Enrichment culture and identification of endophytic methanotrophs isolated from peatland plants

Aerobic methane-oxidizing bacteria (MOB) are an environmentally significant group of microorganisms due to their role in the global carbon cycle. Research conducted over the past few decades has increased the interest in discovering novel genera of methane-degrading bacteria, which efficiently utilize methane and decrease the global warming effect. Moreover, methanotrophs have more promising applications in environmental bioengineering, biotechnology, and pharmacy. The investigations were undertaken to recognize the variety of endophytic methanotrophic bacteria associated with Carex nigra, Vaccinium oxycoccus, and Eriophorum vaginatum originating from Moszne peatland (East Poland). Methanotrophic bacteria were isolated from plants by adding sterile fragments of different parts of plants (roots and stems) to agar mineral medium (nitrate mineral salts (NMS)) and incubated at different methane values (1–20% CH4). Single colonies were streaked on new NMS agar media and, after incubation, transferred to liquid NMS medium. Bacterial growth dynamics in the culture solution was studied by optical density—OD600 and methane consumption. Changes in the methane concentration during incubation were controlled by the gas chromatography technique. Characterization of methanotrophs was made by fluorescence in situ hybridization (FISH) with Mg705 and Mg84 for type I methanotrophs and Ma450 for type II methanotrophs. Identification of endophytes was performed after 16S ribosomal RNA (rRNA) and mmoX gene amplification. Our study confirmed the presence of both types of methanotrophic bacteria (types I and II) with the predominance of type I methanotrophs. Among cultivable methanotrophs, there were different strains of the genus Methylomonas and Methylosinus. Furthermore, we determined the potential of the examined bacteria for methane oxidation, which ranged from 0.463 ± 0.067 to 5.928 ± 0.169 μmol/L CH4/mL/day.


Introduction
Cultivation of methanotrophs was started in 1906 by Söhngen, who isolated Bacillus methanicus (now known as Methylomonas methanica) (Söhngen 1906). Since that time, the number and diversity of cultured methanotrophs have significantly increased. The crucial studies on cultivation of methanotrophs were performed by Whittenbury et al. (1970), who isolated more than 100 strains of these bacteria from various environments. The two mineral media containing nitrate or ammonium mineral salts (NMS and AMS) optimized by Whittenbury et al. are still widely used. So far, methanotrophic bacteria have been detected in different types of ecosystems both natural: soils, deserts, landfills, tundra, wetlands, rice paddies, sediments, lakes, marine (Hanson and Hanson 1996), as well as in the atmosphere (Santl-Temkiv et al. 2013), coal mines (Stępniewska et al. 2006), and mercury mines (Baesman et al. 2015) as well as engineered ecosystems, e.g., waste water treatment plant (Ho et al. 2013).
Wetlands are the main natural source of methane. This emission is the result of the balance between methanogenesis and methanotrophic processes and is actively affected by the composition of wetland plants, which can influence CH 4 production, consumption, and transport in the soil. Studies on this phenomenon indicated a significant role of methanotrophic bacteria in CH 4 emissions, both the free-living microorganisms in the rhizosphere attached to the root surface in the form of a biofilm (rhizoplane) and those living inside host tissues (endophytes) and colonizing parts of plants (Raghoebarsing et al. 2005;Liebner et al. 2011;Stępniewska et al. 2013;Stępniewska and Kuźniar 2013;Putkinen et al. 2014). The mechanism of the association of methanotrophs with peat vascular plants is poorly recognized and is currently the subject of studies undertaken by many researchers throughout the world (Fig. 1).
During the first step in methane oxidation to CO 2 , an important role is played by methane monooxygenase (MMO), which catalyzes the conversion of methane to methanol (Hanson and Hanson 1996). Methane monooxygenase occurs in two forms: membrane-bound, particulate (pMMO) and cytoplasmic soluble (sMMO). sMMO is responsible for the catalysis of oxidation of some hydrocarbons (saturated, unsaturated, linear, branched, and cyclic hydrocarbons, single-and double-ring aromatics, heterocycles, halogenated alkenes, and ethers (Dunfield et al. 1999)). All aerobic methanotrophs contain one or both MMOs. pMMO is coded by a pmoCAB operon and sMMO by genes including mmoXYBZDC. The pmoA and mmoX genes are usually used as functional markers for methanotrophic bacteria. Detection of pmoA or mmoX gene sequences from new isolates combined with determination of the 16S ribosomal RNA (rRNA) gene sequence is a necessary requirement for identification of methanotrophs (Dedysh and Dunfield 2014). mmoX is characteristic of most type II methanotrophs as well as in members of the genus Methylococcus and some Methylomonas strains (Shigematsu et al. 1999), Methylovulum miyakonense HT12 (Iguchi et al. 2011), but not in most of the other type I methanotrophs (Hanson and Hanson 1996).
Methanotrophs are multifunctional, harmless bacteria with promising applications in environmental bioengineering of methane removal and biodegradation of toxic compounds. The biodiversity of the methanotroph habitat especially with an established presence of sMMO gives many opportunities for application of these unique bacteria (Sullivan et al. 1998;Pandey et al. 2014).

