Microbial Ecology of Phototrophic Biofilms
PROEFSCHRIFT
ter verkrijging van de graad van doctor aan de Technische Universiteit Delft,
op gezag van de rector Magnificus prof. dr. ir. J. T. Fokkema, voorzitter van het College voor Promoties,
in het openbaar te verdedigen op dinsdag 11 september 2007 om 15.00 uur.
door
Guus Roeselers
Dit proefschrift is goedgekeurd door de promotor: Prof. dr. ir. M. C. M. van Loosdrecht
Toegevoegd promotor: Dr. G. Muyzer
Samenstelling promotie commissie:
Prof. K. C. A. M. Luyben, Technische Universiteit Delft, voorzitter
Prof. dr. ir. M. C. M. van Loosdrecht, Technische Universiteit Delft, promotor Dr. G. Muyzer, Technische Universiteit Delft, toegevoegd promotor
Prof. dr. J. G. Kuenen, Technische Universiteit Delft Prof. dr. W. Admiraal, Universiteit van Amsterdam Prof. dr. L. J. Stal, Universiteit van Amsterdam Prof. dr. H.J. Laanbroek, Universiteit Utrecht Dr. A. Wilmotte, Université de Liège, België
Het in dit proefschrift beschreven onderzoek is uitgevoerd bij de sectie Milieubiotechnologie, afdeling Biotechnolgie, Technische Universiteit Delft, Julianalaan 67, 2628 BC, Delft.
Dit onderzoek is gefinancierd door de Europese Unie (PHOBIA: QLK3-CT-2002-01938) en de Technische Universiteit Delft.
CONTENTS
Chapter 1 General introduction 9
Chapter 2 Phototrophic biofilms and their potential applications 17
Chapter 3 Diversity of phototrophic bacteria in microbial mats from Arctic hotsprings (Greenland)
33
Chapter 4 Heterotrophic pioneers facilitate phototrophic biofilm development
53
Chapter 5 On the reproducibility of microcosm experiments -different community composition in parallel phototrophic biofilm microcosms
67
Chapter 6 Development of a PCR for the detection and identification of cyanobacterial
nifD
genes83
Chapter 7 Diversity and expression of cyanobacterial
hupS
genes in pure cultures and in a nitrogen limited phototrophic biofilm97
Chapter 8 Concluding remarks
Summary Samenvatting About the author List of publications Acknowledgements
1
Chapter 1
10
INTRODUCTION Biofilms
Biofilms can be defined as layered structures of microbial cells and an extracellular matrix of polymeric substances, associated with surfaces and interfaces. Biofilms trap nutrients for growth of the enclosed microbial community and help prevent detachment of cells from surfaces in flowing systems. Moreover, microorganisms embedded in a biofilm are protected against toxic substances, predation, and for that matter host immune responses (Davey and O'Toole G, 2000). However, life in a biofilm has also some disadvantages compared to the planktonic mode of life. Cells growing in a biofilm often experience mass transfer limitation and competition for available resources may be intense. Therefore, microbial growth rates are often found to be lower in a biofilms than in suspension.
Given their ubiquity and importance in the microbial world, it is hardly surprising that biofilms have received much attention from the scientific community. However, the morphology, chemistry, physiology, and ecology of naturally occurring biofilms are as diverse as their constituent microorganisms. For example, pearl-string-like archaeal biofilms found in sulfidic springs (Henneberger et al., 2006) are profoundly different from the plaque formed on air-exposed surfaces of teeth (Kolenbrander and London, 1993).
Phototrophic biofilms and microbial mats
The structure, growth dynamics and physiology of heterotrophic biofilms have been extensively studied. But until recently phototrophic biofilms have received little attention. Phototrophic biofilms occur on contact surfaces in a range of terrestrial and aquatic environments. Phototrophic biofilms can best be described as surface attached microbial communities mainly driven by light as the energy source with a photosynthesizing component clearly present. Eukaryotic algae and cyanobacteria generate energy and reduce carbon dioxide, providing organic substrates and oxygen. The photosynthetic activity fuels processes and conversions in the total biofilm community, including the heterotrophic fraction. Thick laminated multilayered phototrophic biofilms are usually referred to as microbial mats or phototrophic mats. Phototrophic biofilms and microbial mats have been described in extreme environments like thermal springs, hyper saline ponds (Sorensen et al., 2005), desert soil crusts (Garcia-Pichel and Pringault, 2001), and in lake ice covers in Antarctica (Taton et al., 2003). The 3.4-billion-year fossil record of benthic phototrophic communities, such as microbial mats and stromatolites (Des Marais, 1990; Arp et al., 2001), indicates that these associations represent the Earth's oldest known ecosystems. These early ecosystems played a key role in the formation of the present Earth’s oxygenic atmosphere (Knoll, 1996; Hoehler et al., 2001).
Green algae, diatoms and cyanobacteria
Algae are a large and diverse group of eukaryotic organisms containing chloroplasts capable of performing oxygenic photosynthesis. Although most algae are of microscopic size and hence are clearly microorganisms, some macroscopic species known as kelps are over 70 meters in length.
General introduction
11 Diatoms (Bacillariophyta) are another major group of eukaryotic microalgae. Most diatoms are unicellular, although some form chains or simple colonies. They are characterized by their highly ornamented glasslike cell walls consisting of hydrated silicon dioxide. Diatoms have an extensive fossil record going back to the Cretaceous; some rocks are formed almost entirely of fossil diatoms. Many diatoms show gliding motility, giving them the ability to adhere and glide over a substratum. Benthic diatoms are an important component of the microphytobenthos that inhabit intertidal mudflats. They are also an important component of the biofouling community in marine environments.
Cyanobacteria are Gram-negative, aerobic bacteria containing the photosynthetic pigment chlorophyll a and performing oxygenic photosynthesis, fixing CO2 through the reductive pentose phosphate cycle (Castenholz, 2001). This monophyletic group is morphologically and developmentally very diverse (Figure 1).
Figure 1. Micrograph of an Anabaena variabilis filament with a heterocyst. The nitrogenase enzyme complex involved in the fixation of atmospheric N2 is
extremely sensitive to oxygen. Heterocysts are cells specialized in the fixation of N2 under oxic conditions, which differentiate at regular intervals along the
filaments of some diazotrophic cyanobacteria.
Both unicellular and filamentous forms are known, and considerable variation within these morphological types occurs (Rippka et al., 1979; Castenholz, 2001). They are widely distributed in nature in terrestrial, freshwater, and marine habitats. In general they are more tolerant to extreme environmental conditions than eukaryotic algae. Many species grow attached to the surfaces of rocks and soils. Cyanobacteria can form dense microbial mats, especially in extreme environment with low grazing pressure such as hyper saline ponds (Caumette et al., 1994) and hot springs (Ward et al., 1998).
Cyanobacterial nitrogen fixation
Chapter 1
12
Biological N2 fixation involves the activity of the oxygen sensitive multi-component nitrogenase enzyme (Mancinelli, 1996). The conventional nitrogenase is encoded by the nifHDK genes (Figure 2). The nitrogenase subunit genes are like the 16S rRNA genes highly conserved and can be used as phylogenetic markers (Zehr et al., 1995).
Diazotrophic cyanobacteria (cyanobacteria capable of N2 fixation) can be subdivided into three main groups (Stal, 1995). Group I consists of heterocystous cyanobacteria. Heterocysts are differentiated cells specialized in nitrogen fixation. Their thick cell wall contains special lipopolysacharides and forms a diffusion barrier for gases, limiting the entry of oxygen (Figure 1). Heterocysts have lost the capacity of oxygenic photosynthesis and provide the other cells in the filament with nitrogen for biosynthesis. Group II consist of filamentous and unicellular cyanobacteria that do not form heterocysts and only perform N2 fixation under anoxic conditions. Group III cyanobacteria comprise filamentous and unicellular cyanobacteria that are able to fix N2 under fully aerobic conditions. Their mechanism of protecting the nitrogenase complex from O2 inhibition is still not fully understood.
Uptake hydrogenases
Most nitrogenases are catalysts for hydrogen production as they liberate H2 during the reduction of nitrogen to ammonia (Mancinelli, 1996). In cyanobacteria, a NiFe-uptake hydrogenase recycles the H2 effused by the nitrogenase machinery (Figure 2). The recycling of hydrogen has been suggested to have three physiological functions. I) It generates ATP via the oxyhydrogen (Knallgas) reaction, II) it protects the sensitive nif complex from oxydative inactivation, and III) it supplies reducing equivalents (electrons) for N2 reduction and other cell functions (Bothe et al., 1977). Biohydrogen production could be facilitated by the selection of strains, or the production of mutants, deficient in H2 uptake activity.
