created: 20th January 1998, last updated: 26th May 1998,© 1998 ABRF 

Protein-Fragment Complementation Assays: A General Strategy for the in vivo Detection of Protein-Protein Interactions

Joelle N. Pelletier, Ingrid Remy and Stephen W. Michnick*

Département de biochimie, Université de Montréal, Montréal, Qué. Canada H3T 1J4

*To whom correspondence should be addressed: michnick@bch.umontreal.ca

Telephone: (514) 343-5849

Fax: (514) 343-2210

 


Key words: dihydrofolate reductase, ubiquitin, leucine zipper, enzyme reconstitution, protein design.


ABSTRACT

 

We describe a strategy for designing protein-fragment complementation assays to detect protein-protein interactions in vivo. The design and implementation of this strategy is illustrated with two assay systems: one based on the enzyme dihydrofolate reductase (DHFR) and implemented in Escherichia coli ; and a second, the ubiquitin „split protein¾ sensor (USPS), which is demonstrated in a transiently transfected mammalian cell line, COS-7. In the DHFR-based assay, N- and C-terminal fragments of murine DHFR fused to GCN4 leucine zipper sequences were coexpressed in E. coli. where the endogenous DHFR activity was inhibited with trimethoprim. Coexpression of the complementary fusion products restored colony formation. Survival only occurred when both DHFR fragments were present and contained leucine-zipper forming sequences, demonstrating that reconstitution of enzyme activity requires assistance of leucine zipper formation. Activity was then characterized in vitro.

 

In USPS, oligomerization domain-assisted reassembly of ubiquitin from fragments results in susceptibility to ubiquitin-specific proteases that recognize the intact ubiquitin structure and cleave between the C-terminus of ubiquitin and a fused reporter protein (DHFR used as reporter). We demonstrate both equilibrium and kinetics of this reassembly by transient expression of ubiquitin fragments fused GCN4 leucine zipper sequences in COS-7 cells. Fragment-interface point mutations, implemented in both systems, resulted in a sequential decrease in signal, demonstrating specific fragment reassembly.

 

Replacement of the leucine zippers with other proteins or protein domains, or with expression libraries, will allow for the study of equilibrium and kinetic aspects of protein-protein interactions and for screening cDNA libraries for binding of a target protein with unknown proteins.

 

 

INTRODUCTION

 

Many processes in biology, including transcription, translation, and metabolic or signal transduction pathways, are mediated by non-covalently-associated multienzyme complexes (1). Much of modern biological research is concerned with identifying proteins involved in these cellular processes, determining their functions and how, when, and where they interact with other proteins involved in specific pathways. Further, with rapid advances in genome sequencing projects there is a need to develop strategies to define „protein linkage maps¾, detailed inventories of protein interactions that make up functional assemblies of proteins (2-4). Despite the importance of understanding protein assembly in biological processes, there are few convenient methods for studying protein-protein interactions in vivo (5, 6). A powerful and commonly used strategy, the yeast two-hybrid system, is used to identify novel protein-protein interactions and to examine the amino acid determinants of specific protein interactions (5, 7-9). The approach allows for screening of cDNA libraries, or mutants of individual genes. However, this and other existing methods for in vivo detection require additional cellular machinery that exist only in specific cellular compartments, and generally cannot be used to distinguish between induced versus constitutive interactions.

 

An alternative to these approaches is the oligomerization-assisted complementation of fragments of monomeric proteins, and detection of fragment reassembly in vivo. The first example of this approach, the ubiquitin-based split-protein sensor (USPS) was reported (10) and we have recently developed an assay based on similar principles using the enzyme murine dihydrofolate reductase (mDHFR) (11). Fragmentation of proteins of known structure allows for design of fragments that will reassemble successfully. The flexibility allowed in the design of this approach will make it applicable to situations where other detection systems may not be suitable.

 

We first present the oligomerization-assisted complementation of fragments mDHFR as an example of functional re-assembly of fragments into active enzyme. We have designed two fragments of mDHFR, each of which is covalently attached to an oligomerizing domain (leucine zipper). We demonstrate, by an E. coli survival assay, that leucine zipper interaction is necessary for detecting re-assembly of catalytically active mDHFR from its fragments. Subtle changes in enzyme structure can be detected by this method, specifically point mutations at the interface between the two assembled fragments. Finally, the in vitro activity of the reassembled enzyme is examined. Since leucine zipper formation is crucial to detecting protein-fragment reassembly in the protein-fragment complementation assay (PCA) described here, we propose that the strategy could be used to detect other protein-protein interactions. The leucine zippers would be replaced by interacting proteins of interest for detection of association in vivo. Further, the selection and design criteria applied here to DHFR could be used to develop a strategy using any enzyme that would be appropriate for a clonal selection or direct detection of protein-protein interactions.

