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Jul 19, 2005 - Alexander von Humboldt Foundation (to B.R.), National Institutes of. Health Grant GM062989 (to R.M.G.), Swiss National Foundation Grant.
Molecular architecture and assembly mechanism of Drosophila tripeptidyl peptidase II Beate Rockel*†, Ju¨rgen Peters*, Shirley A. Mu¨ller‡, Go¨nu¨l Seyit*, Philippe Ringler‡, Reiner Hegerl*, Robert M. Glaeser§, and Wolfgang Baumeister* *Department of Molecular Structural Biology, Max Planck Institute of Biochemistry, D-82152 Martinsried, Germany; ‡Maurice E. Mu¨ller Institute, Biozentrum, University of Basel, CH-4056 Basel, Switzerland; and §Department of Molecular and Cell Biology, University of California, Berkeley, CA 94720-3206

In eukaryotes, tripeptidyl peptidase II (TPPII) is a crucial component of the proteolytic cascade acting downstream of the 26S proteasome in the ubiquitin-proteasome pathway. It is an amino peptidase belonging to the subtilase family removing tripeptides from the free N terminus of oligopeptides. The 150-kDa subunits of Drosophila TPPII assemble into a giant proteolytic complex of 6 MDa with a remarkable architecture consisting of two segmented and twisted strands that form a spindle-shaped structure. A refined 3D model has been obtained by cryoelectron microscopy, which reveals details of the molecular architecture and, in conjunction with biochemical data, provides insight into the assembly mechanism. The building blocks of this complex are apparently dimers, within which the 150-kDa monomers are oriented head to head. Stacking of these dimers leads to the formation of twisted single strands, two of which comprise the fully assembled spindle. This spindle also forms when TPPII is heterologously expressed in Escherichia coli, demonstrating that no scaffolding protein is required for complex formation and length determination. Reciprocal interactions of the N-terminal part of subunits from neighboring strands are probably involved in the formation of the native quaternary structure, lending the TPPII spindle a stability higher than that of single strands. 3D reconstruction 兩 cryoelectron microscopy 兩 scanning transmission electron microscopy

I

ntracellular proteolysis must be subject to spatial and temporal control to avoid havoc. Self-compartmentalization serves as a key stratagem, enabling the cell to confine proteolysis to selfassembled nanocompartments. As a consequence, a number of large homo- or heterooligomeric proteolytic complexes have evolved; the prime example is the 26S proteasome, which degrades ubiquitinated proteins in an ATP-dependent manner (1, 2). However, proteases that can degrade the peptides released by the proteasome further, such as the Tricorn protease in prokaryotes (3, 4) or tripeptidyl peptidase II (TPPII) in eukaryotes (5), are also very large megadalton complexes. TPPII has recently attracted attention because, in addition to its role in general intracellular protein turnover, it seems to be capable of complementing 26S proteasome function to some extent under conditions in which the latter is inhibited (6–9). TPPII plays a role in pathological conditions such as muscle sepsis (10, 11) and in apoptosis (12–14) and, furthermore, is instrumental in the N-terminal trimming that occurs in the late stages of antigenic peptide processing for the MHC class I system (15–17). A membrane-bound form exists in addition to cytosolic TPPII, and this form is responsible for the degradation of cholecystokinin (18, 19), one of the most abundant neurotransmitters in the brain and an important signal factor for the peripheral and central nervous systems (20). TPPII has only been found thus far in eukaryotes, in which it exists as a large homooligomeric assembly with a molecular mass far beyond 1 MDa. It is a serine protease of the subtilisin type carrying a 200-aa insert between its catalytic Asp-44 and His-264 www.pnas.org兾cgi兾doi兾10.1073兾pnas.0504569102

