Riverbed methanotrophy sustained by high carbon conversion efficiency
Mark Trimmer*1, Felicity C. Shelley1, Kevin J. Purdy2, Susanna T. Maanoja1, PanagiotaMyrsini Chronopoulou1 & Jonathan Grey1.
5
1
School of Biological and Chemical Sciences, Queen Mary University of London, London, E1
4NS, UK
2
School of Life Sciences, University of Warwick, Coventry, CV4 7AL, UK
*Correspondence: M. Trimmer, E-mail m.trimmer@qmul.ac.uk
10
Supplementary Information
Supplementary Methods.
Organic carbon by wet oxidation to CO2. An adaptation of a standard wet-oxidation
method (no. 505C; (APHA, 1995)) was used to recover and quantify the yield of bulk
15
13
C-
organic carbon from the harvested gravels. A subsample (~150 mg) of freeze-dried gravels
was transferred into a serum bottle (20 mL) which was then acidified to remove carbonates
(4 mL H3PO4 6% v/v) and left overnight. After, an additional aliquot of concentrated acid (300
µL H3PO4 concentrated) was added to check for any residual effervescence. Finally,
potassium persulfate (5 mL of 0.15 M) was added to each serum bottle, which were then
20
purged for 10 minutes with N2 gas and autoclaved (121 C, 3 h). Once cool, the headspace
was analysed for CO2, first by GC/FID (after catalytic reduction to CH4 over hot nickel), to
quantify the TOC, and then by CF/IRMS for
13
C-CO2 to quantify the labelled fraction.
Procedural blanks (4 mL acid) were prepared and analysed in every sample batch.
The performance of the wet oxidation assay was verified by standard addition. Three
25
sets of vials were prepared: one set contained only the organic standard potassium biphthalate (0–2 mg C), while the other two contained ~150 mg gravel, plus the same
respective amount of standard. Varying amounts of the standard were pipetted into the vials
and the volume made up to 3.7 mL with ultra-pure water. The oxidation of the potassium biphthalate to CO2 was perfectly linear (r2 = 1) over a wide range (Supplementary Fig. 2) and
1
30
its recovery from spiked gravels was also complete (102% ± 4 s.e., n = 10; One sample ttest, t = 0.59, d.f. = 9, P > 0.05). After the success of the performance test the wet-oxidation
method was used to analyse the experimental gravels, with the total amount of organic
carbon on the gravels being calculated against a calibration curve (0.2–3 mg carbon)
prepared from the organic standard.
35
Initially the efficiency of different acids and acidification time on removal of
carbonates from the gravel was tested by acidifying samples separately with either 6 %
H3PO4 or 6 % H2SO4 (w/w) over 24h. Two sets of samples were left to stand for 24h and then
purged for 10 min with N2. There were also two samples that were not acidified at all. After
acidification the samples were treated as described earlier. Digestion with 6 % H2SO4 for 24
40
h failed to fully dissolve all of the inorganic carbon, resulting in distinctly less negative δ13C
values for the carbon. Digestion with 6 % H3PO4 for 24 h removed all of the inorganic carbon
and was used throughout.
Quantifying the yield of
45
gravels
that
would
13
C-lipids. A crude lipid extract was performed on the freeze-dried
have
included
other
hydrophobic
compounds,
such
as
lipopolysaccharides (Gurr and Harwood, 1991; Parrish et al., 2000). Glassware was acid
washed and flushed (×3) with HPLC grade methanol and chloroform. Solvents and samples
were kept on ice while being continuously flushed with N2 (Gurr and Harwood, 1991). An
aliquot (4.5 mL) of extracting solution (2 mL CHCl3, 1 mL MeOH, 1 mL CHCl3/MeOH mixture
50
[2:1] and 0.5 mL UHP water) was added to 1 g of gravel, sonicated (Ultrasonic Cleaner 45
kHz, VWR USC200T, England), vortexed (1 min) and centrifuged (2 min at 715 g) to
separate the organic and aqueous layers. The organic layer was recovered and the
procedure repeated twice more, the extracts pooled and concentrated (to 250 µl) under N2.