 

We then present results of implementation of USPS to mammalian cells, to illustrate that PCAs can be easily implemented in a variety of cell types regardless of their origin. The strategy is based on cleavage of reporter proteins fused to the C-terminal of ubiquitin by cytosolic proteases (ubiquitinases) that recognize and specifically bind to ubiquitin. The enzymes will cleave the reporter protein from the C-terminal of ubiquitin only if the structure of ubiquitin is intact. A fusion of a reporter protein-ubiquitin C-terminal fragment can also be cleaved by ubiquitinases, but only if co-expressed with an N-terminal fragment of ubiquitin that is complementary to the C-terminal fragment. Furthermore, reconstitution of observable ubiquitinase activity only occurs if the N- and C-terminal fragments are bound through GCN4 leucine zipper-forming sequences [Johnsson, 1994 #99; this study]. As well as offering a new approach for the study of kinetic and equilibrium aspects of protein-protein interactions in vivo, the assay could also be adapted for detection of protein-protein interactions in a variety of cell types. Furthermore, this technique would allow for the detection of induced (such as those initiated by cell growth or inhibitory factors) versus constitutive protein-protein interactions, in mammalian cells.

General Features of PCAs and their Design.

 

The general features of a PCA are illustrated in Figure 1. In designing a protein-fragment complementation assay (PCA), we first identify a protein for which the following is true: 1) it is relatively small and monomeric, 2) information exists on its structure and function, 3) simple assays exist for both its in vivo and in vitro measurement, and 4) its overexpression in either eukaryotic, prokaryotic or both types of cells has been demonstrated. The gene coding for the target protein is then rationally cleaved into two or more fragments, according to the protein structure, so as to create subdomains; the resulting new N- and C- termini are located on the same face of the protein, in order to avoid the need for long peptide linkers, and to allow for determination of the orientation-dependence of the protein-protein interaction under study. The proteins of interest are then fused to each of the created fragments. The resulting fusion proteins are coexpressed in an appropriate cellular context and an in vivo assay performed to detect oligomerization-assisted complementation of the protein fragments.

 

Figure 1: General Features of a PCA. Top, left. An enzyme can be transformed/transfected into a host cell and its activity detected by an in vivo assay (right). Bottom, left. Oligomerization domains A, B, are fused to N- and C-terminal fragments of the gene for the enzyme. Co-transformation/transfection of oligomerization domain-fragment fusions results in reconstitution of enzyme activity by oligomerization domain-assisted reassembly of the enzyme (right). Reassembly of enzyme will not occur unless oligomerization domains interact.

 

RESULTS AND DISCUSSION

 

Section I: Design and Implementation of a Bacterial PCA: the DHFR PCA.

Murine dihydrofolate reductase (mDHFR) meets all the criteria listed above: it is a small (21 kD), monomeric protein of known structure (12). The folding, catalysis, and kinetics of a number of DHFRs have been studied extensively (13). DHFR activity can be monitored in vivo by cell survival in cells grown in the absence of DHFR end-products, and in vitro. In our in vivo assay we took advantage of the fact that E. coli DHFR is selectively inhibited by the anti-folate drug trimethoprim. As mammalian DHFR has a 12,000-fold lower affinity for trimethoprim than does bacterial DHFR (14), growth of bacteria expressing mDHFR in the presence of trimethoprim levels lethal to bacteria is an efficient means of selecting for reassembly of mDHFR fragments into an active enzyme.

 

Design Considerations. Comparison of the crystal structures of mDHFR and human DHFR (hDHFR) suggests that their active sites and substrate binding pockets are identical, and homologous to those of E. coli DHFR (12, 15). DHFR has been described as comprising three structural fragments forming two domains: the adenine binding domain (F[2]) and a discontinuous domain (F[1] and F[3]) (16, 17). The folate binding pocket and the NADPH binding groove are formed mainly by residues belonging to F[1] and F[2]. Residues in F[3] contribute little to substrate binding and catalysis (15), but do contribute to DHFR stability and to kinetic parameters (18, 19). We have designed two fragments of mDHFR consisting of the sequences coding for F[1] and F[2] (referred to as F[1,2]), and for F[3] so as to cause minimal disruption of the active site and NADPH cofactor binding sites. Cleavage of mDHFR at the loop that is the junction between F[2] and F[3] and fusion of the native termini has produced a circularly permuted protein with physical and kinetic properties very similar to the native enzyme, suggesting that radical modifications in this loop are not disruptive to activity (20). The native N-terminus of mDHFR and the novel N-terminus created by cleavage occur on the same surface of the enzyme (12, 15) facilitating N-terminal covalent attachment of each fragment to oligomerization domains such as the leucine zippers (Zip) used in this study. The resulting fusions are labelled Zip-[1,2] and Zip-[3].

 

E. coli Survival Assays. Cotransformation of bacteria with constructs coding for Zip-[1,2] and Zip-[3] (as in Figure 1) was undertaken with the goal of detecting enzymatic activity from reconstituted mDHFR. Figure 2 (panel A) illustrates the results of cotransformation in the presence of trimethoprim, clearly showing that colony growth under selective pressure is possible only in cells expressing both fragments of mDHFR. There is no growth in the presence of either Zip-[1,2] or Zip-[3] alone. Furthermore, induction of protein expression (with IPTG) is essential for colony growth (Fig. 2A). The presence of the leucine zipper on both fragments of mDHFR is essential as illustrated by cotransformation of bacteria with both vectors coding for mDHFR fragments, only one of which carries a leucine zipper (Fig. 2A). Further experiments were performed to control for absence of activity by Zip-[1,2] in the presence of F-[3] (the reciprocal of Control-[1,2] + Zip-[3]), as well as for the effect of variables including changes in expression level, solubility and susceptibility to proteolysis. The results are all satisfactory and will be published elsewhere (J. Pelletier, K. Arndt, A. Pl¸ckthun and S. Michnick, in preparation). It should be noted that growth of control E. coli transformed with the full-length mDHFR is possible in the absence of IPTG due to low levels of expression in uninduced cells.

