1. Structural Biology and Molecular Biophysics

Cryo-EM structures of mitochondrial respiratory complex I from Drosophila melanogaster

  1. Ahmed-Noor A Agip
  2. Injae Chung
  3. Alvaro Sanchez-Martinez
  4. Alexander J Whitworth  Is a corresponding author
  5. Judy Hirst  Is a corresponding author
  1. The Medical Research Council Mitochondrial Biology Unit, University of Cambridge, The Keith Peters Building, Cambridge Biomedical Campus, United Kingdom
https://doi.org/10.7554/eLife.84424
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Cite this article
  1. Ahmed-Noor A Agip
  2. Injae Chung
  3. Alvaro Sanchez-Martinez
  4. Alexander J Whitworth
  5. Judy Hirst
(2023)
Cryo-EM structures of mitochondrial respiratory complex I from Drosophila melanogaster
eLife 12:e84424.
https://doi.org/10.7554/eLife.84424
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Abstract

Respiratory complex I powers ATP synthesis by oxidative phosphorylation, exploiting the energy from NADH oxidation by ubiquinone to drive protons across an energy-transducing membrane. Drosophila melanogaster is a candidate model organism for complex I due to its high evolutionary conservation with the mammalian enzyme, well-developed genetic toolkit, and complex physiology for studies in specific cell types and tissues. Here, we isolate complex I from Drosophila and determine its structure, revealing a 43-subunit assembly with high structural homology to its 45-subunit mammalian counterpart, including a hitherto unknown homologue to subunit NDUFA3. The major conformational state of the Drosophila enzyme is the mammalian-type 'ready-to-go' active resting state, with a fully ordered and enclosed ubiquinone-binding site, but a subtly altered global conformation related to changes in subunit ND6. The mammalian-type 'deactive' pronounced resting state is not observed: in two minor states, the ubiquinone-binding site is unchanged, but a deactive-type π-bulge is present in ND6-TMH3. Our detailed structural knowledge of Drosophila complex I provides a foundation for new approaches to disentangle mechanisms of complex I catalysis and regulation in bioenergetics and physiology.

Editor's evaluation

This important article advances our understanding of respiratory complex I. The cryoEM data are convincing and the interpretation of different conformational states will stimulate discussions in the field. The work introduces Drosophila melanogaster as a model organism to study respiratory complex I and will be of interest to researchers studying respiratory enzymes, the evolution of respiration and mitochondrial diseases.

https://doi.org/10.7554/eLife.84424.sa0

Introduction

Mitochondrial complex I (NADH:ubiquinone oxidoreductase) is a crucial enzyme in cellular metabolism, central to NAD+ homeostasis, respiration, and oxidative phosphorylation, and a key contributor to the production of cellular reactive oxygen species (ROS) (Hirst, 2013; Parey et al., 2020). By catalyzing NADH oxidation in the mitochondrial matrix coupled to ubiquinone reduction in the inner membrane, it regenerates the oxidised NAD+ pool to sustain crucial metabolic processes, including the tricarboxylic acid cycle and β-oxidation, and provides reducing equivalents to the downstream complexes of the electron transport chain. The energy from NADH:ubiquinone oxidoreduction is harnessed to transport four protons across the inner membrane (Jones et al., 2017), supporting the proton motive force (Δp) that drives ATP synthesis and transport processes. These central roles of complex I in both metabolism and oxidative stress make complex I dysfunctions, induced by genetic, pharmacological, and environmental factors, some of the most frequent primary causes of mitochondrial diseases, as well as a contributor to many socially and economically important diseases common in ageing populations (Fassone and Rahman, 2012; Fiedorczuk and Sazanov, 2018; Padavannil et al., 2022). For example, ROS production by complex I operating in reverse, during ‘reverse electron transfer’ (RET, Δp-driven ubiquinol:NAD+ oxidoreduction) (Pryde and Hirst, 2011), is a major contributor to the tissue damage that occurs in strokes and heart attacks, during ischaemia-reperfusion (IR) injury (Chouchani et al., 2016; Chouchani et al., 2014; Dröse et al., 2016; Yin et al., 2021).

Mammalian complex I is a 1 MDa asymmetric assembly of 45 subunits, encoded on both the nuclear and mitochondrial genomes (Hirst, 2013; Hirst et al., 2003; Zhu et al., 2016). Fourteen of them (seven nuclear and seven mitochondrial) are the core subunits conserved in all complex I homologues that are essential for catalysis, whereas the other 31 subunits are supernumerary subunits that are involved in enzyme assembly, stability, and regulation, or that have independent roles within the cell (Hirst et al., 2003; Padavannil et al., 2022; Zhu et al., 2016). Bioinformatic analyses have indicated how the cohort of supernumerary subunits has been augmented gradually throughout the evolution of the eukaryotic complex (Gabaldón et al., 2005), and an increasing range of structural analyses of different species of complex I now illustrates the diversity of the supernumerary subunit cohorts that have developed in different eukaryotic lineages (Klusch et al., 2021; Parey et al., 2021; Soufari et al., 2020; Zhou et al., 2022).

For mammalian complex I, the form of the enzyme most relevant in medicine, single-particle electron cryomicroscopy (cryo-EM) has yielded detailed structural information on multiple different states of the complex (Chung et al., 2022a; Kampjut and Sazanov, 2022; Parey et al., 2020). However, detailed structure–function studies are limited for the mammalian enzyme due to substantial challenges in creating and studying genetic variants in representative mammalian model systems, such as mouse. Whereas simpler model systems, such as α-proteobacteria or yeast species (Jarman et al., 2021; Jarman and Hirst, 2022; Kravchuk et al., 2022; Parey et al., 2019), allow far greater opportunities for genetic studies, the protein compositions of their complex I vary substantially from the mammalian enzyme, fail to recapitulate key characteristics and behaviour of the mammalian complex such as the ‘active/deactive transition’ (Babot et al., 2014; Kotlyar and Vinogradov, 1990; Maklashina et al., 2003; Vinogradov, 1998), and the physiological environments in which the variant complexes can be studied are very restricted. Most relevant here, the active and deactive states of mammalian complex I are two biochemically and structurally characterised resting states of the complex (Agip et al., 2018; Blaza et al., 2018; Chung et al., 2022a; Chung et al., 2022b; Zhu et al., 2016): the ‘active’ ready-to-go resting state and the ‘deactive’ pronounced resting state. They differ both in their global conformations and in the status of local structural features. In particular, the ubiquinone-binding site in the active state is fully enclosed and sealed, whereas in the deactive state disorder in the enclosing loops opens the site to the matrix (Agip et al., 2018; Blaza et al., 2018; Chung et al., 2022a; Chung et al., 2022b; Zhu et al., 2016). The active and deactive resting states have also been referred to as the ‘closed’ and ‘open’ states of the mammalian enzyme on the basis of changes in the apparent angle between their membrane and hydrophilic domains (Kampjut and Sazanov, 2020). Finally, we note that there is currently substantial controversy about the biochemical and physiological relevance of the open states of the mammalian complex (Chung et al., 2022a), which have recently been proposed to include not only the deactive resting state but also on-cycle catalytic intermediates (Kampjut and Sazanov, 2020; Kravchuk et al., 2022).

The fruit fly, Drosophila melanogaster, is a powerful genetically tractable model organism for metazoa. Drosophila encodes a complex I with a composition that closely resembles that of the mammalian complex (Gabaldón et al., 2005; Rhooms et al., 2020), with clear homologues to 42 of the 44 mammalian subunits identified. Therefore, in addition to providing an additional model system for studying the mechanism of complex I catalysis (also accessible in simpler unicellular models), variants in Drosophila complex I can be studied for their effects on regulation and assembly (Cho et al., 2012; Garcia et al., 2017; Murari et al., 2020). Furthermore, Drosophila can potentially be exploited to investigate features of complex I function that are observed for mammalian complex I, but not universal features of the enzyme in simpler organisms, such as the active/deactive transition, RET, and the involvement of complex I in supercomplexes (Garcia et al., 2017; Scialò et al., 2016; Shimada et al., 2018). For instance, studies in Drosophila have proposed that RET-ROS increase lifespan (Scialò et al., 2016) and Drosophila are remarkably resistant to hypoxic or anoxic exposure (Haddad, 2006; Zhou and Haddad, 2013), which might provide insights into pathological mechanisms of RET-mediated IR injury. Furthermore, with substantial tissues, such as indirect flight muscles, highly enriched with mitochondria, Drosophila represent an attractive animal model for the analysis of basic mitochondrial biology, offering a complex physiological system for the generation and study of complex I genetic variants at the whole organism or tissue-specific level, as well as the involvement of complex I in differing physiological conditions.

To date, no detailed molecular studies of Drosophila complex I have been pursued to confirm its structural and functional similarity with the mammalian enzyme, or exploit its potential as a metazoan model system. Therefore, we sought here to structurally and biochemically evaluate Drosophila as a model system for mammalian complex I. We determine structures for three distinct conformational states of the Drosophila enzyme and compare them to well-characterised resting states of the mammalian complex, leading to new insights into the mammalian active/deactive transition and enhancing understanding of the conformational link between the ubiquinone-binding site and the proximal membrane domain. We thus present detailed knowledge of Drosophila complex I at the molecular level and confirm and define its relationships to the mammalian enzyme.

Results

The 43-subunit structure of Drosophila complex I

Complex I was isolated from mitochondrial membranes prepared from whole adult Drosophila by detergent extraction from the membrane followed by anion-exchange and size-exclusion chromatography, according to a small-scale protocol developed previously for mammalian complex I (Agip et al., 2018). The complex eluted from the size-exclusion column in a homogeneous peak consistent with the expected ~1 MDa mass of the monomeric complex (Figure 1—figure supplement 1). The highest concentration peak fraction (3.4 mg mL–1), which exhibited an NADH oxidoreductase activity comparable to mammalian complex I of 7.3 ± 0.3 µmol min–1 mg–1 (ca. 120 NADH s–1), was collected and frozen onto thiol-modified gold cryo-EM grids (Blaza et al., 2018; Meyerson et al., 2015; Russo and Passmore, 2014). The grids were imaged using a 300 KeV Titan Krios microscope equipped with a Gatan K2 camera and GIF Quantum energy filter (Table 1), and 63,471 particles images were selected and processed using RELION (Zivanov et al., 2020; Zivanov et al., 2018) into three major classes (Figure 1—figure supplement 2). The highest resolution map reached an estimated global resolution of 3.3 Å with consistent local resolution, and the two smaller subclasses reached estimated global resolutions of 3.7 and 4.0 Å (Figure 1—figure supplement 3). Example densities are shown in Figure 1—figure supplements 4–6.

