Download Role of cryo-ET in membrane bioenergetics research

Bioenergetics in Mitochondria, Bacteria and Chloroplasts
Role of cryo-ET in membrane bioenergetics
research
Karen M. Davies*1 and Bertram Daum*
*Department of Structural Biology, Max Planck Institute of Biophysics, Max-von-Laue Strasse 3, 60438 Frankfurt am Main, Germany
Abstract
To truly understand bioenergetic processes such as ATP synthesis, membrane-bound substrate transport or
flagellar rotation, systems need to be analysed in a cellular context. Cryo-ET (cryo-electron tomography)
is an essential part of this process, as it is currently the only technique which can directly determine the
spatial organization of proteins at the level of both the cell and the individual protein complexes. The need
to assess bioenergetic processes at a cellular level is becoming more and more apparent with the increasing
interest in mitochondrial diseases. In recent years, cryo-ET has contributed significantly to our understanding
of the molecular organization of mitochondria and chloroplasts. The present mini-review first describes the
technique of cryo-ET and then discusses its role in membrane bioenergetics specifically in chloroplasts and
mitochondrial research.
Introduction
Membrane bioenergetics is the study of energy-converting
processes involving biological membranes, including ATP
production, active transport of substrates and flagellar
rotation. Key to these processes is the use of electrochemical
gradients to drive energy-demanding reactions. The sources
of these gradients are often distinct from the consumer and
their effect on reaction rates is a fundamental question in
bioenergetics research.
ATP is the universal energy currency of all living
organisms. Its synthesis is catalysed primarily by the
membrane-bound enzyme F1 Fo -ATP synthase. In eukaryotes, this enzyme is located in the inner membrane of
mitochondria and in the thylakoid membranes of plant
chloroplasts. The F1 Fo -ATP synthase of these organelles
uses the energy stored in an electrochemical gradient of
protons to power the conversion of ADP and Pi into
ATP by rotary catalysis [1]. The proton gradients are
formed by either oxidative phosphorylation (mitochondria)
or photosynthesis (chloroplasts) using proteins distinct from
the F1 Fo -ATP synthase. These proteins either pump protons
across membranes while transferring electrons along a redox
pathway (NADH dehydrogenase, cytochrome c reductase
and cytochrome c oxidase in mitochondria, and cytochrome
b6 f in chloroplasts) or generate protons by catalysing the
oxidation of water [PSII (Photosystem II) in chloroplasts]
(Figure 1).
As early as the 1970s with the advent of freeze–fracture,
low-angle shadowing techniques in EM, it became evident
that the protein complexes involved in ATP synthesis by
photosynthesis were spatially separated [2,3]. Membrane
Key words: chloroplast, cryo-electron tomography, membrane bioenergetics, membrane protein
organization, mitochondrion.
Abbreviations used: cryo-ET, cryo-electron tomography; PSI, Photosystem I; PSII, Photosystem
II.
1
To whom correspondence should be addressed (email [email protected]).
Biochem. Soc. Trans. (2013) 41, 1227–1234; doi:10.1042/BST20130029
fracture planes that showed both stacked and unstacked
thylakoid membranes of chloroplasts revealed a non-random
distribution of protein complexes. Proteins on the unstacked
membrane surfaces were larger and more dispersed than
those on the stacked membranes. Through a combination of
immunolabelling and biochemical methods, it was concluded
that PSII was located primarily on the stacked membranes,
whereas PSI (Photosystem I), along with ATP synthase, were
located on the unstacked membranes [4,5].
Similar experiments to determine the distribution of
proteins complexes involved in ATP synthesis by oxidative
phosphorylation were not reported until 1989 when Allen
et al. [6] used the technique of freeze–fracture deep-etch
EM to analyse protein distribution in mitochondria from the
protist Paramecium multimicronucleatum. Here two rows of
interdigitating particles forming a ‘zipper-like’ arrangement
were seen along the outer edge of helical cristae membranes
and a single row of 13-nm-wide particles along the inner edge.
These particles were thought to be ATP synthase and NADH
dehydrogenase (Complex I) on the basis of their size and
shape, but no further proof was obtained. Similar experiments
with mitochondria from other species were unsuccessful, but
proteomic studies [7,8] and 2D class averaging by singleparticle EM [9,10] have shown that both ATP synthase
and the respiratory chain complexes do form higher-order
assemblies in the cristae membranes of various species.