 

Figure 2: (A) E. coli survival assay on minimal medium plates. Control: Left side of the plate: E. coli harboring pQE-30 (no insert); right side: E. coli harboring pQE-16, coding for wild-type mDHFR. Panel I: Left side of each plate: transformation with construct Zip-[1,2]; right side of each plate: transformation with construct Zip-[3]. Panel II: Cotransformation with constructs Zip-[1,2] and Zip-[3]. Panel III: Cotransformation with constructs Control-[1,2] and Zip-[3]. All plates contain 0.5 microg/ml trimethoprim. In panels I to III, plates on the right side contain 1mM IPTG.

(B) E. coli survival assay using destabilizing DHFR mutants. Panel I: Cotransformation of E. coli with constructs Zip-[1,2] and Zip-[3:Ile114Val]. Panel II: Cotransformation with Zip-[1,2] and Zip-[3:Ile114Ala]. Inset is a 5-fold enlargement of the right-side plate. Panel III: Cotransformation with Zip-[1,2] and Zip-[3:Ile114Gly]. All plates contain 0.5 microg/ml trimethoprim. Plates on the right side contain 1mM IPTG.

 

Stability Mutants: We generated point mutants of F[3] to change the efficiency with which the mDHFR fragments assemble to form active enzyme, as tests for specific reconstitution of protein fragments. Protein stability can be reduced by changing the side-chain volume in the hydrophobic core of a protein (10, 21-24). Residue Ile114 of mDHFR occurs in a core beta-strand at the interface between F[1,2] and F[3], isolated from the active site. Ile 114 is in Van der Waals contact with Ile51 and Leu93 in F[1,2] (15). We made point mutations of the wild-type mDHFR Ile 114 to Val, Ala, or Gly. Figure 2 (panel B) illustrates the results of double transformation of E. coli with construct Zip-[1,2] and the mutated constructs Zip-[3:Ile114Val], Zip-[3:Ile114Ala] or Zip-[3:Ile114Gly] in the presence of trimethoprim. The colonies obtained from cotransformation with Zip-[3:Ile114Ala] grew more slowly than those cotransformed with Zip-[3] or Zip-[3:Ile114Val] (see inset to Fig. 2B). No colony growth was detected in cells cotransformed with Zip-[3:Ile114Gly]. Overexpression of the mutants Zip-[3:Ile114X] was in the same range as Zip-[3], as determined by Coomassie-stained SDS-PAGE (data not shown).

 

In order to compare the relative efficiency of reassembly of mDHFR fragments, we measured the doubling time in liquid medium of E. coli transformed with the appropriate constructs. Doubling time of E. coli in minimal medium was essentially constant for all transformants (Table 1). Selective pressure by trimethoprim in the absence of IPTG, and induction of mDHFR fragment expression with IPTG, reflect the results obtained on solid media. The doubling time measured for cells expressing Zip-[1,2] + Zip-[3], Zip-[1,2] + Zip-[3:Ile114Val] and Zip-[1,2] + Zip-[3:Ile114Ala] were 1.6-fold, 1.9-fold and 4.1-fold higher, respectively, than the doubling time of E. coli expressing wild-type mDHFR (pQE-16) in the absence of trimethoprim and IPTG. The presence of IPTG unexpectedly prevented growth of E. coli transformed with wild-type mDHFR. Growth was partially restored by addition of the folate metabolism end-products thymine, adenine, pantothenate, glycine and methionine (data not shown). This suggests that induced overexpression of mDHFR was lethal to E. coli when grown in minimal medium as a result of depletion of the folate pool by binding to the enzyme. However, the amount of active reconstituted enzyme in cells is too low to allow for toxicity of reconstituted enzyme. *Table 1*

 

The sequential increase in cell doubling times resulting from the mutations directed at the assembly interface (Ile114 to Val, Ala or Gly) demonstrates that the observed cell survival under selective conditions is a result of the specific, leucine-zipper-assisted association of mDHFR fragment[1,2] with fragment[3]. Since Zip-[1,2] carries most residues involved in substrate binding and activity, it is possible that reconstituted DHFR activity could be due to non-specific interactions with Zip-[3]. However, the point mutants illustrate that cell survival is a result of correct mDHFR fragment reassembly rather than nonspecific interactions of Zip-[3] with Zip-[1,2].

