Introduction
Methane (CH4), the second-most important anthropogenic greenhouse gas after
CO2, is the most abundant reduced organic compound in the atmosphere
and plays a central role in atmospheric chemistry (IPCC, 2013; Kirschke et
al., 2013; Lelieveld et al., 1998). The mixing ratio of CH4 in the
atmosphere has been increasing from preindustrial values of around 715 ppbv
(parts per billion by volume) to about 1800 ppbv in 2010 (Kirschke et al.,
2013). In total, annual CH4 emissions from natural and anthropogenic
sources amount to 500–600 Tg (1012 g) yr-1. They derive from
various terrestrial and aquatic sources and are balanced primarily by
photochemical oxidation in the troposphere (≈ 80 %), diffusion
into the stratosphere, and microbial CH4 oxidation in soils.
Until recently, natural sources of atmospheric CH4 in the biosphere have been considered to originate solely from strictly anaerobic microbial
processes in wetland soils and rice paddies, the intestines of termites and
ruminants, human and agricultural waste, and from biomass burning, fossil
fuel mining, and geological sources including mud volcanoes, vents and seeps.
However, more recent studies have suggested that terrestrial vegetation,
fungi, and mammals may also produce CH4 without an input from
methanogens and under aerobic conditions (Bruhn et al., 2012; Ghyczy et al.,
2008; Keppler et al., 2006; Lenhart et al., 2012; Wang et al., 2013b; Liu et
al., 2015). A fraction of these vegetation-derived emissions might be
released directly by in situ formation in plants (Bruhn et al., 2012; Keppler et
al., 2009; Wang et al., 2013a), and it is now apparent that several pathways
exist by which CH4 is generated under aerobic conditions (Bruhn et al.,
2014; Messenger et al., 2009; Wang et al., 2013b). Hence, the biogeochemical
CH4 cycle appears to be even more complex than previously thought.
In particular, the biogeochemical cycle of CH4 in the oceans is still
far from being understood. The world's oceans are considered to be a minor
source of CH4 to the atmosphere with approximately 0.6–1.2 Tg CH4 yr-1 (Rhee et al., 2009). Concentrations of CH4 in near-surface
waters are often 5–75 % supersaturated with respect to the atmosphere, implying a net flux from the ocean to the atmosphere (Conrad, 2009;
Reeburgh, 2007; Scranton and Brewer, 1977). Because the surface ocean is
also saturated or slightly supersaturated with oxygen, which does not favor
methanogenesis, the observed CH4 supersaturation has been termed the
oceanic methane paradox (Kiene, 1991). To explain the source of CH4 in
surface waters, it has been suggested that methanogenesis takes place in
anoxic microenvironments of organic aggregates (Grossart et al., 2011; Karl
and Tilbrook, 1994; Bogard et al., 2014), the guts of zooplankton or fish
(de Angelis and Lee, 1994; Oremland, 1979), and inside bacterial cells (Damm
et al., 2015). It has also been shown that contrary to the conventional
view, some methanogens are remarkably tolerant to oxygen (Angel et al.,
2011; Jarrell, 1985).
A potential substrate for methanogenesis in such anoxic microniches is
dimethylsulfoniopropionate (DMSP) (Damm et al., 2008, 2015; Zindler et al.,
2013), an algal osmolyte that is abundant in marine
phytoplankton and serves as a precursor of dimethyl sulfide (DMS) and
dimethyl sulfoxide (DMSO) (Stefels et al., 2007; Yoch, 2002) For example, Zindler et al. (2013) measured concentrations of DMS, DMSP, DMSO, and
CH4, as well as various phytoplankton marker pigments in the surface
ocean along a north–south transit from Japan to Australia. Positive
correlations between DMSP (dissolved) and CH4, and DMSO (particulate
and total) and CH4, were found along the transit. Based on their data, they concluded that DMSP and DMSO and/or their degradation products serve as
substrates for methanogenic archaea in the western Pacific Ocean.
Damm et al. (2010) hypothesized that under N limitation and a concomitant
availability of phosphorus, marine bacteria use DMSP as a carbon source and
thereby release CH4 as a by-product and its production could yield
energy under aerobic conditions. Methanethiol, a further potential
degradation product of DMSP, may act as a direct precursor of methane in
aerobic environments. By reason of thermodynamic calculations the authors
considered it possible for microorganisms to yield energy from the pathway of
methanethiol formation operating in its reverse direction, whereby methane
is formed.
An alternative non-biological CH4 formation pathway in seawater might
occur via a photochemical pathway due to the formation of methyl radicals;
however, photochemical production of CH4 in oceans is thought to be
negligible under oxic conditions (Bange and Uher, 2005).