residue. The size of its subunit varies from 138 kDa in mammals to 150 kDa in plants, insects, and worms (5, 21). Although most members of the subtilisin family are endopeptidases (22), the basic activity of TPPII is the release of tripeptides from the free N terminus of peptides, but endoproteolytic activity has been measured also (7, 16). TPPII occurs as a spindle-shaped structure (28 ⫻ 60 nm) that is formed by two segmented and twisted strands. Although this seems to be the preferred conformation in vivo, the size of TPPII particles seems to be very variable in vitro, as indicated by size-exclusion chromatography (23) and electron microscopy (24, 25). This variance is attributable, in principle, to the stacking of segments into strands for which there is no obvious termination. Thus, the architecture of TPPII is fundamentally different from that of barrel-shaped proteolytic complexes such as the 20S proteasome (which has a ring structure confined to four rings), that of the archaeal functional TPPII homologue Tricorn [a hexamer that can assemble into an icosahedral capsid (26)], or that of TET [an archaeal aminopeptidase that assembles into a tetrahedron (27–29)]. Mainly because of the apparent structural heterogeneity of purified TPPII, which may be attributable to its concatameric architecture and a result of its lability during purification, progress has been slow in elucidating its structure; as a result, its function is not at all understood on the molecular level. Thus, the dependence of TPPII activity on the oligomeric state as well as the reasons for its enormous size have remained an enigma, as have the factors determining the unique quaternary structure that this complex assumes. Because the inherent propensity of purified preparations of TPPII to form heterogeneous structures in vitro hampers crystallization, we recently set out to study the structure of this protease by cryoelectron microscopy (25). Here we report an improved cryoelectron-microscopy structure of TPPII. This structure provides clues on subunit arrangement and segment boundaries and thus enables us to map the orientation of the segments within a strand and deduce the spatial arrangement of the monomers. Using an N-terminally tagged mutant, we also show that the N-terminal region of the TPPII monomer is essential in the assembly of the holocomplex. Finally, we present a model for the assembly of the spindle-shaped complex that can explain its ability to assemble into an oligomer of defined length. Materials and Methods Electron Microscopy. TPPII was purified from Drosophila embryos (25). Protein solution (4 ␮l) was applied to lacey carbon grids, and the grids were subsequently washed twice by using buffer without glycerol before being plunged into liquid ethane. Focal pairs were recorded on Kodak SO-163 films with a JEOL 4000EX microscope operated at 400 kV and a calibrated magAbbreviations: TPPII, tripeptidyl peptidase II; MBP, maltose-binding protein. †To

whom correspondence should be addressed. E-mail: [email protected].

© 2005 by The National Academy of Sciences of the USA

PNAS 兩 July 19, 2005 兩 vol. 102 兩 no. 29 兩 10135–10140

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Communicated by Pamela J. Bjorkman, California Institute of Technology, Pasadena, CA, June 2, 2005 (received for review April 20, 2005)

nification of ⫻37,440. Defocus values for the close-to-focus images ranged from ⫺3.4 to ⫺1.3 ␮m. Images were scanned by using a Nikon Super Coolscan 8000 ED scanner at a pixel size of 12.5 ␮m. 3D Reconstruction of Ice-Embedded TPPII Particles. Far-from-focus

images were aligned to close-to-focus images, and 6,123 particle images were selected interactively by using the EMAN software program (30). Particles were boxed out from the close-to-focus images according to the resulting position lists. Images were contrast transfer function–corrected by using the ‘‘ctfit’’ option of EMAN. After visual inspection of the resulting particle stack for distance of the particles from the edge of the hole in the carbon film, particle preservation, and contamination, 4,809 particles were kept for 3D alignment, which was carried out by projection matching using our previous 3D reconstruction of TPPII at 3.3-nm resolution (25) as reference and imposing C2 symmetry assuming antiparallel orientation of the strands. Additionally, a solid double bow (25) was used as reference without imposing any symmetry. In both cases, no additional improvement of the resulting 3D reconstruction was noticeable after five cycles of alignment. The resolution of the reconstructions was estimated by Fourier shell correlation. According to the 0.5 criterion, the resolution was 2.2 nm for the 3D reconstruction with imposed C2 symmetry and 2.7 nm for the nonsymmetrized 3D reconstruction. 3D-Image Analysis. Additional image analysis of both 3D reconstructions was carried out by using the EM software package (31). The two strands comprising the TPPII molecule were segmented out from the C2-symmetrized and nonsymmetrized 3D reconstructions. A small 3D box containing a segment was cut out from the center of the strands and used as a search model to find the positions and orientations of the other segments. Strand– strand distances were calculated by using the x and y shifts of each segment with respect to the centered reference segment. The tilt angle (␪) and the rotation (␾ ⫹ ␺) around an axis parallel to the longitudinal axis of the TPPII complex running through the center of a segment were calculated by using segments five and six as references. Mass Determination by Scanning Transmission Electron Microscopy. A

