Aerobic methanotrophic bacterial communities in sediments of Lake Constance

sediments of Lake Constance

Dissertation

zur Erlangung des akademischen Grades des Doktors der Naturwissenschaften (Dr. rer. nat.) an der

Universität Konstanz, Fachbereich Biologie

vorgelegt von

Monali Rahalkar Konstanz 2006

Tag der mündlichen Prüfung: 21. Februar 2007

Referent: Prof. Dr. Bernhard Schink Referent: Prof. Dr. Andreas Brune

Dedicated To

My Dear Parents

Who have really done a lot for me….

This work was carried out under the supervision of Prof. Bernhard Schink in the laboratory of Microbial Ecology and Limnology chaired by him in the Faculty of Biology, University of Konstanz, from April 2003 to December 2006.

This work is a part of the Sonderforschungsbereich (SFB) 454 program-B1 subproject, supported by the Deutsche Forschungsgemeinschaft.

Words of thanks are less to express my gratitude towards Prof. Schink for giving me an opportunity to work with him as his PhD student. He is a great ‘Guru’!!

(Teacher: in sanskrit). His continuous enthusiasm about my work, support and deep knowledge in various aspects of Microbiology has been a driving force for me. I will always remember the scientific discussions with him and his suggestions throughout my life.

Next, I would like to thank Ingeborg Bussmann for helping me to learn different techniques in the cultivation of methanotrophs, especially gradient

cultivation. Her contribution to the methane and oxygen measurements and fruitful discussions is gratefully acknowledged.

I take the opportunity to thank Dr. Michael Hoppert, University of Göttingen, for performing electron microscopy of our strain. I am grateful to Alfred Sulger and his colleagues from the Limnology section for their help in sample collections.

I would further like to thank Prof. Peter Kroth for allowing me to use the Real time PCR machine and also for his help from time to time. I would also express my sincere thanks to Prof. Adamska for allowing me to use glass desiccators, which were really precious for growing my bacteria. Also the help from the members of her group is gratefully acknowledged.

spite of his busy schedule. I would also thank him for his advices and suggestions when he was there in Konstanz.

I would like to thank Uli Stingl, Dirk Schmitt-Wagner and Oliver Geissinger for their help when I was learning to use ARB. Thanks to Claudia Wilderer for her help during agar shakes. I also would like to thank Katrin Styp von Rekowskii for her help when I checked if my methanotroph was producing any AHLs. I also thank Christian Bruckner for his help during the nitrogenase assay.

Thank you Paula for your friendly advice, support and laugh. Sascha, thanks for the friendly conversations and your sense of humor. Further I would also like to thank Jörg Deutzmann for sharing his ideas, discussions and for translating the summary in german. Thanks to Ansgar Gubner from A G Kroth for the help in translation of some difficult sentences. I thank Britta for her last minute tips during the thesis preparation. I also thank Simone Gerhardt for her encouragement and wishes.

I express my sincere thanks to all my present and previous lab members and Bodo Philipp, from our group for creating a nice environment and for timely help.

Thanks to all my Indian friends especially Shanmu and Geetika for their help and of course to Farzana, for being always there with a helping hand.

Here I want to express my love and gratitude to my parents, my brother, in- laws and other family members and friends. Their constant support, wishes and care have been very important for me. |Last but not the least; it’s my dear husband, Rahul.

Without his care, support and help in everything, this thesis was just not possible.

pMMO – particulate methane mono-oxygenase sMMO – soluble methane mono-oxygenase pmoA – gene for alpha subunit of the pMMO

16S rDNA – gene coding for small subunit of ribosomal ribonucleic acid MOB – methane oxidizing bacteria

EPS – extracellular/exo- polymeric substance LO – cells grown under low oxygen tension HO – cells grown under high oxygen tension MO – cells grown under moderate oxygen tension ICM – intracytoplasmic membrane

MS – membrane stacks

ANME – anaerobic methane oxidizing archaea SRB – sulfate reducing bacteria

Bacteria are among the most fascinating organisms in the world. The word

‘incredible’ is appropriate for them, because they can do those processes which modern machines or reactors cannot do. With the advent of sequencing their whole genomes many exciting facts have been revealed. Several metabolic pathways, suite of enzymes are well packaged and controlled within the genomes within a small cell of about one µm diameter. This quality makes them the key players in element transformations, e.g. C, N, P, etc. Bacteria are the silent actors, transforming different forms of elements back and forth and cycling them, making life on Earth feasible.

Microbiology started with cultivation of bacteria in the laboratory and by studying their biochemical properties, so as to predict what metabolic processes they carried out in nature. Now with the help of molecular biological tools like cloning, analyzing metagenomes, many things can be studied or predicted without cultivation.

Cultivation remains still the method of choice because it opens the doors to infinite amount of information lying within the bacterium. With the notion that only 1% or less of the total bacteria can be cultivated the remaining 99% uncultured bacteria remain a challenge for microbiologists. It can be said that amidst all the difficulties in cultivation, these uncultivated groups are like a ‘mirage’ which microbiologists like to follow. My studies on methanotrophic or methane oxidizing bacteria (MOB) thus focus on improving the cultivation strategies of ecologically relevant methanotrophs by providing them more natural conditions. In addition, I have used cultivation independent approach and tried to address the question of “which methanotrophs are actually there and doing the job of methane oxidation in sediment of Lake

1

Methane 1

Methanotrophs or Methane Oxidizing Bacteria (MOB) 2

Phylogeny of Methanotrophs 3

Overview of genera 4

Aerobic Methane Oxidation 5

Enzymes for Methane Oxidation 7

Methanotrophic communities in Lakes 9

Cultivation-Independent Approaches 14

Cultivation of methanotrophs 20

Anaerobic Oxidation of Methane (AOM) 22

Perspective and Concept of this thesis 24

2

25

Abstract 25

Introduction 26

Materials and Methods 28

Results 36

Discussion 47

3.

nov., sp. nov., enriched by gradient cultivation from littoral sediment of Lake Constance

55

Summary 55

Introduction 57

Methods 58

Results and 64

Discussion

4.

74

Abstract 74

Introduction 75

Materials and Methods 76

Results 78

Discussion 82

5.

85

Abstract 85

Introduction 86

Materials and Methods 88

Results 94

Discussion 103

6.

Abstract 112 Introduction 113

Materials and Methods 115

Results 118 Discussion 126

7.

8.

9.

References 157

Contributions to this thesis 170

Appendix 171

A

a study from epilithic biofilms in Lake Constance

172

Abstract 172 Introduction 173

Materials and Methods 175

Results 179 Discussion 185

Chapter 1

Introduction

Methane

Methane gas is produced by both natural and anthropogenic sources and it is 26 times more effective as a greenhouse gas as compared to CO2 (Lelieveld et al., 1993). The discovery of methane gas goes back to 1776, a gas which was collected from stirred sediment and which ignited readily (Dalton, 2005). Methane gas has a strong absorbance of infrared radiations which are not able to escape from the Earth’s atmosphere, leading to global warming. The concentration of methane has increased from 0.75 – 1.75 ppm in the last 300 years at an enormous rate and might reach 4.0 ppm till 2050 (Ramanathan et al., 1985). The release of methane to the atmosphere results in an increase in global warming and causes changes in the chemical

composition of the atmosphere (Lelieveld et al., 1993). It has been predicted that increased levels of methane in the atmosphere will decrease OH radical

concentrations and thus increase the lifetime of methane in the atmosphere (Lelieveld et al., 1993) and finally would result in decrease in the tropospheric ozone

concentrations.

The major natural and anthropogenic sources of methane include natural wetlands, paddy fields, ruminants, termites lakes and oceans, landfills, oil recovery operations, and methane hydrates (Dalton, 2005).