In addition, Karl et al. (2008) suggested that CH4 is produced
aerobically as a by-product of methylphosphonate (MPn) decomposition when
aerobic marine organisms use methylphosphonic acid as a source of phosphorus
when inorganic sources of this element are limited. Furthermore, a mechanism
has been identified that leads to the formation of CH4 from MPn via
enzyme-catalytic cleavage of the C–P bond (Kamat et al., 2013). The
critical issue with this pathway is that MPn is not a known natural product nor has it been detected in natural systems. However, it was recently shown
that the marine archaeon Nitrosopumilus maritimus encodes a pathway for MPn biosynthesis and that it
produces cell-associated MPn esters (Metcalf et al., 2012). They argued that
these cells could provide sufficient amounts of MPn precursor to account for
the observed CH4 production in the oxic ocean via the C-P lyase-dependent scenario suggested by Karl et al. (2008). However, it was not
possible to explain the supersaturation state of CH4 in oxic surface
water by the quantification of produced CH4 from dissolved MPn under
natural conditions (del Valle and Karl, 2014).
Overview of sample collection during the incubation of E. huxleyi.
Day
0
1
2
3
4
5
6
7
8
9
10
Headspace
CH4
x
x
x
x
x
x
x
δ13CH4
x
x
x
x
x
x
x
Water
cell density
x
x
x
x
x
x
x
x
x
It remains uncertain whether CH4 formation from MPn (Karl et al., 2008)
or the metabolism of DMSP by methanogens in anoxic microenvironments (Damm et al.,
2008, 2015; Zindler et al., 2013) is sufficient to provide a
permanent increase in the concentration of CH4 in oxygenated surface
waters or whether other pathways are also required to fully explain the CH4 oversaturation in oxic waters. In this context it is important to
note that almost 40 years ago researchers (Scranton and Brewer, 1977; Scranton
and Farrington, 1977) already mentioned the possibility of in situ formation of
CH4 by marine algae. These scientists measured CH4 saturation
states in open-ocean surface waters of the west subtropical North Atlantic.
They observed 48–67 % higher CH4 concentrations in surface waters
than estimated from atmospheric equilibrium concentration, with a narrow
maximum of CH4 concentration in the uppermost part of the pycnocline.
Since the loss of CH4 from the surface to the atmosphere was calculated to be
much larger than diffusion from CH4 maxima of the pycnocline into the
mixed layer, an in situ biological CH4 formation process within the mixed
layer was hypothesized (Scranton and Farrington, 1977; Scranton and Brewer,
1977). However, direct evidence of algae-derived CH4 formation from
laboratory experiments with (axenic) algae cultures is still lacking, and
the accumulation of CH4 in the upper water layer has not yet been
directly related to production by algae.
The aim of our study was to quantify in situ CH4 formation from marine algae
such as coccolithophores and to identify precursor compounds of CH4 via
13C labeling techniques. Therefore, we used Emiliania huxleyi, a widely distributed,
prolific alga. The coccolithophore blooms including E. huxleyi are the major regional
source of DMS release to the atmosphere (Holligan et al., 1993). Specific
goals in this study were (I) to measure the CH4 production of a
biogeochemically important marine phytoplankton, (II) to screen for
methanogenic archaea or bacteria, and (III) to identify methyl sulfides, such
as the amino acid methionine, which play a role in metabolic pathways of
algae, as possible precursors for CH4.
Materials and methods
Culture media and culture conditions
Monoclonal cultures of E. huxleyi (RCC1216; http://roscoff-culture-collection.org/) were grown in full-batch mode
(Langer et al., 2013) in sterile filtered (0.2 µm) seawater
(Helgoland, North Sea) enriched with phosphate, nitrate, trace metals, and
vitamins according to F/2 (Guillard and Ryther, 1962). Main cultures were
inoculated with 3500 cells mL-1, sampled from a pre-culture grown in
dilute-batch mode (Langer et al., 2009). Final cell densities of the main
cultures were approximately 1×106 cells mL-1.
Experimental setup: the potential precursors of
CH4, 13C-labeled bicarbonate (13C-Bic) or a position-specific
13C-labeled methionine (13C-Met) were added to the flasks
containing either a culture of E. huxleyi or seawater
only.
To investigate algae-derived CH4 formation a closed-chamber system was
used. Hence, 2 L flasks (Schott, Germany) filled with 1800 mL sterile filtered
seawater and with 480 mL headspace volume were used in our investigations.
The flasks were sealed with lids (GL 45, PP, 2 port, Duran Group) equipped
with two three-way ports (Discofix®-3, B-Braun), where one
port was used for water and the other port (fitted with a sterile filter,
0.2 µm; PTFE, Sartorius) for gas sampling. The cells were grown on a
day–night cycle of 16 and 8 h at 20 ∘C and a light intensity of
≈ 450 µE over a 10-day period. The initial dissolved inorganic
carbon (DIC) of the culture medium was 2235 µmol L-1 (for
details on DIC measurements, see Langer et al., 2009).
