1. Biochemistry and Chemical Biology
  2. Structural Biology and Molecular Biophysics
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Structural basis of proton translocation and force generation in mitochondrial ATP synthase

  1. Niklas Klusch
  2. Bonnie J Murphy
  3. Deryck J Mills
  4. Özkan Yildiz
  5. Werner Kühlbrandt  Is a corresponding author
  1. Max Planck Institute of Biophysics, Germany
Research Article
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Cite as: eLife 2017;6:e33274 doi: 10.7554/eLife.33274

Abstract

ATP synthases produce ATP by rotary catalysis, powered by the electrochemical proton gradient across the membrane. Understanding this fundamental process requires an atomic model of the proton pathway. We determined the structure of an intact mitochondrial ATP synthase dimer by electron cryo-microscopy at near-atomic resolution. Charged and polar residues of the a-subunit stator define two aqueous channels, each spanning one half of the membrane. Passing through a conserved membrane-intrinsic helix hairpin, the lumenal channel protonates an acidic glutamate in the c-ring rotor. Upon ring rotation, the protonated glutamate encounters the matrix channel and deprotonates. An arginine between the two channels prevents proton leakage. The steep potential gradient over the sub-nm inter-channel distance exerts a force on the deprotonated glutamate, resulting in net directional rotation.

https://doi.org/10.7554/eLife.33274.001

Introduction

Mitochondrial ATP synthase uses the energy of the electrochemical proton gradient across the inner mitochondrial membrane to produce ATP from ADP and phosphate by rotary catalysis (Abrahams et al., 1994; Gresser et al., 1982). ATP synthases consist of the catalytic F1 head and the Fo subcomplex in the membrane (von Ballmoos et al., 2009). Rotation is driven by protons flowing down the membrane gradient through the Fo subcomplex. Understanding how this fundamental process generates rotary force requires an atomic model of the proton pathway. Until now, no high-resolution structure of an intact, functionally competent mitochondrial ATP synthase has been reported. The recent cryo-EM structure of the Fo subcomplex dimer isolated from yeast mitochondria (Guo et al., 2017) indicated the positions of key residues in the proton pathway. We have determined the structure of the complete mitochondrial ATP synthase dimer from the unicellular green alga Polytomella sp. Our structure reveals two prominent aqueous channels, each spanning one half of the membrane, that conduct protons to and from the conserved glutamates in the rotor ring. Protonation and deprotonation of these glutamates drives ring rotation and ATP synthesis.

Rotor rings of F-type ATP synthases consist of 8 (Watt et al., 2010; Zhou et al., 2015) to 15 (Pogoryelov et al., 2009) identical c-subunits that each form a hydrophobic helix hairpin. Mammalian mitochondria have a c8-ring, while yeasts (Hahn et al., 2016; Stock et al., 1999) and Polytomella (Allegretti et al., 2015) have 10 c-ring subunits, which we refer to as cA to cJ. A conserved glutamate (cGlu111 in Polytomella) serves as the c-subunit proton-binding site (Meier et al., 2005; Pogoryelov et al., 2009). The previous 6.2 Å cryo-EM map of the Polytomella ATP synthase dimer indicated two long, membrane-intrinsic helix hairpins in subunit a (Allegretti et al., 2015), but did not resolve sidechains. The helix hairpins run roughly at right angles to the c-ring helices. The longest helix bends around the c-ring, positioning the strictly conserved aArg239 and other key subunit a residues next to the c-subunit protonation site. Subsequent structures of F-type (Guo et al., 2017; Hahn et al., 2016; Morales-Rios et al., 2015) and V-type ATPases (Mazhab-Jafari et al., 2016) at 3.7 to 7 Å resolution have shown that the long membrane-intrinsic helix hairpins are a conserved and apparently essential feature of all rotary ATPases (Kühlbrandt and Davies, 2016), but the reason for this was not understood until now. Two proton channels were proposed to provide access to the c-ring protonation sites (Vik and Antonio, 1994) and first observed in the 6.2 Å Polytomella structure (Allegretti et al., 2015). Like the a-subunit helix hairpins themselves, the channels appear to be conserved in all rotary ATPases (Kühlbrandt and Davies, 2016).

Results and discussion

Structure determination and atomic model

We performed single-particle electron cryo-microscopy (cryo-EM) on ATP synthase dimers of the colourless unicellular alga Polytomella sp. to reveal the proton translocation pathway in atomic detail. With a molecular mass of ~1.6 MDa and its bulky peripheral stalk of subunits ASA 1-9 (ATP synthase associated proteins 1–9) (Figure 1; Figure 1—figure supplement 1) (Vázquez-Acevedo et al., 2006), the robust, V-shaped Polytomella dimer is well suited to high-resolution cryo-EM.

Figure 1 with 3 supplements see all
Cryo-EM structure of the Polytomella sp.

F1 Fo ATP synthase dimer at 4.1 Å resolution. Subunit a, blue; c-ring, yellow; ASA 6, brick; other subunits, cyan; detergent micelle, grey.

https://doi.org/10.7554/eLife.33274.002

A total of 90,142 particle images (Figure 1—figure supplement 2A) from 9,518 movies recorded with a direct electron detector in counting mode were aligned and refined to yield a map at 4.1 Å resolution (Table 1; Figure 1—figure supplement 2B), in which most sidechains of the ~14,000 residue dimer are visible (Figure 1; Video 1), except in the F1 heads and central stalks, where blending of multiple rotational states reduces map definition. Subvolume masking of the bulky peripheral stalk and associated Fo subunits improved the local resolution to 3.7 Å (Figure 1—figure supplement 2C). The best-defined region of the assembly includes subunit a, the c-ring interface and ASA 6, which was built de novo (Figure 2; Figure 2—figure supplement 1).

Figure 2 with 1 supplement see all
3.7 Å map of Polytomella Fo subcomplex with fitted atomic models.

