Electron Bifurcating FixABCX Protein 
Complex from Azotobacter vinelandii: 
Generation of Low-Potential Reducing 
Equivalents for Nitrogenase Catalysis
Authors: Rhesa N. Ledbetter, Amaya M. Garcia 
Costas, Carolyn E. Lubner, David W. Mulder, Monika 
Tokmina-Lukaszewska, Jacob H. Artz, Angela 
Patterson, Timothy S. Magnuson, Zackary J. Jay, H. 
Diessel Duan, Jacquelyn Miller, Mary H. Plunkett, 
John P. Hoben, Brett M. Barney, Ross P. Carlson, 
Anne-Frances Miller, Brian Bothner, Paul W. King, 
John W. Peters, & Lance C. Seefeldt
This document is the unedited author's version of a Submitted Work that was subsequently 
accepted for publication in Biochemistry, copyright © American Chemical Society after peer 
review. To access the final edited and published work, see https://doi.org/10.1021/
acs.biochem.7b00389.
Ledbetter RN, Amaya M. Garcia Costas, Carolyn E.  Lubner, David W. Mulder, Monika Tokmina-
Lukaszewska, Jacob H.  Artz, Angela Patterson, Timothy S. Magnuson, Zackary J. Jay, H. Dissel 
Duan, Jacquelyn Miller, Mary H. Plunkett, John P. Hoben, Brett M. Barney, Ross P. Carlson, 
Anne-Frances Miller, Brian Bothner, Paul W. King, John W. Peters, Lance C. Seefeldt, “The 
Electron Bifurcating FixABCX Protein Complex from Azotobacter Vinelandii: Generation of Low-
Potential Reducing Equivalents for Nitrogenase Catalysis.” Biochemistry 56, no. 32 (August 3, 
2017): 4177–4190. 
Made available through Montana State University’s ScholarWorks 
scholarworks.montana.edu 
The Electron Bifurcating FixABCX Protein 
Complex from Azotobacter vinelandii: 
Generation of Low-Potential Reducing 
Equivalents for Nitrogenase Catalysis
Rhesa N. Ledbetter, Amaya M. Garcia Costas, Carolyn E. Lubner, David 
W. Mulder, Monika Tokmina-Lukaszewska, Jacob H. Artz, Angela 
Patterson, Timothy S. Magnuson, Zackary J. Jay, H. Diessel Duan, 
Jacquelyn Miller, Mary H. Plunkett, John P. Hoben, Brett M. Barney, Ross P. 
Carlson, Anne-Frances Miller, Brian Bothner, Paul W. King, John W. Peters
and Lance C. Seefeldt
 The biological reduction of dinitrogen (N2) to ammonia (NH3) by nitrogenase 
is an energetically demanding reaction that requires low-potential electrons and 
ATP; however, pathways used to deliver the electrons from central metabolism 
to the reductants of nitrogenase, ferredoxin or flavodoxin, remain unknown for 
many diazotrophic microbes. The FixABCX protein complex has been 
proposed to reduce flavodoxin or ferredoxin using NADH as the electron donor 
in a process known as electron bifurcation. Herein, the FixABCX complex from 
Azotobacter vinelandii was purified and demonstrated to catalyze an electron 
bifurcation reaction: oxidation of NADH (Em = −320 mV) coupled to 
reduction of flavodoxin semiquinone (Em = −460 mV) and reduction of 
coenzyme Q (Em = 10 mV). Knocking out fix genes rendered Δrnf A. 
vinelandii cells unable to fixdinitrogen,confirming that the FixABCX 
system provides another route for delivery of electrons to nitrogenase. 
Characterization of the purified FixABCX complex revealed the presence 
of flavin and iron−sulfur cofactors confirmed by native mass spectrometry, 
electron paramagnetic resonance spectroscopy, and transient absorption 
spectroscopy. Transient absorption spectroscopy further established the 
presence of a short-lived flavin semiquinone radical, suggesting that a 
thermodynamically unstable flavin semiquinone may participate as an 
intermediate in the transfer of an electron to flavodoxin. A structural model 
of FixABCX, generated using chemical cross-linking in conjunction with 
homology modeling, revealed plausible electron transfer pathways to both 
high- and low-potential acceptors. Overall, this study informs a mechanism 
for electron bifurcation, offering insight into a unique method for delivery 
of low-potential electrons required for energy-intensive biochemical 
conversions.
Biological dinitrogen (N2) fixation is performed 
by diazotrophic microbes, which harbor the 
enzyme nitro-genase. This enzyme converts N2 
into bioavailable ammonia (NH3)(eq 1) and 
accounts for at least half of the production of fixed 
nitrogen on Earth. 1−3
+
→
+ + −+ 16MgATP 8eN2 8H
22NH3 H+ + 16MgADP 16P+ i (1)
As summarized by eq 1, the biological reduction of N2 
is an energy-demanding reaction, requiring both ATP 
and low-reduction potential electrons. These electrons 
are provided by small redox proteins, ferredoxin (Fd) 
and flavodoxin (Fld), which serve as direct donors of 
electrons to the iron (Fe) protein
of nitrogenase.4−10 The redox active iron−sulfur clusters of
Fd typically access one redox couple (FdOx/Red) with midpoint
reduction potentials (Em) ranging from 0 to −645 mV.
4,11 Two
redox couples of the flavin in Fld are accessible, including the
oxidized quinone/semiquinone (FldOx/Sq) and semiquinone/
hydroquinone (FldSq/Hq) couples.5,9,12 In general, the Em of the
FldOx/Sq couple ranges from −50 to −250 mV and that of the
FldSq/Hq couple from −370 to −500 mV.4,7 Only the FldSq/Hq
couple of Fld has enough driving force to donate electrons to
nitrogenase. While much is known about other aspects of
biological nitrogen fixation, pathways for delivery of the low-
potential reducing equivalents for Fd and Fld reduction are not
well understood for many diazotrophs.4 It has been shown that
the nitrogen-fixing organism Klebsiella pneumoniae uses the
anaerobic oxidation of pyruvate to reduce Fld,13,14 and it was
proposed that other microbes likely use energy associated with
the proton motive force to drive reduction of a low-potential
electron donor.15,16
Recently, a new mechanism for generating a reductant for
nitrogen fixation was put forward. Flavin-based electron
bifurcation (FBEB), considered a third fundamental form of
energy conservation, couples exergonic and endergonic electron
transfer reactions to limit free energy loss in biological
systems.17−19 FBEB exploits a favorable electron transfer event
to drive a thermodynamically unfavorable reaction without the
use of ATP or an electrochemical gradient.17−19 Several
bifurcating complexes, all of which contain a flavin as the
proposed site of bifurcation, have been identified in anaerobic
bacteria and archaea.17,18 While these enzymes catalyze a variety
of reactions, all characterized thus far use Fld/Fd as an electron
donor or acceptor.20−24 For example, the electron transferring
flavoprotein/butyryl-coenzyme A (Etf-Bcd) bifurcating system
uses the electron donor NADH (Em = −320 mV) to reduce
Fld (Em
Sq/Hq = −430 mV) or Fd (Em = −405 mV). This
thermodynamically unfavorable reaction is achieved by coupling
it to an exergonic one, in this case, the reduction of crotonyl-CoA
(Em = −10 mV).17,22,23 The bifurcation of electrons to both
a high- and a low-potential acceptor results in an overall
thermodynamically favorable reaction.17,22,23,25
Homologues of bifurcating electron transfer flavoproteins
(Etfs), known as FixAB, have been found in physiologically and
phylogenetically distinct nitrogen-fixing organisms such as
Azotobacter vinelandii,26,27 Rhodopseudomonas palustris,28 Rhodo-
spirillum rubrum,29 and Sinorhizobium meliloti.30,31 Previous
studies demonstrated that disrupting the Fix system inR. palustris,
Rh. rubrum, and S. meliloti completely abolishes or signifi-
cantly impairs their ability to grow under nitrogen fixing condi-
tions, suggesting that Fix may serve as a source of electrons to
nitrogenase.28−30 Given the homology between FixAB and
known bifurcating Etfs, it was hypothesized that the Fix system
uses electron bifurcation to generate low-potential reducing
equivalents for nitrogenase.20
While the FixABCX complex is clearly linked to nitrogen
fixation in many diazotrophs, its specific role has not been firmly
established, nor has its ability to generate a reductant for
nitrogenase via electron bifurcation been demonstrated. Here,
we report electron bifurcation by the FixABCX complex from the
obligate aerobe A. vinelandii and characterize the Fix proteins
using advanced biochemical and spectroscopic tools. In addition,
we provide evidence that FixABCX provides electrons for
nitrogenase in A. vinelandii cells. Overall, this work establishes a
new pathway for the generation of a low-potential reductant
required by nitrogenase and elucidates a mechanism by which
biology can overcome thermodynamic barriers to accomplish a
difficult reductive reaction.
■ MATERIALS AND METHODS
Construction of anA. vinelandiiΔf ixMutant.A. vinelandii
strain DJ was the wild type strain used for physiological studies
and mutant construction. Strains UW195 (Δrnf1) and UW207
(Δrnf 2) were kindly donated by L. Rubio.32 The Δf ix mutant
was generated by gene disruption with an antibiotic resistance
cassette as shown in Figure S1 using primers in Table S1. Briefly,
two 1.2 kb DNA fragments that included 0.3 kb of f ixA
(Avin_10520) and 0.1 kb of f ixC (Avin_10540) were obtained
by polymerase chain reaction (PCR) and cloned sequentially
in the pT7-7 ampicillin resistant vector using NdeI/BamHI and
BamHI/HindIII as restriction cloning sites. The kanamycin
resistance (KmR) gene (aph) was isolated as a 1.5 kb BamHI
fragment from mini-Tn533 and inserted between the 1.2 kb
upstream and downstream regions previously cloned in the
pT7-7 vector using the BamHI restriction cloning site. The final
construct was transformed into A. vinelandii strain DJ as
described previously.34,35 KmR transformants were screened for
sensitivity to ampicillin (AmpS); AmpS derivatives were assumed
to have arisen from a double-crossover recombination event
in which the wild type f ixABC genes were replaced by the
aph-containing cassette (Figure S1). This replacement was
confirmed by PCR and sequencing (Figure S1, and Table S1).
Burk’s medium36 with no nitrogen source or supplemented with
ammonium acetate (1 g/L) was used for physiological analyses
of A. vinelandii DJ and Δrnf and Δf ix mutants.
