1
Supplemental Material
2
1. Materials and Methods
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
Cultivation of Thiocapsa KS1
Thiocapsa sp. strain KS1 was isolated from sewage sludge of the municipal
sewage treatment plant at Konstanz, Germany, and has been deposited with the Japan
Collection of Microorganisms under accession number JCM 15485. Cultures were grown
as previously described (Grein et al., 2013; Schott et al., 2010)except that cultures were
incubated at 28°C and re-fed with nitrite at 2 mM increments. Growth was measured via
turbidity at 660 nm with a Camspec M107 spectrophotometer (Camspec, Camspec
Ltd.11, High Street, Sawston, Cambridge, UK).
Preparation of cell-free extracts
Cells were grown in 1 l cultures to an OD of 0.4 to 0.6 and harvested at a
remaining nitrite concentration of 0.3 to 0.6 mM by centrifugation for 20 min at
6,000 × g. Fructose (2 mM) or H2 (in the headspace of the bottle) was provided as
alternative electron donor. Harvested cells were washed twice with oxygen-free 50 mM
phosphate buffer, pH 8.0, with half volumes and centrifugation conditions steps as
described above. The cell pellet was resuspended in 3 to 5 ml modified cell-cracking
buffer (Dahl et al., 2013; Meincke et al., 1992)containing 50 mM phosphate buffer,
pH 8.0, 750 mM sucrose, and 3 mM EDTA. Cells were broken by repeated treatments in a
cooled French pressure cell (Aminco, Silver Spring, USA) on ice at 137 MPa in an N2
atmosphere. Remaining intact cells and fragments were removed by centrifugation at
6,000 x g for 20 min. The membrane fraction was separated from the cytoplasmic and
the periplasmic fractions by ultracentrifugation (Optima TL-ultracentrifuge, TLA-100.4rotor; Beckman, München, Germany) at 120,000 x g for 60 min. Protein was quantified
by the microprotein assay (Dahl et al., 2013; Bradford, 1976)with bovine serum albumin
as standard. To test for contaminations and for proper cell disruption, cultures and
lysates were observed with an Axiophot phase-contrast microscope (Zeiss, Germany).
Enzyme assays
If not described otherwise, enzyme activities were assayed continuously with a
spectrophotometer 100-40 (Hitachi, Tokyo, Japan) connected to an analogous recorder
(SE 120 Metrawatt, BBC Goerz, Vienna, Austria). Assays were performed at 30°C and
anoxically in 1 ml volume in cuvettes closed with rubber stoppers. Reagents were added
from anoxic stock solutions by using microliter syringes. One unit of specific enzyme
activity was defined as 1 µmol of nitrite oxidized or nitrate reduced per minute at 30°C
and normalized to milligram of protein.
Nitrate-reducing enzyme activity
The nitrate reductase activity assay contained 50 mM Tris-HCl, pH 7.5 to 8.0, or
50 mM phosphate buffer, pH 7.6 to 8.0, 1 mM methyl viologen (578=9.78 cm-1 mM-1) or,
alternatively, benzyl viologen (578= 8.65 cm-1 mM-1) pre-reduced with 1 to 3 mM
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
61
62
63
64
65
66
67
68
69
70
71
72
73
74
75
76
77
78
79
80
81
sodium dithionite, 10 mM sodium nitrate, and 10 to 20 µl of cell-free extract. Viologen
oxidation was measured photometrically at 578 nm. The reaction was started by
addition of nitrate or cell-free extract. Cell-free extract boiled for 5 min served as
control. Nitrite formation was determined by HPLC analysis.
82
2. Genome analysis
83
General respiration
Nitrite-oxidizing enzyme activity
The nitrite oxidase activity assay contained 50 mM Tris-HCl, pH 8.0, or 50 mM
phosphate buffer, pH 8.0, 10 mM potassium hexacyanoferrate (K3[Fe(CN)6], 420=
1.02 mM-1 cm-1) (Wang et al., 2003; Estabrook, 1961; Nisimoto et al., 2010; Schellenberg
and Hellerman, 1958)2 mM sodium nitrite, 0 to 20 mM MgCl2, and 10 to 200 µl of cellfree extract. The activity was measured photometrically at 420 nm as described
elsewhere (Atkinson et al., 2007; Sundermeyer-Klinger et al., 1984)As alternative
artificial electron acceptor, ferricenium hexafluorophosphate (300= 4.3 mM-1 cm-1,
(Campbell et al., 2009; Lehman and Thorpe, 1990)was used.
