ISSN 0006-2979, Biochemistry (Moscow), 2024, Vol. 89, No. 2, pp. 257-268 © Pleiades Publishing, Ltd., 2024.
257
Safari with an Electron Gun:
Visualization of Protein and Membrane Interactions
in Mitochondria in Natural Environment
Semen V. Nesterov
1,a
*, Konstantin S. Plokhikh
1
,
Yuriy M. Chesnokov
1
,
Denis A. Mustafin
1
, Tatyana N. Goleva
1
, Anton G. Rogov
1
,
Raif G. Vasilov
1
, and Lev S. Yaguzhinsky
2
1
National Research Center “Kurchatov Institute,” 123182 Moscow, Russia
2
Belozersky Research Institute for Physico-Chemical Biology, Lomonosov Moscow State University,
119992 Moscow, Russia
a
e-mail: semen.v.nesterov@phystech.edu
Received September 24, 2023
Revised February 6, 2024
Accepted February 7, 2024
Abstract— This paper presents new structural data about mitochondria using correlative light and electron mi-
croscopy (CLEM) and cryo-electron tomography. These state-of-the-art structural biology methods allow studying
biological objects at nanometer scales under natural conditions. Non-invasiveness of these methods makes them
comparable to observing animals in their natural environment on a safari. The paper highlights two areas of
research that can only be accomplished using these methods. The study visualized location of the Aβ42 amy-
loid aggregates in relation to mitochondria to test a hypothesis of development of mitochondrial dysfunction in
Alzheimer’s disease. The results showed that the Aβ42 aggregates do not interact with mitochondria, although
some of them are closely located. Therefore, the study demonstrated that mitochondrial dysfunction is not direct-
ly associated with the effects of aggregates on mitochondrial structure. Other processes should be considered as
sources of mitochondrial dysfunction. Second unique area presented in this work is high-resolution visualization
of the mitochondrial membranes and proteins in them. Analysis of the cryo-ET data reveals toroidal holes in the
lamellar structures of cardiac mitochondrial cristae, where ATP synthases are located. The study proposes a new
mechanism for sorting and clustering protein complexes in the membrane based on topology. According to this
suggestion, position of the OXPHOS system proteins in the membrane is determined by its curvature. High-res-
olution tomography expands and complements existing ideas about the structural and functional organization
ofmitochondria. This makes it possible to study the previously inaccessible structural interactions of proteins with
each other and with membranes invivo.
DOI: 10.1134/S0006297924020068
Keywords: membrane, mitochondria, oxidative phosphorylation, cryo-electron microscopy, supercomplex, ATP
synthase, respirasome, Aβ42, amyloid aggregates
Abbreviations: ATP,adenosine triphosphate; CLEM,correlative light and electron microscopy; cryo-EM,cryogenic electron
microscopy; cryo-ET, cryogenic electron tomography; OXPHOS, oxidative phosphorylation; TEM, transmission electron
microscopy.
* To whom correspondence should be addressed.
INTRODUCTION
In recent years, cryogenic-electron microscopy
(cryo-EM) techniques have undergone significant im-
provements in both hardware and software. These im-
provements have led to a breakthrough in structural
biology with increased availability and comprehen-
sive image processing capabilities [1-3]. While many
NESTEROV et al.258
BIOCHEMISTRY (Moscow) Vol. 89 No. 2 2024
microscopy studies are investigating isolated proteins
with high resolution, we will narrow our focus on
macromolecular organization of the enzyme systems
in mitochondrial membranes in whole mitochondria
without isolating proteins from them. This article
demonstrates capabilities of the correlative light and
electron microscopy (CLEM) [4]. CLEM, in combination
with genetic engineering, allows visualization of inter-
actions between any proteins of interest, as well as be-
tween proteins and membranes, with a resolution of a
few nanometers. CLEM outperforms solely light-based
techniques in terms of resolution, including super-res-
olution microscopy such as STED microscopy [5]. STED
microscopy allows visualization of membranes but not
individual proteins within them [6]. In this work, we
present our new results, deeply analyze a number of
tomographic data obtained in our previous works, and
discuss the literature devoted to “photoelectron hunt-
ing” of mitochondrial proteins in ‘wild’ natural con-
ditions of their existence in protein–lipid membranes.
Despite the considerable amount of structural
data on mitochondrial proteins, the data on their na-
tive positions in the membrane and matrix are very
limited, which makes it difficult to form an integral
model of the work of these organelles. Recently, cryo-
EM has helped to fill this gap in our understanding.
For example, it has been found that adenosine triphos-
phate (ATP) synthases in the mammalian mitochon-
dria are located at the folds of the inner membrane
(cristae), while components of the respiratory chain
are located in the less curved parts of the membrane.
Additionally, much information has emerged on the
structural interaction of respiratory chain complexes
(respirasome). However, many questions still remain
unanswered. Standard laboratory methods for extract-
ing respirasomes with a mild detergent cannot provide
reliable information on the percentage of respirato-
ry chain complexes that form respirasomes. This is
because some complexes disintegrate during the ex-
traction process, and some could be not extracted from
the membrane. Simultaneously, a detailed analysis of
cryo-EM tomograms enables us to solve this problem.
For instance, we demonstrated that all complexes I
and III are part of respirasomes in the rat heart [7].
Data from numerous scientific groups indicate that
proton transfer between the proton pumps and ATP
synthase occurs not due to electrochemical gradient
between the matrix and intermembrane space vol-
umes, but rather occurs locally with protons moving
laterally along the membrane surface for short dis-
tances. A detailed study of the structure of mitochon-
drial membranes and proteins can provide unique
information necessary for testing existing hypotheses
and modeling proton transfer processes in the oxida-
tive phosphorylation system (OXPHOS) [8]. Additional-
ly, many structural aspects of interaction between mi-
tochondria and other organelles or various aggregates
are still unknown. Even interaction of amyloid aggre-
gates with mitochondrial membrane remains poorly
investigated despite high prevalence of Alzheimer’s
disease. However, colocalization of the fluorescent tags
does not provide sufficient resolution for unambigu-
ous conclusions. In this study, we were able to answer
this question using correlative light-electron microscopy.
MATERIALS AND METHODS
Cultivation of yeast cells. Yarrowia lipolytica Po1f
yeast expressing eGFP-Aβ42 construct [9] were used
in this work. Cells were grown in 250-ml Erlenmeyer
flasks at 28°C on a rotary shaker (220rpm) in 50ml of
semi-synthetic medium containing 1.3% succinate as a
carbon and energy source. Cells were harvested in the
early exponential growth phase (OD
600
=1.0).