General introduction
13 The PHOBIA project
The research described in this thesis was partly performed within the frame of the European Union funded project PHOBIA (Phototrophic biofilms and their potential applications: towards the development of a unifying concept). This 36-month project has been funded within the 5th EU Framework Programme 'Quality of Life', Key action 3 'The Cell Factory'. A consortium with the Centre for Environmental Research, Magdeburg (Germany), Institute of Chemical and Biological Technology, Oeiras (Portugal), University of Copenhagen (Danmark), The Netherlands Institute of Ecology (NIOO-KNAW), University of Rome “Tor Vergata” (Italy), and Delft University of Technology investigated the microenvironment, physiology, and spatial development of a number of aquatic phototrophic biofilms.
The objectives of PHOBIA were (1) to comprehend how aquatic biofilms adhere to submerged surfaces, how the biofilms grow and function; (2) to make a unifying conceptual model for phototrophic biofilms, based on artificial neural networks. Such self-learning models can predict how environmental conditions structure the biofilm (both in spatial terms and in terms of biodiversity/phylogeny), and how the biofilm interacts with the environment and the substrata. The knowledge of these biofilms will provide a basis for applied research in the field of water bioremediation and development of new antifouling agents.
Within the PHOBIA project, various aspects of phototrophic biofilm growth were studied in separate work packages. Additional to biofilm structure and architecture, physical and chemical gradients, surface interactions, and biofilm strength, an analysis of the microbial species in the biofilm communities was made. This included (1) a taxonomic analysis of the major species, using light- and electron microscopy (University of Rome); (2) fingerprints of membrane phospholipid fatty acids (PFLA) that are specific for various groups of microbes (heterotrophic bacteria, cyanobacteria, green algae, diatoms), using GC-MS and fingerprints of group-specific photosynthetic pigments via HPLC-PDA (NIOO-KNAW); (3) phylogenetic analysis of the biofilm species (photo- and heterotrophs) using molecular techniques such as PCR-denaturing gradient gel electrophoresis (DGGE) (Delft University of Technology).
Phototrophic biofilm incubator
In several chapters of this thesis results are presented that were obtained with aquatic phototrophic biofilms cultivated in a temperature controlled flow-lane incubator (Fig. 4) which was designed by collaborating partners in the PHOBIA consortium and constructed by the workshop at the Centre for Environmental Research in Magdeburg.
This flow-lane incubator system contained four separate flow channels (LC1, LC2, LC3 and LC4) through which a volume of 4 L mineral medium circulated over a surface covered with 47 polycarbonate slides (76 x 25 x 1 mm). Polycarbonate slides were used as a substratum for biofilm adhesion. Each light chamber contained an adjustable light source and the circulation speed of the culture medium could be regulated precisely.
Chapter 1
14
Figure 3. Schematic representation of the phototrophic biofilm incubator.
General introduction
15 OUTLINE
Chapter 2 provides a brief introduction in the ecology of phototrophic biofilms and discusses their actual and potential applications in wastewater treatment, bioremediation, fish-feed production, biohydrogen production, and soil improvement and their role in biofouling.
Chapter 3 describes the diversity of phototrophic bacteria in hot spring microbial mats found on the east coast of Greenland.
Chapter 4 focuses on the successional changes in community composition of freshwater phototrophic biofilms growing under different light intensities. The role of heterotrophic bacteria in phototrophic biofilm development is further explored
I n Chapter 5 t h e incubator system is used for an unprecedented microcosm reproducibility experiment.
Chapter 6 demonstrates that nifD gene sequences can be used to detect and identify diazotrophic cyanobacteria in natural communities. PCR products generated using primers homologous to conserved regions in the cyanobacterial nifD genes were subjected to DGGE and clone library analysis in order to determine the genetic diversity of diazotrophic cyanobacteria in environmental samples.
Chapter 7 describes the development of PCR primers targeting conserved regions within the cyanobacterial h u p S gene family. We analyzed hupS diversity and transcription in cultivated phototrophic biofilms by the direct retrieval and analysis of mRNA that was reverse transcribed, amplified with hupS specific primers, and cloned. In Chapter 8 the main findings presented in this thesis are summarized and evaluated.
REFERENCES
Arp, G., Reimer, A., and Reitner, J. (2001) Photosynthesis-induced biofilm calcification and calcium
concentrations in Phanerozoic oceans. Science 292: 1701-1704.
Beijerinck, M.W. (1901) On oligonitrophilous bacteria. In KNAW Proceedings volume 3; 1900-1901. Amsterdam, pp. 586-595.
Bothe, H., Tennigkeit, J., and Eisbrenner, G. (1977) The utilization of molecular hydrogen by the
blue-green alga Anabaena cylindrica. Arch Microbiol 114: 43-49.
Castenholz, R.W. (2001) Phylum BX. Cyanobacteria. Oxygenic photosynthetic bacteria. In Bergey's Manual of Systematic Bacteriology. Boone, D.R., Castenholz, R.W., and Garrity, G.M. (eds). New York, USA: Springer, pp. 474-487.
Caumette, P., Matheron, R., Raymond, N., and Relexans, J.C. (1994) Microbial Mats in the Hypersaline
Ponds of Mediterranean Salterns (Salins-De-Giraud, France). Fems Microbiology Ecology 13: 273-286.
Davey, M.E., and O'Toole G, A. (2000) Microbial biofilms: from ecology to molecular genetics. Microbiol Mol Biol Rev 64: 847-867.
Des Marais, D.J. (1990) Microbial mats and the early evolution of life. Trends Ecol Evol 5: 140-144.
Garcia-Pichel, F., and Pringault, O. (2001) Microbiology. Cyanobacteria track water in desert soils.
Nature 413: 380-381.
Henneberger, R., Moissl, C., Amann, T., Rudolph, C., and Huber, R. (2006) New insights into the
lifestyle of the cold-loving SM1 euryarchaeon: natural growth as a monospecies biofilm in the subsurface. Appl Environ Microbiol 72: 192-199.
Hoehler, T.M., Bebout, B.M., and Des Marais, D.J. (2001) The role of microbial mats in the production
of reduced gases on the early Earth. Nature 412: 324-327.
Knoll, A.H. (1996) Breathing room for early animals. Nature 382: 111-112.
Chapter 1
16
Lewis, L.A., and Mccourt, R.M. (2004) Green algae and the origin of land plants. American Journal of Botany 91: 1535-1556.
Mancinelli, R.L. (1996) The nature of nitrogen: an overview. Life Support Biosph Sci 3: 17-24.
Rippka, R., Deruelles, J., Waterbury, J.B., Herdman, M., and Stanier, R.Y. (1979) Generic
assignments, strain histories and properties of pure cultures of cyanobacteria. J Gen Microbiol 111: 1–61.
Sorensen, K.B., Canfield, D.E., Teske, A.P., and Oren, A. (2005) Community composition of a
hypersaline endoevaporitic microbial mat. Appl Environ Microbiol 71: 7352-7365.
Stal, L.J. (1995) Physiological ecology of cyanobacteria in microbial mats and other communities. New Phytologist 131: 1-32.
Taton, A., Grubisic, S., Brambilla, E., De Wit, R., and Wilmotte, A. (2003) Cyanobacterial diversity in
natural and artificial microbial mats of Lake Fryxell (McMurdo Dry Valleys, Antarctica): a morphological and molecular approach. Appl Environ Microbiol 69: 5157-5169.
Ward, D.M., Ferris, M.J., Nold, S.C., and Bateson, M.M. (1998) A natural view of microbial biodiversity
within hot spring cyanobacterial mat communities. Microbiol Mol Biol Rev 62: 1353-1370.
Zehr, J.P., Mellon, M., Braun, S., Litaker, W., Steppe, T., and Paerl, H.W. (1995) Diversity of
2
Phototrophic biofilms and their potential applications
G. Roeselers, M.C.M. van Loosdrecht, and G. Muyzer
Chapter 2
18
ABSTRACT
Phototrophic biofilms occur on contact surfaces exposed to light in a range of terrestrial and aquatic environments. Oxygenic phototrophs like diatoms, green algae and cyanobacteria are the major primary producers that generate energy and reduce carbon dioxide, providing the system with organic substrates and oxygen. Photosynthesis fuels processes and conversions in the total biofilm community, including the metabolism of heterotrophic organisms. A matrix of polymeric substances secreted by phototrophs and heterotrophs enhances the attachment of the biofilm community.