(A) Side view with fitted subunit a, trans-membrane helices of ASA 6 and c-ring. (B) Subunit a and ASA 6 seen from the c-ring. (C) subunit a, c-ring and ASA 6 seen from the matrix. Subunit a, blue; c-ring, yellow; ASA 6, brick; other subunits, light cyan.

https://doi.org/10.7554/eLife.33274.006
Table 1
Cryo-EM data collection parameters, image processing, and refinement statistics.
https://doi.org/10.7554/eLife.33274.008
Data collection
 Electron MicroscopeJEOL JEM-3200FSC
 CameraK2 Summit
 Voltage300 kV
 Energy filter slit width20 eV
 Nominal Magnification30,000 x
 Calibrated physical pixel size1.12 Å
 Pixel size after mag. distortion corr.1.105 Å
 Total exposure82.5 e-2
 Exposure rate11.5 e-/(pixel x s)
 Number of frames45
 Defocus range−0.4 to −5 μm (95% between -0.9 and -2.5 µm)
Image Processing
 Motion correction softwareUnblur/MotionCor2 (with mag. distortion corr.)
 CTF estimation softwareCTFFind4, Gctf (for per-particle CTF)
 Particle selection softwaree2boxer (EMAN2)
 Micrographs used9,518
 Particles selected117,281
 3D map classification and refinement softwareRelion2
 Particles contributing to final map90,142
 Applied symmetryC2
 Global resolution (FSC = 0.143)3.68 Å
 Applied B-factor-125 Å
Model Building
 Modeling softwareCoot
 Refinement softwarePhenix (phenix.real_space_refine)
 Number of residues built1,039
 RMS (bonds)0.01 Å
 RMS (angles)1.09°
 Ramachandran outliers0.0%
 Ramachandran favoured94.88%
 Rotamer outliers0.27%
 Clashscore7.5
 EMRinger score1.42
Video 1
Three-dimensional map of the 1.6 MDa mitochondrial ATP synthase dimer from Polytomella sp.

showing the two c-rings (yellow), subunits a (blue) and ASA 6 (brick). The remaining eight ASA subunits in the peripheral stalks, the central stalks and catalytic F1 heads are shown in transparent cyan.

https://doi.org/10.7554/eLife.33274.009

Sequences of functionally important subunit a regions are highly conserved (Figure 1—figure supplement 3) and likely to have similar structures. In yeasts and mammals, subunit a is mitochondrially-encoded (known as ATP6 in human mitochondria), whereas in Chlamydomonas reinhardtii and, presumably, its close relative Polytomella, it is nuclear-encoded (Funes et al., 2002). N-terminal sequencing indicates that Polytomella subunit a has a 94-residue mitochondrial targeting sequence (Vázquez-Acevedo et al., 2006). The polypeptide forms a total of six α-helices H1 to H6. The mostly hydrophobic residues of the mature gene product are clearly resolved (Figure 2—figure supplement 1), except for the 11-residue loop connecting H4 and H5. The N-terminal H1 on the matrix-facing membrane surface is amphipathic, whereas in fungal and mammalian a-subunits, the first helix crosses the membrane (Guo et al., 2017; Hahn et al., 2016; Zhou et al., 2015). The four-helix bundle of hairpins H3/H4 and H5/H6 is immersed in the hydrophobic membrane interior. The 52 residues of H5 include the essential (Mitome et al., 2010) aArg239 and seven other charged or polar sidechains, interspersed with hydrophobic residues in a striking, strongly conserved pattern (Figure 1—figure supplement 3). Our structure puts these residues into a functional context of proton translocation and force generation.

The lumenal channel conducts protons to the c-ring rotor through a helix hairpin in the membrane

Two prominent aqueous channels span half of the Fo assembly, one from each side of the membrane (Figures 3,4). The lumenal channel enables proton access to cGlu111 from the crista lumen. Its entrance is a 23 by 37 Å funnel between the c-ring, the loop connecting the membrane-intrinsic H3/H4 hairpin, and the two trans-membrane helices of subunit ASA 6 (Figure 3A,C,D). Although there is no detectable sequence homology, ASA 6 appears to take the place of the peripheral stalk subunit b in yeasts and mammals (Guo et al., 2017; Hahn et al., 2016; Zhou et al., 2015). Four lipid acyl chains and densities that accommodate two phosphatidyl head groups are resolved at the rim of the lumenal funnel (Figure 4—figure supplement 2A), consistent with a cardiolipin molecule mediating close contacts between ASA 6 and subunit a in this position.

Two aqueous channels in Fo.

(A) Lumenal channel seen from the crista lumen. (B) Matrix channel seen from the matrix. (C) Side view of both channels seen from the c-ring, with outer c-ring helices in transparent yellow. Lumenal channel, left; matrix channel, right. The strictly conserved aArg239 in H5 separates the lumenal and matrix channels. (D) The lumenal channel passes through the H5/H6 hairpin at the small sidechains aAla246, aGly247 (H5) and aAla292 (H6) (green). (E) H4, the N-terminal half of H5 and the connecting H4/H5 loop at the matrix channel. Subunit a, blue; c10-ring, yellow; ASA 6, brick. Channels are shown as potential surfaces (red, negative; blue, positive; grey, neutral). (A) and (B) display a 5 Å slice of the c10-ring at the level of the protonated cGlu111.

https://doi.org/10.7554/eLife.33274.010

In the protein interior, the channel is lined by conserved charged, polar and hydrophobic sidechains (Figure 4; Figure 4—figure supplement 3A; Videos 2 and 3). The lumenal channel extends to a cluster of buried, closely spaced glutamates and histidines (aGlu172, aHis248, aHis252, aGlu288; Figure 4A; Figure 5A; Videos 2 and 3) that appears to serve as a local reservoir for protons to be fed to the c-ring glutamates. At aGlu288 about 20 Å below the lumenal membrane surface, the channel narrows to 4 by 5 Å and changes direction by 90° towards the c-ring (Figure 3D; Figure 4A). In the hydrophobic membrane interior, the strictly conserved polar sidechains of aAsn243 (H5) and aGln295 (H6) that face one another would stabilise the H5/H6 hairpin (Figure 5B). The channel passes through the hairpin at the small, conserved sidechains of aAla246, aGly247 (H5) and aAla292 (H6) (Figure 3C,D; Figure 4A; Figure 5B; Figure 1—figure supplement 3). Bulky hydrophobic sidechains close by on H5 and H6 keep the helices apart and the channel open (Figure 4A; Figure 5B). From aGlu288 the proton may jump to either of two c-ring glutamates. The path to cAGlu111 is 4 Å longer than the ~12 Å path to cBGlu111 (Figure 5C) but includes the hydrophilic sidechains of aAsn243 and cBSer112. Therefore, in our static structure the path to cAGlu111 is more favourable for proton transfer. In a rotating c-ring, the estimated minimum distance from aGlu288 to cGlu111 is ~11 Å. Note that these distance estimates are subject to an error margin of 3 to 4 Å, since the carboxyl groups of the glutamate sidechains are not visible in the map, as is usual in high-resolution cryo-EM structures due to radiation damage (Allegretti et al., 2014).

