7Commentary

IntroductionCellular activities rely on the concerted actions of macromolecularcomplexes that function within dynamic networks. For instance,the cell cytoskeleton is remodeled within fractions of a second, thusmodulating cell shape and function. Consequently, at the molecularlevel, cellular architecture rapidly changes during the cell cycle andthroughout various biological processes. Despite the wealth ofinformation that exists on cellular components and their dynamicproperties, our current understanding of the functional interactionsthat lead to a given cellular process is rather limited. Developingthe experimental tools necessary for the analysis of complex andvariable supramolecular structures inside cells is crucial to rectifythe situation. Microscopical imaging techniques, which now offerresolution at the previously unattainable nanometer scale, wouldthus be expected to provide novel insight into the local and globalorganization of functional modules and networks inside cells(Robinson et al., 2007).

Microscopy techniques have traditionally been central in drivingcell biology forward. Fluorescence microscopy and confocal laser-scanning microscopy revolutionized our thinking and opened up alarge set of possible strategies for investigating cellular processes(reviewed by Schliwa, 2002). In particular, the introduction of greenfluorescent protein (GFP) and its analogs has allowed kineticmeasurement of proteins in living cells (Tsien, 1998). Additionally,the development of ultra-high-resolution fluorescence microscopy,such as PALM (Betzig et al., 2006), STED (Hell, 2003) and structuredillumination (Schermelleh et al., 2008), allowed the visualization ofindividual macromolecular complexes such as the nuclear porecomplex (NPC) (Schermelleh et al., 2008). However, such approachescan focus on only a limited number of proteins at a time, dependingon the number of fluorophores available. The molecular architecturecan, however, be reconstructed in three dimensions and at highresolution by electron microscopy (EM), particularly cryo-electrontomography (cryo-ET), therefore complementing fluorescence-microscopy techniques.

In this Commentary, we focus on the principles and implementationof cryo-ET in the field of cell biology. We demonstrate the possibility of resolving three-dimensional (3D) cytoskeleton networkswithin intact cells by showing actin cytoskeleton networks in thenative context of membranes, vesicles and other molecular complexes.The potential of using cryo-ET for following viruses as they infectcells will be shown and discussed. We also consider how theapplication of cryo-ET to visualize intact nuclei, in combination with3D-averaging procedures, has yielded a 3D structure of active NPCs.Finally, we consider future prospects for cryo-ET. We focus on theapplication of this technique to eukaryotic cells; however, it shouldbe noted that cryo-ET has been successfully used for the study ofprokaryotes and viruses (Borgnia et al., 2008; Briggs et al., 2006;Gan et al., 2008; Grunewald et al., 2003; Kurner et al., 2005; Lieberet al., 2009; Liu et al., 2008; Morris and Jensen, 2008).

Cryo-ET: technical notesUniquely among imaging techniques, cryo-ET can generate 3Dinformation concerning the macromolecular architecture of cells inan unperturbed state (Cyrklaff et al., 2007; Li et al., 2007; Nickellet al., 2006). Using this technique, one can depict unique cellularstates and reconstruct molecular networks. Through vitrification byrapid freezing, biological material can be physically fixed, ensuringclose-to-life conditions in samples prepared for cryo-ET (Dubochetet al., 1988). Because neither chemical fixation nor staining isneeded, the delicate cellular landscape is preserved during samplepreparation and accurately depicts in vivo conditions.

In cryo-ET, 3D structures of specimens are retrieved from 2Dmicrographs. Owing to the large depth of focus, electronmicrographs are essentially two-dimensional (2D) projections of a3D object in the direction of the electron beam. Consequently,features of the sample are superimposed and cannot be separated,in contrast to the situation in confocal laser-scanning microscopy.Nevertheless, the three-dimensionality of an object can be retrievedby recording a series of projections at varying angles and

Visualizing cellular processes at the molecular levelby cryo-electron tomographyKfir Ben-Harush1, Tal Maimon1, Israel Patla1, Elizabeth Villa2 and Ohad Medalia1,*1Department of Life Sciences and the National Institute for Biotechnology in the Negev, Ben-Gurion University, Beer-Sheva, 84105 Israel2Max-Planck-Institute for Biochemistry, D-82152 Martinsried, Germany*Author for correspondence (omedalia@bgu.ac.il)