5-␮l aliquot was adsorbed to glow discharged thin carbon film that spanned a thick fenestrated carbon layer covering a goldplated copper microscopy grid. The grid was left unstained, blotted, washed four times with 10 mM ammonium acetate, and freeze-dried at ⫺80°C and 5䡠10⫺8 torr (1 torr ⫽ 133 Pa) overnight in the microscope. A HB-5 scanning transmission electron microscope (Vacuum Generators, East Grinshead, U.K.) interfaced to a modular computer system (Tietz Video and Image Processing Systems GmbH, Gauting, Germany) and operated at 80 kV was used for the mass measurements. Series of digital dark-field images were recorded (nominal magnification: ⫻100,000; dose, 69.2 ⫾ 8.5 electrons per nm2). Grids prepared for negative-stain microscopy were similarly washed and stained with 2% uranyl acetate. Dark-field images were recorded at 100 kV (nominal magnification: ⫻500,000; dose: ⬇5,450 electrons per nm2). The mass-measurement images were evaluated as described (32) by using the program package IMPSYS (33). The resulting values were scaled according to the mass measured for tobacco mosaic virus. No correction was required for beam-induced mass-loss, because in the dose range used, it is ⬍1% (34). Recombinant Expression and Purification of Wild-Type and MaltoseBinding Protein-Tagged TPPII. Wild-type TPPII was expressed from

the vector pET 30 containing the structural gene for TPPII using BL21 (DE3) cells. Protein N-terminally tagged with maltosebinding protein (MBP) was obtained from TB1 cells containing 10136 兩 www.pnas.org兾cgi兾doi兾10.1073兾pnas.0504569102

the gene cloned in the pMAL vector. In both cases, induction was carried out at 18°C for 24 h and in the presence of 0.1 mM isopropyl thiogalactoside. Wild-type protein was purified as follows: crude extract was subjected to two passages through a cell disrupter and centrifuged for 20 min at 20,000 ⫻ g, and the sediment was resuspended with 100 mM potassium phosphate (pH 7.5)兾5% glycerol and again passed through the cell disrupter. After centrifugation for 20 min at 20,000 ⫻ g, the supernatant was collected and separated on a Superose 6 size-exclusion column as described for native TPPII (25). MBP-tagged protein was separated on amylose resin, followed by chromatography on Superose 6. The buffer routinely used was 80 mM potassium phosphate (pH 7.5)兾1 mM DTT. Finally, 5% glycerol and 10 mM DTT were added to facilitate storage at room temperature and freezing. Activity was assayed as described (25). MBP-Tag Removal Using Proteinase K. Because factor Xa did not

cleave MBP from MBP-tagged TPPII, proteinase K was used. MBP-tagged TPPII was dissolved in 80 mM potassium phosphate兾1 mM DTT, incubated with proteinase K (proteinase K兾TPPII ratio of 1:250 by weight) on ice for 30 min, and separated on Superose 6 as described above. The fraction eluting at 7.8–8.8 ml was subjected to ammonium sulfate precipitation (1.5 vol) on ice, and the sediment was rechromatographed. The high-molecular-weight fraction was collected again and scrutinized by electron microscopy after the addition of 5% glycerol using uranyl acetate as stain. Results 3D Structure of TPPII at 2.2-nm Resolution. The majority of projec-