The different treatments and the number of replicates are provided in Table 1 and Fig. 1.
To increase the detectability of CH4 formation and to exclude a
possible contamination with CH4 from the surrounding air,
13C-labeled bicarbonate (NaH13CO3, 99 % purity,
Sigma-Aldrich, Germany) was added to the cultures. Bicarbonate (Bic) was
used as a C source for biomass production. To gain a 13C enrichment of 1 % of the total inorganic C (CO2, HCO3-, and
CO32-), 22.35 µmol L-1 NaH13CO3 was added,
leading to a theoretical δ13C value of 882 ‰.
We used two different control treatments: (1) algae cultures without
13C-Bic and (2) seawater with 13C-Bic.
To test methionine (Met) as a precursor of algae-derived CH4, Met with only the sulfur-bound methyl group 13C labeled
(R-S-13CH3, 99 % enriched, 1 µmol L-1) was added
to the cultures. Met has previously been identified as a methyl-group donor
for CH4 biosynthesis in higher plants and fungi (Lenhart et al., 2012,
2015). Moreover, marine algae use Met to produce DMSP, DMS, and DMSO,
substances that can be released into seawater and are known to act as precursors
for abiotic CH4 production.
Sample collection and analysis
Samples were taken daily from day 4 until day 10 (see Table 1). Prior to day
4, algae biomass was too low to allow the measurement of changes in CH4
mixing ratio.
For gas chromatography (GC) and continuous-flow isotope ratio mass spectrometry (CF-IRMS) analysis samples of headspace (30 mL) were taken
from each flask. GC samples were measured within 24 h after sampling, while
GC-IRMS samples were stored in 12 mL exetainers until 13C-CH4
measurements were carried out.
After gas sampling, samples of medium (25 mL) from each flask were also
taken for cell density determination. These samples were supplemented with
0.15 mL Lugol solution (Utermöhl, 1958) and stored in 50 mL Falcon tubes
at 4 ∘C. In order to maintain atmospheric pressure within the
flask, the surrounding air was allowed to enter via an orifice fitted with a
sterile filter to avoid bacterial contamination. Variable amounts of water
and headspace volume as well as the inflow of surrounding air were all taken
into consideration when CH4 production rates were calculated.
Cell density was determined via a hemocytometer (Thoma-Kammer with 256
fields, 0.0025 mm2 × 0.1 mm; Laboroptik Ltd, UK).
Gas chromatography
Gas samples were analyzed for CH4 mixing ratio within 24 h on a gas
chromatograph (Shimadzu GC-14B, Kyoto, Japan) fitted with a flame ionization
detector (FID) operating at 230 ∘C with N2 as carrier gas
(25 mL min-1) (Kammann et al., 2009). The GC column (PorapakQ, Fa.
Millipore, Schwallbach, mesh 80/100) was 3.2 m long and 1/8 inch in
diameter. The length of the precolumn was 0.8 m. The GC gas flow scheme and
automated sampling was that of Mosier and Mack (1980) and Loftfield (1997),
and peak area integration was undertaken with the software PeakSimple,
version 2.66. The standard deviation (SD) of the mean of six atmospheric
air standard samples was below 0.2 % for CH4.
CF-IRMS for measurement of
δ13C values of CH4
Headspace gas from exetainers was transferred to an evacuated sample loop
(40 mL). Interfering compounds were separated by GC and CH4 trapped on
Hayesep D. The sample was then transferred to the IRMS system
(ThermoFinnigan Deltaplus XL, Thermo Finnigan, Bremen, Germany) via an
open split. The working reference gas was carbon dioxide of high purity
(carbon dioxide 4.5, Messer Griesheim, Frankfurt, Germany) with a known
δ13C value of -23.64 ‰ relative to Vienna
Pee Dee Belemnite (V-PDB). All δ13C values of CH4 were corrected using three CH4 working standards (isometric
instruments, Victoria, Canada) calibrated against IAEA and NIST reference
substances. The calibrated δ13C-CH4 values of the three
working standards were -23.9±0.2 ‰, -38.3±0.2 ‰, and -54.5±0.2 ‰.
Samples were routinely analyzed three times (n=3) and the average
standard deviations of the CF-IRMS measurements were in the range of 0.1 to
0.3 ‰.
All 13C / 12C-isotope ratios are expressed in the conventional
δ notation in per mil (‰) vs. V-PDB,
using the following equation (Eq. 1):
δ13C=((13C/12C)sample/(13C/12C)standard)-1.
To determine the δ13C signature of the CH4 source, the
Keeling-plot method was applied (Keeling, 1958).