This review discusses the actual and potential applications of phototrophic biofilms in wastewater treatment, bioremediation, fish-feed production, biohydrogen production, and soil improvement and their role in biofouling.
INTRODUCTION
Phototrophic biofilms can best be described as surface attached microbial communities driven by light energy with a photosynthesizing component clearly present. Oxygenic phototrophic microorganisms such as benthic diatoms (centric, pennate, unicellular and filamentous), unicellular and filamentous cyanobacteria, and benthic green algae generate energy and reduce carbon dioxide, providing organic substrates and oxygen.
The photosynthetic activity fuels processes and conversions in the total biofilm community. For example, heterotrophs derive their organic C and N requirements from excreted photosynthates and cell lysates, while nutrient regeneration is enhanced by heterotrophs (Bateson and Ward, 1988).
The microorganisms produce extracellular polymeric substances (EPS) that hold the biofilm together (Flemming, 1993; Wimpenny et al., 2000). Thick laminated multilayered phototrophic biofilms are usually referred to as microbial mats or phototrophic mats (Guerrero et al., 2002; Roeselers et al., 2007; Stal et al., 1985; Ward et al., 1998). The top layer of microbial mats is typically dominated by oxygenic phototrophs, such as cyanobacteria (Castenholz, 2001a) and diatoms, with underlying or intermixed layers of anoxygenic phototrophs, i.e. green and purple sulfur bacteria (GSB and PSB) (Martinez-Alonso et al., 2005) and chloroflexi like bacteria (Castenholz, 2001b; Ruffroberts et al., 1994).
Steep vertical redox and chemical gradients (~ microns to millimeters) that establish in phototrophic biofilms and mats enforce these stratifications in the microbial community (Figure 1). Light intensity decreases with depth, restricting phototrophic activity to the upper layer of the mat. Oxygenic photosynthesis results in a steep oxygen gradient that restricts most anoxygenic phototrophs and anaerobic chemotrophs to the lower parts of the mat. However, recent studies have also shown examples of anaerobes thriving in the oxic zone of microbial mats (Cypionka, 2000; Schaub and Van Gemerden, 1994). The utilization of CO2 during photosynthesis results in a pH gradient (Revsbech et al., 1983).
Phototrophic biofilms and mats are formed on surfaces in a range of terrestrial and aquatic environments (Chan et al., 2003; Ferris et al., 1997; Ortega-Morales et al., 2000). The oldest fossilized phototrophic biofilm or mat like structures date back approximately 3.5 billion years (Des Marais, 1990).
Phototrophic biofilms and their potential applications
19 biohydrogen production (Prince and Kheshgi, 2005; Tsygankov et al., 1999), and in the development of antifouling agents (Bhadury and Wright, 2004; Callow and Callow, 2002). The present review will give a brief introduction in various actual and potential biotechnological applications of phototrophic biofilms.
Figure 1. This cross section reveals the stratified structure of a thick cultivated freshwater phototrophic biofilm. The dark top layer consists predominantly of Oscillatoria like cyanobacteria. Scale bar indicates 1 mm.
Wastewater treatment
The application of oxygenic phototrophs in the treatment of waste streams that are relatively rich in nutrients and low in organic carbon has many advantages. In a heterotrophic biofilm O2 transfer by diffusion is limited to approximately 20 nmol cm
-2 min-1. However, the areal net oxygen production in an active phototrophic biofilm at a light intensity of 1000 mol photons m-2 s-1 is approximately a factor two higher (Epping and Kuhl, 2000). Hence, the oxygen that is produced by phototrophs can cover a great part of the oxygen demand of bacterial nitrification and the heterotrophic consumption of organic carbon. In adition, oxygenic phototrophs assimilate nutrients for building biomass with carbon dioxide as carbon source. In contrast to wastewater treatment by bacterial nitrification and denitrification, where a large part of the nitrogen escapes as N2 to the atmosphere, the nitrogenous compounds are in this case retained in algal biomass.
Chapter 2
20
Table 1. Nitrogen and phosphorus removal rates obtained with different phototrophic biofilm based wastewater treatment systems.
System N-removal
ratea
P- removal
rateb Reference
Algal turf scrubber (ATS). 1110 730 Craggs et al, 1996 Periphyton-fish system mesocosm. 108 27 Rectenwald and Drenner,
2000
Secondary effluent clarifier 1900 160 Davis et al., 1990 ATS fed with 1% dairy manure. 720 330 Pizarro et al., 2002 Phototrophic biofilms in natural streams 648 117 Davis and Minshall, 1999
a
Average phosphorus removal rates (mg P m-2
day-1
), b
Average nitrogen removal rates (mg N m-2
day-1
)
The capability of nutrient removal in the absence of organic carbon has often been used for wastewater treatment in algal pond systems (Davis et al., 1990; Garcia et al., 2000). A major disadvantage using suspended algae is the high secondary organic pollution caused by algae biomass in the effluent of the ponds (Racault, 1993). Biomass can be removed by filtration, sedimentation with centrifugation or with decantation but most of these methods are costly. By using immobilized phototrophic biofilms the problem of separation of suspended algal biomass and water can be avoided and the nitrogenous compounds retained in algal biomass can be harvested and used as a fertilizers in agriculture (Schumacher and Sekoulov, 2002).
An interesting feature of some cyanobacteria is that they can accumulate inorganic phosphorus and store them internally as polyphosphates (Kromkamp, 1987). However, this aspect has hardly been explored in the context of wastewater treatment.
The photosynthetic activity in phototrophic biofilms results in an increasing pH due to the change of the carbon dioxide equilibrium in water. This increase in pH causes precipitation of dissolved phosphates, in addition to phosphorus removal by assimilation. This photosynthesis induced pH increase has also shown potential for the reduction of faecal coliform bacteria in wastewater streams (Schumacher et al., 2003).
European Union regulations have led to more stringent effluent standards for sewage treatment facilities located in ecologically sensitive areas. Phototrophic biofilms can be applied for the additional nutrient removal from secondary effluents of wastewater treatment plants. Nutrient removal in so-called constructed wetlands, which show potential for small scale wastewater treatment in developing countries, depends also to a large extend on the activity of epiphytic phototrophic biofilms growing on reed stems (Larsen and Greenway, 2004; Ragusa et al., 2004). Table 1. shows the nitrogen and phosphorus removal rates obtained with several phototrophic biofilm based wastewater polishing systems.
For process control and system optimization, it is important to define the right operational conditions. Craggs et al. (1996) described several parameters that determine the efficiency of nutrient removal in an experimental phototrophic biofilm system. An important parameter is the applied flow velocity. At higher flow velocities there is a trade-off between reduced colonization and shear stress versus increased metabolism in the established biofilms by reduced boundary layers and increased water mixing.
Phototrophic biofilms and their potential applications
21 water with a limited flow-velocity, the reduced debit can lead to conditions were the biofilm becomes nutrient limited instead of light limited. Hence, optimal water depths will also depend on nutrient loads and local or seasonal light conditions.
Figure 2. Discharge of the municipal wastewater treatment facility (WWTF) of Sint Maartensdijk (the Netherlands) is polished in a constructed wetland system (I= inlet, O = outlet). Nitrate is primarily assimilated by epiphytic phototrophic biofilms growing on reed stems.
Removal of heavy metals
Biosorption consists of several mechanisms, mainly ion exchange, chelation, adsorption, and diffusion through cell walls. These “passive” mechanisms can take place at the cellular level and at the microbial community level. The active mode of metal uptake and concentration is called bioaccumulation. This process is dependent on the cellular metabolism.
Chapter 2
22
Bhaskar and Bhosle, 2006; Liu et al., 2001; Wang et al., 1998). Parker et al. (2000) showed that mucilage sheaths isolated from the cyanobacteria Microcystis aeruginosa and Aphanothece halophytica exhibit strong affinity for heavy metal ions such as copper, lead and zinc. In addition to biosorption and bioaccumulation, the elevated pH inside photosyntetically active biofilms may favor removal of metals by precipitation (Liehr et al., 1994).
Major advantages of metal removal by biosorption include, low cost, and high efficiency of heavy metal removal from diluted solutions. However, in order not to simply displace heavy metal pollution, methods will have to be developed to extract heavy metals easily from biomass (Kratochvil and Volesky, 1998).