Preparation of samples for cryo-EM. Cells were
rinsed with 50 mM phosphate buffer (pH = 5.5) to re-
move the growth medium. Next, they were incubat-
ed with 500 nM MitoTracker Red CMXRos for 30 min.
Afterward, the cells were washed to remove the dye
and concentrated to OD
600
= 25. To minimize formation
of ice crystals, 5% glycerol was added to the suspen-
sion before plating it on a cryo-EM grid [10]. Electron
microscopy grids underwent hydrophilization using
a Pelco easiGlow unit (USA) at 25 mA current and
0.26 mBar pressure for 30 s. Next, a 5-μl aliquot of cell
suspension was applied to a microscopy grid. Extra liq-
uid from the grids with the test objects was removed
from both sides using filter paper and next grids
were subjected to quick freezing in a liquefied ethane
cooled to liquid nitrogen temperature. When applying
the sample to the grid, temperature in the chamber
of the Thermo Fisher Scientific Vitrobot system (USA)
was maintained at 4°C, and humidity was at least 95%.
If homogenate of cardiac tissue or isolated rat heart
mitochondria were used, a similar grid preparation
procedure was performed, but without the use of glyc-
erol, as described in the previously published works
[7, 11]. Briefly, 3 µl of homogenized Wistar rat heart
tissue or isolated cardiac mitochondria (isolated us-
ing differential centrifugation, without the use of de-
tergents or proteases, as previously described [12])
were applied to a microscopy grid and vitrified using
aVitrobot system.
Fluorescence microscopy. Vitrified yeast samples
were placed in a cryogenic chamber of a Leica THUNDER
Imager EM Cryo CLEM fluorescence microscope (Ger-
many) equipped with a Leica HC PL APO 50×/0.90 CRYO
CLEM objective (Germany). Fluorescence of eGFP-Aβ42
and MitoTracker Red was recorded using appropriate
filters, and colocalization of eGFP-Aβ42 and mitochon-
drial aggregates were assessed. The Leica Thunder
MITOCHONDRIAL PROTEINS AND MEMBRANES IN NATURAL ENVIRONMENT 259
BIOCHEMISTRY (Moscow) Vol. 89 No. 2 2024
algorithm was used to perform 3D reconstruction of
yeast cells, followed by image deconvolution with the
aid of embedded software [13]. Images of the entire
grid provided a “map” for orientation during lamella
etching. Regions of interest were selected based on the
images obtained from the entire grid.
Preparation of lamellae by focused ion beam
(FIB). The vitrified sample on microscopic grids was
transferred to a Thermo Fisher Scientific Versa 3D
scanning electron ion microscope (USA) equipped with
a cryoprobe and a Quorum 3100P liquid nitrogen tem-
perature sample loading system. To reduce charge ac-
cumulation and protect surface from radiation dam-
age during the experiment, a gas-injection system was
used to sputter a protective layer of platinum on the
entire grid surface. Based on the surface morphology
and fluorescence microscopy maps, we selected areas
for preparing thin cell slices using cryo-FIB (Fig. 1).
We have chosen regions closer to the center of the
grid where the cells were arranged in a single layer.
The lamellae were cut in these squares with expec-
tation of passing along a few cells located at a suffi-
cient distance from the metal framework of the grid.
Gallium ions with an energy of 30 keV were used to min-
imize radiation damage to the near-surface layer. The
current was sequentially reduced from 1nA to 30pA.
This approach produced object slices with 150-250nm
thickness. The slice plane was inclined at an angle of
10-12° with respect to the plane of the microscopic grid.
Cryo-EM and cryo-ET. Cryo-EM studies were per-
formed using a Thermo Fisher Scientific Titan Krios
60-300 cryogenic transmission electron microscope
(USA) at accelerating voltage of 300 kV, equipped with
a Falcon II direct electron detection system. For each
microscopic grid, sections were selected to obtain ro-
tating series of images. Each dataset consisted of 56
images taken by tilting the sample between –50° and
60° in 2° angular increments. Data acquisition was
performed automatically using a Thermo Fisher Scien-
tific Tomography 4 software (USA) in low dose mode,
which minimized radiation damage while preserving
native structure of the studied objects. Magnification
of 18,000× (pixel size 0.37 nm and image size 1516 nm)
with a defocus value in the range [–5 μm; –8 μm] was
used to set the tomographic series. Total electron dose
passing through a unit area of the sample during the
entire exposure time did not exceed ~120e
–
/Å
2
.
Data processing. Tomographic reconstruction
procedure using the Weighted Back Projections (WBP)
method in the IMOD software package [14] was applied
to the obtained data. Alignment of the rotated series
of cell images was performed by calculating cross-cor-
relation between the regions of the tomographic series
(the “patch tracking” procedure [15]), since it is impos-
sible to introduce colloidal gold nanoparticles into the
sample. In the case of studying fragments of the iso-
lated mitochondria, colloidal gold nanoparticles with
10 nm diameter were used to achieve higher resolu-
tion [7, 16].
In order to reduce noise and resolution anisotro-
py in different planes of the obtained tomograms, fil-
tering in the IsoNet program was used [17]. This tool
uses a U-net neural network and is trained on small
sections of tomograms with addition of extra noise.
Segmentation of mitochondrial membranes was per-
formed in an automatic mode using the program
TomoSegMemTV [18]. Errors in the automatic segmen-
tation were corrected manually using the original to-
mogram data.
The tomograms after filtering were used in man-
ual mode to determine positions of different molecu-
lar complexes. Positions and orientations of various
macromolecules were also determined automatically
using three-dimensional templates of these molecules
in the Dynamo [19] or WARP [20] programs. Subsequent
iterative 3D classification was performed to eliminate
incorrectly selected coordinates. To increase the sig-
nal-to-noise ratio and spatial resolution [21] of mac-
romolecules, averaging of small sections of the tomo-
gram (called subtomograms) containing individual
macromolecules aligned with each other was used [22].
This process, called subtomographic averaging, simul-
taneously provides information about orientation of
macromolecules, which is used for subsequent visual-
ization of the object by inserting a model of the aver-
aged structure in the desired orientation.
RESULTS
Visualization of amyloid aggregates in cells.