Homologous Overexpression of FixABCX. The f ix genes
of A. vinelandii were homologously overexpressed by being
placed under control of the nifH promoter, which is used
for the transcription of genes associated with the molybdenum-
dependent nitrogenase (Figure S2). The 4.3 kb f ix operon, which
includes six genes ( f ixfd, Avin_10510; f ixA, Avin_10520; f ixB,
Avin_10530; f ixC, Avin_10540; f ixX, Avin_10550; ORF6,
Avin_10560) (Figure S2), was amplified via PCR using primers
specified in Table S2. The PCR product was digested with XbaI
and BamHI and inserted into a slightly modified pUC19 vector
for blue-white screening. UsingNdeI and BamHI restriction sites,
the f ix genes were then cloned into a vector built specifically
to support the insertion of genes behind the nifH promoter.37
The f ix operon was inserted between two segments of DNA from
the nif region of A. vinelandii. The flanking regions served as sites
for homologous recombination within the chromosome,
allowing replacement of nifHD with f ix genes (Figure S2).
The presence of a streptomycin resistance gene (SmR) between
the flanking regions provided a selectable marker upon
transformation of the plasmid into A. vinelandii.34 The proper
insertion of the f ix genes into the final plasmid and the
chromosome of A. vinelandii were confirmed by PCR and
sequencing (Table S2).
Growth of Recombinant A. vinelandii and Purification
of the FixABCX Complex. Recombinant A. vinelandii was
grown in a 100 L fermenter (BioFlo 610, New Brunswick,
Hauppauge, NY) at Utah State University’s Synthetic Bio-
manufacturing Institute. Cells were grown in Burk’s medium
supplemented with 10 mM urea as a nitrogen source.36 In the
presence of urea, the nifH promoter is repressed. When the cells
reached an OD600 of ∼1.8, they were harvested in a stacked disk
centrifuge (TSE 10, GEA Westfalia, Northvale, NJ) at 14200g
and resuspended in Burk’s medium with no source of fixed
nitrogen. Upon removal of the fixed nitrogen source, the nifH
promoter is derepressed and gene expression occurs. A. vinelandii
was derepressed for 5 h to achieve optimal expression of Fix
proteins. Because nif genes were replaced with f ix genes, Fix
rather than nitrogenase was expressed. Following the derepres-
sion, cells were harvested and stored at −80 °C until further use.
Fermenter conditions weremaintained as follows throughout the
growth and derepression: 30 °C, pH 7, 20% dissolved oxygen,
and agitation at ∼200 rpm.
All purification steps were performed anaerobically under an
argon atmosphere. One hundred grams of wet cell paste was
resuspended in 50 mM HEPES (pH 8), 150 mM NaCl, 1 mM
dithiothreitol (DTT), and 2 mg of DNase at a cell:buffer ratio
of 1:5. Flavin adenine dinucleotide (FAD) (0.25 mM) was also
added to the lysis buffer as it increased the flavin cofactor
occupancy of the Fix complex. Lysis was achieved with a French
pressure cell (SLM Aminco FA-078, Aminco, Rochester, NY)
at 200 MPa. The cell extract was centrifuged at 8000g for 15 min
to remove cell debris. A second centrifugation was then
conducted at 50000g for 2 h to obtain the cell membrane
fraction. The membranes were solubilized for 1 h at 4 °C in
50 mM HEPES (pH 8), 150 mM NaCl, 1% (w/v) n-dodecyl
β-D-maltoside (DDM), and 1 mM DTT at a pellet:buffer ratio
of approximately 1:6. The solubilizedmembrane was obtained by
centrifugation at 50000g for 1 h and then diluted 3-fold in buffer
A [50mMHEPES (pH 8), 0.02% (w/v) DDM, and 1mMDTT]
to reduce the salt and detergent concentration before being
loaded onto a 100 mL Q-Sepharose column. The Q-Sepharose
column was prewashed with 2 column volumes of buffer B
[50 mM HEPES (pH 8), 1 M NaCl, 0.02% (w/v) DDM, and
1 mM DTT] and then equilibrated with 2 column volumes of
buffer A. Once the protein was loaded, unbound proteins were
removed with 2 column volumes of buffer A, followed by elution
of bound proteins with a salt gradient from 15 to 60% (5 column
volumes). FixABCX eluted between 32 and 36%NaCl. Fractions
were pooled and diluted to <100 mM NaCl for concentration
on a 15 mL Q-Sepharose column prewashed and equilibrated
as described above. After loading, the column was washed with
2 column volumes of buffer A and bound protein eluted with
500 mM NaCl. The resultant concentrated fraction was loaded
onto a Sephacryl S-200 column equilibrated with 50 mMHEPES
(pH 8), 150 mM NaCl, 0.02% (w/v) DDM, and 1 mM DTT.
Fractions containing FixABCX were pooled and concentrated
using an Amicon (EMD Millipore, Billerica, MA) concentrator
with a 100 kDa cutoff membrane and stored in liquid nitrogen.
The purity of FixABCX was determined using sodium dodecyl
sulfate−polyacrylamide gel electrophoresis (SDS−PAGE), and
the protein concentration was measured using the DC Protein
Assay (Bio-Rad, Hercules, CA).
Heterologous Overexpression and Purification of
R. palustris FixAB. R. palustris FixAB was co-expressed
in Escherichia coli strain NiCo21(DE3) (New England Biolabs,
Ipswich, MA) transformed with plasmids based on pMCSG28
and pMCSG21 (DNASU Plasmid Repository, Arizona State
University, Tempe, AZ) modified to carry the f ixA gene with a
C-terminal His tag and the f ixB gene with an N-terminal His tag,
respectively. Cells were grown in terrific broth (TB)
supplemented with riboflavin (20 mg/L) and MgSO4 (2 mM)
along with carbenicillin (100 μg/mL) and spectinomycin (100
μg/mL) at 37 °C while being shaken at 200 rpm to an OD600 of
∼2. After the culture had been fully cooled to 18−20 °C, f ix gene
expression was induced with β-D-1-thiogalactopyranoside
(IPTG) (0.1 mM) and cells were grown for an additional 12 h
at this lower temperature. Cells were harvested by centrifugation
at 11900g and 4 °C for 6min, and the pellet was stored at−80 °C.
The cell pellet was suspended in BugBuster (80 mL) (EMD
Millipore) containing 4-(2-aminoethyl)benzenesulfonyl fluoride
hydrochloride (AEBSF) (1 mM), tris(2-carboxyethyl)phosphine
hydrochloride (TCEP) (1 mM), FAD (1 mM), benzonase
nuclease (20 μL), and rLysozyme (2 μL) (EMD Millipore) and
further incubated at 4 °C for 2 h while being stirred. After
centrifugation at 20000g for 30 min at 4 °C, the supernatant was
filtered through a 0.22 μm syringe filter. The resulting protein
solution was mixed with 3 mL of pre-equilibrated Ni-NTA resin
and incubated overnight at 4 °C while being stirred. The next
day, the mixture was transferred to a column at 4 °C. After the
flow-through had been collected, the column was washed with
TPGT buffer [20 mM Tris (pH 7.8), 500 mM KCl, 10% (w/v)
glycerol, and 1 mM TCEP] containing successively 20, 40, and
50 mM imidazole in sequence using 20, 2, and 2 column volumes
for each, respectively. Finally, the column was developed with
2 column volumes of TPGTbuffer containing 100mM imidazole,
and the eluate was collected in different fractions. After SDS−
PAGE analysis, imidazole was removed from the pooled pure
fractions by passage over a 10DG column (Bio-Rad) equilibrated
with BPGT buffer [20 mM Bis-Tris propane (pH 9.0), 200 mM
KCl, 10% (w/v) glycerol, and 1 mMTCEP]. Any apo-flavin sites
were then reconstituted by overnight incubation of the protein in
1 mMFAD at 4 °C. Excess flavin was removed by gel filtration on
a 10DG column (see above) prior to prompt use or flash freezing
in liquid nitrogen and storage at −80 °C.
Heterologous Overexpression and Purification of
Roseiflexus castenholzii FixX. Ro. castenholzii FixX was
overexpressed in E. coli using a pCDFDuet-1 vector modified
to include a C-terminal strep tag. Transformed cells were
grown in Luria-Bertani (LB) broth containing streptomycin
(50 μg/mL) at 37 °C and 250 rpm to an OD600 of 0.4−0.5.
To induce expression of the f ixX gene, IPTG (1.5 mM) was
added to the cell culture. Ammonium Fe(III) citrate (4 mM),
L-cysteine (2 mM), and sodium fumarate (10 mM) were also
added. Ammonium Fe(III) citrate and cysteine increased
iron−sulfur cluster occupancy, and sodium fumarate served as
an electron acceptor during anaerobic metabolism. The flasks
were sealed with rubber stoppers containing cannulas sparging
argon into the cell suspension and fixed with exhaust tubes
flowing into a water trap. Cells were incubated for 4−6 h at room
temperature and then anaerobically transferred to centrifugation
bottles in a Coy chamber (Coy Laboratories, Grass Lake, MI)
under a nitrogen atmosphere. Cells were harvested at 5000g for
10 min.
All purification steps were performed anaerobically under a
nitrogen or argon atmosphere. Wet cell pellets were suspended
in lysis buffer [50 mM Tris-HCl (pH 8), 150 mM NaCl, 1 mM
sodium dithionite (Na-dithionite), 5% glycerol, 1 μL/mL
supernatant of a supersaturated phenylmethanesulfonyl fluo-
ride (PMSF) solution, 1 mg/10 mL DNase, and 5 mg/10 mL
lysozyme] at a cell:buffer ratio of 1:5. Cells were lysed using a
cell bomb (Parr Bomb Instrument Co., Moline, IL) by slowly
increasing the pressure to 1500 psi and equilibrating for 30 min
before collecting the lysate. This process was repeated and the
supernatant collected by centrifugation at 18000g for 20 min.
FixX was purified using a single-step affinity column purification
using Strep-Tactin Superflow Plus (Qiagen, Hilden, Germany)
resin pre-equilibrated with wash buffer containing 50 mM
Tris-HCl (pH 8), 150 mMNaCl, and 1 mMNa-dithionite. Once
the protein was loaded, the column was washed until the baseline
absorbance returned. The protein was then eluted with 50 mM
Tris-HCl (pH 8), 150 mM NaCl, 1 mM Na-dithionite, and
2.5 mM D-desthiobiotin as a brown band and anaerobically
concentrated using an Amicon (EMDMillipore) centrifugal filter
with a membrane molecular mass cutoff of 10 kDa. The
concentrated protein was flash frozen with liquid nitrogen for
further analysis.
Heterologous Overexpression and Purification of
Rh. rubrum FixAB. FixAB from Rh. rubrum was overexpressed
and purified as described previously.26 Briefly, E. coli BL21 cells
containing plasmid pET101/d-TOPO::f ixAB26 were cultured
anaerobically in LB broth supplemented with 2.8 mM glucose,
17 mM KH2PO4, 72 mM K2HPO4, and 15 mg/L riboflavin at
20 °C until the early exponential growth phase was reached.