Figure 4 with 3 supplements see all
Proton pathway through the Fo subcomplex.

(A) In the lumenal channel, protons (red arrow) pass via the local proton reservoir of aGlu172, aHis248, aHis252 and aGlu288 (dashed red ellipse) through the H5/H6 helix hairpin at the small sidechains of aAla246, aGly247 (H5) and aAla292 (H6) (green) to cGlu111 in the rotor ring c-subunits (red circles). (B) aArg239 (blue circle) is located halfway between the lumenal channel on the left and the matrix channel on the right, forming a seal to prevent proton leakage. c-ring helices (transparent yellow) with cGlu111 are seen in the foreground. (C) In the matrix channel, protons (dashed red arrow) can pass straight from the deprotonated cGlu111 to the pH 8 matrix. Subunit a, blue; adjacent c-ring helices, transparent yellow; aqueous channels, translucent grey; residues in stick representation. Figure 4—figure supplement 1 shows the fitted model together with the map density in stereo.

https://doi.org/10.7554/eLife.33274.011
Figure 5 with 1 supplement see all
3.7 Å map of functionally important a-subunit residues with fitted atomic model.

(A) Proton reservoir formed by aGlu172, aHis248, aHis252, aGlu288 (dashed red ellipse) in the lumenal channel; (B) Interaction of aAsn243 (H5) and aGln295 (H6) stabilises the H5/H6 hairpin. The space between aGlu288 (H6) and aAsn243 (H5) marks the lumenal channel (dashed red ellipse). (C) Protons in the lumenal channel can pass from aGlu288 to cGlu111 of c-subunit A near aArg239 (H5) via cSer112 and aAsn243. (D) aTrp189 (H4), aThr193 (H4) and aTyr229 (H5) act as wedges between H4 and the N-terminal end of H5, forming two sides of a triangle.

https://doi.org/10.7554/eLife.33274.015
Video 2
Three-dimensional arrangement of subunit a (blue), c-ring (yellow) with lumenal channel in pink and matrix channel in light blue.
https://doi.org/10.7554/eLife.33274.017
Video 3
Arrangement of channel-lining sidechains for the lumenal channel.

Sidechains in stick representation are coloured as: subunit a, blue; c-ring, yellow; ASA 6, brick; lipids, grey. Channels are shown as potential surfaces (red, negative; blue, positive; grey, neutral).

https://doi.org/10.7554/eLife.33274.018

The strictly conserved aArg239 is positioned halfway between the matrix and lumenal channels (Figures 3A–C,4; Video 2), forming a positively charged seal that prevents proton leakage from the lumen to the matrix. The map density for this arginine sidechain is particularly well defined (Figure 2—figure supplement 1). Its position and orientation do not suggest a salt bridge with the deprotonated c-ring glutamate, which would impede ring rotation. aAsn243, cSer112, the protonated cGlu111 and the two aqueous channels provide a local hydrophilic environment. The shortest distance between the lumenal and matrix channels on either side of aArg239 is ~6 Å.

The matrix channel at the subunit a/c interface forms the proton exit pathway

The matrix channel (Figure 3B,C,E, 4C; Figure 4—figure supplement 3B; Videos 2 and 4) is defined by the N-terminal half of H5 (residues 221–239), the C-terminal half of H6 (301–312) and residues cTyr102 to cGlu111 of the outer helices of c-subunits A and J. Its deepest point is a 4 Å by 7 Å cavity next to cJGlu111, ~25 Å below the c-ring surface. The channel widens to 7 by 13 Å at aArg232 (H5) and cJLeu104 and continues straight to the 36 by 30 Å exit funnel on the matrix side. H4 and H5 describe two sides of a triangle that is wedged open by the conserved aromatic sidechains of aTrp189 (H4) and aTyr229 (H5), which forms a hydrogen bond to aThr193 (H4) (Figure 5D). The aromatic wedge of aTrp189 and aTyr229 induces a change in direction of H5 to follow the curvature of the c-ring. Like the lumenal channel, the matrix channel includes conserved charged and polar sidechains (Figure 4C; Figure 1—figure supplement 3; Figure 4—figure supplement 3B), notably aGlu225 (H5) and aGlu309 (H6), which form a salt bridge with aArg232 (H5). At a distance of 7.4 Å, aGlu225 is in a good position to receive protons from cGlu111. The protons pass via aGlu309 and aHis312 near the channel exit into the matrix.

Video 4
Arrangement of channel-lining sidechains for the matrix channel.

Sidechains in stick representation are coloured as: subunit a, blue; c-ring, yellow; ASA 6, brick. Channels are shown as potential surfaces (red, negative; blue, positive; grey, neutral).

https://doi.org/10.7554/eLife.33274.019

The relevance of residues at the matrix channel to human health is highlighted by a number of mutations in ATP6, the a-subunit of human ATP synthase, that result in severe and, at present, incurable diseases. Several of these mutations map to H4, H5 and H6 (Figure 5—figure supplement 1). Sequence comparison (Figure 1—figure supplement 3) indicates that functionally important subunit a residues are conserved in ATP6. A change of atp6Leu156 (aLeu236 in Polytomella) to Arg or Pro reduces ATP production by 70%. Both mutations result in Maternally Inherited Leigh Syndrome (MILS) or in Neuropathy, Ataxia and Retinitis Pigmentosa (NARP) Syndrome (Cortés-Hernández et al., 2007; Holt et al., 1990; Kucharczyk et al., 2009). aLeu236 marks the point where H5 bends around the c-ring. A change of the nearby Trp (aTrp189 in Polytomella) in H4 to arginine causes Bilateral Striatal Lesions (De Meirleir et al., 1995). aTrp189 appears to be crucial for keeping H4, H5 and H6 apart and the matrix channel open. Replacing these residues by an arginine or proline would disrupt the interaction of H5 with the c-ring rotor, impairing proton translocation and ATP synthesis. Mutations of atp6Leu217, atp6Leu220 and atp6Leu222 (aLeu302, aVal305 and aVal307 in Polytomella) on H6 in the same region of the long helix hairpin also result in Leigh Syndrome and reduced ATPase activity (Castagna et al., 2007; Moslemi et al., 2005; Thyagarajan et al., 1995) (Figure 5—figure supplement 1). Our structure thus provides direct new insights into the cause of serious mitochondrial diseases.