Plant samples
Plant materials were collected from the Moszne Lake area located in the northwestern part of the Poleski National Park (51°23′ N, 23°63′ E), situated in the province of Lublin, in the Polish part of Polesie, where vegetation was dominated by Sphagnum magellanicum, Sphagnum fuscum, Sphagnum fallax, Aulacomium palustre, Polytrichum strictum, and Drosera rotundifolia with dense colonies of Vaccinium species, Eriophorum vaginatum, Eriophorum angustifolium, Carex rostrate, Carex nigra, and Carex gracilis.

Methanotrophic bacteria isolation and culture
Methanotrophic bacteria were isolated from plants C. nigra (C), Vaccinium oxycoccus (V), and E. vaginatum (E) by adding sterile fragments of each part of plant: roots (R) and stems (S) to agar mineral medium (NMS) (Whittenbury et al. 1970) and incubated in methane atmosphere 10% (v/v) CH 4 . Fragments of plants were sterilized by sequential immersion in 70% (v/v) ethanol for 5 min, and 0.9% (w/v) sodium hypochlorite solution for 20 min, and then surface-sterilized samples were washed in sterile water for three times to remove surface sterilization agents. Single colonies were streaked on new NMS agar media and, after incubation, transferred to liquid NMS medium. The cultures were incubated at 30°C and 180 rpm (Innova 42R, New Brunswick Scientific, USA) under atmospheric conditions supplemented with 1, 5, 10, and 20% (v/v) CH 4.
Bacterial growth dynamics was studied by absorbance and methane consumption. Changes in the methane concentrations were controlled during incubation by the gas chromatography technique. The headspace concentrations of gases (CH 4 , CO 2 , O 2 , N 2 ) were analyzed by a gas chromatograph Shimadzu GC 2010 (Japan) equipped with a flame ionization detector and a thermal conductivity detector, after CH 4 , CO 2 , O 2 , and N 2 calibrations. Bacterial growth dynamics was assessed by measuring absorbance at 600 nm (A 600 ) (UV1800, Shimadzu, Japan).

Calculation and statistical analysis
The methanotrophic activity of cultures was calculated from the slope of the regression line of the measured CH 4 concentration vs. time. Adjustments were made with r 2 ≤ 0.95 with Sigma Plot 10.0 software (USA) and written as micromole per liter CH 4 per milliliter of culture per day. MMO enzyme activity was characterized by a curve of initial velocity vs. substrate concentration, referred to as a Michaelis-Menten plot, with an inset Lineweaver-Burk plot. The maximum specific methane oxidation rate (Vmax) and saturation constant (Km) were determined using GraphPad Prism 6 software (USA).