Journal of Cell Science 123, 7-12 Published by The Company of Biologists 2010doi:10.1242/jcs.060111

SummaryThe cellular landscape rapidly changes throughout the biological processes that transpire within a cell. For example, the cytoskeletonis remodeled within fractions of a second. Therefore, reliable structural analysis of the cell requires approaches that allow for instantaneousarrest of functional states of a given process while offering the best possible preservation of the delicate cellular structure. Electrontomography of vitrified but otherwise unaltered cells (cryo-ET) has proven to be the method of choice for three-dimensional (3D)reconstruction of cellular architecture at a resolution of 4-6 nm. Through the use of cryo-ET, the 3D organization of macromolecularcomplexes and organelles can be studied in their native environment in the cell. In this Commentary, we focus on the application ofcryo-ET to study eukaryotic cells – in particular, the cytoskeletal-driven processes that are involved in cell movements, filopodiaprotrusion and viral entry. Finally, we demonstrate the potential of cryo-ET to determine structures of macromolecular complexes insitu, such as the nuclear pore complex.

Key words: Actin, Cytoskeleton, Cryo-electron tomography, Herpes simplex virus, Nuclear envelope

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synthesizing these projections into a 3D density map – that is, atomogram (Fig. 1) (Frank, 1992). In practice, the different projectionimages are collected by tilting the specimen incrementally arounda single axis inside the electron microscope that is perpendicularto the optical axis of the electron beam (Fig. 1A). This tilt seriesis then aligned to a common frame of reference, followed bycalculation of the tomogram (Fig. 1A), most commonly by usinga ‘weighted back-projection’ algorithm (Radermacher, 1992).

The resolution of a tomogram is directly dependent on the angularincrement between two adjacent projections and on the total numberof images that are obtained (Koster et al., 1997). Therefore, the aimis to collect as many tilted projections as possible, covering thewidest possible angular range, but keeping the electron dose at asub-critical level. Thus, the cumulative electron dose in the entiretilt series must be kept within tolerable limits, typically notexceeding ~6000 e–/nm2, to prevent radiation damage to thebiological specimen. Furthermore, because of technical limitations,the tilt series cannot cover the entire spectrum of views and is limitedto ±70°. In practice, a typical tilt series consists of 80-100 exposuresand covers only 120°-140° of the 180° angular range. Consequently,elongation of features along the beam axis is evident because of amissing ‘wedge’ in the 3D Fourier space (Frank et al., 2002).Overall, to minimize the exposure time and to increase the accuracyof the process, data acquisition must be fully automated, and relieson computer control (Dierksen et al., 1993; Dierksen et al., 1992).

A further limitation is that the application of cryo-ET toeukaryotic cells is restricted to relatively thin regions. When theobject is thicker than the mean free path of an electron [200 nmand 350 nm for 120 keV and 300 keV (acceleration voltage ofelectrons in the electron microscope), respectively (Grimm et al.,1998)] multiple scattering events substantially degrade the image

quality, despite the use of high to medium acceleration voltage(300 keV) and an energy filter to minimize this effect (Grimm et al., 1996). As a consequence, samples thicker than 1 m canbarely be studied in toto, and require cryosectioning before theycan be subjected to tomographic analysis. Several laboratories havedevoted substantial effort to establishing freeze-hydratedcryosectioning procedures. The feasibility of this approach has beenshown using cryosectioned rat liver cells, mouse epidermis, andhuman epidermis and cardiomyocytes (Al-Amoudi et al., 2004;Al-Amoudi et al., 2007; Castano-Diez et al., 2007; Gruska et al.,2008; Hsieh et al., 2002; Salje et al., 2009), but it is still technicallydemanding and often gives rise to sectioning artifacts (Al-Amoudiet al., 2005).