Exactly 100 years ago, in 1906, the dutch microbiologist N. L. Sohngen isolated from pond water and aquatic plants a first bacterium growing on methane,

and named it Bacillus methanicus (Sohngen, 1906). This organism was unfortunately lost and not many methanotrophs were discovered until 1950 -1960s.

In 1970, Whittenbury and his coworkers (Whittenbury et al., 1970) started a new era of research on methanotrophs by isolating more than 100 strains which enabled the scientists to investigate their biochemical properties and pathways of aerobic methane oxidation in much detail.

Methanotrophs or Methane Oxidizing Bacteria (MOB)

Methanotrophs or Methane Oxidizing Bacteria (MOB) are a subset of

methylotrophs (Hanson & Hanson, 1996) and have the unique ability to use methane as the sole source of C and energy. Methylotrophs are capable of utilizing a wide variety of C-1 compounds including methane, methanol, methylamines, etc. MOB are specialized, as they utilize mainly methane and sometimes also methanol as their sole source of C and energy, and use molecular oxygen as the terminal electron acceptor (Hanson & Hanson, 1996). Methylotrophs are phylogenetically widespread and include archaea, eubacteria and yeasts. On the other hand, known MOB belong only to the α and γ subclass of Proteobacteria. With only one proven exception, i.e. the genus Methylocella, MOB are obligately methylotrophic and cannot utilize complex C compounds such as sugars or organic acids (Bowman, 2000). The evolutionary and physiological reasons for this, remain speculative (Dedysh et al., 2005). In Type I methanotrophs this could be due to the lack of certain enzymes in the Krebs cycle like α ketogulatarate dehydrogenase. In the recently published genome of Methylococcus

capsulatus it was shown that the genes coding all the necessary enzymes of the TCA cycle were present, but still this methanotroph cannot utilize organic compounds.

Another reason could be the lack of specific transporters for complex C compounds

Phylogeny of Methanotrophs

Whittenbury (Whittenbury et al., 1970) and his colleagues derived a

classification scheme for methanotrophs based on morphology, arrangement of intra- cytoplasmic membranes, pathway of C assimilation, nitrogen fixation ability,

presence of cysts or spores, colony color and motility, and grouped them into five genera, i.e. Methylosinus, Methylocystis, Methylomonas, Methylobacter and Methylococcus.

Using the following criteria, i.e. - numerical taxonomy, DNA-DNA

hybridization, PLFA analysis, physiological properties and phylogenetic relationships, Bowman and his colleagues (Bowman et al., 1993) (Bowman et al., 1995)

reorganized the phylogeny of methanotrophs. According to their scheme, the family Methylococcaceae (composed of Type I and Type X methanotrophs) should contain Methylococcus (Type X) and Methylomonas, Methylobacter, Methylomicrobium (Type I). The family Methylocystaceae (Type II methanotrophs) comprised the closely related groups Methylosinus and Methylocystis. After these rearrangements in phylogeny, seven more genera were added till 2006, which include Methylocaldum (Type X), Methylosphaera, Methylosarcina, Methylothermus, Methylohalobium (Type I), Methylocella and Methylocapsa (Type II).

Type I methanotrophs have intracytoplasmic membranes arranged in bundles of vesicular discs, 16-carbon phospholipid fatty acids, and assimilate formaldehyde by the ribulose monophosphate (RuMP) pathway. Type II methanotrophs are

characterized by assimilation of formaldehyde by the serine pathway, they have 18- carbon phospholipid fatty acids and paired intracellular membranes aligned to the periphery of the cell. The membrane arrangements of the type II genera Methylocella and Methylocapsa are, however, different from the other Type 2 methanotrophs. The

characteristics of methanotrophs with respect to their phylogeny, membrane arrangements and phospholipid fatty acids are the key characteristics in the identification of methanotroph isolates obtained from the environment (Bowman, 2000).

Overview of genera

A total of 13 genera of methanotrophs have been described so far :

Methylobacter, Methylomonas, Methylomicrobium, Methylosarcina, Methylothermus, Methylohalobium; and Methylosphaera belong to the Type I ; Type X have two genera Methylococcus, Methylocaldum, and Type II MOB include Methlyosinus, Methylocystis, Methylocella and Methylocapsa. Among the newly isolated genera, most are extremophilic. Methylocapsa and Methylocella (Dedysh et al., 2000) are acidophilic, Methylocaldum (Bodrossy et al., 1997) and Methylothermus are

thermotolerant and thermophilic, respectively, Methylosphaera (Bowman et al., 1997) is psychrophilic and Methylohalobius is halophilic.

Crenothrix polyspora : a well known bacterium, is a methanotroph!!

Crenothrix polyspora, a well known conspicuous filamentous bacterium with a complete lifecycle was discovered 135 years ago by Ferdinand Cohn (the founder of Bacteriology). Recently it has been discovered to be a methane oxidizer with an unusual methane mono-oxygenase (Stoecker et al., 2006). This bacterium is infamous for mass development in drinking water systems and wells and remained

uncharacterized till date. Characterization of 16S rDNA, methane-utilizing capacity and presence of pmoA sequences supported that it is a methanotroph.

Aerobic Methane Oxidation

Fig.1. Aerobic oxidation of methane. Abbreviations :FADH-formaldehyde dehydrogenase.

FDH-formate dehydrogenase, CytC-cytochrome c. Taken from: (Hanson & Hanson, 1996)

Pathways for the oxidation of methane and assimilation of formaldehyde

The first step in aerobic methane oxidation (Fig.1) takes place with the help of specialized enzymes known as methane mono-oxygenases (MMOs). These enzymes break the O-O bond in the dioxygen molecule by utilizing two reducing equivalents (Hanson & Hanson, 1996). One of the oxygen atoms is converted to water and the other one is incorporated into methane to form methanol. Methanol is further oxidized to formaldehyde by methanol dehydrogenase (MDH). Formaldehyde is the central metabolite in the anabolic and catabolic pathways. In the catabolic pathway, HCHO is further converted to formate and then to CO2. HCHO assimilation takes place by two different pathways in Type I and Type II methanotrophs. In Type I methanotrophs, it takes place by the ribulose monophosphate (RuMP) pathway for assimilation of formaldehyde.

The pathway is shown below: (Fig. 2)

Fig.2. RuMP pathway for HCHO assimilation in Type I methanotrophs

Taken from (Hanson and Hanson 1996).

The Type II methanotrophs use the serine pathway for assimilation of HCHO.

Fig.3. Serine pathway for assimilation of HCHO in Type II methanotrophs

Taken from (Hanson and Hanson 1996).

Enzymes for Methane Oxidation

Two methane mono-oxygenases are known to be present in methanotrophs, i.e. soluble and particulate (sMMO and pMMO). Almost all known methanotrophs possess pMMO except Methylocella, whereas the soluble MMO is not present in all methanotrophs. The sMMO utilizes NADH + H ⁺ as the electron donor and is a non- heme, iron-containing enzyme complex consisting of three components: hydroxylase (245 kDa in size), protein B, a regulatory protein and C is the reductase component.

