Constance
Melanie Hempel, Hans–Peter Grossart & Elisabeth M. Gross
ABSTRACT: Submerged macrophytes provide vast surfaces for the settling of microorganisms. Those biofilms contribute substantially to the overall nutrient cycling in the littoral zones and interact in multiple ways with macrophytes and other epiphytes. Although studies on aquatic biofilms are numerous, aquatic macrophytes and their heterotrophic biofilms have rather been neglected. During two seasons, we investigated the bacterial community composition (BCC) on two submerged macrophytes with DGGE and with a clone library. We followed the spatial BCC on Myriophyllum spicatum in Lower Lake Constance in 2005 and found differences between the apices and the leaves. These differences lessened towards autumn. In 2006, we investigated the substrate–specific BCC on M. spicatum, Potamogeton perfoliatus and polypropylene sheets. The comparison between all substrates also exhibited a distinct BCC on M. spicatum apices. On the leaves of both plants, the BCC was rather similar, also compared to the artificial substrate. A comparison between the BCC on mesocosm M. spicatum and artificial substrates gave similar results as in the field. Bacterial sequences from excised DGGE bands and the clone library were mainly affiliated with yet uncultured clones originating from various freshwater habitats. We also found bacteria capable of degrading phenolic and aromatic compounds. Our results indicate that the BCC on M. spicatum apices is rather unique and may point to specific bacterial functions in this microenvironment.
To better elucidate these functions, further studies are needed, preferentially with cultivation–based approaches, which allow for more detailed physiological studies of biofilm bacteria.
Keywords: Myriophyllum spicatum, Potamogeton perfoliatus, DGGE, phenolic
INTRODUCTION
Several studies have shown that plants are no neutral substrates for epiphytic algae and cyanobacteria, and that some plants might influence the density and composition of their autotrophic biofilm, while a dense biofilm formation may severely hamper the growth of submerged macrophytes (Phillips et al. 1978, Eminson & Moss 1980, Blindow 1987). Comparable studies for the heterotrophic biofilm on freshwater macrophytes are scarce. We were interested whether substrate–specific differences exist in the heterotrophic bacterial community composition (BCC) between artificial and natural substrates. We chose the submerged macrophytes Myriophyllum spicatum L. (Haloragaceae) and Potamogeton perfoliatus L. (Potamogetonaceae) as the two natural substrates. Both are common in lakes of different trophic state, also in the oligo– to mesotrophic Lower Lake Constance, Germany. Myriophyllum spicatum is known for its allelopathic activity against algae and cyanobacteria caused by hydrolysable polyphenols (Gross et al.
1996, Leu et al. 2002). Many species of the Potamogetonaceae, among them P. perfoliatus, contain antibacterial compounds, although none has been identified so far for P. perfoliatus (Bushmann & Ailstock 2006).
Both plant species differ in their leaf structure, growth form, life–cycle and chemical composition. Myriophyllum spicatum has pinnate leaves with individual filaments of about 1 mm diameter. In Lake Constance, it forms small and distinct stands of 10 – 15 shoots with a length of 30 – 100 cm. Potamogeton perfoliatus has laminar, oval leaves and grows in stands of several square meters up to the surface with shoot length up to 4 m, resembling ‘underwater forests’. In Lake Constance, M. spicatum starts growing in late June and declines not before November.
Potamogeton perfoliatus emerges already in May and is senescing in September to October. Our long–term analysis shows that the chemical composition of M. spicatum in Lake Constance exhibits seasonal patterns and a gradient of macro– and micronutrients and phenolic compounds from the apices to the older leaves, while P. perfoliatus contains very little phenolic compounds (Choi et al. 2002), and in
general exhibits no pronounced gradients in macronutrients (Gross et al., unpublished; this study).
Submerged macrophytes do not only provide shelter and nutrition for other organisms, they also structure the littoral zone, prevent sediments from re–
suspension and may change the nutrient composition of the water column (Stoner 1980, Jeppesen et al. 1998). They provide a vast surface for the settlement of microorganisms, algae and meio– and macrofauna. Bacteria in biofilms may have beneficial effects for the plants: they can attract zoospores, enhance and restore growth or growth form of plantlets and produce metabolites beneficial for the plant (Joint et al. 2000, Marshall et al. 2006). Secondary metabolites of bacteria associated with terrestrial plants may reduce herbivory (Hoagland 2001, Sturz & Kimpinski 2004). Yet, dense layers of autotrophic epiphytes decrease the light and nutrient availability of submerged macrophytes (Sand-Jensen 1990). Further, sedimentation and particle retention are a source of nutrients both for macrophytes and epiphytes.