3D visualization of cytoskeletal networksFor decades, EM of actin cytoskeletons was performed on sectionsof chemically fixed, detergent-extracted cells shadowed with metal(Brown et al., 1976; Svitkina et al., 1997), or by using adherentcells, which have an apical surface that has been mechanicallyremoved, for EM analysis (Hartwig et al., 1989; Heuser andKirschner, 1980). Although these methods provided importantinsight into the architecture of actin networks (Small et al., 1994),the spatial resolution of the structures revealed by thesemethodologies was limited, especially in the third dimension. Forinstance, connections between actin filaments were difficult toresolve in detail after metal decoration or replica formation.Moreover, distortion of the actin network by detergent treatmentdoes not permit the study of anchorage of actin filaments tomembranes. Most importantly, by means of total internal reflectionfluorescence (TIRF) microscopy using specific probes for actinnucleation, growth and branching (e.g. Arp2/3) it was shown that,

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Fig. 1. Applying cryo-ET to eukaryotic cells. (A)2Dprojections at different tilt angles for individual 3D objects,such as an intact eukaryotic cell, are recorded by tilting thespecimen holder; the projections typically cover ~120°. Theholder is tilted incrementally around an axis that isperpendicular to the electron beam. All tilted projections aresynthesized into a 3D density map, typically by applying a‘weighted back-projection’ algorithm (Radermacher, 1992).Shown are projections and a reconstructed volume of aD. discoideum cell, adapted from Medalia et al. (Medalia etal., 2002). Scale bar: 300 nm. (B)Surface-rendering view ofthe reconstructed volume shown in A shows the actinfilaments (red), cell membrane (purple) and largemacromolecular complexes, mostly ribosomes (green). Thesurface-rendering views were segmented semi-automatically.Colors were chosen subjectively. Branches of actin filamentsas found at the cell cortex are shown in the lower panel.(C)Surface-rendering view of the reconstructed volume of ahuman fibroblast cell shows all cytoskeletal elements. Corticalactin (red) is located along the cell membrane (purple),whereas intermediate filaments (turquoise) are localizedfurther into the cell interior; these present a wider diameter(~10 nm), a different texture and a lower persistence lengththan actin. In addition, one microtubule (pink) is found in theupper left corner of the tomogram in close proximity to acluster of ribosomes (green). Scale bar: 100 nm (also for B).(B)Adapted from Medalia et al. (Medalia et al., 2002).

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in highly motile cells, the proteins associated with cytoskeletalnetworks have half-lives in the order of 7-10 seconds (Bretschneideret al., 2004). Therefore, it is to be expected that actin-bundling and-crosslinking proteins, which are crucial for maintaining thearchitecture of the actin network, might dissociate from actinfilaments and redistribute within a specimen during the course oftraditional-preparation EM procedures.

In live cells, actin structures rapidly reorganize during motility,endocytosis and cytokinesis (Dalous et al., 2008; Kaksonen et al.,2004; Pantaloni et al., 2001; Pellegrin and Mellor, 2007; Walpitaand Hay, 2002). Because of these dynamics, immediate arrest ofthe cellular processes in intact cells is a prerequisite for obtainingfaithful information regarding actin networks. Electron tomographyof vitrified but otherwise unaltered cells has proven to be a keytechnique for the 3D reconstruction of actin architecture (Medaliaet al., 2002). It was demonstrated that cryo-ET of intactDictyostelium discoideum cells could reveal the connections of theactin-filament network with the plasma membrane (Fig. 1B, upperpanel), as well as massive actin filaments branching at various angles(Fig. 1B, lower panels), without the need for chemical fixation orheavy-metal decoration (Medalia et al., 2002). Similar views canbe obtained when cells of higher eukaryotes are studied (Fig. 1C).

In the future, our understanding of cell motility and othercytoskeleton-dependent processes will be increased by investigatingthe 3D organization of the actin system in relation to specificfunctional states, such as during consecutive steps of particle uptakeby a phagocyte or at cell-adhesion sites. The possibility of arrestingcells instantly also allows the investigation of rapid cellularprocesses such as filopodial protrusion, which is discussed below.

Zooming in on actin remodeling duringfilopodium formationFilopodia are finger-like plasma-membrane protrusions that areinvolved in adhesion to the extracellular matrix (ECM), sensing theenvironment and cell-cell signaling (Chhabra and Higgs, 2007; Mattilaand Lappalainen, 2008). Filopodia also represent an excellent modelsystem for describing the process of actin-driven membraneprotrusion. These structures grow at their tips through the assemblyof actin and are stabilized along their lengths by a core of bundledactin filaments (Pollard and Borisy, 2003). Given their relatively lowthickness (150-400 nm), filopodia are also excellent cellular structuresfor study by cryo-ET. With this approach, the organization of actinfilaments and membranes to which the actin network is anchored canbe carefully analyzed in an unperturbed state.