Mechanism and energetics of ring rotation

Rotation of the c-ring is driven by the proton-motive force (pmf) across the inner mitochondrial membrane, which has a chemical component (ΔpH ≅ 0.8 units, equivalent to ~50 mV) and an electrostatic component (ΔΨ ≅ 150 mV). The higher concentration of protons acts to protonate c-subunit A in the lumenal channel, whereas in the matrix channel, the incoming protonated c-subunit J would lose its proton to the pH 8 matrix (Figure 3A,B; Figure 6; Video 2). Protonation in the lumenal channel neutralises the residue and favours the buried sidechain conformation observed in x-ray structures of isolated c-rings (Pogoryelov et al., 2010). This would allow c-subunit A to move to position B, where it partitions into the hydrophobic environment of the membrane by counter-clockwise rotation, as seen from the matrix. At the same time, deprotonation of c-subunit J in the matrix channel renders its glutamate negatively charged, attracting it to the positively charged aArg239 halfway between the channels.

In a minimal model of rotary ATPases, torque is generated simply by stochastic bidirectional movement of the rotor, biased by the free energy inherent in the pmf (Junge et al., 1997). Other authors propose that the electrostatic field between the channels acts on the negatively charged glutamate, causing the ring to rotate (Miller et al., 2013). The atomic coordinates of our structure allow us to quantify this field. In a vacuum, a potential difference of 200 mV over the minimum 6 Å distance between the channels would generate a local electrostatic field of 330 million V/m, in a direction parallel to the membrane plane. Given a cGlu111-cGlu111 c-ring diameter of 4.2 nm, this electrostatic field would exert a torque of 110 pN nm on the single negative charge of the deprotonated cGlu111. The local dielectric of the protein reduces this value by more than 50%, in good agreement with the experimentally determined torque generated by the F1 head of E. coli ATPase in ATP hydrolysis mode of 40 to 60 pN nm (Kinosita et al., 2000; Pänke et al., 2001; Spetzler et al., 2006). Considering that, on the molecular scale, catalysis in the ATP synthase must be reversible, the estimated torque generated by the transverse electrostatic field between the two aqueous channels is in the expected range for ATP synthesis, though this does not exclude the possibility that thermal energy plays a role in overcoming activation barriers to rotation. Note that, without the channels, the electrostatic field across the ~35 Å hydrophobic membrane core would be about six times weaker and in the wrong direction (perpendicular to the membrane plane), and hence not able to drive the production of ATP by rotary catalysis. Our structure, in particular the small distance between channels, supports the notion that the force generated by the electrostatic field acting on the deprotonated cGlu111 gives rise to directional c-ring rotation. The structure of the Polytomella Fo ATP synthase thus explains how electrochemical energy is converted into the mechanical torque that powers ATP synthesis, as one of the most fundamental life processes. Theoretical studies, made possible by the atomic coordinates that are now available, will be necessary to evaluate the energetics of ATP synthesis in detail.

The hydrophobic membrane environment (circular arrow in Figure 6) disfavours movement of negatively charged, deprotonated c-ring subunits via the longer of two possible routes between the channels. Conversely, the short route past aArg239 at the a/c interface must discriminate against the passage of a protonated c-ring subunit. Failure to do so would result in proton leakage and dissipation of the pmf. aArg239 is bound to play a key role in this process. Conceivably, the positive charge on the flexible tether of the arginine sidechain associates with the deprotonated cGlu111 on its passage between the aqueous channels, smoothing the energy profile of c-ring rotation.

c-ring rotation is powered by the potential gradient between the lumenal channel (pink) and matrix channel (light blue).

The c-ring (yellow) and the membrane-intrinsic four-helix bundle of subunit a (blue) drawn to scale as seen from the matrix. Protons (red) pass from the crista lumen below the projection plane through the lumenal channel between H5 and H6 to protonate cGlu111 of c-subunit A, while c-subunit J is deprotonated by the higher pH of the matrix channel. The positively charged aArg239 is likely to interact with the deprotonated cGlu111 during its short passage to the lumenal channel. The lumenal and matrix channels approach one another to within 5–7 Å. A pmf of 200 mV between the closely spaced channels creates a local electrostatic field in the range of 40 million to 100 million V/m, depending on the protein dielectric. The field exerts a force on the deprotonated cGlu111 that results in net counter-clockwise rotation of the c-ring (grey arrow). Scale bar, 10 Å.

https://doi.org/10.7554/eLife.33274.020

Conclusion

We determined the structure of a 1.6 MDa mitochondrial F1Fo ATP synthase dimer by single-particle electron cryo-microscopy. At 3.7 Å resolution, all Fo subunits, helices, loops, most sidechains and some lipids are well resolved. Two prominent aqueous channels defined by subunit a and the 10-subunit c-ring rotor conduct protons to protonate and deprotonate a c-subunit glutamate in the middle of the membrane. Protons enter from the ~pH 7.2 crista lumen through the wide funnel-like opening of the lumenal channel that is lined by conserved polar or charged subunit a residues. A cluster of glutamates and histidines in the channel about 20 Å below the membrane surface serves as a local proton reservoir. At this point, the channel narrows and turns by 90° towards the c-ring, passing between the long, membrane-intrinsic subunit a helices H5 and H6. The channel ends at cGlu111 of the proximal c-ring subunit. cGlu111 is protonated and partitions into the hydrophobic membrane environment. Upon a ~320° revolution of the c-ring, the protonated cGlu111 encounters the aqueous matrix channel defined by a triangle of the membrane intrinsic helices H4, H5 and H6, and the proton escapes to the pH 8 matrix. The positively charged, strictly conserved aArg239 in H5 separates the lumenal and matrix channels, preventing proton leakage. The sub-nm distance between the aqueous channels results in a steep potential gradient, acting on the deprotonated c-ring glutamate to generate net directional rotation. The structure explains the fundamental process that drives ATP synthesis in all forms of life. Our atomic model will be essential in evaluating the energetics of proton translocation and force generation in ATP synthases.