tions seen in electron micrographs of ice-embedded TPPII particles are side views that are elongated in shape and characterized by navette- (pointed oval), dumbbell-, or fish-shaped outlines (Fig. 1A). Less abundant are the more compact projections of particles that that are oriented perpendicular to the ice film. Using a total of 4,809 particle views and imposing C2 symmetry, we calculated a 3D reconstruction of the TPPII complex of Drosophila melanogaster at a resolution of 2.2 nm (Fig. 1B). The spindle-shaped protease (60 ⫻ 28 nm) is composed of two segmented strands (12 nm in width), which are closely apposed at the spindle poles but ⬇4 nm apart midway between them (Fig. 1C). In contrast to our previous reconstruction at 3.3-nm resolution, segment boundaries can be drawn easily, and the building blocks comprising the two twisted strands are revealed as kinked and angular segments. In each of the two strands comprising the TPPII spindle, 10 such segments are assembled linearly (Fig. 1C). To obtain the translation vectors and rotation matrices that describe the transformation of segment n into segment n ⫹ 1, we used a box containing one of the two central segments (i.e., either segment five or six) as a 3D search template. Because of the interdigitation of the segments within the strands, this box also contains two half-segments that intrude from below and above. To detect effects that were possibly induced by the imposed C2 symmetry, we also performed the analysis with a model that was reconstructed without imposing any symmetry. The parameters obtained were basically similar for both reconstructions (Fig. 1D). The vertical distance between the segments in a strand was ⬇5.5 nm, and the segment-to-segment rotation (i.e., the rotation around an axis parallel to the longitudinal axis of the TPPII complex and running through the center of the segment) was ⬇25°. Additionally, neighboring segments differed in their local tilt angle: Tilt angles calculated relative to the 3D search template ranged from ⫺10° to ⫹10° depending on the position of the segment and showed segment-to-segment increments of ⬇2° (Fig. 1D). This locally varying tilt angle underlines the nonhelical structure of TPPII and is probably responsible for the Rockel et al.

gradual reduction of the strand-to-strand distance in the direction of the spindle poles (Fig. 1). TPPII Is a 6-MDa Complex. The mass values for TPPII given in the literature vary from ⬎1 MDa (5) to 5–9 MDa (7) and are rough estimates derived from size-exclusion chromatography. Therefore, we used scanning transmission electron microscopy to obtain a more accurate value of the particle mass of the TPPII spindle. The quality of the preparation after the washes required to remove buffer salts and glycerol before mass measurement was documented by images recorded from grids that were negatively stained after the wash steps (Fig. 2B). Mass determination was performed on corresponding unstained, freeze-dried samples. The masses measured for the TPPII particles fell into two categories. As shown by the corresponding images, the smaller particle comprised single TPPII strands with a mass of 2.12 ⫾ 0.82 MDa (n ⬇ 170; standard error, ⫾0.06 MDa; overall uncertainty, ⫾0.12 MDa; Fig. 2 A and C). The fully assembled complexes, readily identifiable by their dumbbell-, navette-, or fish-shaped projections, formed the major category and had masses of 5.66 ⫾ 1.03 MDa (n ⬇ 480; standard error, ⫾0.05 MDa; overall uncertainty, ⫾0.29 MDa; Fig. 2 A and C). Al-

Fig. 2. Mass determination by scanning transmission electron microscopy. (A) Histogram displaying the mass values measured for an unstained freeze-dried TPPII sample. The Gaussian peaks fall at masses of 2.12 and 5.66 MDa, respectively. (B) Scanning transmission electron microscopy dark-field images of the TPPII complex recorded from grids that had been washed as for the mass measurements and negatively stained with 2% uranyl acetate. (C) Galleries showing particles with masses in the range of 1.30 –2.90 MDa (Left; almost the whole range of the first peak) and 5.46 –5.86 MDa (Right; central region of the second peak). The projections clearly distinguish the particles as single- and double-stranded (intact) complexes, respectively. (Scale bars: 25 nm in B and 50 nm in C.)