Based on fluorescence microscopy data, it has previous-
ly been suggested that the beta-amyloid (Aβ42) aggre-
gates can interact with mitochondrial membranes and
thereby disrupt bioenergetics [9]. For the experiment,
we have chosen a previously used model based on the
aerobically metabolizing yeast Y. lipolytica expressing
the eGFP-Aβ42 construct [9]. It was shown earlier that
expression of Aβ42 or of eGFP-Aβ42 reporter gene con-
struct led to dysfunction and fragmentation of yeast
mitochondria [9]. Colocalization of amyloid aggregates
and mitochondria revealed in this work [9] led to the
hypothesis of direct physical contact of amyloid ag-
gregates with the mitochondrial membrane. To test
this hypothesis, a CLEM microscopy protocol was
performed, culminating in cryo-electron tomography
(cryo-ET) of the selected regions of interest. It is im-
portant to note that cryo-ET is the only suitable meth-
od of investigation for this task, because during the
vitrification process and subsequent microscopic ex-
amination, native structure of the samples is practi-
cally not disturbed, which is an important parameter
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because fluorescent objects are located at a small dis-
tance and even a slight change in their colocalization
due to the use of standard technique of fixing and con-
trasting the sample for electron microscopy could lead
to the appearance of artifacts.
The cryogenic CLEM protocol involved using flu-
orescence microscopy to examine the vitrified sam-
ple for the most representative fields of view where
MitoTracker Red-stained mitochondria and fluorescent
eGFP-Aβ42 aggregates were expected to colocalize.
Once a region of interest was identified, z-stack imag-
ing was performed to reconstruct 3D location of the
areas containing mitochondria and eGFP-Aβ42 aggre-
gates. Based on this information, FIB was adjusted to
obtain a lamella containing the region of interest for
which cryo-electron tomograms were obtained. Fig-
ure1 shows the intermediate results of each stage of
the study.
As a result, a series of detailed tomograms of the
regions of interest were obtained, superimposing both
mitochondrial fluorescence (red channel, MitoTracker
Red) and amyloid aggregates (green channel, eGFP-
Aβ42). At the same time, resolution of the cryo-elec-
tron tomograms was significantly higher than that
of the fluorescence microscope, allowing us to distin-
guish both membranes and individual proteins in the
mitochondria that have characteristic structural fea-
tures, such as ATP synthases (Fig. 1d). The eGFP fluo-
rescence signal allowed to localize amyloid aggregates,
which could not be clearly identified without fluores-
cent labeling and using CLEM. The aggregates were
shown not to interact with the mitochondrial mem-
brane. As can be seen in Fig. 1, some of the aggregates
do not localize near the mitochondria at all, which can
also be seen in the fluorescence microscopy images.
As for the ambiguous areas where the fluorescence
signals overlap, statistical data were collected for sev-
eral dozen of aggregates in these areas (Fig.2) and it
was shown that the vast majority of aggregates were
Fig. 1. Workflow on a cell sample during CLEM microscopy
combined with cryo-ET. a)Photograph of a grid with a vitri-
fied sample under a transmission light microscope. b)Magni-
fied image of the area highlighted in panel(a) with sputtered
platinum layer and an area cut out by cryogenic focused ion
beam method in which there is a thin lamella (highlighted
with a red frame). Scanning electron ion microscopy image.
c) Correlative light and electron microscopy (CLEM) of the
lamella. Overlay of the fluorescence image region correspond-
ing to the etched lamella. Red fluorescence is MitoTracker Red
(mitochondrial marker), green fluorescence is eGFP (from
the eGFP-Aβ42 construct). d) Slice of the tomogram plotted
against the region inside the lamella showing two mitochon-
dria as well as other cell components (cell wall, ribosomes,
amyloid aggregates and various vesicular structures). Inset
shows an enlarged fragment of a mitochondrial crista, with
red arrows pointing at F1 subunits of ATP synthase protrud-
ing from the membrane.
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BIOCHEMISTRY (Moscow) Vol. 89 No. 2 2024
Fig. 2. Visualization and estimation of distances between Aβ42 aggregates and mitochondrial outer membrane on multiple to-
mograms. a)Example of computer reconstruction of a tomogram with automatically labeled mitochondrial membranes (blue),
amyloid aggregates (yellow) and ribosomes (red). b)Distribution of distances between the aggregates and the mitochondrial
outer membrane assembled from the tomographic data.
Fig. 3. Structures of the respiratory chain supercomplex of mitochondria from different organisms. a,c)Different projections of
the respirasome surface structure from Arabidopsis sp. mitochondria obtained by single-particle cryo-EM (resolution about2 Å,
PDBID: 8BPX) [34]. b,d)Different projections of the respirasome structure from rat heart mitochondria obtained by subtomo-
graphic averaging without extraction of the respirasome from the membrane according to [7]. In panels(a), (b), (c) different
complexes within the supercomplex [blue– complexI, purple– complexIII, green– complexIV (only present in rat respira-
somes)] are highlighted by different colors. In panel(c) color mapping is done according to the surface hydrophilicity (blue–
hydrophilic, yellow– hydrophobic) to better illustrate presence of the membrane curvature.
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BIOCHEMISTRY (Moscow) Vol. 89 No. 2 2024
located at a distance of 30-300 nm from the mitochon-
drial membrane surface.
The closest aggregates had no physical contacts
with the mitochondrial membrane. This finding is
important for understanding the mechanisms of Alz-
heimer’s disease development, as it suggests that the
disturbance in mitochondrial bioenergetics that cor-
relates with aggregate accumulation [23, 24] is not
caused by the direct effect of amyloid aggregates on
mitochondrial structure and function, but is mediated
by other factors.
Study of the topology of the inner mitochondri-
al membrane and proteins of the OXPHOS system
within it. The single-particle cryo-EM method, based
on computer classification and averaging of huge ar-
rays of images obtained by transmission electron mi-
croscopy(TEM), allows us to study structures of large
protein complexes and supercomplexes. Presence or
absence of supercomplexes has long been a contro-
versial topic in bioenergetics, yet this fact is extremely
important. Presence of supercomplexes implies direct
transfer of metabolites between them with minimal
involvement of diffusion in this process. Such arrange-
ment is much more resistant to stress, reduces electron
leakage and ROS generation, making the whole system
more energy efficient. Existence of large respiratory
chain supercomplexes in mitochondria prior to the
use of cryo-EM was demonstrated by the presence of
common migration of their components during native
electrophoresis after protein extraction from mem-
branes with mild detergents [25]. Nowadays, existence
of the respiratory supercomplexes– respirasomes – is
no longer in doubt due to cryo-EM imaging of single
particles, as their structures from different organisms
and tissues have been demonstrated in a large number
of studies [26-33]. Nevertheless, more detailed issues,
such as exact configuration and composition of super-
complexes under certain conditions, their functional,
regulatory and structural significance for bioenerget-
ics, remain controversial. In particular, mechanisms of
regulation by respirasomes, existence of any structur-
al interaction between them and ATP synthases, and
role of the membrane in these processes are not fully
understood. To address these issues, we analyzed the
data obtained in our previous studies together with
the new literature data. Figure 3 shows structures of
the respirasomes from the evolutionarily distant Ara-
bidopsissp. mitochondria and rat heart mitochondria,
which surprisingly turned out to be not only similar in
structure, but also to have a similar curvature of the
membrane, creating its kink at the point of contact of
complexesI andIII.
