A family of about 40 related proteins make up the intermediate filaments (IFs), ()a major class of cytoskeletal elements in most eukaryotic cells. There are several types of IF proteins, based on protein and gene structure: Types I and II are the keratins, which form IFs in epithelial cells; vimentin and desmin are Type III IF proteins, as are peripherin and glial fibrillar acidic protein, two proteins found in the nervous system. Vimentin is expressed in cells of mesenchymal origin, and desmin is muscle-specific. The neurofilament triplet proteins, NF-L, NF-M, NF-H, and -internexin, are found only in neurons and are Type IV proteins. Type V IF proteins are the nuclear lamins and Type VI consists of a single protein, nestin. Several new IF proteins of limited tissue distribution, filensin, phakinin, and tanabin, also have been identified recently (reviewed by Fuchs and Weber(1994) and Klymkowsky (1995)). The unifying structural principle of this family of proteins is the presence of a tripartite motif, a central, 310-residue long, -helical rod domain and flanking non--helical head and tail domains.

The assembly of 10-nm wide filaments from these highly asymmetric molecules is a complex process. Because the protein molecules are extended, and are packed in a roughly parallel (or anti-parallel) but staggered fashion within the filament, a large number of potential interactions between adjacent molecules is possible. Only a few of these interactions have been identified or characterized. Another complicating factor is that, although several intermediate assembly steps, e.g. the formation of dimers and tetramers, have been identified experimentally (reviewed by Parry and Steinert(1992) and Stewart(1993)), recent detailed structural studies (Steinert, 1991b) suggest that a continuum of oligomers exists beyond the dimer stage. This effectively increases the number of possible protein-protein interactions that may occur within an assembled IF.

The most well known interaction during IF assembly is the lateral association of the -helical rod domains via hydrophobic interactions to form dimeric coiled-coils. This association is thought to lead to formation of the filament backbone, and is made possible by the interaction of hydrophobic amino acids located at the first and fourth positions of a 7-residue repeat present in the primary sequence of the rod domain. Additionally, the rod domain contributes to filament structure on a higher level of organization. The external surface of the dimeric coiled-coil exhibits alternate zones of positive and negative charges (Parry et al., 1977; McLachlan and Stewart, 1982), much like those found in other -helical proteins such as the myosin rod (Atkinson and Stewart, 1992) and tropomyosin (Hitchcock-DeGregori and Varnell, 1990). Recent experimental evidence, obtained by a deletion mutagenesis approach, strongly suggests that electrostatic interactions involving these charged zones are a major driving force for the formation of tetramers and/or higher order structures (Meng et al., 1994).

Additionally, less well recognized interactions involving the head and tail domains also contribute to IF assembly (Birkenberger and Ip, 1990; Eckelt et al., 1992; Herrmann et al., 1992; Makarova et al., 1994; Rogers et al., 1995). It is generally agreed that the head domain of IF proteins is essential for filament assembly, and it has been demonstrated in numerous studies that experimental removal of this domain results in assembly incompetence. Involvement of the tail domain is a matter of some controversy. While it seems clear that removal of the tail does not appreciably hinder in vitro assembly of certain IF proteins, in other cases it has been reported to result in structural aberrations in vitro, in abortive assembly in cells (Kaufmann et al., 1985; Quinlan et al., 1989; Makarova et al., 1994; Bader et al., 1991), or in unusual localization of the IF protein to the nucleus (Rogers et al., 1995).