Rockel et al.

though single strands should give one half the mass of a fully assembled complex, the averaged value obtained is lower and reflects their frequent partial fragmentation (see below). Indeed, there was a population of much smaller particles present (data not shown). With 10 segments per strand and 20 segments per TPPII spindle, the mass calculated for an individual segment is 283 ⫾ 52 kDa, indicating that the subunits are dimers (2 ⫻ 150 kDa). The combined mass of such a 40-mer, 6 MDa, is in good agreement with the measured value of 5.66 MDa. Arrangement of the Monomers Within the Dimers. Using the rotation

matrices and translation vectors that were obtained by the 3D search with reference segment six, we calculated averages over the 10 segments of one strand for both the symmetrized and nonsymmetrized TPPII models. Both averages exhibit a twofold rotational axis, which is most obvious at the distal surface of the average (i.e., the surface away from the axis of the spindle), where a vertical density links the two levels of one segment like a handle (Fig. 3 A and B). The two levels, which are of globular shape, seem to be associated diagonally at the proximal side of the average (i.e., the side that is oriented toward the neighboring strand in the TPPII model). A dimer was computationally excised from the average. It is obvious that the front and back views of the dimer differ explicitly, suggesting that the monomers are arranged in a head-to-head fashion (Fig. 3C).

Fig. 3. Averaged TPPII dimers. Average over the 10 segments of one strand of the C2-symmetrized (A) and nonsymmetrized (B) TPPII reconstruction shown as a distal surface representation and as a projection (Upper). (Lower) Proximal surface representation. The probable locations of the monomers are denoted with M and M⬘. The threshold value for the proximal surface representation was set higher than for the distal surface representation to stress the diagonal linkage of the globular domains at the proximal side of the average. (C) Excised TPPII dimer in different orientations. The threshold value for the surface representation of the excised dimer was set to include a mass of 300 kDa is included. (Scale bars: 15 nm.) PNAS 兩 July 19, 2005 兩 vol. 102 兩 no. 29 兩 10137

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Fig. 1. 3D structure of TPPII. (A) TPPII particles embedded in vitreous ice, recorded at 400 kV and at a defocus of ⫺5 ␮m. (B) Fourier shell correlation (FSC). (C) Surface representations of C2-symmetrized TPPII model in navette (Left) and dumbbell (Right) orientations. Segment numbers are given on the side of the strands. Segment six is highlighted in purple and enclosed by a box. (Scale bar: 30 nm.) (D Upper) Tilt angles (␪) determined for segments 1–10 of both strands plotted versus their rotation angle around the longitudinal axis of the complex (␾ ⫹ ␺) using segments six (closed symbols) and five (open symbols) as references. Note that this rotation angle (␾ ⫹ ␺) corresponds to the z height of the respective segment. (D Lower) Segment-to-segment distances plotted versus the rotation angle (␾ ⫹ ␺). Note that the shift between the curves results from the different z height of reference segment five versus six. Squares, C2-symmetrized 3D reconstruction; circles, nonsymmetrized 3D reconstruction.

Fig. 4. TPPII expressed in E. coli. (A) TPPII particles expressed in E. coli embedded in vitreous ice. (B–E) Enlarged particles excised from A: TPPII spindles (B); breakdown products (C); long single strands (D); and extended spindles (E).

From the twofold rotational symmetry of the averaged segments obtained from both the C2-symmetrized and the nonsymmetrized reconstruction at the present resolution, we conclude that the top and bottom surfaces of the dimers are basically identical. Consequently, the strands, which are built from stacked dimers, do not show any obvious polarity, which in turn might facilitate the observed assembly of single strands into spindles (see below). The Double Clamp: A Means of Stabilization and Length Determination? Native TPPII, isolated from Drosophila, is only stable and

active for prolonged periods of time if it is assembled into spindles. Single strands do occur as well and are normally shorter than spindles, whereas longer strands are extremely rare. The specific activity toward the substrate Ala-Ala-Phe-4-methylcoumarin-7-amide is the same for single-stranded and doublestranded TPPII. The specific activity of single-stranded MBPtagged TPPII (see below) and wild-type TPPII (both native and heterologously expressed in E. coli) is 21,000 ⫾ 1,000 pmol䡠␮g⫺1䡠min⫺1, and the activity of intact spindles is indistinguishable from the activity of dissociated spindles that are still assembled to longer strands. However, single strands seem to be more fragile and prone to dissociation. When the 150-kDa subunits of Drosophila TPPII are expressed in E. coli, fully active and fully assembled TPPII complexes occur, but long single strands and breakdown products coexist with extended complexes (Fig. 4). This observation demonstrates that the TPPII complex can assemble without the aid of helper proteins or cofactors and that it is the spatial packing arrangement of the dimers within the strands that causes the characteristic spindle shape. To determine which parts of the dimers participate in the strand–strand interactions, we placed the averaged dimers at the positions found by the 3D alignment (Fig. 5A). It is obvious that a close contact region is established between the two globular domains of dimer one in the first strand and one globular domain of each of dimers one and two in the second strand. In the second strand, the scenario is analogous: Dimer one of the second strand is in close contact with dimers one and two of the first strand (Fig. 5B). The resulting structure thus forms a double clamp in which the terminal dimer of one strand ‘‘locks’’ the two terminal dimers of its neighboring strand and vice versa, probably resulting in a higher stability for the complex than can exist for single strands. In contrast to the single strands observed in the preparations, which vary in the number of dimeric subunits, the spindles are of defined length. Thus, the double clamp may also be crucial in length determination. Spindle Formation Must Be Driven by a Free-Energy Gain Resulting from Interstrand Contacts. There are no obvious limitations to the