Cryo-ET (cryo-electron tomography) allows us to
reconstruct entire membrane regions by determining
location of the proteins with characteristic structur-
al features. Such proteins include ATP synthases and
complexesI, which have unique shape of a large hydro-
philic part protruding from the membrane (Fig. 4, a, b).
In addition, it is possible to reconstruct three-dimen-
sional structure of the inner mitochondrial membrane
with resolution that significantly exceeds that of to-
mography, based on analysis of the slices of fixed mi-
tochondrial samples. As a result, cryo-ET shows the
inner membranes of cardiac mitochondria in greater
detail and are not represented as wave-like lamellar
structures, as interpreted from the TEM ultrathin sec-
tions [35], but as a complex structure of many small
flat fragments interspersed with many tubular junc-
tions (Fig. 4). This structure answers the question of
how ATP synthase dimers are arranged in the inner
membrane of cardiac mitochondria. The most abun-
dant oligomers of ATP synthases are found at the
bends of the highly curved cristae [36], but there are
fewer long clusters of ATP synthases in the regions
with lower curvature, where a less ordered structure
is created in which dense packing of respirasomes
and ATP synthases is impossible (Fig.4b, right row of
ATP synthases).
On a smaller scale (without visualization of indi-
vidual membrane proteins), it is possible to perform a
detailed reconstruction of the inner membrane of car-
diac mitochondria using high resolution images gener-
ated with cryo-ET. We have recently performed such
reconstruction to visualize position of the large dehy-
drogenase complexes relative to the membrane under
natural conditions [11]. The obtained data suggest
(Fig. 4) that the cardiac mitochondria are a complex
maze of highly curved membranes with a lamellar-
tubular structure. At the same time, ATP synthase di-
mers cannot be located in the completely lamellar flat
areas [36]. From the point of view of ensuring optimal
structure and functionality of mitochondria, the lamel-
lar structure of the membrane seems to be unsuitable
for accommodating the key components of OXPHOS.
This issue can be partially resolved by formation of the
wave-like curved structures of the inner membrane bi-
layer, on the curved areas of which ATP synthases can
be localized. Such topology has been indeed observed
in some TEM images [37]. However, the membrane
structures are often visualized as lamellar, which pre-
vents formation even of the strained structures with
ATP synthase dimers, one of which is shown in Fig.4b.
The paradox of the presence of lamellae visible by
classical electron microscopy was resolved by using
higher resolution of cryo-ET. The computer recon-
struction showed that toroidal holes are formed in the
lamellar regions at the edges of which ATP synthase di-
mers could be located, and close to them in the regions
with less curvature – respirasomes. In contrast to the
cristae edges shown in Fig.3, the toroidal openings at
the edges could accommodate not only one row of ATP
synthase dimers, but even two rows of the tightly at-
MITOCHONDRIAL PROTEINS AND MEMBRANES IN NATURAL ENVIRONMENT 263
BIOCHEMISTRY (Moscow) Vol. 89 No. 2 2024
Fig. 4. Reconstruction of the complex structure of the mitochondrial membranes of the heart. a)Reconstruction of a tomogram
of a large area in a whole intact mitochondrion of the rat heart. Blue– cores of ketoglutarate dehydrogenase or branched-chain
ketoacid dehydrogenase complexes, green– cores of pyruvate dehydrogenase complexes. b,d)Fragments illustrating presence
of holes of toroidal topology. c)Reconstruction of a fragment of a tomogram of a rat heart mitochondrial crista. ATP synthases
are shown in yellow, complexesI in blue, dimers of complexesIII in purple and complexesIV in green. The membrane shown
in grey is not transparent and covers hydrophobic parts of the complexes. e)Thin section of part of the crista at the location
indicated by the arrows in panel(c). The membrane is shown transparent, allowing visualization of the hydrophobic parts of
thecomplexes. The image created based on the tomographic data obtained in the course of previous work [7, 11].
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BIOCHEMISTRY (Moscow) Vol. 89 No. 2 2024
Fig. 5. Schematic model of putative location of protein complexes of the OXPHOS system in the vicinity of toroidal holes in the
pseudolamellar regions of cristae. The model was constructed based on manual processing of cryo-ET images of several toroidal
holes with labelling of the location of F1 subunits of ATP synthases in the inner membrane of rat heart mitochondria. Yellow
indicates ATP synthase dimers, blue indicates respirasomes, dark grey indicates membrane and light grey indicates intermem-
brane space.
tached ATP synthases. This ensures the most efficient
use of the rather small (compared to the total area of
the cristae) curved regions of the membrane for locat-
ing ATP synthase dimers on them. An oligomeric struc-
ture of the ATP synthase dimers similar to that shown
in Fig.4 has been observed in the toroidal holes in the
region with high curvature. However, the preliminary
manual analysis shows a larger number of structures
in some of the toroidal holes that correspond in size to
the F1 subunits of ATP synthases, suggesting that the
two rows of ATP synthase dimers may be present at
the membrane fold at the same time. Due to the high
protein density, which prevents high resolution, accu-
rate computer reconstruction using averaging of ATP
synthases in these regions has not yet been possible.
Schematic representation of the proposed arrange-
ment of the OXPHOS complexes in such structures is
shown in Fig.5.
This complex topology of mitochondrial mem-
branes may be maintained by high concentration of
the cone-shaped lipids such as cardiolipin and phos-
phatidylethanolamine in the membrane. Presumably,
the membrane junctions are so enriched in these lip-
ids, and their structure is so different from the bilayer,
that they have been defined by NMR as a non-bilayer
lipid phase [37, 38]. Given the necessity of cardiolipin
for operation of the OXPHOS system [39] and optimiza-
tion of its structure to ensure proton transport across
the membrane [8], we can assume that the propensity
of respirasomes and ATP synthases to generate mem-
brane curvature may, at least in part, be evolutionarily
determined by the need for cardiolipin concentration
at the interface of OXPHOS system proteins.