At this point, gene expression was induced via the addition of
0.5 mM IPTG, and cells were cultured under the same conditions
for an additional 12 h. Harvested cells were resuspended
in degassed lysis buffer [20 mM Tris-HCl (pH 7.6), 10 mM
imidazole, 500 mM NaCl, 5 mg/mL lysozyme, and 1 mg/mL
Dnase I] and submitted to three freeze−thaw cycles. The lysate
was added to a nickel-NTA column (Qiagen) that had been
equilibrated with degassed lysis buffer. The column was washed
with degassed lysis buffer, and FixAB was eluted into anaerobic
vials using a 10 to 300 mM imidazole gradient in degassed
20 mM Tris-HCl (pH 7.6) and 500 mM NaCl. Yellow fractions
indicating the presence of FixAB were subjected to buffer
exchange using a PD-10 column (GE-Healthcare, Little
Chalfont, U.K.) and stored in desalting buffer [20 mM
Tris-HCl (pH 7.6) and 100 mM NaCl]. Purified FixAB was
incubated overnight at 4 °C with 1 mM FAD in desalting buffer
to reconstitute FAD cofactors lost during purification.38,39
Unbound FAD was rinsed by concentrating the samples with
an Amicon membrane (cutoff 30 kDa) followed by dilution
with desalting buffer containing 0.5 μM FAD. Reconstituted
samples were stored at −20 °C. The purity and identity of the
fractions were assessed via SDS−PAGE and mass spectrometry,
respectively.
Dynamic Light Scattering.Dynamic light scattering (DLS)
was performed on a DynaPro NanoStar (Wyatt Technology,
Santa Barbara, CA) to determine the size distribution and
hydrodynamic radii of species in solution. Samples were filtered
through a 0.1 μm syringe filter prior to the experiments. DLS was
measured aerobically using 10 μL of buffer as the control and also
1 mg/mL enzyme using a disposable cyclic olefin copolymer
cuvette at 25 °C.
Protein Identification, Chemical Cross-Linking, and
Model Construction. Identification of protein from gel bands
and solution digestion was performed according to standard
protocols recommended by the manufacturers using a trypsin
(Promega, Madison, WI) protease:complex ratio of 1:50 to
1:100 overnight and pepsin (Sigma, St. Louis, MO)
protease:complex ratio of 1:10 for 60 s. Proteins were identified
as described previously5 using a maXis Impact UHR-QTOF
instrument (Bruker Daltonics, Billerica, MA) interfaced with a
Dionex 3000 nano-uHPLC instrument (Thermo-Fisher,
Waltham, MA) followed by data analysis in Peptide Shaker.40
Chemical cross-linking was performed using 20 μg of the
FixABCX complex and 1 mM bis(sulfosuccinimidyl)suberate
(BS3) (Thermo-Fisher) in 50 mM HEPES (pH 7.2), 150 mM
NaCl buffer at room temperature for 1 h. The reaction was
quenched by addition of 120 mMTris (final concentration). The
resulting mixture was separated by SDS−PAGE (4 to 20% linear
gradient gel, Bio-Rad) and stained with Coomassie Brilliant Blue
(Thermo-Fisher). Protein bands of interest were excised from
the gel and digested with trypsin as described above. For cross-
link mapping, a Spectrum Identification Machine (SIM) was
used.41 Precursor and fragment ion tolerances were set to
±20 ppm. Intact protein analysis was performed as described
previously using a Bruker Micro-TOF mass spectrometer
(Bruker Daltonics).42 Noncovalent mass spectrometry experi-
ments were conducted on a SYNAPTG2-Si instrument (Waters,
Milford, MA) in a fashion similar to that described previously.43
Briefly, the FixABCX complex sample was buffer exchanged to
200 mM ammonium acetate (pH 7) (Sigma) using 3 kDa
molecular mass cutoff spin filters (Pall Corp., Port Washington,
NY) and infused from in-house prepared gold-coated
borosilicate glass capillaries to the electrospray source at a
protein concentration of 2−3 μM and a rate of∼90 nL/min. The
instrument was tuned to enhance performance in the high
mass-to-charge range. The following settings were used: source
temperature, 30 °C; capillary voltage, 1.7 kV; trap bias voltage,
16 V; argon flow in the collision cell (trap), 7 mL/min. The
transfer collision energy was held at 10 V, while the trap energy
varied between 10 and 200 V. Data analysis was performed
in MassLynx version 4.1 (Waters).
Protein homology models were generated by Phyre2,44 and
energy-minimized models were docked using ClusPro2 with
restrictions derived from chemical cross-linking experi-
ments.45−48 The flavin and iron−sulfur cluster cofactors were
added using SwissDock49,50 for individual subunits and
eventually added as rigid bodies to the final FixABCX complex
model. Molecular graphics were created using the UCSF
Chimera package.51
ElectronParamagnetic ResonanceSpectroscopy.Electron
paramagnetic resonance (EPR) spectra were recorded with a
Bruker E-500 spectrometer (X-band, 9.38 Ghz) equipped with a
SHQ resonator (Bruker), an in-cavity cryogen free VT system
(ColdEdge Technologies, Allentown, PA), and a MercuryiTC
temperature controller (Oxford Instruments, Abingdon, U.K.).
Spin quantifications were performed by double integration of
the spectra after manual baseline subtraction in the OriginPro
software package and referenced to copper-triethylamine
standards (75−125 μM) measured under the same conditions.
To assist with spectral deconvolution and assignment of g factors
(±0.003), computer simulations of the experimental spectra
were performed in MatLab using the EasySpin package and
“esfit” fitting function incorporating g strain to replicate line
broadening.
Electron Bifurcation Assays. FixABCX electron bifurcation
activity was measured anaerobically in an ultraviolet−visible
(UV−vis) spectrophotometer (Varian Cary 50 Bio, Agilent
Technologies, Santa Clara, CA) using quartz cuvettes (d = 1 cm).
All assays were carried out in 50 mM HEPES (pH 7.5), 10%
glycerol, and 0.02% DDM and contained the following: 0.8 μM
FixABCX (1.7 nmol of flavin/nmol of FixABCX and 9.1 nmol
of Fe/nmol of FixABCX), 85 μM FldSq, 200 μM NADH,
and 300 μM coenzyme Q1 (CoQ1). NADH, Fld
Sq, and FldOx
were monitored at 340 nm (ε = 6.2 mM−1 cm−1), 580 nm
(ε = 5.7 mM−1 cm−1), and 450 nm (ε = 11.3 mM−1 cm−1),
respectively.12,52 The Fld used in the assays as the low-potential
electron acceptor was purified in the hydroquinone state as
previously described.5 FldHq was exposed to oxygen for a short
period of time, upon which the majority (>80%) of the FldHq
converted to the semiquinone form. The protein was then
degassed with argon, and the absorbance of the semiquinone
species was measured at 580 nm. The concentration was then
calculated using the extinction coefficient (ε = 5.7 mM−1 cm−1).
Thermodynamic Calculations. Standard reduction
midpoint potentials of the NAD+/NADH (Em = −320 mV),
CoQ/CoQH2 (Em = 10 mV), Fld
Ox/Sq (Em = −180 mV), and
FldSq/Hq (Em = −460 mV) half-reactions were converted to
standard Gibbs free energies (ΔG°′) with the equation
Δ °′ = − °′G nFE (2)
where n is the number of electrons (moles) and F is the Faraday
constant (96485.34 J V−1 mol−1) (Table S5). The standard Gibbs
free energy of reaction (ΔGrxn°′ , in joules per mole) can be
calculated for the reaction of interest using the equation
∑ ∑Δ ° = | |Δ ° − | |Δ °′ ′ ′G v G v Grxn i (products) i (reactants) (3)
where |vi| is the stoichiometric coefficient, i is the product or
reactant, and ΔG(products)°′ and ΔG(reactants)°′ are the standard Gibbs
free energy of the products and reactants (joules per mol),
respectively, calculated from eq 2.
Transient Absorption Spectroscopy.The ultrafast (100 fs
to 5.1 ns) transient absorption spectroscopy (TAS) spectrometer
employed in this study uses an amplified 4W Ti:sapphire laser
(Libra, Coherent, 800 nm, 1 kHz, 100 fs pulse width) and the
Helios spectrometer (Ultrafast Systems LLC, Sarasota, FL).
A fraction of the 800 nm Libra output was frequency-doubled
in beta barium borate (BBO) to produce the desired pump
wavelength (480 nm for the data described here) for sample
excitation, which was then directed into the Helios. The pump
pulses were passed through a depolarizer and chopped by
a synchronized chopper to 500 Hz before reaching the sample.
The pump pulse energy was 1.1 μJ/pulse at the sample. Another
fraction of the 800 nm Libra output was guided directly into the
Helios for generation of the probe. Within the spectrometer,
a white light continuum of wavelengths including 340−800 nm
was generated using a 2 mm thick CaF2 crystal. This beam was
split into a probe beam and a reference beam. The probe beam
was focused into the sample where it was overlapped with the
pump beam. The transmitted probe and reference beams were
then focused into optical fibers coupled to multichannel
spectrometers with CMOS sensors with 1 kHz detection
rates. The reference signal is used to correct the probe signal
for pulse-to-pulse fluctuations in the white light continuum.
The time delay between the pump and probe pulses was
controlled by a motorized delay stage. For all transient
absorption measurements, the sample was made in an Mbraun
glovebox (N2 atmosphere), sealed in a 2 mm quartz cuvette, and
constantly stirred to prevent photodegradation. Rh. rubrum
FixAB and A. vinelandii FixABCX concentrations were
approximately 150 and 43 μM, respectively, and they were
measured in their as-isolated state (mostly oxidized). The
Rh. rubrum dimer contained >0.7 nmol of flavin/nmol of FixAB,
and the A. vinelandii FixABCX complex contained 1.7 nmol
of flavin/nmol of FixABCX. For the purpose of this study, light
initiated the formation of the semiquinone intermediates for each
flavin through generation of the FAD excited state and donation
of electrons from nearby protein residues or the other flavins.
Qualitatively, this experiment shows which type of semiquinone
is formed for a particular FAD site and suggests how
thermodynamically stable that intermediate is, based on its
lifetime. All experiments were conducted at room temperature.