Table 1
Cryo-EM data collection, refinement, and validation statistics for the three states of Drosophila complex I.
Drosophila melanogaster complex I dataset
Data collection and processing
Magnification130,000
Voltage (kV)300
Electron exposure (e–/Å2)42
Defocus range (μm)–1.0 to –2.0
Nominal pixel size (Å)1.07
Calibrated pixel size (Å)1.048
Symmetry imposedC1
Initial particle images (no.)194,538
Final particle images (no.)63,471
ClassesDm1 [Active]Dm2 [Twisted]Dm3 [Cracked]
EMD-15936EMD-15937EMD-15938
PDB-8B9ZPDB-8BA0
Final particle images (no.)37,60812,34313,520
Map resolution (Å)
 FSC threshold: 0.1433.283.683.96
Map resolution range (Å)2.98–6.193.33–9.403.51–11.35
Map sharpening B-factor (Å2)01520
Model statistics
Initial model (PDB ID)6G2J6G2J
Model resolution (Å)
 FSC threshold: 0.53.413.92
Model composition
 Non-hydrogen atoms66,97065,912
 Protein residues8,1788,136
 Ligands3922
Average B-factors (Å2)
 Protein99110
 Ligand103115
Root mean square deviations
 Bond lengths (Å)0.0060.007
 Bond angles (°)0.7260.783
MolProbity score1.602.04
All-atom clash score6.2410.90
EMRinger score3.181.68
Rotamer outliers (%)0.000.00
Ramachandran plot
 Favoured (%)96.1891.97
 Allowed (%)3.808.01
 Outliers (%)0.020.02

Figure 1 shows the overall structure of Drosophila complex I, which consists of 43 subunits: 14 core subunits (Figure 1a) and 29 supernumerary subunits (Figure 1b). The 14 core subunits comprise the canonical heart of the enzyme that is conserved throughout all species of complex I, with the core subunits of the Drosophila and mammalian (bovine, PDB ID: 7QSK; Chung et al., 2022b) enzymes exhibiting an overall root mean square deviation (RMSD) of 1.065 Å. The 29 supernumerary subunits all correspond to supernumerary subunits found in mammalian complex I, confirming the close relationship between them. However, two supernumerary subunits present in mammalian complex I are absent from the Drosophila complex (Figure 1c): subunits NDUFC1 and NDUFA2 (to aid comparisons to the mammalian enzyme, we use the human nomenclature throughout; see, e.g., Rhooms et al., 2020 for a list of the corresponding gene names in Drosophila). NDUFC1 is a short, single transmembrane helix (TMH)-containing subunit in the membrane domain that is peripherally associated with the mammalian complex through its interaction with subunit NDUFC2, and subunit NDUFA2 binds to subunit NDUFS1 at the top of the hydrophilic domain in the mammalian complex. The absence of NDUFC1 was expected since no orthologue was identified in the Drosophila genome by bioinformatic analyses (Gabaldón et al., 2005), and in Drosophila the N-terminus of NDUFC2 is displaced by the C-terminal extension of NDUFA11 (see Figure 1—figure supplement 5). Based on the same bioinformatic analyses, subunit NDUFA3 was also expected to be absent, but a matching subunit (Dmel gene CG9034) was detected by mass spectrometry in our preparation (see 'Materials and methods') and is clearly present in our density map in the location of mammalian-NDUFA3 in the membrane domain (see Figure 1—figure supplement 6). However, the sequence homology is weak and the structures of the two proteins diverge in the C-terminal membrane-extrinsic domain, with the obtuse-angled ‘turn’ that follows the TMH in the mammalian protein sterically blocked by the marginally extended C-terminal TMH of ND1 in Drosophila. NDUFA2, which has a characteristic thioredoxin fold and is widely conserved in eukaryotic complex I, is surprisingly absent from our Drosophila structure despite a highly conserved homologue in the Drosophila genome (Gabaldón et al., 2005). However, NDUFA2 interacts with only subunit NDUFS1 in the mammalian complex, and inspection of the (otherwise highly conserved) region of interaction in the Drosophila enzyme shows local disorder in a specific helix in the Drosophila NDUFS1 subunit (residues 673–684) that binds NDUFA2 in the mammalian enzyme. Although this result suggests NDUFA2 is associated with Drosophila complex I in vivo but has been lost during enzyme purification, detailed transcriptomic analyses (Brown et al., 2014; Leader et al., 2018) show that NDUFA2 (Drosophila ND-B8) expression is restricted principally to the male germline, and therefore the NDUFA2 protein is unlikely to be a constitutive component of complex I in somatic tissues.

Figure 1 with 6 supplements see all
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The architecture of complex I from Drosophila melanogaster.

(a) The 14 core subunits are shown in colour and labelled accordingly, and the 29 supernumerary subunits are shaded in grey. (b) The 29 supernumerary subunits are shown in colour and labelled accordingly, and the 14 core subunits are shaded in grey. (c) Drosophila complex I shown in transparent colour (as in a) with NDUFA2 (purple), NDUFC1 (green), and ND5-TMH1 (red), which are absent in Drosophila but present in mammalian complex I, indicated in solid colour from the structure of bovine complex I (PDB ID: 7QSK) (Chung et al., 2022b). The NADH-binding site at the flavin mononucleotide (FMN) cofactor, iron-sulphur clusters, the ubiquinone-binding site (Q9; purple), and the proton-pumping domain are indicated. All structures are of the Dm1 active-state Drosophila cryo-EM map, shown at a map threshold of 0.013 in UCSF ChimeraX (Pettersen et al., 2021). MM, mitochondrial matrix; IMS, intermembrane space; IMM, inner mitochondrial membrane; TMH, transmembrane helix; Q, ubiquinone; QH2, ubiquinol.

Overall, Drosophila complex I is remarkably similar in its composition and structure to the mammalian enzyme, underlining expectations of the value of Drosophila as a model system for complex I research. Only further minor differences are present in some of the subunits (Figure 1—figure supplement 5). ND5-TMH1 is absent from the Drosophila subunit: although ND2, ND4, and ND5 have a canonical 14-TMH core structure, truncation of the N-terminal TMHs appears tolerated, consistent with them lacking specific catalytically active residues or features, and demonstrated by the 11-TMH form of subunit ND2 in bilateria that lacks the three N-terminal TMHs found in lower organisms (Birrell and Hirst, 2010). In addition, the structures of supernumerary subunits NDUFB6 and NDUFB1 are noticeably different in the Drosophila enzyme (Figure 1—figure supplement 5). Notably, all the substantial differences in the membrane domain (absence of NDUFC1 and ND5-TMH1, variations in NDUFA3, NDUFB6 and NDUFB1, extension of NDUFA11) are located on the ‘right’ side of the boot-shaped enzyme, perhaps because there is less evolutionary pressure on the right side than on the left, where interactions with complexes III and IV are central to the stabilisation of respiratory chain supercomplexes (Milenkovic et al., 2017).

Three distinct states of Drosophila complex I

Cryo-EM particle classification identified three distinct states in our preparation of Drosophila complex I, which we refer to as Dm1, Dm2, and Dm3 (Figure 1—figure supplement 2). The Dm1 class is the dominant class, containing ~60% of the particles, whereas the two minor classes, Dm2 and Dm3, each contain ~20%. On a global scale (Figure 2), the Dm2 state is ‘twisted’ relative to the Dm1 state: with the two models aligned on subunit ND1 in the ‘heel’ of the complex, the hydrophilic and membrane domains twist in opposite directions (there is no apparent opening or closing motion between the domains). A similar twisting relationship was identified between the active and deactive resting states of mammalian complex I (Zhu et al., 2016). However, standard biochemical assays used to detect the presence of the mammalian deactive state did not detect any deactive Drosophila enzyme, even after incubation at 37°C to promote deactivation (Figure 2—figure supplement 1), indicating that Dm2 is not directly comparable to the mammalian-type deactive state. In the mammalian deactive state, the equivalent residue to ND3-Cys41 (we use Drosophila numbering throughout) is exposed to solution and can be derivatised by N-ethylmaleimide (NEM), preventing reactivation of the deactive enzyme and its return to catalysis, whereas in the active state ND3-Cys41 is buried (Galkin et al., 2008). Our assays suggest that either ND3-Cys41 is buried and inaccessible to derivatisation in all three Dm1, Dm2, and Dm3 states, or that ND3-Cys41 is exposed in one or more state that is completely inactive, being unable to either reactivate or catalyse. For the Dm3 state, the most obvious global feature (Figure 2) is that the membrane domain appears ‘cracked’ at the interface between ND2 and ND4; the density for the adjacent subunit NDUFA11 is disordered, along with the adjacent N-terminus of NDUFS2 (Figure 2—figure supplement 2). These characteristics resemble those of the ‘slack’ state of bovine complex I (Chung et al., 2022b; Zhu et al., 2016), which is of uncertain biochemical and physiological relevance and which may result from destabilisation of the membrane-intrinsic domain following extraction from the membrane and delipidation by detergents during purification. To evaluate the three states of ‘resting’ Drosophila complex I further, we first focus on the Dm1 state and its relationship with known resting states of the mammalian enzyme.

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Global comparison between the three states of Drosophila complex I.

Side views of the three Drosophila complex I cryo-EM maps identified by 3D classification are shown with global motions between the three states indicated. States Dm1 and Dm2 are related by a twisting motion of the hydrophilic and membrane domains about the ND1-containing ‘heel’ subdomain. States Dm2 and Dm3 are related by ‘cracking’ open of the ND2–ND4 interface in Dm3. Cryo-EM densities are shown at map thresholds of 0.013 (Dm1), 0.014 (Dm2), and 0.015 (Dm3) in UCSF ChimeraX (Pettersen et al., 2021).

Dm1 is the active resting state of Drosophila complex I

In addition to differing in their global conformations, the mammalian active and deactive states are differentiated by the status of a set of local features in the core subunits (Agip et al., 2018; Blaza et al., 2018; Chung et al., 2022a; Chung et al., 2022b; Zhu et al., 2016). In the Dm1 state of Drosophila complex I, these features are all unambiguously in the active state (Figure 3a). First, ND6-TMH3 is clearly α-helical, it does not contain the π-bulge that is characteristic of the deactive state, and ND1-TMH4 is clearly in the ‘bent’ conformation of the active state (with Tyr149 pointing toward the E-channel), not the straight conformation of the deactive state (with Tyr149 pointing away from the E-channel). Second, the densities for the NDUFS2-β1−β2 loop that carries the His97 ligand to bound ubiquinone, the ND3-TMH1−2 loop that carries Cys41 (the biochemical marker of the mammalian active/deactive states), and the ND1-TMH5−6 loop, are all well-defined in the Dm1 density map and their conformations match the mammalian active-state conformations (they are not disordered as in the deactive state). ND3-Cys41, NDUFS2-His93, and ND1-Tyr134 meet in a trigonal junction at the top of ND1-TMH4 (Grba and Hirst, 2020), as they do in the active state (Figure 3c). Finally, the FRASPR motif in NDUFS7 that includes Arg119 matches its conformation in the mammalian active, not deactive, state.

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Local structural elements show that Dm1 is the active resting state of Drosophila complex I.

(a) The local elements in the core subunits that show the Dm1 complex is in the active state are individually compared against an active-state bovine structure (transparent purple; PDB ID: 7QSK) (Chung et al., 2022b). Active state-specific key local features are indicated in square brackets. The same features are highlighted and labelled in (b) and (c), respectively, showing that subunits NDUFS2, NDUFS7, ND1, ND3, and ND6 encapsulate a fully structured and sealed Q-binding cavity (aquamarine surface; detected by CASTp; Tian et al., 2018) with a Q9 molecule bound. The Coulomb potential density for Q9 is shown in Figure 1—figure supplement 4. The box in (c) denotes the trigonal junction (ND3-Cys41, NDUFS2-His93, and ND1-Tyr134).