Microbial investigations
DNA extraction and real-time PCR
Samples for DNA extraction were taken from the stem culture (RCC 1216)
during the stationary growth phase (2×106 cells mL-1).
After DNA extraction, real-time polymerase chain reaction (qPCR) was used to detect mcrA genes, which are
solely found in methanogenic archaea. As positive proof, aliquots of the
samples were supplemented with a defined cell density of
Methanothermobacter marburgensis (either 104 or 107 cells mL-1).
The DNA extraction was carried out according to (Bürgmann et al., 2001).
A total of 1 mL of the algae culture was transferred into a 2 mL vial containing
200 µL of zirconia–silica beads (Roth) and centrifuged for 20 min
(1.3×104 U min-1; 20 ∘C). Afterwards, 850 µL of the supernatant was replaced with extraction buffer
(Bürgmann et al., 2001) and beaten for 50 s (Retsch, type MM2). After
centrifugation the supernatant was transferred to another vial (2 mL,
Eppendorf, Germany), mixed with 850 µL
phenol–chloroform–isoamyl-alcohol solution (Roth) and again centrifuged for
5 min (1.3×104 U min-1; 20 ∘C). The water
phase was supplemented with 800 µL phenol, mixed, and centrifuged
again. Afterwards, the water phase was transferred in a new vial, mixed with
800 µL precipitating buffer (polyethylene glycol, PEG) and centrifuged for 60 min (1.3×104 U min-1; 20 ∘C). The pellet was washed
with 800 µL ethanol (75 %; -20 ∘C; centrifuged for 10 min
at 1.3×104 U min-1, 20 ∘C) and air-dried in
the laboratory. For elution and storage of the pellet, we used 20 µL
nuclease-free water.
Real-time PCR was carried out according to Kampmann et al. (2012) with a
Rotor-Gene 3000 (Corbett Research, Australia) by using
ABsolute™ QPCR SYBR® Green Mix (ABgene). For the
detection of mcrA genes, we used a primer
(ML forward:5′ GGTGGTGTMGGATTCACACARTAYGCWACAGC-3′; ML reverse:
5′ AACTAYCCWAACTAYGCAATGAA-3′), which encodes the α-subunit of the
methyl-CoM reductase, which solely occurs in methanogenic archaea (Luton et
al., 2002).
The real-time PCR reference standards were produced according to Kampmann et al. (2012). By using the standard solution (5.5×107 DNA
copies µL-1), dilution with nuclease-free water was accomplished
down to 5.5×101 copies per µL-1. All standards
and regular samples taken from the flasks were analyzed with four
repetitions.
Quality assurance of the real-time PCR product was achieved by melt curve
analysis and gel electrophoresis using the fluorescent stain GelRedTM
(Biotium).
Cultivation approach
In addition to real-time PCR, a cultivation and enrichment procedure (Kampmann
et al., 2012) was conducted to screen for methanogenic archaea in algae
cultures. The enrichment medium (Widdel and Bak, 1992) was modified for
marine conditions by adding 320 mmol L-1 NaCl, 16 mmol L-1
MgCl2, and 1 mmol L-1 NaHCO3. At day 10, an aliquot (5 mL) of
each cultivation flask was transferred into injection flasks (Ochs,
Bovenden-Lenglern, Germany) with the enrichment medium (50 mL) and acetate
(10 mM), methanol (5 mM) was added, and in the gas phase H2 and CO2
(90:10) were provided as substrates. Incubation was carried out over a period
of 6 weeks at 20 ∘C in the dark.
CH4 mass
The mass of CH4 (mCH4) per flask was calculated via the ideal
gas law from the corrected CH4 mixing ratio (ppmv), where the changing
volume of water and headspace and the inflow of surrounding air were all
considered, according to Eq. (3):
mCH4=pR×T×cCH4×V×MCH4,
where p is pressure, T is temperature, R is ideal gas constant, V is
volume, and MCH4 is mol. weight of CH4. The solubility of
CH4 in the water phase was calculated according to Wiesenburg and
Guinasso (Wiesenburg and Guinasso Jr., 1979) based on the headspace-CH4
mixing ratio, temperature and salinity of the water phase.
Calculation of CH4 production
The low CH4 mixing ratios produced by E. huxleyi during the exponential growth
phase precluded the determination of CH4 production during this period.
Therefore, we calculated production from day 7 to day 10, a period
representing the transition from exponential to stationary phase. This
growth phase features changing growth rates and cellular CH4 quotas,
rendering the dilute-batch method of calculating production inapplicable
(Langer et al., 2013). We followed the recommendation of Langer et al. (2013)
and calculated incremental (daily) CH4 production:
Pinc=qinc×μinc,
where Pinc is incremental CH4 production (ng CH4 cell-1
day-1), qinc is incremental cellular CH4 quota (ng CH4
cell-1), and μinc is incremental growth rate
(day-1).
