Macromolecular Architecture inEukaryotic Cells Visualized by

Cryoelectron TomographyOhad Medalia, Igor Weber, Achilleas S. Frangakis,

Daniela Nicastro, Gunther Gerisch, Wolfgang Baumeister*

Electron tomography of vitrified cells is a noninvasive three-dimensional im-aging technique that opens up new vistas for exploring the supramolecularorganization of the cytoplasm.We applied this technique toDictyostelium cells,focusing on the actin cytoskeleton. In actin networks reconstructed withoutprior removal of membranes or extraction of soluble proteins, the cross-linkingof individualmicrofilaments, their branching angles, andmembrane attachmentsites can be analyzed. At a resolution of 5 to 6 nanometers, single macromol-ecules with distinct shapes, such as the 26S proteasome, can be identified inan unperturbed cellular environment.

Visualizing the three-dimensional (3D) orga-nization of eukaryotic cells, with their dy-namic organelles, cytoskeletal structures, andmolecular machines in an unperturbed con-text, requires a noninvasive, high-resolutionimaging technique combined with a methodof arresting cells in their momentary state offunction. Electron tomography (ET) has aunique potential for this purpose (1, 2). Con-ventional electron microscopic images areessentially 2D projections of the object: Fea-tures are superimposed upon one another inthe direction of the electron beam. Tomo-graphic techniques, in contrast, acquireprojections of the object as viewed from dif-ferent directions and then merge them com-putationally into a 3D reconstruction, the to-mogram. In ET this is done by tilting thespecimen holder incrementally around anaxis perpendicular to the electron beam.

Although ET is not a new concept [see,e.g. (3)], formidable experimental problemsstood in the way of practical applications formore than three decades. Basically, there aretwo conflicting requirements, which are dif-ficult to reconcile. On the one hand, resolu-tion and quality of a tomogram are directlydependent on spacing of the projections andon the angular range covered. Therefore, datamust be collected over a tilt range as wide aspossible, with increments as small as possi-ble. On the other hand, the exposure to theelectron beam must be minimized to preventradiation damage (4, 5). The development ofelaborate automated data-acquisition proce-dures enabled us to reduce beam exposuredramatically (6, 7) and to take advantage ofthe principle of dose fractionation (8, 9).With these improvements, ET could be ap-

plied to radiation-sensitive specimens, suchas biological materials embedded in amor-phous ice (10–12). Vitrification by rapidfreezing ensured close-to-life preservation ofcellular structures, avoiding the risk of arti-facts associated with chemical fixation, stain-ing, or dehydration.

Specimen thickness is a major limitationin ET. When it exceeds the mean free path ofelectrons, multiple inelastic scattering beginsto degrade the quality of images substantial-ly, despite the use of higher voltages (300 to400 keV) and energy filters to temper theseeffects (13). In consequence, samples thickerthan 1 �m cannot be studied in toto, but mustbe (cryo)sectioned before tomographic anal-ysis. Many prokaryotic cells are smallenough to be examined by cryo-ET withoutthe need of sectioning, and some eukaryoticcells can be grown flat enough to apply cryo-ET at least locally in a noninvasive manner.Here we apply this method to the highlymotile cells of Dictyostelium discoideum. Aprerequisite for studying intact eukaryoticcells by cryo-ET is their ability to spread onelectron microscope grids (see supporting on-line methods) (14). Dictyostelium cells mov-ing on a copper grid coated by a plain carbonfilm are shown in Movie S1. These cellsassume their normal shape, migrate, and evendivide as on a glass surface. The grids withthe cells enclosed in a thin aqueous layer areplunge-frozen to achieve vitrification.

Zooming into intact vitrified cells. Thecellular regions that we examined by cryo-ETvaried between 300 and 600 nm in thickness.Two-dimensional projection images of suchregions, obtained by conventional cryoelec-tron microscopy (cryo-EM), are hard to inter-pret because of the superposition problemnoted above. Only membranes and other fea-tures that are continuous through most of thevolume are clearly visible (Fig. 1A). Becausein computing a tomogram the information