The higher the level of filament organization, the less is known about how oligomeric subunits are arranged within the filament, and what forces drive their formation. For example, although there is strong evidence indicating that two -helices form a coiled-coil dimer by interactions between hydrophobic residues, what causes the monomers to be aligned in-register and in parallel is not known. Also not yet identified are the forces (and sequences) that drive the formation of tetramers, and filament elongation. At which level of organization does a subunit stagger occur to generate the characteristic 22-nm repeat seen in all IFs? How are IF assembly and disassembly regulated in vivo? While it is widely believed that these interactions are functions of individual IF protein domains, testing of this hypothesis has been primarily carried out by a limited number of approaches, in vitro mutagenesis to look for loss of function in transfected cells or transgenic animals, or in vitro approaches such as reassembly and chemical cross-linking studies. Given that significant differences have been observed between the assembly behavior of some IF proteins in vivo and in vitro (Raats et al., 1991; Eckelt et al., 1992; McCormick et al., 1993), it seems desirable that these interactions be studied in an alternative context. In this paper, we describe our recent efforts in identifying such interactions that are involved in the assembly and maintenance of IF structure, using the two-hybrid system (Fields and Song, 1989). Our results have allowed us to examine more closely certain interactions that have been suggested to occur by current structural models.

MATERIALS AND METHODS

Yeast Strain and Media

Yeast Transformation and -Galactosidase Assay

-Galactosidase activity was measured fluorometrically, using 4-methylumbelliferyl -D-galactopyranoside (4-MUG, Fluka) as substrate. Yeast culture samples (100 µl) from MinGal medium were pipetted into a 1.5-ml Eppendorf tube and pelleted in a microcentrifuge at top speed. The supernatants were discarded. The yeast pellets were then frozen in liquid nitrogen for 2 min and thawed at room temperature. 350 µl of reaction buffer (0.01 M phosphate, pH 7.0, 1 mM MgCl, 0.1% bovine serum albumin, 0.1% NaN) and 50 µl of MUG solution (0.001 M MUG in 0.01 M phosphate buffer, pH 7.0, 1 mM MgCl) were then added to the yeast cells, mixed well, and incubated at 37 °C for 60 min. The reactions were terminated by addition of 400 µl of stop solution (0.1 M glycine, pH 10.3). The amount of 4-MU released from 4-MUG was determined using a Gilford Fluoro IV spectrofluorometer (Gilford, Oberlin, OH), with excitation at 360 nm and emission at 450 nm. Results reported were obtained from at least three independent determinations; variations were less than 20%.

For rapid mass screening for -galactosidase positive yeast colonies, plates were replica-plated onto nitrocellulose membrane, which were then stained with 5-bromo-4-chloro-3-indoyl -D-galactoside (X-gal) (Breeden et al., 1985). On occasion, -galactosidase activity in crude yeast extracts was determined colorimetrically by using O-nitrophenyl--D-galactopyranoside as substrate, as described by Ausubel et al.(1990).

Yeast Vector and Plasmid Construction

All plasmid constructions ( Fig. 1and Table 1) were performed in Escherichia coli strain XL-1 Blue (Stratagene, La Jolla, CA) and bacteria were grown in LB medium. The wild type full-length vimentin cDNA was inserted between the unique SalI and NotI cloning sites of both yeast fusion vectors, pPC62 and pPC86. A sense oligonucleotide, V-P-5 (5`-GCCATGTCGACCAGGT-3`) and the T7 promoter primer (Life Technologies, Inc., Grand Island, NY) in the antisense orientation, were used to amplify the vimentin cDNA and, at the same time, create a unique SalI site at the 5` terminus. After restriction with SalI and NotI and ligation, the plasmids, p62-wtVim and p86-wtVim, were obtained.

Figure 1: Domains of vimentin whose interactions were examined by two-hybrid cloning. wtVim, the full-length vimentin molecule, which consists of a head domain, a rod domain, and a tail domain. VimH, the non-helical head domain; VimH1B, a fragment encompassing the head and the entire helix 1; Vim1B, helix 1B only; Vim2B, helix 2B only; VimT, the non-helical tail domain.

A SalI/PstI fragment encoding the vimentin head domain (amino acids 1-106) was introduced into pPC62 to yield p62-VimH. From this resulting plasmid, a SalI/NotI fragment was excised and subcloned into pPC86 to yield p86-VimH.