Incremental growth rate was calculated according to
μinc=LN(t1)-LN(t0),
where t1 is cell density on the day qinc was determined and t0 is cell density on the previous day. We present average Pinc (SD).
In order to compare CH4 production to literature data it was necessary
to normalize to cellular particulate organic carbon (POC) quota as opposed
to cell. The POC-normalized CH4 production is termed “methane emission
rate” in the following. Since it was not possible to measure cellular POC
quota on a daily basis, we used a literature value determined for the same
strain under similar culture conditions, i.e., 10.67 pg POC cell-1
(Langer et al., 2009). We are aware of the fact that the cellular POC quota is
likely to change alongside other element quotas when approaching the stationary
phase, but this change is well below an order of magnitude (Langer et al., 2013). For our purpose this method is therefore sufficiently accurate to
determine POC-normalized CH4 production.
Statistics
To test for significant differences in cell density, CH4 mixing ratio,
and CH4 content between the treatments, two-way analysis of variance (ANOVA) (considering
repeated measurements) and a post hoc test (Fisher least significant difference (LSD) test; alpha 5 %)
were used.
Results
Algae growth
Cell density and growth of the cultures are presented in Fig. 2a, b over
the whole incubation period for all treatments. The initial cell density at
time 0 (t0) was 3.5×103 cells mL-1 in all flasks.
At day 10 cell density reached its maximum value with 1.37×106 cells mL-1 (algae), 0.82×106 cells mL-1
(“algae + 13C-Bic”), and 1.24×106 cells mL-1 (“algae
+ 13C-Met”). The exponential growth rates (μ) were 0.85±0.2 d-1 for algae + 13C-Met, 0.98±0.1 d-1 for
algae + 13C-Bic, and 1.06±0.02 d-1 for the control
“algae” (n.s., p=0.286). Significant differences in cell density between
the treatments only occurred at days 9 and 10, where the cell density of the
control algae was higher than in the treatments where 13C-Bic or
13C-Met was added.
Methane mixing ratio
Initial headspace-CH4 mixing ratios measured at day 4 were in the range
of 1899 to 1913 ppbv for all treatments including the controls without
algae. From day 4 to day 7 headspace-CH4 mixing ratios slightly
increased in all flasks. Therefore, no significant differences in the
CH4 mixing ratios occurred between the treatments. After day 8 CH4
mixing ratios in the flasks containing algae were significantly higher
compared to the controls without algae (Fig. 2c, d). The highest CH4
mixing ratios at day 10 corresponded to 2102 ± 62 ppbv (algae
+13C-Met), 2138 ± 42 ppbv (algae +13C-Bic), and 2119±25 ppbv (algae).
Culture cell density when algae grown in seawater (n=2)
supplemented with (a) Bic or (b) Met (n=3) and headspace-CH4 mixing
ratio for cultures supplemented with (c) Bic or (d) Met. δ13CH4 values after addition of (e) 13C-Bic and (f)
13C-Met (n=3; error bars mark the standard deviation). Stars mark
the significance between algae + 13C-Bic and “seawater +
13C-Bic” or between algae +13C-Met and “seawater +
13C-Met”, with *p≤0.05; **p≤0.01; ***p≤0.001.
Hence, from day 4 to day 10 the CH4 mixing ratios increased by about
192 ppbv (algae + 13C-Met), 49 ppbv (seawater + 13C-Met),
235 ppbv (algae + 13C-Bic), and 67 ppbv (seawater + 13C-Bic).
Stable carbon isotope values of methane
The δ13C signature of headspace CH4 (δ13CH4 value) is presented in Fig. 2e and f. The addition of
13C-Bic did not affect CH4 production of algae, but the δ13CH4 value was clearly different from that of the
control algae. The initial value of -47.9 ± 0.2 ‰ increased to 44 ± 13 ‰, whereas in the controls “seawater + 13C-Bic” and algae no
change in the δ13CH4 value was observed.
The addition of 13C-Met did not affect algal CH4 formation, but it
increased the δ13CH4 signature from -46.35 +0.84 ‰
to 59.1 ± 25.3 ‰ (day 8). In
the treatment “13C-Met”, where only isotopically labeled Met was
added to sterile filtered seawater, a small increase from -48.0 ± 0.3
to -38.1 ± 2.3 ‰ (at day 10) was observed.
Based on the initial amount of 13C-Bic and the total amount of
13CH4 at the end of the incubation period, 88.3 ± 17.2 pmol of 22.4 µmol 13C-Bic were converted to 13CH4. For
Met, this was 78.5 ± 18.6 pmol of the initial 1.8 µmol
13C-Met.