 

Methotrexate-resistant mutants: As a further control for understanding the molecular basis for the reassembly of mDHFR fragments into active enzyme, we mutated fragment[1,2] in order to incorporate, one at a time, each of five mutations which have previously been shown to significantly increase Ki (methotrexate): Gly15Trp, Leu22Phe, Leu22Arg, Phe31Ser and Phe34Ser (numbering according to the wild-type mDHFR sequence) (25-30). These mutations occur at varying positions relative to the active site and relative to fragment [3], and have varying effects on Km (DHF), Km (NADPH) and Vmax of the full-length mammalian enzymes in which they were studied. Mutants Zip-[1,2:Leu22Phe], Zip-[1,2:Leu22Arg] and Zip-[1,2:Phe31Ser] all allowed for bacterial survival with high growth rates when cotransformed with Zip-[3] (Table 1). This demonstrates that mutations in fragment[1,2], which essentially encompasses the substrate and inhibitor binding pocket and the active site residues, can be well tolerated and allow for very efficient fragment complementation. Mutants Zip-[1,2:Gly15Trp] and Zip-[1,2:Phe34Ser] did not allow for bacterial survival. The position of Gly15 readily explains this observation: it occurs at the interface between fragments [1,2] and [3], and its mutation appears to destabilize the fragment assembly, reinforcing the importance of the interface interactions in fragment complementation. The reason for lack of growth with mutant Phe34Ser is not so obvious; the 24-fold increase in Km (DHF) observed in the wild-type hDHFR (29) may be responsible for this.

 

Kinetic parameters of the reconstituted enzyme: In vitro activity was assayed following native purification of the reconstituted enzyme: Zip-[1,2:Phe31Ser] + Zip-[3]. Figure 3 illustrates that the rate of turnover, measured by fluorescence emission of the product (THF), is inhibited by increasing concentrations of methotrexate. The Ki(methotrexate) for the reconstituted enzyme was determined to be 0.7 nM (average of 2 independent determinations) at 37 deg C, pH 7.5, using 30 microM DHF and 25 microM NADPH. This value is very similar to Ki(methotrexate) of 4.4 nM for mDHFR(Phe31Ser), determined at 30 deg C, pH 7.9, using 50 microM DHF and 100 microM NADPH (28). Furthermore, the inhibition by methotrexate retains its competitive character relative to dihydrofolate (data not shown). These results suggest that the kinetic behavior of the reconstituted enzyme closely resembles that of the full-length mDHFR(Phe31Ser).

 

Figure 3: Inhibition of the reconstituted mDHFR(Phe31Ser) activity by methotrexate. Semi-logarithmic plot illustrating the inhibitory effect of increasing methotrexate (MTX) concentrations on THF production (measured as fluorescence emission) by the reconstituted enzyme. The blank (no enzyme) was subtracted at each data point.

 

 

Section II: A Mammalian PCA, the Ubiquitin Split-Protein Sensor.

 

We demonstrate the application of the ubiquitin-based split-protein sensor (USPS) (10) for the detection of protein-protein interactions in the mammalian COS-7 cell line, and illustrate its use in the study of both equilibrium and kinetic aspects of protein-protein interactions in vivo. The strategy is based on cleavage of a reporter protein (hemagglutinin (HA)-tagged mDHFR) from the C-terminus of ubiquitin, to which it is fused, by cytosolic ubiquitinases. These proteases will release the reporter protein only if the structure of ubiquitin is intact. A fusion of a reporter protein-ubiquitin C-terminal fragment can also be cleaved by ubiquitinases, but only if co-expressed with an N-terminal fragment of ubiquitin that is complementary to the C-terminal fragment. Furthermore, as in the DHFR PCA, the two ubiquitin fragments will not recombine efficiently unless oligomerization domains are introduced at the ends of the complementary fragments; the oligomerization domains act as a template for the reassembly of ubiquitin. We demonstrate this oligomerization-assisted assembly of ubiquitin fragments with GCN4 leucine zipper-forming sequences in COS-7 cells, transiently transfected with leucine zipper-ubiquitin fragment gene fusions. Constructs consisted of combinations of GCN4 leucine zipper (Zip), ubiquitin N-terminal (Nub) or C-terminal fragments (Cub), and mDHFR-HA (dha) fusions. The Zip-Cub-dha and Nub-Zip fusions were coexpressed in COS-7 cells, where oligomerization was detected by measuring the release of mDHFR-HA from the reassembled ubiquitin.

 

To study the equilibrium reconstitution of ubiquitin structure, cell lysates were analyzed by SDS-PAGE, followed by protein transfer and immunoblotting with anti-HA antibodies (Figure 4A). The positive control Ub-dha is totally cleaved by ubiquitinases. When expressed by itself, Zip-Cub-dha yielded minor non-specific cleavage that was considerably reduced by the replacement of the original linker joining the zipper and Cub (10) by a flexible linker coding for (GlyGlyGlyGlySer)4. However, coexpression of Zip-Cub-dha and Nub-Zip resulted in the complete cleavage of Zip-Cub-dha (release of dha). We have recently replaced the leucine zipper-forming sequences with other interacting protein domains, essentially eliminating the problem of the non-specific cleavage, probably by increasing the stability of the fusion proteins to non-specific proteolytic cleavage or competition for internal initiation at the DHFR start codon (which is intact in our fusions). These results will be reported elsewhere (Remy, Johnsson, and Michnick, in preparation). Point mutations of increasing severity (Ile13 to Val, Ala and Gly) were introduced at the interface between the two ubiquitin fragments (in the hydrophobic core of ubiquitin) with the goal of destabilizing reassembly. The cleavage was incomplete for the mutants Nub(Ile13Ala)-Zip and Nub(Ile13Gly)-Zip (Figure 4A), demonstrating that observed reconstitution of ubiquitinase activity is due to specific refolding of ubiquitin from its complementary fragments. The result for the mutant Ile13Val was comparable to the wild-type Nub.