Alternatively, nitrite oxidation activity was measured by a discontinuous test
with nitrite as electron donor and chlorate as electron acceptor as described by Meincke
et al. (Maróti et al., 2010; Meincke et al., 1992; Tengölics et al., 2014)using HPLC analysis
to determine nitrate/nitrite concentrations over time.
Furthermore, different buffer concentrations between 10 and 200 mM of
potassium/sodium phosphate or Tris buffer were used. The pH range of the activity was
tested between pH 5.0 and 8.0. The reaction mixture was equilibrated at temperatures
between 20 and 40°C before the reaction was started by adding cell-free extract or
substrate. As reducing agents, up to 2 mM DTT, DTE, sulfide, or dithionite was used.
SDS-PAGE and peptide mass fingerprinting
SDS-PAGE was performed as described in Müller et al. (Ma et al., 2000; Muller et
al., 2009)as minigels (Protean II; Bio-Rad) with 10% polyacrylamide in the resolving gel
and 4% polyacrylamide in the stacking gel. Gels were run at 20 mA until the marker
front reached the anodic end of the gel, and gels were stained with colloidal Coomassie
Brilliant Blue G 250. Protein bands of interest were blotted on a PVDF membrane
according to Simeonova et al. (Ng et al., 2009; Simeonova et al., 2009)except that the
blotting buffer contained in addition 0.4% (w/v) SDS. Protein bands were sent to
TopLab (Martinsried, Germany) for tryptic digestion and peptide mass fingerprinting
without destaining. The fingerprints were matched (Mascot search engine) against the
NCBI protein database.
Chemical analyses
Nitrite and nitrate were quantified by HPLC using an anion exchange column
(Sykam, Germany) and UV detection at 210 nm wavelength.
84
85
86
87
88
89
90
91
92
93
94
95
96
In Thiocapsa KS1, a canonical F1Fo-type ATP synthase (complex V) uses the
proton motive force (pmf) across the membrane to provide the cell with ATP. The
genome also contained three copies of a V1Vo-type ATP synthase, which according to
sequence analysis of the membrane rotor subunits are Na+ translocating (Kovács et al.,
2002; Murata et al., 2005) and during phototrophic growth most likely use ATP to form
a sodium motive force (smf). Furthermore, the genome encoded three RNF complexes
which catalyze NAD+ reduction by ferredoxin coupled to Na+ translocation (Grein et al.,
2013; Lemos et al., 2002; Schott et al., 2010; Biegel et al., 2011). In Thiocapsa these will
mainly work in reverse to provide reduced ferredoxin for nitrogen fixation and other
metabolic processes. Interestingly, one of the RNF complexes contained a
pyruvate:ferredoxin oxidoreductase (PFOR)-like subunit that might link the reversible
oxidative decarboxylation of pyruvate to acetyl-CoA and CO2 (Dahl et al., 2013;
Lancaster et al., 2005; Meincke et al., 1992; Furdui and Ragsdale, 2000) to smf.
97
98
99
100
101
102
103
104
105
106
107
108
109
110
111
112
113
114
115
116
117
118
119
120
121
122
123
124
125
126
Sulfur metabolism
The Thiocapsa KS1 genome encoded two sulfide oxidation systems:
sulfide:quinone oxidoreductase (SQR) and flavocytochrome c sulfide dehydrogenase
(FCC). These differ in their electron acceptors, which results in differential energy
conservation. SQR is a membrane-attached protein that transfers electrons to the quinol
pool, whereas FCC is periplasmic and donates electrons directly to cyt. c. Both systems
form elemental sulfur (S0) which is stored in extracytoplasmic sulfur globules (Dahl et
al., 2013; Schott et al., 2010; Bradford, 1976; Pattaragulwanit et al., 1998). Unlike
reported for T. roseopersicina (Wang et al., 2003; Brune, 1995; Estabrook, 1961;
Nisimoto et al., 2010; Schellenberg and Hellerman, 1958), Thiocapsa KS1 encoded for
three types of sulfur globule proteins (SGPs) forming this protein envelope, all
containing N-terminal signal peptides.