Materials and methods

Cultures of Polytomella sp. (198.80, E.G. Pringsheim) from the alga collection at the Sammlung Algenkulturen Göttingen, Germany were grown aerobically with agitation at room temperature (23 ± 2˚C) in MAP medium (Atteia et al., 2000). Mitochondria and mitochondrial ATP synthase dimers were isolated as described (Allegretti et al., 2015; van Lis et al., 2005) with modifications. Briefly, mitochondrial membranes (50 mg) were resuspended in solubilisation buffer (10 mM Tris-HCl, pH 8.0, 1 mM MgCl2, 50 mM NaCl, 2% (w/v) n-dodecyl-β-D-maltoside (DDM)) in a total volume of 5 ml. After 30 min at 4°C, unsolubilised material was removed by centrifugation at 20,000 x g for 15 min at 4°C and the supernatant was loaded onto a POROS GoPure HQ column equilibrated in buffer A (10 mM Tris-HCl, pH 8.0, 1 mM MgCl2, 50 mM NaCl and 0.015% (w/v) DDM) on an Äkta purifier (GE Healthcare). The column was washed with buffer A + 100 mM NaCl and ATP synthase dimers were eluted with a linear gradient of 100 mM to 300 mM NaCl in buffer A. Fractions containing ATP synthase dimers were pooled, concentrated to 50 µl in Vivaspin 500 columns with 100,000 molecular weight cutoff and loaded onto a Superose-6 size exclusion column (PC 3.2/30) equilibrated in buffer B (10 mM Tris-HCl, pH 8.0, 1 mM MgCl2, 20 mM NaCl, 0.05% (w/v) DDM) on an Ettan purifier (GE Healthcare). Fractions containing ATP synthase dimers were collected. 250 µM of the substrate analogue AMP-PNP, 5 µM ADP and 0.02% (w/v) sodium azide were added 30 min before freezing to inhibit rotation.

For cryo-EM, 3 µl aliquots of a 1.5 mg/ml inhibited ATP synthase dimer solution were applied to C-flat-MH-4C grids with tobacco mosaic virus (TMV) for spreading, or to Quantifoil R2/2 grids covered with self-perforating hydrogel nanomembranes without TMV (Scherr et al., 2017). Grids were plunge-frozen in a Vitrobot (FEI) at 70% humidity, 10°C with 5.5 s (CF-MH-4C) or 9 s (hydrogel membranes) blotting time.

Images were acquired in electron counting mode with a K2 Summit direct electron detector on a JEM-3200FSC field emission cryo-TEM (JEOL, Tokyo) with in-column electron energy filter (slit width 20 eV) at 300kV. The nominal magnification was 30,000x, resulting in a specimen pixel size of 1.12 Å. 45-frame dose-fractionation movies were recorded manually at a defocus range of −0.4 to −5.0 µm (with 95% of micrographs being in the range −0.9 to −2.5 µm) with 0.2 s per frame at an electron flux of 11.5 e-/pixel/s.

Specimen movement between movie frames was corrected using Unblur (Brilot et al., 2012), followed by MotionCor2 with magnification distortion correction (Zheng et al., 2017), resulting in a corrected pixel size of 1.105 Å. Combining these two programs gave better results than either program alone, as judged by quality of a 3D reconstruction for a subset of the micrographs. ATP synthase dimers were picked manually with e2boxer. Micrograph CTF parameters were calculated using CTFFind4.1.5 (Mindell and Grigorieff, 2003), and per-particle defocus values were refined using gctf (Zhang, 2016). Further processing was carried out in Relion2.0 (Kimanius et al., 2016; Scheres, 2012). Dimer images were extracted and refined to a 40 Å lowpass-filtered reference with a soft mask surrounding the entire dimer. Particle polishing was carried out, and polished images were classified in 3D. Four of five classes (90,142 of 117,281 particles) were selected and combined for further refinement with a soft mask surrounding the dimer, yielding an overall resolution of 4.14 Å. Focussed refinement using a mask around the peripheral stalk and Fo region gave a better-resolved map of this region (3.68 Å resolution). The map was sharpened using a B-factor of −125 Å2 and low-pass filtered to 3.68 Å with the automated post-processing utility. Local resolution was calculated with the LocalRes utility in Relion.

The quality of the cryo-EM map of the Polytomella ATP synthase dimer enabled manual de novo modelling of all subunits. For the c-ring rotor an initial model was based on the structure of a c10-ring in the proton-unlocked state at pH 8.3 (pdb code 3U2F). Each c-subunit was fitted as a rigid body in UCSF Chimera (Goddard et al., 2007). The model was fitted and built manually in Coot (Emsley and Cowtan, 2004) with additional rounds of real-space refinement in PHENIX (Adams et al., 2010) (Table 1). The final model was converted into a density map using the molmap command in UCSF Chimera to calculate the map-to-model FSC (Figure 1—figure supplement 2C). Water-accessible channels were traced with the program Hollow (Ho and Gruswitz, 2008).

Data availability

The cryo-EM map has been deposited to the Electron Microscopy Data Bank (accession number EMD-4176). Atomic models have been deposited to the Protein Data Bank (accession number 6F36).

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Decision letter

  1. Sjors HW Scheres
    Reviewing Editor; MRC Laboratory of Molecular Biology, United Kingdom

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "Structural basis of proton translocation and force generation in mitochondrial ATP synthase" for consideration by eLife. Your article has been favorably evaluated by John Kuriyan (Senior Editor) and three reviewers, one of whom, Sjors HW Scheres (Reviewer #1), is a member of our Board of Reviewing Editors. The following individual involved in review of your submission has agreed to reveal their identity: Henning Stahlberg (Reviewer #3).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

This paper describes a cryo-EM structure to an overall resolution of 4.1A of the intact FoF1 ATP synthase dimer from the alga Polytomella sp. Focussed refinement on the dimeric Fo region yielded a map to 3.7A resolution, in which an atomic model could be built. The manuscript focuses on the description of this part of the complex. After a description of the yeast Fo dimer by another group, this manuscript claims to be the first describing the Fo dimer in a functionally competent dimer. The manuscript describes the proton pathway, with detailed descriptions of the half-channels on either side of the c-ring, and provides a mechanism of ring rotation. All three reviewers agreed that this paper represents an exciting step forwards in understanding these molecular machines (with one reviewer envisioning seeing it in future textbooks) and all three reviewers recommended publication.