Importantly, as expected from the ordered states of the loop structures that form the ubiquinone-binding site, the site is sealed and closed from the matrix (Figure 3b and c) as in the mammalian active state, not open to the matrix as in the deactive state (Agip et al., 2018; Blaza et al., 2018; Chung et al., 2022a; Chung et al., 2022b; Zhu et al., 2016). This observation indicates that the Dm1 state is a catalytically competent state, ready to bind and reduce the extended and hydrophobic ubiquinone-9 or ubiquinone-10 substrate (referred to as Q9 for brevity). Indeed, density for Q9 is observed within the site, although the Q9 is only partially inserted, with its ubiquinone-headgroup in the central section of the channel, rather than ligated to the two proton-donor ligands NDUFS2-His97 and NDUFS2-Tyr146 as required for its reduction (Baradaran et al., 2013; Tocilescu et al., 2010). Partially inserted ubiquinones have been observed previously in several different species and states of complex I (Gu et al., 2022; Kampjut and Sazanov, 2020; Kravchuk et al., 2022; Parey et al., 2019; Soufari et al., 2020), but the headgroup typically sits slightly lower down the channel than observed here, in an array of positions distributed largely around the hydrophilic ‘kink’ of the channel (Figure 3—figure supplement 1). The Drosophila Q9 headgroup is bound 11 and 14 Å away, respectively, from its proposed ligating partners NDUFS2-His97 and NDUFS2-Tyr146, between the ‘1F’ site described in Sus scrofa complex I (Gu et al., 2022) and the ‘Qm’ site described in Escherichia coli (Kravchuk et al., 2022). The wide spectrum of headgroup positions identified in different complex I structures suggests that substrates may shuttle in a step-wise manner, occupying numerous sites of localised energy minima (Chung et al., 2021; Hoias Teixeira and Menegon Arantes, 2019; Warnau et al., 2018). Thus, although the site observed here is clearly separated from the reactive site, it is only broadly defined and not a highly specific site.

Modified domain disposition between the Drosophila and mammalian active states

In the mammalian complex, two subunits, NDUFA5 on the hydrophilic domain and NDUFA10 on the membrane domain, meet in the corner of the L-shape forming an interface between the two domains. Upon deactivation of the mammalian enzyme, the altered disposition of the hydrophilic and membrane domains changes the NDUFA5/NDUFA10 interface and decreases their contact area (Agip et al., 2018; Zhu et al., 2016). The nature and extent of the interface thereby provide an easy way to evaluate the active/deactive status of mammalian structures. Both subunits are present in the Drosophila enzyme, and Figure 4a compares their relative positions in the Dm1 active state to their relative positions in the active and deactive mammalian states. With the structures aligned to subunit NDUFA10, subunit NDUFA5, which is dominated by a three-helix bundle, clearly lies in an intermediate position in the Dm1 state, it does not overlay its position in the mammalian active state. The N-terminus of hydrophilic-domain subunit NDUFS2 that runs along the top of the membrane domain is also in an intermediate position. However, the NDUFA5/NDUFA10 contact area still matches closely to that observed in the mammalian active state (388 Å2 vs. 354 and 131 Å2 in the bovine active and deactive states [Chung et al., 2022b], respectively), and the NDUFA5/NDUFA10 interface is clearly different as it incorporates the N-terminus of the NDUFS4 subunit (residues 33–49; Figure 4b), which is substantially extended relative to in mammalian species (Figure 4c). Contacts between the NDUFS4 N-terminal ‘tether’ and subunits NDUFA5 and NDUFA10 of 182 and 427 Å2, respectively, further stabilise the interface and, by extension, the relative disposition of the hydrophilic and membrane domains in the Drosophila Dm1 active state.

Figure 4
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The domain interface between subunits NDUFA5, NDUFA10, and NDUFS4 in Drosophila complex I.

(a) The interface between subunits NDUFA5 (mint) and NDUFA10 (turquoise) of Drosophila complex I is compared against the active (purple; PDB ID: 7QSK) and deactive (light pink; PDB ID: 7QSM) states of bovine complex I (Chung et al., 2022b), displaying an ‘intermediate’ conformation. Triangles indicate positions of the three-helix bundles in NDUFA5. The structures are aligned to subunit NDUFA10. (b) The extended N-terminal loop of NDUFS4 (salmon) specific to Drosophila complex I is tethered between NDUFA5 and NDUFA10, locking them in place. Inset shows the positions of the three subunits in complex I. The active-state (Dm1) Drosophila complex I map is shown at a threshold of 0.013 in UCSF ChimeraX (Pettersen et al., 2021). (c) Sequence alignment of NDUFS4 across a selection of NDUFA5/NDUFA10-containing organisms. Residues are coloured by similarity. Known mitochondrial targeting sequences are highlighted in yellow, and the modelled N-terminal extension of NDUFS4 in the Dm1 active-state structure is highlighted in salmon. UniProt IDs used for the alignment in Clustal Omega 1.2.4 (Sievers et al., 2011): Drosophila melanogaster, Q9VWI0, Homo sapiens, O43181, Bos taurus, Q02375, Mus musculus, Q9CXZ1.

Further comparison of the mammalian and Drosophila active-state structures showed that subunits NDUFS2, NDUFS7, and ND1 that constitute the ubiquinone-binding site (Figure 3), overlay closely (RMSD 0.574 between Dm1 and the bovine active state [PDB ID: 7QSK]; Chung et al., 2022b) but that the structures then diverge along the membrane domain, shifting the position and orientation of subunit ND2 (Figure 5a). As NDUFA5 is bound to NDUFS2 and NDUFA10 to ND2, their relative positions thus also change. Within the connecting subdomain between ND1 and ND2 that contains subunits ND3, ND6, and ND4L, the arrangement of the TMHs in subunit ND6 (Figure 5b) clearly differs between the Drosophila and mammalian enzymes (despite them both containing a fully α-helical ND6-TMH3). In particular, ND6-TMH4 is markedly displaced, enhancing a cleft between ND6 and ND1 in which three phospholipid molecules are observed (Figure 1—figure supplement 4). The position of ND6-TMH4 is remarkably variable between different species and states of complex I, suggesting that it is not functionally important (Figure 5—figure supplement 1). Structures around ND6-TMH4 also vary (Figure 5b and c): (i) the ND6-TMH4–5 loop is restructured and its β-hairpin disrupted by neighbouring ND4L-TMH1, (ii) ND6-TMH1 is displaced away from TMH4 to avoid steric clashes, (iii) the ND6-TMH3–4 loop is restructured to accommodate the movement of TMH4, and (iv) the C-terminal loop of NDUFA9, located just above the reordered ND6-TMH3–4 loop, is retracted. The ND3-TMH1–2 loop, which is also adjacent to the restructured region, remains ordered with ND3-Cys41 occluded (Figure 5c). Strikingly, the residues of the central axis that link the terminus of the E-channel to the start of the first antiporter-like subunit (ND2) are not affected by the altered connecting subdomain structure, which thus adjusts the relative disposition of subunits in the hydrophilic and membrane domains in the Drosophila Dm1 state without affecting the catalytic machinery of the active state structure.

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The structure of subunit ND6 and the connecting subdomain between subunits ND1 and ND2 alters the relative domain dispositions in the Drosophila active state relative to the mammalian active state.

(a) The structures of subunits NDUFS2, NDUFS7, NDUFA5, and ND1 are tightly conserved between the Drosophila (Dm1, solid cartoon) and mammalian (PDB ID: 7QSK [Chung et al., 2022b], transparent cartoon) active states, forming a rigid subdomain. Subunits ND3, ND6, and ND4L form a connecting subdomain that differs, shifting the position and orientation of the mobile subdomain containing subunits ND2 and NDUFA10. The altered connecting domain changes the domain interface between NDUFA5 and NDUFA10. The models for the Drosophila and mammalian complexes are aligned on subunit NDUFS2 and shown alongside a flat surface representation of the Drosophila model. (b, c) Changes to the structure of the ND6 subunit, plus structural changes in adjacent subunits. The models for the Drosophila (Dm1, coloured) and mammalian (PDB ID: 7QSK [Chung et al., 2022b], white) active states are overlaid on subunit ND6.

Minor states with restricted deactive characteristics

Inspection of the set of local features in the core subunits (Agip et al., 2018; Blaza et al., 2018; Chung et al., 2022a; Chung et al., 2022b; Letts et al., 2019; Zhu et al., 2016) that differentiate the mammalian active and deactive states in the two minor states revealed that both the Dm2 and Dm3 states also most closely correspond to the mammalian active state. Figure 6a shows that, in the Dm2 state, two features of the mammalian deactive state are present: a π-bulge has formed in ND6-TMH3 and ND1-TMH4-Tyr149 has ‘flipped’ its conformation (Chung et al., 2022b; Grba and Hirst, 2020; Kampjut and Sazanov, 2020). However, all the other key elements remain in their active states, including ND1-TMH4, which remains in its bent conformation, and as a result the ubiquinone-binding site remains enclosed and sealed from the matrix (Figure 6b and c). The same is true for the Dm3 state (Figure 6—figure supplement 1). Notably, the trigonal junction between ND3-Cys41, NDUFS2-His93, and ND1-Tyr134 (Grba and Hirst, 2020) is preserved in all three states, occluding the Cys from the matrix and explaining why a mammalian-type deactive state of Drosophila complex I could not be trapped in biochemical assays (Figure 2—figure supplement 1). We conclude that, in contrast to the mammalian enzyme, Drosophila complex I does not form an ‘open’ resting state, it rests only in ‘closed’ conformations with the ubiquinone-binding site enclosed and sealed from the matrix.

Figure 6 with 1 supplement see all
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Local structural elements show that the Dm2 state of Drosophila complex I most closely resembles the mammalian active state, with only two deactive-like features in the membrane domain.

(a) The local elements in the core subunits, individually compared against an active-state bovine structure (transparent purple; PDB ID: 7QSK) (Chung et al., 2022b), show that all ubiquinone-binding site features of the Dm2 complex are in the active state, whereas in the membrane domain ND1-Tyr149 and ND6-TMH3 match the bovine deactive state (transparent pink; PDB ID: 7QSM) (Chung et al., 2022b). Active state-specific key local features are indicated in square brackets in purple and deactive state-specific features in pink. The same features are highlighted and labelled in (b) and (c), respectively, showing that subunits NDUFS2, NDUFS7, ND1, ND3, and ND6 encapsulate a fully structured and sealed Q-binding cavity (aquamarine surface; detected by CASTp; Tian et al., 2018). The box in (c) denotes the trigonal junction (ND3-Cys41, NDUFS2-His93, and ND1-Tyr134).

Further comparison of the Dm1 and Dm2 states (Figure 7a and b) shows that the ‘twisting’ motion that relates their global conformations originates in changes in the ‘connecting subdomain’ between the rigid and mobile subdomains described in Figure 5a, where the deactive-like elements of Dm2 are located (the π-bulge in ND6-TMH3 and ND1-TMH4-Tyr149). The twisting motion displaces the N-terminus of subunit NDUFS2 (adjacent to ND6-TMH5) and changes the NDUFA5/NDUFA10 interface, displacing and disordering the N-terminus of subunit NDUFS4 (Figure 7c) and causing a small decrease in the interface area (from 388 Å2 in Dm1 to 333 Å2 in Dm2, relative to 354 and 131 Å2 in the active and deactive states of bovine complex I; Chung et al., 2022b). Inspection of the region around the π-bulge in ND6-TMH3 revealed a further striking change between the Dm1 and Dm2 states. In Dm1, the tail of a phosphatidylcholine molecule is intercalated into the structure, sterically obstructing the rotation of bulky residues on ND6-TMH3 around the helical axis to form the π-bulge (Figure 7a and b). It is absent from the Dm2 state (and also from Dm3, where the local protein conformation matches Dm2; Figure 6—figure supplement 1). In the Dm1 state, the phosphatidylcholine headgroup stabilises the ND6-TMH3–4 loop at the top of ND6-TMH4, whereas its absence in Dm2 allows the TMH3–4 loop and TMH4 to move, along with a further adjustment to the adjacent C-terminal loop of NDUFA9 and displacement of ND3-TMH2–3. The lipid may either have been ejected during relaxation of ND6-TMH3 into a π-bulge structure, or removed during detergent extraction, promoting π-bulge formation.

Figure 7
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Differences between the Dm1 and Dm2 states at the connecting subdomain and at the domain interface between subunits NDUFA5, NDUFA10, and NDUFS4.