All other truncations of vimentin were constructed by polymerase chain reaction-based cloning strategies. For construction of p62-Vim1B, primers V-P-6 (5`-GATAAGTCGACGTACGAGGAGGAGATGCGG-3`) and V-P-7 (5`-GCGCAACTAGTGGCAGCTCCTGGATCTCTTC-3`) were used to amplify the appropriate fragment from BS-Vim. Similarly, a 735-base pair fragment for VimH1B was obtained by amplification using primers V-P-5 and V-P-7. Vim2B, a 363-base pair fragment, was amplified from BS-Vim using a sense primer V-P-8 (5`-GCTTCGTCGACGTACAAGTCCAAGTTTGCT-3`) and an antisense primer, V-P-12 (5`-GCGGA CTAGTAAATCCTGCTCTCCTC-3`). The resulting fragment was subcloned between the SalI and SpeI sites of pPC62 and pPC86 to yield p62-Vim2B and p86-Vim2B, respectively. Finally, VimT, a 165-base pair fragment that encodes the vimentin tail domain, was similarly constructed, using primer V-P-9 (5`-GCTTCGTCGACGTCTCTGCCTCTGCCAACC-3`) (which added a SalI site at the 5` terminus) and the T7 polylinker primer. After restriction with SalI/NotI and ligation into pPC62 and pPC86, p62-VimT and p86-VimT were obtained.

The keratin K8 encoding plasmids, p62-K8 and p86-K8, were obtained by polymerase chain reaction amplification using PUC-K8 as template. The 5` primer, K8-P-10 (5`-GATAAGTCGACGATGTCCATCAGGGTGACC-3`) was used to insert sequence encoding a SalI restriction site 5` to the K8 start site. The 3` primer, the M13(-20) reverse primer (Promega, Madison, WI), adds two restriction sites, XbaI and EcoRI, from the PUC18 vector polylinker, to the 3` end of the keratin cDNA. The SalI/XbaI fragment was then subcloned into pPC62 to obtain p62-K8, and the SalI/EcoRI fragment from the same polymerase chain reaction product was subcloned into pPC86 to obtain p62-K8. Similarly, using BS-K18 as template, primer K18-P-11 (5`-GATAAGTCGACGATGAGCTTCACCACTCGC-3`) and the T3 promoter primer (Life Technologies, Inc.), a polymerase chain reaction fragment containing a 5` unique SalI site, the K-18 coding region, and a 3` XbaI site was obtained. The SalI/XbaI fragment was introduced into pPC62 to yield p62-K18. From this, a SalI/NotI fragment was subcloned into pPC86 to yield p86-K18.

Preparation of Cell Extract and Immunoblotting

Immunodetection on Western blots was performed essentially according to the method of Towbin et al.(1979). A rabbit antiserum, Vim11091, raised against full-length murine vimentin expressed in E. coli, was used as the primary antibody at a dilution of 1:200. Alkaline phosphatase-conjugated affinity-purified, goat-anti-rabbit IgG (Kirkegaard & Perry Laboratories, Gaithersburg, MD) was used as the secondary antibody at a dilution of 1:2000.

RESULTS

Strategy and Controls

We used two vectors, pPC62 (the DB vector) and PPC86 (the TA vector) (Chevray and Nathans, 1992). Complementary DNAs encoding full-length IF proteins or individual domains were subcloned into these vectors, and the fusion plasmids were used pairwise to transform yeast strain PCY2, which carried a genomic copy of the GAL1 promoter driving expression of lacZ. Initially, interactions were detected qualitatively by staining nitrocellulose replicas of yeast plates with 5-bromo-4-chloro-3-indoyl -D-galactoside. Quantitation was carried out by expanding positive yeast colonies and assaying for -galactosidase activity in cell homogenates derived from such colonies, using the fluorescent substrate, 4-MUG. In transformations involving pairs of IF protein domains, the experiments were always repeated by interchanging inserts and vectors, to control for potential steric effects that could result from differences in spatial juxtaposition of the IF protein domain relative to the GAL4 domain when they were expressed as a fusion protein. In most cases, swapping of vectors and inserts in this manner resulted in differences in -galactosidase activity of only 10-20%, but more importantly, it did not alter the strength of the interaction relative to others that were studied.