DISCUSSION
Lack of direct interaction of amyloid aggre-
gates with mitochondrial membranes. Advanced
models of chronic diseases associated with mitochon-
drial dysfunction, based on the aerobic metabolic yeast
Y. lipolytica [9], have proven to be a powerful tool to
reveal intracellular actions of the markers of these dis-
eases, which is an important task in the case of pathol-
ogies associated with mitochondrial dysfunction [40],
since mitochondria are quite deeply involved in the
cell signaling systems, and it is often difficult to under-
stand whether their dysfunction is primary or medi-
ated by pathological changes in the cellular environ-
ment. The yeast Y. lipolytica, as a unicellular organism,
lacks complex system of intercellular interactions and,
in addition, possesses a branched system of fully func-
tional “animal-type” mitochondria containing all the
complexes of the respiratory chain.
It was shown [9] that the Aβ42 expression in the
yeast caused mitochondrial dysfunction, which is char-
acteristic for the neurons at the early stages of Alzhei-
mer’s disease development. In addition, colocalization
of mitochondria and amyloid aggregates suggested the
presence of physical contact between the mitochondria
and amyloid aggregates, as assumed in some other work
[41]. Since vitrification of the samples for cryogenic mi-
croscopy is the least destructive method of object fixa-
tion, the data obtained in this work could be considered
as the most reliable showing absence of the contact be-
tween the amyloid protein aggregates and mitochondria.
Nevertheless, distribution of the distances between
the aggregates and mitochondrial membrane indicates
MITOCHONDRIAL PROTEINS AND MEMBRANES IN NATURAL ENVIRONMENT 265
BIOCHEMISTRY (Moscow) Vol. 89 No. 2 2024
concentration of aggregates near mitochondria, which
may be essential for the processes of mitochondrial
substrate transport or for the effects of aggregates on
the protein biosynthesis due to the large number of ri-
bosomes also located near mitochondria.
Model of topological coupling of supercomplex-
es in the mitochondrial membrane. An interesting
peculiarity that was identified owing to cryo-ET was
the fact that the ATP synthases not only dimerize (this
can be detected by native electrophoresis), but also form
oligomeric arrays that deform the membrane and in-
duce a significant curvature on it [16, 42, 43]. It turns
out that respirasomes also affect membrane curvature,
as shown in the cryo-EM images of respirasomes from
different organisms [30, 34]. Furthermore, interaction
of the complex I and complex III dimers in respira-
some is conserved, and structure of the respirasome is
highly similar between the species [44] suggesting that
interaction of the respirasome with the membrane is
also conserved. This implies that association of respi-
rasomes with ATP synthases shown in [7] (Fig. 4c)
could be achieved largely through the mechanical prop-
erties of the membrane, which tend to compensate for
excessive tension. The ATP synthase dimers create
a region of maximum membrane curvature close to
them [36], thereby creating tension in the bilayer and
distorting it from the energetically optimal structure
that would be present if the lipids self-assembled in
the absence of the protein. In the region away from
ATP synthases, the membrane tends to approach the
equilibrium topology of a lamellar bilayer. At the same
time, respirasomes also generate membrane tension,
but the lipid bilayer around them has much less cur-
vature than near the ATP synthase dimers. In order
for respirasomes not to create a separate region of
membrane curvature, it is advantageous for them to
be located closer to ATP synthases in the part of the
membrane where the curvature is already optimal for
them and no additional energy is required to mechani-
cally distort the lipid bilayer. In such case, preferential
orientation of respirasomes along one axis should be
observed, which is indeed confirmed by the experi-
ments with cardiac mitochondria [7]. Thus, the force
attracting respirasomes to ATP synthase has an entro-
pic topological nature and is determined by the mem-
brane, which tends to minimize its deviation from the
approximate equilibrium bilayer structure. It should
also be noted that the integral protein complexes that
create curvature of the mitochondrial membrane also
create around themselves membrane domains (rafts)
of cardiolipin and other lipids that have conical shape,
since they are the ones best suited to create curvature
in the lipid bilayer [45]. Binding of ATP synthases and
respiratory complexes (both individually and assem-
bled into respirasomes) to cardiolipin has been repeat-
edly confirmed in experiments [38, 46-53]. Ability of
cardiolipin to form membrane domains has also been
experimentally demonstrated [54].
Sorting of the proteins based on their interaction
with membrane lipids is well known – such sorting
system based on rafts and nanorafts operates in the
cell [55]. Sorting of the proteins in membrane rafts
is mainly based on coincidence or mismatch of the
thickness of their hydrophobic membrane zone with
the thickness of a certain cluster of membrane lipids
(hydrophobic mismatch principle). The above data on
mitochondrial membranes allow us to add to the well-
known principle of clustering of proteins with similar
hydrophobic zone another sorting principle based on
affinity of the protein complexes to the zones of differ-
ent membrane curvature. Complex topology of the in-
ner membrane of cardiac mitochondria and presence
of the non-bilayer phases in it suggest that self-orga-
nization of the OXPHOS system complexes in the car-
diac mitochondria could proceed via the mechanism
similar to the in meso crystallization of membrane
proteins[56].
Thus, the above data suggest that the structure
of respirasomes is evolutionarily selected to ensure
localization of respirasomes close to ATP synthase di-
mers and thus clustering of the entire OXPHOS system
through interaction with the membrane lipids.
CONCLUSION
This work demonstrates experimental application
of one of the most advanced methods in structural bi-
ology to study interaction of protein aggregates with
membranes, topology of the membranes themselves,
and mutual arrangement of membrane proteins in mi-
tochondria under native conditions. A series of new
experiments using CLEM have shown that amyloid
aggregates do not interact with the outer mitochon-
drial membrane and do not enter the mitochondria.
In addition, our own recent data and recent literature
data obtained using cryogenic-electron transmission
microscopy and tomography of mitochondria and mi-
tochondrial membranes have been analyzed in more
detail. Complex arrangement and intricate topology of
the rat heart mitochondrial membranes, enriched in
large supercomplexes that influence membrane cur-
vature, are shown. In complete agreement with this, it
was shown that the cardiac mitochondrial membrane
has only small regions where it is close to a flat bilayer,
whereas most of the membrane is a complex network
of highly curved junctions on which rows of ATP syn-
thase dimers and respiratory chain supercomplexes
are located. The regions of the membrane that appear
lamellar on the TEM thin sections are actually dotted
with a multitude of holes containing tightly packed
clusters of protein complexes of the OXPHOS system.
