THE JOURNAL OF BIOLOGICAL CHEMISTRK (0 1991 by The American Society for Biochemistry and Molecular Biology, Inc.

Vol. 266, No. 36, Issue of December 25, pp. 24650-24656.1991 Printed in U. S. A.

Cryoelectron Microscopy of Mammalian Pyruvate Dehydrogenase Complex*

(Received for publication, April 1, 1991)

Terence WagenknechtSSlI, Robert GrassucciS, Gary A. Radkell, and Thomas E. Rochell From the f Wadsworth Center for Laboratories and Research, New York State Department of Health and the §School of Public Health Sciences, Department of Biomedical Sciences, State University of New York, Albany, New York 12201 -0509 and the (1 Department of Biochemistry, Kansas State University, Manhattan, Kansas 66506

Cryoelectron microscopy has been performed on fro- zen-hydrated pyruvate dehydrogenase complexes from bovine heart and kidney and on various subcomplexes consisting of the dihydrolipoyl transacetylase-based (E2) core and substoichiometric levels of the other two major components, pyruvate dehydrogenase (El) and dihydrolipoyl dehydrogenase (E3). The diameter of frozen-hydrated pyruvate dehydrogenase complex (PDC) is 50 nm, which is significantly larger than previously reported values. On the basis of micro- graphs of the subcomplexes, it is concluded that the El and E3 are attached to the E2-core complex by ex- tended (4-6 nm maximally) flexible tethers. PDC con- structed in this manner would probably collapse and appear smaller than its native size when dehydrated, as was the case in previous electron microscopy studies. The tether linking El to the core involves the hinge sequence located between the El-binding and catalytic domains in the primary sequence of E2, whereas the tether linking E3 is probably derived from a similar hinge-type sequence in component X. Tilting of the E2- based cores and comparison with model structures con- firmed that their overall shape is that of a pentagonal dodecahedron. The -6 copies of protein X present in PDC do not appear to be clustered in one or two regions of the complex and are not likely to be symmetrically distributed.

Pyruvate dehydrogenase complexes (PDCs)’ are among the largest enzymes known, having molecular masses approaching 10 million daltons in mammals (for a recent review see Patel and Roche, 1990). One of the component enzymes in PDC, dihydrolipoyl transacetylase (E2), forms a multisubunit core assembly to which are bound noncovalently multiple copies of the other component enzymes. In eukaryotes, the E2 core consists of about 60 polypeptide chains and has the overall shape of a pentagonal dodecahedron (Oliver and Reed, 1982). The other major components are pyruvate dehydrogenase

* This work was supported by National Institutes of Health Grants GM38161 (to T. W.) and DK18320 (to T. E. R.) and by Contribution 91-439-5 from the Kansas State Agriculture Experiment Station (to T. E. R.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

ll To whom correspondence should be addressed Wadsworth Cen- ter for Laboratories and Research, New York State Department of Health, P. 0. Box 509, Albany, NY 12201-0509.

The abbreviations used are: PDC, pyruvate dehydrogenase com- plex; El, pyruvate dehydrogenase; E2, dihydrolipoyl transacetylase; E3, dihydrolipoyl dehydrogenase; E2. X, complex of E2 and compo- nent X; E2. X. K, complex of E2, component X, and kinase.

(El) , an a& tetramer present in 20-30 copies, and dihydro- lipoyl dehydrogenase (E3), a homodimer present in about six copies. Additionally, there are one to three copies of a regu- latory El-specific kinase and a few copies of a complementary, loosely bound phosphatase. Finally, about six copies of a poorly understood polypeptide, designated component X, which is involved in binding E3, are tightly associated to the E2 core (Gopalakrishnan et al., 1989; Powers-Greenwood et al., 1989). E 3 binding by component X has also been estab- lished for yeast PDC (Lawson et al., 1991).

Electron microscopy has played a major role in elucidating the three-dimensional architecture of PDCs from prokaryotic and eukaryotic organisms (for a review, see Oliver and Reed, 1982). However, electron microscopy studies of mammalian PDCs have been hampered by the tendency of the complexes to partially dissociate into subunits when prepared by conven- tional techniques such as negative staining. Thus, in most previous studies, chemical cross-linking (Junger and Rei- nauer, 1972) or freeze-etching techniques (Junger and Bach- mann, 1977) were used to minimize dissociation. I t is not known to what degree native structure has been preserved in these studies. PDCs are probably not rigid structures (Perham et al., 1987; Perham and Packman, 1989; Wagenknecht et al., 1990) and so they might be particularly sensitive to distortions caused by intramolecular cross-linking, dehydration, or ad- sorption to surfaces. The E2 polypeptide has a segmented structure (Fig. 1) in which specialized, folded domains are linked to one another by hinge sequences, typically 20-30 residues in length, that are rich in alanine, proline, and charged amino acids (Thekkumkara et al., 1988; Coppel et al., 1988). The hinge sequences are conformationally mobile and are thought to have extended structures in bacterial PDCs (for reviews, see Perham et al. (1987) and Perham and Pack- man (1990)). The El-binding domains are linked to the E2 core by typical hinge segments, and thus the El tetramers could be particularly sensitive to distortions of the type de- scribed.