The following points should be addressed in a revised version:

1) Atomic model validation metrics: this paper needs map-to-model FSC analyses, a molprobity report that includes a clashscore, Ramachandran statistics, and an EMRINGER quantification of rotamer placement. The paper doesn't mention any model refinement approach, but at this resolution, that should improve the statistics, so a model refinement should probably be included in the paper.

2) When describing the mechanism of ring rotation, the authors claim that the field across the sub-nanometer distance between the half-channels "would exert a significant force […], resulting in a net directional ring rotation", but provide no data on how "significant" this force would be. How large will this force be? How significant is it compared to the alternative model of stochastic bidirectional movement? What is missing here is a specifically stated conclusion about which of the two models (stochastic bidirectional movement or potential gradient force) is most likely to be correct.

3) Many of the figures that used a mesh for the cryoEM density, especially the stereo-images, were too busy for easy interpretation. A transparent isosurface, or removal of the density altogether (perhaps while showing zoomed in regions separately in supplementary figures?) may be easier for readers to digest, especially with all of the residue labels.

4) The readability for the wider readership of eLife may be improved by adding a figure with an overview sketch, showing the enzyme subunits, their names, and where they are placed in the entire dimeric structure. Something along the lines of Figure 6 in this manuscript http://www.plantphysiol.org/content/144/2/1190.long would do it.

Please find the individual reviewers' comments below. It would probably be beneficial to the paper if most of the other comments are also incorporated, but provided the ones above are addressed, the manuscript will most likely be ready for acceptance.

Reviewer #1:

This paper describes a cryo-EM structure to an overall resolution of 4.1A of the intact FoF1 ATP synthase dimer from the alga Polytomella sp. Focussed refinement on the dimeric Fo region yielded a map to 3.7A resolution, in which an atomic model could be built. The manuscript focuses on the description of this part of the complex. After a description of the yeast Fo dimer by another group, this manuscript claims to be the first describing the Fo dimer in a functionally competent dimer. The manuscript describes the proton pathway, with detailed descriptions of the half-channels on either side of the c-ring, and provides a mechanism of ring rotation. Overall, the findings are interesting and worthy of publication in eLife, provided the following criticism is taken into account:

In the cover letter the authors describe that the F1 heads adopt multiple conformations and that more data is needed. This statement seems to conflict with the one that most side chains in the dimer are visible. It would be better to include a statement in the main text on the heterogeneity in the F1 heads to explain why the manuscript focuses on the Fo monomer structure.

Wouldn't it be interesting to also comment on the dimerisation interface of the Fo subunits? Both in the membrane and in the stalk region? The yeast manuscript describes these interfaces in detail. Does this interface change in this functionally competent dimer? In general, it would be interesting to know what are the differences and similarities with the yeast structure.

When describing the mechanism of ring rotation, the authors claim that the field across the sub-nanometer distance between the half-channels "would exert a significant force […], resulting in a net directional ring rotation", but provide no data on how "significant" this force would be. How large will this force be? How significant is it compared to the alternative model of stochastic bidirectional movement?

The atomic model was built and fitted manually in Coot. However, at this resolution, it should also be refined (e.g. using Phenix or Refmac). This will improve the fit to the density, and the resulting statistics about the fit (please insert FSC model-vs-map curves) and the geometry (please include MolProbity and perhaps EM-ringer stats) will provide the reader with a means to assess its quality.

Reviewer #2:

The report by Klusch et al. is an exciting step forward in our understanding of the mechanism of proton motive force-driven rotary ATP catalysis that I would like to see published without undue delay. I can easily imagine seeing their Figure 6 reproduced in textbooks in the near future. The key message of this paper concerns the path of protons through F1F0 complex of a green alga to drive rotation by directional protonation and deprotonation of glutamate residues found in the c-ring. The proton path depends on the fascinating arrangement of a helical hairpin found in the α subunit stator. In addition, these authors identify a critical Arg residue "seal" that prevents leaking between adjacent aqueous paths and provide new insight into how the complex dimerizes to bend the cristae membrane. Of course, similar insights were recently described in Guo et al. (Science Oct 26th) based on a similar structure of the yeast F1F0 complex. Therefore, I hope to see this paper published in 2017 if possible, provided the authors can redress some deficiencies:

1) Atomic model validation metrics.

I would like to see map-to-model FSC analyses, a molprobity report that includes a clashscore, Ramachandran statistics, and an EMRINGER quantification of rotamer placement.

Reviewer #3:

The authors describe the 3.7A structure of an entire F-ATPase dimer. Their high resolution allows resolving the aqueous proton access channels through the a-subunit, and quantitatively explain the pathway of protons onto the c-ring, and which forces are involved in driving the enzyme.

The manuscript is nicely written, has very impressive data and findings, and is an important progress in the field of F-ATPase. The figures are clear, beautiful, and contribute significantly to the understandability of the manuscript.

However, manuscript may be a bit specialized for the general life sciences readership of eLife. To make the Results section and the Discussion more understandable to the general audience, the authors should include an overview sketch, showing the enzyme subunits, their names, and where they are placed in the entire dimeric structure. Something along the lines of Figure 6 in this manuscript http://www.plantphysiol.org/content/144/2/1190.long would do it.

In the last paragraph of the subsection “Structure determination and atomic model”, the authors first mention "the 11-residuel loop connecting H4 and H5", and only later introduce the a-subunit as being composed of six alpha-helices termed H1 to H6. The order here should be inverted.

In the subsection “Mechanism of ring rotation”, the authors discuss two different rotation models. In model 1 by Junge et al. 1997 of a stochastic bidirectional movement that is biased towards one direction by a proton-motive force (i.e., chemical plus electric gradient force), which is also called a ratchet motor, I believe. In model 2 by Miller et al., 2013, a net electrostatic field pulls a negatively charged glutamate forward, cause direct rotation, which may be called a power stroke.

The authors then calculate that the electric potential of 200mV over the short distance between the two aqueous channels onto the C-ring would correspond to 100'000'000 Volt/m perpendicularly to the membrane normal, and that this would exert a significant force, resulting in net directional rotation. What is missing here is a specifically stated conclusion. I interpret the author's statement as that they argue to have proven Model 2 above. However, this statement is missing.