The hydroxylase has three subunits, α, β and γ, of 60, 45 and 20 kDa respectively, which are arranged in a α2β2γ2 configuration. The α subunit contains a non-heme centre at the active site of the enzyme, where methanol is formed from methane and oxygen. The crystal structure of this enzyme is known and contains a carboxylate- bridged di-iron centre (Lieberman & Rosenzweig, 2005). The genes encoding sMMO have been cloned andsequenced from several methanotrophs, including

Methylococcus capsulatus (Bath) and Methylosinus trichosporium OB3b. In methanotrophs, these genes are clustered on the chromosome. mmoX, mmoY and mmoZ encode the α, β and γ subunits of the hydroxylase, respectively; mmoB and mmoC code for protein B and protein C , respectively. This enzyme has a broad substrate specificity and can co-oxidize a number of other aromatic compounds and hydrocarbons (Hanson & Hanson, 1996).

pMMO is expressed only when the copper supply in the culture medium is high (Prior & Dalton, 1985). All known methanotrophs except Methylocella palustris harbor pMMO (Dedysh et al., 2005). Unlike sMMO, pMMO has relatively narrow substrate specificity and can oxidize alkanes and alkenes of up to five carbons in length. However, it could be useful for biotransformation as well (DiSpirito et al., 1992). pMMO is composed of three subunits with molecular masses of 47 kDa (β,

pmoB subunit), 26 kDa (α, pmoA subunit) and 23 kDa (γ, pmoC subunit), encoded by pmoA, pmoB and pmoC genes, respectively (Semrau et al., 1995). Most of the

biochemistry, structure and mechanism of this predominant methane monooxygenase had remained unanswered in spite of the numerous efforts done in the last 20 years, until the crystal structure of pMMO was obtained (Lieberman & Rosenzweig, 2005).

The crystal structure of pMMO from Methylococcus capsulatus (Bath) was obtained to a resolution of 2.8 Aº. The enzyme is now known to be a trimer with α3β3γ3

polypeptide arrangement. Two metal centres modeled as mononuclear copper and dinuclear copper, are located in the soluble regions of each pmoB subunit, which resembles cytochrome c oxidase subunit II. A third metal centre, occupied by zinc in the crystal (coming from the buffer used), is located in the membrane and it is not clear what could be there in the original structure in the place of zinc.

There are two nearly identical copies of the genes encoding pMMO

(pmoCAB) in the chromosome of Methylococcus capsulatus (Bath) (Semrau et al., 1995; Stolyar et al., 1999) and a third, separate copy of pmoC has also been identified (Stolyar et al., 1999). Comparison of pMMO and ammonia monooxygenase (AMO) gene sequences suggests that pMMO and AMO could be evolutionarily related (Holmes et al., 1995). The pmoA, which encodes the α subunit of pMMO, has been shown to be evolutionary highly conserved among methanotrophs (Holmes et al., 1995). In methanotrophs such as Methylosinus trichosporium OB3b, which possess both pMMO and sMMO, there is a metabolic switch mediated by copper ions. When cells are starved forcopper, and the copper-to-biomass ratio of the cell is low, sMMO is expressed. Cells grown under conditions of excess copper express pMMO and there is no detectable sMMO expression (Murrell et al., 2000).

Habitats

Methanotrophs are ubiquitous, i.e. present in nearly all samples of swamps, sediment, rice paddies, oceans, ponds, soils from meadows, deciduous forest, streams, sewage sludge, etc. Wetlands and rice paddies are important sources of methane and oxic layers of sediments and soils act as biofilters for methane which is produced in the anoxic soil or sediment (Hanson & Hanson, 1996).

Methanotrophic communities in Lakes

Freshwater lakes contribute to a major portion, i.e. from 6 to 16 % of the global methane emissions, then thought before. Methane oxidation in lakes is an important process to prevent the escape of the methane produced in deeper anoxic sediment layers (Bastviken et al., 2004; Rudd & Hamilton, 1978; Thebrath et al., 1993) to the atmosphere, and eventually controls global warming.

In mesotrophic or oligotrophic lakes which are oxic down to the sediment, aerobic methane oxidation occurs at the sediment-water interface (Frenzel et al., 1990; Lidstrom & Somers, 1984). The availability and rate of production of methane, as well as the availability of oxygen and nitrogen, determines the location and rates of methane oxidation (Hanson & Hanson, 1996). In stratified, eutrophic lakes, methane oxidation occurs near the bottom of the chemocline (metalimnion), where dissolved- oxygen levels are relatively low compared with levels in the epilimnion during summer stratification. Very little methane is found in the oxygenated epilimnion, and most of the methane produced in the anoxic sediments is stored in the hypolimnion, which is also devoid of oxygen, or is oxidized in the metalimnion (Hanson & Hanson, 1996).

Methane oxidation in freshwater lakes was considered exclusively to be an aerobic process (Kuivila et al., 1988), (Eller et al., 2005a), (Frenzel et al., 1990) but

recently in Lake Pluβsee (Eller et al., 2005b), also anaerobic methane oxidation has been described.

Lake Washington

Lake Washington is a monomictic, mesotrophic freshwater lake close to Seattle. Different aspects of methane oxidation (Kuivila et al., 1988; Lidstrom &

Somers, 1984) as well as methanotrophic and methylotrophic communities have been studied in the profundal sediment of this lake (62 m deep site) (Auman et al., 2000;

Auman & Lidstrom, 2002; Costello & Lidstrom, 1999; Costello et al., 2002;

Kaluzhnaya et al., 2006; Nercessian et al., 2005). Cultivation-independent studies were focused on the natural communities of MOB where maximal methane oxidation occurred, i.e. the top 1 cm (Costello & Lidstrom, 1999). Later, both Type I and Type II MOB were isolated from this layer (Auman et al., 2000).

Further, the abundances of methanotrophic bacteria were determined by slot blot hybridization with Type I, Type II 16S rDNA and pmoA gene probes, PLFA (phospholoipid fatty acid) analysis, and methane oxidation rates (Costello et al., 2002). This was the first report where three independent quantitative studies were done with methanotrophs which were estimated to be about 10⁸ to 10⁹ cells per g dry sediment. Type I methanotrophs were found to be dominant, and at least one order of magnitude higher than Type II. Moreover the sMMO containing Methylomonas sp.

dominated the whole community (Auman & Lidstrom, 2002). Thus, a particular type of methanotroph could be dominant in a particular habitat, based upon its

characteristics.

The peak of methane oxidation was located in the top 7-8 mm (Kuivila et al., 1988), although the methane oxidation potential in 0.5 cm sections of the top 1.5 cm

suggesting that the number of methanotrophs in these layers did not differ

significantly. Recently, in order to identify the communities active in metabolism of different C1 compounds (methanol, methylamine, HCHO and formate), two novel approaches were used, analysis of mRNA and rRNA by using RT (reverse

transcription) PCR-based technique and DNA-SIP (Stable Isotope Probing) (Nercessian et al., 2005).

Lake Constance

Lake Constance is a pre-alpine lake and the second largest one in Europe.

Various aspects of methane oxidation in Lake Constance have been studied in great detail over the past years (Bosse et al., 1993; Bussmann et al., 2004; Bussmann, 2005;

Bussmann et al., 2006; Frenzel et al., 1990; Pester et al., 2004; Schulz & Conrad, 1995). Different from the profundal sediment, the littoral sediment is subject to changing environmental factors such as wind and waves, seasonal fluctuations of water level and temperature, and day-night cycles of light, and oxygen.

Aerobic methane oxidation in the surface layer of the profundal sediment (147 m) of Lake Constance has been studied (Frenzel et al., 1990). It was found that 93%

of the total methane was consumed under aerobic conditions and this contributed to around 9% of total O2 consumption. Further, it was found that phytoplankton blooms are responsible for seasonal variation in the concentration of the acetate, as the dominant substrate for methanogenic archaea (Schulz & Conrad, 1995). Another study was done to investigate activity and survival of bacteria from the deeper parts of the lake (Rothfuss et al., 1997). In this study, cultivable methanotrophic bacteria were found down to 32 cm depth but spontaneous methane oxidation (without induction with methane) was observed only in the upper 7.5 cm layer of the sediment. The deeper layers i.e. below 7.5 cm were active in methane oxidation after induction by

incubation with methane in the headspace. Thus, it was proven that the

methanotrophic bacteria might have survived many years of anoxic conditions and methane starvation. The role of spores or cysts has been predicted to be very important in this situation.

In the littoral sediment of Lake Constance, CH4 production rates varied from 5 mmolm^-2d^-1 in December to 95 mmolm^-2d^-1 in September (Thebrath et al., 1993).