(a, b) Changes to the structure of the ND6 subunit, plus structural changes in adjacent subunits. The intercalated phosphatidylcholine molecule is present in the Dm1 state only. (c) The interface between subunits NDUFA5 (mint), NDUFA10 (turquoise), and NDUFS4 (slate) of the Dm1 state is compared against Dm2 (white). Triangles indicate positions of the three-helix bundles in NDUFA5. The N-terminal NDUFS4 tether occupies the interface in Dm1 only, and the NDUFA5/NDUFA10 interface area is decreased in Dm2. The models for the Dm1 (coloured) and Dm2 (white) states are overlaid on subunit ND6 in (a) and (b), and on subunit NDUFA10 in (c).

Discussion

The structures determined here for Drosophila complex I confirm its close relationships with mammalian complex I and thus its potential as a powerful genetically tractable model system for studying mammalian-specific aspects of complex I biology. The close-to-identical subunit compositions and structures of the mammalian and Drosophila enzymes now enable genetic approaches to be applied to elucidate, for example, the roles of the supernumerary subunits, the assembly pathway, and the detrimental effects of clinically identified pathological point mutations. Importantly, these aspects can be studied in physiologically relevant in vivo environments and in specific cell types and tissues, extending the scope of earlier studies in cultured mammalian cells (Guerrero-Castillo et al., 2017; Stroud et al., 2016). However, our structures also reveal limitations in Drosophila as a model organism for complex I, as the Drosophila enzyme, despite its remarkable similarity to the mammalian enzyme, does not undergo the full mammalian-type active/deactive transition. Our cryo-EM analyses revealed the major class of enzyme particle (Dm1) in the active resting state, with all the characteristics of the mammalian active state enzyme. Two minor states (Dm2 and Dm3) also more closely resemble the active state, and we were unable to either detect a mammalian-type deactive resting state in biochemical assays, or to generate one by incubation of the enzyme at 37°C (the method used to deactivate mammalian complex I). However, we note that our biochemical assay relies on the availability of ND3-Cys41, just one characteristic that distinguishes the mammalian active and deactive states; the functional consequences of conversion to the Dm2 state are currently unknown, most notably whether it is (like the mammalian active state) able to catalyse RET, or (like the mammalian deactive state) unable to do so.

Comparison of the Dm1 (active) and Dm2 (twisted) structures of Drosophila complex I determined here suggests that the Dm2 state is a relaxed state, which may be considered a structurally curtailed form of the full mammalian-type deactive transition (Agip et al., 2018; Blaza et al., 2018; Chung et al., 2022a; Chung et al., 2022b; Zhu et al., 2016). In changes that also occur in the mammalian transition, a π-bulge forms in ND6-TMH3, and the nearby sidechain of ND1-TMH4-Tyr149 flips in conformation. Although water molecules cannot be resolved in our structures, these changes are expected to alter the connectivity between the E-channel and the central axis of charged residues along the membrane domain as reported previously for mammalian, yeast, and bacterial species (Agip et al., 2018; Blaza et al., 2018; Chung et al., 2022b; Grba and Hirst, 2020; Kampjut and Sazanov, 2020; Kravchuk et al., 2022; Parey et al., 2021). Furthermore, the limited and local changes we observe in the ND6 region result in a limited twisting of the global conformation in the Dm2 state, a motion that qualitatively resembles (but to a much lesser extent) the twisting of the deactive enzyme. However, the cascade of changes that also occurs in the mammalian-type deactive transition does not follow: ND1-TMH4 does not straighten its conformation, the trigonal junction between ND3-Cys41, NDUFS2-His93, and ND1-Tyr134 is preserved, and so the conformational change from ND6-TMH3 does not propagate to the ubiquinone-binding site, which remains fully ordered, sealed from the matrix, and in its active state (Figure 6). This lack of direct correlation between the status of the α-helix/π-bulge and the structuring of the ubiquinone-binding site (Figure 8) argues against a concerted and structurally enforced connection between them being crucial for catalysis (Kampjut and Sazanov, 2022; Kampjut and Sazanov, 2020; Kravchuk et al., 2022). Although structures of complex I from other (non-mammalian) species have also been reported with a π-bulge in ND6-TMH3 but without the ‘opening’ of the ubiquinone-binding site observed in the mammalian deactive state (Chung et al., 2022a), our Drosophila Dm2 structure is the first example in which the π-bulge and a fully ordered, active ubiquinone-binding site have been observed together. Our structures are consistent with the elements that change during the mammalian deactive transition (such as the π-bulge) being mobile during catalysis, but do not suggest that they move in a coherent and coordinated transition, with the ubiquinone-binding site open to the matrix, during catalysis.

Figure 8
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Schematic representation of the status of local active/deactive elements in the (a) Dm1 and (b) Dm2 states of Drosophila complex I.

Local elements that change conformation in the mammalian active/deactive transition are shown and labelled as A for active and D for deactive, respectively. In the Dm2 state, ND6-TMH3 and ND1-TMH4-Tyr149 are in the D conformation. The boundary of the A and D regions is marked with a dashed line. In the mammalian deactive state, the top section of ND1-TMH4 moves, straightening the helix and resulting in loss of the trigonal junction (ND3-Cys41, NDUFS2-His93, and ND1-Tyr134), destructuring of the NDUFS2, ND3, and ND1 loops and restructuring of the NDUFS7 loop and NDUFS7-Arg119. ND3-Cys41, the derivatisable marker of the deactive state in mammalian/eukaryotic complex I (Galkin et al., 2008), is indicated with a yellow circle.

The observation, together, of the active Dm1 state and the ‘curtailed-deactive’ Dm2 state raises two questions: what causes the π-bulge to form in Dm2, and why does the conformational cascade to the mammalian-type deactive state not occur in the Drosophila enzyme (Figure 8)? First, it is possible that delipidation of the complex during detergent extraction removes the intercalated phospholipid that obstructs π-bulge formation in the Dm1 state, allowing conversion to Dm2. However, a similar intercalated phospholipid has not been observed in any mammalian active-state structure, so it may only bind when catalysis stops, or be an artefact of enzyme purification. Indeed, if ND6-TMH3 converts between its π-bulge and α-helical structures during catalysis (Agip et al., 2018; Kampjut and Sazanov, 2020; Kravchuk et al., 2022; Parey et al., 2021; Röpke et al., 2021), then the intercalating phospholipid is very unlikely to be present in the α-helical state, moving repeatedly in and out. Alternatively, it is possible that enzyme twisting, induced by loss of the NDUFS4 tether from the NDUFA5/NDUFA10 interface during purification, causes the π-bulge to form: this possibility may be addressed in future by genetic truncation of the NDUFS4 tether from the N-terminus of the mature subunit. Second, if formation of the π-bulge in Drosophila represents a curtailed-deactive transition, then conversion to a full mammalian-type deactive state would be accompanied by further twisting, disruption of the NDUFA5/NDUFA10 interface, and destructuring of the ubiquinone-binding site. That these changes are not observed in Drosophila complex I is likely due to the modified domain disposition in the Dm1 state that is stabilised by the structure of the connecting subdomain and accommodating changes in linked structures such as the NDUFA5/NDUFA10 interface. We propose that the stable domain disposition is resistant to further twisting and so resists the local changes that accompany it in the mammalian deactive transition. Computational simulations of the Dm1 structure may help further elucidate the answer to this question in future. Notably, our proposal implies high activation energy barriers for the ‘opening’ of the ubiquinone-binding site to the matrix in the Drosophila enzyme, arguing against opening and closing of the site during catalysis (Kampjut and Sazanov, 2020; Kravchuk et al., 2022).

The Dm3 ‘cracked’ state is not discussed in detail here as we suspect it is an artefact resulting from detergent-induced loss of stability in the distal membrane domain of the Dm2 state. Similar opening and relaxation of the ND2–ND4 interface has also been observed in the ‘slack’ state of bovine complex I (Chung et al., 2022b; Zhu et al., 2016), as well as in a catalytically inactive state of complex I from rhesus macaque (Agip et al., 2019), and in pronounced open states of the ovine complex (Kampjut and Sazanov, 2020). In all cases, opening of the ND2–ND4 interface is linked to loss of density for nearby subunit NDUFA11, and to changes in the C-terminal section of the ND5 transverse helix and anchor helix (Figure 2—figure supplement 2). It may result from delipidation during enzyme purification, most likely removal of phospholipids from the interface on both sides of the complex, including ‘behind’ the transverse helix (Figure 1—figure supplement 4b). Consistent with this picture, treatment of the mammalian enzyme with zwitterionic detergents or prolonged incubation in detergent solution leads to fractionation at this interface (Hirst et al., 2003; Zhu et al., 2015).

The deactive transition and RET are linked in mammalian complex I biology, as deactivation protects against the burst of ROS production that occurs upon reperfusion by RET (RET-ROS), driven by oxidation of the reduced succinate pool that accumulates during ischaemia, leading to IR injury (Dröse et al., 2016; Galkin and Moncada, 2017; Wright et al., 2022; Yin et al., 2021). The deactivation of complex I minimises the RET-ROS burst and tissue damage upon reperfusion because the deactive state of mammalian complex I is unable to catalyse RET (Kotlyar and Vinogradov, 1990; Wright et al., 2022; Yin et al., 2021). An elegant demonstration is provided by the ND6-P25L variant of mouse complex I, which deactivates much more rapidly than the wild-type enzyme, preventing RET-ROS catalysis and thereby protecting against IR injury (Yin et al., 2021). Strikingly, while Drosophila do not appear to adopt a mammalian-type deactive state, they are able to survive long periods of hypoxia followed by reoxygenation (Haddad, 2006; Zhou and Haddad, 2013), raising the question of whether they are protected by a corresponding mechanism. ROS production by RET has been described in studies of Drosophila mitochondria (although not demonstrated directly in the isolated enzyme) (Scialò et al., 2016), and the ability of Drosophila complex I to catalyse RET is consistent with it persisting in the active state (Dm1) when catalysis stops, rather than deactivating. Alternative mechanisms are therefore required to explain the resistance of Drosophila to hypoxia−reoxygenation challenges, such as greater robustness to oxidative stress from a RET-ROS-induced stress-responsive transcriptional programme (Scialò et al., 2020) and/or metabolic adaptations (Perkins et al., 2012). Future genetic studies that exploit structural insights will illuminate these mechanisms and provide new perspectives on the mechanisms of mammalian complex I.