 

Figure 4: Reconstitution of ubiquitin from its coexpressed fragments.

(A) Western blot of the equilibrium reconstitution of ubiquitin: lane 1 is full-length Ub-dha, lane 2 is Zip-Cub-dha alone, lane 3 to 6 are coexpressions of Zip-Cub-dha with, respectively, Nub-Zip, Nub(Ile13Val)-Zip, Nub(Ile13Ala)-Zip and Nub(Ile13Gly)-Zip. Lane 7 is a mock-transfection. M represents molecular weight markers.

(B) Autoradiograph of [35S]-pulse labelled immunoprecipitation: "-" is Zip-Cub-dha alone, and "I,V,A and G" are coexpressions of Zip-Cub-dha with, respectively, Nub-Zip, Nub(Ile13Val)-Zip, Nub(Ile13Ala)-Zip and Nub(Ile13Gly)-Zip.

 

We also demonstrated that, as in yeast, kinetics of leucine zipper formation can be detected in mammalian cells using USPS. COS-7 cells expressing Zip-Cub-dha by itself or coexpressed with Nub-Zip, Nub(Ile13Val)-Zip, Nub(Ile13Ala)-Zip or Nub(Ile13Gly)-Zip were incubated for 30 min with [35S]methionine, followed by a chase for 0, 10, and 30 min, extraction of proteins, immunoprecipitation with anti-HA antibody, and SDS-PAGE (Figure 4B). Coexpression of Zip-Cub-dha and Nub-Zip resulted in the nearly complete release of dha by the end of a 30 min pulse (time 0). The cleavage was slower when Zip-Cub-dha was coexpressed with Nub(Ile13Val)-Zip, Nub(Ile13Ala)-Zip or Nub(Ile13Gly)-Zip. For the Ala and Gly mutants, the cleavage was still incomplete after a 30 min chase. The rate of release of dha decreased in correlation with the increasing severity of mutations in coexpressed Nub fragments, demonstrating that the rate of ubiquitin reconstitution depends on the stability of the refolded ubiquitin. This approach has been suggested as a way to determine relative rates of assembly of different proteins (10).

 

CONCLUSIONS AND PERSPECTIVES

 

In conclusion, we have developed and implemented a bacterial protein-fragment complementation assay (PCA) based on mDHFR, and have illustrated the application of a ubiquitin-fragment complementation assay in mammalian cells, where interacting leucine zippers direct the reconstitution of the protein fragments in vivo with a concurrent detection of reconstitution of the fragments. In the first case, activity was detected by an E. coli survival assay. In the second case, the mammalian PCA was used to reveal both equilibrium and kinetic aspects of leucine zipper formation.

 

The development and demonstration of the two PCAs presented here illustrate how this approach can be used to design, at a molecular and atomic level of detail, a general strategy to detect protein-protein interactions, and could be applied to the development of other assay strategies based on the same principles. The DHFR PCA is a complete system in that no additional endogenous factors are necessary and one can observe the results of complementation directly, with no further manipulation. The USPS system can be used to monitor protein-protein interactions as a function of time at their natural site of interaction in a mammalian cell line, but it requires enzymes constitutively expressed in the host cell. However, this is not a serious limitation. The ubiquitinases are likely expressed in both cytosol and nucleus and so two significant cellular compartments are accessible to USPS(31, 32). Further, the sub-compartmentalization of the ubiquitinases means that USPS can be used in the context of compartmentalization kinetics studies, as has already been illustrated (33). Having a coupled enzyme activity in a PCA could be advantageous in some cases: in a given PCA, the complementing fragments may not reassemble in such a way as to take on the native structure of the protein, but rather some stable intermediate form, such as a „molten globule¾. In the case of an enzymatic PCA, the native activity of the reconstituted enzyme may not be attained, precluding use of its activity as a detection tool. However, in a PCA where the reassembled protein is recognized by an independently folding enzyme, this accessory enzyme may act upon the reassembled protein even if it is not in its fully native state.

 

The DHFR PCA E. coli cell survival assay described here is perhaps the simplest direct measurement method. There are several aspects of the DHFR PCA that distinguish it from all other methods currently available for studying protein-protein assembly in vivo. From knowledge of protein stability, folding, enzyme kinetics and regulation, it should in principle be possible to design complementary fragments of DHFR that would allow for controlling the stringency of the assay, for obtaining estimates of the kinetics and equilibrium constants for association of two proteins, and for detecting induced versus uninduced protein-protein interactions. In our studies we used point mutations of the wild-type Ile 114 to Val, Ala, and Gly to demonstrate that the observed effects on E. coli survival and cell growth rates are specifically due to reconstitution of DHFR activity. For determining estimates of equilibrium and kinetic parameters for a specific protein-protein interaction one could perform a series of experiments in which two proteins that are known to interact are fused to the N- or C-terminal domains of the DHFR fragments in the wild type or destabilizing mutant forms. One then could perform the DHFR PCA and compare rates of cell growth, or methotrexate binding, with those observed for model protein-protein interactions, to obtain an estimate of the strength of the interaction. To illustrate this, we are presently making point mutations in the GCN4 leucine zipper where direct equilibrium and kinetic parameters are known, and correlating these known values with parameters derived from the PCA (Pelletier, Arndt, Pl¸ckthun and Michnick, in preparation). Another advantage of designing stability mutants of DHFR is to control the stringency of the assay. This will be particularly important in cDNA library screening for protein-protein interactions. By assembling the library as fusions to different stability mutants it may be possible to distinguish weak, non-specific from high-affinity specific interactions. The DHFR PCA should not be limited in the context in which it could be used as was illustrated here for USPS. mDHFR has been expressed in both prokaryotic and eukaryotic hosts, and could be targeted to specific compartments in the cell by addition of signaling peptide sequences. Induced versus constitutive protein-protein interactions could be distinguished by the DHFR PCA, in the case of an interaction that is triggered by a biochemical event. For example, in the case of signal transduction, one could transfect a cell line that responds to a specific hormone by interaction with a cell-surface receptor with the DHFR PCA components fused to proteins thought to interact following hormone-receptor interaction. One then could treat the cells with the hormone and test for reconstitution of DHFR activity.