Water hardness is a crucial factor that influences metal uptake efficiency because cations such as Ca2+ and Mg2+ compete with trace metals for binding sites on cell membranes and extracellular polysaccharides (Fortin et al., 2007). Meylan et al., (2003) showed that different concentrations of dissolved manganese affected the intracellular accumulation of zinc and copper by phototrophic biofilms. In addition to cation concentrations, metal uptake is affected by light intensity, pH, biofilm density, the presence of metal binding humic substances, and the tolerance of individual algal species to specific heavy metals. (Fortin et al., 2007; Vymazal, 1984)
Oil degradation
The volumes of petroleum-based products transported across the world are enormous and the risk of oil spillage is significant. The volume of spills usually exceeds the inherent remediation capacity for any given environment, resulting in a significant ecological impact (Cohen, 2002). It has been suggested that microbial mats can play a role in the biodegradation of oil. In the years after the massive oil spills during the first Gulf war of 1991, it was observed that dense mats of cyanobacteria formed on contaminated beaches (Sorkhoh et al., 1992). It has been shown that in particular Oscillatoria spp. are able to cope with heavy oil pollution (Abed et al., 2006; Cohen, 2002; van Bleijswijk and Muyzer, 2004).
Although there is no direct evidence that cyanobacteria are directly involved in the degradation of petroleum products, they probably facilitate degradation by sulfate-reducing bacteria (Edwards et al., 1992) and aerobic heterotrophs (Benthien et al., 2004; Cohen, 2002; Sorkhoh et al., 1995). Previous studies have shown that the addition of nitrogen supplements enhances microbial assimilation of carbon from oil (Coffin et al., 1997). Cyanobacterial N2 fixation could provide sufficient nitrogen compounds for heterotrophic oil degradation. Free radicals formed during oxygenic photosynthesis could indirectly enhance photochemical oil degradation (Nicodem et al., 1997).
Microcosm studies examining the initial response of phototrophic biofilms exposed to petrochemical compounds revealed signs of acute toxicity (Nayar et al. 2004). Phototrophic biofilms are ubiquitous and dominant primary producers forming the base of aquatic food webs. Therefore, it has been suggested that phototrophic biofilms are applicable as sensitive bioindicators of petrochemical pollution and for ecotoxicology tests (Nayar et al. 2004).
Agriculture
Phototrophic biofilms and their potential applications
23 Cyanobacteria can also be applied for in situ soil fertilization via N2 fixation. Much work has been done on the fertilization of rice paddy fields with nitrogen fixing cyanobacteria (Ariosa et al., 2004; Habte and Alexander, 1980; Lem and Glick, 1985).
In addition, EPS produced by algae and cyanobacteria can improve the soil water-holding capacity and prevent erosion (Barclay and Lewin, 1985; Rao and Burns, 1990). Mazor et al. (1996) showed that addition of 0.5 mg of Microcoleus sp. EPS per g sand retained approximately 30% of the water-holding capacity of the sand after 24 h of desiccation at 55°C while sand samples without EPS dried out completely.
Elevated soil salinity, which is increasing world wide, has a major impact on soil quality and agricultural production. In many coastal areas, salinity is an inherent situation, but inefficient water management, i.e. excess recharging of groundwater and accumulation through concentration often leads to secondary salinization of farmlands. Early 1950 a biological approach to the problem of saline soils using cyanobacteria was proposed (Singh, 1950). It has been shown that inoculation of soil surfaces with a suspension of halotolerant cyanobacteria leads to a salinity reduction (Apte and Thomas, 1997; Kaushik and Venkataraman, 1982). This amelioration of soil salinity is probably caused by a temporal entrapment of Na+ ions in cyanobacterial EPS sheaths, resulting in a restricted Na+ influx in the plant roots (Ashraf et al., 2006). Permanent removal of Na+ from the soil may not be possible, since Na+ is released back into the soil subsequent to the death and decay of the cyanobacteria.
Aquaculture
Effluent discharges of intensive fish production systems may cause significant nutrient pollution. Fish farmers have a stake in regulating nutrient pollution, because poor water quality can reduce aquaculture productivity. On a small scale phototrophic biofilm based systems can be used to reduce ammonia and nitrate concentrations in aquaculture effluents (Bender and Phillips, 2004).
In fish aquacultures, commercial feeds, consisting mostly of fishmeal and oil, may account for more than 50% of the total production costs (Elsayed and Teshima, 1991). Only about 15-30% of the nutrient input in feed-driven pond systems is converted into harvestable products (Gross et al. 2000). Therefore, there is a growing interest for substitution of commercial feeds with alternative protein sources. The cost savings and the reduction of ecological impact by using phototrophic biofilms for fish feed production and the possible simultaneous effluent treatment may be significant (Elsayed and Teshima, 1991; Naylor et al., 2000; Phillips et al., 1994).
Tilapia (Oreochromis niloticus) can consume microalgae as a major, even exclusive source of its feed requirements. Due to their omnivorous diet and rapid growth, species of tilapia are highly suitable for aquaculturing in fertized pond systems. In fertilized ponds, organic and inorganic fertilizers are used to increase productivity. Nutrients are incorporated into algal biomass and, through a complex food web ultimately incorporated into fish biomass.
Azim et al. (2003) showed that by adding substrate for biofilm adherence to fertilized aquaculture ponds; the conversion of nutrients into harvestable products could be optimized. Tilapia growth was significantly higher and nitrogen retention doubled in substrate ponds compared with control ponds. The potential of fish production based on phototrophic biofilms was reviewed in detail by van Dam et al. (2002).
Chapter 2
24
consumption (Magalhaes et al., 2001; Wiegand and Pflugmacher, 2005). The occurrence of toxins has often been related to rapid planctonic cyanobacterial biomass development during algal blooms but this link is much less common in benthic assemblages (Blaha et al., 2004).
Biohydrogen
Hydrogen is a clean alternative to fossil fuels as its combustion only generates water as a byproduct. Biological production of hydrogen could provide a renewable source of energy. Cyanobacteria are highly promising microorganisms for biological photohydrogen production. Two cyanobacterial enzymes are capable of hydrogen production. The bidirectional hydrogenase complex can either produce or oxidize H2 in the presence of suitable electron donor or acceptor. The physiological role of bidirectional hydrogenases is still unclear and the enzyme is absent in a significant number of strains.
Hydrogen evolution is also catalyzed by nitrogenases (Mancinelli, 1996; Tamagnini et al., 2002; Zehr and Turner, 2001). The nitrogenase machinery releases at least one mol H2 per mol N2 reduced to ammonia, which represents a significant loss of energy. However, most diazotrophic cyanobacteria posses an enzyme called uptake hydrogenase that serves to recycle some of the electrons lost in the form of H2. Several studies with bioreactors have demonstrated the feasibility of cyanobacterial biohydrogen production (Lindberg et al., 2004; Schutz et al., 2004; Tsygankov et al., 1999). As mentioned, N2 reduction has a high ATP requirement and this reduces the potential conversion of solar energy considerably. An advantageous aspect of this is that ATP hydrolysis provides a relatively strong thermodynamic driving force pushing hydrogen evolution, which is not true for bidirectional hydrogenases (Prince and Kheshgi, 2005). This allows the generation of a higher partial H2 pressure in potential bioreactors.
Ideally, hydrogen producing cyanobacteria should invest a minimal amount of ATP in growth, have a high metabolism and should be restricted in place. Cyanobacterial biofilms or mats are of great interest because they meet up to these requirements. The efficiency of hydrogen production could be increased by constructing defined biofilm assemblages containing a range of desired cyanobacterial species or genetically modified strains with a reduced uptake hydrogenase activity (Lindberg et al., 2004; Tsygankov et al., 1999). In order to select an “optimal genetic background” for the construction of genetically engineered cyanobacteria, future studies should focus on the natural molecular variation of strains that have the potential to produce hydrogen. Currently, this technology is in its infancy and as yet not ready for commercial adaptation and exploitation.
Anoxygenic phototrophs and sulfide removal
Sulfide-containing waste streams are usually treated in reactors by chemotrophic sulfide-oxidizing microorganisms using either oxygen or nitrate as the ultimate electron acceptor. In many bioreactors, sulfide is transformed into sulfate by aerobic sulfur-oxidizing bacteria, such as different species of the genus Thiobacillus.