Thus an optimum between nutrient supply and light availability for photosynthesis has to be kept (Schulz et al. 2003).
However, also the plant may influence its epiphytes. Secondary metabolites of red algae interfering with quorum sensing may inhibit bacterial attachment (Maximilien et al. 1998, Nylund et al. 2005). Plant polyphenols similar to those present in M. spicatum apparently hamper the quorum sensing regulated biofilm formation (Huber et al. 2003). The antimicrobial properties of polyphenols are due to their ability to chelate proteins, nutrients and iron (Scalbert 1991). These allelochemicals also inhibit gut microbiota of herbivores feeding on M. spicatum (Walenciak et al.
2002). If polyphenols would interfere with epiphytic bacteria, this could lead to a reduced biofilm formation and in turn to an increased light and nutrient availability for the plant.
So far, only a few studies dealt with the heterotrophic biofilm composition on freshwater macrophytes. They often focus on marine and brackish species or are restricted to certain bacterial functions such as nitrification and denitrification
(Eriksson & Weisner 1999). On the chlorophytes Desmidium grevillii, Hyalotheca dissiliens and Spondylosium pulchrum bacteria belonging to the CFB phylum and alpha–, beta– and gammaproteobacteria were found (Fisher et al. 1998), and the obtained sequences were not closely related to known taxa. Bacteria belonging to the Flavobacteria–Sphingobacteria group were most abundant on marine diatoms (Grossart et al. 2005). Bacterial biofilm community composition and the reaction of individual taxa were selectively influenced by the type of dissolved organic matter provided (Olapade & Leff 2006). Submerged plants as producers of dissolved organic matter might thus influence bacterial community composition. Cultivation–based assays detected gammaproteobacteria and Cytophaga–Flavobacteria in the biofilm on M. spicatum (Chand et al. 1992). The biofilm community composition on Potamogeton crispus in comparison to cellulose filters yielded variable numbers of alpha–, beta–
and gammaproteobacteria (Hong et al. 1999). Molecular studies investigating aquatic surfaces have mostly been on artificial substrates (Brummer et al. 2000, Olapade &
Leff 2006), algae (Rao et al. 2006), among them diatoms (Grossart et al. 2005). Others investigated the heterotrophic biofilm on macrophytes with cultivation dependent techniques (Chand et al. 1992), which can be rather selective depending on the growth conditions applied. In the last decade, our knowledge on microbial communities increased continuously, mainly due to the establishment of new molecular methods such as FISH (fluorescence in situ hybridisation) and DGGE (denaturing gradient gel electrophoresis). These methods allow the investigation of bacterial communities without cultivation (Head et al. 1998).
By using DGGE and clone library, we investigated whether the biofilm BCC differed between different parts of the same plant, different macrophytes and/or substrates. We expected differences between the apices and the lower leaves on M. spicatum reflecting the unique chemical gradient of this plant (Hempel & Gross, submitted, Gross et al., unpublished). Further, we propose that the biofilm BCC on the polyphenol–rich M. spicatum is distinct from the polyphenol–free P. perfoliatus and an artificial substrate. Plant quality differences between plants and upper and
lower leaves was analysed by measuring carbon, nitrogen, phosphorus and chlorophyll content as well as total phenolic content, anthocyanins and, only in M. spicatum, the hydrolysable polyphenol tellimagrandin II. In 2005 we investigated the spatial differences in biofilm BCC between younger and older plant parts of M. spicatum. In summer 2006, we extended our study to the comparison of the BCC on three different substrates: M. spicatum, P. perfoliatus and polypropylene sheets, with extensive plant chemistry. To get a more detailed community composition we constructed a clone library of the biofilm on the apices of M. spicatum.
MATERIALS AND METHODS
Sampling site. We sampled submerged macrophytes during the growing season in 2005 and 2006 near the Isle of Reichenau in Lower Lake Constance, Germany (N47°42.247, E9°02.289). In July, August and October 2005, we investigated whether the biofilm community composition on Myriophyllum spicatum differed between younger and older plant parts. Triplicates of three different plant stands were analyzed. In 2006, we investigated the BCC on the two submerged macrophytes M. spicatum and Potamogeton perfoliatus and polypropylene sheets as an artificial substrate. The artificial substrates were deployed in 2.6 m depth two weeks before the sampling campaign started on 17 July. They were made out of 0.3 mm thick polypropylene sheets cut to the size of 9.7×1.2 cm and punctuated at each end with a hole–puncher. A floater was tied to one end for an upright position; the other was fixed with a lace to a plastic bar fixed to the ground by tent pegs. Myriophyllum spicatum, Potamogeton perfoliatus and artificial substrates were sampled in triplicates by snorkelling in a depth of 1.5 – 2 m every two weeks between 17 July and 09 October 2006. Plants and artificial substrate samples were stored individually in sterile 50 ml polyethylene tubes at 4 °C until processing started at latest 24 hours later. Additionally, we deployed artificial substrates in an outdoor mesocosm densely stocked with M. spicatum. Samples of the mesocosm plants (apex and leaf)
and artificial substrates were taken on the same dates as in the field. Only samples from September and October were analysed.