Extracts were kept at -18 C until later combustion and elemental analysis coupled to
55
CF/IRMS (Trimmer et al., 2009). Analytical controls of just UHP water were added to every
batch.
2
Nucleic acid extraction, PCR amplification and 454 sequencing. Nucleic acids were
extracted from 0.8 g of fresh gravel from all 8 rivers. In order to assess intra-river variation in
60
the methanotroph community additional gravel samples were collected from six points along
a 250 m reach (~50 m apart) of the River Lambourn which were first extracted individually,
then the gravel samples were homogenized and DNA extracted from this mixed sample. The
gravel was added to 2 ml sterile tubes with zirconia/silica beads, 0.1mm (supplied by
Stratech Scientific Limited) and suspended in 500 µl of 0.1 M potassium phosphate buffer
65
(pH 8.0), and 0.5 ml of phenol-chloroform-isoamyl alcohol (25:24:1 v/v). Cells were lysed by
bead beating using a TissueLyser LT, Qiagen, at 50 Hz for 1 min. The rest of the extraction
process was as described by (McKew et al., 2007).
Amplification of the pmoA gene was performed in a Bio-Rad MJ Research PTC-220
DYAD Thermal Cycler using the primers A189/A682 (Holmes et al., 1995). DNA was diluted
70
two times and 1 µl of this dilution was used as template in triplicate PCR reactions. The 50 μl
reaction mixture contained 0.4 mM of each primer, 200 μM deoxynucleoside triphosphates,
20 ng μl-1 BSA, 1.25 U of Taq DNA polymerase and 5 ml of the reaction buffer supplied with
the enzyme (New England Biolabs). PCR conditions were as follows: 94°C for 5 min, 35
cycles of 94°C for 30 s, 57°C for 1.5 min and 72°C for 30 s, with a final elongation of 72°C for
75
10 min. PCR triplicate products were pooled and purified using the QIAquick PCR
Purification Kit (Qiagen) according to the manufacturer’s protocol. The purified products were
sent for 454 pyrosequencing of the pmoA gene, using the Roche 454 FLX/FLX+ platform at
the Research and Testing Laboratory (http://www.researchandtesting.com/index.php). 454
libraries were constructed using the A189 as forward primer in conjunction with the mb661
80
primer (internal of A682) (Costello and Lidstrom, 1999).
Processing of 454 sequences and phylogenetic analysis. 454 reads were analyzed using
the QIIME pipeline and its associated modules (Caporaso et al., 2010). Initially, all
sequences were checked for the presence of correct pyrosequencing adaptors, 8 bp
85
barcodes (unique for each sample) and pmoA-specific primers, and those containing errors
in these regions were removed. Sequences <200 bp, those with low quality scores (< 25),
3
and sequences containing homopolymers (>6 bp) were also removed. All 454 reads that
passed the above mentioned quality controls were clustered into operational taxonomic units
(OTUs here referred to as CS i.e. Chalk Stream taxonomic unit) at the 90% similarity level
90
(given the 3.5-times-higher nucleotide substitution rate of the pmoA compared to the 16S
rRNA gene (Pester et al., 2004), using the USEARCH algorithm (Edgar, 2010). The
associated de novo chimera checker UCHIME was used to detect and remove all chimeras
(Edgar et al., 2011). Representative sequences from each CS were assigned taxonomy
using BLAST (Altschul et al., 1990) against the National Center for Biotechnology Information
95
(NCBI) database. Principal Coordinate Analysis (PCoA) was performed using the Unifrac
distance metric (Lozupone et al., 2011), after random subsampling so that all samples
contained the same number of sequences (i.e. normalizing to the 930 sequences collected
from the Ver sample). All representative sequences and their closest relatives from BLAST
were aligned using the muscle algorithm (Edgar, 2004) and edited in the Bioedit program. A
100
phylogenetic tree was constructed using the FastTree method (Price et al., 2010) and edited
in Dendroscope (Huson et al., 2007).