from many statistically noisy projections iscombined, the signal-to-noise ratio is signif-icantly improved as compared with individu-al projections. Nevertheless, there is substan-tial residual noise because data sets need tobe recorded under the constraint of a subcriti-cal cumulative electron dose. But even with-out further “denoising” or averaging of seg-mented features, some structural elements areclearly visible in a tomogram displayed in theform of 60-nm-thick x-y slices along the zaxis, in particular microfilaments and a pop-ulation of high-density particles roughly 30nm in size (Fig. 1, B to D). As an example ofintracellular organelles that can be studied bycryo-ET in their native environment, we fo-cus in Fig. 1, E and F, on the rough endo-plasmic reticulum (ER). Dictyostelium cellshave a profusely developed ER, which ex-tends from the nucleus up to the inner face ofthe cortical actin layer (15). Figure 1E showsa series of x-y slices through a portion of therough ER located within the peripheral regionof a cell (framed white in Fig. 1A). Themembrane is decorated with an array of glob-ular complexes, which resemble 80S ribo-somes in size (�27 nm) and shape (Fig. 1F).Their high contrast is consistent with a parti-cle containing mostly RNA. We assume thatthe high-density particles in the cytoplasm,often nestled in the actin cytoskeleton, arealso ribosomes. The interior of the ER ishighly crowded, making it difficult to identi-fy distinct features in this compartment.

Visualizing a macromolecular complexin situ: the 26S proteasome. The less-crowded environment in the cytoplasm al-lows one to search the tomograms interac-tively for complexes recognizable throughtheir structural signature, i.e., their size andshape. A particularly striking example of alarge protein complex residing in the cyto-plasm is the 26S proteasome, the most down-stream element of the ubiquitin pathway ofprotein degradation (16). This molecular ma-chine of 2.5 MD has a characteristic elongat-ed shape (45 nm in length) reminiscent of a“double dragon head” (17–19). When sub-tomograms containing this motif were ex-tracted (Fig. 2A) and the slices containingthis structure were summed, the resultingprojection image (Fig. 2B) correspondedclosely to an average derived from purifiedXenopus 26S proteasomes (Fig. 2C). Thereare a few subtle differences in the regionconnecting the base and the lid of the 19Sregulatory subcomplex, but this is known tobe a weak and flexible linkage (20). Despitethe relatively low resolution of the tomogram(�5 to 6 nm), the similarity with the 26Scomplex from Xenopus is striking and theidentification is unambiguous.

Capturing unperturbed actin networkswithout removal of membranes. Cell mo-tility, endocytosis, and cytokinesis rely on the

Max Planck Institute for Biochemistry, D-82152 Mar-tinsried, Germany.

*To whom correspondence should be addressed. E-mail: baumeist@biochem.mpg.de

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spatially and temporally controlled assemblyand disassembly of actin filaments. Light andelectron microscopy have provided importantinsights into actin-filament organization and itschanges in response to external signals. A va-riety of electron microscopic techniques havebeen used to study cytoskeletons; some arebased on fixed and sectioned material, and oth-ers use detergent-extracted or mechanically un-roofed cells (21–24). The cytoskeletons can bevisualized by negative staining or by freeze-drying in conjunction with platinum replication(21). As recently shown, artifacts associatedwith freeze-drying can be avoided by directobservation of frozen cytoskeletons from deter-gent-extracted cells (24).

In the cell cortex, actin filaments areinterconnected in various modes, forming net-works anchored to the inner face of the plasmamembrane. Polymerization and depolymeriza-tion, cross-linking, and branching of actin fila-ments are regulated by different proteins thatact together in determining cell shape and mo-tility (25–27). Actin organization in Dictyoste-lium cells is similar to that in neutrophils andother fast-moving animal cells, and many of theproteins identified in mammalian cells havehomologs in Dictyostelium (28–32). These in-clude the Arp2/3 complex and a filamin-likeF-actin cross-linker (33, 34).

In highly motile cells, like those of Dictyo-stelium, the actin filament system is reorganized

within seconds (35). The structure of these net-works in any part of the cell is transient andreflects their momentary role in cell motility,endocytosis, or mitotic cell division. Because ofthe dynamic nature of the actin cytoskeleton, itis essential to combine ET with cryotechniques,which allow one to vitrify the sample, avoidingperturbation of the delicate balance of regula-tory proteins in this system.

Here, we report that cryo-ET allows visual-ization of the actin filament network withoutany pretreatment and in its native associationwith cellular membranes. Thus, two major risks

are avoided. First, actin-binding proteins mayrapidly dissociate from, or artificially bind to,the actin network during the treatment of cellswith detergent. In both cases, the connectionsbetween actin filaments and thus the structureof the network will be altered. Second, distor-tions of the network caused by its detachmentfrom membranes are prevented and, by thesame token, the sites of membrane attachmentcan be studied.