For chemical analysis, plants were stored in plastic bags, three replicates consisting of at least five plants from one stand. Samples were stored at 4 °C until analysis the next day. At each sampling date, temperature, oxygen, conductivity and pH were measured in the water column 20 cm below water surface.
Detachment of epiphytic biofilm. In the laboratory, the plant length was measured and the overall state of the plant was recorded. Artificial substrates were documented by photography and approximately 2 cm of the middle part were used for biofilm analysis. The apex and 13 leaves of the 11 – 25 cm shoot section (lower leaf) of both plant species were taken and transferred to 15 ml sodium pyrophosphate (0.1 M Na4P2O7×10 H2O). Since the leaf surface of Potamogeton perfoliatus was much larger, only five leaves were sampled. The biofilm of all samples was detached by 1 min ultrasonication (Laboson 200 ultrasonic bath, Bender &
Hobein, Germany), 15 min of shaking (18.3 Hz, horizontal shaker Eppendorf, Germany) and subsequent ultrasonication for 1 min. The detachment of epiphytic bacteria had been optimized before as described in (Hempel et al. 2008). The suspension containing the detached biofilm was filtered onto ME 24 membrane filters (0.2 µm; Ø 45 mm, Schleicher & Schuell) for DNA extraction, and stored at –20
°C until use.
DNA extraction. Filters were cut into small pieces and DNA was extracted following a standard phenol/chloroform protocol with an additionally lysozyme step (8 mg ml–1; 260 µl sample–1; 30 min at 65 °C; (Walenciak 2004)). Extracted DNA was dried, taken up in 40 µl of filter sterilized Millipore water and quantified.
Polymerase chain reaction (PCR). PCR was performed in a Thermocycler T–
Gradient (Biometra, Germany). We used the primers 341f [5`–CCT ACG GGA GGC AGC AG–3`(Muyzer et al. 1993)] and 907r [5`–CCG TCA ATT CMT TTG AGT TT–
3`(Lane et al. 1985)]. To perform DGGE, primer 341f was supplemented with a GC–
clamp [5`–CGC CCG CCG CGC CCC GCG CCC GTC CCG CCG CCC CCG CCC–3´
(Muyzer et al. 1995)]. One 50 µl PCR reaction contained 5 µl PCR–buffer (10×Taq–
buffer, Eppendorf, Germany), 5 µl dNTP–Mix (500 mM, Eppendorf, Germany), 0.5 µl of the forward primer at 25 pmol µl–1, 0.5 µl of the reverse primer at 25 pmol µl–1, 3 µl MgCl2 (25 mM, Eppendorf, Germany), 10 µl 6 mg ml–1 BSA (Sigma) and 1 U Taq polymerase (0.2 µl, Eppendorf, Germany). The following protocol was used for amplification: (1) 5 min 95 °C; (2) 1 min 95 °C; (3) 1 min 55 °C; (4) 2 min 72 °C; repeat (2) – (4) 29×; (5) 15 min 72 °C. We did not retrieve PCR products from all replicates, probably due to the high polyphenol content in M. spicatum plants resulting in variable replicate numbers.
Denaturing gradient gel electrophoresis (DGGE). DGGE was performed in an INGENY PhorU system (Ingeny). For better comparison of DGGE banding patterns, equal amounts of PCR products (ca 50 ng) were loaded to the gel and an external standard was used. DGGE was performed in a 7% (v/v) polyacrylamide gel with a denaturing gradient of 40 to 70% urea and formamide, and run at 60 °C for 20 hours.
Gels were stained with 1× SybrGold (Invitrogen), washed in deionised water and documented with an AlphaImager 2200 Transilluminator (Biozym) under UV light.
Bands were excised from the gel with a sterile scalpel and immediately transferred to a sterile PCR cup, in which DNA was eluted with sterile water. DNA was amplified using the primer pair 341f – 907r (without GC–clamp) and conditions as described above. Sequencing was performed by 4base lab, Reutlingen, Germany. DGGE gels were analyzed with the software GelCompar II version 3.5 (Applied Maths, Belgium). Cluster analysis was performed with Pearson correlation using the unweighted pair group method with arithmetic mean (UPGMA).