Mammalian component X is thought to have a similar structure to that shown for E2 in Fig. 1 (Patel and Roche, 1990), except that it has only one lipoyl domain (Rahmatullah et al., 1989a, 1989b). Proteolytic cleavage suggests a three- domain structure with the inner domain binding to the inner domain of E2 (Rahmatullah et al., 1989b). The presence of an intervening E3-binding domain and its role in binding E3 has been demonstrated for yeast component X (Lawson et al., 1991). The degree of relatedness of the yeast and mammalian component X has not been established.

Cryoelectron microscopy of frozen-hydrated solutions of macromolecules is a relatively new method that is free from most of the artifacts associated with conventional electron microscopy techniques (Adrian et al., 1984; Dubochet et al.,

24650

Cryoelectron Microscopy of Pyruvate Dehydrogenase Complex 24651

1987). The method is capable of preserving macromolecular structure to near-atomic resolution (Taylor and Glaeser, 1976; Chiu, 1986; Jaffe and Glaeser, 1987), although for noncry- stalline specimens it is usually not possible to retrieve the high resolution information owing to the low contrast of the images and radiation-induced damage. Recently, this tech- nique was applied to PDC and the related enzyme complex, 2-oxoglutarate dehydrogenase, from Escherichia coli. Evi- dence was found that these enzymes are indeed fragile struc- tures, which appear to be well preserved in the frozen-hy- drated state (Wagenknecht et al., 1990). Therefore, we have investigated the structure of mammalian PDC by cryoelectron microscopy. Portions of this work were described in abstract form (Grassucci et al., 1990).

EXPERIMENTAL PROCEDURES

Materials-Bovine heart and kidney PDC and their components were isolated by procedures described previously (Rahmatullah and Roche, 1987; Roche and Cate, 1977; Linn et al., 1972; Stepp et al., 1983). The loosely bound phosphatase is lost during these procedures. Reconstituted subcomplexes consisting of the E2.X (or E2.X.K) and E l or E3 were prepared by incubation of the components at room temperature for 30 min. The subcomplexes were stored at 4 “C prior to preparation for cryoelectron microscopy. A buffer consisting of 0.05 M potassium phosphate (pH 7.0), 1.0 mM MgCl,, 0.1 mM EDTA, 0.005% sodium azide was used unless stated otherwise.

Cryoelectron Microscopy-Specimens were applied at 0.05-0.2 mg/ ml to holey carbon films supported by 300-mesh copper grids, blotted, and immediately plunged into supercooled liquid ethane (Lepault et al., 1983; Milligan et al., 1984; Wagenknecht et al., 1990). A modifi- cation of this procedure was used for PDC from bovine kidney, which showed significant dissociation of subunits (probably E l ) from the E2 core when prepared for microscopy as outlined above. Kidney PDC was maintained at a concentration of 0.75 mgjml at room temperature immediately prior to application to a grid. A 1-p1 aliquot was removed and injected into a 4-pl droplet of buffer at 4 “C, which was applied in advance to a grid already clamped into the plunging apparatus. Within a few seconds after injection of the PDC, the grid was blotted and frozen in the usual way. Micrographs prepared by this modified procedure showed a marked reduction in disassembly of PDC.

A Philips EM420T transmission electron microscope equipped with a low dose kit, goniometer stage, a Gatan model 651N auxiliary anticontaminator, and a Gatan model 626 cryotransfer system was used. Cryospecimens were maintained at -170 to -180 “C, and the net electron dose to record a micrograph was less than 2000 electrons/ nm’. Most micrographs were recorded at a nominal magnification of 50,OOOX and with the objective lens underfocused by 1.5-2 pm in order to generate phase contrast. The magnification of the microscope was determined with tobacco mosaic virus as a calibration standard; the axial spacing of the third layer line, 1/2.3 nm”, in the computed Fourier transform of selected virions was used to determine the pixel size of the digitized images (see below).

Analysis of Images-Micrographs were digitized using a Perkin- Elmer PDS microdensitometer operated with a 25-rm square scan- ning aperture corresponding to 0.5 nm on the specimen. The micro- graphs were displayed on the video monitor of a VAX3100 work station, and individual enzyme complexes were selected interactively and stored as image files. All image processing was done using the SPIDER program package implemented on a VAX6210 computer

E2:

FIG. 1. Schematic representation of the domain structure of mammalian E2. The two lipoyl domains (LI and L2), the El- binding domains ( B ) , and the inner domain ( I ) are arranged sequen- tially along the primary sequence of E2, with the amino terminus at the left. The hinge peptides that connect the domains are represented by the wavy lines. Lipoic acid residues are shown projecting from the upper surfaces of the lipoyl domains. Adapted from Pate1 and Roche (1990) and based on sequences determined by Coppel et al., 1988 and Thekkumkara et al., 1988. Component X has a similar domain struc- ture but contains only one lipoyl domain.