The Keeling plots to determine the 13C values of the CH4 source
are presented in Fig. 3. For the bicarbonate treatment (algae + 13C-Bic), the mean δ13CH4 value of the CH4 source was 811.9 ± 89.9 ‰, which is close to
the calculated δ13C value of 881.5 ‰ after
the addition of NaH13CO3.
For the treatment algae + 13C-Met, we applied the Keeling-plot
method only for the period from day 5 to day 7, as the increase in the
δ13C values were not linear after day 7. For this treatment,
the δ13C values of the CH4 source range between 967 and
2979 ‰.
Keeling plots for the treatment (a) algae + 13C-Bic and
(b) algae + 13C-Met, where f(0) refers to the 13C value
of the CH4 source.
Correlation between cell density per flask and CH4 content (sum
of headspace and water phase) for the coccolithophore E. huxleyi
(a) in seawater only (n=2; light green) and supplemented with
13C-labeled bicarbonate (Bic; dark green) or (b) methionine
(Met) (n=3); error bars mark the standard deviation; d is day of
incubation.
The correlation between the growth of the algae cultures and the total
amount of CH4 in the flasks (headspace + water phase) is presented in
Fig. 4. For the treatment algae + 13C-Bic (Fig. 4a), there is an
exponential correlation between cell density and CH4 content (r2=0.994), whereas for the treatment algae + 13C-Met (Fig. 4b), a
linear correlation was observed (r2=0.995).
The daily CH4 content in the flasks for days 8, 9, and 10 is shown in
Fig. 5. For all flasks the CH4 content exceeded the CH4 content
of the respective control, with a continuous increase in the CH4
content in the flasks containing algae. At day 10, the difference between
algae + 13C-Bic and seawater + 13C-Bic and between
algae + 13C-Met and “seawater + 13C-Met” was 65 ± 16 and 54 ± 22 ng, respectively.
The CH4 production of algae presented in Table 2 shows no major
differences between the treatments. Furthermore, for all treatments, the
daily CH4 production rates did not change over time (Fig. 6).
Microbial investigations
Via real-time PCR no mcrA genes could be detected in the flasks containing
the CH4-producing algae cultures, whereas in the positive control in which
the algae culture was supplemented with 104 and 107 cells mL of the methanogenic archaea Methanothermobacter marburgensis,
9.4×104 and 4.6×106 mcrA-gene copies mL-1 have been detected, respectively.
With the cultivation approach, where an aliquot of each flask was taken at
day 10 and transferred to the media for the enrichment of methanogenic archaea,
no CH4 production was observed after the 6-week incubation period. In
the case of a successful enrichment of methanogenic archaea, the CH4-mixing
ratio in the headspace would increase over time.
Discussion
Our results of the CH4 mixing ratio and stable isotope measurements
provide unambiguous evidence that E. huxleyi produces CH4. In the following we
will discuss the relationship between CH4 production and the growth of the
algae, stable isotope measurements, potential precursor compounds, and the
exclusion of methanogenic archaea. Finally, we will discuss the implications
of our results for the methane paradox in oxic waters.
Mean daily CH4 production rates of E. huxleyi (*n=2; **n=3)
determined between days 7 and 10; ag: attogramm (10-18).
Treatment
CH4
CH4
(ag cell-1 d-1)
(µg g-1 POC d-1)
E. huxleyi+ 13C-Bic**
6.8 ± 4.1
0.63 ± 0.39
E. huxleyi+ 13C-Met**
9.3 ± 2.6
0.88 ± 0.24
E. huxleyi*
6.1 ± 3.7
0.57 ± 0.35
Mean CH4 content (sum of headspace and water phase) in the
flasks of E. huxleyi supplemented with either bicarbonate or
methionine (n=3) or the respective control without algae (n=2)
measured at days 8, 9 and 10; error bars show the standard deviation.
Growth and CH4 production
Over the course of the exponential growth phase headspace-CH4 mixing
ratios in treatments containing E. huxleyi were not measurably different from the
control treatments. Therefore, it was not possible to determine CH4
production in the exponential growth phase. However, we conclude that E. huxleyi
produces CH4 throughout all growth phases as will be detailed in the
following. In the transitionary growth phase leading up to the stationary
phase, we calculated incremental CH4 production (daily). The transitionary
phase features a declining growth rate and often increasing cellular carbon
quotas (Langer et al., 2013). Cellular CH4 quotas also increased (data not shown). On the other hand, CH4 production remained constant
within the measurements of error, displaying a slight downward trend when
approaching stationary phase (Fig. 6). Therefore, we conclude that CH4
production is not a feature of senescent cells only but is probably operational in all growth phases. This is interesting in the context of the
ecology and biogeochemistry of E. huxleyi. Contrary to the traditional assumption that
E. huxleyi production in the field is dominated by late summer bloom events, it was
recently shown that non-bloom production in spring contributes significantly
to yearly average production and therefore bloom events are not
exceptionally important in biogeochemical terms (Schiebel et al., 2011).