Maximum amounts of CH4 were produced in the littoral zone in 2 – 5 cm depth (Thebrath et al., 1993), a large part of methane was found to be lost by ebullition.

Methane oxidation in the littoral sediment was recorded in the surface layers of the sediment (0-1.5 cm); although a large methane oxidation potential was also found to be in the deeper layers down to 5 cm. The inhibitory effect of ammonium ions on the surface sediment from the littoral zone of Lake Constance was observed (Bosse et al., 1993).

Recently, the effect of sediment resuspension by waves or bioturbation was studied in the littoral sediment and was found to be a major reason for methane release through the sediment (Bussmann, 2005). Methane profiles were recently recorded using a new methane sensor (Bussmann & Schink, 2006) based on (Rothfuss

& Conrad, 1994). This method has enabled us to measure methane profiles with more precision.

In littoral sediment of Lake Constance, MOB have been investigated by both culture-independent and cultivation-dependent methods. Here a stable and diverse community of both type I and type II MOB, and an apparent dominance of type I MOB could be documented with a T-RFLP and pmoA clone library approach (Pester et al., 2004). Attempts to optimise the cultivation conditions by modification of the

counts, but the diversity of the cultivated MOB still did not reflect the diversity of methanotrophs in this sediment (Bussmann et al., 2004).

Mono Lake California

Mono Lake, an alkaline salt lake located in California that undergoes seasonal stratification, is characterized by high salinity, and the methane is produced

biogenically as well as thermogenically. Two studies were conducted to correlate the patterns of methane oxidation and associated methane oxidizing communities (Carini et al., 2005; Lin et al., 2005). A temporarily shifting zone of methane oxidation was consistently associated with a micro-oxic zone that migrated upwards in the water column as stratification progressed. Although a stable number of methanotrophs over depth and over time was observed, the change in methane oxidation zones were attributed to small shifts in the activity and ratios of the MOB communities. The study on the community structure was based on pmoA clone library and DGGE at 4

different depths. Methylobacter- related methanotrophs dominated the clone libraries, and Methylocystis appeared to be stratified at the zone of highest methane oxidation.

Plußsee

Plußsee is a eutrophic lake and has a stable thermal stratification in summer and anoxia in the hypolimnion (Eller et al., 2005b). ¹³C signature and methane, oxygen and sulfide profiles determined the zones of aerobic and anaerobic methane oxidation.

The finding that aerobic and anaerobic methane oxidation occurs together was further supported by the detection of a relatively high number of aerobic MOB as well as a significant number of ANME in anoxic water column by fluorescent in situ

hybridization (FISH) (Eller et al., 2005b). This seems to be the first report where anaerobic oxidation of methane (AOM) in a freshwater lake has been investigated.

Other lakes and Chironomid Larvae

A comparative study has been done on methane oxidation in two lakes in Northern Germany, i.e. Holzsee and Groβer Binnensee (Eller et al., 2005a). Stable isotope analysis showed that MOB could be an important food source for chironomid larvae because of the highly depleted ¹³C signatures. These two lakes are heavily inhabited by chironomid larvae. It was shown that the larval ¹³C depletion was not due to differences in the community structure but due to the activities and sizes of the MOB communities.

Cultivation-Independent Approaches

Due to the difficulties in cultivating methanotrophs (Bowman, 2000), various efforts have been undertaken to explore the methanotrophic diversity by cultivation- independent approaches. Mainly the 16S rDNA sequences of methanotrophs, and some of the functional genes, pmoA, mmoX and mxaF have been used as phylogenetic markers. The diversity of methanotrophs has been assessed by cloning, T-RFLP, DGGE, and micro-array approach. For quantification of methanotrophs, fluorescence in situ hybridization (FISH), use of PLFA (phospholipid fatty acid) profiles, real time PCR, etc. have been used. Stabe isotope probing (SIP), an attractive method to link bacteria to their functions, has been used also used extensively, in relation to aerobic methane oxidation.

Community analysis:

Clone Libraries

In this method, a gene or a partial gene of interest is amplified from the DNA of an environmental sample and is cloned to generate a set of clones or a clone library. Molecular systematics of bacteria is based on the rRNA of the ribosomal

construction. Similar to DNA, RNA can be isolated, reverse transcribed and the cDNA clone libraries can be prepared. In case of methanotrophs, primers have been designed to specifically amplify partial 16S rDNA from Type I and Type II

methanotrophs (Wise et al., 1999). Clone libraries of 16S rDNA and functional genes like pmoA (subunit A of pMMO), mmoX (α subunit of sMMO), mxaF (methanol dehydrogenase), etc. have been used to study methanotrophs from a variety of habitats including rice fields (Horz et al., 2001), landfills (Wise et al., 1999), freshwater lakes (Costello & Lidstrom, 1999), marine environment (Holmes et al., 1995) and soil (Knief et al., 2003).

DGGE

Denaturing gradient gel electrophoresis (DGGE) works by applying a small sample of DNA (or RNA) to an electrophoresis gel that contains a denaturing agent.

Certain denaturing gels are capable of inducing DNA to melt at various stages. As a result of this melting, the DNA spreads through the gel and can be analyzed for single components. Sequence differences in otherwise identical fragments often cause them to partially melt at different positions in the gradient and therefore stop at different positions in the gel. TGGE (Temperature gradient gel electrophoresis) is based on the same principle, except it provides a temperature gradient instead of a chemical

gradient.

These are valuable fingerprinting techniques for studying microbial community structure. They separate a mix of PCR amplified gene fragments based on sequence differences and allow large number of samples to be analyzed simultaneously. They are particularly useful when, the dynamics of microbial communities is influenced by environmental changes. DGGE has been used for community analyses of 16S and

functional genes of methanotrophs in rice fields (Henckel et al., 1999) and (Eller &

Frenzel, 2001), in freshwater wetland communities (Bodelier et al., 2005).

T-RFLP

Terminal Restriction Fragment Length Polymorphism (T-RFLP) is a

molecular biological technique initially developed for characterizing natural bacterial communities. The technique works by PCR amplification of DNA using primer pairs in which one of the primers is labeled with fluorescent tags. The PCR products are then digested using RFLP enzymes and the resulting patterns visualized using a DNA sequencer or a gel. The results are analyzed either by simply counting and comparing bands or peaks in the T-RFLP profile, or by matching bands from one or more T- RFLP runs to a database (base clone library) of known species.

The T-RFLP approach is similar to DGGE and can be used to monitor changes or differences within different communities. (Horz et al., 2001) used this technique for the detection of methanotrophic diversity on roots of rice, coupled with clone libraries of mxaF, mmoX and type I methanotrophs. The only disadvantage of this method is that it requires a base clone library for comparison and cannot be used like DGGE directly. This technique has been used to study methanotrophs from Lake Constance (Pester et al., 2004). It was useful for analyzing the differences in methanotrophic communities in the littoral and profundal sediment of Lake Constance, and in addition the temporal and spatial (depth wise) differences. This technique was also used for monitoring the methanotrophic community growing in a cultivation-based approach (Bussmann et al., 2004) and comparing the T-RFLP profiles of the growing bacteria to the original community in the littoral sediment of Lake Constance.

Microarray approach

The database of pmoA genes has been used to create microarray analysis of the methanotrophs. This technique requires a large amount of efforts in the initial

establishment, but can be used as an efficient tool for analyzing the diversity of methanotrophs in new habitats. This microarray consisted of a set of 59 probes that covered the entire diversity of pmoA sequences available and could detect less dominant bacteria up to 5% of the total community (Bodrossy et al., 2003).