(Frank et at., 1981). To improve the contrast of the image files, they were low pass filtered to a limiting resolution of l/2.9 nm” and density thresholded such that the minimum and maximum densities were within two standard deviations of the mean density of the unprocessed image.

Images of E2.X cores in orientations displaying 2-, 3-, or 5-fold symmetry (as verified by tilting experiments; see Fig. 4) were aligned and averaged (Frank et al., 1978, 1985; Wagenknecht et al., 1988).

Measurements of positions of bound El and E3 molecules were made interactively from enlarged, contrast-enhanced images of sub- complexes. A cursor was manually positioned at the center of each bound unit and at the center of the E2. X . K core. The coordinates were stored in a SPIDER system “document” file and recalled later for determination of the distances and associated statistical quan- tities. For the E2.X.K-E3 subcomplexes, the center coordinates of the E2 .X. K component were refined by cross-correlation with a rotationally averaged E2. X. K image. An accuracy better than one pixel (0.5 nm) was achieved by this procedure, and we found that the coordinates of most of the particles’ centers as determined interac- tively agreed with those determined by the cross-correlation proce- dure to within two pixels.

Images of E2 .X and E2. X. K cores were compared with projections of a computer-generated, icosahedrally symmetric three-dimensional model that was obtained as follows. Spheres were placed at coordi- nates corresponding to the vertices of a pentagonal dodecahedron, and these were connected by cylinders, which formed the edges. The spheres and cylinders were scaled such that the total volume of the model corresponded to the volume contributed by the inner domains of the E2 polypeptide (assuming 1.3 X nm3/Da). Although this model cannot be expected to account for the fine structural details of the images, it was nevertheless useful for interpreting the tilting experiments in which the various views of the E2.X core were interconverted (Fig. 4).

RESULTS

Morphology of Native PDC-A portion of an electron mi- crograph of frozen-hydrated PDC from bovine heart is shown in Fig. 2.4. The molecules are not adsorbed to a support film (e.g. carbon) in this micrograph, but rather are suspended in a thin film of frozen phosphate buffer (pH 7). The background ice surrounding the PDC particles is largely free of smaller particles, indicating that dissociation of subunits from the complex has not occurred. The E2 core is often visible at the center of the images, and in some, its characteristic 5-, 3-, and 2-fold rotational symmetries are recognizable (cf. Fig. 3). In the contrast-enhanced images of selected PDC molecules (Fig. 2 B ) , individual morphological units, most of which cor- respond to El subunits, are visible surrounding the E2 core, but they do not appear to be distributed in a symmetric pattern. Sometimes the peripherally located morphological units appear to be separated from the surface of the E2 by a gap of several nanometers.

The average diameter of frozen-hydrated PDC is about 50 -1- 2 nm. On several of the images in Fig. 2B, a circle of radius corresponding to 25 nm has been superimposed, from which it is apparent that material occasionally extends even beyond this radius. Previously reported measurements of the diameter of mammalian PDC, which were obtained using negatively stained or freeze-dried/metal-coated specimens, were in the range of 40-45 nm (Oliver and Reed, 1968; Junger and Rei- nauer, 1972; Junger and Bachmann, 1977). It is likely that the dehydration associated with these techniques resulted in partial contraction of the PDC and, hence, a smaller diameter than we have obtained for frozen-hydrated specimens (see “Discussion”). Even lower values for the diameter have been reported for negatively stained PDC in which chemical cross- linking was not employed to reduce partial dissociation to subunits (Ishikawa et al., 1966; Hayakawa et al., 1969).

Selected images of PDC from bovine kidney are shown in Fig. 2C. The only apparent difference from heart PDC is that the E2 core is often more clearly resolved in the kidney PDC,

24652 Cryoelectron Microscopy of Pyruvate Dehydrogenase Complex

FIG. 2. Cryoelectron microscopy of bovine PDC. A, a field of heart PDC. A few phospholipid vesicles, which were added to some preparations (Wagen- knecht et al., 1990) were also visible (ar- rowhead). Scale bar, 100 nm. B and c, selected contrast-enhanced images of heart and kidney PDC, respectively. A circle of a diameter corresponding to 50 nm is superimposed at the centers of the right-most three images in B and C.

A

Bl C

probably because of the lesser amount of El present than in heart PDC (20 uersus 30 tetramers (Barrera et al., 1972)). The kidney PDC also showed a tendency to partially disso- ciate, which was not observed for heart PDC (see “Experi- mental Procedures”).