linear extension of a single or double strand of TPPII. Therefore, because the spindle conformation is nevertheless clearly pre10138 兩 www.pnas.org兾cgi兾doi兾10.1073兾pnas.0504569102

Fig. 5. The double clamp. (A) Surface representation of the C2-symmetrized TPPII model into which the dimers were placed according to their alignment parameters. The two strands are color-coded yellow and blue. The box denotes the segments participating in double-clamp formation that are shown enlarged in B. (B) Double clamp formed by segments one and two (dark and light blue, respectively) from strand one and segments one and two (red and yellow, respectively) from strand two. (B Right) The clamp after a 180° rotation around the longitudinal axis of the complex.

ferred and the complex can assemble autonomously both in the E. coli host and in purified form, this state must be thermodynamically favored either by sterical factors or through the binding energy gained by interstrand contacts. Lateral contact between two strands through the proximal globular domains is a prerequisite for formation of the spindleshaped TPPII complex. As a consequence, a bulky tag such as MBP (35) (molecular mass, 43 kDa) at the contact area is likely to prevent the formation of double bows. To map these contact regions, we fused MBP to the N terminus of TPPII from D. melanogaster and observed that long single strands were formed in E. coli, which exhibited the same specific molar activity for hydrolysis of Ala-Ala-Phe-4-methylcoumarin-7-amide as does wild-type TPPII. However, no spindle-shaped assemblies were observed (Fig. 6A). When the MBP tag was cut off enzymatically and the sample was concentrated by ammonium sulfate precipitation, the majority of TPPII particles appeared as spindles, but single strands were still present, as were spindles with single strands extending beyond their poles (Fig. 6B). Even double-stranded spirals longer than 200 nm were observed occasionally. Spindles carrying strand extensions were also found when untagged TPPII was expressed in E. coli and concentrated with ammonium sulfate (Fig. 6D). Thus, the formation of spindles does not preclude additional extension of strands on steric terms. Cleaving the MBP tag from TPPII by proteinase K treatment affected neither the specific activity nor the structure of the complex (see Fig. 6 B and C). Furthermore, the appearance of wild-type TPPII after treatment with proteinase K also did not hint at any truncation as judged by electron microscopy and according to activity measurements. Only the probable loop in the C-terminal region, which is partially cleaved in vivo with both native and recombinant protein and gives rise to a weak band at an apparent molecular mass of 115 kDa (Fig. 6E), was now cleaved completely. This cleavage resulted in a band at 32 kDa, in addition to the 45-kDa band attributable to MBP. The MBP construct (192 kDa; apparent molecular mass, ⬇200 kDa; Fig. 6E) showed an additional band at ⬇170 kDa, which confirmed the assumed C-terminal nicking. After titration of recombinant complexes of TPPII, such as those shown in Fig. 6B with guanidine, a stage of disassembly was reached, where spindle-shaped complexes were strongly enriched. Single-stranded fragments disappeared, being disassembled into smaller complexes, which were then separated efficiently from spindles by molecular sieve chromatography (Fig. 6C). At the same time, appendixes at the ends of the spindles were also trimmed, leaving intact spindles and separable small fragments. The guanidine concentration required for optimal Rockel et al.