A disadvantage of using aerobic sulfur-oxidizing bacteria is that sulfide removal cannot be combined with sewage treatment by anaerobic digestion. Sulfide oxidation has to take place in a separate reactor in order to avoid exposure of strictly anaerobic methanogens to inhibitory levels of oxygen. Hence, anaerobic oxidation by phototrophic sulfur bacteria (Chloroflexi, GSB and PSB) has been proposed as an alternative method for sulfide removal (Kim et al., 1990).
Phototrophic biofilms and their potential applications
25 of anoxygenic phototrophic bacteria showed promising results (Ferrera et al., 2004; Kobayashi et al., 1983; Syed and Henshaw, 2003).
Only very few processes based on substratum-irradiated biofilms have been employed for large scale treatment of sulfide-containing waste streams (Hurse and Keller, 2004; Jensen and Webb, 1995).
Phototrophic biofilm nuisances
From a human point of view, phototrophic biofilms can develop in wrong places and at the wrong times (Flemming, 2002). Biofouling is known to cause widespread problems in industrial fluid processing applications. Since phototrophic biofilms require light they can cause problems at surfaces exposed to daylight. Phototrophic biofouling causes technical failure or damage to power plant cooling systems, aquaculture systems, fishing nets, ship hulls, marine infrastructures and historical buildings (Ortega-Morales et al., 2000). It constitutes a serious problem for seagoing ships due to increased resistance and fuel costs that can rise by as much as 40 percent (Townsin, 2003). In order to prevent or control phototrophic biofouling, efforts have been made to develop anti-fouling agents.
The most common compounds in anti-fouling agents are tri-butyl-tin (TBT). TBT is highly effective but it is also toxic to non-target organisms and it is not biodegradable. Since, the use of the TBT based coatings will be completely banned by January 2008, there is an urgent need for sustainable alternatives.
Surprisingly, some marine algae could serve in the prevention of biofouling. Certain cyanobacteria and eukaryotic algae produce biogenic compounds such as lipopeptides, amides, alkaloids, terpenoids, lactones, pyrroles and steroids with antibacterial, anti-algal, and antifungal properties, which could be applied in the development of environmentally friendly anti-fouling agents (Bhadury and Wright, 2004; Xu et al., 2005).
Successional community changes during the colonization of new habitats or after environmental disturbances have been widely studied in the macro-ecology, but also for communities of planktonic and benthic microalgae (Helbling et al., 2005; Schäfer et al., 2001), and more recently in bacterial communities (Ferris et al., 1997; Martiny et al., 2003; Roeselers et al., 2004). Previous studies have shown that heterotrophic bacteria play an important role as early colonizers during phototrophic biofilm development (Chan et al., 2003; Roeselers et al., 2007). A valuable strategy for the development of anti-fouling agents could lie in the identification and characterization of pioneer microorganisms responsible for the initial surface colonization that leads to biofilm formation (Zhang et al., 2006).
A critical remark at this point could be that the despite of the promising results presented by scientists, yet there has hardly been any development of commercially applicable alternatives for TBT.
Conclusions and future perspectives
The field of microbiology has come to accept the universality of biofilms. Researchers in the fields of clinical, food and water, and environmental microbiology have begun to investigate microbiologic processes from a biofilm perspective. There are multiple examples of genotypic and physiological differences between microorganisms growing planktonic or in biofilms. Until recently, applied phycological studies have focused mainly on the planktonic mode of life.
Chapter 2
26
In the context of potential applications, the most important features of these biofilm systems are versatility and adaptability, i.e. they have a broad spectrum of capabilities. This makes it possible to link different end uses within the same process; for example nitrate and phosphate removal combined with production of fish feeds (Bender and Phillips, 2004).
The complexity in terms of species richness is an important aspect determining the metabolic biodiversity, and adaptability of phototrophic biofilms (Boles et al. 2004; Girvan et al. 2005). In addition, applications of pure culture or defined community biofilms seem less attractive due to the high costs associated with control of the culture performance and equipment sterilization and isolation to prevent contaminations. Hence, the focus of application development should be on the use of open mixed culture systems. Therefore, a clear understanding of the ecology of phototropic biofilm communities is essential in order to optimize their cultivation for specific biotechnological applications.
Depending on the application it can be essential to select for phototropic biofilms containing specific species and strains, e.g. strains with a high polyunsaturated fatty acid content, strains that do or do not secrete harmful secondary metabolites, strains producing EPS with high metal sorption capacities, or strains with a high tolerance for petroleum-based compounds.
The efficiency and reliability of phototrophic biofilm applications depend to a great extent on the possibility to select and maintain desired community compositions. Future studies should focus on successional changes during biofilm development (Chan et al., 2003; Roeselers et al., 2007), susceptibility to viral and grazing pressure (Simek and Chrzanowski, 1992; Thingstad, 2000), and the mechanisms that determine the structural and functional responses to abrupt perpetuations, and seasonal fluctuations on community composition and productivity (Kaufman, 1982).
Molecular ecological techniques that allow detailed "in situ" characterization of community compositions and activities provide an important tool for future research. Non-culture based molecular methods such as DGGE, clone library analysis, quantitative PCR and stable isotope probing can be used to obtain the phylogeny, relative abundance and genetic activity of individual members of a biofilm community (Omoregie et al., 2004; Roeselers et al., 2006; Steunou et al., 2006). In particular functional genomics approaches will offer important clues about phototrophic biofilm biology.
An important consideration for applications based on phototrophic activity in general is that they require much surface. These systems are primarily fueled by light, which differs from all other resources because it cannot be mixed. The unidirectional nature of photons requires that biofilms are cultivated on surfaces exposed to direct solar radiation. Therefore, high land prices could be a major hurdle for applications in for instance the treatment of municipal wastewater in densely populated areas. Combining different end uses within processes could compensate for the cost of these relatively large footprints. Phototrophic biofilms would also be suitable for the development of inexpensive treatment methods for developing countries, where land values are relatively low and where the bulk of domestic and industrial wastewater is still discharged without any treatment.
ACKNOWLEDGMENTS
Phototrophic biofilms and their potential applications
27 REFERENCES
Abed, R. M., Al-Thukair, A. and de Beer, D. (2006). Bacterial diversity of a cyanobacterial mat
degrading petroleum compounds at elevated salinities and temperatures. FEMS Microbiol Ecol 57: 290-301.
Apte, S. K. and Thomas, J. (1997). Possible amelioration of coastal soil salinity using halotolerant
nitrogen-fixing cyanobacteria. Plant and Soil 189: 205-211.
Ariosa, Y., Quesada, A., Aburto, J., Carrasco, D., Carreres, R., Leganes, F. and Fernandez Valiente, E.
(2004). Epiphytic cyanobacteria on Chara vulgaris are the main contributors to N(2) fixation in rice fields. Appl Environ Microbiol 70: 5391-5397.
Ashraf, M., Hasnain, S. and Berge, O. (2006). Effect of exo-polysaccharides producing bacterial
inoculation on growth of roots of wheat (Triticum aestivum) plants grown in a salt-affected soil. IJEST 3: 43-51.
Azim, M.E., Verdegem, M.C.J., Singh, M., van Dam, A.A., and Beveridge, M.C.M. (2003) The effects of
periphyton substrate and fish density on water quality, phytoplankton, periphyton and fish growth. Aquac Res 34:685-695.
Barclay, W. R. and Lewin, R. A. (1985). Microalgal polysaccharide production for the conditioning of
agricultural soils. Plant and Soil 88: 159-169.
Bateson, M. M. and Ward, D. M. (1988). Photoexcretion and fate of glycolate in a hot spring
cyanobacterial mat. Appl Environ Microbiol 54: 1738-1743.
Bender, J., Rodriguez-Eaton, S., Ekanemesang, U. M. and Phillips, P. (1994). Characterization of
metal-binding bioflocculants produced by the cyanobacterial component of mixed microbial mats. Appl Environ Microbiol 60: 2311-2315.
Bender, J. and Phillips, P. (2004). Microbial mats for multiple applications in aquaculture and
bioremediation. Bioresour Technol 94: 229-238.
Benthien, M., Wieland, A., García de Oteyza, T., Grimalt, J. O. and Kühl, M. (2004). Oil-contamination
effects on a hypersaline microbial mat community (Camargue, France) as studied with microsensors and geochemical analysis. OPHELIA 58: 135-150.
Bhadury, P. and Wright, P. C. (2004). Exploitation of marine algae: biogenic compounds for potential
antifouling applications. Planta 219: 561-578.
Bhaskar, P. V. and Bhosle, N. B. (2006). Bacterial extracellular polymeric substance (EPS): a carrier of
heavy metals in the marine food-chain. Environ Int 32: 191-198.