A series of quantitative controls were undertaken, using MUG fluorescence measurements, to ensure that positive -galactosidase activity truly reflected interactions between IF domains (Table 1). These included co-transformation with one or both vectors lacking insert, and transformation with only one vector. Without exception, such transformations resulted in extremely low levels of -galactosidase activity, the highest of which amounted to 0.13% of that reached by co-transformation with both vectors encoding full-length vimentin (see below).

Full-length IF Proteins Expressed in Yeast Cells Interact Strongly

Figure 2: Yeast cells transformed with p62wtVim and p86wtVim express vimentin protein. Immunoblot analysis of a cytoplasmic homogenate of yeast transformed with the two vectors encoding full-length vimentin as fusion proteins with the TA and DB domains of GAL4. Left lane, molecular mass markers at 205, 94, 68, and 45 kDa. Center lane, PCY2 cell homogenate probed with a polyclonal anti-vimentin antibodies, revealing a reactive product at about 80 kDa, consistent with the size of GAL4-vimentin fusion polypeptides. Right lane, purified recombinant vimentin.

An additional measure of confidence that the IF protein interactions detected by using the two-hybrid were specific was provided by transformation experiments using keratins K8 and K18 (Table 2). Keratin IFs are obligate heteropolymers, i.e. they must contain one Type 1 and one Type 2 keratin. Thus, one would expect interaction to occur only when yeast cells were co-transformed with plasmids encoding one Type 1 and one Type 2 keratins, but not in a co-transformation with both plasmids encoding either Type 1 or Type 2 keratins. This was indeed the case. Strong interaction was detected when co-transformation was carried out with one plasmid encoding K8, a Type 2 keratin, and the other encoding K18, a Type 1 keratin (Table 2, rows 1 and 2), but when both plasmids encoded either K8 or K18 (Table 2, rows 3 and 4), -galactosidase activities fell to about 3% of the K8/K18 co-transformation. When one keratin was omitted from the transformation (Table 2, rows 5 and 6), -galactosidase activity fell to baseline levels. Interestingly, K18, but not K8, interacted quite strongly with full-length vimentin (Table 2, rows 7-10). A similar interaction between an acidic keratin, K14, and vimentin has also been reported (Steinert et al., 1993c). However, as suggested by these investigators, this interaction probably does not proceed beyond the dimer stage and is likely to be non-productive for filament formation.

IFP Dimers and Tetramers Are Both Formed When IF Proteins Interact in Yeast

Results from a sample taken from a co-transformation with both plasmids encoding helix 1B is shown in Fig. 3, lane 2. Two immunoreactive bands, with molecular mass of 55 and 100 kDa are found. The former has a size appropriate for a DB-Vim1B/TA-Vim1B dimer, while the latter has a size appropriate for a tetramer. A similar sample, from a co-transformation with helix 2B-encoding vectors, is shown in lane 3 of Fig. 3. Again, immunoreactive bands of sizes compatible with DB-Vim2B/TA-Vim2B dimers and tetramers are evident. Thus, we conclude that the -galactosidase activity we detected in our transformations was indicative of interactions between IF proteins at the level of both the dimer and the tetramer. We do not have direct information as to whether our GAL4 domain-IF fusion proteins further assemble into 10-nm filaments in PCY2 cells; however, because the fusion proteins contain sizable amounts of TA or DB domains (110 and 144 residues, respectively), which are probably sufficiently large to interfere sterically with subunit packing during filament assembly, it is unlikely that filaments as such are formed.