EBCore Complex-Core complexes from PDC, referred to as E2. X. K cores because they contain component X and the El-specific kinase (K) in addition to E2, were obtained by resolution of PDC and examined by cryoelectron microscopy (Fig. 3A). It has been possible to extract component X from the core complex only by using conditions that disrupt the quaternary structure of the complex, indicating that compo- nent X might itself be an integral constituent of the core (Powers-Greenwood et al., 1989). The kinase can be quanti- tatively removed to yield E2 * X cores, which appear essentially identical with the E2. X. K cores (Fig. 3B). Treatment of E2. X cores with trypsin, which removes the outer domains (see Fig. 1) from both E2 and X (Rahmatullah et al., 1989b), results in micrographs that appear qualitatively similar to the E2 X. K and E2 X images shown in Fig. 3 (data not shown). Thus, it is likely that only the structure formed by the inner domains of E2. X is being resolved reliably in the micrographs, and we hereafter refer to this structure as the E2 - X inner core.

Individual images of the E2 .X (or E2 .X. K) cores show three principal views, which appear to possess either 5-, 3-, or 2-fold symmetry (Fig. 3, B-D). Usually, 5-fold symmetric images occur more frequently than 2- or 3-fold ones. This behavior may result from interactions of the cores with the air-buffer interface present prior to freezing. Similar types of images have been observed previously with negatively stained specimens (Reed and Oliver, 1968; Junger and Reinauer, 1972), and have been interpreted in terms of an icosahedrally symmetric model for the core in which most of the mass from the inner domains is positioned near the 3-fold vertices so as to form a pentagonal dodecahedron.

We have tested the hypothesized icosahedral symmetry of the E2 inner core by determining whether the various sym- metric views interconvert as predicted by the model when the

cores are tilted in the electron microscope. Fig. 4 shows pairs of images consisting of a 5-fold untilted view and the corre- sponding image obtained following a tilt of 35” about a vertical axis. This amount of tilting should convert the 5-fold views to views that are approximately along 3- or %fold symmetry axes, depending on the azimuthal orientation of the &fold view with respect to the tilt axis. The experimental 5-fold views do indeed convert to views having 2- or 3-fold symmetry or views intermediate to the 2- and 3-fold symmetric ones (Fig. 4, A-C, respectively). Also shown with each pair of experimentally determined images is a projection of a com- puter-generated icosahedrally symmetric model of the E2 e X inner core (see Fig. 3G). The model projection was obtained by rotationally aligning the model in the &fold orientation with the 5-fold symmetric experimental image and then tilting it about a vertical axis (corresponding to the experimental tilt axis) and computing its projection image. Clearly, the exper- imental images agree well with those predicted by the icosa- hedrally symmetric model. Despite this evidence for icosahed- ral symmetry, it is still possible that the true symmetry of the E2 core is only quasi-icosahedral because component X might reduce the symmetry of the core complex (see “Discussion”).

Fig. 3E shows averaged images of E2.X in the three char- acteristic orientations. The orientations of the individual images from which the averages were determined were con- firmed by tilting as in Fig. 4. The averaged images show the morphology of the E2 - X core more clearly than do the indi- vidual images. The diameter of the E2. X core in the 5-fold orientation is 22.5 k 1 nm, and the central hole (which arises by superimposition of pentagonal faces on opposite sides of the core) appears to have a diameter of 5 f 1 nm. The averaged images qualitatively resemble the projections (Fig. 3F) that were computed from the model (Fig. 3G). The largest devia- tions of the experimental data from the model occur for the 5-fold symmetric orientation. Here, the model actually shows 10-fold symmetry because the subunits from which it is con- structed are not asymmetric, as are the subunits comprising the E2.X core. The averaged 5-fold image shows 10 sectors of mass emanating from a centrally located circular density.

Cryoelectron Microscopy of Pyruvate Dehydrogenase Complex 24653

A

FIG. 3. S t ruc tu re of t h e E2. X core. A, a field of frozen-hydrated E 2 . X . K complexes. Some of the complexes appear to have one or a few peripherally located morphological units that might correspond to the kinase (arrowheads). Scale bar, 100 nm. B-D, selected contrast- enhanced images of E 2 . X cores in approximately 5-, 3-, and 2-fold symmetric orientations, respectively. E, averaged images of E 2 . X cores in 5-fold (left, n = 61), 3-fold (center, n = 15), and 2-fold (right, n = 21) orientations. F, projections of the three-dimensional model shown in C in orientations corresponding to those in E. G, three- dimensional model of the E 2 . X complex (see "Experimental Proce- dures"). In B-E, the frame width corresponds to 63 nm.

Additional mass a t higher radii appears between alternating pairs of sectors, giving the averaged projection its &fold character. Again, this is due to the subunits of the E2 - X core having a more complex shape than the simple spheres (ver- tices) and cylinders (edges) from which the model is con- structed.