We used cryo-ET to track the length and relative position ofindividual filaments within D. discoideum filopodia, which alloweda quantitative analysis of actin filaments. The data revealed thatactin filaments in these fast-moving cells are not continuousthroughout the entire protrusion (Fig. 2) (Medalia et al., 2007).Importantly, it was shown that the filopodial tip comprises manyshort filaments that interact with the membrane at their distal andproximal ends. These filaments, arranged at the tip in a cone shape,have been suggested to provide the driving force that pushes themembrane forward (Gerisch and Weber, 2007). Their location andlength support the notion that sites of de novo actin-filamentnucleation and growth are confined to the tip of the filopodia (Faixet al., 1992; Faix and Grosse, 2006; Medalia et al., 2007). It isnoteworthy that no vesicles are found within filopodia that arevisualized by cryo-ET, implying that the membranes needed for the formation of protrusions are supplied at a distance from thefilopodial tip. The presence of Dia2, a formin with actin-nucleating

activity that is important for filopodia formation and maintenance(Schirenbeck et al., 2005), at the tips of filopodia in D. discoideumimplies that actin filaments undergo nucleation through filopodiaprotrusion. When filopodia protrude, the actin filaments grow andare then bundled while laterally connecting to the membrane alongthe filopodial shaft (Medalia et al., 2007). In general, we found thatD. discoideum filopodia are characterized by a discontinuity of actinfilaments along the filopodial axis (Fig. 2C). However, sometransverse filaments connect to the membrane and the shaft filamentsare found in the tip-shaft zone.

This analysis can thus be explained by the ‘sequential-nucleationmodel’ (Medalia et al., 2007), which proposed that the sites of denovo nucleation and growth of actin filaments are confined to thefilopodium tip. The short filaments located at the tip detach fromand reattach to the cell membrane with their distal and/or proximalends, thus enabling actin polymerization. The growing filamentsare then bundled and laterally connect to the cell membrane alongthe filopodia shaft. Within the shaft zone, actin filaments are bundledand axially oriented (Fig. 2C). The unique organization andarrangement of D. discoideum filopodia can presumably beattributed to the fast motility of these cells. That is, the apparentdiscontinuity of actin filaments might be a property of filopodia in

Fig. 2. The architecture of D. discoideum filopodia. (A)A 50-nmtomographic slice through a filopodium demonstrates the discontinuity of thefilopodial actin filaments. Short filaments are found at the tip of thefilopodium, and these are distinct from the transverse and straight filamentsfound along the filopodial shaft. Scale bar: 200 nm. (B)Surface-renderingview of the filopodium (boxed area in A) reveals the overall organization ofthe actin network (red) and the interaction of actin filaments with the plasmamembrane (blue) at the filopodial tip. Macromolecular complexes are shownin green. (C)The ‘sequential-nucleation model’ (see text) is illustrated. At thetip of the filopodium, de novo nucleation and growth of actin filaments occur(Medalia et al., 2007). In the shaft zone, actin filaments are bundled andaxially oriented along the filopodium and, within the tip-shaft zone, growingactin transverse filaments are connected to the membrane and to the shaftfilaments. Adapted from Medalia et al. (Medalia et al., 2007).

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fast motile cells, which differs from filopodia in other adherent cells,which are characterized by continuous actin filaments.

Remodeling of actin networks during viral entrySeveral viruses use an endocytic mechanism to enter cells prior toremodeling cortical actin (Greber, 2002; Munter et al., 2006). Inaddition, filopodia and other cellular protrusions are susceptible toviral docking, which eventually leads to cell infection (Clement et al.,2006). Cryo-ET can provide a unique tool for studying infected cells,supplying unprecedented information on the stages of viral assemblyand maturation within cells. In a pioneering study, Maurer et al. showedsnapshots of the entry of Herpes simplex virus 1 (HSV-1) into cells(Maurer et al., 2008). By means of cryo-ET, the authors showed thatcytosolic capsids of HSV-1 were located between actin bundles withinPtK2 cells. They also identified capsids between individual actinfilaments in the cell cortex. Furthermore, the data revealed that thevirus does not induce local depolymerization of actin in its vicinity,on the basis of morphological appearance of the cell cortex, but rathermight be involved in remodeling the dense cytoskeletal network asshown by Clement et al. (Clement et al., 2006) (Fig. 3).