NESTEROV et al.266
BIOCHEMISTRY (Moscow) Vol. 89 No. 2 2024
This paper also discusses theoretical framework that
explains clustering of the membrane proteins of the
OXPHOS system and cardiolipin by minimization of
the strain energy of the lipid part of the membrane.
This principle of membrane protein organisation is
proposed to be termed topological.
Contributions. S.V.N. planned and performed ex-
periments, manually analyzed tomograms, prepared
the manuscript; D.A.M., T.N.G., and A.G.R. planned and
conducted CLEM experiments with yeast; K.S.P. and
Yu.M.Ch. conduct TEM and CLEM experiments, perform
computer processing of the tomographic data. L.S.Y.
and R.G.V. developed the study concept, supervised the
study. All authors discussed the results and contribute
to the manuscript editing.
Funding. This work was financially supported
by the NRC Kurchatov Institute (thematic plan 1f.4.1
“Study of energy generation, transfer and distribution
processes in living organisms aimed at finding new
approaches to the development of therapeutic agents,
new bioenergetic devices and artificial photosynthesis
systems”).
Ethics declarations. This work does not contain
any studies involving human and animal subjects.
The authors of this work declare that they have no
conflicts of interest.
REFERENCES
1. Saibil, H. R. (2022) Cryo-EM in molecular and cel-
lular biology, Mol. Cell, 82, 274-284, doi: 10.1016/
j.molcel.2021.12.016.
2. Guaita,M., Watters, S.C., and Loerch,S. (2022) Recent
advances and current trends in cryo-electron micros-
copy, Curr. Opin. Struct. Biol., 77, 102484, doi:10.1016/
j.sbi.2022.102484.
3. Chua, E.Y.D., Mendez, J.H., Rapp,M., Ilca, S.L., Tan,
Y.Z., Maruthi,K., Kuang,H., Zimanyi, C.M., Cheng,A.,
Eng, E.T., Noble, A.J., Potter, C.S., and Carragher, B.
(2022) Better, faster, cheaper: recent advances in
cryo-electron microscopy, Annu. Rev. Biochem., 91,
1-32, doi:10.1146/annurev-biochem-032620-110705.
4. Hoffman, D.P., Shtengel,G., Xu, C.S., Campbell, K.R.,
Freeman, M., Wang, L., Milkie, D. E., Pasolli, H. A.,
Iyer, N., Bogovic, J. A., Stabley, D. R., Shirinifard, A.,
Pang,S., Peale,D., Schaefer,K., Pomp,W., Chang, C.-L.,
Lippincott-Schwartz,J., Kirchhausen,T., Solecki, D.J.,
Betzig, E., and Hess, H. F. (2020) Correlative three-
dimensional super-resolution and block-face electron
microscopy of whole vitreously frozen cells, Science,
367, eaaz5357, doi:10.1126/science.aaz5357.
5. Liu, T., Stephan, T., Chen, P., Keller-Findeisen, J.,
Chen, J., Riedel, D., Yang, Z., Jakobs, S., and Chen, Z.
(2022) Multi-color live-cell STED nanoscopy of mito-
chondria with a gentle inner membrane stain, Proc.
Natl. Acad. Sci. USA, 119, e2215799119, doi: 10.1073/
pnas.2215799119.
6. Wang,C., Taki,M., Sato,Y., Tamura,Y., Yaginuma,H.,
Okada, Y., and Yamaguchi, S. (2019) A photostable
fluorescent marker for the superresolution live im-
aging of the dynamic structure of the mitochondrial
cristae, Proc. Natl. Acad. Sci. USA, 116, 15817-15822,
doi:10.1073/pnas.1905924116.
7. Nesterov,S., Chesnokov, Y., Kamyshinsky,R., Pantele-
eva,A., Lyamzaev,K., Vasilov,R., and Yaguzhinsky,L.
(2021) Ordered clusters of the complete oxidative
phosphorylation system in cardiac mitochondria,
Int.J. Mol. Sci., 22, 1462, doi:10.3390/ijms22031462.
8. Nesterov, S. V., Yaguzhinsky, L. S., Vasilov, R. G.,
Kadantsev, V.N., and Goltsov, A.N. (2022) Contribution
of the collective excitations to the coupled proton and
energy transport along mitochondrial cristae mem-
brane in oxidative phosphorylation system, Entropy,
24, 1813, doi:10.3390/e24121813.
9. Epremyan, K.K., Rogov, A.G., Goleva, T.N., Lavrushki-
na, S.V., Zinovkin, R.A., and Zvyagilskaya, R.A. (2023)
Altered mitochondrial morphology and bioenergetics
in a new yeast model expressing Aβ42, Int.J. Mol. Sci.,
24, 900, doi:10.3390/ijms24020900.
10. Bischof, J., Hunt, C. J., Rubinsky, B., Burgess, A., and
Pegg, D. E. (1990) Effects of cooling rate and glyc-
erol concentration on the structure of the frozen
kidney: Assessment by cryo-scanning electron mi-
croscopy, Cryobiology, 27, 301-310, doi:10.1016/0011-
2240(90)90029-4.
11. Plokhikh, K.S., Nesterov, S.V., Chesnokov, Y.M., Rogov,
A.G., Kamyshinsky, R.A., Vasiliev, A.L., Yaguzhinsky,
L.S., and Vasilov, R.G. (2023) Association of 2-oxoac-
id dehydrogenase complexes with respirasomes
in mitochondria, FEBS J., 291, 16965, doi: 10.1111/
febs.16965.
12. Nesterov, S.V., Skorobogatova, Y.A., Panteleeva, A.A.,
Pavlik, L. L., Mikheeva, I. B., Yaguzhinsky, L. S., and
Nartsissov, Y. R. (2018) NMDA and GABA receptor
presence in rat heart mitochondria, Chem. Biol. Inter-
act., 291, 40-46, doi:10.1016/j.cbi.2018.06.004.
13. Sibarita, J.-B. (2005) Deconvolution microscopy, in Mi-
croscopy Techniques (Rietdorf,J., ed) Springer, Berlin,
Heidelberg, pp.201-243, doi:10.1007/b102215.
14. Kremer, J.R., Mastronarde, D.N., and McIntosh, J.R.
(1996) Computer visualization of three-dimension-
al image data using IMOD, J. Struct. Biol., 116, 71-76,
doi:10.1006/jsbi.1996.0013.