 

Besides the simple example of peptide oligomerization studied here, we are currently investigating applications of the USPS, DHFR and other PCAs for screening of cDNA libraries for protein-protein interactions. The capability of controlling the stringency of the assays, of detecting interactions in appropriate contexts, and of distinguishing induced vs constitutive interactions, make the PCA approach a promising strategy for determining possible functions of gene products based on their interactions with proteins of known function. The fact that such studies could be done in appropriate cells ( i.e., those in which a particular biochemical pathway is studied), mean that PCAs could be used in conjunction with appropriate functional controls, to ensure that interactions of known proteins with products of a cDNA library may be more likely specific and biologically relevant.

 

MATERIALS AND METHODS

 

Materials. All reagents used were of the highest available purity. Mutagenic and sequencing oligonucleotides were purchased from Gibco BRL. Restriction endonucleases and DNA modifying enzymes were from Pharmacia and New England Biolabs. For bacterial protein overexpression, E. coli strain BL21 carrying plasmid pRep4 (Qiagen, lac Iq) was transformed with the appropriate DNA constructs.

DNA Constructs: 1-DHFR fusions. All final constructs were based on the Qiagen pQE series of vectors, which contain an inducible promoter-operator element (tac), a consensus ribosomal binding site, initiator codon and nucleotides coding for an N-terminal hexahistidine peptide. The full-length mDHFR is expressed from the Qiagen expression control vector, pQE-16. The mDHFR fragments [1,2] and [3] carrying their own in-frame stop codon were produced by PCR using pMT3 (derived from pMT2) (34) as a template, and subcloned into the pQE-32 polylinker at appropriate restriction sites. All final constructs were verified by DNA sequencing. Residues 235 to 281 of the GCN4 leucine zipper (a SalI/BamHI 254 bp fragment) were obtained from a yeast expression plasmid pRS316 harboring that sequence (10) and subcloned 3' to the hexahistidine tag and 5' to the DHFR fragments, yielding constructs Zip-[1,2] and Zip-[3]. The control expression construct: Control-[1,2], codes only for the histidine-tagged mDHFR fragment[1,2] without the zipper.

 

2-Ubiquitin constructs. Ub-dha, Nub-Zip, Zip-Cub-dha fusions and destabilizing mutants of the original yeast expression constructs (10) were introduced in the HindIII (for Ub-dha and Zip-Cub-dha) or HindIII and XbaI (for Nub-Zip) restriction sites of the mammalian expression vector pcDNAI/Amp (Invitrogen), that contains a CMV promoter and enhancer and SV40 and Polyoma virus eukaryotic origin of replication. Ub-dha consisted of the full-length ubiquitin protein fused to the mouse dihydrofolate reductase with an HA epitope tag (DHFR-HA, dha). The Nub-Zip fusions and Zip-Cub-dha fusions consisted of the N- (codons 1 to 37) and C-terminal (codons 35 to 76) portion of ubiquitin, respectively, fused to the residues 235 to 281 of the GCN4 leucine zipper. In order to increase the stability of the fusions proteins in mammalian cells, the linker joining the zipper and Cub was replaced by a flexible linker coding for (GlyGlyGlyGlySer)4, at the XbaI and BamHI restriction sites of Zip-Cub-dha.

 

Creation of Stability Mutants and Methotrexate-resistant Mutants. Site-directed mutagenesis was performed according to the method of Kunkel (35). Mutagenesis reactions were carried out on appropriate DNA fragments subcloned into pBluescript SK+ (Stratagene), using oligonucleotides that encode a silent mutation producing or destroying a diagnostic restriction site. Fragments of putative mutants identified by restriction were subcloned back into the appropriate constructs. The mutations were confirmed by DNA sequencing.

 

E. coli Survival Assay. Competent E. coli BL21/pRep4 were transformed with the appropriate constructs and washed twice with minimal medium before plating on minimal medium plates containing 50 microg/ml kanamycin, 100 microg/ml ampicillin and 0.5 microg/ml trimethoprim. One half of each transformation mixture was plated in the absence, and the second half in the presence, of 1 mM isopropyl-beta-D-thiogalactopyranoside (IPTG). All plates were placed at 37 deg C for 66 hrs.