Progress in assessing the effect of protein distribution on
the rates of enzyme reactions that involve electrochemical
gradients requires a technique which can directly determine
protein identity and distribution at nanometre resolution
in an unperturbed cellular context. Although fluorescence
microscopy has recently broken the theoretical resolution
limit of light microscopy {STED (stimulated emission depletion), PALM (photo-activated localization microscopy),
etc. [11]}, the currently obtainable resolution (20–100 nm)
is still below that required for assessing the distribution
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Figure 1 ATP synthesis in mitochondria and chloroplasts
Upper panel: in mitochondria, ATP is generated by oxidative phosphorylation. Electrons are transferred from the electron
donors NADH + and FADH + to O2 via NADH dehydrogenase (Complex I, blue), ubiquinone (UQ), cytochrome c reductase
(Complex III, orange), cytochrome c (black), and cytochrome c oxidase (Complex IV, green). During electron transfer,
Complexes I, III and IV pump protons across the inner mitochondrial membrane generating the electrochemical proton
gradient used by ATP synthase (Complex V, cream) to produce ATP. Additional electrons from succinate oxidation enter
the electron-transfer pathway via succinate dehydrogenase (Complex II, pink), which reduces ubiquinone, but does not
pump protons. Lower panel: in chloroplasts, ATP is generated by photosynthesis. Electrons derived from water oxidation
are transferred to NADP + via PSII (blue), plastoquinone (PQ), cytochrome b6 f (orange), plastocyanin (pink), PSI (green),
ferredoxin (Fd, brown) and ferredoxin–NADP reductase (FNR, deep red). Light energy captured by the light-harvesting
complexes (LHCI, LHCII, cyan, and minor LHC, lilac) and transmitted to PSI or PSII promotes electron transfer by exciting
electrons to a higher-energy state. During electron transfer, cytochrome b6 f pumps protons across the thylakoid membrane
and, together with water oxidation, catalysed by PSII, forms the electrochemical proton gradient used by ATP synthase
(cream) to produce ATP. Membrane bilayers are represented by blue bars. Figure created with Chimera using the PDB codes
3M9S, 1NTK, 3AEF, 2Y69, 2O01, 1Q90, 1A70, 1GJR, 2BHW, 3BQV, 3ARC and 3PL9. Protein density maps (except ATP synthase)
were created in CCP4 by calculating MTZ files from PDB files with a resolution cut-off of 30 Å (1 Å = 0.1 nm). The ATP
synthase density map was obtained from the EM database (emd-1357) and fitted with PDB codes 3V3C and 1FX0 or 4B2Q.
of neighbouring proteins in crowded environments such as
thylakoids or cristae membranes. Cryo-ET (cryo-electron
tomography), on the other hand, generates 3D volumes of
biological samples in the resolution range 2–5 nm. We have
thus used this technique to determine the distribution of
proteins involved in ATP synthesis in both chloroplast and
mitochondria [12–15].
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Authors Journal compilation Cryo-ET
Cryo-ET is a relatively new technique that can image cellular
features in three dimensions as well as the distribution and,
in favourable cases, the structure of protein complexes in
situ. The technique has been used to investigate biological
structures ranging from isolated protein and viral particles to
whole cells and organelles [16–19]. The principles of electron
Bioenergetics in Mitochondria, Bacteria and Chloroplasts
tomography were worked out some time ago [20–22], but
its routine application to cryo-specimens has depended
on the development of suitable preparation techniques,
cryo-capable electron microscopes, sufficiently large digital
cameras and automated data collection software [23–26].
Cryo-ET combines the principles of 3D volumetric
imaging (tomography) with cryo-preservation of samples.
The process of collecting a tomogram works in a similar
way to 3D medical imaging in CAT (computerized axial
tomography) or MRI. The sample is placed inside an
appropriate instrument and a series of projection images are
acquired at different angles. These images are then aligned
relative to each other in the computer and re-projected along
the acquisition angle to generate a 3D volume [19].
Sample preparation
To image protein structures, samples have to be prepared
without the use of destructive stains or fixatives, protected
from dehydration in the high vacuum of the electron
microscope and imaged with low electron doses to limit
radiation damage. For cryo-EM and cryo-ET, samples are
rapidly cooled to liquid nitrogen temperature at freezing
rates greater than 105 K·s − 1 to avoid the formation of ice
crystals [23]. Samples that can be thinly spread on to EM
grids, such as purified proteins, organelles, virus suspensions
or small bacteria, are usually plunge-frozen. Solutions are
applied to EM grids covered with a 100–200-nm-thick holey
carbon support film, blotted to remove excess liquid and
immediately plunged into liquid ethane using a guillotine
device [27]. Larger cells, which produce thin protrusions of
interest, e.g. axons, synapses or filopodia, can also be frozen
using this method [28–30].