15. Wan, W., and Briggs, J. A. G. (2016) Cryo-Electron To-
mography and Subtomogram Averaging, 1st Edn.,
Elsevier, doi:10.1016/bs.mie.2016.04.014.
16. Nesterov, S.V., Chesnokov, Yu.M., Kamyshinsky, R.A.,
Yaguzhinsky, L.S., and Vasilov, R.G. (2020) Determin-
ing the structure and location of the ATP synthase
in the membranes of rat’s heart mitochondria using
MITOCHONDRIAL PROTEINS AND MEMBRANES IN NATURAL ENVIRONMENT 267
BIOCHEMISTRY (Moscow) Vol. 89 No. 2 2024
cryoelectron tomography, Nanotechnol. Russia, 15,
83-89, doi:10.1134/S1995078020010139.
17. Liu, Y.-T., Zhang,H., Wang,H., Tao, C.-L., Bi, G.-Q., and
Zhou, Z. H. (2022) Isotropic reconstruction for elec-
tron tomography with deep learning, Nat. Commun.,
13, 6482, doi:10.1038/s41467-022-33957-8.
18. Martinez-Sanchez, A., Garcia, I., Asano, S., Lucic, V.,
and Fernandez, J. J. (2014) Robust membrane de-
tection based on tensor voting for electron tomog-
raphy, J. Struct. Biol., 186, 49-61, doi: 10.1016/j.jsb.
2014.02.015.
19. Castaño-Díez, D., Kudryashev, M., Arheit, M., and
Stahlberg, H. (2012) Dynamo: A flexible, user-friend-
ly development tool for subtomogram averaging of
cryo-EM data in high-performance computing envi-
ronments, J. Struct. Biol., 178, 139-151, doi: 10.1016/
j.jsb.2011.12.017.
20. Tegunov,D., and Cramer,P. (2019) Real-time cryo-elec-
tron microscopy data preprocessing with Warp,
Nat. Methods, 16, 1146-1152, doi: 10.1038/s41592-
019-0580-y.
21. Bharat, T. A. M., and Scheres, S. H. W. (2016) Resolv-
ing macromolecular structures from electron cryo-
Tomography data using subtomogram averaging in
RELION, Nat. Protoc., 11, 2054-2065, doi: 10.1038/
nprot.2016.124.
22. Asano,S., Fukuda,Y., Beck,F., Aufderheide,A., Först-
er,F., Danev,R., and Baumeister,W. (2015) A molecular
census of 26S proteasomes in intact neurons, Science,
347, 439-442, doi:10.1126/science.1261197.
23. Ashleigh, T., Swerdlow, R. H., and Beal, M. F. (2023)
Therole of mitochondrial dysfunction in Alzheimer’s
disease pathogenesis, Alzheimers Dement. J. Alzhei-
mers Assoc., 19, 333-342, doi:10.1002/alz.12683.
24. Bhatia,S., Rawal,R., Sharma, P., Singh, T., Singh, M.,
and Singh,V. (2022) Mitochondrial dysfunction in Alz-
heimer’s disease: opportunities for drug development,
Curr. Neuropharmacol., 20, 675-692, doi: 10.2174/
1570159X19666210517114016.
25. Eubel,H., Heinemeyer,J., and Braun, H.-P. (2004) Iden-
tification and characterization of respirasomes in
potato mitochondria, Plant Physiol., 134, 1450-1459,
doi:10.1104/pp.103.038018.
26. Chaban,Y., Boekema, E.J., and Dudkina, N.V. (2014)
Structures of mitochondrial oxidative phosphory-
lation supercomplexes and mechanisms for their
stabilization, Biochim. Biophys. Acta, 1837, 418-426,
doi:10.1016/j.bbabio.2013.10.004.
27. Dudkina, N.V., Kouřil,R., Peters,K., Braun, H.-P., and
Boekema, E.J. (2010) Structure and function of mito-
chondrial supercomplexes, Biochim. Biophys. Acta,
1797, 664-670, doi:10.1016/j.bbabio.2009.12.013.
28. Bultema, J. B., Braun, H.-P., Boekema, E. J., and Kou-
ril, R. (2009) Megacomplex organization of the oxi-
dative phosphorylation system by structural anal-
ysis of respiratory supercomplexes from potato,
Biochim. Biophys. Acta, 1787, 60-67, doi: 10.1016/
j.bbabio.2008.10.010.
29. Dudkina, N. V., Kudryashev, M., Stahlberg, H., and
Boekema, E. J. (2011) Interaction of complexes I,
III, and IV within the bovine respirasome by sin-
gle particle cryoelectron tomography, Proc. Natl.
Acad. Sci. USA, 108, 15196-15200, doi: 10.1073/pnas.
1107819108.
30. Mühleip, A., Flygaard, R. K., Baradaran, R., Haapa-
nen, O., Gruhl, T., Tobiasson, V., Maréchal, A., Shar-
ma,V., and Amunts, A. (2023) Structural basis of mi-
tochondrial membrane bending by the I-II-III2-IV2 su-
percomplex, Nature, 615, 934-938, doi:10.1038/s41586-
023-05817-y.
31. Guo,R., Zong,S., Wu,M., Gu,J., and Yang, M. (2017)
Architecture of human mitochondrial respirato-
ry megacomplex I2III2IV2, Cell, 170, 1247-1257.e12,
doi:10.1016/j.cell.2017.07.050.
32. Gu, J., Wu, M., Guo, R., Yan, K., Lei, J., Gao, N., and
Yang, M. (2016) The architecture of the mammali-
an respirasome, Nature, 537, 639-643, doi: 10.1038/
nature19359.
33. Vercellino,I., and Sazanov, L.A. (2021) Structure and
assembly of the mammalian mitochondrial super-
complex CIII2CIV, Nature, 598, 364-367, doi: 10.1038/
s41586-021-03927-z.
34. Klusch,N., Dreimann,M., Senkler,J., Rugen,N., Kühl-
brandt, W., and Braun, H.-P. (2023) Cryo-EM struc-
ture of the respiratory I + III2 supercomplex from
Arabidopsis thaliana at 2 Å resolution, Nat Plants., 9,
142-156, doi:10.1038/s41477-022-01308-6.
35. Kühlbrandt, W. (2015) Structure and function of mi-
tochondrial membrane protein complexes, BMC Biol.,
13, 89, doi:10.1186/s12915-015-0201-x.
36. Strauss, M., Hofhaus, G., Schröder, R. R., and Kühl-
brandt, W. (2008) Dimer ribbons of ATP synthase
shape the inner mitochondrial membrane, EMBO J.,
27, 1154-1160, doi:10.1038/emboj.2008.35.