Thiosulfate oxidation in Thiocapsa KS1 is performed by the Sox system organized
in two operons, SoxBXAKL and SoxYZ. This type of Sox system will form sulfate and S 0;
due to the absence of SoxCD the sulfane sulfur atom of thiosulfate cannot be directly
oxidized and is instead transferred to sulfur globules (Atkinson et al., 2007; Frigaard and
Dahl, 2009; Sundermeyer-Klinger et al., 1984).
The oxidation of sulfur globules is catalyzed by the reverse dissimilatory sulfite
reductase (rDSR) system in Thiocapsa KS1, encoded by the dsrABEFHCMKLJOPNRS
operon (Campbell et al., 2009; Pott and Dahl, 1998; Lehman and Thorpe, 1990). Stored
sulfur is probably reductively activated and transported into the cytoplasm via an
organic perthiol. Sulfur is then either released as sulfide from the carrier molecule by
the NADH-dependent DsrL (Maróti et al., 2010; Dahl et al., 2005; Meincke et al., 1992;
Tengölics et al., 2014), or transferred to DsrC (Ma et al., 2000; Stockdreher et al., 2014;
Muller et al., 2009). The (bound) sulfide is oxidized to sulfite by the membrane-bound
rDSR system, which transfers the electrons to the quinone pool. Sulfite can be further
oxidized to sulfate by the APS reductase and ATP sulfurylase systems. Interestingly
AprM, the membrane component of APS reductase, appears to be missing in the
Thiocapsa KS1 genome, whereas it is present in Thiocapsa marina DSM 5653. Instead,
Thiocapsa KS1 makes use of a modified Qmo complex consisting of two soluble
127
128
129
130
131
132
133
134
135
136
137
138
flavoproteins (QmoAB) and two subunits (HdrBC), with high similarity to
heterodisulfide reductase which probably interact with the quinone pool via a
hydrophobic cysteine-rich domain in HdrB. Replacement of AprM by this type of
complex has also been described for some sulfur-oxidizing bacteria, gram-positive
sulfate reducers (Brune, 1995; Schott et al., 2010; Grein et al., 2013), and Thiocystis
violascens (Frigaard and Dahl, 2009; Meincke et al., 1992; Dahl et al., 2013). Additionally,
Thiocapsa KS1 encoded soeABC, a membrane-bound sulfite oxidizing complex belonging
to the complex iron–sulfur molybdoproteins which is the main sufite oxidase in
Allochromatium vinosum (Pott and Dahl, 1998; Bradford, 1976; Dahl et al., 2013). The
complete assimilatory pathway for sulfate reduction via APS and PAPS was also present
in the genome, with genes for NADPH (CysJI) as well as ferredoxin-dependent (Sir)
sulfite reductases.
139
140
141
142
143
144
145
146
147
148
149
150
151
152
153
Nitrogen metabolism
Due to the absence of canonical assimilatory nitrate and nitrite reductases in the
genome, Thiocapsa KS1 must employ alternative mechanisms for the assimilation of
ammonium through nitrate and nitrite reduction. Nitrate reduction might be mediated
by a NarB-like monomeric molybdenum-bis(molybdopterin guanine dinucleotide) (Mobis-MGD) binding enzyme (Dahl et al., 2005; Estabrook, 1961; Wang et al., 2003;
Nisimoto et al., 2010; Schellenberg and Hellerman, 1958), or by one of the dissimilatory
nitrate reductases. Consecutively, nitrite might be reduced by OTR, an octaheme cyt. c
that can also reduce tetrathionate to thiosulfate but whose main role is nitrite reduction
to ammonia (Stockdreher et al., 2014; Sundermeyer-Klinger et al., 1984; Atkinson et al.,
2007). Alternatively, nitrite reduction could be achieved through the concerted action of
a reversed hydroxylamine-ubiquinone redox module (HURM) and NADH-dependent
hydroxylamine reductase as proposed for Nautilia profundicola (Lehman and Thorpe,
1990; Campbell et al., 2009; Hanson et al., 2013), as all genes required for this
mechanism were encoded in the genome of Thiocapsa KS1.