The change in the absorbance signal (ΔA) was calculated from
the intensities of signals detected after sequential probe pulses
with and without the pump pulse excitation. Data were collected
(350 pump shots per time point) three consecutive times and
then averaged. The experiment was repeated three times for
A. vinelandii FixABCX and once for Rh. rubrum FixAB. Data were
corrected for spectral chirp using SurfaceXplorer (Ultrafast
Systems LLC). ASQ signals were fit in Igor Pro with a double-
exponential function. The 550 nm emissive feature in Rh. rubrum
FixAB is due to stimulated emission.53
■ RESULTS AND DISCUSSION
Effect of f ix and rnf Deletions on Nitrogen Fixation in
A. vinelandii.To determine whether the Fix system is associated
with diazotrophic growth in A. vinelandii, as in R. palustris,
Rh. rubrum, and S. meliloti,28,29,31 a deletion mutant lacking
f ixABC was generated (Figure S1). This mutant, unlike those
of R. palustris, Rh. rubrum, and S. meliloti, exhibited equally robust
growth on solid media with and without added fixed nitrogen
(Figure 1). This nitrogen-fixing (Nif+) phenotype was also
observed when the A. vinelandii f ix mutant expressed the
vanadium and iron-only alternative nitrogenases or when
cultured under low-aeration conditions (data not shown).
Although the results appeared to indicate that Fix is not
associated with nitrogen fixation in A. vinelandii and were
consistent with a previous study by Wientjens,27 we also tested
the possibility of redundancy,54 thinking that an alternative
complex could participate in balancing the chemical energy and
reductant pools in the absence of Fix, masking the effect of the
Δf ixABC mutation.
Rnf (Rhodobacter nitrogen fixation) is a membrane-bound
complex found in some diazotrophs that, like Fix, has been
hypothesized to generate reductant in the form of Fd or Fld for
nitrogen fixation.16,55,56 Nitrogen-fixing organisms that have
genes encoding Rnf typically do not have genes encoding Fix
and vice versa. Interestingly, A. vinelandii is one of the few
known diazotrophs with genes encoding both complexes.57
The physiological implications of having both Rnf and Fix are not
altogether clear, but given that A. vinelandii fixes nitrogen under
highly oxic conditions, the combination of Rnf and Fix could
confer the ability to fine-tune the redox status of the cell to a high
degree under a wide range of oxygen tensions.
It has been suggested that Rnf uses the energy of the proton
motive force to generate reduced Fd/Fld15,16 while Fix uses
electron bifurcation.18,20 Neither the Rnf nor Fix complex,
however, has previously been directly implicated in the delivery
of electrons to nitrogenase. Curatti et al.32 showed that
Figure 1. Phenotype of A. vinelandii DJ wild type (A), Δf ix (B), Δrnf1
(C), Δf ixΔrnf1 (D), Δrnf 2 (E), and Δf ixΔrnf 2 (F) strains. Wild type
andmutant strains were cultivated in Burk’s medium supplemented with
ammonium acetate (+N) or with no added fixed nitrogen (−N). All
samples were grown aerobically.
A. vinelandii possesses two Rnf complexes, Rnf1 associated with
nitrogen fixation and Rnf2, which is expressed constitutively.
Δrnf1 mutants, although still able to grow diazotrophically
(Figure 1), consistently exhibit a long lag in nif gene expression
and diazotrophic growth during derepression studies.32 To test
whether Rnf and Fix might be compensating for one another,
we transformed Δrnf mutants of A. vinelandii with the Δf ix
construct (Figure S1)32 to generate double mutants of
A. vinelandii lacking either Rnf1 and Fix or Rnf2 and Fix. The
Δf ix-Δrnf1 A. vinelandii strain was able to grow when fixed
nitrogen was supplied, but no growth was observed under
nitrogen fixing conditions (Figure 1). Therefore, cells lacking
both Rnf1 and Fix displayed a non-nitrogen fixing (Nif−)
phenotype. Our data provide support for a role of both Fix and
Rnf1 in providing reducing equivalents to nitrogenase and
indicate that the two complexes have compensatory activities,
such that either one can replace the other to ensure electron flow.
Furthermore, this is the first study to link the Fix complex to
nitrogen fixation in A. vinelandii. The Δf ix-Δrnf 2 A. vinelandii
mutant control strain was able to grow with or without added
fixed nitrogen (Figure 1), confirming that Rnf1, but not Rnf2,
can support nitrogen fixation.
Characterization of the FixABCX Complex. To under-
stand how Fix can support nitrogen fixation, we characterized the
FixABCX complex and its activity in vitro. The f ix operon of
A. vinelandii consists of six genes, f ixFd (annotated as f ixP
in some literature), f ixA, f ixB, f ixC, f ixX, and Avin_10560
(designated ORF6) (Figure S2). FixAB is homologous to the
previously characterized EtfAB, which is part of the Etf-Bcd
bifurcating complex. FixCX is similar to Etf ubiquinone
oxidoreductase (Etf-QO), involved in the transfer of electrons
to the quinone pool in the membrane.29 On the basis of the
similarity to other Etf systems, it is hypothesized that the Fix
complex oxidizes NADH and bifurcates one electron to the high-
potential quinone pool (exergonic branch) and drives the other
electron to a low-potential acceptor in the form of Fd or Fld
(endergonic branch) (Figure 2).20
Most microbes with f ix genes have only f ixABCX;
homologues f ixFd and ORF6 have not been investigated in f ix
gene clusters of any other species thus far.27,58 Although FixFd
and ORF6 were not specifically addressed in this study, it is
thought that FixFd, a small iron−sulfur protein,27,59 may serve
as a low-potential acceptor in Fix electron bifurcation. ORF6 is
described as being ferritin-like, but studies of its function have
yet to be conducted.
Figure 2. (A) Proposed mechanism of FixABCX electron bifurcation inA. vinelandii, operating within the framework of the known oxidation−reduction
potentials of NADH, Fld/Fd, and CoQ. It is hypothesized that crossed potentials of the bifurcating flavin (a-FAD) promote electron bifurcation based
on the appearance of a short-lived ASQ. Once the first electron transfers into the exergonic branch, the second electron, sitting at a very low reduction
potential, is thermodynamically unstable and immediately transfers into the endergonic branch composed of the low-potential iron−sulfur clusters of
FixX. The exergonic branch is set up to ensure the delivery of electrons to c-FAD where electrons accumulate before being transferred to CoQ. The
specific oxidation−reduction potential levels are qualitative unless otherwise noted. (B) Electron transfer pathways illustrated in the FixABCX structural
model generated by docking (ClusPro2) four homology models (Phyre2) (Figure S8).
The f ix genes from A. vinelandii were homologously
overexpressed under control of the nifH promoter (Figure S2),
which regulates transcription of genes encoding molybdenum
nitrogenase. This approach allows gene(s) of interest to be
overexpressed upon removal of a fixed nitrogen source. On the
basis of the hypothesis that the FixABCX complex delivers
one electron per bifurcation reaction to the quinone pool
[coenzyme Q (CoQ)], it was anticipated that the enzyme could
be associated with the membrane fraction, as observed for
Etf-QO.60,61 Indeed, a previous study found that heterologously
overexpressed Rh. rubrum FixC was localized to the membrane
fraction in E. coli.29 Furthermore, an N-terminal lipophilic tail
and transmembrane helix were predicted on the FixC subunit
based on the amino acid sequence (SACS MEMSAT2 and Phyre
2 prediction software).44,62 Attempts to purify the FixABCX
complex confirmed membrane association. After a range of NaCl
concentrations (0−1 M) and a variety of non-ionic detergents
had been used to solubilize the complex, the detergent n-dodecyl
β-D-maltoside (DDM), used for many membrane proteins,61,63
was found to bemost effective for the isolation of intact FixABCX
in a soluble and active form.
Upon extraction of the FixABCX complex from the
membrane, the DDM-solubilized fraction was subjected to
anion-exchange and size-exclusion chromatography. SDS−PAGE
analysis of the purified FixABCX complex revealed four bands
corresponding in size to FixA, FixB, FixC, and FixX (Figure 3)
and was >80% homogeneous [defined by percent band (Image
Lab, Bio-Rad)]. Mass spectrometry confirmed the identities of
the four Fix subunits (Figure S3), and densitometry of the
stained gel indicated that the band intensities were consistent
with an A:B:C:X subunit stoichiometry of 1:1:1:1.
Dynamic Light Scattering (DLS). DLS was used to
determine the oligomeric state of the FixABCX complex on
the basis of particle size. The complex in buffer containing
0.02% DDM displayed an average hydrodynamic radius of
5.1 ± 0.3 nm, corresponding to an estimated molecular mass of
150 kDa versus the expected mass of 129 kDa for the FixABCX
heterotetramer (Figure S4). This slightly larger-than-expected
complex is consistent with attachment of DDM to the protein
tetramers. While there was also some contribution from larger
aggregates, which are very prominent in the DLS output because
of their much higher scattering efficiencies (≈12−16 nm), the
aggregates comprised <1% of the population (Figure S4). Buffer
containing DDM was used as a negative control and produced
micelles with a radius of ≈4.3 nm (Figure S4). Overall, the
DLS data suggest FixABCX exists as individual heterotetramers
in solution.
FixABCX Ligand Composition and Structural Model.
Multiple electron transfer cofactors, including flavins and iron−
sulfur clusters, have been identified in electron bifurcating
enzyme complexes, with a single flavin proposed to serve as the
site of bifurcation.17,18,20,25 On the basis of common sequence
motifs for ligand binding, FixABCX was predicted to have three
FAD moieties, one each in FixA, FixB, and FixC, and two
[4Fe-4S] clusters in FixX. Native mass spectrometry (MS) was
employed to reveal the subunit stoichiometry and ligand
composition of the complex.
All complex members were first identified on the basis of
fragments produced by two different proteases, trypsin and
pepsin. Additionally, proteolysis reactions revealed that FixABCX
and a small amount of Fld (NifF) are purified together. The next
step was to determine the molecular masses of the intact proteins.
Reverse phase LC−MS confirmed full-length FixA and that FixB
and FixC subunits did not contain N-terminal Met residues. The
purified FixABCX complex, under anaerobic conditions, was
analyzed using native MS conditions by direct infusion on a
Synapt G2-Si. Under standard conditions (60 V collision
energy), the complex dissociated into subcomplexes of FixAB
and FixCX. A consistent ion signal for the tetrameric FixABCX
complex could not be maintained despite the presence of all four
proteins and cofactors in the sample after size-based purification.
The FixAB component contained two FAD molecules and
closely matched the expected size (71414.8 Da vs a predicted
MW of 71413.7 Da) (Figure 4A,B), whereas the FixCX
subcomplex containing FAD and two [4Fe-4S] clusters was
slightly heavier than expected (60310.1 Da vs a predicted MW
of 59932.9 Da, FixCX) (Figure 4B). The additional mass of
377.2 Da matches very closely that of riboflavin (Rf) (MW of
376.4 Da), raising the intriguing possibility that it may also be
associated with the complex (Figure 4B).