retention of intact spindles depended on the protein concentration and temperature and was ⬇500 mM. At concentrations beyond ⬇650 mM, spindles also disassembled, as was obvious from size-exclusion chromatography, activity measurements, and electron microscopy, which testifies to an increased stability of spindles over nonspindles. Thus, both the formation of spindles from segments and the breakdown of extended complexes into spindles are thermodynamically favored. From the strict dependence of double-strand formation on the removal of the bulky tag, we conclude that the N terminus of the TPPII monomer is situated in the proximal globular domain and oriented toward the strand-to-strand interface in the TPPII spindles. This orientation is corroborated further by tomographic reconstructions of ice-embedded MBP-tagged TPPII strands, in which additional globular masses (probably representing the MBP tag) are visible along the backbone of the single strands (data not shown). Discussion TPPII complexes isolated from Drosophila embryos are giant spindle-shaped 28 ⫻ 60-nm particles consisting of two segmented and twisted strands. Individual segments are likely to represent dimers within which the monomers are arranged in a head-to-head and tail-to-tail fashion. Because the TPPII strands are composed of a linear assembly of dimers, the interfaces of the two strands are identical; yet, the native, fully assembled TPPII complexes isolated from mammals and Drosophila are of defined length. In TPPII preparations from Drosophila we have never observed any particle among thousands that carried an extension beyond one of the spindle poles, unless the samples had been concentrated by ultrafiltration. Additionally, the spindle form always clearly predominated over single strands and broken single strands, provided optimal buffer conditions [phosphate buffer (pH 7.0–8.2); absence of chloride] had been maintained throughout. In Drosophila egg lysates, there is no significant fragmentation of TPPII complexes into small oligomeric complexes, as judged by Western blotting of crude lysate separated by size-exclusion chromatography (data not shown). Intact TPPII spindles can also be expressed in E. coli, although the assembly seems to be less complete with respect to intact spindle formation. As shown in the present work, the N-terminal domain of TPPII is involved in the strand-to-strand contacts that result in the formation of the spindle; N-terminally MBP-tagged TPPII only assembles into single strands, and no spindles are observed until the tag is removed. The mechanism of spindle formation, in principle, may be envisaged in three different scenarios: (i) A scaffolding protein that also provides length determination aids the assembly. (ii) An initiation complex (first spindle pole) is formed from two tetramers. Starting from this initiation complex, two strands grow in parallel by the addition of dimers until Rockel et al.

they meet to form the second spindle pole. Additional extension is prevented by steric factors (‘‘steric clash’’). Alternatively, a kind of ‘‘terminating structure’’ could be formed at the spindle poles. (iii) No initiation complex is formed. Instead, single strands grow separately by the progressive addition of dimers until a length corresponding to ⱖ10 dimeric subunits has been attained (step 1). Subsequently, single strands of sufficient length assemble laterally into double strands (step 2). This assembly leads to complexes having extensions of different length beyond the contact sites. Driven by free-energy minimization, the extensions are eventually lost (step 3) and go through steps 1–3 until, ideally, all complexes are present as spindles. The first possibility can be finally dismissed by the successful expression of TPPII spindles in E. coli and, moreover, by their reconstitution in vitro using ammonium sulfate precipitation. The second mechanism requiring an initiation complex and the parallel growth of the two strands is unlikely, because the experiments with MBP-tagged TPPII show that isolated single strands can indeed form double strands as long as the contact regions are accessible. Also, precisely synchronous growth would be required, and spindles having a terminal extension of any kind should never be observed. However, we have observed a plethora of different double-bow extensions in vitro (see Fig. 6 B and D), all of which should not exist according to this mechanism. The third mode of assembly involving the formation of more or less extended complexes is the only one compatible with our observations as well as experimental evidence showing that spindles are more resistant to guanidine than any other conformation, both fragmented and extended. By postulating the ‘‘double clamp,’’ we offer a plausible structural explanation for the experimentally observed stabilization of spindles. The dimerto-dimer interfaces of the terminal tetramers are reciprocally stabilized; moreover, additional stabilization is probably gained through the increased conformational rigidity resulting from double-bow formation (see also Movie 1, which is published as supporting information on the PNAS web site, for the proposed assembly mechanism of TPPII). Although we now seem to understand why discrete spindles are the preferred conformation of TPPII in vivo, the relevance of the enormous particle size for its role in the degradation machinery is unclear. The smallest active unit of human TPPII has been described as the dimer, which has approximately one 10th of the specific activity of the oligomer (36), although there is one report stating that the monomeric subunits might be active as well (37). Our own data obtained with heterologously expressed TPPII corroborate these results and reveal a similar relative activity of dimers (G.S., unpublished data). Thus, in vivo activation of TPPII seems to proceed by dimerization, in analogy to the serine protease HCMG (38), followed by assembly into strands and spindles as sketched above. PNAS 兩 July 19, 2005 兩 vol. 102 兩 no. 29 兩 10139