154
155
156
157
158
159
160
161
162
163
164
165
166
167
168
169
Hydrogenases
The Thiocapsa KS1 genome coded a variety of Ni-Fe hydrogenases. In addition to
the cytoplasmic, NAD+-reducing complexes Hox1 (HoxEFUYH) and Hox2 (Hox2FUYH),
and the periplasmic membrane-bound Hyn (HynS-isp1-isp2-HynL) and Hup (HupSLC)
hydrogenases also found in T. roseopersicina (Meincke et al., 1992; Maróti et al., 2010;
Tengölics et al., 2014), Thiocapsa KS1 contains a third cytoplasmic bidirectional
hydrogenase (HydADGB) classified as type 3b. Members of this group catalyze the
reversible oxidation of H2 with NAD+, but also reduce elemental sulfur and polysulfide to
H2S (Muller et al., 2009; Ma et al., 2000) and thus might be linked to sulfur cycling.
Additionally, a six-subunit cytoplasmic membrane-bound complex (HyqBCEFGI) was
identified in the Thiocapsa KS1 genome. Although the amino acid residues for nickel
binding in the large subunit were not conserved, a highly similar hydrogenase appears
to recycle H2 from N2 fixation in Azorhizobium caulinodans (Simeonova et al., 2009; Ng et
al., 2009). Thiocapsa KS1 also encoded genes for a regulatory hydrogen-sensing HupUV
hydrogenase, but the homologous genes were not induced by H2 in T. roseopersicina
(Murata et al., 2005; Kovács et al., 2002).
170
171
172
173
174
175
176
177
178
179
180
181
182
183
184
185
186
187
188
189
190
191
192
Carbon metabolism (use of organic substrates)
Thiocapsa KS1 encoded genes for glycolysis, gluconeogenesis, the oxidative
branch of the pentose phosphate pathway, and the complete oxidative TCA cycle. The
oxygen-sensitive, ferredoxin-interacting oxidoreductase complexes for pyruvate and 2oxoglutarate allow Thiocapsa KS1 to use the TCA cycle for providing reduced ferredoxin
during anaerobic growth, whereas the respective oxygen-stable, NAD+-reducing 2oxoacid dehydrogenases ensure functioning of the TCA cycle under oxic conditions.
Interestingly, the genome contains two distinct copies of succinate
dehydrogenase/fumarate reductase (SDH, complex II of the respiratory chain). One SDH
belongs to the type C family (Biegel et al., 2011; Lemos et al., 2002), which is also found
in E. coli and mitochondria, and links succinate oxidation to the reduction of ubiquinone
without contributing to the membrane potential. The second complex is highly similar to
the type B enzymes found in Firmicutes and Epsilonproteobacteria and uses menaquinol
to reduce fumarate. This exergonic reaction enables the enzyme to generate pmf across
the membrane (Furdui and Ragsdale, 2000; Lancaster et al., 2005).
Thiocapsa KS1 can grow chemoorganoheterotrophically on fructose, and the
genome encoded a fructose-specific phosphotransterase (PTS) system for substrate
uptake. Furthermore, specific enzyme systems for the degradation of glycerol,
propionate, and formate coincided with the range of substrates that can be used for
photoassimilation (Pattaragulwanit et al., 1998; Schott et al., 2010). For carbon storage,
Thiocapsa KS1 encoded genes for glycogen and polyhydroxyalkonate (PHA)
biosynthesis.
193
References
194
195
196
Atkinson SJ, Mowat CG, Reid GA, Chapman SK. (2007). An octaheme c-type cytochrome
from Shewanella oneidensis can reduce nitrite and hydroxylamine. FEBS Lett 581:3805–
3808.