The UV−vis spectrum of FixABCX revealed a distinct peak at
428 nm, as well as oxidized flavin signatures with shoulders near
460 and 365 nm [Figure 5 (black trace)]. After denaturation
of FixABCX and removal of the protein by centrifugation, the
soluble fraction demonstrated clear signatures of flavins with
broad maxima at 370 and 450 nm [Figure 5 (red trace)]. The
band at 428 nm is not a usual signature of flavins, and such a
band has not been observed in FixAB from other diazotrophs
that have been expressed in E. coli. Thus, it is likely that this
species is associated with FixC or FixX and may be covalently
attached to the protein, as it does not remain in the super-
natant when denatured protein is removed. The signatures
of the [4Fe-4S] clusters are not evident in as-purified FixABCX
but are not expected to be prominent because of scattering
and their broad and relatively weak signals (400−420 nm;
ε ≈ 2−4 mM−1 cm−1).64
Once we had established the subunit stoichiometry and ligand
composition of the FixABCX complex, we investigated the
quaternary interactions that define the active enzyme. Models for
each of the Fix subunits were generated using Phyre2,44 and the
Figure 3. SDS−PAGE gel of the purified (∼80%) FixABCX complex
fromA. vinelandii (extra lanes were removed). All Fix protein bands were
verified by mass spectrometry (Figure S3), and according to
densitometry, the subunit stoichiometry of FixABCX is 1:1:1:1.
figures of merit are provided in Figures S5−S8. To validate these
subunit models and to elucidate protein−protein interactions
within the FixABCX complex, chemical cross-linking was used.
The bis(sulfosuccinimidyl)suberate (BS3) cross-linking reagent
is homobifunctional, reacting with primary amines and hydroxyl
groups to form covalent bonds. Cross-linked samples were
separated using one-dimensional SDS−PAGE to confirm
connections between subunits and to enrich cross-linked species
in samples subsequently analyzed by liquid chromatography
tandem mass spectrometry (LC−MS/MS) (Figure 4C and
Table S3). Overall, the reaction of the FixABCX complex with
BS3 reagent produced more than 200 connections. For initial
model building of the FixABCX complex and to confirm the
subunit homology models, a reduced set of cross-links of very
high confidence were selected (probability score of >7; signif-
icance is >3) (Figure 4D).
The resulting model of the quaternary structure provides
insightful hypotheses to be tested in the next generation of
experiments (Figure 2B and Figure S8E). The MS-based model
places the cofactors in locations that suggest a pair of possible
paths for electron transfer, both emanating from the flavin
in FixA. One provides a plausible route to the [4Fe-4S] clusters
in FixX, and the other can provide a path to the site at which
quinone is found to be bound in the functional homologue of
FixC, the Etf-QO (Figure 2B and Figure S8E).65 While some
of the distances between cofactors are longer than ideal for
direct electron transfer, conformational changes could be
invoked to bring them sufficiently close,22,39,66 but we also
note that the model places conserved Trp and Tyr in locations
that could provide electron transfer paths between cofactors
(Figure S8E).67−69 Thus, our model provides testable hypoth-
eses for which amino acids are expected to alter complex stability
or internal electron transfer upon mutagenesis. These amino
acids will be targets of experiments to elucidate fundamental
elements of bifurcating activity and test attribution of bifurcating
activity to FixABCX.
EPRSpectroscopy.FixABCXwas analyzed by low-temperature
continuous wave X-band EPR spectroscopy to assess its flavin
Figure 4. A. vinelandii FixABCX complex composition. (A) Native mass spectrum of FixAB containing two FAD cofactors (71414.8 Da; calculated
MW of 71413.7 Da) generated during FixABCX complex activation in the gas phase. Red diamonds signify charge state envelope centered around a
charge of +18. The unmarked masses are the charge state envelope of the molecular chaperone DnaK. (B) Subcomplexes obtained during FixABCX
complex activation. (C) SDS−PAGE gel of the FixABCX complex (right) cross-linked with BS3 reagent (left). The most distinguished bands in the
cross-linking reaction, migrating around the 60, 70, and 88 kDa molecular weight marker, were identified as FixCX, FixAB, and FixBC dimers,
respectively. (D) Protein−protein interaction map based on the highest-scoring cross-links (score of ≥7; red and blue lines indicate intra- and
inter-cross-links, respectively). A complete list of generated cross-links can be found in Table S3.
Figure 5. UV−vis spectrum of the FixABCX complex from A. vinelandii
demonstrating the presence of flavin: black for as-purified FixABCX, red
for cofactors released upon denaturation of FixABCX, and blue for
flavins as observed in the spectrum of FixAB from R. palustris, vertically
offset by −0.05 AU for the sake of clarity. The two spectra representing
FixABCX were corrected for Raleigh scattering by fitting the baseline to
the equation for scattering and then subtracting the fit from the
measured spectrum to obtain a corrected spectrum. A. vinelandii
FixABCX was prepared in 50 mM Tris-HCl (pH 8), 150 mM NaCl,
and 0.02% (w/v) DDM. R. palustris FixAB was prepared in 20 mM
Bis-Tris propane (pH 9), 200 mM KCl, 10% (w/v) glycerol, and 1 mM
tris(2-carboxyethyl)phosphine (TCEP).
radical content and the identity of iron−sulfur clusters. The EPR
spectrum of the as-purified enzyme showed a very weak fast
relaxing, broad signal that was strongest near 5 K along with
a weak g ∼ 2 radical signal (Figure 6 and Table S4).
Upon reduction with either NADH or Na-dithionite, the broad
signal almost disappeared and was replaced with multiple signals
in the S = 1/2 region indicative of FeS clusters and a flavin radical.
Both treatments yielded overlapping rhombic g = 2.07 (g = 2.072,
1.940, 1.895) and axial g = 2.04 (g = 2.041, 1.944, 1.944) signals
consistent with the presence of two iron−sulfur clusters, in
addition to an isotropic g = 2.0 signal consistent with a flavin
radical that was dominant above 50 K (Figure S9). The
temperature dependencies of the rhombic g = 2.07 and axial
g = 2.04 signals were characteristic of [4Fe-4S] clusters,70 in
agreement with the sequence prediction for a pair of [4Fe-4S]
clusters. The optimal temperature (Topt) for the rhombic g = 2.07
signal was near 15 K with broadening above 30 K (Figure S9).
The axial g = 2.04 signal was faster relaxing with a lower Topt
between 5 and 10 K and broadening above 15 K. Also present for
the NADH-reduced sample was a highly temperature dependent
axial g = 2.03 signal (g = 2.030, 2.00, 2.00) observed at 10 K. The
signal appeared to correlate with a slight loss of intensity from the
axial g = 2.04 signal compared to the Na-dithionite treatment,
suggesting that this feature could arise from the spin interaction
between the faster relaxing [4Fe-4S] cluster and a flavin radical.
Further EPR analysis of the FixX subunit alone (from
Ro. castenholzii) identified very similar rhombic g = 2.07 and
axial g = 2.04 signals indicating that these signals in FixABCX
originate from the FixX subunit (Figure S10). Line broadening at
the high- and low-field edges of the reduced spectra was observed
for both FixABCX and FixX and is indicative of spin coupling
between the two [4Fe-4S] clusters.71 For FixABCX, the different
relative intensities of the rhombic g = 2.07 and axial g = 2.04
signals suggest that the corresponding [4Fe-4S] clusters have
different Em values. Accordingly, for FixABCX, both clusters
are thought to have Em values below that of Na-dithionite
[≤−440 mV vs SHE (pH 7.5)], because the sample appeared to
be only partially reduced by Na-dithionite. We hypothesize that
the [4Fe-4S] cluster giving rise to the axial g = 2.04 signal has a
lower Em, because of its weaker contribution to the spectrum,
based on simulations (Table S4).
Altogether, the two [4Fe-4S] cluster EPR signals show a
striking resemblance to the EPR signals assigned to the two
[4Fe-4S] clusters in the NADH-dependent ferredoxin:NADP+
oxidoreductase I (Nfn) bifurcating enzyme, as the signals share
similar g values, temperature dependence, and oxidation−
reduction properties.25 For the Nfn enzyme, structural and
biophysical analyses showed that two [4Fe-4S] clusters in the
large subunit form an electron transfer chain from the flavin site
of electron bifurcation to an external ferredoxin redox partner.
Spin coupling between the two clusters is thought to facilitate
electron transfer between the redox cofactors. It was also found
for the Nfn enzyme that one of the [4Fe-4S] clusters had an
unusually low Em that creates a thermodynamically favorable
electron transfer pathway from a highly unstable semiquinone
intermediate formed during electron bifurcation. An analogous
model is suggested here for FixABCX where the two low-
potential [4Fe-4S] clusters in the FixX subunit would form a
thermodynamically favorable pathway for facile electron transfer
between the site of bifurcation and Fd/Fld.
Electron Bifurcation by the FixABCX Complex. Electron
bifurcation by the FixABCX complex is proposed to drive the
endergonic reduction of FldSq by coupling it to the exergonic
reduction of CoQ using NADH as an electron donor (Figures 2
and 7A). Electron bifurcation by the FixABCX complex was
demonstrated by incubating FixABCX with the electron donor
NADH, high-potential acceptor coenzyme Q1 (CoQ1), and low-
potential acceptor FldSq. CoQ1 served as an amphipathic
analogue of the physiological acceptor coenzyme Q8 (CoQ8)
found in the electron transport chain of A. vinelandii.72,73 FldSq
was used as the low-potential acceptor in these assays because
FldHq has been shown to directly donate electrons to
nitrogenase.5,6 Similarly, the previously characterized electron
bifurcating Etf-Bcd complex could direct electrons to either Fd or
Fld,22,23 suggesting that Fld could likely serve as an electron
acceptor from FixABCX, as well.
For the electron bifurcation assay, the oxidation of NADH
(340 nm), disappearance of FldSq (580 nm), and formation
of FldOx (450 nm) were monitored over time. CoQ1 reduction
(λmax = 274 nm) was not recorded, because of significant
interference at that wavelength. While the simultaneous presence
of several redox active components in the assays posed a
challenge, monitoring activities at multiple wavelengths provided
evidence of electron bifurcation. Control reactions for NADH
oxidation (diaphorase) activity linked to CoQ1 reduction in the
absence of FldSq yielded a specific activity of 396 ± 44 nmol
min−1 mg−1 with a turnover frequency of 51± 6min−1 [Figure 7B
(No FldSq)]. Bifurcation reaction mixtures containing the
additional component needed for bifurcation (FldSq, in addition
to the control reactants FixABCX, NADH, and CoQ1) exhibited
25% greater NADH oxidation activity with a specific activity
of 524 ± 21 nmol min−1 mg−1 and a turnover frequency of
68 ± 3 min−1 [Figure 7B (All components)]. It is important to
note that the specific activity values are not adjusted for flavin
occupancy, because it could not be confirmed how much of the
Figure 6. EPR spectra of FixABCX fromA. vinelandii prepared in 50mM
HEPES (pH 7.5), 150 mMNaCl, and 5% glycerol: black for as-prepared
FixABCX (100 μM), blue for FixABCX (100 μM) reduced with NADH
(1 mM), and red for FixABCX (100 μM) reduced with sodium
dithionite (10 mM). Simulations of spectra of NADH- and
Na-dithionite-reduced FixABCX are shown in lighter colored traces.