Blaha, L., Sabater, S., Babica, P., Vilalta, E. and Marsalek, B. (2004). Geosmin occurrence in riverine
cyanobacterial mats: is it causing a significant health hazard? Water Sci Technol 49: 307-312.
Blanco, A., Sanz, B., Llama, M. J. and Serra, J. L. (1999). Biosorption of heavy metals to immobilised
Phormidium laminosum biomass. J Biotechnol 69: 227–240.
Boles, B.R., Thoendel, M., and Singh, P.K. (2004) Self-generated diversity produces "insurance
effects" in biofilm communities. Proc Natl Acad Sci U S A 101: 16630-16635
Callow, M. E. and Callow, J. E. (2002). Marine biofouling: a sticky problem. Biologist (London) 49: 10-14.
Castenholz, R. W. (2001a). Phylum BX. Cyanobacteria. Oxygenic photosynthetic bacteria. In Bergey's Manual of Systematic Bacteriology, pp. 474-487. Edited by D. R. Boone, R. W. Castenholz and G. M. Garrity. New York, USA: Springer.
Castenholz, R. W. (2001b). Class I: Chloroflexi. In Bergey's Manual of Systematic Bacteriology, pp. 427. Edited by D. R. Boone, R. W. Castenholz and G. M. Garrity. New York: Springer-Verlag.
Chaillan, F., Gugger, M., Saliot, A., Coute, A. and Oudot, J. (2006). Role of cyanobacteria in the
biodegradation of crude oil by a tropical cyanobacterial mat. Chemosphere 62: 1574-1582.
Chan, B. K., Chan, W. K. and Walker, G. (2003). Patterns of biofilm succession on a sheltered rocky
shore in Hong Kong. Biofouling 19: 371-380.
Coffin, R. B., Cifuentes, L. A. and Pritchard, P. H. (1997). Assimilation of oil-derived carbon and
remedial nitrogen applications by intertidal food chains on a contaminated beach in Prince William Sound, Alaska. Mar Environ Res 44: 27-39.
Cohen, Y. (2002). Bioremediation of oil by marine microbial mats. Int Microbiol 5: 189-193.
Craggs, R. J., Adey, W. H., Jessup, B. K. and Oswald, W. J. (1996). A controlled stream mesocosm for
Chapter 2
28
Craig, R. J., Adey, W. H., Jenson, K. R., St. John, M. S., Green, F. B. and Oswald, J. (1996).
Phosphorus removal from wastewater using an algal turf scrubber. Water Sci Technol 33: 191–198.
Cypionka, H. (2000). Oxygen respiration by desulfovibrio species. Annu Rev Microbiol 54: 827-848.
Davis, L. S., Hoffmann, J. P. and Cook, P. W. (1990). Production and Nutrient Accumulation by
Periphyton in a Waste-Water Treatment Facility. Journal of Phycology 26: 617-623.
Davis, J.C., and Minshall, G.W. (1999) Nitrogen and phosphorus uptake in two Idaho (USA) headwater
wilderness streams. Oecologia 119:247-255.
Des Marais, D. J. (1990). Microbial mats and the early evolution of life. Trends Ecol Evol 5: 140-144.
Edwards, E. A., Wills, L. E., Reinhard, M. and Grbic-Galic, D. (1992). Anaerobic degradation of toluene
and xylene by aquifer microorganisms under sulfate-reducing conditions. Appl Environ Microbiol 58: 794-800.
Elsayed, A. F. M. and Teshima, S. I. (1991). Tilapia nutrition in aquaculture. Rev Aquat Sci 5: 247-265.
Epping, E. and Kuhl, M. (2000). The responses of photosynthesis and oxygen consumption to
short-term changes in temperature and irradiance in a cyanobacterial mat (Ebro Delta, Spain). Environ Microbiol 2: 465-474.
Ferrera, I., Sanchez, O. and Mas, J. (2004). A new non-aerated illuminated packed-column reactor for
the development of sulfide-oxidizing biofilms. Appl Microbiol Biotechnol 64: 659-664.
Ferris, M. J., Nold, S. C., Revsbech, N. P. and Ward, D. M. (1997). Population structure and
physiological changes within a hot spring microbial mat community following disturbance. Appl Environ Microbiol 63: 1367-1374.
Flemming, H. C. (1993). Biofilms and Environmental-Protection. Water Sci Technol 27: 1-10.
Flemming, H. C. (2002). Biofouling in water systems--cases, causes and countermeasures. Appl Microbiol Biotechnol 59: 629-640.
Flores, E. and Herrero, A. (2005). Nitrogen assimilation and nitrogen control in cyanobacteria. Biochem Soc Trans 33: 164-167.
Fortin, C., Denison, F.H. and Garnier-Laplace, J. (2007) Metal-phytoplankton interactions: Modeling the
effect of competing ions (H+, Ca2+, and Mg2+) on uranium uptake. Environmental Toxicology and Chemistry 26: 242-248.
Garcia, J., Mujeriego, R. and Hernandez-Marine, M. (2000). High rate algal pond operating strategies
for urban wastewater nitrogen removal. J of Appl Phycol 12: 331-339.
Girvan, M.S., Campbell, C.D., Killham, K., Prosser, J.I., and Glover, L.A. (2005) Bacterial diversity
promotes community stability and functional resilience after perturbation. Environ Microbiol 7: 301-313
Gross, A., Boyd, C.E., and Wood, C.W. (2000) Nitrogen transformations and balance in channel catfish
ponds. Aquacultural Engineering 24: 1-14
Guerrero, R., Piqueras, M. and Berlanga, M. (2002). Microbial mats and the search for minimal
ecosystems. Int Microbiol 5: 177-188.
Habte, M. and Alexander, M. (1980). Nitrogen fixation by photosynthetic bacteria in lowland rice culture.
Appl Environ Microbiol 39: 342-347.
Havens, K. E., East, T. L., Meeker, R. H., Davis, W. P. and Steinman, A. D. (1996). Phytoplankton and
periphyton responses to in situ experimental nutrient enrichment in a shallow subtropical lake. J Plankton Res 18: 551-566.
Helbling, E. W., Barbieri, E. S., Marcoval, M. A., Goncalves, R. J. and Villafane, V. E. (2005). Impact of
solar ultraviolet radiation on marine phytoplankton of Patagonia, Argentina. Photochem Photobiol 81: 807-818.
Hurse, T. J. and Keller, J. (2004). Reconsidering the use of photosynthetic bacteria for removal of
sulfide from wastewater. Biotechnol Bioeng 85: 47-55.
Jensen, A. B. and Webb, C. (1995). Treatment of H2s-Containing Gases - a Review of Microbiological
Alternatives. Enzyme Microb Technol 17: 2-10.
Kaushik, B. D. and Venkataraman, G. S. (1982). Reclamative capacity of blue-green algae in saline
and sodic soils. In Proceedings of the National Symposium on Biological Nitrogen Fixation, pp. 378–389. Bombay: Department of Atomic Energy.
Kaufman, L.H. (1982) Stream aufwuchs accumulation - disturbance frequency and stress resistance
and resilience. Oecologia 52: 57-63
Kim, B. W., Kim, I. K. and Chang, H. N. (1990). Bioconversion of hydrogen sulfide by free and
Phototrophic biofilms and their potential applications
29 Kobayashi, H. A., Stenstrom, M. and Mah, R. A. (1983). Use of Photosynthetic Bacteria for
Hydrogen-Sulfide Removal from Anaerobic Waste Treatment Effluent. Water Research 17: 579-587.
Kratochvil, D. and Volesky, B. (1998). Biosorption of Cu from ferruginous wastewater by algal biomass.
Water Research 32: 2760-2768.
Kromkamp, J. (1987). Formation and Functional-Significance of Storage Products in Cyanobacteria. N Z J Mar Freshwater Res 21: 457-465.
Larsen, E. and Greenway, M. (2004). Quantification of biofilms in a sub-surface flow wetland and their
role in nutrient removal. Water Sci Technol 49: 115-122.
Lem, N. W. and Glick, B. R. (1985). Biotechnological uses of cyanobacteria. Biotechnol Adv 3: 195-208.
Liehr, S. K., Chen, H. J. and Lin, S. H. (1994). Metals removal by algal biofilms. Water Sci Technol 30: 59-68.
Lindberg, P., Lindblad, P. and Cournac, L. (2004). Gas exchange in the filamentous cyanobacterium
Nostoc punctiforme strain ATCC 29133 and Its hydrogenase-deficient mutant strain NHM5. Appl Environ Microbiol 70: 2137-2145.