Clone library. In May 2006, apices of M. spicatum grown in an outdoor mesocosm (a concrete basin of 2×2×1 m) were used to analyze the BCC with a clone library. The mesocosm had a flow–through with Lake Constance water of approx. 20 l h–1, and the plants were subjected to the same climatic conditions as in the field except that the water level was kept constant. Biofilm detachment, DNA extraction and PCR conditions were the same as described above, except that PCR was performed with
primers 27f (5`–AGA GTT TGA TCC TGG CTC AG–3`) and 1492r (5`–TAC GGY TAC CTT GTT ACG ACT T–3`). We used the TA Cloning Kit by Invitrogen. Instead of sequencing each single clone, we separated distinct clones by performing a restriction fragment length polymorphism (RFLP) with MspI (0.08 U sample–1, 37 °C over night). Only clones exhibiting different banding patterns were sequenced (4base lab, Reutlingen, Germany).
Statistics. We assumed that plant chemistry and environmental conditions might influence the biofilm community composition. Thus, biofilms experiencing comparable conditions should display a similar community composition. We related the DGGE banding patterns to plant chemistry (Hempel & Gross, submitted) and environmental conditions with a BEST–ENV analysis to see which factors explain the differences between both plants best. First of all, the DGGE banding patterns were transformed into a presence-absence matrix. With these data a dissimilarity matrix was calculated based on Bray Curtis dissimilarity. A second dissimilarity matrix was calculated for standardised environmental data with Euclidean distance. For the plant chemistry, we chose tissue nitrogen, carbon, phosphorus, chlorophyll and total phenolic content and as environmental factors water level, temperature, conductivity and pH. The data were normalised to allow a comparison between different units.
This means, that all data are placed on a common scale by subtracting the mean of each variable from each value and divide the product by the standard deviation. This yields values in the range of –2 to +2. The ranks of both matrices were compared by Spearman rank coefficient (ρ) to find the best match between them. To provide statistic validation, 999 permutations were carried out. These analyses were performed with Primer 6 (Version 6.1.6, Primer E Ltd.).
RESULTS
Environmental variables and plant condition
Environmental conditions changed during the sampling period from July to October (Table 3.1). The temperature dropped from the beginning to the end by about 10 °C. The water level at the sampling dates was more or less constant at 319 cm, but was at maximum 25 cm higher or 27 cm lower. Conductivity and pH were also relatively constant (267 ± 14 µS cm–1 and 8.3 ± 0.2, respectively).
Throughout the sampling period in 2006, Myriophyllum spicatum shoots were 30 – 45 cm long and exhibited dark green leaves, the typical red coloured stems, and were not densely colonized with epiphytic algae. Potamogeton perfoliatus shoots were 20 – 50 cm long and during summer had intact, brightly green coloured leaves. Both plant species did not show severe signs of grazing. While M. spicatum did not show any sign of senescence throughout the sampling period, P. perfoliatus had brownish leaves at the last two sampling dates. At the beginning of the sampling period, the artificial substrates were covered with a thin layer of bacteria and algae and with increasing exposition time by several layers of Dreissena polypmorpha (zebramussel;
Mollusca).
Table 3.1. Environmental variables measured at the sampling dates.
Water level data are of the water gauge at the harbour in Konstanz.
Sampling
15. August 3 18.7 324 265 8.53
29. August 4 17.6 321 263 8.27
12. September 5 20.0 324 263 8.06
22. September 6 18.9 329 274 8.41
09. October 7 16.2 322 286 8.01
23. October 8 15.6 292 282 8.23
Denaturing gradient gel electrophoresis (DGGE)
Biofilm bacterial community composition on apices and leaves of Myriophyllum spicatum (summer 2005). In July and August, the DGGE banding pattern differed between apices and leaves. Most of the apices samples clustered together separately from the leaf samples (Figure 3.1A and B). In both month, however, some replicates diverged and clustered with the respective other plant part. In October 2005, the differences between apex and leaves were less apparent (Figure 3.1C) indicating changes in the plants’ physiology. In July, the bacterial diversity based on the banding pattern seemed to be highest on M. spicatum leaves. In contrast, the BCC of the apices was less diverse, and exhibited a higher variability of the banding pattern than that of the leaves (Figure S3.1A, see supplementary). These differences seemed to fade when macrophytes aged (Figure S3.1B, see supplementary).
Figure 3.1. Cluster analysis of DGGE banding patterns of Myriophyllum spicatum samples in 2005.
A) July B) August C) October as determined by unweighted pair group method with arithmetic mean (UPGMA).