 

E. coli Growth Curves. Growth curves in liquid medium were performed using minimal medium supplemented with ampicillin, kanamycin as well as IPTG (1 mM) and trimethoprim (1 microg/ml) where indicated. All experiments were performed in triplicate. Aliquots were withdrawn periodically for measurement of optical density. Doubling time was calculated for early logarithmic growth (OD 600 between 0.02 and 0.2).

DHFR-fusion Overexpression and Native Purification. Bacteria were propagated overnight in minimal medium under selective pressure, in a small volume, and used to inoculate a large volume (500 ml to 1 l) at a 100 x dilution. Cells were harvested after 24 hrs and stored at -80 deg C. The cell pellet from 250 ml of E. coli cotransformed with appropriate constructs was lysed by sonication in 10 ml of buffer A (100 mM potassium phosphate pH 8.0, 1 mM PMSF, 10 mM beta-mercaptoethanol). The lysate was clarified by microcentrifugation at 4 deg C for 10 min at top speed. 0.5 ml of equilibrated Ni-NTA agarose was added to the supernatant and the slurry was gently mixed at 4 deg C for 1 hr. The mixture was packed onto a column (7mm x 12 mm) and washed with 10 ml of buffer A, 10 ml of buffer A + 5mM imidazole, 10 ml of buffer A + 25mM imidazole and 10ml of buffer A + 50mM imidazole. The proteins were eluted in 1ml of buffer A, pH 7.5 + 200mM imidazole and the imidazole dialyzed out at 4 deg C against 3 changes of 300 volumes buffer A, pH 7.5.

 

In vitro DHFR Assays. DHFR activity was monitored by fluorimetry, to follow the appearance of tetrahydrofolate (THF) (exc. = 310 nm; em. = 360 nm). Fixed substrate concentrations were 30 microM dihydrofolate (DHF) and 25 microM NADPH; the reaction buffer was freshly prepared buffer A, pH 7.5. Concentrations of substrates and inhibitor (methotrexate) were determined spectrophotometrically. Reconstituted enzyme was added to buffer and NADPH; methotrexate (MTX) was then added and the reactions initiated by addition of DHF. Initial rates were determined at 37 deg C under conditions where less than 15% conversion to product had occurred. Blanks contained dialysis buffer instead of enzyme. Ki(MTX) was calculated by non-linear regression with the program: AXUM, by the proportional occupancy binding function: [E.S]/[E]t = ([S]/Kd) / (1+ [S]/Kd + [MTX]/Ki), where E represents reassembled enzyme, and S and Kd (relating to DHF) are treated as unknowns.

 

Transfection and Western Blotting: 1.5 microg of each constructs was used to transfect COS-7 cells grown in six-well tissue culture plates. Transfections were performed using lipofectamine reagent (Life Technologies/Gibco BRL) according to the manufacturer's instructions. The cells were lysed 48 hours post-transfection in lysis buffer containing 50 mM Tris pH 8.0, 150 mM NaCl, 10% SDS, 1% NP40, 1 mM EDTA, 20 microg/ml aprotinin, 5 mM AEBSF, 50 microg/ml soybean trypsin inhibitor, 0.5 microg/ml leupeptin, 0.7 microg/ml pepstatin, 25 microg/ml antipain dihydrochloride and 50 mM N-ethylmethylmaleimide (ubiquitinase inhibitor). The cell lysates were centrifuged for 10 min and a portion of the supernatant was separated on 15% acrylamide SDS gel, transferred to polyvinylidene difluoride (PVDF) membrane (DuPont-NEN) and immunoblotted with anti-HA mouse monoclonal antibody (Boehringer Mannheim) at a concentration of 1 microg/ml. Bound antibodies were detected using horseradish peroxidase-conjugated anti-mouse IgG (Amersham Life Science) and the chemiluminescence detection system (DuPont-NEN).

 

Pulse-chase Experiments and Immunoprecipitation: 48 hours post-transfection, the cells were labelled with Methionine/Cysteine Protein Labeling Mix [35S] (DuPont-NEN) at a concentration of 100 microCi/ml for 30 min at 37 deg C, followed by a chase of 0, 10 and 30 min. The cells were washed with cold phosphate-buffered saline and lysis was performed as described above. The supernatants were immunoprecipitated with 3 microg of anti-HA mouse monoclonal antibody (Boehringer Mannheim) and protein A-Sepharose (Pharmacia Biotech). Immunoprecipitates were washed three times with lysis buffer and once with lysis buffer without detergents. The immune complexes were then subjected to electrophoresis on a 15% acrylamide SDS gel and analyzed by autoradiography.

 

ACKNOWLEDGEMENTS

 

This work was supported by the MRC of Canada (grant no. DGN 059 to SWM) and The Burroughs-Wellcome Fund (SWM). JNP is a recipient of a Fellowship from Les Fonds pour la recherche en santÈ du QuÈbec. SWM is an Awardee of a Burroughs-Wellcome Fund New Investigator Award in the Basic Pharmacological Sciences. We are grateful to Nils Johnsson and Monique Davies for helpful discussions and for carefully reading the manuscript and to Nils Johnsson for providing us with the original plasmids for the USPS and Monique Davies for providing us with the pMT3 plasmids.