DNA isolation and amplification
Bacterial DNA was isolated from the cultures with the method of Sambrook with own modifications. (Sambrook 1989). Cultured cells were harvested by centrifugation at 17500g for 5 min and subjected to lysis using 5 mol/L guanidine thiocyanate, 100 mmol/L EDTA, and 0.5% sarcosyl (pH 8.0). DNA was purified by extraction with ice-cold 7.5 mol/L ammonium acetate (10 min) and, subsequently, using a chloroform/3methyl-1-butanol (24:1, v/v) mixture (1 min). The two-phase mixture was centrifuged at 17500g for 10 min. The upper layer was collected into a new tube. DNA was precipitated at −20°C with 0.8 volumes of 2-propanol for 1 h. The pellet was separated by centrifugation at 17,500g for 30 min, rinsed five times with 70% (v/v) ethanol, dried under vacuum (RVC 2_18, Christ, Germany), and resuspended in 30 mL of ultrapure, DNAse-free water. The purity and concentration of the isolates were evaluated spectrophotometrically (UV_1800, Shimadzu, Japan). 16S rRNA genes were amplified from enrichment culture DNA using primers 27f (AGAGTTTG ATCMTGGCTCAG) and 1492r (TACGGYTACCTTGT TACGACTT), (DeLong 1992). The presence of the mmoX gene in the isolates encoding sMMO, which are key enzymes in the bacterial methane metabolism pathway, was used for authentication of the isolates. mmoX gene was detected due to the oxidation of methane in sMMO is critical for understanding the microbial applications of C-H activation in one-carbon substrates and limited diversity of sMMO-containing methanotrophs (Lee 2016). The presence of genes in the isolates was detected by partial amplification of the genes using specific primers. The following primer pairs were used for mmoX (mmoX1 CGGTCCGC TGTGGAAGGGCATG AAGCGCGT and mmoX2 -GGCTCGACCTTGAA CTTGGAGCCA TACTCG) (Miguez et al. 1997). The PCR reaction mixture (50 μL) contained PCR Master Mix (2×): 0.05 U/μL TaqDNA polymerase, reaction buffer, 4 mmol/L MgCl 2 and 0.4 mmol/L of each dNTP, (Thermo Scientific); forward and reverse primers at 0.1 mmol/L and 3 μL of template DNA. PCR amplification was performed in a TProfessional gradient system thermocycler (Biometria, Germany) using the following conditions: initial denaturation at 95°C (1 min); 30 cycles consisting of denaturation 95°C (0.30 min), annealing 55°C (1 min) for 16S rRNA, 62.5°C (0.55 min) for mmoX primers; extension 72°C (1 min); and final extension 72°C (5 min). Isolated DNA and PCR products were analyzed by gel electrophoresis on MiniSubR Cell GT (Bio_Rad Laboratories Ltd., UK), stained with ethidium bromide, and visualized in UV light in the Red™ Imaging System (Alpha Innotech, USA).

DNA sequence analysis
The identification of the cultured plant endophytes was performed after PCR product sequencing (Genomed S.A., Poland) and compared with sequences stored in NCBI (USA) using the BLASTN algorithm.  (Table 1).

FISH from endophyte cultures
FISH was performed according to Eller et al. (2001) with minor modifications using cells growing in the logarithmic phase. Cells were harvested by centrifugation and resuspended in 0.5 mL of phosphate-buffered saline (PBS). The suspensions were then mixed with 1 mL of 4% (w/v) paraformaldehyde and fixed for 3 h. The fixed cells were collected by centrifugation (6000g for 1 min) and washed twice with PBS. The resulting pellet was resuspended in 50% ethanol in PBS and stored at −20°C until use. The hybridization was carried out on slides where 15 μL of fixed cells suspensions were transferred and left to dry. Dehydration was performed by washing the slides in 50, 80, and 96% (v/v) ethanol for 3 min each. A 50-mL polypropylene falcon tube containing a slip of filter paper soaked in hybridization buffer was used as a hybridization chamber. Ten-microliter hybridization buffer (1 mol/L Tris-HCl (pH 8.0), 5 mol/L NaCl, 10% sodium dodecyl sulfate and 20% formamide), and 1 μL of a fluorescent probe solution was placed on each spot of the fixed cells. The specificity of the probes applied is presented in Table 2. The chamber was incubated for 90 min at the hybridization temperature (47°C). Then, the slides were washed at the hybridization temperature for 15 min in washing buffer (1 mol/L Tris-HCl, 0.5 mol/L EDTA, and 5 mol/L NaCl) and rinsed twice with distilled water. The slides were air-dried and stained with Vectashield Mounting medium containing DNA-staining DAPI (Vector Laboratories, USA). The slides were analyzed using fluorescence with a Nikon Eclipse 80i microscope. The pictures were taken with a Digital Sight camera (Nikon) and processed with software provided by the manufacturer.

