OTUs 15 and 16, grouped exclusively sequences from the midterm and late decomposition stages.

image

Figure 2. Rarefaction curves for amino acid-derived LmPH sequences at a cutoff value of 10% dissimilarity obtained for the three leaf litter decomposition stages (●) initial, (○) midterm, and (▼) late. Pairwise distances among sequences were conducted using the Poisson correction model. The coding data were translated assuming a standard genetic code table. All positions containing gaps and missing data were eliminated. There were a total of 138 amino acid positions in the final sequences.

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The potential functional differences between communities over the course of leaf decomposition were investigated by deducing kinetic properties of bacterial phenol hydroxylases. LmPH genes can be assigned to different functional groups according to changes at selected positions of the amino acid sequence (Futamata et al., 2001). Key amino acid residues at positions 217, 252, and 253 (position numbering based on the Pseudomonas sp. CF600 dmpN gene sequence) may facilitate the prediction of theoretical Michaelis–Menten semi-saturation constants for most uncultured microorganisms (Viggor et al., 2008). Most of the retrieved sequences (86) belonged to the betaproteobacteria low-Ks LmPH group, previously defined by Futamata et al. (2001) and grouped separately into clusters A and E (Fig. 3). LmPH sequences in cluster A showed significant similarities (> 80%) to phcN, tbc1D, and afpN genes from Comamonas testosteroni, Burkholderia cepacia, and Alcaligenes faecalis, respectively. On the other hand, cluster E contained LmPH sequences with high similarity with phenol-degrading genes from Comamonas sp. and Alicycliphilus sp. Sequences from the three stages appeared in both clusters, although those from the late stage were less abundant in cluster E.

image

Figure 3. Neighbor-joining phylogenetic tree based on the derived amino acid sequences of LmPH genes from samples of Platanus acerifolia leaves at different stages of leaf decomposition at the Fuirosos stream. Bootstrap values (> 50%) based on 1000 trials are indicated at nodes. Phylogenetic trees were reconstructed using neighbor-joining and maximum-likelihood methods yielding similar tree topologies. Phylogenies were reconstructed using the amino Poisson correction and complete deletion methods. Reference LmPH sequences (bold) were obtained from blast searches within the GenBank reference genomic sequences database. Clusters A, B, C, D, and E were defined according to potential activity of deduced amino acid residues at positions 217, 252, and 253 based on the Pseudomonas sp. CF600 dmpN gene sequence (accession number P19732). Terminal tree nodes containing LmPH sequences obtained in this study have been collapsed and indicated by the OTU number. The number of sequences of the initial (INI), midterm (MID), and late (LATE) samples in every OTU is indicated.

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All sequences in cluster B except one (LATE13_E10) were retrieved from the initial and midterm stage samples. Sequences in this cluster exhibited high sequence diversity and grouped into eight different OTUs. Higher similarities (84–94%) were found to LmPH sequences retrieved from noncultured microorganisms from benzene-contaminated soils or trichloroethylene-contaminated aquifers. Maximum similarities to isolated representatives (< 84%) were obtained with Methylibium petroleiphilum strain PM1, a Betaproteobacterium that degrades methyl tert-butyl ether (Hristova et al., 2007). Although degradation of phenolic compounds has not been studied in detail in PM1, exposure of this strain to MTBE induces additional pathways for the degradation of aromatics such as benzene, toluene, and xylene. In a recent study employing PCR-denaturing gradient gel electrophoresis (DGGE) analysis of reverse-transcribed rRNA, active M. petroleiphilum was shown to accumulate in soils contaminated with penta-chlorophenol (Cáliz et al., 2011).

The specific Variovorax group (cluster C) was also represented by two sequences obtained from the midterm stage (sequences MID06_F3 and MID06_G7, OTU 7). Nevertheless, these two sequences were < 85% similar to Variovorax sp. HAB30. The ecological relevance of Variovorax sp. relies in the presence of a characteristic LmPH type, corresponding to highly active phenol-degrading enzymes with high semi-saturation constants according to determinations of kinetic parameters using isolated cultures (Futamata et al., 2005). Cluster D grouped sequences belonging to Gammaproteobacteria with a high-Ks LmPH, including Pseudomonas putida relatives. A single sequence from the initial stage (sequence INI06_A3, OTU 1) was found in cluster D. Interestingly, this sequence contained the typical signature of low-Ks phenol hydroxylases at amino acid positions 252 and 253, and position in the high-Ks group should be confirmed by incubation experiments with isolated cultures.

Changes in the phenol-degrading bacterial community

The number of bacterial OTUs remained at relatively low values (from 5 to 10) in the three samples analyzed. The bacterial community at the initial and midterm stages of decomposition showed a greater richness, greater diversity (Shannon's H′), and greater evenness (E) of LmPH gene compared to the late stage (Table 1). The significant decrease in richness and diversity values suggests a major specificity of phenol-degrading bacteria in the late-stage community. The results from the phenol-degrading bacterial community analysis showed a highest degree of specialization at the late decomposition stage. All LmPH genes obtained at the late stage, except for one, grouped in clusters A and E together with sequences belonging to known high-affinity phenol degraders (Watanabe et al., 1996). On the contrary, at the initial stage, the lower bacterial biomass and weaker phenol oxidase activity may indicate that decomposition of the large recalcitrant plant molecules had not yet begun (Fig. 1, Artigas et al., 2011). At this first stage, bacterial communities are supposed to be defined by environmental conditions of the stream and random colonization of the leaf surface (Harrop et al., 2009; Marks et al., 2009).