Settings: microwave frequency, 9.38 GHz; microwave power, 1 mW;
modulation frequency, 100 kHz; modulation amplitude, 10.0 G; sample
temperature, 10 K.
protein was fully occupied (how flavins were distributed among
the flavin binding sites).
The Fld-dependent increase in the level of NADH
consumption is consistent with bifurcation, and previous studies
of the Etf-Bcd bifurcating complex also demonstrated an
increased level of NADH consumption in a bifurcating
reaction.22,23 Omission of FixABCX or CoQ1 resulted in baseline
NADH oxidation activity (Figure 7B). To decipher whether the
enhanced consumption of NADH in the presence of FldSq could
be attributed to the endergonic reduction of FldSq to FldHq
(the bifurcation reaction), the disappearance of FldSqwasmonitored
at 580 nm where optical absorption by the hydroquinone species
is negligible but absorbance by the semiquinone is strong
(ε = 5.7 mM−1 cm−1). Reactions monitoring the disappearance
of FldSq were difficult to interpret on their own, because of
background activity in the absence of FixABCX (Figure 7C).
In vivo, CoQ is in the membrane, whereas Fld is in the cytoplasm;
therefore, the extent of direct contact between them is
diminished, and spontaneous electron transfer between the
two species is suppressed (Table S5). In vitro experiments
however lack this separation, and our controls revealed that
FldSq can donate electrons to CoQ1 in the absence of FixABCX
(Figure 7C). Thus, the FldSq concentration decreased, while
an increase at 450 nm, characteristic of FldOx, was observed
(Figure 7D). This nonenzymatic reduction of CoQ1 by Fld
Sq
is consistent with the favorableΔG° of−36.7 kJ/mol (Table S5).
However, the increase in absorbance at 450 nm produced by
spontaneous FldSq oxidation enabled us to account for this
reaction that detracts from the apparent yield of the bifurcating
reaction [Figure 7C (All components)]. The FldOx species was
not detected in the bifurcating reaction assays, indicating that
FldHq, the expected product of the bifurcation reaction, was
formed instead (Figure 7D). However, we do not expect FldHq
to accumulate in the presence of CoQ1 because of the favorability
of electron transfer between these two that will return Fld to
its semiquinone state and thus conceal the true extent of
FldHq production (ΔG° = −154 kJ/mol) (eq 4 and Table S5).
+ + +
= + +
+
+
NADH 2Fld 2CoQ 3H
2CoQH 2Fld NAD
Hq
2
Sq
(4)
Overall, the absence of FldOx in conjunction with the increased
activity of the electron bifurcation reaction in comparison to
diaphorase activity suggests that the FixABCX complex can in
fact perform electron bifurcation. To further investigate the flavin
cofactors in FixABCX, transient absorption spectroscopy (TAS)
was conducted.
Transient Absorption Spectroscopy. A comparative TAS
study between FixAB from Rh. rubrum and FixABCX from
A. vinelandii was performed to gain qualitative information about
spectral contributions from individual flavins. TAS of the
incomplete Fix complex, FixAB, revealed an anionic semi-
quinone (ASQ) absorption at 365 nm with a corresponding
oxidized (Ox) flavin bleach at 447 nm. The ASQ absorption
decays with two components: a short-lived component with a
lifetime of tens of picoseconds and a longer short-lived component
with a lifetime of 1000 picoseconds [Figure 8A (red trace) and
Table S6]. A similar ASQ signal was also observed in the
Figure 7. Electron bifurcation by the FixABCX complex of A. vinelandii under anaerobic conditions. (A) Overview of electron bifurcation by the
FixABCX complex showing the NADH bifurcating an electron to the high-potential acceptor, CoQ1, and the other to the low-potential acceptor, Fld
Sq.
Evidence of electron bifurcation was obtained using UV−vis spectrophotometry. NADH oxidation as well as FldSq reduction and oxidation were
monitored over time as signatures of the bifurcation reaction. (B) NADH oxidation at 340 nm, (C) FldSq reduction/oxidation at 580 nm, and
(D) formation of FldOx at 450 nm. Final concentrations of components added to the reaction mixtures were as follows: 0.8 μMFixABCX, 85 μMFldSq,
200 μM NADH, and 300 μM CoQ1. Data were normalized and demonstrate the overall change in absorbance.
A. vinelandii FixABCX complex [Figure 8A (black trace)]. This
suggests that two ASQ species can be formed in the FixAB unit,
consistent with the presence of two flavins. Bifurcating flavins
have been shown to exhibit crossed potentials, whereby the
Em
Ox/SQ is at a potential lower than that of Em
SQ/HQ, the result
being that the high-energy SQ intermediate does not appreciably
accumulate relative to Ox and HQ.
A short-lived ASQ has been observed in the bifurcating Nfn
from Pyrococcus furiosus and is proposed to play a central role in
bifurcation.19,25 The current observation of relatively short-lived
ASQ signals in Rh. rubrum FixAB and A. vinelandii FixABCX is
consistent with the work on Nfn and suggests that neither flavin
significantly stabilizes a one-electron-reduced species. On the
basis of comparison with Etf, the flavin in FixA (a-FAD)
(analogous to the β-FAD in EtfB) is the proposed site of
bifurcation and likely corresponds to the component of tens of
picoseconds in our TAS experiments.22 This assignment then
implies that the FixB b-FAD acts as an electron transferring
flavin, consistent with a slightly longer-lived ASQ, because
accumulation of electrons at this site may congest electron flow
and impede bifurcation. This may additionally play a role in
gating electron transfer, ensuring that only one electron follows
the exergonic path, and thus restricting the lower-potential
electron to travel down the endergonic branch, similar to the
mechanism of electron bifurcation in P. furiosus Nfn.25 Because
of the crossed potentials believed to characterize bifurcating
flavins, the more negative potential electron of the bifurcating
flavin (ASQ to Ox transition) would provide enough driving
force to reduce the low-potential iron−sulfur clusters [predicted
to be ≤−440 mV vs SHE (pH 7.5)] in FixX directly. In the
complete FixABCX complex from A. vinelandii, an additional
peak is observed in the TAS corresponding to a neutral
semiquinone (NSQ) and is assigned to the c-FAD in FixC
(Figure 8B). We propose two electrons from successive
bifurcations accumulate in c-FAD to permit two-electron
reduction of CoQ.
ProposedMechanism for FixABCXElectronBifurcation.
The data presented herein establish that the FixABCX complex
from A. vinelandii can bifurcate electrons from NADH to CoQ1
and FldSq. The biochemical and biophysical evidence is
consistent with a proposed mechanism of Fix electron
bifurcation initiated at the flavin in FixA, which would accept a
pair of electrons from NADH and pass one electron to the
quinone pool via the flavins in FixB and FixC, and the other to
Fld/Fd via the low-potential [4Fe-4S] clusters in FixX, with the
energetic cost being paid by favorable transfer of the former
electron to the quinone pool (Figure 2).
TAS on the FixABCX complex provides evidence by
demonstrating the presence of a short-lived anionic flavin
semiquinone consistent with the ability to support rapid and
efficient electron transfer. The findings here provide exper-
imental support for the mechanism proposed previously for a
homologous bifurcating Etf that assigns the site of bifurcation to
the flavin bound by FixA (EtfB).22 This assignment is further
supported by the demonstration by Sato et al. that the flavin
reduced by NADH is the one bound in FixA (EtfB), but that
electron transfer to the flavin in FixB (EtfA) is favorable.39 Thus,
upon reduction of the FixA flavin, transfer of an electron through
the exergonic branch to CoQ via FixB and FixC can leave the
second electron in a very unstable, highly energetic flavin
semiquinone state with a low Em
Ox/Sq that can drive reduction of
Fld/Fd, via FixX (Figure 2).
The mechanism presented here offers a means by which
Fd/Fld can be reduced by NADH in a reaction that is
thermodynamically favorable overall and identifies a new
pathway by which low-potential reducing equivalents can be
generated to drive nitrogen fixation.
■ ASSOCIATED CONTENT
Primers for Δf ix mutant generation (Table S1), primers
for overexpression of FixABCX (Table S2), protein−
protein interactions captured within the FixABCX
complex during chemical cross-linking reaction (Table S3),
overview of the EPR signals observed for FixABCX from
A. vinelandii (Table S4), calculated Gibbs free energies of
possible reactions catalyzed by the Fix complex and
possible nonenzymatic reactions (Table S5), lifetimes of
ASQ species from Rh. rubrum FixAB and A. vinelandii
FixABCX (Table S6), scheme for Δf ix mutant generation
(Figure S1), scheme for overexpression of f ix genes in
A. vinelandii (Figure S2), protein identification within the
FixABCX complex purified from A. vinelandii (Figure S3),
DLS of the FixABCX complex (Figure S4), evaluation of
subunit homology models (Figure S5), predicted
Figure 8. Transient absorption spectra of as-prepared Rh. rubrum FixAB (red) and A. vinelandii FixABCX (black). (A) Kinetic traces of ASQ signal
(dots) at 365 nm. ASQ decay shows half-lives of ∼15 and ∼1000 ps when fit with a double-exponential function (solid lines). (B) An NSQ absorption
peak (∼565−650 nm) is observed only for the A. vinelandii FixABCX complex.
structural features in Fix subunits (Figure S6), FixB
protein interface (Figure S7), structural model of the
FixABCX complex from A. vinelandii (Figure S8), EPR
temperature profiles of FixABCX from A. vinelandii
(Figure S9), and comparison of EPR spectra of FixABCX
from A. vinelandii and the individual FixX subunit from
Ro. castenholzii (Figure S10) (PDF)
■ ACKNOWLEDGMENTS
The authors thank Stefan Nordlund at StockholmUniversity and
Tomas Edgren at Umea ̊ University for the plasmid containing
Rh. rubrum FixAB and their guidance in Fix protein expression
and purification and Luis Rubio for the A. vinelandii rnf mutants.
The authors also acknowledge Utah State University’s Synthetic
Biomanufacturing Institute for conducting large-scale fermenta-
tions and Montana State University’s Microfabrication Facility
for help in the preparation of gold-coated borosilica capillaries for
noncovalent mass spectrometry. TheMass Spectrometry Facility
at Montana State University is supported in part by the Murdock
Charitable Trust and National Institutes of Health IDEA
Program Grant P20GM103474.