Materials and methods

Key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Gene (Drosophila melanogaster)NDUFA3, Dmel gene CG9034FlyBaseUniProt ID: Q9W380
FlyBase ID: FBgn0040931
Biological sample (D. melanogaster)isogenic w1118Bloomington Drosophila Stock CenterRRID:BDSC_6326
Chemical compound, drugFatty acid-free bovine serum albuminMerck MilliporeCAS number:
9048-46-8
Chemical compound, drugEDTA-free cOmplete protease inhibitor cocktailRocheCOEDTAF-RO
Chemical compound, drugDodecyl-β-D-maltoside (DDM)Merck MiliporeCAS number:
69227-93-6
Chemical compound, drugAsolectin from soy beanAvantiCAS number:
8030-76-0
Chemical compound, drugCHAPS (3-((3-cholamidopropyl) dimethylammonio)–1-propanesulfonate)CalbiochemCAS number:
75621-03-3
Chemical compound, drugNADHMerck MilliporeCAS number:
606-68-8
Chemical compound, drugDecylubiquinone (dQ)Merck MilliporeCAS number:
55486-00-5
Chemical compound, drugN-ethylmaleimide (NEM)Merck MilliporeCAS number:
128-53-0
Chemical compound, drug11-Mercaptoundecyl hexaethylene glycolSensoPath TechnologiesSPT-0011P6
Software, algorithmEPUThermo Fisher Scientific
Software, algorithmRELION-3.0Zivanov et al., 2018RRID:SCR_016274
Software, algorithmRELION-3.1Zivanov et al., 2020RRID:SCR_016274
Software, algorithmMotionCor2Zheng et al., 2017RRID:SCR_016499
Software, algorithmCTFFIND-4.1Rohou and Grigorieff, 2015RRID:SCR_016732
Software, algorithmcrYOLO 1.5.3Wagner et al., 2019https://cryolo.readthedocs.io/en/stable/
Software, algorithmUCSF ChimeraXPettersen et al., 2021RRID:SCR_015872
Software, algorithm3DFSCTan et al., 2017https://github.com/LyumkisLab/3DFSC; LyumkisLab, 2019 version 3.0
Software, algorithmSWISS-MODELWaterhouse et al., 2018RRID:SCR_018123
Software, algorithmUCSF ChimeraPettersen et al., 2004RRID:SCR_004097
Software, algorithmMODELLERWebb and Sali, 2016RRID:SCR_008395
Software, algorithmCootCasañal et al., 2020RRID:SCR_014222
Software, algorithmISOLDECroll, 2018https://isolde.cimr.cam.ac.uk/
Software, algorithmPyMOL 2.5.2Schrodinger, 2022RRID:SCR_000305
Software, algorithmPhenix 1.18.2–3874Liebschner et al., 2019RRID:SCR_014224
Software, algorithmMapQPintilie et al., 2020
Software, algorithmMolProbityChen et al., 2010RRID:SCR_014226
Software, algorithmEMRingerBarad et al., 2015
Software, algorithmCASTpTian et al., 2018http://sts.bioe.uic.edu/castp/
OtherUltrAuFoil gold gridsQuantifoil; 
Russo and Passmore, 2014		R 0.6/1 Gold foil on Gold 300 mesh grid

Drosophila stocks and husbandry

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Flies of a common wild type-equivalent genotype, isogenic w1118 (RRID:BDSC_6326), were obtained from Bloomington Drosophila Stock Center (RRID:SCR_006457), raised and kept under standard conditions in a temperature-controlled incubator with a 12 hr:12 hr light:dark cycle at 25°C and 65% relative humidity, on food consisting of agar, cornmeal, molasses, propionic acid, and yeast. Approximately 5500 mixed adults collected 5 days after eclosion were used for the preparation of the cryo-EM sample.

Preparation of Drosophila complex I

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All experimental procedures were carried at 4°C unless otherwise stated. One volume of w1118 flies (1 mL is equivalent to ~100 flies) was mixed with five volumes of homogenisation buffer containing 20 mM Tris-HCl pH 7.8, 250 mM sucrose, 2 mM EDTA, 2 mM EGTA, 1% (w/v) fatty acid-free bovine serum albumin (BSA, Merck) and 1× EDTA-free cOmplete protease inhibitor cocktail (Roche) (one tablet per 50 mL of buffer). Mitochondria were prepared using a differential centrifugation method. Briefly, 20 mL of fly suspension were homogenised by 10 strokes with a motor-driven Teflon pestle at 1300 rpm in a 30 mL Wheaton glass homogeniser. The homogenate was then centrifuged at 1000 × g for 5 min, and the supernatant filtered through a muslin cloth to remove cuticles. The same process was repeated a second time. Then, mitochondria were pelleted at 3000 × g for 10 min and washed with 5 mL of homogenisation buffer but without BSA. Finally, mitochondria were collected by centrifugation at 7000 × g for 10 min and resuspended in 5.8 mL (56 mg of protein) of resuspension buffer containing 20 mM Tris-HCl (pH 7.8), 20% glycerol (v/v), 2 mM EDTA, 2 mM EGTA, 1% and 1× EDTA-free cOmplete protease inhibitor cocktail (one tablet per 50 mL of buffer). Isolated mitochondria were stored at –80°C until further use. Mitochondrial membranes were prepared as described previously for mouse samples (Agip et al., 2018). Defrosted mitochondria were diluted to 5 mg mL–1 in resuspension buffer then ruptured on ice with a Q700 Sonicator (Qsonica) in three intervals (5 s bursts each followed by a 30 s pause) at an amplitude setting of 65%. Membranes (38 mg of protein) were collected by centrifugation at 75,000 × g for 1 hr, then resuspended to 4.9 mg mL–1 in the same buffer and stored at –80°C.

Purification of Drosophila melanogaster complex I followed the same procedure as previously described for mouse complex I, with minor adjustments (Agip et al., 2018). While being continuously stirred on ice, mitochondrial membranes (7.6 mL at 4.9 mg mL–1) were solubilised for 30 min by the drop-wise addition of dodecyl-β-D-maltoside (DDM) to a final concentration of 0.75% from a 10% stock solution. The solubilised membranes were then centrifuged at 48,000 × g for 30 min and the clarified supernatant loaded on to a Hi-Trap Q HP anion exchange column (1 mL; Cytiva) pre-equilibrated with elution buffer A (20 mM Tris-HCl pH 7.8°C, 2 mM EDTA, 2 mM EGTA, 0.1% DDM, 10% ethylene glycol [v/v, VWR], 0.005% asolectin [Avanti], and 0.005% CHAPS [Calbiochem]) and operated at a flow rate of 0.3 mL min–1. The column was washed with several column volumes of buffer A until the 280 nm absorbance reached the baseline. Unwanted proteins were eluted with seven column volumes of 20% buffer B (buffer A + 1 M NaCl), then complex I was eluted with an additional seven column volumes of 35% buffer B. Fractions containing complex I (ca. 3 mL) were pooled and concentrated to ca. 100 µL using an Amicon-Ultra filter device (100 kDa molecular weight cut-off; Amicon, Millipore). The concentrated sample was then injected onto a Superose 6 Increase size exclusion column (150 × 5 mm; Cytiva) pre-equilibrated in buffer C (20 mM Tris-HCl pH 7.8°C, 150 mM NaCl, and 0.05% DDM) operated at a flow rate of 0.03 mL min–1. All chromatographic procedures described were carried out using an ÄKTA micro FPLC system (Cytiva) with elution monitored at 280 and 420 nm. Complex I concentrations were estimated at 280 nm (ε = 0.2 mg mL–1 mm–1). The total collected protein was estimated at 0.6 mg, and the peak concentration was 3.4 mg mL–1.

Kinetic activity measurements

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All activity measurements were measured on a 96-well Spectramax 384 plate reader at 32°C. For NADH:decylubiquinone (dQ) oxidoreductase activities, NADH (200 µM final concentration) was used to initiate catalysis by complex I (0.2 µg mL–1) with 200 µM dQ, 0.15% (w/v) asolectin, and 0.15% (w/v) CHAPS in 20 mM Tris-HCl (pH 7.55). NADH oxidation was monitored at 340–380 nm (ε = 4.81 mM–1 cm–1), and was confirmed to be sensitive to rotenone and piericidin A. The cryo-EM sample had an activity of 7.3 ± 0.3 µmol min–1 mg–1 (mean ± SD; n = 4).

For evaluation of the active/deactive state ratio of Drosophila complex I using the N-ethylmaleimide (NEM) assay (Galkin et al., 2008; Yin et al., 2021), 4 mg mL–1 mitochondria were incubated with 2 mM NEM or the equivalent volume of DMSO on ice for 20 min., before determining the NADH:O2 oxidoreductase activity. The mitochondria had been frozen for storage before measurement. To attempt to deactivate the complex, the mitochondria were incubated at 37°C for 30 min (equivalent to, or longer than, the treatments used to deactivate complex I in mammalian mitochondrial membranes [Agip et al., 2018; Blaza et al., 2018]). NADH:O2 oxidoreductase activities were measured in 20 mM Tris-HCl (pH 7.55) using 10 µg mL–1 mitochondria and 10 µg mL–1 alamethicin, and initiated using 200 µM NADH. NADH oxidation was monitored as described above.

Cryo-EM grid preparation and image acquisition

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UltrAuFoil gold grids (0.6/1, Quantifoil) (Russo and Passmore, 2014) were prepared for Drosophila complex I as described previously (Blaza et al., 2018). First, the grids were glow discharged with plasma under vacuum for 90 s at 20 mA then incubated for 7 days under anaerobic and room temperature conditions in a solution of 5 mM 11-mercaptoundecyl hexaethylene glycol (SPT-0011P6, SensoPath Technologies) (Meyerson et al., 2015). Grids were then washed several times in ethanol and left to dry. Complex I (3.4 mg mL–1) was then applied (3 µL per grid) to the treated grids in a Vitrobot Mark IV (Thermo Fisher Scientific) set to 4°C and 100% relative humidity. Grids were blotted for 10 s with a force setting of –10, before being plunged into liquid ethane. Frozen grids were then stored in liquid nitrogen before screening and data collection.

Both cryo-EM screening and high-resolution image collection were carried out on a Titan Krios (Thermo Fisher Scientific) at University of Cambridge cryo-EM facility. The Titan Krios microscope for data collection was operating at an accelerated voltage of 300 kV and equipped with a Gatan K2 detector utilising a GIF quantum energy filter with a slit width of 20 eV. The microscope was operated in electron counting mode with a nominal sampling rate of 1.07 Å pix–1 (nominal magnification of 130,000) and a dose rate of ca. 4.18 electrons Å–2 s–1. The specimen was radiated for 10 s over 40 frames with a total exposure amounting to ca. 42 electrons Å–2. A 100 µm and 50 µm objective and C2 aperture, respectively, were inserted during high-resolution imaging. The microscope was operated with EPU software and the defocus range was set to –1.0 to –2.0 µm, with an autofocus routine run every 5 µm.