E2- X. K-E3 Subcomplexes"E2. X. K-E3 subcomplexes were prepared by reconstitution in uitro from purified com- ponents. A field of frozen-hydrated subcomplexes (Fig. 5 A ) shows images of E2-X. K in which morphological units (av- erage of three per core in the 5-fold orientation) are visible a t the periphery, most of which we presume are E3. The back- ground concentration of unbound E3 is sufficiently low that

FIG. 4. Interconversion of images of E 2 . X cores by tilting. 5-fold symmetric views (left-most image in each triplicate) were chosen from a micrograph recorded with the specimen grid untilted, and then the same molecules were selected from a micrograph ob- tained with the specimen grid tilted by 35" (second image in each pair). Axis of tilting is the vertical direction. The third image in the sets was obtained by projecting the model of the E 2 . X core (Fig. 3G) after orienting it in the following manner. The model was first projected in the 5-fold orientation and then rotationally aligned with respect to the image from the untilted specimen (Le. the left-most image in a set). The aligned model was then tilted about the same axis as was the specimen, and its projection was determined. A-C show, respectively, two examples of tilt interconversions in which 2- fold views, 3-fold views, and views intermediate to 2- and %fold were obtained.

there is little uncertainty in classifying E3 molecules as being bound or free. Selected images of E2. X. K-E3 complexes are shown in Fig. 5B.

Examination of E2. X. K-E3 subcomplex images gives the impression that the E39 do not occupy fixed positions. The radial positions of the bound E3s vary, and often there is an apparent gap of variable size ( ~ 0 - 4 nm) between the surfaces of the E3 and E2. X. K core. In &fold symmetric images, it is difficult to ascertain the azimuthal position of the bound E3s relative to the E2. X. K cores owing to uncertainties in iden- tifying structural details as points of reference in the latter. In 2- or %fold symmetric views, this is a lesser problem, but the azimuthal positions of the E3 units relative to the cores do not seem consistent with a single type of binding location (e.g. at vertices or across faces). Overall, one is left with the impression that the E3 molecules are attached to the E2. X. K cores by flexible, extended tethers. A more quantitative description of this behavior is provided by Fig. 6A, which shows a histogram of measured distances of E3 molecules from the centers of the E2. X. K cores (&fold views) to which they are associated. The average radial position of the E3 is 17 nm, and, more importantly, the distribution of radii is broader, by about 4 nm, than would be expected from the error associated with a single determination, =1 nm.

Component X is responsible for binding E3 to the E2. X. K core (Powers-Greenwood et al., 1989; Gopalakrishnan et al., 1989). Thus, knowledge of the E3 locations in PDC should also help localize component X. However, this determination will be hampered if the E3 is indeed flexibly tethered to the E2. X. K core. Nevertheless, it should still be possible to test some models of the spatial distribution of component X. For example, component X polypeptides might be clustered to- gether in the core, perhaps as trimeric morphological units forming a few of the 20 vertices. To investigate this possibility, the azimuthal relationships among pairs of bound E3 mole- cules as they occur in subcomplexes (5-fold symmetric views) were characterized. A histogram of angles between pairs of E3 molecules (Fig. 7) shows no evidence for clustering; on the contrary, the frequency distribution shows a relative scarcity of inter-E3 angles between 0 and -30". Otherwise the distri-

24654 Cryoelectron Microscopy of Pyruvate Dehydrogenase Complex

A

B

FIG. 5. Electron microscopy of E2 .X.K-E3 subcomplexes. A, a field of complexes prepared by reconstitution in vitro a t a ratio of 12 mol of dimeric E3/mol of E2.X. K core. Scale bar, 100 nm. B- D, selected contrast-enhanced images of E2. X. K-E3 subcomplexes in 5-, 3-, and 2-fold orientations, respectively. Some of the morpho- logical units attributed to E3 are indicated by arrowheads. The width of a single frame corresponds to 63 nm.

bution is rather flat. A part of the explanation for the low frequency of angles less than 30” could be that E3s that are close together overlap and are therefore not distinguishable, but it is unlikely that this could be the entire explanation. Based upon this evidence that component X does not occur as clusters in the core and the limited number of component X polypeptides present, estimated a t about six per core (Jilka et al., 1986), we suggest a structural model in which compo- nent X is inserted either in a random manner or binding of one component X tends to hinder further X molecules binding a t neighboring regions on the E 2 inner core (cf. “Discussion”).

E2. X. K - E l Subcomplexes-Fig. 8 shows a micrograph and selected contrast-enhanced images of E2. X . K-E1 complexes that were prepared by reconstitution in vitro. As was observed for the E 2 . X . K - E 3 subcomplexes above, there is often an apparent gap of up to 4-6 nm between the E 2 - X - K-bound El molecules. Measurements of the distances between the centers of the E 2 . X . K core (&fold symmetric views) and the bound El unit reveal a broad distribution (Fig. 6B) consistent with a mode of attachment via thin, flexible polypeptide tethers.

A , O r - - - -

1 I i bh]

70 30

RADIUS (nm)

FIG. 6. Frequency distributions of radial coordinates of E2. X . K core-bound E3 and E l . A, distribution for E2.X.K-E3 subcomplexes (see Fig. 5). n = 320. B, distribution for E2.X.K-El subcomplexes (see Fig. 8). n = 309. See “Experimental Procedures.”