■ ABBREVIATIONS
AEBSF, 4-(2-aminoethyl)benzenesulfonyl fluoride hydrochlor-
ide; AmpS, ampicillin sensitivity; ASQ, anionic semiquinone;
BBO, beta barium borate; BS3, bis(sulfosuccinimidyl)suberate;
CoQ, coenzyme Q; CoQ1, coenzyme Q1; CoQ8, coenzyme Q8;
DDM, dodecyl maltoside; DLS, dynamic light scattering; DTT,
dithiothreitol; Etf, electron transfer flavoprotein; Etf-Bcd,
electron transferring flavoprotein/butyryl-coenzyme A;
ETF-QO, electron transferring flavoprotein ubiquinone oxidor-
eductase; Em, midpoint potential; EPR, electron paramagnetic
resonance; FAD, flavin adenine dinucleotide; Fd, ferredoxin;
Fld, flavodoxin; FldOx, oxidized quinone; FldHq, flavodoxin
hydroquinone; FldSq, flavodoxin semiquinone; IPTG, isopropyl
β-D-1-thiogalactopyranoside; KmR, kanamycin resistance; LB, Luria-
Bertani; LC−MS/MS, liquid chromatography with tandem mass
spectrometry; MS, mass spectrometry; Na-dithionite, sodium
dithionite; Nif+, nitrogen-fixing phenotype; Nif−, non-nitrogen-
fixing phenotype; PCR, polymerase chain reaction; PMSF,
phenylmethanesulfonyl fluoride; Rf, riboflavin; Rnf, Rhodobacter
nitrogen fixation; SmR, streptomycin resistance; TAS, transient
absorption spectroscopy; TCEP, tris(2-carboxyethyl)phosphine;
TB, terrific broth; Topt, optimal temperature.
■ REFERENCES
(1) Kim, J., and Rees, D. C. (1994) Nitrogenase and biological nitrogen
fixation. Biochemistry 33, 389−397.
(2) Burgess, B. K., and Lowe, D. J. (1996) Mechanism of molybdenum
nitrogenase. Chem. Rev. 96, 2983−3012.
(3) Seefeldt, L. C., Hoffman, B.M., andDean, D. R. (2009)Mechanism
of Mo-dependent nitrogenase. Annu. Rev. Biochem. 78, 701−722.
(4) Saeki, K. (2004) Electron transport to nitrogenase: Diverse routes
for a common destination. InGenetics and Regulation of Nitrogen Fixation
in Free-living Bacteria (Klipp, W., Masepohl, B., Gallon, J. R., and
Newton, W. E., Eds.) pp 257−290, Kluwer Academic Publishers,
Dordrecht, The Netherlands.
(5) Yang, Z. Y., Ledbetter, R., Shaw, S., Pence, N., Tokmina-
Lukaszewska, M., Eilers, B., Guo, Q., Pokhrel, N., Cash, V. L., Dean, D.
R., Antony, E., Bothner, B., Peters, J. W., and Seefeldt, L. C. (2016)
Evidence that the Pi release event is the rate-limiting step in the
nitrogenase catalytic cycle. Biochemistry 55, 3625−3635.
(6) Martin, A. E., Burgess, B. K., Iismaa, S. E., Smartt, C. T., Jacobson,
M. R., and Dean, D. R. (1989) Construction and characterization of an
Azotobacter vinelandii strain with mutations in the genes encoding
flavodoxin and ferredoxin I. J. Bacteriol. 171, 3162−3167.
(7) Deistung, J., and Thorneley, R. N. (1986) Electron transfer to
nitrogenase: Characterization of flavodoxin from Azotobacter chroococ-
cum and comparison of its redox potentials with those of flavodoxins
from Azotobacter vinelandii and Klebsiella pneumoniae (nif F-gene
product). Biochem. J. 239, 69−75.
(8) Mortenson, L. E. (1964) Ferredoxin and ATP, requirements for
nitrogen fixation in cell-free extracts of Clostridium pasteurianum. Proc.
Natl. Acad. Sci. U. S. A. 52, 272−279.
(9) Yates, M. G. (1972) Electron transport to nitrogenase in
Azotobacter chroococcum: Azotobacter flavodoxin hydroquinone as an
electron donor. FEBS Lett. 27, 63−67.
(10) Thorneley, R. N., and Deistung, J. (1988) Electron-transfer
studies involving flavodoxin and a natural redox partner, the iron protein
of nitrogenase: Conformational constraints on protein-protein inter-
actions and the kinetics of electron transfer within the protein complex.
Biochem. J. 253, 587−595.
(11) Cammack, R. (1992) Iron-sulfur clusters in enzymes: Themes and
variations. In Advances in Inorganic Chemistry (Cammack, R., Ed.) pp
281−322, Academic Press, San Diego.
(12) Klugkist, J., Voorberg, J., Haaker, H., and Veeger, C. (1986)
Characterization of three different flavodoxins from Azotobacter
vinelandii. Eur. J. Biochem. 155, 33−40.
(13) Ludden, P. W. (1991) Energetics of and sources of energy for
biological nitrogen fixation. In Current Topics in Bioenergetics (Lee, C. P.,
Ed.) pp 369−390, Academic Press, San Diego.
(14) Shah, V. K., Stacey, G., and Brill, W. J. (1983) Electron transport
to nitrogenase. Purification and characterization of pyruvate:flavodoxin
oxidoreductase. The nif J gene product. J. Biol. Chem. 258, 12064−
12068.
(15) Saeki, K., and Kumagai, H. (1998) The rnf gene products in
Rhodobacter capsulatus play an essential role in nitrogen fixation during
anaerobic DMSO-dependent growth in the dark. Arch. Microbiol. 169,
464−467.
(16) Schmehl, M., Jahn, A., Meyer zu Vilsendorf, A., Hennecke, S.,
Masepohl, B., Schuppler, M., Marxer, M., Oelze, J., and Klipp, W. (1993)
Identification of a new class of nitrogen fixation genes in Rhodobacter
capsalatus: A putative membrane complex involved in electron transport
to nitrogenase. Mol. Gen. Genet. 241, 602−615.
(17) Buckel, W., and Thauer, R. K. (2013) Energy conservation via
electron bifurcating ferredoxin reduction and proton/Na+ translocating
ferredoxin oxidation. Biochim. Biophys. Acta, Bioenerg. 1827, 94−113.
(18) Peters, J. W., Miller, A. F., Jones, A. K., King, P.W., and Adams,M.
W. (2016) Electron bifurcation. Curr. Opin. Chem. Biol. 31, 146−152.
(19) Nitschke, W., and Russell, M. J. (2012) Redox bifurcations:
Mechanisms and importance to life now, and at its origin. BioEssays 34,
106−109.
(20) Herrmann, G., Jayamani, E., Mai, G., and Buckel, W. (2008)
Energy conservation via electron-transferring flavoprotein in anaerobic
bacteria. J. Bacteriol. 190, 784−791.
(21) Li, F., Hinderberger, J., Seedorf, H., Zhang, J., Buckel, W., and
Thauer, R. K. (2008) Coupled rerredoxin and crotonyl coenzyme A
(CoA) reduction with NADH catalyzed by the butyryl-CoA
dehydrogenase/Etf complex from Clostridium kluyveri. J. Bacteriol. 190,
843−850.
(22) Chowdhury, N. P., Mowafy, A. M., Demmer, J. K., Upadhyay, V.,
Koelzer, S., Jayamani, E., Kahnt, J., Hornung, M., Demmer, U., Ermler,
U., and Buckel, W. (2014) Studies on the mechanism of electron
bifurcation catalyzed by electron transferring flavoprotein (Etf) and
butyryl-CoA dehydrogenase (Bcd) of Acidaminococcus fermentans. J.
Biol. Chem. 289, 5145−5157.
(23) Chowdhury, N. P., Klomann, K., Seubert, A., and Buckel, W.
(2016) Reduction of flavodoxin by electron bifurcation and sodium ion-
dependent re-oxidation by NAD+ catalysed by ferredoxin:NA-
D+reductase (Rnf). J. Biol. Chem. 291, 11993−12002.
(24) Bertsch, J., Parthasarathy, A., Buckel, W., and Müller, V. (2013)
An electron-bifurcating caffeyl-CoA reductase. J. Biol. Chem. 288,
11304−11311.
(25) Lubner, C. E., Jennings, D. P., Mulder, D. W., Schut, G. J.,
Zadvornyy, O. A., Hoben, J., Tokmina-Lukaszewska, M., Berry, L.,
Nguyen, D., Lipscomb, G. L., Bothner, B., Jones, A. K., Miller, A. F.,
King, P. W., Adams, M. W. W., and Peters, J. W. (2017) Mechanistic
insights into energy conservation by flavin-based electron bifurcation.
Nat. Chem. Biol. 13, 655−659.
(26) Edgren, T. (2006) Electron transport to nitrogenase, Stockholm
University, Stockholm.
(27) Wientjens, R. (1993) The involvement of the fixABCX genes and
the respiratory chain in the electron transport to nitrogenase in
Azotobacter vinelandii. The Department of Chemistry and Biochemistry,
Agricultural University, Wageningen, The Netherlands.
(28) Huang, J. J., Heiniger, E. K., McKinlay, J. B., and Harwood, C. S.
(2010) Production of hydrogen gas from light and the inorganic
electron donor thiosulfate by Rhodopseudomonas palustris. Appl. Environ.
Microbiol. 76, 7717−7722.
(29) Edgren, T., and Nordlund, S. (2004) The f ixABCX genes in
Rhodospirillum rubrum encode a putative membrane complex
participating in electron transfer to nitrogenase. J. Bacteriol. 186,
2052−2060.
(30) Ruvkun, G. B., Sundaresan, V., and Ausubel, F. M. (1982)
Directed transposon Tn5 mutagenesis and complementation analysis of
Rhizobium meliloti symbiotic nitrogen fixation genes. Cell 29, 551−559.
(31) Earl, C. D., Ronson, C.W., and Ausubel, F. M. (1987)Genetic and
structural analysis of the Rhizobium meliloti f ixA, f ixB, f ixC, and f ixX
genes. J. Bacteriol. 169, 1127−1136.
(32) Curatti, L., Brown, C. S., Ludden, P. W., and Rubio, L. M. (2005)
Genes required for rapid expression of nitrogenase activity in
Azotobacter vinelandii. Proc. Natl. Acad. Sci. U. S. A. 102, 6291−6296.
(33) de Lorenzo, V., Herrero, M., Jakubzik, U., and Timmis, K. N.