Figure 3: Dimers and tetramers are both formed when IF proteins interact in PCY2 cells. Immunoblot analysis of two cytoplasmic homogenates of PCY2 cells transformed such that they expressed vimentin helix 1B-GAL4 (lane 2) and helix 2B-GAL4 fusion proteins (lane 3). Lane 1, molecular mass markers at 205, 94, 68, and 45 kDa. Lane 4, Commassie Blue-stained gel of the homogenate used for lane 2; that used for lane 3 was virtually identical. The homogenates were cross-linked with glutaraldehyde before immunoblotting to stabilize dimers and tetramers. The lower bands in lanes 2 and 3 are dimers of the fusion proteins and the upper bands are tetramers of the same species.

Interactions between IF Protein Domains

Several co-transformations that resulted in low levels of -galactosidase activity are also noteworthy. Among these are ones to screen for interaction between helices 1 and 2B, predicted to overlap in several models of IF structure (Stewart et al., 1989; Geisler et al., 1992; Heins et al., 1993; Steinert et al., 1993a). When co-expressed in PCY2 yeast cells, neither helix 1B alone, nor domain H1B, encompassing the entire amino half of the vimentin polypeptide, interacted significantly with helix 2B, which contained the most highly conserved COOH-terminal end of the rod domain (Table 3, rows 5-8). The level of -galatosidase activity attained in these transformations was less than 1% of that produced by homotypic interaction between full-length vimentin polypeptides. Also of considerable interest is the apparent lack of interaction between vimentin head domains, between tail domains, and between head and tail domains (Table 3, rows 9-18). Interaction among these pairs of domains resulted in 2% or less of the activity attained by interaction between full-length vimentin polypeptides.

DISCUSSION

Specificity and Utility of the Approach

It is also worth noting that this approach complements other methods of structural analysis in specific ways. While sequencing of cross-linked protein fragments provides information concerning nearest neighbors within a filament, whether or not such neighbors in fact interact to effect filament assembly cannot be determined by this approach. In contrast, a two-hybrid screen gives a positive readout only if the two protein domains under study actually interact. Likewise, two-hybrid analyses complement DNA transfection studies because of its ability to specify the oligomeric state of the IF proteins when the interaction takes place, whereas in transfection studies it is only possible to look for the end point of the interaction, i.e. whether the proteins, modified or not, can assemble into a filamentous network in cells.

Implications Concerning IF Structure

Figure 4: Models of IF structure and domain alignments specified by them. This figure is redrawn from Geisler et al.(1992) and Steinert et al. (1993a, 1993b, 1993c) to illustrate the domain alignments specified by current IF models. G is the model of Geisler et al.(1992) and A, A, A, and A are the alignments of Steinert et al. The model of Heins et al.(1993) is closely similar to the ones presented here. Domains shaded in each alignment are ones whose interaction was examined by two-hybrid cloning. The high level of interaction between helices 1B and between helices 2B to form tetramers, revealed by two-hybrid cloning, is compatible with the hypothesis that the G, A, and A alignments are the consequence of true domain binding. Conversely, the lack of interaction between helix 1A and helix 2B suggests that the A and A alignments do not contribute to forces that hold subunits together within a filament.

An important question arises when considering these structural data: which of the domain overlaps represent actual protein-protein interactions that bring the molecules together to form the filament, and which of them represent nearest neighbors that were identified as a result of the proximity of cross-linkable lysine and/or cysteine residues within their sequences? Because we were able to determine whether or not given pairs of IF protein domains interact in a cytoplasmic context, it is possible to examine this question in some detail.