Stable Isotope Probing (SIP)

SIP is a technique that is used to identify the organisms in an environmental sample which use a particular substrate (Radajeweski et al., 2000) (Dumont & Murell, 2005). Thus, this method relates the identity of bacteria with their metabolic role in the environment. In this method, a substrate which is highly enriched in a stable isotope such as ¹³C is incorporated into the cellular components of the bacteria

utilizing it. Usually DNA/RNA being the most informative biomarkers are used which can be separated by density gradient centrifugation. The nucleic acids are further analyzed by amplification of 16S rDNA or functional genes followed by cloning, or more advanced techniques like metagenomic analysis can be combined to unravel the metabolic pathways of the active bacteria.

Methanol and methane were amongst the earliest and most ideal substrates used. SIP has been used for determining active methano- or methylotrophic populations in different habitats, like peat soil microcosms (Morris et al., 2002), Transbaikal soda lakes (Lin et al., 2004), in acidic forest soils (Radajeweski et al., 2002), freshwater lakes (Nercessian et al., 2005), Movile cave (Hutchens et al., 2004). The drawbacks of this method are 1. Longer incubation times can result in labeling those bacteria

which are cross feeding on the intermediates and 2. High substrate concentrations can select for the fast growing bacteria.

SIP coupled to Metagenomics

Metagenomics is the study of genomes recovered from environmental sample.

This relatively new field of genetic research allows the genomic study of organisms that are not easily cultured in a laboratory. SIP enables to link organisms in the environment to functions as described above. If the isolated heavy DNA is used for metagenomic analysis, i.e. isolation of large inserts from uncultivated bacteria, entire operons or set of genes can be analyzed from an environment. Recently, such an approach has been used for a forest soil sample, for which a prior SIP experiment was done (Radajeweski et al., 2002). In this study, an entire methane mono-oxygenase operon was found on an insert of a 15 kb clone (Dumont et al., 2006).

Fluorsecence in situ hybridization-flow cytometry-cell sorting-based method (FISH-FACS)

In this method FISH is carried out after extracting the cells from an environmental sample, followed by cell sorting. This method was used as a novel approach for the separation of Type I and Type II methanotrophs from the surface sediment of Lake Washington (Kaluzhnaya et al., 2006). This is a promising tool for genetic and genomic characterization of yet uncultivated and uncultured microbes.

Further advances in this method could employ mRNA based cell sorting followed by whole genome amplification (WGA).

Abundance distribution FISH

Fluorescence in situ hybridization (FISH) has been a breakthrough for the

1990). For identification and quantification of methanotrophs, FISH probes were designed by Eller et al (Eller et al., 2001). Although, before this study there were a few probes already existing which distinguished between γ proteobacterial and α proteobacterial methanotrophs (Holmes et al., 1995). In this study (Eller et al., 2001) a pair of probes for γ proteobacterial methanotrophs (Mγ705 and Mγ84).and a probe Mα450 for the detection of α proteobacteria was designed. These probes have fewer mismatches to the sequences of available methanotrophs and less false positive strains, and are most widely used. FISH has been used for detection and / or enumeration of methanotrophs from various habitats such as rhizoplane samples (Eller & Frenzel, 2001), in Sphagnum peat bogs (Dedysh et al., 2001) and in sediment of lakes and chirinomid larvae (Eller et al., 2005a).

Lipid Biomarkers

The use of specific phospholipids fatty acid biomarkers (PLFA) has been proven to be useful for determining the abundance of methanotrophs (Bowman, 2000). In methanotrophs, 16:1 ω8c is a characteristic biomarker used for Type I methanotrophs and 18:1 ω8c for Type II methanotrophs. The detection of signature fatty acids rely on the conformation of double bond geometry and isomeric state using gas chromatography coupled to mass spectrometry (GC-MS). This method lacks sensitivity, i.e., it is reliable only when the communities are (>1%) (Bowman, 2000).

Real Time PCR

Real Time PCR was developed basically for the quantitative determination of methanotrophs from soil because the MPN method or FISH are cumbersome and require large manual efforts (Kolb et al., 2003), whereas methods like lipid

biomarkers have limited ability to distinguish between subgroups of methanotrophs.

Since pmoA and 16S rDNA clusters in phylogenetic trees are congruent (Holmes et

al., 1995) and pMMO is present in all methanotrophs except Methylocella. These assays were based on pmoA with a detection limit of 10-100 cells (Kolb et al., 2003).

Recently, for quantification of aerobic and anaerobic methanotrophs from the black sea water column, real time PCR assays were developed specifically for Type I and Type II methanotrophs and ANME (anaerobic methane oxidizing archaea) (Schubert et al., 2006). Real time PCR assays have also been developed for unusual pmoA genes of Crenothrix polyspora (Stoecker et al., 2006) and for mmoX genes in Methylocella palustris (Dedysh et al., 2005).

Cultivation of methanotrophs

Cultivation of methanotrophs goes back to around 100 years, when Sohngen isolated the first methanotroph. The real landmark in the cultivation of methanotrophs was when Whittenbury and his colleagues isolated more than 100 methanotrophs.

Traditional methods

Whittenbury et al (1970) mainly used microscopy to detect tiny colonies of methanotrophs on the plates which were uncontaminated with heterotrophs and obtained > 100 cultures of methanotrophs. They also developed two media for cultivation of methanotrophs and named them NMS (Nitrate Mineral Salt medium) and AMS (Ammonium Mineral Salt medium). Till date this has remained to be the classical method and the classical media for isolation of methanotrophs and has yielded many new strains. Small variations have been made in the methodology e.g.

use of microtitre plates.

Non-conventional cultivation methods Gradient cultivation

Since methanotrophs are difficult to cultivate by classical plating-based

developed –‘Gradient cultivation’. Here, methanotrophs are grown in opposing gradients of methane and oxygen in agarose diffusion columns (Amaral & Knowles, 1995). Gradient cultivation allows both substrates, i.e., methane and oxygen to diffuse, and bacteria can develop wherever these gradients meet and grow as a band.

In this study, Type I methanotrophs were found at low methane and high oxygen concentrations (by hybridization studies) whereas Type II were found where methane concentrations were high and oxygen concentrations were low. When ammonium – based medium was used, nitrite was formed below the band creating toxic conditions.

In nitrate-based medium, denitrifying conditions were found below the band creating anoxic conditions and supplying nutrients to denitrifying bacteria (Amaral et al., 1995).

Thus, this method was basically designed to isolate novel methanotrophs which do not grow in conventional cultivation methods and offers some advantages with a possibility of chemical measurements, morphological description and nucleic acid studies. Although only one methanotroph has been isolated by this study,

Methylobacter sp. T20 (Ren et al., 2000), this method actually formed the basis of our cultivation strategy.

Soil Membrane Culture Method

This is a new method for isolation of methanotrophs from soil (Svenning et al., 2003). Soil samples are plated on polycarbonate membranes which were incubated in a methane-air atmosphere using a non-sterile soil suspension as a medium. The membrane was permeable and the soil acted as a buffer absorbing the excess of metabolites excreted, allowing the growth of methanotrophs not allowing other contaminants to grow. Both Type I and Type II methanotrophs were cultivated by this study as revealed by FISH.

Anaerobic Oxidation of Methane (AOM) Coupled to sulfate reduction

Anaerobic methane oxidation (AOM) is the major sink of methane in marine sediments. AOM is assumed to be a reversal of methanogenesis coupled to reduction of sulfate to sulfide and is believed to involve methanogenic archaea (ANME) and sulfate reducers (SRB) as syntrophic partners (Boetius et al., 2000).