Since senescent cells in field samples are mainly a feature of late bloom
stages, the exclusive production of CH4 by such cells would confine any
contribution of E. huxleyi to the oceanic CH4 budget to a relatively short, and
biogeochemically less important, period. However, from results found in this
study we would propose that E. huxleyi produces CH4 during all growth phases as
part of its normal metabolism. If our findings are confirmed and supported
by other research groups, this has considerable implications as it would
render this species a prolific aerobic producer of CH4, on a par with,
for example, terrestrial plants (Bruhn et al., 2012).
Daily CH4 production of E. huxleyi for days 7 to 10
(a, c, e) on a per-cell basis and (b, d, f) relative to
particulate organic carbon (POC) separately for the treatments (a, b) E. huxleyi +
13C-Bic (n=3), (c, d)
E. huxleyi + 13C-Met (n=3), and (e, f) E. huxleyi (n=2). Values are presented as means with the standard deviation.
Methane emission rates
To calculate CH4 emission rates of E. huxleyi, we normalized CH4 production
to cellular POC content (see Material and
Methods). The CH4 emissions were 0.7 µg POC g-1 d-1,
or 30 ng g-1 POC h-1 (mean for all treatments, n=8).
In this study the main aim was (as a proof of principle) to unambiguously
provide evidence that E. huxleyi are able to produce methane under aerobic conditions
and without the help of microorganisms.
However, we suggest that CH4 emission rates of E. huxleyi algae are different
under changing environmental conditions, e.g., temperature, light intensity,
or nutrient supply. The effect of changing environmental parameters should
be the focus of future investigations.
For comparison CH4 emission rates presented so far for terrestrial
plants range from 0.3 to 370 ng g-1 DW (dry weight) h-1 (Keppler
et al., 2006; Wishkerman et al., 2011; Lenhart et al., 2015; Brüggemann
et al., 2009).
Inorganic and organic precursors of CH4
Based on the addition of bicarbonate (13C-Bic, 1 % enrichment),
which is the principal carbon source for the growth of algae, and the
measurements of δ13CH4 values it was possible to clearly
identify bicarbonate as the principal carbon precursor of CH4 in E. huxleyi.
In the flasks where algae were supplemented with 13C-Bic, a significant
increase in δ13CH4 values occurred over the incubation
period, which shows that algae use bicarbonate as precursor carbon (C) for
CH4 production. As expected, in the controls flasks algae where no
13C-Bic was added and the control seawater + 13C-Bic
without algae, no change in δ13CH4 values was observed.
The initial δ13C value of the bicarbonate in the treatment
algae + 13C-Bic (+882 ‰) is within the
range of the source δ13CH4 values obtained via the
Keeling-plot method (+812 ± 90 ‰). Even though
there might be kinetic isotope fractionations involved in each of the
several steps during organic matter formation, these data clearly indicate
that bicarbonate is the principle inorganic carbon precursor of CH4
produced in algae.
Bicarbonate is taken up by the algae via autotrophic C fixation (Burns and
Beardall, 1987) and might therefore – during several steps of metabolism, i.e., the formation of organic compounds – lead to the formation of CH4.
It will probably be used as an unspecific C source in many different
metabolic pathways, e.g., the synthesis of lignin, pectin, and cellulose
(Kanehisa et al., 2014) – components already known as CH4 precursors
from terrestrial plants, where CH4 can be
produced via methyl group cleavage (Keppler et al., 2008; Bruhn et al., 2009; Vigano et al., 2009).
However, lignin and pectin are not commonly found in marine algae such as
E. huxleyi. For these organisms, sulfur-bonded methyl groups such as thioethers,
sulfoxides, and sulfonium salts (methionine, S-adenosylmethionine (SAM),
adenosylmethionine DMSP, DMSO, DMS) are of much more interest. For our experiments, we used 13C
positionally labeled Met where only the sulfur-bond methyl group
(–S-CH3) was 99 % enriched in 13C. Our choice of this compound
was partly due to its commercial availability but more importantly because
it is known to be involved in a number of metabolic pathways and
transmethylation reactions (Stefels, 2000; Bruhn et al., 2012).
In contrast to the ubiquitous C-source bicarbonate – which can also be used
to build Met in algae (Stefels, 2000) – Met is incorporated in specific
metabolic pathways. Algae use part of the Met for protein synthesis; in E. huxleyi it
is also involved in the synthesis of DMSP, a main precursor of DMS and DMSO.