0 30 60 90 120 150 180

ANGLE (deg) FIG. 7. Frequency distribution of relative azimuthal loca-

tions of pairs of bound E3 molecules in E2. X . K-E3 subcom- plexes. n = 486.

DISCUSSION

The technique of cryoelectron microscopy of frozen-hy- drated macromolecular solutions has not previously been ap- plied to eucaryotic PDC. Unlike conventional electron micro- scopic techniques, such as negative staining, the native archi- tecture of the enzyme complex is expected to be preserved in the frozen state. Indeed, we have detected several differences in the structure of PDC as compared with previous studies (Reed and Oliver, 1968; Junger and Reinauer, 1972; Junger and Bachmann, 1977). Frozen-hydrated PDC has a diameter of 50 f 2 nm, which is significantly larger than the 40-45 nm or less reported in earlier studies by electron microscopy and agarose gel electrophoresis (Easom et al., 1989). Recent quas- ielastic light scattering studies on bovine heart and kidney PDC indicate a diameter of 49-51 nm for the native enzyme complex (Roche et al., 1991). Our fitting of model projections to tilted images and to additive images of known tilt strongly support 532 icosahedral symmetry originally proposed by Reed and Oliver (1968). These fits indicate the dihedral point

Cryoelectron Microscopy of Pyruvate Dehydrogenase Complex 24655

A

B

C

D

FIG. 8. Electron microscopy of frozen-hydrated E 2 . X . K - E l subcomplexes. Reconstitution performed at k 2 . X . K:I;l molar ratio of about 1%. A, field of subcomplexes. Scale bur, 100 nm. R-D, selected contrast-enhanced images in 5-, 3-, and 2-fold orientations, respectively. Some of the bound subunits attributed to El are indi- cated by arrowheads. The width of a single frame corresponds to 63 nm.

group 522 suggested by Oliver and Reed (1982) as a possible alternative symmetry for the inner core is not correct.

Another finding of this study is that the El and E3 mole- cules are apparently not bound directly to the surface of the E2 X inner core. This is consistent with the larger size of the native complex. The 20-30 molecules of bound El comprise most of the mass located exterior to the E2.X inner core complex and, hence, effectively determine the size of PDC. If the El were bound directly to the surface of the E2.X inner core, then the overall size of the complex would not be greater than the sum of the diameter of the inner core complex and twice that of the El. The diameter of the frozen-hydrated E2 .X inner core in the &fold orientation is 22.5 * 1 nm, a value that agrees well with most previously reported values, which range from 21-24 nm (Reed and Oliver, 1968; Junger and Reinauer, 1972). The El molecules vary in apparent diameter between 6 and 9 nm, reflecting differing orientations of the molecule. Values of 7-10 nm for the dimensions of El have been reported previously (Hayakawa et al., 1969; Junger and Reinauer, 1972). From these values for the size of El and the E2. X inner core, the diameter of PDC should be between

33 and 44 nm if no gaps were present. This is considerably less than we have observed. The gaps observed between the bound El (and E3) and the inner core in reconstituted sub- complexes account for the diameter observed for PDC. The maximum size of the gap is 4-6 nm, which appears to be sufficient to account for the 50-nm diameter of PDC.

Owing to the limited resolution of the electron micrographs and the low contrast of frozen-hydrated protein, there is often no visible mass connecting the bound El or E3 to the E2. X. K core. The gaps observed between the bound El and the core (Fig. 8), which range in size from 0 to about 6 nanometers, are likely bridged by polypeptide deriving primarily from the hinge regions. The hinge regions are located between the 5- kDa El-binding and inner domain in the amino acid sequence of E2 (Fig. 1). A similar hinge-type structure is thought to connect the E3-binding domain of X to its inner domain (Pate1 and Roche, 1990; Lawson et al., 1991). The hinge regions are especially susceptible to proteolysis, and some of the amino acids of the hinges are characterized by unusually high mobility as detected by nuclear magnetic resonance spectroscopy (Perham et al., 1987; Perham and Packman, 1989). This mobility is probably responsible for movement of lipoyl domains among active sites, but neither the scale of the motions allowed by the hinge regions or the locations of the components bound through hinge-linked domains were re- vealed in these studies. The variable gaps observed between the El or E3 and the inner E2.X core are consistent with a structural model of the hinge regions that allows movements of several nanometers by tethered components. Further sup- port for this model comes from images of frozen-hydrated PDC in which the morphological units (mainly El and E3) that are bound to the core do not give the impression of being arranged in an orderly manner and do not reflect the under- lying symmetry of the core.