Liu, Y., Lam, M. C. and Fang, H. H. (2001). Adsorption of heavy metals by EPS of activated sludge.
Water Sci Technol 43: 59-66.
Magalhaes, V. F., Soares, R. M. and Azevedo, S. M. (2001). Microcystin contamination in fish from the
Jacarepagua Lagoon (Rio de Janeiro, Brazil): ecological implication and human health risk. Toxicon 39: 1077-1085.
Mancinelli, R. L. (1996). The nature of nitrogen: an overview. Life Support Biosph Sci 3: 17-24.
Martinez-Alonso, M., Van Bleijswijk, J., Gaju, N. and Muyzer, G. (2005). Diversity of anoxygenic
phototrophic sulfur bacteria in the microbial mats of the Ebro Delta: a combined morphological and molecular approach. FEMS Microbiol Ecol 52: 339-350.
Martiny, A. C., Jorgensen, T. M., Albrechtsen, H. J., Arvin, E. and Molin, S. (2003). Long-term
succession of structure and diversity of a biofilm formed in a model drinking water distribution system.
Appl Environ Microbiol 69: 6899-6907.
Mazor, G., Kidron, G. J., Vonshak, A. and Abeliovich, A. (1996). The role of cyanobacterial
exopolysaccharides in structuring desert microbial crusts. FEMS Microb Ecol 21: 121-130.
Mehta, S. K. and Gaur, J. P. (2005). Use of algae for removing heavy metal ions from wastewater:
progress and prospects. Crit Rev Biotechnol 25: 113-152.
Meylan, S., Behra, R., and Sigg, L. (2003) Accumulation of copper and zinc in periphyton in response to
dynamic variations of metal speciation in freshwater. Environ Sci Technol 37:5204-5212.
Naylor, R. L., Goldburg, R. J., Primavera, J. H. and other authors (2000). Effect of aquaculture on world
fish supplies. Nature 405: 1017-1024.
Nayar, S., Goh, B.P.L., and Chou, L.M. (2004) The impact of petroleum hydrocarbons (diesel) on
periphyton in an impacted tropical estuary based on in situ microcosms. J Exp Mar Bio Ecol 302:213-232
Nicodem, D. E., Fernandes, M. C. Z., Guedes, C. L. B. and Correa, R. J. (1997). Photochemical
processes and the environmental impact of petroleum spills. Biogeochemistry 39: 121-138.
Omoregie, E. O., Crumbliss, L. L., Bebout, B. M. and Zehr, J. P. (2004). Determination of
nitrogen-fixing phylotypes in Lyngbya sp. and Microcoleus chthonoplastes cyanobacterial mats from Guerrero Negro, Baja California, Mexico. Appl Environ Microbiol 70: 2119-2128.
Ortega-Morales, O., Guezennec, J., Hernandez-Duque, G., Gaylarde, C. C. and Gaylarde, P. M.
(2000). Phototrophic biofilms on ancient Mayan buildings in Yucatan, Mexico. Curr Microbiol 40: 81-85.
Parker, D. L., Mihalick, J. E., Plude, J. L., Plude, M. J., Clark, T. P., Egan, L., Flom, J. J., Rai, L. C. and
Kumar, H. D. (2000). Sorption of metals by extracellular polymers from the cyanobacterium Microcystis aeruginosa f. flos-aquae strain C3-40. J Appl Phycol 12: 219-224.
Pizarro, C., Kebede-Westhead, E., and Mulbry, W. (2002) Nitrogen and phosphorus removal rates
using small algal turfs grown with dairy manure. J Appl Phycol 14:469–473.
Phillips, P., Russell, A., Bender, J. and Muñoz, R. (1994). Management plan for utilization of a floating
microbial mat with its associated detrital gelatinous layer as a complete tilapia Oreochromis niloticus feed system. Bioresource Technol 47: 239-245.
Prince, R. C. and Kheshgi, H. S. (2005). The photobiological production of hydrogen: potential
Chapter 2
30
Racault, Y. (1993). Pond malfunction: case study of three plants in the south-west of France. Water Sci Technol 28: 183–192.
Ragusa, S. R., McNevin, D., Qasem, S. and Mitchell, C. (2004). Indicators of biofilm development and
activity in constructed wetlands microcosms. Water Res 38: 2865-2873.
Rao, D. L. N. and Burns, R. G. (1990). The effect of surface growth of blue-green-algae and bryophytes
on some microbiological, biochemical, and physical soil properties. Biol Fertil Soils 9: 239-244.
Rectenwald, L.L., and Drenner, R. W. (2000) Nutrient removal from wastewater effluent using an
ecological water treatment system. Environ Sci Technol 34:522-526.
Revsbech, N. P., Jorgensen, B. B., Blackburn, T. H. and Cohen, Y. (1983). Microelectrode Studies of
the Photosynthesis and O-2, H2s, and Ph Profiles of a Microbial Mat. Limnol Oceanogr 28: 1062-1074.
Roeselers, G., van Loosdrecht, M. C. M. and Muyzer, G. (2004). Biodiversity of phototrophic biofilms. In
European Symposium on Environmental Biotechnology; ESEB 2004, pp. 161-164.
Roeselers, G., Zippel, B., Staal, M., van Loosdrecht, M. and Muyzer, G. (2006). On the reproducibility
of microcosm experiments - different community composition in parallel phototrophic biofilm microcosms. FEMS Microbiol Ecol 58: 169-178.
Roeselers, G., Norris, T., Castenholz, R., Rysgaard, S., Glud, R., Kuhl, M. and Muyzer, G. (2007).
Diversity of phototrophic bacteria in microbial mats from Arctic hot springs (Greenland). Environ Microbiol 9:26-38
Roeselers, G., van Loosdrecht, M. C. M. and Muyzer, G. (2007). Heterotrophic pioneers facilitate
phototrophic biofilm development. Microb Ecol. DOI: 10.1007/s00248-007-9238-x
Ruffroberts, A. L., Kuenen, J. G. and Ward, D. M. (1994). Distribution of cultivated and uncultivated
cyanobacteria and Chloroflexus-like bacteria in hot-spring microbial mats. Appl Environ Microbiol 60: 697-704.
Schäfer, H., Bernard, L., Courties, C. and other authors (2001). Microbial community dynamics in
Mediterranean nutrient-enriched seawater mesocosms: changes in the genetic diversity of bacterial populations. FEMS Microbiol Ecol 34: 243-253.
Schaub, B. E. M. and Vangemerden, H. (1994). Simultaneous phototrophic and chemotropic growth in
the purple sulfur bacterium Thiocapsa roseopersicina M1. FEMS Microbiol Ecol 13: 185-195.
Schumacher, G. and Sekoulov, I. (2002). Polishing of secondary effluent by an algal biofilm process.
Water Sci Technol 46: 83-90.
Schumacher, G., Blume, T. and Sekoulov, I. (2003). Bacteria reduction and nutrient removal in small
wastewater treatment plants by an algal biofilm. Water Sci Technol 47: 195-202.
Schutz, K., Happe, T., Troshina, O., Lindblad, P., Leitao, E., Oliveira, P. and Tamagnini, P. (2004).
Cyanobacterial H(2) production - a comparative analysis. Planta 218: 350-359.
Simek, K., and Chrzanowski, T.H. (1992) Direct and indirect evidence of size-selective grazing on
pelagic bacteria by fresh-water nanoflagellates. Appl Environ Microbiol 58: 3715-3720
Singh, R. N. (1950). Reclamation of 'Usar' lands in India through blue-green algæ. Nature 165: 325-326.
Sorkhoh, N., Al-Hasan, R., Radwan, S. and Höpner, T. (1992). Self-cleaning of the Gulf. Nature 359: 109.
Sorkhoh, N. A., al-Hasan, R. H., Khanafer, M. and Radwan, S. S. (1995). Establishment of
oil-degrading bacteria associated with cyanobacteria in oil-polluted soil. J Appl Bacteriol 78: 194-199.
Stal L. J., Vangemerden, H. and Krumbein, W. E. (1985). Structure and Development of a Benthic
Marine Microbial Mat. FEMS Microbiol Ecol 31: 111-125.
Steunou, A. S., Bhaya, D., Bateson, M. M., Melendrez, M. C., Ward, D. M., Brecht, E., Peters, J. W.,
Kuhl, M. and Grossman, A. R. (2006). In situ analysis of nitrogen fixation and metabolic switching in unicellular thermophilic cyanobacteria inhabiting hot spring microbial mats. Proc Natl Acad Sci U S A
103: 2398-2403.