The NPC: combining cryo-ET and single-particle approachesCryo-ET is primarily a static tool. However, by collecting a largenumber of datasets and correlating them with pre-existinginformation, one can acquire information about a dynamic processat the molecular level, as has been described above for filopodiaand virus-infected cells. Thus, the remodeling and structural changesof macromolecular complexes that transpire during cellularprocesses can be examined. Additionally, a detailed 3Dreconstruction of macromolecular complexes in situ can be achievedby combining cryo-ET with 3D-averaging procedures (Bartesaghiet al., 2008; Bostina et al., 2007; Forster et al., 2005).

Such a hybrid technique (cryo-ET and 3D-averaging approach)was applied to NPCs, which are large molecular machines that areembedded in the nuclear envelope and connect the nucleoplasm withthe cytoplasm by means of an aqueous channel. NPCs, which arecomposed of hundreds of proteins (Alber et al., 2007) arranged inpseudo-eightfold rotational symmetry, function as a selective barrier

(Rout et al., 2000) by allowing small molecules and ions to diffusefreely while mediating the passage of large molecules in an energy-dependent manner. Although work on the NPC structure using EMbegan in 1950 (Callan and Tomlin, 1950), only at the end of thetwentieth century were the main components of the structure revealed(Akey and Radermacher, 1993; Brohawn et al., 2009; Yang et al.,1998). Despite dimensional differences between species, the basicarchitecture of the NPC is conserved; the consensus structure consistsof a central spoke ring that is confined by a cytoplasmic and anucleoplasmic ring. Eight cytoplasmic filaments and a nuclear basketcomposed of eight filamentous structures join to form a distal ring.Owing to its sheer size, the NPC presents a major challenge forstructural determination. Applying cryo-ET to intact nuclei ensuresthat the NPC is arrested in its active form, as is evident from thepreservation of a nucleocytoplasmic gradient of Ran-GTP in isolatednuclei (Becskei and Mattaj, 2003; Becskei and Mattaj, 2005; Gorlichet al., 2003). The nuclear envelope can be viewed as an ellipsoid

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Fig. 3. Surface-rendering view of a Ptk2 cell infected by HSV-1. Two capsidsof recently entered virions (light blue) were found to reorganize and modify theactin bundles, but not depolymerize actin, upon viral entry. On the upper leftside, the virus-derived glycoprotein spikes (yellow) can be seen emerging fromthe membrane. Viral tegument (orange), cell and viral membrane (dark blue),actin (dark red; upper part cut away), and cellular vesicles (purple) are shown.Adapted from Maurer et al. (Maurer et al., 2008). Scale bar: 100 nm.

Fig. 4. Cryo-ET of the nuclear envelope. (A)A 32-nm tomographic slicethrough the nuclear envelope of a human fibroblast shows a central slice throughan NPC, which fuses the inner and outer nuclear membranes (INM and ONM,respectively). Scale bar: 100 nm. (B)Stereo-view representation of an averagedreconstructed volume of the human NPC. The central spoke ring is flanked bythe cytoplasmic ring (arrowheads) and the nuclear ring. (C)Schematicrepresentation of the nuclear envelope on the basis of cryo-ET of intact nuclei.The ONM is decorated with ribosomes (red), and the ONM and INM (yellow)are fused at the NPC (blue). Nuclear lamins (purple) are seen underlying theINM and interacting with the NPC via the nuclear basket (green). The structureof the lamin filaments was adapted from an in vitro cryo-ET study of theCaenorhabditis elegans lamin filaments (Ben-Harush et al., 2009).

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with NPCs embedded in its surface in all possible orientations; thus,extracting these elements in silico, followed by 3D alignment andaveraging, results in isotropic resolution, i.e. in all three dimensions.Using cryo-ET, we have resolved the NPC to 8-9 nm, in which someof the flexible filaments of the NPC were observed, in addition tothe scaffolding features of the complex (Beck et al., 2004). Animprovement in resolution (<6 nm) was later achieved when thestructure of the D. discoideum NPC was resolved without imposingeightfold symmetry (Beck et al., 2007). In this study, snapshots ofthe trajectory of cargo transported through the NPC were visualized.Recently, work from our laboratory has shown that the applicationof a similar approach using nuclei from human fibroblasts (Fig. 4)yielded the first insight into the structure of the human NPC (Eladet al., 2009). Although the resolved structures of the D. discoideumNPC and the preliminary structure of the human NPC share severalcommon features, such as the outer and inner diameters (~120 nmand ~50 nm, respectively), they differ in height and protein-densitydistribution, suggesting differences in protein positions (Elad et al.,2009). Currently, analysis of intact nuclear envelopes by means ofcryo-ET is limited to a resolution of 5-8 nm, which restricts structuralinterpretation to the level of determining the position of subcomplexes.Increasing the resolution of tomograms and the application ofspecimen-thinning techniques, such as cryosectioning (see above) andfocused ion beam (see below), will enable a step forward inunderstanding the functional organization of the nuclear envelope.