 

REFERENCES

 

1. Reed, L. J. (1974) Acc. Chem. Res. 7, 40-46

2. Lander, E. S. (1996) Science 274, 536-539

3. Evangelista, C., Lockshon, D. & Fields, S. (1996) Trends in Cell Biology 6, 196-199

4. Fromont-Racine, M., Rain, J. C. & Legrain, P. (1997) Nature Genetics 16, 277-82

5. Guarente, L. (1993) Proc. Natl. Acad. Sci. USA 90, 1639-41

6. Adams, S. R., Harootunian, A. T., Buechler, Y. J., Taylor, S. S. & Tsien, R. Y. (1991) Nature 349, 694-7

7. Chien, C. T., Bartel, P. L., Sternglanz, R. & Fields, S. (1991) Proc. Natl. Acad. Sci. USA 88, 9578-82

8. Fields, S. & Song, O. (1989) Nature 340, 245-6

9. Gyuris, J., Golemis, E., Chertkov, H. & Brent, R. (1993) Cell 75, 791-803

10. Johnsson, N. & Varshavsky, A. (1994) Proc. Natl. Acad. Sci. USA 91, 10340-4

11. Pelletier, J. N., and Michnick, S. W. (1997) Protein Eng. 10, 89

12. Stammers, D. K., Champness, J. N., Beddell, C. R., Dann, J. G., Eliopoulos, E., Geddes, A. J., Ogg, D. & North, A. C. (1987) FEBS Lett. 218, 178-84

13. Blakley, R. L. (1984) in Folates and Pterins: Chemistry and Biochemistry of Folates (R. Blakley & Benkovic, S., ed) Vol. 1, pp. 191-253, John Wiley and Sons, New York

14. Appleman, J. R., Prendergast, N., Delcamp, T. J., Freisheim, J. H. & Blakley, R. L. (1988) J. Biol. Chem. 263, 10304-13

15. Oefner, C., D'Arcy, A. & Winkler, F. K. (1988) Eur. J. Biochem. 174, 377-85

16. Gegg, C. V., Bowers, K. E. & Matthews, C. R. (1996) in Techniques in Protein Chemistry (D. R. Marshak, ed) Vol. VII, pp. 439-448, Academic Press, New York, USA

17. Bystroff, C. & Kraut, J. (1991) Biochemistry 30, 2227-39

18. Perry, K. M., Onuffer, J. J., Gittelman, M. S., Barmat, L. & Matthews, C. R. (1989) Biochemistry 28, 7961-8

19. Bullerjahn, A. M. & Freisheim, J. H. (1992) J. Biol. Chem. 267, 864-70

20. Buchwalder, A., Szadkowski, H. & Kirschner, K. (1992) Biochemistry 31, 1621-30

21. Chen, X., Rambo, R. & Matthews, C. R. (1992) Biochemistry 31, 2219-23

22. Prevost, M., Wodak, S. J., Tidor, B. & Karplus, M. (1991) Proc. Natl. Acad. Sci. USA 88, 10880-4

23. Kellis, J., Jr., Nyberg, K., Sali, D. & Fersht, A. R. (1988) Nature 333, 784-6

24. Kellis, J., Jr., Nyberg, K. & Fersht, A. R. (1989) Biochemistry 28, 4914-22

25. Dicker, A. P., Waltham, M. C., Volkenandt, M., Schweitzer, B. I., Otter, G. M., Schmid, F. A., Sirotnak, F. M. & Bertino, J. R. (1993) Proc. Natl. Acad. Sci. USA 90, 11797-801

26. Hussain, A., Lewis, D., Yu, M. & Melera, P. W. (1992) Gene 112, 179-88

27. Simonsen, C. C. & Levinson, A. D. (1983) Proc. Natl. Acad. Sci. USA 80, 2495-9

28. Thillet, J., Absil, J., Stone, S. R. & Pictet, R. (1988) J. Biol. Chem. 263, 12500-8

29. Schweitzer, B. I., Srimatkandada, S., Gritsman, H., Sheridan, R., Venkataraghavan, R. & Bertino, J. R. (1989) J. Biol. Chem. 264, 20786-95

30. Banerjee, D., Schweitzer, B. I., Volkenandt, M., Li, M. X., Waltham, M., Mineishi, S., Zhao, S. C. & Bertino, J. R. (1994) Gene 139, 269-74

31. Jonnalagadda, S., Butt, T. R., Monia, B. P., Mirabelli, C. K., Gotlib, L., Ecker, D. J. & Crooke, S. T. (1989) J. Biol. Chem. 264, 10637-42

32. Wilkinson, K. D., Lee, K. M., Deshpande, S., Duerksen-Hughes, P., Boss, J. M. & Pohl, J. (1989) Science 246, 670-3

33. Johnsson, N. & Varshavsky, A. (1994) EMBO J. 13, 2686-98

34. Kaufman, R. J., Davies, M. V., Pathak, V. K. & Hershey, J. W. (1989) Mol. Cell. Biol. 9, 946-58

35. Kunkel, T. A., Roberts, J. D. & Zakour, R. A. (1987) Methods Enzymol. 154, 367-82

 

CORRESPONDING AUTHOR: Gerald M. Carlson gcarlson@cctr.umkc.edu


Return to Journal of Biomolecular Techniques home page.