Cryo-EM data processing

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All 3082 collected movies were subjected to processing by RELION-3.0 and 3.1 (Zivanov et al., 2020; Zivanov et al., 2018) except where stated otherwise. Micrographs were motion-corrected using MotionCor2 (Zheng et al., 2017) with 5 × 5 patches, and contrast transfer function (CTF) parameters estimated using CTFFIND-4.1 (Rohou and Grigorieff, 2015) in RELION-3.0. In parallel, the motion-corrected micrographs were exported and subjected to particle autopicking using a general model without training in crYOLO 1.5.3 (Wagner et al., 2019), resulting in 194,538 picked particles. Ice-contaminated micrographs were removed to give 180,342 particles from 2852 micrographs. Particles were extracted with an initial downscaling to 6.0 Å pixel–1 (box size of 80) and subjected to initial 2D and 3D classification steps to remove junk particles, yielding a total of 93,332 particles. These particles were re-extracted at the nominal pixel size of 1.07 Å pixel–1 (box size of 450) and used to reconstruct a 3.74 Å resolution map using the 3D autorefinement procedure in RELION, at the calibrated pixel size of 1.048 Å pixel–1 (Spikes et al., 2020). The active state map of mouse complex I (EMD-4345) (Agip et al., 2018) was used as a reference map for the 3D reconstruction. Bayesian polishing (Zivanov et al., 2019) was then applied and CTF parameters, including astigmatism, defocus, and beam tilt, estimated using the CTF refinement procedure in RELION-3.0. Particles were subjected to additional rounds of classifications, to further remove junk and bad complex I particles. From hereon, all data processing was performed in RELION-3.1, at the nominal pixel size of 1.07 Å pixel–1, then corrected to the calibrated pixel size of 1.048 Å pixel–1 at the postprocessing or local resolution stages. The particles were subject to iterative rounds of CTF refinement (Zivanov et al., 2020), to estimate anisotropic magnification, beam tilt, trefoil, fourth-order aberration, and per-particle defocus, astigmatism and B-factor parameters. Particles with an rlnNrOfSignificantSamples value greater than 3000 were removed to give 65,864 particles. Using a complex I mask (generated from a working model using RELION MaskCreate) and with solvent flattening, the global resolution of the 3D refined map was 3.23 Å (according to a gold-standard Fourier shell correlation [FSC] of 0.143; Rosenthal and Henderson, 2003). 3D classification (number of classes, K = 5, local angular search to 0.2° sampling) was then performed, and three complex I classes, Dm1, Dm2, and Dm3, were identified and retained, containing 37,608, 12,343, and 13,520 particles, respectively, a ratio of roughly 3:1:1. Clear cryo-EM densities for Dm2-specific local features, including the ‘flipped’ ND1-TMH4-Tyr149 and the ND6-TMH3 π-bulge, revealed no evidence for Dm1 contamination in the Dm2 population. The ‘Focus-Revert-Classify’ classification strategy (Letts et al., 2019), applied using the regularisation parameter t = 8 and K = 5, yielded comparable population distributions (three complex I classes matching Dm1, Dm2, and Dm3, plus two junk classes) whilst 3D classification without alignment using t = 20 and K ≤ 12 yielded two <4 Å complex I classes, with the major class matching Dm1 and the minor class an apparent mixture of Dm2 and Dm3. The 3D classification approach with local angular sampling was therefore employed to give the final set of Dm1, Dm2, and Dm3 particles as described above. Using model-generated (Dm1 and Dm2) or map-generated (Dm3) masks and solvent-flattening, the three classes refined to 3.28, 3.68, and 3.96 Å resolution, respectively. Global resolutions were estimated from two independent half maps using a gold-standard FSC of 0.143 (Rosenthal and Henderson, 2003) in RELION postprocess. The final map was globally sharpened (or blurred) in RELION postprocess using user-provided B-factor values. The model-generated or map-generated mask used for 3D refinement procedures and resolution estimation was generated in UCSF ChimeraX (Pettersen et al., 2021) using the molmap (Dm1 and Dm2) or vop threshold (Dm3) functions, before being low-pass filtered to 15 Å and having a 6-pixel soft cosine edge added using RELION MaskCreate. Local resolution was estimated using RELION LocRes. Mollweide projections were plotted using Python and Matplotlib, and the degree of directional resolution anisotropy calculated using the 3DFSC program suite (Tan et al., 2017).

Model building, refinement, and validation

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Model coordinates were built into the 3.3 Å resolution Dm1 Drosophila complex I map, with a published mouse complex I structure (PDB ID: 6G2J) (Agip et al., 2018) serving as a homology model. SWISS-MODEL (Waterhouse et al., 2018) was used to generate an initial model for each subunit and the models rigid-body fitted into the map using Chimera (Pettersen et al., 2004). MODELLER (Webb and Sali, 2016) was used to generate models for subunits NDUFB6, NDUFB9, and NDUFB8 as the homology models were unsatisfactory. The sequence for the Drosophila NDUFA3 subunit (UniProt ID: Q9W380; Dmel gene CG9034; FlyBase ID: FBgn0040931) was identified in routine peptide mass spectrometry analyses of the purified enzyme. The evidence for Q9W380 relies on a single peptide (LGYVVYR, 9.1% coverage) but the MASCOT score is 43, well above 30 (the 99% confidence limit), and inspection of the distribution of tryptic cleavage sites suggests no further peptides would be expected to be detected. Models for subunits NDUFA2 and NDUFC1 were deleted due to the lack of corresponding densities in the cryo-EM maps, and N- and C-terminal extensions were built where necessary. It was noted that densities for the N-termini of ND1 and ND5 were extended beyond the reviewed UniProt sequences (P18929 and P18932, respectively). Therefore, to incorporate the correct translation start site, UniProt IDs C7DZL9 and C7DZL4 (Stewart and Beckenbach, 2009) were used to build models for subunits ND1 and ND5, respectively. UniProt ID A0A024E3A5 was used for subunit NDUFA9 to include a L174F mutation supported by the Drosophila cryo-EM density features. Notably, the N-terminus of core subunit NDUFS7 is resolved for the first time in Drosophila complex I (Figure 1—figure supplement 5). Preliminary model building and real-space refinements were carried out in Coot 0.9-pre (Casañal et al., 2020). Then GPU-powered ISOLDE 1.0 (Croll, 2018), which implements a molecular dynamic approach to model refinement, was used to iterate through the model, improving the map-to-model fit, resolving clashes and maintaining good protein stereochemistry. Emerging modelling errors were monitored using a real-time validation functionality present in ISOLDE and corrected. Densities for existing and additional phospholipid molecules were identified with the Unmodelled blobs tool in Coot 0.9.6.2 (Casañal et al., 2020). All non-cardiolipin phospholipids were modelled as phosphatidylethanolamines, the largest component of the phospholipid composition of the Drosophila mitochondrial membranes (Jones et al., 1992), unless density features indicated phosphatidylcholine to be more likely. Lipid tails were clipped where necessary using the delete tools in Coot and PyMOL 2.5.2 (Schrodinger, 2022). dGTP was modelled in subunit NDUFA10 (Molina-Granada et al., 2022). The model was then Curlew all-atom-refined using Coot and real-space refined against the active-state map using phenix.real_space_refine in Phenix 1.18.2–3874 (Liebschner et al., 2019) with custom geometry restraints. Ligand restraints were generated using Phenix eLBOW. No secondary structure restraints were used during real-space refinement of the Dm1 model. The model was checked manually in Coot, new resolvable regions built, and rotameric and/or Ramachandran outliers corrected. Atom resolvabilities (Q-scores) were calculated using MapQ (Pintilie et al., 2020) and any persisting outliers identified and corrected. The model was then real-space refined in Phenix as described above to produce the final Dm1 model.

To build the Dm2 model, the Dm1 Drosophila model was rigid-body fitted into the Dm2 map using the Fit in map tool in UCSF ChimeraX (Pettersen et al., 2021) followed by rigid-body fitting by subunit in Phenix 1.18.2–3874, and Curlew all-atom-refined using Coot 0.9.6.2. The Dm2 model was manually inspected and new resolvable regions, less resolved regions, and/or conformationally different regions built or deleted manually in Coot, and locally refined in ISOLDE 1.4 (Croll, 2018). Q-scores were calculated using MapQ, and any outliers identified and corrected. Existing lipids were checked against their densities and deleted where appropriate; lipid tails were similarly clipped where necessary as described above. The model was then real-space refined against the Dm2 map in Phenix 1.18.2–3874 with custom geometry restraints and secondary structure restraints (identified by ksdssp). Iteratively, rotameric and Ramachandran outliers were corrected manually in Coot and real-space refined in Phenix. The final real-space refinement for the Dm2 model was performed without secondary structure restraints in Phenix. The model statistics for the Dm1 and Dm2 classes (Table 1) were produced by Phenix, MolProbity (Chen et al., 2010), and EMRinger (Barad et al., 2015). Model-to-map FSC curves were generated using phenix.validation_cryoem in Phenix.

Individual subunits from the Dm2 model were rigid-body fitted into the Dm3 map in UCSF ChimeraX (Pettersen et al., 2021) to generate a tentative model for the Dm3 state for visualisation.

Cryo-EM model analyses

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RMSD calculations between models were performed using the Align command in PyMOL 2.5.2 (Schrodinger, 2022). Buried surface area between subunits were calculated using the measure buriedarea command in UCSF ChimeraX (Pettersen et al., 2021). The interior surface of the ubiquinone-binding channel was predicted using CASTp (Tian et al., 2018), which computes a protein surface topology from a PDB model. The default 1.4 Å radius probe was used and the results were visualised in PyMOL using the CASTpyMOL 3.1 plugin and by UCSF ChimeraX.

Data availability

Structural data have been deposited in the EMDB and PDB databases under the following accession codes: EMD-15936 and 8B9Z (Dm1; active), EMD-15937 and 8BA0 (Dm2; twisted), and EMD-15938 (Dm3; cracked).

The following data sets were generated
    1. Agip A-NA
    2. Chung I
    3. Sanchez-Martinez A
    4. Whitworth AJ
    5. Hirst J
    (2022) RCSB Protein Data Bank
    ID 8B9Z. Drosophila melanogaster complex I in the Active state (Dm1).
    https://www.rcsb.org/structure/8B9Z
    1. Agip A-NA
    2. Chung I
    3. Sanchez-Martinez A
    4. Whitworth AJ
    5. Hirst J
    (2022) EMDataResource
    ID EMD-15936. Drosophila melanogaster complex I in the Active state (Dm1).
    https://www.ebi.ac.uk/emdb/EMD-15936
    1. Agip A-NA
    2. Chung I
    3. Sanchez-Martinez A
    4. Whitworth AJ
    5. Hirst J
    (2022) RCSB Protein Data Bank
    ID 8BA0. Drosophila melanogaster complex I in the Twisted state (Dm2).
    https://www.rcsb.org/structure/8BA0
    1. Agip A-NA
    2. Chung I
    3. Sanchez-Martinez A
    4. Whitworth AJ
    5. Hirst J
    (2022) EMDataResource
    ID EMD-15937. Drosophila melanogaster complex I in the Twisted state (Dm2).
    https://www.ebi.ac.uk/emdb/EMD-15937
    1. Agip A-NA
    2. Chung I
    3. Sanchez-Martinez A
    4. Whitworth AJ
    5. Hirst J
    (2022) EMDataResource
    ID EMD-15938. Drosophila melanogaster complex I in the Cracked state (Dm3).
    https://www.ebi.ac.uk/emdb/EMD-15938

References

Decision letter

  1. Volker Zickermann
    Reviewing Editor; Goethe University Frankfurt, Germany
  2. Volker Dötsch
    Senior Editor; Goethe University, Germany
  3. Yongchan Lee
    Reviewer; Max Planck Institute of Biophysics, Germany
  4. Hans-Peter Braun
    Reviewer; Leibniz Universität Hannover, Germany

Our editorial process produces two outputs: (i) public reviews designed to be posted alongside the preprint for the benefit of readers; (ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Decision letter after peer review:

Thank you for submitting your article "Cryo-EM structures of mitochondrial respiratory complex I from Drosophila melanogaster" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Volker Dötsch as the Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Yongchan Lee (Reviewer #2); Hans-Peter Braun (Reviewer #3).

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

Essential revisions:

The reviewers requested to address a few issues to improve the manuscript. I would like to highlight and add a few points here.

1. The activity of the Dm complex I preparation is somewhat hidden in the Materials and methods section. Please report the inhibitor sensitive activity in the text. To allow better comparison with other preparations please give the value in µmol min-1 mg-1 and electrons s-1.

2. The NEM assay alone might not be sufficient to exclude that the A/D transition is completely absent in Dm complex I, especially because Dm2 is considered to be a relaxed or deactive-like state. Please show the traces of activity measurements or report if a lag phase exists or not. The A/D transition should be also tested by monitoring the impact of divalent cations when added before or after start of the activity measurement.

3. Reviewer 1 pointed out that the assignments of Dm1 and Dm2 need more discussion. This is a critical point. Dm2 shows two features of complex I in the deactive state, but you exclude that Drosphila complex I undergoes an A/D transition. These two statements are difficult to reconcile. Isn´t that at least suggesting that the pi-bulge of TMH3 of ND6 is actually not a hallmark of the D state? Please discuss.

4. It is discussed that in Dm2, a lack of transmission from the pi-bulge to the Q binding site argues against a direct link between the two being crucial for catalysis. This appears not fully justified because very likely the water chain connecting Q tunnel and hydrophilic axis is interrupted. It is clear that at the given resolution this cannot be observed here, but maybe you want to add a sentence on this point.

Reviewer #1 (Recommendations for the authors):

– The authors describe Dm1 as the active resting state. What makes it resting? How would it differ from a 'turnover' state?