The clear increase in δ13CH4 values of headspace-CH4
in the treatment algae + 13C-Met (Fig. 2e, f) shows that the
methyl thiol group of Met is a direct CH4 precursor. The Keeling-plot
results (Fig. 3) show higher variability for Met than for Bic. However, Met
is almost certainly not the only precursor of CH4, as the
headspace-CH4 mixing ratios increased (Fig. 2d), while the 13C
values of headspace-CH4 showed a saturation curve (Fig. 2f). This
indicates either a shift from Met to other CH4 precursors or to the
use of newly synthesized, non-labeled Met. Based on the initial amount and
the total amount of 13CH4 formed at the end of the
incubation, only a small fraction (79 pmol, i.e., 4.0 ‰)
of the initial added 13C-Met (1.8 µmol) was converted to
13CH4. The formation of CH4 from 13C-Met explains
roughly about 3 % of the total amount of CH4 formed throughout the
incubation period. Possibly, the formation of potential precursors of
CH4 may change under various climatic conditions, leading to varying
CH4 production rates in different pathways.
This observation is in line with the findings of Lenhart and colleagues, who
demonstrated that the sulfur-bound methyl group of Met was a precursor of CH4 in plants (Lenhart et al., 2015) and fungi (Lenhart et al., 2012).
The linear increase in headspace-CH4 mixing ratio (Fig. 2d) together
with the nonlinear increase in δ13CH4 signature (Fig. 1f)
indicates that the pool of 13C-Met was either exhausted or was diluted
by newly synthesized, non-13C-enriched Met.
In addition, we also found an indication for a chemical CH4 formation
pathway in the seawater with Met as methyl-group donor as a small increase
in 13CH4 values in the control treatment seawater + 13C-Met was observed (Fig. 2f). This CH4 formation pathway is
approximately 10-fold lower when compared to the treatment algae + 13C-Met and is only observed in the isotopic experiment but not when
only the CH4 mixing ratio is considered (Fig. 2d). However, this
observation is in line with some previous findings (Althoff et al., 2010, 2014), who showed that the abiotic formation of CH4 due to
the degradation of methionine or ascorbic acid by light or oxidants such as
iron minerals is possible. In the case of methionine, it was shown that the
sulfur-bound methyl group of Met was the carbon precursor of CH4
(Althoff et al., 2014).
Potential implications for the occurrence of CH4 in oxic marine
waters
Several hypotheses with regard to the occurrence of the seasonal and spatial
CH4 oversaturation in oxic surface waters (Bange et al., 1994; Forster
et al., 2009; Owens et al., 1991) have been postulated. They include
CH4 formation from methanogenic archaea in anoxic microsites (Karl and
Tilbrook, 1994) or CH4 formation via the C-P-lyase pathway from
methylphosphonate (Karl et al., 2008).
In the ocean, both CH4 production by methanogens and consumption via
methanotrophic bacteria occur simultaneously. Therefore, CH4 production
can exceed estimated CH4 production rates when based solely on CH4
mixing ratio measurements (Reeburgh, 2007). To provide a noteworthy
contribution to oceanic CH4 production, precursors must either be
available in high abundance or be continually synthesized. Algae-derived
methylated sulfur compounds such as Met, DMSP, DMS, and DMSO are ubiquitous
in the ocean but show a high spatial and temporal variability with high
mixing ratios in algal blooms. Therefore, they are potential compounds that
might be involved in CH4 formation in the oceans (Keppler et al., 2009;
Althoff et al., 2014). The involvement of methyl moieties from methylated
sulfur compounds in CH4 biosynthesis might therefore play an important
role in pelagic CH4 production. Mixing ratios of DMS and DMSP in seawater during algal blooms were reported in the range of 0.82 to 8.3 nmol L-1 and 1.25 to 368 nmol-1, respectively (Matrai and Keller,
1993).
The CH4 emission rates of E. huxleyi may also occur by a second formation
pathway, where DMSP is first converted to DMS and subsequently oxidized to
DMSO (Bentley and Chasteen, 2004).
However, several studies have afforded evidence for a CH4 formation
pathway via methyl radicals (Althoff et al., 2014; Eberhardt and Colina,
1988; Herscu-Kluska et al., 2008), leading to the hypothesis that
algae-derived DMSO can also act as a precursor of CH4 in oxic seawater
(Althoff et al., 2014). A correlation between Met and DMSP synthesis was
provided by Gröne and Kirst (1992), who showed that the supplementation of
Tetraselmis subcordiformis with 100 µg L-1 Met yielded a 2.6-fold increase in DMSP. For
E. huxleyi, DMSO mixing ratios in the stationary growth phase can reach 0.1 pg per
cell (Simo et al., 1998). Assuming that a similar DMSO mixing ratio were to
be found in our study, this would mean that in every 4×103
DMSO molecules per day must be transferred to CH4 to explain the
observed increase in CH4. Moreover, a positive correlation was observed
between chlorophyll a and CH4, as well as between DMSP or DMSO and
CH4 (Zindler et al., 2013).