Thicker samples, such as tissues, whole cells or large
cellular compartments such as chloroplasts, have to be frozen
at high pressure to obtain good freezing rates to depths of a
few hundred microns. Samples are placed in protective metal
carriers and exposed to high-pressure jets of liquid nitrogen
[31,32]. The frozen samples are then sliced with a cryomicrotome into 50–250-nm-thick sections and transferred to
an EM grid, all at liquid-nitrogen temperature [33].
Identification of proteins in tomograms
The greatest difference between conventional roomtemperature EM of plastic sections and cryo-ET is the level
of observable detail. Figures 2(A) and 2(B) show plastic
sections of mitochondria and a chloroplast. In both images,
the membranes show up clearly, but no molecular detail is
visible. By contrast, slices through tomographic volumes of
plunge frozen mitochondria (Figure 2C) or high-pressure
frozen chloroplasts (Figure 2D) show large protein complexes
such as ATP synthase (yellow arrowheads) and PSII (red
arrowheads) in the membranes.
Protein densities in tomograms can be identified by various
techniques. Large protein complexes with characteristic
shapes such as ATP synthase, PSII and ribosomes can
usually be detected by eye (Figures 2C and 2D), but for
unbiased identification, template matching is often employed.
For this approach, atomic models of known structure are
filtered to a resolution of 40–60 nm and used as a template
to search entire tomographic volumes [34]. This technique
has been used to identify large protein complexes such as
ribosomes in whole cells [35–37]. Success is highly dependent
on sample thickness, contrast and information content, as
well as the size and abundance of the target protein. The
accuracy of template matching falls not only with protein
size and abundance, but also with increasing molecular
crowding [34,37]. Therefore this method is currently not
appropriate for the analysis of protein-dense organelles such
as mitochondria or chloroplasts.
Smaller proteins or proteins of unknown or uncharacteristic shape have to be identified by labelling. Electron-dense
tags such as colloidal gold or quantum dots conjugated to
primary or secondary antibodies are easily visible in cryotomograms. Protein densities within a ∼23 nm radius of these
tags are likely to be the protein of interest. Using this method,
we have identified NADH dehydrogenase in mitochondrial
membranes [13] and have determined the orientation and
subunit topology of the pre-protein translocase in the
chloroplast outer membrane (TOC) [38]. Electron-dense
fusion tags, akin to GFP for light microscopy, are currently
being developed for cryo-ET. One promising candidate is
metallothionein, a 6 kDa heavy-metal-binding protein, which
appears as a 2 nm black density in tomograms. This tag has
been used to identify proteins associated with microtubules
and intermediate filaments [39].
Protein structure determination and
organization
Subtomogram averaging is a technique which can both
identify known proteins and determine previously unknown
protein structures within tomograms [14,16–18,40] (Figure 2E). Individual subvolumes containing the protein of
interest are extracted from the tomogram, aligned with each
other and averaged together using specialized software. By
subvolume averaging, characteristic features of a particular
protein are amplified, whereas random features around it are
averaged out increasing the signal-to-noise ratio and hence
observable structural detail. During the alignment procedure,
a list of co-ordinates and rotation angles are produced
describing how the final average aligns with each extracted
subvolume. The resulting average or fitted atomic model
can then be positioned back into the tomographic volume
to provide information about the spatial organization of the
target protein within the cell or organelle. Using this method,
we have determined the arrangement of ATP synthase dimers
in mitochondrial cristae and PSII in thlyakoid membranes
[12,14] (Figures 2G and 2H).
Cryo-ET and membrane bioenergetics
Using cryo-ET, our laboratory has uncovered the structure
and distribution of proteins involved in ATP synthesis
in both mitochondria and chloroplasts. One of our most
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Figure 2 EM of mitochondria and chloroplasts
Image of (A) mitochondria from the yeast Pichia pastoris, and (B) a chloroplast from Marchantia, prepared by conventional
EM on resin-embedded samples. Membranes are clearly visible in both images, but molecular information is lacking. (C)
Slice through a tomogram of a cryo-preserved mitochondrion from the fungus Podospora anserina, and (D) a chloroplast
from spinach. Membranes and protein densities such as ATP synthase (yellow arrowheads) and PSII (red arrowheads)
are clearly visible. (E) Subtomogram average of ATP synthase dimers picked from mitochondrial membranes of the
fungus P. anserina (left) and fitted with the atomic X-ray model PDB code 4B2Q (right). (F) Comparison of the protein
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densities attributed to PSII in thylakoid membranes (right) and the atomic model of PSII, PDB code 2O01 filtered to 30 Å (1 Å
= 0.1 nm) (surface view, left; slice through volume, middle). Both densities are roughly the same size, two-fold symmetric
and have two kidney-shaped protrusions on one side of the complex. (G) Surface-rendered volume of (C) showing the
location of ATP synthase dimers (yellow spheres) in the cristae membranes. Rows of dimers are always observed along
the highly curved membrane ridges of lamellar cristae. (H) Tomographic slice (top left) and corresponding diffraction pattern
(top right) of a PSII crystal in stacked thlyakoid membranes. PSII–LHCII (light-harvesting complex II) supercomplex [51] was
positioned into the tomographic volume by aligning the two protruding densities described in (F) to reveal the packing of
PSII within (bottom left) and between (middle right) adjacent thylakoid membranes. PSII–LHCII supercomplexes are oriented
in the same direction within a membrane, but rotated 90◦ in neighbouring membranes. Connections between adjacent
membranes are mediated by the negatively charged N-termini of LHCII (bottom right, black arrowhead, see [12] for details).