Transformation with both vectors encoding helix 1B-GAL4 fusion proteins produced high levels of -galactosidase activity (Table 3, rows 2 and 3), indicating that helices 1B bind strongly to one another. Accompanying Western blotting experiments showed that helices 1B form tetramers, consistent with the aforementioned observation that helix 1 tetramers could be isolated from proteolyzed wool keratin (Gruen and Woods, 1981; Woods and Inglis, 1984). Thus, we conclude that helix 1B is a site along the molecule at which active interaction occurs to stabilize the structure of an IF. Interestingly, the amino-terminal halves of vimentin, encoded by the VimH1B construct, interacted considerably more strongly than helices 1B alone. As the head domain did not interact with either another head domain or the rod (Table 3, rows 9-12), the strength of the homotypic interaction between the amino-terminal halves of vimentin likely stems from the presence of helix 1A, which might have enhanced molecular alignment. An additional transformation using a helix 1 (1A plus 1B) construct should provide a definitive answer.

In light of the fact that a tetramer composed of helix 2 has not been isolated, the finding that helices 2B interacted strongly and produced tetrameric complexes in the two-hybrid system was somewhat surprising. Nonetheless, this interaction is consistent with the A alignment of Steinert et al. (1993a, 1993b), and is a logical consequence of the A interaction discussed above. This is because the alignment of helices 1B between two rows of dimers as specified by the A interaction would necessarily place their helices 2B in close proximity also (see Fig. 4). The larger size of helix 2, and the presence of more proteolytic cleavage sites, may account for the failure to detect intact tetramers composed of helix 2. The fact that both helix 1B and helix 2B exist as tetramers does not allow one to discriminate between the anti-parallel staggered alignment of dimers, favored by all three models of IF structure, and the parallel unstaggered alignment not favored by any. However, it does argue against active interaction between dimers arranged in an A alignment, because this would have resulted in dimers but not tetramers in an interaction screen.

A second result that suggests a lack of true interactions between dimers in the A and A types of alignment is the low levels of -galactosidase activity produced by transformations using the relevant constructs (Table 3, rows 7 and 8). VimH1B, which encompassed essentially the amino terminus half of the vimentin polypeptide and includes helix 1A, did not interact with Vim2B, which encompasses the rod's carboxyl terminus, as would have been predicted by the A as well as the A modes of alignment (Fig. 4). Although there is little doubt that these regions of adjacent vimentin molecules are situated very close to one another within the filament, as evidenced by the positional data provided by chemical cross-linking (Steinert et al., 1993c), it would seem that the cross-links represent juxtaposing, but not interacting, regions of the molecules.

Finally, equally interesting was the observation that interactions involving the vimentin end domains were rather minimal (Table 3, rows 9-18). In the two-hybrid system, the end domains formed neither homodimers nor heterodimers, suggesting that they do not bind one another to contribute to filament structure in vivo. Thus, it may be inferred that growth in filament length and girth, and stabilization of filament structure, are functions of overlap of the rod alone. Also of some interest is that neither end domain binds to the full-length vimentin molecule avidly (Table 3, rows 9 and 14). Thus, the binding of tail sequences to other parts of the IF protein molecule as a mechanism of modulating filament assembly, suggested by several investigators including ourselves (Eckelt et al., 1992; Kouklis et al., 1991, 1993; McCormick et al., 1993; Makarova et al., 1994; Rogers et al., 1995), is likely to be a transient or weak interaction, or perhaps one that depends on post-translational modification not found in yeast.

FOOTNOTES

*

This work was supported in part by Grant CB104 from the American Cancer Society and National Institutes of Health Grant AR35973. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§

Supported by an Established Investigator Award from the American Heart Association(1989-1994). To whom correspondence should be addressed: Dept. of Cell Biology, Neurobiology, and Anatomy, University of Cincinnati College of Medicine ML0521, 231 Bethesda Ave., Cincinnati, OH 45267-0521. Tel.: 513-558-3614; Fax: 513-558-4454.

()

The abbreviations used are: IF, intermediate filament: MUG, 4-methylumbelliferyl -D-galactopyranoside; DB, DNA binding domain; TA, transcription activating domain.

ACKNOWLEDGEMENTS

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