Also, the force by 100e6 V/m on one charge with the given C-ring radius would have to be compared to the force of Brownian motion on the entire C-ring, in order to be able to state that a purely electrostatic power stroke mechanism drives the ring, and not biased Brownian motion. A quantitative comparison, and a clear final statement are missing.

In the Materials and methods section, the authors describe that they first used Unblur, and then followed by MotionCor2 with magnification distortion correction, which changed the pixel size from 1.12 to 1.105 A. This is a very unusual motion correction, which should accumulate interpolation errors from the two drift correction runs. The authors could add one sentence, explaining why they used two consecutive motion correction procedures, and why they believe the iterative drift correction was required.

https://doi.org/10.7554/eLife.33274.026

Author response

The following points should be addressed in a revised version:

1) Atomic model validation metrics: this paper needs map-to-model FSC analyses, a molprobity report that includes a clashscore, Ramachandran statistics, and an EMRINGER quantification of rotamer placement. The paper doesn't mention any model refinement approach, but at this resolution, that should improve the statistics, so a model refinement should probably be included in the paper.

The model-to-map FSC is now included in Figure 1—figure supplement 2C. We uploaded the PDB validation report and EM map deposition as supplementary information in the eLife submission system. The clashscore, Ramachandran statistics and EMRinger score are now shown in the new Table 1 that summarizes the cryoEM data acquisition, data processing and model statistics. The final model has been refined using PHENIX (phenix.real_space_refine). The procedure is described in the Materials and methods section of the revised manuscript.

2) When describing the mechanism of ring rotation, the authors claim that the field across the sub-nanometer distance between the half-channels "would exert a significant force […], resulting in a net directional ring rotation", but provide no data on how "significant" this force would be. How large will this force be? How significant is it compared to the alternative model of stochastic bidirectional movement? What is missing here is a specifically stated conclusion about which of the two models (stochastic bidirectional movement or potential gradient force) is most likely to be correct.

We are happy to include a quantitative estimate of the force exerted on the negatively charged c-ring glutamate, as requested by the reviewers. The “significant force” in vacuum would be 53 pN. At the 2.1 nm radius of the glutamate sidechain, the resulting torque is 112 pN nm. Since the protein dielectric would reduce the local field strength to less than 50% of that in vacuum, the net torque is somewhere around 50 pN nm, in remarkably good agreement with the torque generated by the E. coli F1 ATPase, determined experimentally as 40 to 60 pN nm by three independent groups. This is now explained in the revised Discussion subsection “Mechanism and energetics of ring rotation” (second paragraph), where the three papers reporting the experimentally determined F1 torque are cited. Since the basic mechanisms of torque generation and ATP synthesis are highly conserved and essentially identical in rotary ATPases of mitochondria, chloroplasts and bacteria, torque measurements made with the E. coli ATP synthase are relevant for the mitochondrial ATP synthase as well.

The stochastic bidirectional model does not predict an explicit force or torque acting on the c-ring glutamates. As the two models are not mutually exclusive, it would be wrong to say that one of them is incorrect. Even if the ring moves back and forth stochastically, the local electrostatic field across the channels we describe would provide a strong bias towards ring rotation in the experimentally observed counter-clockwise direction (as seen from the matrix) in ATP synthesis mode.

3) Many of the figures that used a mesh for the cryoEM density, especially the stereo-images, were too busy for easy interpretation. A transparent isosurface, or removal of the density altogether (perhaps while showing zoomed in regions separately in supplementary figures?) may be easier for readers to digest, especially with all of the residue labels.

The 3D structure of the Fo ATP synthase is fairly complex and not easy to display in two-dimensional figures. We tried to do our best but realized the result was not perfect. Presumably, anyone interested in the details of the structure and mechanism will look at the three-dimensional map and model with a suitable molecular graphics program, once the pdb coordinates are released, which they will be on acceptance of the manuscript. Nevertheless we have simplified Figure 4 as requested, which is now non-stereo and the map has been omitted (except for the channel surfaces). The original figure with the map is now Figure 4—figure supplement 1. A supplementary movie for Figure 4—figure supplement 3 was added (Video 3 and Video 4).

4) The readability for the wider readership of eLife may be improved by adding a figure with an overview sketch, showing the enzyme subunits, their names, and where they are placed in the entire dimeric structure. Something along the lines of Figure 6 in this manuscript http://www.plantphysiol.org/content/144/2/1190.long would do it.

Thank you for this suggestion. The new overview sketch is Figure 1—figure supplement 1.

Please find the individual reviewers' comments below. It would probably be beneficial to the paper if most of the other comments are also incorporated, but provided the ones above are addressed, the manuscript will most likely be ready for acceptance.

Reviewer #1:

[…] Overall, the findings are interesting and worthy of publication in eLife, provided the following criticism is taken into account:

In the cover letter the authors describe that the F1 heads adopt multiple conformations and that more data is needed. This statement seems to conflict with the one that most side chains in the dimer are visible. It would be better to include a statement in the main text on the heterogeneity in the F1 heads to explain why the manuscript focuses on the Fo monomer structure.

Good point. We have expanded first sentence of the Results: “… in which most sidechains of the ~14,000 residue dimer are visible (Figure 1; Video 1), except in the F1 heads and central stalk, where blending of multiple rotational states reduces map definition.”

Wouldn't it be interesting to also comment on the dimerisation interface of the Fo subunits? Both in the membrane and in the stalk region? The yeast manuscript describes these interfaces in detail. Does this interface change in this functionally competent dimer? In general, it would be interesting to know what are the differences and similarities with the yeast structure.

It would indeed be interesting, but since the dimerization interfaces of the yeast and Polytomella dimers are completely different, a detailed comparison of the dimer interface would go well beyond the scope of the present manuscript and delay its publication unreasonably. With the exception of subunit a and the c-ring, none of the yeast Fo and peripheral stalk subunits are even remotely similar. This is remarkable in itself, since the key mechanism of torque generation appears to be completely conserved. These and other important differences, such as the rotary position of the c-ring relative to subunit a, will be the subject of another paper to be submitted in due course.

When describing the mechanism of ring rotation, the authors claim that the field across the sub-nanometer distance between the half-channels "would exert a significant force […], resulting in a net directional ring rotation", but provide no data on how "significant" this force would be. How large will this force be? How significant is it compared to the alternative model of stochastic bidirectional movement?

See detailed response to comment 2 above.