Scale bars: (A) 200 nm, (B) 500 nm, (C, D, G and H) 100 nm. (D, F and H) Reproduced from Daum, B., Nicastro, D., Austin,
J., McIntosh, J.R. and Kühlbrandt, W. (2010) Arrangement of Photosystem II and ATP Synthase in Chloroplast Membranes of
c American Society of Plant Biologists. (G) Reproduced
Spinach and Pea. Plant Cell 22 (4): 1299–1312. www.plantcell.org. with kind permission from Davies, K.M., Strauss, M., Daum, B., Kief, J.H., Osiewacz, H.D., Rycovska, A., Zickermann, V.,
Kühlbrandt, W. (2011) Macromolecular organization of ATP synthase and complex I in whole mitochondria. Proc. Natl. Acad.
Sci. U.S.A. 108(34): 14121–14126.
striking findings was the long rows of dimeric ATP synthase,
which were observed in cristae membranes from mammals,
fungi and plants [13,15], but not in chloroplasts [12] or
the plasma membranes of bacteria or archaea (B. Daum
and A. Mühleip, unpublished work). Although rows of
mitochondrial ATP synthase dimers were first reported by
Allen et al. [6] over two decades ago, cryo-ET has provided
the first direct 3D visualization of these rows and has shown
by subtomogram averaging that the particles are, in fact,
ATP synthase [13,14]. In addition, we have shown directly
that ATP synthase dimers introduce sharp local membrane
curvature in the cristae membranes [13,14].
ATP synthase dimer rows are mostly found on the tightly
curved ridges of lamellar cristae. Disruption of the ATP
synthase dimers in Saccharomyces cerevisiae, through the
deletion of the ATP synthase subunits e or g, led to a
profound change in cristae morphology in which sharp
local membrane curvature was no longer observed [14].
Instead of the lamellar cristae typical for wild-type yeast
strains, mitochondria of these mutants contained a number
of inner membrane vesicles, which occasionally formed
balloon-shaped protrusions similar to those predicted by
mathematical modelling [41]. In these mutants, ATP synthase
monomers were randomly distributed over the entire inner
membrane surfaces, but no dimers were observed. Yeast
mutants lacking subunits e and g have longer generation times,
a lower membrane potential and decreased decoupling rates
compared with wild-type mitochondria, but no alteration in
the functionality of the ATP synthase complex [42,43]. The
reduced viability of these mutants is therefore most likely to
be due to the disruption of the ATP synthase organization in
the membrane rather than a loss of enzyme functionality.
In contrast with ATP synthase, most other proteins
involved in mitochondrial ATP synthesis are either too small
(Complex III and Complex IV) or cannot easily be identified
in tomographic volumes (Complex I). Nevertheless, using antibody labelling, we were able to identify Complex I in cristae
membranes [13]. This complex appeared to be randomly
distributed, at either side of the rows of ATP synthase dimers,
rather than forming respiratory strings or patches predicted
by proteomics and EM on detergent-solubilized protein complexes [7–9]. Complex I, however, does form supercomplexes
with Complex III and IV in some species [44], but higherorder associations, e.g. Complex I dimers in the membrane,
have not been found. The occurrence of supercomplexes
appears to be related to the increased energy requirements of
certain tissues or organisms, but the random distribution
of Complex I to ATP synthase is a common feature of all
organisms studied. A consequence of this organization is
that the proton pumps (Complex I) are physically separated
from the proton sinks (ATP synthase) (Figure 3A). The effect
of this separation on ATP synthesis rates is unknown, but its
disruption e.g. by e and g mutants described above, does
appear to reduce the cell’s fitness [14,42,43].