– Also Dm2 structurally resembles the active state of the mammalian Complex I, but with some characteristic of the deactive form, such as the pi-bulge but not the ND3 loop or NDUFS2. Why is Dm2 not the active resting state?

– I believe that general reader might find the discussion of active / deactive / deactive-type / opening / twisted confusing and would benefit from a clarification in the Discussion section.

– What interactions stabilize the quinone headgroup in the modelled position? Are homologous residues present in the other isoform (with different resolved quinone positions)?

– "there is no apparent opening or closing of the angle between the domains" – could the authors estimate the angles to show that they are the same?

– Purification: could DDM bind to the quinone cavity and lead to partial inhibition of the activity or block expected conformational changes? I believe that this has been seen in other Complex I structures. Did the authors characterize the activity in other detergents?

– Previous data indicate that cardiolipin binds to Complex I, but I could not see any modeled cardiolipin molecules. Could the authors please comment on this?

Related to this point, the method section say: "The methods say all non-cardiolipin molecules were modelled", but there is no discussion of the cardiolipin molecules.

– Motion around NDUFA5/10 – the authors discuss that these subunits have different conformation in the mammalian active and deactive states, but it is not evident to me why it is expected that these subunits would affect e.g. the twisting shown in Figure 2?

– Activity: for the general reader, please provide a comparison to activities for example for the mammalian enzyme in the active and deactive states. Is the measured activity sensitive to known Complex I inhibitors?

– I understand that no water molecules could be resolved at this resolution, but the authors could comment on whether they find evidence that the conformational changes in Dm1 and Dm2 could lead to hydration changes and in this way affect the proton translocation?

Reviewer #2 (Recommendations for the authors):

– Would sample pre-treatment, such as heat and the addition of NADH or ubiquinone, change the ratio of Dm1 and Dm2 states, which might inform about the nature of these conformations?

https://doi.org/10.7554/eLife.84424.sa1

Author response

Essential revisions:

The reviewers requested to address a few issues to improve the manuscript. I would like to highlight and add a few points here.

1. The activity of the Dm complex I preparation is somewhat hidden in the Materials and methods section. Please report the inhibitor sensitive activity in the text. To allow better comparison with other preparations please give the value in µmol min-1 mg-1 and electrons s-1.

We have amended the Results section to include this information on page 5: “The highest concentration peak fraction (3.4 mg ml-1), which exhibited an NADH oxidoreductase activity comparable to mammalian complex I of 7.3 ± 0.3 µmol min-1 mg-1 (ca. 120 NADH s-1) was collected and frozen…” As stated in Materials and methods we confirmed that the activity is sensitive to both rotenone and piericidin.

2. The NEM assay alone might not be sufficient to exclude that the A/D transition is completely absent in Dm complex I, especially because Dm2 is considered to be a relaxed or deactive-like state. Please show the traces of activity measurements or report if a lag phase exists or not. The A/D transition should be also tested by monitoring the impact of divalent cations when added before or after start of the activity measurement.

The NEM assay is only sufficient to conclude that Cys41 on the ND3-TMH1-2 loop is not exposed in Drosophila complex I, and that it does not become exposed upon heat treatments that are sufficient to deactive the mammalian enzyme. The NEM assay does not probe any other elements of the mammalian deactive transition, such as the ND6-TMH3 π-bulge. The NEM assay is useful for investigating the active/deactive status of the mammalian enzyme because there is a single transition between the two states, in which all the elements convert together. We agree that, unfortunately, the NEM assay is therefore not suitable for probing the conversion between Dm1 and Dm2 and we have now checked our manuscript to ensure that we have been clear on this point (notably, on page 12). We are aware, of course, that two other biochemical characteristics are associated with the mammalian deactive state. First, we have now revisited our assay traces for Drosophila complex I in mitochondria, membranes and isolated in detergent, and we can discern no clear evidence of any catalytic lag phase. Although we regard this as consistent with the fully formed ubiquinone-binding sites in all the Dm1, Dm2 and Dm3 states (no ‘refolding’ lag phase is required) we have not conducted sufficiently focussed investigations to make a definitive statement. Second, we have not yet carried out investigations of the effects of divalent cations on catalysis by Drosophila complex I. We are aware that divalent cations have been used successfully with complex I from Yarrowia lipolytica, but in our experiments on the mammalian enzyme we have not found this characteristic easy to reproduce or to distinguish from inhibition satisfactorily, and our experiments on the Drosophila enzyme were severely limited by the amount of material that was reasonably available. Unfortuantely, we do not currently have the resources available to produce the quantitites of enzyme that would be required for this study.

3. Reviewer 1 pointed out that the assignments of Dm1 and Dm2 need more discussion. This is a critical point. Dm2 shows two features of complex I in the deactive state, but you exclude that Drosphila complex I undergoes an A/D transition. These two statements are difficult to reconcile. Isn´t that at least suggesting that the pi-bulge of TMH3 of ND6 is actually not a hallmark of the D state? Please discuss.

As we have noted in our answer to reviewer 1, Dm1 contains all the features of the mammalian active resting state (Agip, 2018; Blaza et al., 2018; Chung et al., 2022b, 2022a; Zhu et al., 2016) and therefore we are confident in this assignment. Dm2 differs from Dm1 by the presence of a π-bulge in ND6-TMH3 and a flipped Tyr149 in ND1-TMH4, which are two features of the mammalian deactive resting state. However, the mammalian deactive resting state also exhibits a disordered ND3-TMH1-2 loop (i.e. exposed ND3-Cys39), disordered NDUFS2 β1-β2 loop, disordered ND1-TMH5-6 loop, bent ND1-TMH4 as well as a decreased interface area between NDUFA5 and 10 – none of these features are present in Dm2. Dm2 can therefore be considered intermediate between the mammalian active and deactive states, and we therefore chose to give it a different name (‘Twisted’). We described the Twisted state as a ‘restricted’ or ‘curtailed’ deactive state to attempt to capture the suggestion that it has partially converted to a mammalian-type deactive state. Whether Dm2 should be referred to as the ‘Drosophila-deactive state’ and whether we say Dm2 is formed in a ‘Drosophila-A/D transition’ is a matter of semantics. Furthermore, structural elements that move in the mammalian A/D transition, including the π-bulge, are also likely mobile during catalysis and so (although it is not our preferred explanation) we cannot discount that Dm2 is the result of pausing catalysis at a different point on the catalytic cycle than Dm1.

4. It is discussed that in Dm2, a lack of transmission from the pi-bulge to the Q binding site argues against a direct link between the two being crucial for catalysis. This appears not fully justified because very likely the water chain connecting Q tunnel and hydrophilic axis is interrupted. It is clear that at the given resolution this cannot be observed here, but maybe you want to add a sentence on this point.

We apologise that our text was not clear on this point. We intended to observe that the π-bulge can form without changing the structure of the ubiquinone-binding site, as demonstrated by comparison of our Dm1 and Dm2 structures. This is contrary to, for example, the mechanism proposed by Kravchuk et al. that requires a clean switch between two states (destructured Q-site elements plus π-bulge vs. structured and closed Q-site plus α-helix) in order to avoid the formation of the destructured:α-helix combination, which it is proposed would incur proton leak. Although we have not observed this particular state, our Dm2 state demonstrates that a clean switch between the two states described above is not structurally enforced. We have now amended our text on page 13 to be more clear on this point.

Reviewer #1 (Recommendations for the authors):

– The authors describe Dm1 as the active resting state. What makes it resting? How would it differ from a 'turnover' state?

No substrates (NADH or quinone) were added to our cryo-EM sample so that the complex I is not undergoing catalysis and therefore is ‘at rest’. We use the term resting state to clearly differentiate the complex I structures in our preparation from the catalytic intermediates that have been described in the literature to be present in ‘turnover’ samples in the presence of substrates. As we describe, the active resting state refers to the ‘ready-to-go’ resting state; this is the terminology that is used throughout the complex I field.

– Also Dm2 structurally resembles the active state of the mammalian Complex I, but with some characteristic of the deactive form, such as the pi-bulge but not the ND3 loop or NDUFS2. Why is Dm2 not the active resting state?

Whereas Dm1 contains all the features of the mammalian active resting state (Agip, 2018; Blaza et al., 2018; Chung et al., 2022b, 2022a; Zhu et al., 2016), Dm2 deviates from it by the presence of a π-bulge in ND6-TMH3 and a flipped Tyr149 in ND1-TMH4. Dm2 is therefore not equivalent to the active resting state (or to the deactive resting state) and we therefore chose to give it a different name (‘Twisted’).

– I believe that general reader might find the discussion of active / deactive / deactive-type / opening / twisted confusing and would benefit from a clarification in the Discussion section.

It is unfortunate that nomenclature in the complex I field has recently become more confusing because different groups have opted to use different naming systems and have also made different assignments to biochemically characterised states. First, we have clearly described the active/deactive and closed/open states, and the relationships between them, in our introduction (paragraph 3). As noted above, our twisted state is intermediate between the active and deactive resting states. We realise that our use of ‘deactive-type state’ in the Discussion was confusing, and no longer use this term.

– What interactions stabilize the quinone headgroup in the modelled position? Are homologous residues present in the other isoform (with different resolved quinone positions)?

The quinone observed in the Dm1 structure, including its headgroup, is largely stabilised by hydrophobic interactions as indicated in Figure 3 —figure supplement 1. There are no other isoforms present in our structures. We note that no quinones were resolved in the Dm2 or Dm3 states, and that the region of the ubiquinone-binding site in question is predominantly populated by hydrophobic residues in all complex I structures.

– "there is no apparent opening or closing of the angle between the domains" – could the authors estimate the angles to show that they are the same?

‘Opening’ and ‘Closing’ are poorly defined descriptions of complex I conformational transitions that are widely used in the complex I field. The conformational transitions are in fact not represented by a single two dimensional angle but are the result of complicated three dimensional motion that, nevertheless, looks from the side like an opening and closing. We therefore opted for the term ‘apparent opening and closing’. To clarify this further we now no longer specifically refer to the angle between the domains.

– Purification: could DDM bind to the quinone cavity and lead to partial inhibition of the activity or block expected conformational changes? I believe that this has been seen in other Complex I structures. Did the authors characterize the activity in other detergents?

So far, DDM binding has only been observed in the ubiquinone-binding site of the deactive state in bovine (Chung et al., 2022b) and yeast (Grba and Hirst, 2020) complex I. Based on the cryo-EM densities observed in the ubiquinone-binding sites here, there is no clear evidence of DDM binding to any of the three Drosophila complex I structures. In addition, all three states contain a closed ubiquinone-binding site, and no DDM has been observed bound to a fully closed state. The question of whether DDM may bind to inhibit catalysis in other states (that are not present in our structural analysis) is hard to answer but the activity of our complex I purified in DDM was substantial (7.3 µmol min-1 mg-1), offering no support for it being inhibited. Please also see the comment below that specifically addresses the activity value.

– Previous data indicate that cardiolipin binds to Complex I, but I could not see any modeled cardiolipin molecules. Could the authors please comment on this?

Related to this point, the method section say: "The methods say all non-cardiolipin molecules were modelled", but there is no discussion of the cardiolipin molecules.

Cardiolipin molecules have been modelled in the structure(s), but were not referred to in any particular detail because they were found in regions already observed in other complex I structures and not of obvious functional or catalytic significance. We have now indicated the modelled cardiolipins in Figure 1 —figure supplement 4.

– Motion around NDUFA5/10 – the authors discuss that these subunits have different conformation in the mammalian active and deactive states, but it is not evident to me why it is expected that these subunits would affect e.g. the twisting shown in Figure 2?