The cortical region of the cell shown inFig. 1, A to D, is dominated by the network ofactin filaments, with ribosomes and other

Fig. 1. Peripheral re-gion of a vitrified Dic-tyostelium cell ana-lyzed by cryo-ET. (A)Conventional trans-mission electron mi-crograph (2D projec-tion) of the regionused for recording atilt series. Thicknessof the cell in this re-gion varied from 200to 350 nm. (B to D)Series of successivetomographic x-y slic-es of 60-nm thicknessfrom bottom to topof the substrate-at-tached region of thecell. In (B) and (C),portions of the plas-ma membrane thatare perpendicular tothe x-y plane can berecognized. In (B), thecortical actin networkis viewed. Detailedimages of the molec-ular complexes andvesicular structuresare shown in (E) andin Fig. 3; white and

black frames in (A) refer to the areas represented in these figures. (E) Gallery of x-y slices through aribosome-decorated portion of the ER. Original tomograms through the volume are displayed without anydenoising. (F) Surface rendering of the volume represented in (E). The image was manually segmented intomembrane (blue) and ribosomes (green). For clarity, the densely packed structures in the lumen have beenomitted.

Fig. 2. Structure of a 26S proteasome as visualized in its cytoplasmic context. (A) x-y slices throughthe volume of the proteasome. (B) Density profile of a 2D projection of the volume represented in(A). (C) Reference image of the averaged 2D structure of a purified and negatively stained 26Sproteasome from Xenopus oocytes (44).

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macromolecular complexes in between. Thisbecomes particularly clear after surface ren-dering of the tomogram (Fig. 3A). Figure 3Bshows an enlargement of a subvolume crowd-ed with actin filaments and displays the net-work in stereo. From this network and fromsimilar ones, we have extracted X-, Y-, and

T-shaped junctions between the actin fila-ments; 12 of them are displayed in the galleryshown in Fig. 4. Among these examples,branching angles vary between 40° and 90°.Some branches show the arm linked to anactin filament out of plane, as illustrated inFig. 4A (top). A special feature of some

branches is visible in Fig. 4B: The side armsare sharply bent close to their site of attach-ment. This is presumably caused by an actin-binding protein that links one end of a fila-ment to the side of another one. The putativelinker protein sticks out of a filament at awide angle. Cross-linking of actin filamentsoccurs often at a 90° angle (Fig. 4C).

Figure 5A shows a series of x-y slicesfrom a tomogram of another cortical regionwith actin filaments predominantly arrangedin parallel; for clarity, the sections were de-noised. Although this tomogram is of inferiorquality owing to the greater local thickness ofthe cell (�600 nm), the arrangement of actinfilaments is clearly visible. Individual fila-ments can be traced over a length of 400 to500 nm, which is twice the reported averagelength (36). Such parallel arrangements aretypically generated and stabilized by bun-dling proteins forming serial bridges alongthe individual filaments. Indeed, at many lo-cations such bridges are discernible in Fig.5B; two of them are displayed in Fig. 5C.

Because in cryo-ET the integrity of the cel-lular membranes is maintained, we were able todepict binding sites of actin filaments on theplasma membrane. Figure 6A shows a corticalactin-filament network locally connected to theplasma membrane, and in Fig. 6, B and C,surface-rendered actin filaments are presentedthat appear to be kinked close to the membranesurface. These images indicate that actin fila-ments are capable of end binding to the plasmamembrane through a connecting element thatforms the kink. This element might be an adap-tor protein or the cytoplasmic domain of anactin-binding transmembrane protein.

In summary, cryo-ET of peripheral re-gions in Dictyostelium L cells depicted twodifferent types of actin-filament arrays: al-most isotropic networks (Fig. 3) and parallelarrangements of actin filaments that are tiedtogether by bundling proteins (Fig. 5). Withinthe actin networks, we have found a broaddistribution of branching angles and differenttypes of cross-linkages (Fig. 4). These obser-vations are consistent with the presence ineukaryotic cells of a large variety of actin-cross-linking and actin-branching proteinsthat connect actin filaments to each other atangles varying from 0° (parallel bundling) to180° (antiparallel bundling). Groups of con-served branching and cross-linking proteinsinclude the Arp2/3 complex, which is char-acterized by a 70° angle of branching (37,38), and the filamin family (with filamin A asa prototype), which has a preference for or-thogonal crosslinkage of actin filaments (23,34). An advantage of cryo-ET is that branch-ing angles can be determined precisely fromthe 3D maps. The quantitative analysis ofnetwork geometries will benefit from auto-mated procedures, allowing the evaluation oflarge tomographic data sets.