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Fig. 6. The MBP tag abolishes double-strand assembly. (A) TPPII with an N-terminal MBP tag expressed in E. coli and separated on a Superose 6 column. (B) Cleavage of the MBP tag with proteinase K, subsequent precipitation with ammonium sulfate, and separation on a Superose 6 column. (C) Treatment of the sample shown in B with guanidine and subsequent separation on a Superose 6 column. (D) Extended TPPII complexes obtained after concentrating TPPII that was expressed in E. coli. (E) SDS-gel monitoring the MBP cleavage. M, marker; lane 1, TPPII expressed in E. coli; lane 2, TPPII expressed in E. coli and treated with proteinase K; lane 3, MBP-tagged TPPII treated with proteinase K; lane 4, MBP-tagged TPPII.

active sites, and at the same time it might protect the substrates from their complete hydrolysis in the cytosol. A similar protective action has been proposed for human TPPII in peptide trimming (41). A more detailed picture of the structure and interactions of the subunits is indispensable for fully understanding how the activity of TPPII depends on its oligomeric state. Thus, we will need to improve the resolution of the reconstruction to accurately localize the active site, which will be a crucial step toward fully understanding the assembly and activation of TPPII and, eventually, the flow of substrates through this fascinating giant protease complex.

TPPII is most active when assembled into single- or doublestranded oligomers. Between these two forms no difference in specific activity was observed, but compared to double strands, single strands are unstable and tend to dissociate. Whether the intriguing shape of TPPII confers more than stability remains to be seen. TET, a functional TPPII homologue from the archaeon Pyrococcus horikoshii, is a tetrahedron-shaped hexamer of dimers. It was suggested that its four narrow entry channels provide a unique mechanism of substrate attraction and orientation (29). Aside from the subtilisin-like catalytic domain, no similarities to proteins other than TPPII homologues have been detected in the database thus far (39, 40), and at the present resolution we cannot dock the crystal structure of subtilisin into our density map of TPPII unambiguously. Still, based on the data available, we propose that the catalytic domain of TPPII is localized within the globular domains that form the inner backbone of the strands. The linear stacking of the dimers into strands aligns the handles to form a channel or an arcade (Fig. 5), and it is tempting to speculate that access to the active sites occurs through the lateral openings. In fact, there is no obvious reason for channeling the relatively small substrates of TPPII longitudinally through the strands; on the contrary, it might even prevent optimal substrate processing. However, the arcade might enable a lateral substrate flow, leading to the saturation of all

We thank Brigitte Ku ¨hlmorgen for recording the images of negatively stained tripeptidyl peptidase II and Vesna Olivieri for recording scanning transmission electron microscopy images of tripeptidyl peptidase II. We also thank Andreas Engel for discussions and enabling the scanning transmission electron microscopy. Furthermore, we thank Julio Ortiz for preparation of the movie for supporting information. Initial calculations for this project were carried out at the National Energy Research Scientific Computing Center at the Lawrence Berkeley National Laboratory. Part of the work was funded by a Feodor Lynen Fellowship of the Alexander von Humboldt Foundation (to B.R.), National Institutes of Health Grant GM062989 (to R.M.G.), Swiss National Foundation Grant NF 31-59 415.99 (to Andreas Engel), and the Maurice E. Mu ¨ller Foundation of Switzerland (to S.A.M.).

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