Conclusions and perspectivesCryo-ET is the method of choice in acquiring an insight into themolecular organization of cells and cellular components, such asfilopodia and actin filaments at the cortex of the cell, and to trackthe entry of viruses into cells. Additionally, it allows determinationof the 3D structure of large supramolecular assemblies in situ, aswe demonstrated for the NPC, at a medium resolution of 4-6 nm.

A major challenge facing cryo-ET concerns the identification ofmacromolecular complexes within a cellular context. The differentorientations of macromolecules and the current resolution of cellulartomograms prohibits unbiased identification of many molecularcomplexes, although some successful template-matching approacheshave been introduced (Frangakis et al., 2002; Ortiz et al., 2006).These procedures aim to identify specific macromolecularcomplexes in vivo on the basis of their structural fingerprint, bysearching for in-vitro-determined structural complexes in atomogram (van Heel et al., 2000). An approach based on electron-dense labeling must be developed to design a clonable tag that canbe genetically conjugated to proteins, which would facilitate theirlocalization in cryo-tomograms – that is, we are in need of a GFPanalog for cryo-electron microscopy. An elegant example of aclonable tag is metallothionein, a cysteine-rich protein that has beenshown to bind to multiple heavy atoms and can be detected by anelectron beam (Mercogliano and DeRosier, 2007). Such a labelingstrategy would provide a general solution for identifying complexeswhose structure is not yet determined or that intimately interact to form large assemblies. To make cryo-ET applicable not only tocellular protrusions and thin regions of the eukaryotic cell but alsoto thicker samples, there is a need to develop a reliable freeze-hydrated artifact-free sectioning technique that can be applied totissues and cells to produce optimal (thinner than 500 nm) biologicalsamples for cryo-ET. Alternative micro-dissection techniques thatinvolve using a focused ion beam to mill frozen samples arecurrently being developed. In these approaches, gallium ions (Ga+)are directed onto the frozen cell or tissue sample at a specific angle

and, by process of ‘sputtering’, selected parts of the sample can beremoved; this is known as ion-beam milling. Notably, this processdoes not cause major artifacts below 30 nm from the upper levelof the milled surface (Marko et al., 2006; Marko et al., 2007).Realization of this technology would open a window for the entryof cryo-ET into other branches of biology and might provide, forinstance, a bridge between structural and developmental biology.

Another direction of technical development lies in correlatingfluorescent and cryo-electron-microscopy images; this would greatlyhelp in identifying attractive locations within the cell forinvestigation by cryo-ET (Sartori et al., 2007; Schwartz et al., 2007),and would allow for the reconstruction of important cellularstructures. This approach would permit the identification of specificstates of cellular processes and would therefore eventually producestructural snapshots of such processes. Moreover, developingautomated fast algorithms would allow for a shortening of currenttime-consuming procedures, making analysis more robust.

In the future, the application of other correlative approaches – suchas combining cryo-ET with atomic force microscopy – would alsoenable correlations between physical changes in the cell, forcemeasurements and structural information. It is expected that suchhybrid methods will lead to platforms that can provide deeper insightinto cellular processes. Complementary information from a varietyof techniques will thus be combined to reconstruct meaningful cellulardensity maps (Robinson et al., 2007). With advanced instrumentation,such as advanced charge-coupled device (CCD)-camera detectors anddual-axis tilting devices, the prospects are good that higher and moreisotropic resolutions of 2-3 nm can be attained. Therefore, we foreseethat cell biology will increasingly rely on high-resolution 3D imagingtechniques, in conjunction with other approaches.

This work was supported by a grant from the German-IsraeliCooperation Project (DIP) (H.2.2), by the Israel Science Foundation(grant 794/06) and by the German-Israel Foundation, to O.M. We thankKay Grünewald and Ulrike Maurer for providing Fig. 3.

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