Table 1. Main community indicators: observed richness (S), estimated richness (SChao, and SAce), Shannon's diversity index (H′), and evenness (E′) of the phenol-degrading bacterial communities in Platanus acerifolia leaves at the initial (day 7), midterm (day 58), and late (day 112) stages of leaf decomposition at the Fuirosos stream
Decomposition stage S S Chao S Ace HE
Initial99.510.21.93 ± 0.190.74
Midterm101010.31.85 ± 0.280.46
Late555.70.98 ± 0.280.40
Total1616.516.82.06 ± 0.150.73

Differences in the community composition of potential phenol-degrading bacteria were tested from the tree topology using UniFrac and parsimony tests. All pairwise comparisons between samples were highly significant indicating a changing bacterial community at the three degradation stages (Table 2). UniFrac distances ranged from 0.298 to 0.607 and were higher between the initial and late stage samples. UniFrac tests have been previously used as a semi-quantitative determination of the similarities between the bacterial communities on the phyllosphere of Populus deltoides sampled at different times (Redford et al., 2010). According to our estimations, major changes in the phenol-degrading bacterial community may occur between the initial and midterm stages of leaf decomposition. At the midterm, the greatest community richness and diversity was found and coincided with increasing phenol oxidase activity and maximum fungal biomass (Artigas et al., 2011). The LmPH sequences from this stage were scattered throughout the phylogenetic tree (in clusters A, B, C, and E), and their corresponding enzymes exhibit different kinetic properties. It is known that bacteria and fungi have complementary roles in leaf litter degradation. Bacteria are thought to increase their contribution only after leaf material has been partially broken down (Baldy et al., 1995), whereas fungi, especially aquatic hyphomycetes, have been recognized as dominant, in terms of both activity and biomass, during early decomposition (Gulis & Suberkropp, 2003; Romaní et al., 2006). However, bacteria may make a greater contribution to leaf litter decomposition particularly when fungal activity is compromised by unfavorable conditions (Pascoal & Cassio, 2004; Kubartova et al., 2009).

Table 2. Values of diversity (weighted and unweighted UniFrac and parsimony tests) of pairwise comparisons of the phenol-degrading bacterial communities in Platanus acerifolia leaves at the initial (day 7), midterm (day 58), and late (day 112) stages of leaves decomposition at the Fuirosos stream
 Initial–midtermMidterm–lateInitial–lateInitial–midterm–late
  1. Tests were conducted using the amino acid-derived sequences phylogenetic tree in Fig. 3.

  2. n.a., Not applicable.

  3. *< 0.01; **< 0.001.

Unweighted UniFrac test scoren.a.n.a.n.a.0.580**
Weighted UniFrac test score0.473**0.298**0.607**n.a.
Parsimony test score13*25*8**n.a.

In conclusion, by analyzing the LmPH gene from different leaf decomposition stages, we have shown that the bacterial community changes significantly over the course of leaf litter degradation in streams. During early decomposition, the bacterial community is rather complex and potentially exhibits a low degree of metabolic specialization in view of the deduced enzyme kinetics. As decomposition progresses, the phenol-degrading bacterial community is dominated by suspected low-Ks type bacteria, with a high similarity to Alcaligenes spp., Comamonas sp., and Ralstonia sp, suggesting a gradual selection of specialized phenol degraders as decomposition progressed. To the best of our knowledge, this work represents the first specific analysis of any functional gene marker and of bacterial and fungal origin, used for investigating microbial communities during the leaf litter decomposition process in streams. Time series analyses of bacterial and fungal communities in leaf litter decomposition have previously been performed using either DGGE or terminal-restriction fragment length polymorphism (T-RFLP) of amplified SSU rRNA fragments (Das et al., 2007; Marks et al., 2009; Kelly et al., 2010), although no general conclusions can be derived from these studies. The relative presence of general and specialized microorganisms on leaf surfaces during litter decomposition has been proposed as a major determinant of diversity (Das et al., 2007). Moreover, leafs of different plant species have been shown to bear specific fungal and bacterial communities during the decomposition process in riverine environments (Marks et al., 2009). In this work, we show that the use of functional genes, as the bacterial LmPH gene, as a proxy to study microbial diversity of relevant microorganisms in leaf litter decomposition is possible. We are confident that the use of other functional genetic markers of bacteria, and its extension to the study of fungi, will provide additional and interesting results to support the idea of changing microbial communities in the process of litter decomposition and increase our understanding of how microorganism interacts in ecosystem processes.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References

The authors acknowledge the contribution of Anna Díez to laboratory work. This research was financially supported by the Spanish Government through projects CGL2009-08338 and CGL2011-30151-C02-01.

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References
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