In chloroplasts, our cryo-ET findings fully support the
lateral heterogeneity of photosynthetic membrane protein
complexes described previously using freeze–fracture EM
[12,45]. In accordance with these studies, we found that
the chloroplast ATP synthase was located solely on the
stroma-exposed thylakoid membranes and PSII in the stacked
grana membranes, where paracrystalline PSII arrays were
occasionally observed [12] (Figures 2D and 2H). In contrast
with the mitochondrial ATP synthase, the chloroplast ATP
synthase is entirely monomeric and randomly distributed
in the flat stroma-exposed membranes. No ATP synthase
complexes were found in the highly curved margins of the
thylakoid membranes [12]. The separation of ATP synthase
from PSII again leads to the segregation of proton sources
and sinks as observed in mitochondrial cristae.
Previous electron tomography of cryo-sections [12] or
serial sections of plastic-embedded chloroplasts [46] has
shown that the unstacked stroma thylakoids wind around
the stacked membranes in a helical fashion and fuse with
successive stacked grana thylakoids by small tubular or
lamellar protrusions [46,47]. The size of these openings
and the number of membranes in a stack appear to be
highly variable [46,48]. As the proton sources (PSII) and
proton sinks (ATP synthase) of the thylakoids are located in
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Figure 3 Distribution of proton sources and sinks in mitochondria and chloroplasts
(A) In mitochondria, ATP synthases (yellow) form rows of dimers along highly curved edges of lamellar cristae, whereas
the respiratory chain complexes (green) are distributed randomly in the flat membrane regions. This distribution segregates
proton sources and sinks, and is thought to result in a directional flow of protons (red spheres) towards the cristae edges (red
arrows). Reproduced with kind permission from Davies, K.M., Strauss, M., Daum, B., Kief, J.H., Osiewacz, H.D., Rycovska, A.,
Zickermann, V., Kühlbrandt, W. (2011) Macromolecular organization of ATP synthase and complex I in whole mitochondria.
Proc. Natl. Acad. Sci. U.S.A. 108(34): 14121–14126. (B) In chloroplasts, the segregation of proton sources and sinks is even
more extreme. ATP synthases (proton sinks, yellow) are located on the stroma-exposed thylakoid membranes and PSIIs
(proton source, blue) are located in the stacked grana membranes. These two membrane compartments are separated by
narrow openings, which are likely to restrict protein and ion movement. Protons (red spheres) generated by water oxidation
activity of PSII must exit the grana membranes via these connections in order to find their sinks (ATP synthase), which are
located up to a few microns away.
different subcompartments of the thylakoid membranes, the
diameter of these putatively dynamic membrane connections
is likely to affect the rate of ATP synthesis under
different environmental conditions by controlling the flow
of ions and proteins between the two compartments
[46] (Figure 3B). To test this hypothesis, ATP synthesis
rates and membrane protein distribution under different
environmental conditions needs to be investigated in parallel.
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Authors Journal compilation This is only possible through the combination of membrane
bioenergetics and cryo-ET.
Conclusion
We have used cryo-ET to show that proton sources
and proton sinks are physically segregated in the
energy-converting membranes of both mitochondria and
Bioenergetics in Mitochondria, Bacteria and Chloroplasts
chloroplasts. This separation is more extreme in chloroplasts
than in mitochondria, but the common principle indicates
a fundamental underlying energetic advantage to this arrangement. We have already seen that disruption of this protein
distribution in mitochondria through the deletion of the
dimer-specific subunits of ATP synthase results in decreased
cell viability, suggesting that the separation of proton sources
and sinks in mitochondria is required for high rates of
ATP synthesis [15,42,43]. This is likely to pertain also
to chloroplast thylakoids, where the membrane proteins
of grana and stroma membranes intermix randomly when
membrane stacking is abolished [49,50]. At present, the
underlying membrane organization can be visualized only by
cryo-ET. Therefore, to truly understand energy-converting
processes at the level of the cell, bioenergetic measurements
must be combined with cryo-ET. These two techniques,
when used in combination, are likely to become a powerful
tool especially when investigating the molecular causes of
mitochondria-related human diseases and aging.
Acknowledgements
We thank Werner Kühlbrandt for helpful comments on the paper,
Friedricke Joos for providing images for Figures 2(A) and 2(B), and
Paolo Lastrico for preparing Figure 3.
Funding
This work was funded by the Max Planck Society (to K.M.D. and
B.D.).
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Received 13 March 2013
doi:10.1042/BST20130029