Results
Endophyte enrichment culture growth Endophytes of C. nigra, V. oxycoccus, and E. vaginatum were grown on 5, 10, and 20% of CH 4 at 30°C. Cultivations of all methane-oxidizing bacteria (MOB) consortia were characterized by the dynamics of the gaseous phase. A steady decrease in the methane and oxygen concentration was shown, as well as accumulation of carbon dioxide (Fig. 2). The CH 4 consumption in the all investigated enrichment culture showed a Rapid growth of all investigated MOB consortia was observed; it was reflected by an increase absorbance in the range from about 0.1 to 0.8-1.8 after 3 to 6 days of cultivation, which was accompanied by a decline in the CH 4 mixing ratio in the headspace, while no growth was observed in the same medium in the absence of methane. The complete consumption of CH 4 by methanotrophic consortium cultures after 3 days under 1-10% CH 4 was demonstrated. At the highest substrate concentration (20% CH 4 ), the presence of 6% of methane was shown after 6 days of cultivation (Fig. 3).

Methanotrophic activity of the endophytes
On the basis of the consumption of methane during culture, the methanotrophic activity of the endophytes was determined. A clear decrease in the CH 4 concentration tested in the headspace was shown after 4-6 days of incubation. Methane consumption by the consortium isolated from the roots of wetland plants ranged from 0.463 ± 0.067 to 5.928 ± 0.169 μmol/L CH 4 /mL/day and from the stems of these plants from 0.499 ± 0.119 to 5.305 ± 0.022 μmol/L CH 4 /mL/day, depending on the initial CH 4 concentration (Fig. 4). The dependence of the methanotrophic activity of isolated methanotrophs was significantly related to the substrate concentration (p < 0.0001) and did not differ significantly between the plant species.

Kinetic parameters of CH 4 oxidation
The CH 4 oxidation rates were written correctly in the substrate saturation curves as a function of initial methane concentrations (Fig. 5) and showed typical Michaelis-Menten characteristics. Lineweaver-Burk plots based on regression of 1/V vs. 1/S are presented as insets in Fig. 6. The apparent Km and Vmax values obtained for the tested enrichment culture are summarized in Table 3. The methane oxidation by the methanotrophic consortium enrichment cultures followed the Michaelis-Menten mechanism. In a majority of the analyzed cases, the specific methane oxidation rates increased in parallel with the initial methane concentration, but only the CS consortium enrichment culture achieved the maximum rate at 10% methane 5.033 ± 0.823 μmol/L CH 4 /mL/day. Based on the Lineweaver-Burk plot (Fig. 5), Vmax, and Km were calculated in the range from 9.521 ± 1.280 to 7.539 ± 1.115 μmol/L CH 4 / mL/day and from 142.3 ± 40.69 to 95.42 ± 34.96 μmol/L CH 4 / mL/day, respectively (Table 3).