Fig. 3. Visualization of actin network, membranes, and cytoplasmic macromolecular complexes. (A)A volume of 815 nm by 870 nm by 97 nm, corresponding to the area of Fig. 1A framed in black,was subjected to surface rendering. Colors were subjectively attributed to linear elements to markthe actin filaments (reddish); other macromolecular complexes, mostly ribosomes (green); andmembranes (blue). (B) Stereo image of an actin layer taken from the upper left portion of (A)reveals a crowded network of branched and crosslinked filaments.

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Prospects of cryo-ET applied to cellbiology. Cryo-ET has the potential to bridgethe gap between cellular and molecular struc-tural studies (39). By combining the power of3D imaging with the close-to-life preservation

achieved by vitrification, we have visualizedmacromolecular complexes and supramolecu-lar structures inside intact eukaryotic cells. Thetomograms of Dictyostelium cells contain animposing amount of information, even at the

present resolution of 5 to 6 nm. Essentially,they are 3D images of the cell’s entire pro-teome, and they represent the network of mac-romolecular interactions that provides the basisfor coordinated cellular activities. Exploitationof this information is confronted with a numberof problems. Despite advanced image-acquisi-tion procedures and the application of denoisingtechniques, cryoelectron tomograms still sufferfrom substantial residual noise and distortionsbecause of missing data. Moreover, the cyto-plasm and organelles are crowded with macro-molecules, which often makes it impossible toperform segmentation and feature extraction onthe basis of visual inspection of the tomograms,except for large and continuous structures suchas membranes and filaments.

The less-crowded cytoplasm of eukaryoticrelative to prokaryotic cells (40) made it pos-sible to visualize 26S proteasomes withintheir cellular environment. For the detectionof smaller complexes and less idiosyncraticstructures, it will be necessary to rely onpattern recognition techniques, such as tem-plate matching (41). To make the search forspecific complexes objective and reproduc-ible, it should be automated and machinebased, not requiring manual intervention.Such methods are under development (42).One current limitation of cryo-ET is the lackof methods that allow the electron microscop-ic data to be correlated with light microscopicimages. Thus, we cannot yet relate the differ-ent patterns of actin arrangement shown inFigs. 3 and 5 to specific activities in the cellcortex. Future developments will aim at com-bining video recording and fluorescence mi-croscopy of green fluorescent protein–taggedproteins with cryo-ET to characterize the dy-namic state and protein inventory of thestructures under investigation.

In applying cryo-ET to dynamic cytoskel-etal arrangements, a major goal will be to assignthe types of branches and cross-linkages tospecific linker proteins. This might be accom-plished by different routes. The structural anal-ysis of mutants that miss specific actin–cross-linking proteins or that lack combinations ofthese proteins will help to identify the individ-ual contributions of major players in the dy-namic organization of actin networks. Anotherpossibility is the combination of cryo-ET withthe labeling of specific proteins by microin-jected gold-conjugated antibodies. Finally, athigher resolution it will be possible to detectand identify cross-linking proteins by a tem-plate-matching approach, as currently exploredfor larger cytoplasmic protein complexes (42).Because prospects for a significant improve-ment in resolution to 1.5 to 2.0 nm are good(43), it will ultimately be possible to generatepseudoatomic maps of large supramolecular as-semblies or even whole organelles by dockinghigh-resolution structures of molecular compo-nents into cellular tomograms.

Fig. 4. Branches andcrosslinks of actin fila-ments. These exam-ples were chosen fromthe cortical regions ofcells as in Fig. 3. Thebranching angles asdisplayed in 2D wereextracted and maxi-mized in 3D. (A) Com-pilation of branchingswith different angles.(B) Examples charac-terized by a kink thatchanges the angleclose to the branchingsite. (C) Cross-linkag-es between filamentsin addition to branches.

Fig. 5. Bundles of actin filamentsin the cell cortex. (A) Two de-noised x-y slices of 10-nm thick-ness through the cortical regionof a cell, sampled at 490 to 510nm above the substrate. (B) Avolume-rendered portion of theregion depicted in (A) showingbridges between filaments andlateral connections of filamentsto the membrane. (C) Represen-tative bridges between actin fil-aments extracted from (B).

Fig. 6. End binding of actin fila-ments to the plasma membrane.(A) Cortical region of 500 nm by300 nm by 20 nm showing mem-brane-associated actin filamentspresented as in Fig. 3. The arrowpoints to the site of actin-mem-brane interaction shown in (B) athigher magnification. (B and C)High-magnification image of cor-tical regions, depicting kinklikestructures close to filament-mem-brane interaction sites. Kink angleswere not maximized, and themembrane in (C) was not perpen-dicular to the actin filament. (B)and (C) were taken from differentcortical regions.