4
Supplementary Results
105
Supplementary Table 1. Summary budget for the recovery of oxidised 13C-CH4 as either dissolved inorganic carbon (∑DI13C= CO2, HCO3- and
CO32-) or total organic carbon (lipid plus bulk organic fraction, see Materials and Methods) in the first eight repeat batch incubations. Note, there
was no significant difference between the recovery in either fraction and both were indistinguishable from 50% and, overall, the total recovery of
13
C-CH4 was ~ 100%.
Batch
Time (hours)
13
CH4 oxidised (nmol g-1)
Cumulative 13CH4 oxidised (nmol g-1)
TO13C (nmol g-1)
∑DI13C (nmol g-1)
1
51
25
25
15
12
2
90
31
56
34
13
3
119
20
75
49
9
4
166
47
122
62
33
5
212
53
176
86
32
6
289
127
302
57
63
7
344
88
390
274
49
8
414
87
477
183
49
Recovered as TOC (%)
Recovered as DIC (%)
Total (%)
61
46
107
61
42
103
65
46
111
51
69
120
49
60
109
19
49
68
70
56
126
38
57
95
mean
52 ± 6
53 ± 3
105 ± 6
5
110
Supplementary Table 2. The individual estimates of the ratio of 13C-DIC produced to 13CCH4 consumed and methane oxidation activity for each of the eight chalk stream gravel
samples (see main text Fig. 1). ANCOVA revealed strong differences in oxidation activity
between the different samples (Methane oxidation x river, P<0.001, n=8) but only marginal
differences in the ratio of 13C-DIC produced to 13C-CH4 consumed (Ratio x river, P=0.047,
n=8). Significant differences between the samples are indicated by different letters (e.g. a≠b)
and *L is for the pooled sample in the Lambourn (see main text Results and Fig. 5).
Chalk stream Ratio of 13C-DIC Methane oxidation
sampled
to 13C-CH4
(nmol CH4 g-1 h-1)
Chess
0.40b
4.4d
Cray
0.46b
4.6d
Darent
0.73a
2.4b
Gade
0.40b
8.4a
Lambourn (*L)
0.49b
1.6c
Mimram
0.68b
2.8b
b
Misbourne
0.57
1.6c
Ver
0.75a
1.4c
115
120
125
130
135
Supplementary Table 3. Identity and relative abundance (%) of the most abundant pmoA
OTUs (~97% of all analyzed sequences; 24 out of 70 OTUs) in the eight independent river
samples. The most abundant OTU in a sample is dark shaded and in bold; pale shaded
OTUs represent >10% of the sequences in a sample; -- no sequences from the OTU in a
sample. Simpson’s diversity index was calculated based on 1-D, with D = Σ(n/N)2, where
n=the number of sequences in each CS and N=the number of sequences in all 70 detected
OTUs. The closer the index is to 1, the higher the diversity in a sample.