The atomic model was built and fitted manually in Coot. However, at this resolution, it should also be refined (e.g. using Phenix or Refmac). This will improve the fit to the density, and the resulting statistics about the fit (please insert FSC model-vs-map curves) and the geometry (please include MolProbity and perhaps EM-ringer stats) will provide the reader with a means to assess its quality.

See detailed response to comment 1 above. The model has been refined and a table of refinement statistics added (Table 1). The model-vs-map FSC has been added to Figure 1—figure supplement 2.

Reviewer #2:

[…] I hope to see this paper published in 2017 if possible, provided the authors can redress some deficiencies:

1) Atomic model validation metrics.

I would like to see map-to-model FSC analyses, a molprobity report that includes a clashscore, Ramachandran statistics, and an EMRINGER quantification of rotamer placement.

See detailed response to comment 1 and corresponding comment of Reviewer 1 above.

Reviewer #3:

[…] However, manuscript may be a bit specialized for the general life sciences readership of eLife. To make the Results section and the Discussion more understandable to the general audience, the authors should include an overview sketch, showing the enzyme subunits, their names, and where they are placed in the entire dimeric structure. Something along the lines of Figure 6 in this manuscript http://www.plantphysiol.org/content/144/2/1190.long would do it.

Thank you for this suggestion. We have added a schematic overview of the Polytomella ATP synthase dimer as Figure 1—figure supplement 1.

In the last paragraph of the subsection “Structure determination and atomic model”, the authors first mention "the 11-residuel loop connecting H4 and H5", and only later introduce the a-subunit as being composed of six alpha-helices termed H1 to H6. The order here should be inverted.

Done, thank you.

In the subsection “Mechanism of ring rotation”, the authors discuss two different rotation models. In model 1 by Junge et al. 1997 of a stochastic bidirectional movement that is biased towards one direction by a proton-motive force (i.e., chemical plus electric gradient force), which is also called a ratchet motor, I believe. In model 2 by Miller et al., 2013, a net electrostatic field pulls a negatively charged glutamate forward, cause direct rotation, which may be called a power stroke.

The authors then calculate that the electric potential of 200mV over the short distance between the two aqueous channels onto the C-ring would correspond to 100'000'000 Volt/m perpendicularly to the membrane normal, and that this would exert a significant force, resulting in net directional rotation. What is missing here is a specifically stated conclusion. I interpret the author's statement as that they argue to have proven Model 2 above. However, this statement is missing.

Also, the force by 100e6 V/m on one charge with the given C-ring radius would have to be compared to the force of Brownian motion on the entire C-ring, in order to be able to state that a purely electrostatic power stroke mechanism drives the ring, and not biased Brownian motion. A quantitative comparison, and a clear final statement are missing.

See detailed response to comment 2 above.

In the Materials and methods section, the authors describe that they first used Unblur, and then followed by MotionCor2 with magnification distortion correction, which changed the pixel size from 1.12 to 1.105 A. This is a very unusual motion correction, which should accumulate interpolation errors from the two drift correction runs. The authors could add one sentence, explaining why they used two consecutive motion correction procedures, and why they believe the iterative drift correction was required.

A sentence to this effect has been added to the Materials and methods of the revised manuscript: “Combining these two programs gave better results than either program alone, as judged by quality of a 3D reconstruction for a subset of the micrographs.”

https://doi.org/10.7554/eLife.33274.027

Article and author information

Author details

  1. Niklas Klusch

    Department of Structural Biology, Max Planck Institute of Biophysics, Frankfurt, Germany
    Contribution
    Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing—original draft, Grew Polytomella cultures and isolated ATP synthase dimers, Prepared cryo-EM specimens and recorded image data, Built the atomic model, Analysed the structure, Drew the figures
    Contributed equally with
    Bonnie J Murphy
    Competing interests
    No competing interests declared
  2. Bonnie J Murphy

    Department of Structural Biology, Max Planck Institute of Biophysics, Frankfurt, Germany
    Contribution
    Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing—original draft, Prepared cryo-EM specimens and recorded image data, Processed image data and generated the 3D map, Built the atomic model, Analysed the structure
    Contributed equally with
    Niklas Klusch
    Competing interests
    No competing interests declared
    ORCID icon 0000-0001-6341-9368
  3. Deryck J Mills

    Department of Structural Biology, Max Planck Institute of Biophysics, Frankfurt, Germany
    Contribution
    Methodology, Prepared cryo-EM specimens and maintained electron microscopes and cameras
    Competing interests
    No competing interests declared
  4. Özkan Yildiz

    Department of Structural Biology, Max Planck Institute of Biophysics, Frankfurt, Germany
    Contribution
    Resources, Formal analysis, Validation, Visualization, Writing—review and editing, Built the atomic model, Analysed the structure; Drew the figures
    Competing interests
    No competing interests declared
    ORCID icon 0000-0003-3659-2805
  5. Werner Kühlbrandt

    Department of Structural Biology, Max Planck Institute of Biophysics, Frankfurt, Germany
    Contribution
    Conceptualization, Resources, Formal analysis, Supervision, Funding acquisition, Investigation, Writing—original draft, Project administration, Writing—review and editing, Initiated and directed the study, Analysed the structure
    For correspondence
    werner.kuehlbrandt@biophys.mpg.de
    Competing interests
    Reviewing editor, eLife
    ORCID icon 0000-0002-2013-4810

Funding

Max-Planck-Gesellschaft

  • Niklas Klusch
  • Bonnie J Murphy
  • Deryck J Mills
  • Özkan Yildiz
  • Werner Kühlbrandt

Deutsche Forschungsgemeinschaft

  • Niklas Klusch
  • Werner Kühlbrandt

European Molecular Biology Organization (ALTF 702–2016)

  • Bonnie J Murphy

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank Gerhard Hummer for discussion and Janet Vonck and Alexander Hahn for reading the manuscript. This work is supported by the Max Planck Society, DFG SFB 807 and an EMBO long-term fellowship to BJM (ALTF 702–2016).

Reviewing Editor

  1. Sjors HW Scheres, Reviewing Editor, MRC Laboratory of Molecular Biology, United Kingdom

Publication history

  1. Received: November 1, 2017
  2. Accepted: December 5, 2017
  3. Accepted Manuscript published: December 6, 2017 (version 1)
  4. Version of Record published: December 29, 2017 (version 2)

Copyright

© 2017, Klusch et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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