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14153 (2002).43. J. M. Plitzko et al., Trends Biotechnol. 20, S40 (2002).44. J. Walz et al., J. Struct. Biol. 121, 19 (1998).45. O.M. thanks the European Commission for a Marie

Curie fellowship. I.W. was supported by a grant fromthe Deutsche Forschungsgemeinschaft (SFB 266 toG.G.). We thank J. Kohler for the movie and A. C.Steven for critical reading of the manuscript.

Supporting Online Materialwww.sciencemag.org/cgi/content/full/298/5596/1209/DC1Materials and MethodsMovie S1

16 July 2002; accepted 9 September 2002

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Superconductivity in DenseLithium

Viktor V. Struzhkin,1* Mikhail I. Eremets,2 Wei Gan,1,3

Ho-kwang Mao,1 Russell J. Hemley1

Superconductivity in compressed lithium isobservedbymagnetic susceptibility andelectrical resistivity measurements. A superconducting critical temperature (Tc) isfound ranging from9 to 16 kelvin at 23 to 80 gigapascals. The pressure dependenceof Tc suggests multiple phase transitions, consistent with theoretical predictionsand reported x-ray diffraction results. The observed values for Tc are much lowerthan those theoretically predicted, indicating that more sophisticated theo-retical treatments similar to those proposed for metallic hydrogen may berequired to understand superconductivity in dense phases of lithium.

Lithium is considered the simplest metal: it isthe lightest of the alkali metals, and undernormal pressure-temperature conditions itsproperties are well described within a nearlyfree electron model. The two inner core elec-trons effectively shield the nucleus, inhibitingelectron-ion attraction and leaving the outerelectrons to move freely within the ion lattice.Because the valence electron is so delocal-ized, lithium is a metal with high conductiv-ity and it assumes a highly symmetric body-centered cubic (bcc) structure. Recent studiesshow that under pressure, however, this sim-

plicity changes radically. Theoretical predic-tions (1) suggest that lithium may undergoseveral structural transitions, possibly leadingto a “paired-atom” phase with low symmetryand near-insulating properties. Though thisprediction is the antithesis of the intuitiveexpectation that pressure favors high-symme-try crystal structures with metallic properties,recent x-ray diffraction studies reveal a struc-ture similar to the predicted paired structure(2). Near 39 GPa, lithium transforms to acubic polymorph with 16 atoms per unit cell(cI16), a recently discovered structure uniqueto lithium (2). Additionally, a minimum inthe electronic density of states close to theFermi energy suggests near-insulator behav-ior in the paired structure.

The ambient-pressure phase of Li at lowtemperature is not bcc but rather a closedpacked rhombohedral 9R structure (3). Underambient pressure, no sign of superconductiv-

ity in lithium has been detected down to 4mK (4). However, it is predicted that lithi-um’s structural changes under pressure mayhave large effects on possible superconduc-tivity in the material (1, 5). Explicit calcula-tions (6) suggest that Tc may reach a maxi-mum of 60 to 80 K in the cI16 phase. Anearly resistivity experiment had indicated apossible superconducting transition in lithium(7) at 7 K and higher pressures; however, theexperiment was inconclusive due to the lackof magnetic signature of the transition. Sim-ilar experiments, performed recently with adiamond anvil cell to higher pressures, con-firmed a resistance drop and established adependence of the drop on external magneticfield but still lacked the magnetic susceptibil-ity measurement necessary for proving super-conductivity (8). Because of the difficultiesin studying the material under pressure [e.g.,sample containment and reactivity (2, 9)], itis essential to apply a variety of probes of thecompressed sample. For superconductivity,this includes the combination of magnetic(10) and electrical (11) techniques.

Magnetic susceptibility measurements wereperformed in two experiments (12). The temper-ature scan at 28 GPa in the second run is shownin Fig. 1. We used a Nb sample placed in thecompensating coil at ambient pressure as a ref-erence in this experiment. The signal from Nb issuperimposed in phase with the background; thesignal from Li is opposite in phase from that ofthe background. The compensating and signalcoils are connected in opposition; thus, the sig-nal from Li has actually the same phase as thatfrom Nb and, therefore, indubitably correspondsto the superconducting transition. We observed

1Geophysical Laboratory, Carnegie Institution ofWashington, 5251 Broad Branch Road, N.W., Wash-ington, DC 20015, USA. 2Max Planck Institut fur Che-mie, Postfach 3060, 55020 Mainz, Germany. 3ThomasS. Wootton High School, Rockville, MD 20850, USA.

*To whom correspondence should be addressed. E-mail: struzhkin@gl.ciw.edu

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