Syed, M. A. and Henshaw, P. F. (2003). Effect of tube size on performance of a fixed-film tubular
bioreactor for conversion of hydrogen sulfide to elemental sulfur. Water Res 37: 1932-1938.
Tamagnini, P., Axelsson, R., Lindberg, P., Oxelfelt, F., Wunschiers, R. and Lindblad, P. (2002).
Hydrogenases and hydrogen metabolism of cyanobacteria. Microbiol Mol Biol Rev 66: 1-20
Thingstad, T.F. (2000) Elements of a theory for the mechanisms controlling abundance, diversity, and
biogeochemical role of lytic bacterial viruses in aquatic systems. Limnol Oceanogr 45: 1320-1328
Phototrophic biofilms and their potential applications
31 Tsygankov, A. A., Borodin, V. B., Rao, K. K., and Hall, D. O. (1999). H(2) photoproduction by batch
culture of Anabaena variabilis ATCC 29413 and its mutant PK84 in a photobioreactor. Biotechnol Bioeng 64: 709-715.
van Bleijswijk, J., and Muyzer, G. (2004). genetic diversity of oxygenic phototrophs in microbial mats
exposed to different levels of oil pollution. OPHELIA 58: 157-164.
van Dam, A.A., Beveridge, M.C.M., Azim, M.E., and Verdegem, M.C.J. (2002) The potential of fish
production based on periphyton. Rev Fish Biol Fish 12: 1-31
Vymazal, J. (1984) Short-term uptake of heavy metals by periphyton algae. Hydrobiologia 119: 171-179.
Vymazal, J., Sladedek, V. and Stach, J. (2001). Biota participating in wastewater treatment in a
horizontal flow constructed wetland. Water Sci Technol 44: 211-214.
Wang, T. C., Weissman, J. C., Ramesh, G., Varadarajan, R. and Benemann, J. R. (1998). Heavy metal
binding and removal by phormidium. Bull Environ Contam Toxicol 60: 739-744.
Ward, D. M., Ferris, M. J., Nold, S. C. and Bateson, M. M. (1998). A natural view of microbial
biodiversity within hot spring cyanobacterial mat communities. Microbiol Mol Biol Rev 62: 1353-1370.
Wiegand, C. and Pflugmacher, S. (2005). Ecotoxicological effects of selected cyanobacterial secondary
metabolites: a short review. Toxicol Appl Pharmacol 203: 201-218.
Wimpenny, J., Manz, W. and Szewzyk, U. (2000). Heterogeneity in biofilms. FEMS Microbiol Rev 24: 661-671.
Xu, Q., Barrios, C. A., Cutright, T. and Newby, B. M. (2005). Assessment of antifouling effectiveness of
two natural product antifoulants by attachment study with freshwater bacteria. Environ Sci Pollut Res Int
12: 278-284.
Zehr, J. P. and Turner, P. J. (2001). Nitrogen fixation: nitrogenase genes and gene expression.
Methods Microbiol 30: 271–285.
Zhang, K., Choi, H., Dionysiou, D. D., Sorial, G. A. and Oerther, D. B. (2006). Identifying pioneer
3
Diversity of phototrophic bacteria in microbial mats
from Arctic hot springs (Greenland)
Guus Roeselers, Tracy B. Norris, Richard W. Castenholz, Søren Rysgaard, Ronnie N. Glud, Michael Kühl, and Gerard Muyzer.
Chapter 3
34
ABSTRACT
We investigated the genotypic diversity of oxygenic and anoxygenic phototrophic microorganisms in microbial mat samples collected from three hot spring localities on the east coast of Greenland. These hot springs harbour unique Arctic microbial ecosystems that have never been studied in detail before. Specific oligonucleotide primers for cyanobacteria, purple sulfur bacteria, green sulfur bacteria and Choroflexus/Roseiflexus-like green non-sulfur bacteria were used for the selective amplification of 16S rRNA gene fragments. Amplification products were separated by denaturing gradient gel electrophoresis (DGGE) and sequenced. In addition, several cyanobacteria were isolated from the mat samples, and classified morphologically and by 16S rRNA-based methods. The cyanobacterial 16S rRNA sequences obtained from DGGE represented a diverse, polyphyletic collection of cyanobacteria. The microbial mat communities were dominated by heterocystous and non-heterocystous filamentous cyanobacteria. Our results indicate that the cyanobacterial community composition in the samples were different for each sampling site. Different layers of the same heterogeneous mat often contained distinct and different communities of cyanobacteria. We observed a relationship between the cyanobacterial community composition and the in situ temperatures of different mat parts. The Greenland mats exhibited a low diversity of anoxygenic phototrophs as compared with other hot spring mats which is possibly related to the photochemical conditions within the mats resulting from the Arctic light regime.
INTRODUCTION
Microbial mats are layered structures composed of physiologically different groups of microorganisms (van Gemerden, 1993). They are present in a variety of environments where grazing is limited, such as hot springs, shallow coastal lagoons, hypersaline ponds and permanently ice-covered lakes (Stal and Caumette, 1994). The top layer of microbial mats is typically dominated by oxygenic phototrophs, such as cyanobacteria and diatoms, with underlying or intermixed layers of anoxygenic phototrophs, i.e. green and purple sulfur bacteria (GSB and PSB) and green non-sulfur bacteria (Nübel et al., 2002; Martinez-Alonso et al., 2005). Cyanobacteria are the most important primary producers in these ecosystems (Stal, 1995). Their photosynthetic activity fuels a variety of mineralization processes and conversions catalysed by heterotrophic and lithotrophic bacteria in the mat community (Ward et al., 1998).
Microbial mats from Greenland
35 species exist in such microbial communities (Castenholz, 1996). Some of the East Greenland springs have been known since the early explorers visited Greenland, and the presence of coloured slimy coatings in the hot springs was mentioned in several early expedition journals (e.g. Nordenskjöld, 1907). Presence of cyanobacteria was also mentioned in the few botanical and geological surveys of the springs that have been reported (Halliday et al., 1974). However, the microbiology of the East Greenland hot springs was never explored in detail.
Chapter 3
36
Table 1. Source water characteristics at the three sampling sites
Spring Source Temperature (oC) Salinity (‰) pH Oxygen ((M)
Kap Tobin 60-62 13-16 7.0-7.5 5-8 Nørrefjord 53-57 9-13 8.2-8.6 6-8 Rømer Fjord 56-61 5-7 9.1-9.5 0
In July 2003 an expedition was carried out to perform the first microbiological field studies in hot springs at three localities (i.e. Nørrefjord, Rømerfjord and Kap Tobin) on the East Greenland coast (Kühl et al., 2004). In these localities, over 70 springs with source water temperatures of > 40°C were found. All springs, irrespective of pronounced differences in water chemistry and temperature, contained characteristic orange, green and brownish coloured gelatinous biofilms covering the sides and the bottom of the springs. Microscopic observation of these biofilms revealed that they were mainly composed of a dense network of filamentous cyanobacteria embedded in a slimy matrix of exopolymers. The biogeochemistry of these microbial mats will be presented elsewhere (M. Kühl, S. Rysgaard and R. N. Glud, unpublished data).
In this study we provide an inventory of the oxygenic and anoxygenic photoautotrophic microorganisms in the microbial mat samples collected during the expedition. The microbial diversity of these unique Arctic ecosystems was studied by genetic fingerprinting analysis of samples and pure cultures of cyanobacteria isolated from various mat samples. This approach enabled identification of the dominant cyanobacteria, PSB, GSB and green non-sulfur bacteria belonging to the Chloroflexaceae in the microbial mat communities.
MATERIALS & METHODS Environmental parameters of the springs
The mat samples used in this study were obtained from East Greenland hot springs at Kap Tobin (70°24.9'N; 21°57.04'W) and Nørrefjord (71°08.3'N; 22°2.1'W) on Jameson Land north of Scoresbysund, and at Rømerfjord (69°43.7'N; 23°41.9'W), on Blosseville's Coast south of Scoresbysund. The source water in the springs was characterized by temperature measurements with an electronic thermometer (Omnitherm, Germany), salinity determination with a calibrated refractometer (Atago, USA), pH determination with a portable pH-meter (Radiometer, Denmark) and oxygen determination by Winkler titration (Strickland and Parsons, 1972).
Sample collection