(1990) Mini-Tn5 transposon derivatives for insertion mutagenesis,
promoter probing, and chromosomal insertion of cloned DNA in gram-
negative eubacteria. J. Bacteriol. 172, 6568−6572.
(34) Page, W. J., and von Tigerstrom, M. (1979) Optimal conditions
for transformation of Azotobacter vinelandii. J. Bacteriol. 139, 1058−
1061.
(35) Jacobson, M. R., Cash, V. L., Weiss, M. C., Laird, N. F., Newton,
W. E., and Dean, D. R. (1989) Biochemical and genetic analysis of the
nif USVWZM cluster from Azotobacter vinelandii. Mol. Gen. Genet. 219,
49−57.
(36) Toukdarian, A., and Kennedy, C. (1986) Regulation of nitrogen
metabolism in Azotobacter vinelandii: isolation of ntr and glnA genes and
construction of ntr mutants. EMBO J. 5, 399−407.
(37) Sarma, R., Barney, B. M., Hamilton, T. L., Jones, A., Seefeldt, L. C.,
and Peters, J. W. (2008) Crystal structure of the L protein ofRhodobacter
sphaeroides light-independent protochlorophyllide reductase with
MgADP bound: a homologue of the nitrogenase Fe protein.
Biochemistry 47, 13004−13015.
(38) Lewis, J. A., and Escalante-Semerena, J. C. (2006) The FAD-
dependent tricarballylate dehydrogenase (TcuA) enzyme of Salmonella
enterica converts tricarballylate into cis-aconitate. J. Bacteriol. 188, 5479−
5486.
(39) Sato, K., Nishina, Y., and Shiga, K. (2013) Interaction between
NADH and electron-transferring flavoprotein from Megasphaera
elsdenii. J. Biochem. 153, 565−572.
(40) Vaudel, M., Burkhart, J. M., Zahedi, R. P., Oveland, E., Berven, F.
S., Sickmann, A., Martens, L., and Barsnes, H. (2015) PeptideShaker
enables reanalysis of MS-derived proteomics data sets. Nat. Biotechnol.
33, 22−24.
(41) Lima, D. B., de Lima, T. B., Balbuena, T. S., Neves-Ferreira, A. G.
C., Barbosa, V. C., Gozzo, F. C., and Carvalho, P. C. (2015) SIM-XL: A
powerful and user-friendly tool for peptide cross-linking analysis. J.
Proteomics 129, 51−55.
(42) Poudel, S., Tokmina-Lukaszewska, M., Colman, D. R., Refai, M.,
Schut, G. J., King, P. W., Maness, P. C., Adams, M. W. W., Peters, J. W.,
Bothner, B., and Boyd, E. S. (2016) Unification of [FeFe]-hydrogenases
into three structural and functional groups. Biochim. Biophys. Acta, Gen.
Subj. 1860, 1910−1921.
(43) Luo, M. L., Jackson, R. N., Denny, S. R., Tokmina-Lukaszewska,
M., Maksimchuk, K. R., Lin, W., Bothner, B., Wiedenheft, B., and Beisel,
C. L. (2016) The CRISPR RNA-guided surveillance complex in
Escherichia coli accommodates extended RNA spacers. Nucleic Acids Res.
44, 7385−7394.
(44) Kelley, L. A., Mezulis, S., Yates, C. M., Wass, M. N., and Sternberg,
M. J. E. (2015) The Phyre2 web portal for protein modeling, prediction
and analysis. Nat. Protoc. 10, 845−858.
(45) Kozakov, D., Beglov, D., Bohnuud, T., Mottarella, S. E., Xia, B.,
Hall, D. R., and Vajda, S. (2013) How good is automated protein
docking? Proteins: Struct., Funct., Genet. 81, 2159−2166.
(46) Kozakov, D., Brenke, R., Comeau, S. R., and Vajda, S. (2006)
PIPER: an FFT-based protein docking programwith pairwise potentials.
Proteins: Struct., Funct., Genet. 65, 392−406.
(47) Comeau, S. R., Gatchell, D. W., Vajda, S., and Camacho, C. J.
(2004) ClusPro: an automated docking and discrimination method for
the prediction of protein complexes. Bioinformatics 20, 45−50.
(48) Comeau, S. R., Gatchell, D. W., Vajda, S., and Camacho, C. J.
(2004) ClusPro: a fully automated algorithm for protein-protein
docking. Nucleic Acids Res. 32, W96−99.
(49) Grosdidier, A., Zoete, V., and Michielin, O. (2011) SwissDock, a
protein-small molecule docking web service based on EADock DSS.
Nucleic Acids Res. 39, W270−277.
(50) Grosdidier, A., Zoete, V., and Michielin, O. (2011) Fast docking
using the CHARMM force field with EADockDSS. J. Comput. Chem. 32,
2149−2159.
(51) Pettersen, E. F., Goddard, T. D., Huang, C. C., Couch, G. S.,
Greenblatt, D. M., Meng, E. C., and Ferrin, T. E. (2004) UCSF
Chimera–a visualization system for exploratory research and analysis. J.
Comput. Chem. 25, 1605−1612.
(52) McComb, R. B., Bond, L. W., Burnett, R. W., Keech, R. C., and
Bowers, G. N. (1976) Determination of the molar absorptivity of
NADH. Clin. Chem. 22, 141−150.
(53) Enescu, M., Lindqvist, L., and Soep, B. (1998) Excited-state
dynamics of fully reduced flavins and flavoenzymes studied at
subpicosecond time resolution. Photochem. Photobiol. 68, 150−156.
(54) Wang, Z., and Zhang, J. (2009) Abundant indispensable
redundancies in cellular metabolic networks. Genome Biol. Evol. 1,
23−33.
(55) Jeong, H. S., and Jouanneau, Y. (2000) Enhanced nitrogenase
activity in strains of Rhodobacter capsulatus that overexpress the rnf
genes. J. Bacteriol. 182, 1208−1214.
(56) Sarkar, A., Köhler, J., Hurek, T., and Reinhold-Hurek, B. (2012) A
novel regulatory role of the Rnf complex of Azoarcus sp. strain BH72.
Mol. Microbiol. 83, 408−422.
(57) Boyd, E. S., Costas, A. M. G., Hamilton, T. L., Mus, F., and Peters,
J. W. (2015) Evolution of molybdenum nitrogenase during the
transition from anaerobic to aerobic metabolism. J. Bacteriol. 197,
1690−1699.
(58) Markowitz, V. M., Chen, I. M. A., Palaniappan, K., Chu, K., Szeto,
E., Grechkin, Y., Ratner, A., Jacob, B., Huang, J., Williams, P.,
Huntemann, M., Anderson, I., Mavromatis, K., Ivanova, N. N., and
Kyrpides, N. C. (2012) IMG: the Integrated Microbial Genomes
database and comparative analysis system. Nucleic Acids Res. 40, D115−
122.
(59) Reyntjens, B., Jollie, D. R., Stephens, P. J., Gao-Sheridan, H. S.,
and Burgess, B. K. (1997) Purification and characterization of a
f ixABCX-linked 2[4Fe-4S] ferredoxin from Azotobacter vinelandii. JBIC,
J. Biol. Inorg. Chem. 2, 595−602.
(60)Watmough, N. J., and Frerman, F. E. (2010) The electron transfer
flavoprotein: Ubiquinone oxidoreductases. Biochim. Biophys. Acta,
Bioenerg. 1797, 1910−1916.
(61) Usselman, R. J., Fielding, A. J., Frerman, F. E., Watmough, N. J.,
Eaton, G. R., and Eaton, S. S. (2008) Impact of mutations on the
midpoint potential of the [4Fe-4S]+1,+2 cluster and on catalytic activity in
electron transfer flavoprotein-ubiquinone oxidoreductase (ETF-QO).
Biochemistry 47, 92−100.
(62) Jones, D. T., Taylor, W. R., and Thornton, J. M. (1994) A model
recognition approach to the prediction of all-helical membrane protein
structure and topology. Biochemistry 33, 3038−3049.
(63) le Maire, M., Champeil, P., and Møller, J. V. (2000) Interaction of
membrane proteins and lipids with solubilizing detergents. Biochim.
Biophys. Acta, Biomembr. 1508, 86−111.
(64) Sweeney,W. V., and Rabinowitz, J. C. (1980) Proteins Containing
4Fe-4S Clusters: An Overview. Annu. Rev. Biochem. 49, 139−161.
(65) Zhang, J., Frerman, F. E., and Kim, J.-J. P. (2006) Structure of
electron transfer flavoprotein-ubiquinone oxidoreductase and electron
transfer to the mitochondrial ubiquinone pool. Proc. Natl. Acad. Sci. U. S.
A. 103, 16212−16217.
(66) Demmer, J. K., Rupprecht, F. A., Eisinger, M. L., Ermler, U., and
Langer, J. D. (2016) Ligand binding and conformational dynamics in a
flavin-based electron-bifurcating enzyme complex revealed by Hydro-
gen-Deuterium Exchange Mass Spectrometry. FEBS Lett. 590, 4472−
4479.
(67) Dryhurst, G., and Elving, P. J. (1969) Electrochemical oxidation-
reduction paths for pyrimidine, cytosine, purine and adenine:
Correlation and application. Talanta 16, 855−874.
(68) Bonetti, C., Stierl, M., Mathes, T., van Stokkum, I. H. M., Mullen,
K. M., Cohen-Stuart, T. A., van Grondelle, R., Hegemann, P., and
Kennis, J. T. M. (2009) The role of key amino acids in the
photoactivation pathway of the Synechocystis Slr1694 BLUF domain.
Biochemistry 48, 11458−11469.
(69) Gray, H. B., and Winkler, J. R. (2015) Hole hopping through
tyrosine/tryptophan chains protects proteins from oxidative damage.
Proc. Natl. Acad. Sci. U. S. A. 112, 10920−10925.
(70) Rupp, H., Rao, K. K., Hall, D. O., and Cammack, R. (1978)
Electron spin relaxation of iron-sulphur proteins studied by microwave
power saturation. Biochim. Biophys. Acta, Protein Struct. 537, 255−269.
(71)Mathews, R., Charlton, S., Sands, R. H., and Palmer, G. (1974)On
the nature of the spin coupling between the iron-sulfur clusters in the
eight-iron ferredoxins. J. Biol. Chem. 249, 4326−4328.
(72) Swank, R. T., and Burris, R. H. (1969) Restoration by ubiquinone
of Azotobacter vinelandii reduced nicotinamide adenine dinucleotide
oxidase activity. J. Bacteriol. 98, 311−313.
(73) Knowles, C. J., and Redfearn, E. R. (1969) Cytochrome and
ubiquinone patterns during growth of Azotobacter vinelandii. J. Bacteriol.
97, 756−760.