We assume that by ‘excepted’, the reviewer means ‘expected’. NDUFA5 and 10 form one of the key interfaces between the hydrophilic and membrane domains. NDUFA5 is part of the hydrophilic domain whereas NDUFA10 is part of the membrane domain. Due to the apparent closing/opening of the two domains, where they move relative to each other, in the mammalian active/deactive transition, the NDUFA5/10 interface and contact surface areas change. In Drosophila, between the active resting state and the twisted state, the hydrophilic and membrane domains also move relative to each other, and therefore the NDUFA5/10 interface and contact surface areas change. Interestingly, the structurally ordered N-terminus of NDUFS4 specific to Drosophila complex I stabilises the NDUFA5/10 interface in Dm1 as shown in Figure 4. Disordering of this N-terminal tether in Dm2 therefore disrupts this stabilised interface, assisting the hydrophilic and membrane domains twisting against each other as shown in Figure 2.

– Activity: for the general reader, please provide a comparison to activities for example for the mammalian enzyme in the active and deactive states. Is the measured activity sensitive to known Complex I inhibitors?

The activity of complex I from Drosophila (7.3 µmol min-1 mg-1) is comparable that of ovine complex I (5-6 µmol min-1 mg-1) (Kampjut and Sazanov, 2020; Letts et al., 2016), but lower than murine (10-12 µmol min-1 mg-1) (Agip et al., 2018) or bovine (20-24 µmol min-1 mg-1) (Chung et al., 2022b; Wright et al., 2022) complex I. We have amended the Results section to include this information on page 5: “The highest concentration peak fraction (3.4 mg ml-1), which exhibited an NADH oxidoreductase activity comparable to mammalian complex I of 7.3 ± 0.3 µmol min-1 mg-1 (ca. 120 NADH s-1), was collected and frozen…”. Due to very limited sample availability from fly preparations we were unable to conduct extensive inhibitor investigations, however, we did confirm that catalysis is sensitive to addition of rotenone and piericidin (now noted on page 18).

– I understand that no water molecules could be resolved at this resolution, but the authors could comment on whether they find evidence that the conformational changes in Dm1 and Dm2 could lead to hydration changes and in this way affect the proton translocation?

The formation of a π-bulge in ND6-TMH3 in the deactive state is known to block the connection between the E-channel and the central hydrophilic axis of charged residues, as reported previously by many structures of complex I, including mammalian, yeast, and bacterial species (Agip et al., 2018; Blaza et al., 2018; Chung et al., 2022b; Grba and Hirst, 2020; Kampjut and Sazanov, 2020; Kravchuk et al., 2022; Parey et al., 2021). In addition, ND1-TMH4-Tyr149 points into (mammalian active state) or away from (mammalian deactive state) the E-channel, affecting the connectivity between the ND1 cavity (extending from the ubiquinone-binding site) and the neighbouring ND1-Glu150 and ND3-Asp68 (towards ND3) (Chung et al., 2022b). Both features are conserved here, between Dm1 and Dm2, and therefore the same changes in hydration and proton connectivity are expected. We have now added a sentence on page 12 to note this: “Although water molecules cannot be resolved in our structures, these changes are expected to alter the connectivity between the E-channel and the central axis of charged residues along the membrane domain as reported previously for mammalian, yeast, and bacterial species (Agip et al., 2018; Blaza et al., 2018; Chung et al., 2022b; Grba and Hirst, 2020; Kampjut and Sazanov, 2020; Kravchuk et al., 2022; Parey et al., 2021).”.

Reviewer #2 (Recommendations for the authors):

- Would sample pre-treatment, such as heat and the addition of NADH or ubiquinone, change the ratio of Dm1 and Dm2 states, which might inform about the nature of these conformations?

Indeed, it is possible that the ratio of Dm1 and Dm2 may change upon heating or addition of substrates, which may inform us of the biochemical relevance of these states. However, this is not within the scope of our study, which focuses only on the resting state enzymes. We agree that it will be interesting to pre-treat Drosophila complex I to change its biochemical status before performing single particle cryo-EM in our future studies.

https://doi.org/10.7554/eLife.84424.sa2

Article and author information

Author details

  1. Ahmed-Noor A Agip

    The Medical Research Council Mitochondrial Biology Unit, University of Cambridge, The Keith Peters Building, Cambridge Biomedical Campus, Cambridge, United Kingdom
    Contribution
    Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Writing - review and editing
    Contributed equally with
    Injae Chung and Alvaro Sanchez-Martinez
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3020-8262

  2. Injae Chung

    The Medical Research Council Mitochondrial Biology Unit, University of Cambridge, The Keith Peters Building, Cambridge Biomedical Campus, Cambridge, United Kingdom
    Contribution
    Data curation, Formal analysis, Validation, Investigation, Visualization, Writing - original draft, Writing - review and editing
    Contributed equally with
    Ahmed-Noor A Agip and Alvaro Sanchez-Martinez
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-2902-4677

  3. Alvaro Sanchez-Martinez

    The Medical Research Council Mitochondrial Biology Unit, University of Cambridge, The Keith Peters Building, Cambridge Biomedical Campus, Cambridge, United Kingdom
    Contribution
    Investigation, Methodology, Writing - review and editing
    Contributed equally with
    Ahmed-Noor A Agip and Injae Chung
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-2728-6251

  4. Alexander J Whitworth

    The Medical Research Council Mitochondrial Biology Unit, University of Cambridge, The Keith Peters Building, Cambridge Biomedical Campus, Cambridge, United Kingdom
    Contribution
    Conceptualization, Supervision, Funding acquisition, Project administration, Writing - review and editing
    For correspondence
    a.whitworth@mrc-mbu.cam.ac.uk
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-1154-6629

  5. Judy Hirst

    The Medical Research Council Mitochondrial Biology Unit, University of Cambridge, The Keith Peters Building, Cambridge Biomedical Campus, Cambridge, United Kingdom
    Contribution
    Conceptualization, Formal analysis, Supervision, Funding acquisition, Investigation, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    jh@mrc-mbu.cam.ac.uk
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-8667-6797

Funding

Medical Research Council (MC_UU_00015/6)

  • Alexander J Whitworth

Medical Research Council (MC_UU_00028/6)

  • Alexander J Whitworth

Medical Research Council (MC_UU_00015/2)

  • Judy Hirst

Medical Research Council (MC_UU_00028/1)

  • Judy Hirst

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

Acknowledgements

We thank D Chirgadze (University of Cambridge Cryo-EM facility) for assistance with grid screening and cryo-EM data collection; T Croll (Cambridge Institute for Medical Research) for assistance with ISOLDE and I M Fearnley and S Ding (MRC MBU) for mass spectrometry analyses. This work was supported by the Medical Research Council (MC_UU_00015/6 and MC_UU_00028/6 to AJW and MC_UU_00015/2 and MC_UU_00028/1 to JH). Drosophila were obtained from the Bloomington Drosophila Stock Center, which is supported by grant NIH P40OD018537.

Senior Editor

  1. Volker Dötsch, Goethe University, Germany

Reviewing Editor

  1. Volker Zickermann, Goethe University Frankfurt, Germany

Reviewers

  1. Yongchan Lee, Max Planck Institute of Biophysics, Germany
  2. Hans-Peter Braun, Leibniz Universität Hannover, Germany

Version history

  1. Received: October 24, 2022
  2. Preprint posted: November 1, 2022 (view preprint)
  3. Accepted: January 6, 2023
  4. Accepted Manuscript published: January 9, 2023 (version 1)
  5. Version of Record published: March 1, 2023 (version 2)

Copyright

© 2023, Agip, Chung, Sanchez-Martinez et al.

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  1. Ahmed-Noor A Agip
  2. Injae Chung
  3. Alvaro Sanchez-Martinez
  4. Alexander J Whitworth
  5. Judy Hirst
(2023)
Cryo-EM structures of mitochondrial respiratory complex I from Drosophila melanogaster
eLife 12:e84424.
https://doi.org/10.7554/eLife.84424

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Further reading

    1. Structural Biology and Molecular Biophysics

    Resting mitochondrial complex I from Drosophila melanogaster adopts a helix-locked state

    Abhilash Padavannil, Anjaneyulu Murari ... James A Letts
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    Respiratory complex I is a proton-pumping oxidoreductase key to bioenergetic metabolism. Biochemical studies have found a divide in the behavior of complex I in metazoans that aligns with the evolutionary split between Protostomia and Deuterostomia. Complex I from Deuterostomia including mammals can adopt a biochemically defined off-pathway ‘deactive’ state, whereas complex I from Protostomia cannot. The presence of off-pathway states complicates the interpretation of structural results and has led to considerable mechanistic debate. Here, we report the structure of mitochondrial complex I from the thoracic muscles of the model protostome Drosophila melanogaster. We show that although D. melanogaster complex I (Dm-CI) does not have a NEM-sensitive deactive state, it does show slow activation kinetics indicative of an off-pathway resting state. The resting-state structure of Dm-CI from the thoracic muscle reveals multiple conformations. We identify a helix-locked state in which an N-terminal α-helix on the NDUFS4 subunit wedges between the peripheral and membrane arms. Comparison of the Dm-CI structure and conformational states to those observed in bacteria, yeast, and mammals provides insight into the roles of subunits across organisms, explains why the Dm-CI off-pathway resting state is NEM insensitive, and raises questions regarding current mechanistic models of complex I turnover.

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    The neuronal calcium sensor NCS-1 regulates the phosphorylation state and activity of the Ga chaperone and GEF Ric-8A

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    The Neuronal Calcium Sensor 1, an EF-hand Ca2+ binding protein, and Ric-8A coregulate synapse number and probability of neurotransmitter release. Recently, the structures of Ric-8A bound to Ga have revealed how Ric-8A phosphorylation promotes Ga recognition and activity as a chaperone and guanine nucleotide exchange factor. However, the molecular mechanism by which NCS-1 regulates Ric-8A activity and its interaction with Ga subunits is not well understood. Given the interest in the NCS-1/Ric-8A complex as a therapeutic target in nervous system disorders, it is necessary to shed light on this molecular mechanism of action at atomic level. We have reconstituted NCS-1/Ric-8A complexes to conduct a multimodal approach and determine the sequence of Ca2+ signals and phosphorylation events that promote the interaction of Ric-8A with Ga. Our data show that the binding of NCS-1 and Ga to Ric-8A are mutually exclusive. Importantly, NCS-1 induces a structural rearrangement in Ric-8A that traps the protein in a conformational state that is inaccessible to Casein Kinase II-mediated phosphorylation, demonstrating one aspect of its negative regulation of Ric-8A-mediated G-protein signaling. Functional experiments indicate a loss of Ric-8A GEF activity towards Ga when complexed with NCS-1, and restoration of nucleotide exchange activity upon increasing Ca2+ concentration. Finally, the high-resolution crystallographic data reported here define the NCS-1/Ric-8A interface and will allow the development of therapeutic synapse function regulators with improved activity and selectivity.

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    Baited reconstruction with 2D template matching for high-resolution structure determination in vitro and in vivo without template bias

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    Previously we showed that 2D template matching (2DTM) can be used to localize macromolecular complexes in images recorded by cryogenic electron microscopy (cryo-EM) with high precision, even in the presence of noise and cellular background (Lucas et al., 2021; Lucas et al., 2022). Here, we show that once localized, these particles may be averaged together to generate high-resolution 3D reconstructions. However, regions included in the template may suffer from template bias, leading to inflated resolution estimates and making the interpretation of high-resolution features unreliable. We evaluate conditions that minimize template bias while retaining the benefits of high-precision localization, and we show that molecular features not present in the template can be reconstructed at high resolution from targets found by 2DTM, extending prior work at low-resolution. Moreover, we present a quantitative metric for template bias to aid the interpretation of 3D reconstructions calculated with particles localized using high-resolution templates and fine angular sampling.