The reaction is thought to be as follows-

CH4 + SO42- + H⁺= CO2 + HS^- + 2H2O (Strous & Jetten, 2004)

ANME archaea belong to three different groups and are known as ANME -, ANME-2 and ANME-3 although new groups also might be discovered, as newer habitats are surveyed. ANME-1 are distantly related to Methanosarcinales and

Methanomicrobiales. ANME-2 group are more closely related to Methanosarcina and ANME-3 to Methanococcoides species. The SRB belong to the Desulfosarcina- Desulfococcus (DSS) group and are known to be in aggregates with ANME 2. SRB of the Desulfobulbus branch have been associated with ANME-3 aggregates (Lösekann, 2006). Even if AOM have been reported to occur in a variety of habitats, none of the partner bacterium have been isolated yet or cultivated, and the enzymes and

biochemical pathways involved in AOM still remain unknown. Very recently, a candidate enzyme (Ni-protein) has been described that may catalyse the reverse methyl CoM reductase reaction (Krüger et al., 2003). ANME were first detected in sediments or cold seeps of various marine habitats, Black Sea sediment and hydrate ridge.

The habitats for ANME 1 and 2 are different. ANME 1 are found in deep parts of the sediment in completely anoxic conditions and ANME 2/DSS clusters are found

Coupled to nitrate reduction

Very recently, it was discovered that AOM can be also coupled to

denitrification (Raghoebarsing et al., 2006). This possibility was already discussed by (Strous & Jetten, 2004), until it was proven by Raghoebarsing and her colleagues that a microbial consortium can actually carry out this process.

The microorganisms that couple the anaerobic oxidation of methane (AOM) to denitrification, is shown in equations (1) and (2).

(Raghoebarsing et al., 2006)

The consortium was isolated from anoxic sediment from a canal in Netherlands, where the sediment was saturated with methane and nitrate

concentrations were up to 1 mM. After 16 months of enrichment, this consortium was completely dominated by only two partners: a bacterium representing a new phylum having no cultured bacterium (80%) and an archaeon (10%).

Perspective and Concept of this thesis

The main aim of my thesis is to explore the aerobic methanotrophic

communities dominant in the sediment of Lake Constance. Though my main focus is on studying the littoral sediment, for comparison purposes profundal sediment was also explored, as these two habitats share the same lake chemistry, but have different environmental conditions.

I have used cultivation-based and cultivation-independent approaches to fulfill this purpose. Various aspects of gradient cultivation enrichment technique, which we used for enrichments of the methanotrophs from the littoral sediment, are addressed in Chapter 2.

These include enrichment of methanotrophs followed by isolation and characterization of the methanotrophic communities in comparison with the methanotrophic communities in the sediment.

Then, a description of a novel strain of methanotroph follows in Chapter 3. This novel Type I methanotroph has been obtained after an initial gradient culture enrichment and we describe it as a new genus and species of Type I methanotrophs (Methylosoma difficile^T).

In the following two chapters (Chapter 3 and 4) I describe the community structure and abundance of methanotrophs occurring in the littoral and profundal sediments by molecular approaches. The abundance of methanotrophs has been compared with the distribution of methane and oxygen in the sediments.

In the last chapter (Chapter 6) I have studied some physiological and structural aspects of the new methanotroph described in Chapter 3 (Methylosoma difficile LC

Chapter 2

Cultivation of methanotrophic bacteria in opposing gradients of methane and oxygen

Ingeborg Bussmann*, Monali Rahalkar, Bernhard Schink

Published in FEMS Microbiology Ecology 56 (2006), 331-344

Abstract

In sediments, methane-oxidizing bacteria live in opposing gradients of methane and oxygen. In such a gradient system, the fluxes of methane and oxygen are

controlled by diffusion and consumption rates, and the rate-limiting substrate is maintained at a minimum concentration at the layer of consumption. Opposing gradients of methane and oxygen were mimicked in a specific cultivation set-up in which growth of methanotrophic bacteria occurred as a sharp band at either approx. 5 mm or approx. 20 mm below the air-exposed end. Two new strains of methanotrophic bacteria were isolated with this system. One isolate, strain LC 1, belonged to the Methylomonas genus (type I methantroph) and contained sMMO. Another isolate, strain LC 2, was related to the Methylobacter group (type I methantroph), as

determined by 16S rRNA gene and pmoA sequence similarities. However, the partial pmoA sequence was only 86% related to cultured Methylobacter species. This strain accumulated significant amounts of formaldehyde in conventional cultivation with methane and oxygen which may explain why it is preferentially enriched in a gradient cultivation system.

Key words: freshwater sediment, gradient cultivation, Lake Constance, methanotrophs, Methylobacter, Methylomonas, pMMO, sMMO

Introduction

Methanotrophic or methane-oxidizing bacteria (MOB) are an important group of bacteria that use CH4 as their sole source of carbon and electrons. There is an increasing interest in MOB because of their importance in greenhouse gas

consumption and their potential application in bioremedial degradation of industrial pollutants, e.g., trichloroethylene (Hanson & Hanson, 1996). MOB need both methane as electron donor and oxygen as co-reactant in the oxygenase reaction and as electron acceptor. In sediments, methane diffuses upwards from deeper sediment layers, and oxygen diffuses from the water column into the sediment. Both gases overlap at very low concentrations in the top few mm below the sediment surface where MOB can live in counter gradients of methane and oxygen. In this narrow zone, methanotrophic growth is limited by the diffusive transport of both substrates.

MOB include species in the Alphaproteobacteria (type II MOB) and in the Gammaproteobacteria (type I MOB) (Bowman, 2000). The oxidation of methane to methanol is catalysed by either a soluble or a membrane-associated form of methane monooxygenase (sMMO and pMMO, respectively) (Hanson & Hanson, 1996). The pMMO genes are almost universal in MOB. One gene of this operon, pmoA, is strongly conserved and can be used as a functional phylogenetic marker for MOB in general (Holmes et al., 1995).

In profundal sediment of Lake Washington, the enrichment of MOB with mineral medium (Whittenbury et al., 1970) led to the isolation of type I and type II MOB at almost equal numbers (5 and 6 strains out of 11, respectively). Two sMMO containing strains were isolated and assigned to the genus Methylomonas although this type of methanotrophs had not been reported before from a pristine environment

population in Lake Washington sediment consists of sMMO-containing

Methylomonas like MOB (type I) (Auman & Lidstrom, 2002). The number of type I MOB in this sediment, as estimated with cultivation-independent methods, is an order of magnitude higher than that of type II MOB (Costello et al., 2002).

In littoral sediment of Lake Constance, MOB have been investigated by both culture-independent and cultivation-dependent methods. Here a stable and diverse community of both type I and type II MOB, and an apparent dominance of type I MOB could be documented with a T-RFLP and pmoA clone library approach (Pester et al., 2004). Attempts to optimise the cultivation conditions by modification of the composition of the medium and the gas atmosphere resulted in increased viable counts, but the diversity of the cultivated MOB still did not represent the diversity of methanotrophs in this sediment (Bussmann et al., 2004).

Intermediates of methane oxidation such as methanol, formaldehyde, and formate have been detected in methanotrophic cultures, and they may reach even inhibitory concentrations (Agrawal & Lim, 1984; Costa et al., 2001). Production of formaldehyde and formate is favoured under unbalanced growth conditions if such bacteria are grown with methanol at high concentrations of oxygen. Removal of these intermediates, e. g., by a methylotrophic partner organism increases the methane oxidation rate of MOB (Wilkinson et al., 1974). Another way to avoid self- intoxication of MOB by possibly excreted toxic intermediates is the cultivation of these bacteria in counter gradients of methane and oxygen as this was first described by the lab of R. Knowles (Amaral et al., 1995; Amaral & Knowles, 1995).

The aim of our present study was to combine the gradient technique and an optimized mineral medium (Bussmann et al., 2004) for cultivation of novel and ecologically relevant methanotrophs from littoral sediment of Lake Constance. The

diversity of MOB growing in the gradients was compared with the total diversity of MOB in Lake Constance sediment on the basis of pmoA clone libraries.