Fluorescence in situ hybridization
Characterization of MOB was performed by fluorescence in situ hybridization (FISH) with Mg705 and Mg84 for type I methanotrophs and Ma450 for type II methanotrophs. Microscopic observations confirmed the presence of methanotrophic microorganisms isolated from the plants and grown on NMS medium. The combination of oligonucleotide probes Mg84 and Mg705 with Ma450 allowed parallel detection of type I and type II methanotrophs with the predominance of type I. Type I cells (shown as green) were short 1.5-3-μm-long rods and were all similar in size and shape. Type II methanotrophs were less numerous but more diverse. These bacterial cells represented at least two distinct morphologies: straight (1-2 μm long) and irregular, curved bacilli (0.5-1 μm). Simultaneously, the staining with DAPI revealed that methanotrophic bacteria were present in the community with non-methanotrophic cells which are almost as numerous as the methanotrophic cells (Fig. 6).

Identification of cultured endophytic methanotrophs
Analysis of the partial 16S rRNA gene sequences of the isolated MOB showed that they clustered closely with homologous sequences from different strains of the genus Methylomonas and numerous uncultured bacteria (Table 4).
It was found that sequences obtained from the endophyte enrichment culture with C. nigra stream (PPN CS1) were related with 99-98% identity to Methylomonas sp.   (Table 5).

Discussion
It was confirmed that methanotrophic bacteria have great potential as microbial sinks for the greenhouse gas methane as well as for industry in different biotechnological solutions (Semrau et al. 2010). However, their application in biotechnology is limited by the number of suitable strains readily accessible, which were not specifically isolated for this purpose, and the lack of the necessary properties for efficient use on an industrial scale (Jiang et al. 2010). Among the disadvantages, the relatively slow growth rate of the available strains may be mentioned (Hoefman et al. 2010). In this context, searching for new methanotroph strains and investigation of methanotrophic bacterial communities takes on a new meaning.
The endophytic bacterial community isolated in our study from vascular peatland plants C. nigra (PPN CS1) and E. vaginatum (PPN ER1) are closely related to Methylomonas sp. belonging to type Ia methanotrophs, known as fast-growing MOB with a short generation time of 3.5 h (Whittenbury et al. 1970). Type II MOB is generally known to grow more slowly, with generation times ranging from 5 h up to several days (Whittenbury et al. 1970;Dedysh et al. 2000Dedysh et al. , 2002Vorobev et al. 2010). The investigated methanotrophic consortium cultures were characterized by rather rapid growth, reaching absorbance 0.8-1.8 after 3-6 days of cultivation, depending on the initial CH 4 concentration. As reported by literature, methanotrophs possessing pMMO show good growth with A 600 reaching 0.8-1.5 within 3-6 days. By contrast, methanotrophs with only sMMO may require up to 2-3 weeks until the cultures reaches A 600 of 0.2-0.3 (Dedysh and Dunfield 2014). The fast growth rate and Fig. 6 Fluorescent in situ hybridization of the consortium from C. nigra roots (a) and stems (b), V. oxycoccus stems (c) and E. vaginatum roots after hybridization with Mg84 and Mg705 probes (type I) green colored, Ma450 probe (type II) red colored, and DAPI staining probes (non-methanotrophic) blue colored   Kox et al. (2016), emphasize that CH 4 -N cycle interactions were found in the methanotrophs associated with submerged mosses. The presence of a methanotrophic diazotrophs and fixation of N 2 (diazotrophic activity) at a level of from 2 to 3 up to 35% of the N assimilated by Sphagnum sp. was demonstrated (Berg et al. 2013;Larmola et al. 2014). A similar interaction was observed in methanotrophic bacteria association with rice root (Minamisawa et al. 2016). These arguments testify to the fact that methanotrophs are an important component of the microflora of peat land. Literature describes a wide range of methanotoph applications and is constantly looking for new efficient strains for biotechnology applications. The richness of nonmethanotrophic bacteria associated with the endophyte community was confirmed by FISH analysis (Fig. 6). However, further studies are needed to determine whether the effect of the heterotroph richness is species specific and can be extrapolated to other methanotrophs.
In conclusion, the results of this study demonstrate that cultivable bacteria existing as endophytes of vascular plants belong to type I methanotrophs and are characterized by fast growth and relatively high methanotrophic activity. Moreover, we showed that the community composed of methanotrophs and non-methanotrophs has potential applications in biotechnology systems.