Component X has a 28-kDa inner domain that remains associated with the 27-kDa E2 inner domains when the outer domains of both components are fully removed (Rahmatullah et al., 1989b). The location of the component X subunits in the E2. X subcomplex remains a mystery, but our cryoelectron microscopy of E2 -X. E3 complexes (Figs. 5 and 6) has pro- vided some new information. Since component X is involved in binding E3 (Powers-Greenwood et al., 1989; Gopalakrish- nan et al., 1989; Lawson et al., 1991), the bound E3 in the subcomplexes serves as a marker of X. The -six copies of X that are present in each core complex appear to be distributed more or less uniformly in the complex, as opposed to being clustered in one or two locations. It is difficult to envision an organization of the E2. X core in which the X subunits would be arranged in a symmetric pattern that is consistent with the apparently icosahedral symmetry of the E2 inner core. Both the mammalian E2 (Powers-Greenwood et al., 1989) and yeast E2 (Lawson et al., 1991) form assembled E2 oligomers without component X. However, it is not clear whether X is bound after assembly, as suggested by Lawson et al. (19911, or whether its inner domain integrates into the inner core during assembly? In the future, cryoelectron microscopy of E2. X cores containing bound antibody specific for the inner domain of X should aid in resolving this issue.

The finding that tethered components are probably not in fixed positions and have significant accessible volumes (cf. below) for movement may have relevance for the function and regulation of the complex. There are 120 E2-lipoyl domains and six X-lipoyl domains in the complex. Such flexibility and

Resolved Component X active in binding E3 does not bind to the assembled E2 core. L. Li, G. A. Radke, K. Ono, and T. E. Roche, submitted for publication.

24656 Cryoelectron Microscopy of Pyruvate Dehydrogenase Complex

movement may allow several lipoyl domains to directly trans- fer reducing equivalents to active sites of the six E3 dimers. Regarding regulation, a full complement of bound El tetra- mers becomes available for phosphorylation by about two to three tightly bound kinase molecules much more rapidly than El dissociates from E2 (Brandt and Roche, 1983). Although the kinase is bound to the lipoyl domain region of E2 (Rah- matullah et al., 1990), flexibility of that region would not be expected to afford sufficient movement to reach all the El tetramers. Bovine kidney and heart PDC contain 20-30 bound El tetramers per complex or, at most, one tetramer per two E2 subunits. However, reconstituted complexes can have up to 55 bound El tetramers (Wu and Reed, 1984), indicating one E2 subunit can bind a tetramer. Unless asymmetry exists in the a& tetramer, it must have the potential to interact with two E2 subunits. Brandt and Roche (1983) suggested that “flexible extensions” connecting a@ units of the a& tetramers may facilitate nondissociative intracore migration of El. This would involve intermediate binding of a tetramer by only one E2 subunit and exchange with another E2 subunit as a consequence of dissociation and association at the other El-binding regions. Such a process would clearly be aided by the freedom of movement of flexibly tethered components that are implicated by our micrographs.

Steric inhibition of motions by the lipoyl domains and components bound to the outer domains of E2 and component X would be minimal due to the sparse packing of these components in PDC. Approximating the overall shapes of PDC and the inner E2 core as spheres of diameters 50 and 22.5 nm, respectively, we estimate the volume exterior to the inner core to be 59,500 nm3. This volume is occupied by El (30 molecules in heart PDC), E3 (six copies), kinase (one to three copies), phosphatase (a few copies), and the outer do- mains of E2 and X. We estimate, based upon the reported masses and estimated stoichiometries of these components (summarized in Patel and Roche, 1990) and assuming a spe- cific volume of 1.2 X nm3/Da, that the total volume occupied by these peripherally located constituents is 9,400 nm3 in heart PDC and 7,800 nm3 in kidney PDC. These values represent only 13-16% of the available volume. We estimate that for the entire heart PDC complex (i.e. including the inner E2 core) about 80% of the volume potentially accessible to the various protein components is actually occupied by sol- vent.

Acknowledgments-We are grateful to Catherine Forneris for tech- nical assistance and critical evaluation of the manuscript, to Dr. Joachim Frank for use of the image processing facilities, and to Dr. Pawel Penczek for the computer modeling of the E2 core.

REFERENCES Adrian, M., Dubochet, J., Lepault, J., and McDowall, A. W. (1984)

Nature 308,32-36 Barrera, C. R., Namihira, G., Hamilton, L., Munk, P., Eley, M. H.,

Linn, T. C., and Reed, L. J. (1972) Arch. Bwchem. Biophys. 1 4 8 , 343-358

Brandt, D. R., and Roche, T. E. (1983) Biochemistry 22 , 2966-2971 Chiu, W. (1986) Annu. Rev. Biophys. Biophys. Chem. 15,237-257 Coppel, R. L., McNeilage, L. J., Surh, C. D., Van de Water, J.,

Spithill, T. W., Whittingham, S., and Gershwin, M. E. (1988) Proc. Natl. Acad. Sci. U. S. A. 8 5 , 7317-7321

Dubochet, J., Adrian, M., Chang, J.-J., Homo, J.-C., Lepault, J.,

McDowall, A. W., and Schultz, P. (1987) Q. Rev. Biophys. 21,129- 228

Easom, R. E., DeBuysere, M. S., Olson, M. S., and Serwer, P. (1989) Proteins 5 , 224-232