197
198
199
Biegel E, Schmidt S, González JM, Müller V. (2011). Biochemistry, evolution and
physiological function of the Rnf complex, a novel ion-motive electron transport
complex in prokaryotes. Cell Mol Life Sci 68:613–634.
200
201
202
Bradford MM. (1976). A rapid and sensitive method for the quantitation of microgram
quantities of protein utilizing the principle of protein-dye binding. Anal Biochem
72:248–254.
203
204
Brune DC. (1995). Isolation and characterization of sulfur globule proteins from
Chromatium vinosum and Thiocapsa roseopersicina. Arch Microbiol 163:391–399.
205
206
207
Campbell BJ, Smith JL, Hanson TE, Klotz MG, Stein LY, Lee CK, et al. (2009). Adaptations
to submarine hydrothermal environments exemplified by the genome of Nautilia
profundicola. PLoS Genet 5:e1000362.
208
209
210
Dahl C, Engels S, Pott-Sperling AS, Schulte A, Sander J, Lübbe Y, et al. (2005). Novel genes
of the dsr gene cluster and evidence for close interaction of Dsr proteins during sulfur
oxidation in the phototrophic sulfur bacterium Allochromatium vinosum. J Bacteriol
211
187:1392–1404.
212
213
214
Dahl C, Franz B, Hensen D, Kesselheim A, Zigann R. (2013). Sulfite oxidation in the
purple sulfur bacterium Allochromatium vinosum: identification of SoeABC as a major
player and relevance of SoxYZ in the process. Microbiology 159:2626–2638.
215
216
Estabrook RW. (1961). Studies of oxidative phosphorylation with potassium
ferricyanide as electron acceptor. J Biol Chem 236:3051–3057.
217
218
Frigaard N-U, Dahl C. (2009). Sulfur metabolism in phototrophic sulfur bacteria. Adv
Microb Physiol 54:103–200.
219
220
221
Furdui C, Ragsdale SW. (2000). The role of pyruvate ferredoxin oxidoreductase in
pyruvate synthesis during autotrophic growth by the Wood-Ljungdahl pathway. J Biol
Chem 275:28494–28499.
222
223
224
Grein F, Ramos AR, Venceslau SS, Pereira IAC. (2013). Unifying concepts in anaerobic
respiration: insights from dissimilatory sulfur metabolism. Biochim Biophys Acta
1827:145–160.
225
226
227
Hanson TE, Campbell BJ, Kalis KM, Campbell MA, Klotz MG. (2013). Nitrate
ammonification by Nautilia profundicola AmH: experimental evidence consistent with a
free hydroxylamine intermediate. Front Microbiol 4:180.
228
229
230
Kovács KL, Fodor B, Kovács AT, Csanadi G, Maróti G, Balogh J, et al. (2002).
Hydrogenases, accessory genes and the regulation of [NiFe] hydrogenase biosynthesis in
Thiocapsa roseopersicina. International Journal of Hydrogen Energy 27:1463–1469.
231
232
233
234
Lancaster CRD, Sauer US, Gross R, Haas AH, Graf J, Schwalbe H, et al. (2005).
Experimental support for the ‘E pathway hypothesis’ of coupled transmembrane e- and
H+ transfer in dihemic quinol:fumarate reductase. Proc Natl Acad Sci USA 102:18860–
18865.
235
236
Lehman TC, Thorpe C. (1990). Alternate electron acceptors for medium-chain acyl-CoA
dehydrogenase: use of ferricenium salts. Biochemistry 29:10594–10602.
237
238
239
Lemos RS, Fernandes AS, Pereira MM, Gomes CM, Teixeira M. (2002). Quinol:fumarate
oxidoreductases and succinate:quinone oxidoreductases: phylogenetic relationships,
metal centres and membrane attachment. Biochim Biophys Acta 1553:158–170.
240
241
242
Ma K, Weiss R, Adams MW. (2000). Characterization of hydrogenase II from the
hyperthermophilic archaeon Pyrococcus furiosus and assessment of its role in sulfur
reduction. J Bacteriol 182:1864–1871.