6
Type I Methanotrophs
8. Darent
7. Cray
6. Mimram
5. Gade
% ID
4. Ver
(Accession Number)
3. Chess
Closest relative
2. Misbourne
(CS Cluster
on tree)
1. Lambourn
OTU
Percentage of sequences in each sample
Clone OK-55 (AB845086)
93
19.6
0.6
2.3
5.4
14.1
80.6
6.8
21.8
CS3
M’coccus capsulatus Bath (AE017282)
99
--
2.0
4.1
11.8
0.2
0.4
1.5
1.9
CS4
Clone LP-II140 (HQ383722)
92
--
--
--
--
12.9
--
--
--
CS5
Clone P222-11 (HQ738562)
99
--
--
--
0.1
--
--
--
12.4
CS6 (5)
Clone PR-II579 (HQ383812)
99
2.5
2.8
--
6.2
2.8
--
0.3
0.3
CS7 (9)
Clone SS-80 (AB845146)
89
0.2
--
0.02
--
--
10.2
--
1.4
CS8 (5)
Clone LR-II489 (HQ383753)
97
4.2
2.5
--
0.1
1.1
--
--
0.2
CS9 (3)
Clone LL_HA_B07 (HE617690)
95
9.7
--
--
--
--
--
--
--
CS10 (1)
Clone 86 (AY355390)
94
1.2
6.0
--
--
--
--
--
--
Clone SS-59 (AB845133)
92
12.7
--
0.02
--
--
--
--
0.1
CS12 (5)
Clone oytpmoaA32 (AB722429)
94
--
5.7
--
2.2
--
--
--
--
CS13 (9)
uncultured M’bacter (EU124863)
92
0.8
--
--
9.0
--
0.6
--
0.3
CS14 (9)
Clone BQ661-10 (AB844965)
90
3.8
--
--
--
0.8
2.9
--
1.1
CS15
Clone B109 (AY488068)
98
--
0.04
1.4
--
--
--
--
--
CS16
Clone oytpmoAA10 (AB722413)
98
--
--
--
10.5
--
--
--
--
CS17 (2)
Clone LR-II472 (HQ383749)
96
0.9
--
1.4
--
--
--
--
--
CS18 (1)
Clone LP-II101 (HQ383718)
99
--
--
--
--
4.8
--
--
--
CS20 (9)
Clone LR-II494 (HQ383755)
94
2.9
--
--
--
--
0.1
--
--
CS30 (9)
Clone HS-36 (AB845028)
92
1.2
--
0.1
0.1
0.8
3.1
0.3
5.2
CS60 (9)
Clone HT2-45 (AB844925)
90
0.2
--
0.1
--
1.6
0.3
0.6
2.3
19.6
9.4
45.5
39.0
98.3
9.5
46.9
CS2 (9)
CS11
Type I total (%) 60.0
CS1
M’cystis sp. 5FB1 (AJ868406)
99
35.3
72.1
85.5
38.4
54.4
0.2
84.4
41.4
CS19
M’cystis sp. LW5 (AF150791)
92
--
--
--
--
--
--
--
2.9
CS26
M’cystis sp. SC2 (HE955767)
99
0.8
1.2
1.7
0.2
1.3
--
4.5
1.2
CS56
M’cystis sp. SC2 (HE955767)
99
--
1.5
3.2
2.6
1.3
--
1.3
0.3
Type II total (%) 36.0
74.8
90.4
41.2
57.0
0.2
90.2
45.8
Overall Total
96.0
94.5
99.8
86.7
96.1
98.5
99.6
92.7
Simpson’s Diversity (1-D)
0.8
0.5
0.3
0.8
0.7
0.3
0.3
0.8
140
7
Supplementary Figure 1. Water quality data for the eight chalk streams. Data come from
the Environment Agency (UK, public access records) monitoring station close to our sample
sites on each river, which were typically sampled once per month and, here, cover the period
145
2007 to 2014. Each box is defined by the 25th and 75th percentiles (spread) with the dotted
line measuring 1.5 times the spread. The thick horizontal line in each box gives the median
value for each river and the horizontal line gives the median value for the overall dataset. We
selected these parameters as they were the best represented in all of the rivers for the given
period and they give a good indication of overall water quality. Both nitrite and ammonia were
150
two orders of magnitude lower than nitrate. The Cray and Darent are notable exceptions with
long-term medium nitrate concentrations lower than all the other rivers. The Chess had
significantly higher SRP and pH, whereas as all rivers had long-term oxygen saturations of >
91%
155
Supplementary Figure 2. Measuring organic carbon in the gravel biofilm by wet oxidation to
CO2. (a) Response of the GC/FID to increasing amounts of organic carbon standard
additions following wet oxidation to CO2 and catalytic reduction to CH4. (b) Recovery of
organic carbon standard additions from gravel. Duplicate determinations in each case.
160
Supplementary Figure 3. Rates of methane oxidation by gravels remaining constant as a
function of the degree of 13C-CH4 (atom %) labelling. Mean ± s.e., n=5.
8
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