37. Garab, G., Yaguzhinsky, L. S., Dlouhý, O., Nesterov,
S. V., Špunda, V., and Gasanoff, E. S. (2022) Structur-
al and functional roles of non-bilayer lipid phases
of chloroplast thylakoid membranes and mitochon-
drial inner membranes, Prog. Lipid. Res., 86, 101163,
doi:10.1016/j.plipres.2022.101163.
38. Gasanov, S. E., Kim, A. A., Yaguzhinsky, L. S., and
Dagda, R. K. (2018) Non-bilayer structures in mito-
chondrial membranes regulate ATP synthase activity,
Biochim. Biophys. Acta, 1860, 586-599, doi: 10.1016/
j.bbamem.2017.11.014.
39. Paradies, G., Paradies, V., De Benedictis, V., Ruggiero,
F.M., and Petrosillo,G. (2014) Functional role of cardio-
lipin in mitochondrial bioenergetics, Biochim. Biophys.
Acta, 1837, 408-417, doi:10.1016/j.bbabio.2013.10.006.
40. Epremyan, K.K., Goleva, T.N., Rogov, A.G., Lavrushki-
na, S.V., Zinovkin, R.A., and Zvyagilskaya, R.A. (2022)
The first Yarrowia lipolytica yeast models expressing
NESTEROV et al.268
BIOCHEMISTRY (Moscow) Vol. 89 No. 2 2024
hepatitisB virusX protein: changes in mitochondrial
morphology and functions, Microorganism, 10, 1817,
doi:10.3390/microorganisms10091817.
41. Völgyi,K., Badics,K., Sialana, F.J., Gulyássy,P., Udvari,
E. B., Kis, V., Drahos,L., Lubec,G., Kékesi, K.A., and
Juhász, G. (2018) Early presymptomatic changes in
the proteome of mitochondria-associated membrane
in the APP/PS1 mouse model of Alzheimer’s disease,
Mol. Neurobiol., 55, 7839-7857, doi: 10.1007/s12035-
018-0955-6.
42. Buzhynskyy, N., Sens, P., Prima, V., Sturgis, J. N., and
Scheuring,S. (2007) Rows of ATP synthase dimers in
native mitochondrial inner membranes, Biophys. J.,
93, 2870-2876, doi:10.1529/biophysj.107.109728.
43. Blum, T.B., Hahn,A., Meier,T., Davies, K.M., and Kühl-
brandt, W. (2019) Dimers of mitochondrial ATP syn-
thase induce membrane curvature and self-assemble
into rows, Proc. Natl. Acad. Sci. USA, 116, 4250-4255,
doi:10.1073/pnas.1816556116.
44. Davies, K.M., Blum, T.B., and Kühlbrandt,W. (2018)
Conserved in situ arrangement of complex I and III2
in mitochondrial respiratory chain supercomplexes of
mammals, yeast, and plants, Proc. Natl. Acad. Sci. USA,
115, 3024-3029, doi:10.1073/pnas.1720702115.
45. Beltrán-Heredia, E., Tsai, F.-C., Salinas-Almaguer, S.,
Cao, F. J., Bassereau, P., and Monroy, F. (2019) Mem-
brane curvature induces cardiolipin sorting, Commun.
Biol., 2, 225, doi:10.1038/s42003-019-0471-x.
46. Arias-Cartin, R., Grimaldi, S., Arnoux, P., Guigliarel-
li, B., and Magalon, A. (2012) Cardiolipin binding in
bacterial respiratory complexes: structural and func-
tional implications, Biochim. Biophys. Acta, 1817,
1937-1949, doi:10.1016/j.bbabio.2012.04.005.
47. Arnarez,C., Marrink, S.J., and Periole,X. (2013) Iden-
tification of cardiolipin binding sites on cytochromec
oxidase at the entrance of proton channels, Sci. Rep.,
3, 1263, doi:10.1038/srep01263.
48. Duncan, A.L., Robinson, A.J., and Walker, J.E. (2016)
Cardiolipin binds selectively but transiently to con-
served lysine residues in the rotor of metazoan ATP
synthases, Proc. Natl. Acad. Sci. USA, 113, 8687-8692,
doi:10.1073/pnas.1608396113.
49. Pfeiffer,K., Gohil,V., Stuart, R.A., Hunte,C., Brandt,U.,
Greenberg, M.L., and Schägger,H. (2003) Cardiolipin
stabilizes respiratory chain supercomplexes, J. Biol.
Chem., 278, 52873-52880, doi:10.1074/jbc.M308366200.
50. Mühleip, A., McComas, S. E., and Amunts, A. (2019)
Structure of a mitochondrial ATP synthase with bound
native cardiolipin, eLife, 8, e51179, doi: 10.7554/
eLife.51179.
51. Mileykovskaya,E., and Dowhan,W. (2014) Cardiolip-
in-dependent formation of mitochondrial respirato-
ry supercomplexes, Chem. Phys. Lipids, 179, 42-48,
doi:10.1016/j.chemphyslip.2013.10.012.
52. Zhang,M., Mileykovskaya,E., and Dowhan,W. (2005)
Cardiolipin is essential for organization of complex-
esIII and IV into a supercomplex in intact yeast mito-
chondria, J.Biol. Chem., 280, 29403-29408, doi:10.1074/
jbc.M504955200.
53. Zhang,M., Mileykovskaya,E., and Dowhan,W. (2002)
Gluing the Respiratory Chain Together cardiolipin
is required for supercomplex formation in the inner
mitochondrial membrane, J. Biol. Chem., 277, 43553-
43556, doi:10.1074/jbc.C200551200.
54. Mileykovskaya,E., and Dowhan,W. (2009) Cardiolipin
membrane domains in prokaryotes and eukaryotes,
Biochim. Biophys. Acta, 1788, 2084-2091, doi:10.1016/
j.bbamem.2009.04.003.
55. Diaz-Rohrer,B., Levental, K.R., and Levental,I. (2014)
Rafting through traffic: Membrane domains in cellu-
lar logistics, Biochim. Biophys. Acta, 1838, 3003-3013,
doi:10.1016/j.bbamem.2014.07.029.
56. Zabara,A., Meikle, T.G., Newman,J., Peat, T.S., Conn,
C.E., and Drummond, C.J. (2017) The nanoscience be-
hind the art of inmeso crystallization of membrane pro-
teins, Nanoscale, 9, 754-763, doi:10.1039/C6NR07634C.
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