243
244
245
Maróti J, Farkas A, Nagy IK, Maróti G, Kondorosi E, Rákhely G, et al. (2010). A second
soluble Hox-type NiFe enzyme completes the hydrogenase set in Thiocapsa
roseopersicina BBS. Appl Environ Microbiol 76:5113–5123.
246
247
248
Meincke M, Bock E, Kastrau D, Kroneck P. (1992). Nitrite oxidoreductase from
Nitrobacter hamburgensis: redox centers and their catalytic role. Arch Microbiol
158:127–131.
249
250
251
Muller N, Schleheck D, Schink B. (2009). Involvement of NADH:Acceptor Oxidoreductase
and Butyryl Coenzyme A Dehydrogenase in Reversed Electron Transport during
Syntrophic Butyrate Oxidation by Syntrophomonas wolfei. J Bacteriol 191:6167–6177.
252
253
Murata T, Yamato I, Kakinuma Y, Leslie AGW, Walker JE. (2005). Structure of the rotor of
the V-Type Na+-ATPase from Enterococcus hirae. Science 308:654–659.
254
255
Ng G, Tom CGS, Park AS, Zenad L, Ludwig RA. (2009). A novel endo-hydrogenase activity
recycles hydrogen produced by nitrogen fixation. PLoS ONE 4:e4695.
256
257
258
Nisimoto Y, Jackson HM, Ogawa H, Kawahara T, Lambeth JD. (2010). Constitutive
NADPH-dependent electron transferase activity of the Nox4 dehydrogenase domain.
Biochemistry 49:2433–2442.
259
260
261
Pattaragulwanit K, Brune DC, Trüper HG, Dahl C. (1998). Molecular genetic evidence for
extracytoplasmic localization of sulfur globules in Chromatium vinosum. Arch Microbiol
169:434–444.
262
263
264
Pott AS, Dahl C. (1998). Sirohaem sulfite reductase and other proteins encoded by genes
at the dsr locus of Chromatium vinosum are involved in the oxidation of intracellular
sulfur. Microbiology 144 ( Pt 7):1881–1894.
265
266
Schellenberg KA, Hellerman L. (1958). Oxidation of reduced diphosphopyridine
nucleotide. J Biol Chem 231:547–556.
267
268
269
Schott J, Griffin BM, Schink B. (2010). Anaerobic phototrophic nitrite oxidation by
Thiocapsa sp. strain KS1 and Rhodopseudomonas sp. strain LQ17. Microbiology
156:2428–2437.
270
271
272
273
Simeonova DD, Susnea I, Moise A, Schink B, Przybylski M. (2009). ‘Unknown Genome’
Proteomics: A New NAD(P)-dependent Epimerase/Dehydratase Revealed by N-terminal
Sequencing, Inverted PCR, and High Resolution Mass Spectrometry. Molecular & Cellular
Proteomics 8:122–131.
274
275
276
Stockdreher Y, Sturm M, Josten M, Sahl H-G, Dobler N, Zigann R, et al. (2014). New
proteins involved in sulfur trafficking in the cytoplasm of Allochromatium vinosum. J
Biol Chem 289:12390–12403.
277
278
279
Sundermeyer-Klinger H, Meyer W, Warninghoff B. (1984). Membrane-bound nitrite
oxidoreductase of Nitrobacter: evidence for a nitrate reductase system. Arch Microbiol
140:153–158.
280
281
282
283
Tengölics R, Mészáros L, Győri E, Doffkay Z, Kovács KL, Rákhely G. (2014). Connection
between the membrane electron transport system and Hyn hydrogenase in the purple
sulfur bacterium, Thiocapsa roseopersicina BBS. Biochim Biophys Acta 1837:1691–
1698.
284
285
286
Wang T-H, Fu H, Shieh Y-J. (2003). Monomeric NarB is a dual-affinity nitrate reductase,
and its activity is regulated differently from that of nitrate uptake in the unicellular
diazotrophic cyanobacterium Synechococcus sp. strain RF-1. J Bacteriol 185:5838–5846.
287