Material and Methods

Study site and sediment sampling

Experiments were carried out with sediment from the lower infralittoral zone ("Litoralgarten", 47°41'N, 9°12'E) of Lake Constance, Germany. At the study site, CH4 concentrations in the sediment ranged from 20 to 90 µM at the sediment surface (Bussmann, 2005). The sediment consisted mainly of fine sand with a porosity of 0.62. Sediment cores (Ø 2.3 cm) were taken by SCUBA diving or with a sediment corer (∅ 8 cm) at 2–5 m water depth.

Cultivation of MOB in liquid or on solid media

Methanotrophs were grown in diluted mineral medium supplemented with 7 vitamin solution (Widdel & Pfennig, 1981) and were incubated at 16°C or 20°C in desiccators under an atmosphere of 17% O₂, 24% CH₄, 2% CO₂ and 57% N₂

(Bussmann et al., 2004). Solid media in plates contained 1.2% agarose. MOB were also grown in liquid medium in microtiter plates. For positive growth the OD595nm had to be 1.5 times more than the OD of a sterile control. To test for non-methanotrophic growth, cultures were streaked on plates with diluted complex medium (Bussmann et al., 2001) or Luria Bertani (LB) agar (Eisenstadt et al., 1984) and incubated without methane. Pure cultures of Methylobacter luteus type I and Methylosinus

trichosporium type II (a gift by Peter Dunfield, MPI Marburg) were grown in liquid NMS medium (Whittenbury et al., 1970).

Cultivation of MOB in gradients

Bacteria were cultivated in glass tubes (inner ∅ 8 mm, length 12 cm) with

Schuell) that were supported by perforated silicone septa. Agarose (0.2% w/v;

Agarose NEEO, Ultra Quality, Roth, Karlsruhe, Germany) was added to the diluted mineral medium to obtain a semi-solid consistency. Tubes were supplied with inoculum; then the anoxic and warm (38°C) medium was added and mixed immediately.

The incubation chamber carried 42 gradient cultivation tubes and consisted of two chambers (6.5 l volume each) separated by an intermediate bottom which held the cultivation tubes in gas-tight rubber seals. The upper chamber was filled with air; and the lower one was flushed for 20 min (to exchange its volume 3 times) with 2% CO₂, 24% CH4 and rest N2. The gas mixture was water saturated by passage through a washing flask to prevent evaporation from the tubes. The incubation temperature was either 16°C or 20°C. Tubes were checked once a week for presence of bands, and the gas mixture was renewed accordingly.

To verify if the observed bands were due to growth of MOB, the distribution of oxygen and methane was determined. Dissolved oxygen was measured with a Clark- type microelectrode (Ox50, Unisense, Aarhus, DenmarK) at 1 mm intervals. Methane concentrations were determined with a methane sensor modified after (Rothfuss &

Conrad, 1994).

Isolation of methanotrophs

Surface sediment (upper 1 cm) was used as inoculum. Two ml of sediment was mixed with 8 ml of mineral medium, and then further diluted (nine to ten steps).

These dilutions (0.3 ml) were used as inoculum for 3 ml of diluted mineral medium in gradient tubes. The final dilutions ranged from 2 × 10^-1 to 1 × 10^-7. Usually 4-5

replicates were set up for each dilution step, along with controls without methane, without oxygen, and without inoculum. After band formation was observed, the tubes

were removed from the box, the agarose column was pushed out with a rubber plunger, and the bands were excised aseptically with a sterile scalpel. Bands from replicate tubes of the last positive dilution were pooled, resuspended in liquid mineral medium, vortexed and around 500 µl was used for inoculation of another 1:10

dilution series with liquid medium. After incubation for 2-3 weeks, the last positive dilution tube was used again for inoculation of another dilution series until finally one single morphotype dominated. The last three positive dilutions were streaked on plates and incubated with and without methane. A binocular microscope was used to pick smaller colonies which were resuspended and streaked on fresh plates until a pure culture was obtained. Isolates were checked frequently for non-methanotrophic contaminants after streaking on complex media. The cultures were examined with a phase contrast microscope (Axiophot; Zeiss, Oberkochen, Germany) and

photographed using a cooled charge-coupled device camera (Magnafire, INTAS, Goettingen, Germany). Isolates were maintained at 4°C under a methane atmosphere for longer storage.

Alcian Blue staining

Polysaccharides produced after growth, in liquid or on solid medium or in gradients for three weeks were stained with Alcian blue (Hilger et al., 2000 ). Cell material was scraped from plates, liquid cultures were used directly, cell material from gradient cultures was cut from bands and suspended in ca. 300 µl of liquid mineral medium. Twenty µl of 1% Alcian blue solution in ethanol was diluted 1:10 with deionised water and mixed with ca. 20 µl of sample. Negative controls were prepared with pure agarose, to check for staining of agarose.

Chemical analyses

Formaldehyde was analyzed in the gas phase of incubation vessels by gas chromatography. Standards were prepared in glass tubes closed with butyl rubber stoppers. Formaldehyde standards were prepared from a fresh 37% (w/v)

formaldehyde solution (Merck, Darmstadt, Germany) ranging from 0.01% to 2%

(v/v). The formaldehyde concentration in the gas phase was estimated according to (Grützner & Hasse, 2004), gas phase- liquid equilibrium were calculated according to (Flett et al., 1976).

DNA extraction and PCR amplification

DNA was extracted from cell material, by a combination of enzymatic lysis (Ohkuma & Kudo, 1996) and bead beating (Henckel et al., 1999) with the following modifications: Cell material from gradient culture bands (200-500 µl) was used for DNA extraction. In case of plates, colonies were scraped or cell pellets were obtained from 1 - 2 ml of liquid cultures after centrifugation for 10 min at 13,000 rpm, 4 °C.

Cell material was suspended in 800 µl buffer (100 mM Tris HCl, pH 8.0, 50 mM EDTA) and homogenized with plastic pestles (Micropistill sticks, Eppendorf,

Hamburg, Germany). The homogenates were transferred into screw-cap tubes with 0.7 g silica beads (0.1 mm diameter) and lysozyme (5 mg / ml). After incubation for 20 min. at 37°C, proteinase K (100 µg / ml) was added and the mixture was incubated again at 37°C for 40 min. After bead-beating (6.5 m/s, 45 s), proteins and debris were removed by two times washing with chloroform:isoamyl alcohol (24:1 v/v) in phase lock gel tubes (Eppendorf). The DNA was finally precipitated with 0.7 volume of isopropanol and harvested by centrifugation at 20800 g for 60 min, followed by removing the salts with 70% (v/v) ethanol and drying. The DNA was resuspended in c. 50 µl 10 mM Tris-EDTA buffer and stored at -20°C. DNA from pure cultures was used for amplification of 16S rRNA genes, using 27f (Edwards et al., 1989)and 1492r (Weisburg et al., 1991) universal primers. DNA from gradient cultures and pure cultures was used for the amplification of partial pmoA gene, using the pmoA primer pair A189f-mb661r (Costello & Lidstrom, 1999), additionally with isolates,

amplification of partial pmoA gene was also checked with primer pair pmoA A189f- A682r (Holmes et al., 1995). For amplification of the mmoX gene, primers mmoXA and mmoXB were used.

For the construction of a sediment pmoA clone library littoral sediment (upper 1cm layer) was collected in August 2005. DNA was extracted with the using

PowerSoil™ DNA Isolation Kit (MO BIO Laboratories, Inc., Solana Beach, CA).

All amplifications were carried out in 50 µl or 25 µl total volume in an Eppendorf thermal cycler using recombinant Taq DNA polymerase (Eppendorf) or FailSafe™ Enzyme Mix and Premix B (Epicentre, Madison, WI) for clone library construction. For amplification of 16S rRNA genes, an initial denaturation at 94°C for 3 min was done, followed by 32 cycles at 94°C for 30 sec, 53°C for 30 sec and 72°C

I and Type II methanotroph specific primers (Wise et al., 1999) as well as for partial pmoA and mmoX genes, the following program was used: Initial denaturation at 94°C for 3min, followed by 32 cycles at 94°C for 30 sec, 56°C for 1min and 72°C for 90 sec, followed by final extension at 72°C for 10 min. PCR products were checked for amplification on 1.5% agarose gel by electrophoresis.