Frank, J., Verschoor, A., and Wagenknecht, T. (1985) in New Meth- odologies in Studies of Protein Configuration (Wu, T. T., ed) pp. 36-39, van Nostrand Reinhold, New York

Frank, J., Goldfarb, W., Eisenberg, D., and Baker, T. S. (1978) Ultramicroscopy 3,283-290

Frank, J., Shimkin, B., and Dowse, H. (1981) Ultramicroscopy 6,

Gopalakrishnan, S., Rahmatullah, M., Radke, G . A. Powers-Green- wood, S. L., and Roche, T. E. (1989) Biochem. Biophys. Res. Commun. 160,717-721

Grassucci, R., Wagenknecht, T., Radke, G. A., and Roche, T . E. (1990) in Proceedings of the Twelfth International Congress on Electron Microscopy, pp. 260-261, San Francisco Press, San Fran- cisco

Hayakawa, T., Kanzaki, T., Kitamura, T., Fukuyoshi, Y., Sakurai, Y., Koike, K., Suematsu, T., and Koike, M. (1969) J. Biol. Chem.

Ishikawa, E., Oliver, R. M., and Reed, L. J . (1966) Proc. Natl. Acad.

Jilka, J. M., Rahmatullah, M., Kazemi, M., and Roche, T. E. (1986) Jaffe, J. S., and Glaeser, R. M. (1987) Ultramicroscopy 2 3 , 17-28

Junger, E., and Reinauer, H. (1972) Biochim. Biophys. Acta 250,

Junger, E., and Bachmann, L. (1977) Biochim. Biophys. Acta 4 8 1 ,

Lawson, J. E., Behal, R. H., and Reed, L. J. (1991) Biochemistry 30,

Lepault, J., Booy, F. P., and Dubochet, J. (1983) J. Microsc. (Oxf.)

Linn, T. C., Pelley, J. W., Pettit, F. H., Hucho, F., Randall, D. D.,

Milkan. R. A.. Brisson. A., and Unwin, P. N. T. (1984) Ultramicro-

343-358

244,3660-3670

Sci. U. S. A. 5 6 , 534-541

J. Biol. Chem. 261, 1858-1867

478-490

364-376

2834-2839

129,89-102

and Reed, L. J. (1972) Arch. Biochem. Biophys. 148,327-342

scopy 1 3 , l - l o Oliver. R. M.. and Reed. L. J. (1982) in Electron Microscopy of

Proteins (Harris, R., edj Vol. 2, pp. 1-48, Academic Press, London Patel, M. S., and Roche, T. E. (1990) FASEB J. 4, 3224-3233 Perham, R. N., and Packman, L. C. (1989) Ann. N. Y. Acad. Sci.

Perham. R. N.. Packman. L. C.. and Radford, S. E. (1987) Biochem.

. .

573 , l -20

SOC. Syrnp. 54,67-81 Powers-Greenwood. S. I,.. Rahmatullah. M.. Radke, G. A., and Roche, T . E (1989) J. B$L. Chem. 264,3655-3657

Rahmatullah, M., and Roche, T. E. (1987) J. Biol. Chem. 262,10265- 10271

Rahmatullah, M., Gopalakrishnan, S., Andrews, P. C., Chang, C. L., Radke, G. A., and Roche, T. E. (1989a) J. Biol. Chem. 264,2221- 2227

Rahmatullah, M., Gopalakrishnan, S., Radke, G. A., and Roche, T. E. (1989b) J. Biol. Chem. 2 6 4 , 1245-1251

Rahmatullah, M., Radke, G. A., Andrews, P. C., and Roche, T. E. (1990) J. Biol. Chem. 2 6 5 , 14512-14517

Reed, L. J., and Oliver, R. M. (1968) Brookhaven Symp. Biol. 21 , 397-412

Roche, T. E., and Cate, R. L. (1977) Arch. Biochem. Biophys. 183 , 664-677

Roche, T. E., Sorensen, C. M., Shi, W., Zhang, W., Powers-Green- wood, S. L., Roche, E. D., and Cox, D. J. (1991) FASEB J. 5, A1202

Stepp, L. R., Pettit, F. H., Yeaman, S. J., and Reed, L. J . (1983) J. Biol. Chem. 258,9454-9458

Taylor, K., and Glaeser, R. M. (1976) J. Ultrastruct. Res. 55, 448- 456

Thekkumkara, T. J., Ho, L., Wexler, I. D., Pons, G., Liu, T.-C., and Patel, M. S. (1988) FEBS Lett. 240, 45-48

Wagenknecht, T., Grassucci, R., and Frank, J. (1988) J. Mol. Biol.

Wagenknecht, T., Grassucci, R., and Schaak, D. (1990) J. Biol. Chem.

Wu, T.-L. and Reed, L. J. (1984) Biochemistry 23, 221-226

199,127-147

265,22402-22408