Biofilm community composition on different substrates. In summer 2006, DGGE banding patterns of plant apices revealed that the bacterial biofilm on M. spicatum differed from that on P. perfoliatus and the artificial substrate (Figure 3.2A). Most of the P. perfoliatus apex and artificial substrate samples clustered together, whereas
apex samples of M. spicatum formed a distinct cluster. Only both plant samples from the end of August differed from all other samples. The BCC on the leaves of both macrophytes and the artificial substrates was rather similar (Figure 3.2B) and a succession in BCC was not observed neither in apices nor leaves or artificial substrates.
In the mesocosm, we observed a distinct cluster of the early fall samples of M. spicatum apices, another cluster of the two leaf samples from October and a third cluster, which contains all artificial substrates and each one apex and leaf sample (Figure 3.2C).
Sequencing of single DGGE bands. Bands cut from the gels were re–amplified and
Figure 3.2. Cluster analysis of DGGE banding patterns of the heterotrophic biofilm community on Myriophyllum spicatum, Potamogeton perfoliatus and the artificial substrates in 2006.
A: Apices of both plant species in comparison to the artificial substrates. B: Lower leaves of both plant species in comparison to the artificial substrate. A & B:
Samples from Lake Constance. C: Comparison between mesocosm M. spicatum and mesocosm artificial substrates MS: M. spicatum, PP: P. perfoliatus, Art:
artificial substrate.
with BLAST (Altschul et al. 1990). Most of the retrieved sequences belonged to the betaproteobacteria (50%), gammaproteobacteria (21%) and the rest (29%) could be only assigned to the domain bacteria (Table 3.2). The closest relatives based on BLAST search are from soil or freshwater habitats. We analysed 16 bands of the leaves, of which 4% belonged to the gammaproteobacteria, 6% each to the actinobacteria, betaproteobacteria, cyanobacteria and chloroplasts. The rest (50%) could only be assigned to the domain bacteria (Table 3.3, Figure S3.2B, see supplementary). Here, the sequences were mostly similar to those from other freshwater studies.
Table 3.2. BLAST analysis of 16S rRNA gene sequences from biofilm on the apices of Myriophyllum spicatum, Potamogeton perfoliatus and the artificial substrate.
Substrate Sampling Datea) Most similar to Identity
(%)
Bacterial group Accession no. Sourceb )
M. spicatum 31. July (18) Uncultured bacterium clone 164ds20 100 Bacteria; environmental samples. AY212616 Equine faecal contamination P. perfoliatus 31. July (19) Uncultured Burkholderiales bacterium clone Hv(lab)_2.15 99 Betaproteobacteria EF667915 Basal metazoan Hydra P. perfoliatus 31. July (16) Methylophilus sp. U33 98 Betaproteobacteria EU375653 Organic pollutants degradation
P. perfoliatus 15. August (14) Acinetobacter sp. HTYC28 98 Gammaproteobacteria EU372908 China sea
Artificial 15. August (15) Ralstonia sp. JB1B3 100 Betaproteobacteria EU375662 Organic pollutants degradation M. spicatum 12. September (20) Uncultured Ideonella sp. clone GASP–MA2S1_A04 98 Betaproteobacteria EF662829 Bacterial soil communities in Michigan P. perfoliatus 12. September (11) Uncultured bacterium clone MA34_2003DFa_B05 90 uncultured bacterium EF378328 Agricultural soil community
M. spicatum 22. September (8) Uncultured bacterium clone 164ds20 93 Bacteria; environmental samples AY212616 Equine faecal contamination P. perfoliatus 22. September (10) Clonothrix fusca strain AW–b 93 Gammaproteobacteria DQ984190 Clonothrix fusca Roze 1896 M. spicatum 09. October (4) Uncultured betaproteobacterium clone CH_02 97 Betaproteobacteria EF562573 Complex organic matter degradation M. spicatum 09. October (12) Uncultured Ideonella sp. clone GASP–MA2S1_A04 97 Betaproteobacteria EF662829 Bacterial soil communities in Michigan
M. spicatum 22. September (8) Uncultured bacterium clone 164ds20 93 Bacteria; environmental samples AY212616 Equine faecal contamination P. perfoliatus 22. September (10) Clonothrix fusca strain AW–b 93 Gammaproteobacteria DQ984190 Clonothrix fusca Roze 1896 M. spicatum 09. October (4) Uncultured betaproteobacterium clone CH_02 97 Betaproteobacteria EF562573 Complex organic matter degradation M. spicatum 09. October (12) Uncultured Ideonella sp. clone GASP–MA2S1_A04 97 Betaproteobacteria EF662829 Bacterial soil communities in Michigan