Clone libraries and restriction fragment length polymorphism (RFLP) pmoA clone libraries from sediment and from gradient culture bands were prepared by cloning the partial pmoA gene product (508 bp), obtained after amplification with primers A189f-mb661r (Costello & Lidstrom, 1999) and

purification of the PCR products. PCR products were purified using the Qiagen PCR purification kit (Qiagen, Hilden, Germany). All cloning steps were done using TA cloning kit (Invitrogen). In case of libraries with gradient culture bands, thirty clones from each clone library were selected randomly and were subjected to tooth pick PCR, using primers A189f-mb661r. The amplified products were digested with Msp I (5 U, MBI Fermentas), separated on 3.5% Nu-Sieve agarose (NuSieve ® 3:1 Agarose, Cambrex Bio Science Rockland Inc., ME), grouped according to their restriction patterns and each clone was assigned to an operational taxonomic unit (OTU) which represented a unique RFLP. Sediment clone libraries were analysed similarly, except more clones i.e. around seventy clones were randomly picked to cover the entire diversity and were digested with MspI/HaeIII (5 U, MBI Fermentas) and grouped as described above.

16S rRNA gene clone libraries were performed with DNA extracted from gradient bands. DNA was PCR amplified with the universal bacterial primers 27f (Edwards et al., 1989) and 1492 r (Weisburg et al., 1991) and cloned separately as

described above. Clones were digested with MspI restriction enzymes, RFLP analysis was done and OTUs were assigned as described above.

Cloning, sequencing, and phylogenetic analysis

With our isolates strain LC 1 and LC 2, complete 16S rRNA gene sequences were obtained by cloning the fragments using the TA cloning kit (Invitrogen). Clones were sequenced with primers 27f (Edwards et al., 1989), 533f (Lane et al., 1985), 1492r (Weisburg et al., 1991), MethT1dR (Wise et al., 1999) and assembled using the DNAStar software (http://www.dnastar.com). Similarly, sequences of the partial pmoA gene of the isolates and the partial mmoX gene of strain LC 1 were obtained by direct sequencing of the PCR products, and a complete sequence was obtained after cloning the fragment using TA cloning kit and sequencing from both ends.

In case of pmoA and 16S rRNA gene clone libraries, representative clones from each OTU group were sequenced. At least 10% of clones from each RFLP group were sequenced. pmoA and mmoX clones were sequenced using M13f and M13r primers.

Representative clones from 16S rRNA gene clone library were either sequenced completely with 27f (Edwards et al., 1989), 1492r (Weisburg et al., 1991) (clones representing dominant OTU groups) or partially with 27f primer (clones which were less frequent). All sequencing reactions were done at GATC Biotech AG (Konstanz, Germany). Blast search was performed at the NCBI site

(http://www.ncbi.nlm.nih.gov/) (Altschul et al., 1990) and closely related sequences were retrieved. All sequences were checked for chimeras by dividing the sequence in two partial sequences and performing blast search. Two chimeras were found in 16S rRNA gene clone libraries.16S rRNA gene sequences of strain LC 1 and strain LC 2 were phylogenetically analysed using the ARB software package (version 2.5b;

ARB database and aligned using the FAST Aligner tool as implemented in ARB.

Alignments were checked and manually corrected where necessary. Sequences with more than 1400 nucleotides were used for alignment. Only those positions which were identical in 50 % of all sequences were used to create a filter. Phylogenetic analysis was done using the maximum likelihood, neighbour-joining and maximum parsimony algorithms as implemented in ARB. Phylogenetic distances were

determined by calculating the similarity matrix within ARB using E.coli 16S rDNA genes as filter.

For phylogenetic analysis of pmoA gene sequences they were translated within ARB to obtain deduced amino acid sequences and phylogenetic distance dendrograms were constructed using different methods such as Neighbour-joining, Desoete, and PHYLIP with the Fitch and Margoliash method (Felsenstein, 1989), using

representative sequences of pmoA clones and isolates obtained in earlier studies done on Lake Constance as well as other studies (Pester et al., 2004).

All sequences have been deposited in GenBank under accession numbers DQ119042- DQ119051 (sequences from gradient clones and isolates) and DQ235456- DQ235470 (sequences from sediment pmoA clone library).

Results

Growth of MOB in gradient cultures

Initial experiments had shown that gradients of oxygen (and we assume also methane gradients) establish within 3 - 4 days in a 4.5 cm agarose column.

Enrichment cultures from sediment developed 0.5 mm thick bands of bacterial growth after 2 -3 weeks of incubation. A narrow, homogeneous band, rather than single colonies, was taken as indication of methanotrophic growth in gradient tubes, as described in (Amaral & Knowles, 1995). If methane was excluded from the tubes such bands were never observed. Bands occurred typically approx. 5 mm below the air-exposed end, ranging from 2 – 5 mm. In some cases, a thin band was observed at approx. 20 mm below the air-exposed end, ranging from 20 – 25 mm. In older

enrichment culture tubes (> 4 weeks) further bands developed like shadows below the first band. The distribution of oxygen and methane in these gradients was also

measured (Fig. 1.). Oxygen penetrated only to the depth of the growth band (5 or 20 mm), and methane was not detectable above the bands. In a sterile control tubes, oxygen penetrated much deeper into the column and methane reached up to the surface. Thus, the growth bands were always observed where the concentrations of oxygen and methane approached zero.

Subsequent transfers of the bands into new gradient tubes were often not successful. After the 4^th transfer, growth or bands could not be observed anymore. To check for possible reasons for this failure we performed various experiments.

Enrichments in the gradient tubes were started with the medium optimised for MOB growing in liquid cultures. Therefore we checked if the MOB growing in the gradient tubes prefer a different medium composition. Cell material from two to three

bands from initial sediment enrichments was pooled, and aliquots were transferred into the same medium again or into modified media. After incubation for three weeks, tubes were checked for growth bands. Each medium modification was tested three times. Increasing the phosphate and nitrate content (150 µM K-Na-PO₄, 50 µM NO₃) to 2-, 5- or 10-fold did not result in better growth. Addition of organic supplements (7 vitamin solution, 0.05% yeast extract, or 0.05% prefermented yeast extract), different buffers (0.01 M MOPS, 0.01 M TES or 0.01 M K-Na-PO4) and different mineral composition (standard medium, full-strength medium according to Widdel (Widdel, 1988), medium according to Whittenbury (Whittenbury et al., 1970)or (Heyer et al., 1984) had no influence on growth of transferred cultures.

Cultures of MOB that had been cultivated always in liquid or solid medium were checked for growth in gradient cultures. Exponentially growing liquid cultures of Methylobacter luteus and Methylosinus trichosporium were inoculated i) into freshly prepared liquid medium with warm agarose, ii) into already solid medium in tubes stored under nitrogen, iii) on the surface of solid medium with a 3 day-old gradient, or iv) 5 mm below the surface of solid medium with a 3 day-old gradient.

Both strains grew in the gradient tubes. Bands looked best (sharp, homogenous) when inoculated into freshly prepared medium.

In an additional experiment, we tested how many cells of MOB were necessary to form a band in gradient tubes, compared to formation of turbidity in a microtiter plate. An exponentially growing culture of M. trichosporium was counted

microscopically and diluted in 1:2 steps down to 3 cells per ml. Aliquots were transferred into gradient tubes (3 ml) and into microtiter plates (240 µl) resulting in the same cell number per vial. Gradient tubes were incubated in the incubation