©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Step-arrest Mutants of Phage Mu Transposase
IMPLICATIONS IN DNA-PROTEIN ASSEMBLY, Mu END CLEAVAGE, AND STRAND TRANSFER (*)

(Received for publication, August 26, 1994; and in revised form, October 19, 1994)

Keetae Kim Soon-Young Namgoong Makkuni Jayaram Rasika M. Harshey (§)

From the Department of Microbiology, University of Texas at Austin, Austin, Texas 78712

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

We describe the isolation and characterization of Mu A variants arrested at specific steps of transposition. Mutations at 13 residues within the Mu A protein were analyzed for precise excision of Mu DNA in vivo. A subset of the defective variants (altered at Asp, Asp, Gly, and Glu) were tested in specific steps of transposition in vitro. It is possible that at least some residues of the Asp-Asp-Glu triad may have functional similarities to those of the conserved Asp-Asp-Glu motif found in several transposases and retroviral integrases. Mu A(D269V) is defective in high-order DNA-protein assembly, Mu end cleavage, and strand transfer. The assembly defect, but not the catalytic defect, can be overcome by precleavage of Mu ends. Mu A(E392A) can assemble the synaptic complex, but cannot cleave Mu ends. A mutation of Gly to aspartic acid within Mu A permits the uncoupling of cleavage and strand transfer activities. This mutant is completely defective in synaptic assembly and Mu end cleavage in presence of Mg. The assembly defect is alleviated by replacing Mg with Ca. Some Mu end cleavage is observed with this mutant in the presence of Mn. When presented with precleaved Mu ends, Mu A(G348D) exhibits efficient strand transfer activity.


INTRODUCTION

The Mu transposase (Mu A) catalyzes the cleavage of Mu DNA ends and their subsequent joining (or strand transfer) to target DNA (see Mizuuchi, 1992). Under standard assay conditions, the cleavage of Mu ends exhibits a strict requirement for DNA supercoiling, Escherichia coli protein HU and divalent metal ions (Mg or Mn). Interactions of monomeric Mu A protein with six att subsites (three at each end of Mu constituting attL and attR) and an internal enhancer element leads to the formation of a tetrameric assembly of Mu A within a high-order nucleoprotein complex. Ca ions can support assembly of this complex, but cannot support strand cleavage at Mu ends (Mizuuchi et al., 1992). The catalytically functional form of Mu A is the tetramer.

The Mu A protein is organized into three principal domains as determined by limited proteolysis experiments (Nakayama et al., 1987). Structure-function studies have mapped att and enhancer DNA binding activities to two separate regions on the NH(2)-terminal domain I (see Fig. 1A; Nakayama et al., 1987; Leung et al., 1989; Mizuuchi and Mizuuchi, 1989). The central, proteolytically stable domain II was inferred to contain the catalytic site since a temperature-sensitive (ts) mutation that allows cleavage but blocks strand transfer maps here (Leung and Harshey, 1991). This domain also shows nonspecific (target?) DNA binding activity (Nakayama et al., 1987). The catalytic domain must extend into part of COOH-terminal domain III as determined by the activity of deletions that are localized here (Harshey and Cuneo, 1986; Bremer et al., 1988; Betermier et al., 1989; Leung and Harshey, 1991; Baker et al., 1993). The distal two-thirds of domain III is required for interaction with the accessory transposition protein Mu B which promotes intermolecular strand transfer (Harshey and Cuneo, 1986; Leung and Harshey, 1991; Baker et al., 1991).


Figure 1: Mu A protein (transposase): domain structure and amino acid alignment with other transposases and a retroviral integrase. A, on the basis of limited proteolysis, three domains were assigned to A protein (see text). Amino acid numbers corresponding to the amino terminus of each major domain are shown beneath the structure. The NH(2)-terminal domain I contains subdomains alpha, beta, and , which encode site-specific DNA binding activity. Ialpha binds to Mu enhancer and Ibeta binds to Mu att sites. Domain II is likely the catalytic domain. It shows nonspecific DNA binding activity, and a sequence stretch resembling the helix-turn-helix DNA binding motif maps here (open box between residues 390 and 409; Harshey et al., 1985). A mutation specifically affecting strand transfer (asterisk) maps toward the COOH-terminal end of this domain (residue 548; Leung and Harshey, 1991). The distal two-thirds of domain III is responsible for important interactions with the Mu B protein, whereas the proximal one-third is important for assembly and catalysis. The amino acid residues mutated in this study are indicated above the domain structure. B, the amino acid alignment shown here is based on the data from Radstrom et al.(1994). A similar but not identical alignment has been deduced by Baker and Luo(1994). Aspartic acid residues at positions 269 and 294 and a glutamic acid residue at position 392 within Mu A correspond to conserved residues in all proteins (shown in outlined letters). A third aspartic acid residue is conserved in all proteins except Mu A. This position is Gly of Mu A in our alignment and Phe of Mu A in the Baker-Luo alignment. Three other positions, all glycines (shown in bold letters) are also conserved in all proteins.



Mu transposition shares many common chemical features with transposition of other transposable elements as well as integration of retroviruses into their host genomes (see Mizuuchi(1992)). Most of these elements share a conserved Asp-Asp-35-Glu, (where 35 indicates the number of residues separating Asp and Glu) motif (Fayet et al., 1990; Rowland and Dyke, 1990; Kulkosky et al., 1992; Radstrom et al., 1994) whose importance in catalysis has been established from mutagenesis experiments (Engelman and Craigie, 1992; Leavitt et al., 1993; van Gent et al., 1993). Sequence comparison of the Mu transposase with the equivalent proteins of the above elements identified residues within domain II of Mu A that aligned with the Asp-Asp-35-Glu motif (Fig. 1B). (^1)By analogy with the exonuclease of E. coli DNA polymerase I which uses a cluster of acidic residues to position metal ions within its active site (Beese and Steitz, 1991; Derbyshire et al., 1991), the Asp-Asp-35-Glu residues are proposed to be involved in coordinating divalent metal ions and in positioning the target phosphodiester and the attacking nucleophile for phosphoryl transfer (Kulkosky et al., 1992; Engelman and Craigie, 1992).

This study focuses on defining the role of potential active site amino acids within the presumed catalytic domain II of Mu A by making site-directed mutations in this region. We show that mutations of Asp (D269V), Asp (D294N), and Glu (E392A) abolish in vivo activity of the transposase. A mutation at Gly (G348D) also resulted in loss of activity in vivo. From a study of the in vitro properties of these mutants, we infer that these residues play critical and distinct roles in organizing the transposition complex and in the catalytic steps. It seems plausible that the acidic residues could be involved in facilitating a two metal ion mechanism for phosphoryl transfer analogous to that used by the 3`-5` exonuclease reaction of E. coli DNA polymerase I.

While this manuscript was in preparation, Baker and Luo(1994) reported an independent mutational analysis of eleven residues of Mu A. Three of our mutations are at similar positions. The properties of one of our mutants, Mu A(E392A), are comparable with those of Mu A(E392Q) isolated by Baker and Luo(1994). However, the behavior of a second mutant MuA(D269V) characterized by us is distinct from that of Mu A(D269N) described by Baker and Luo(1994). In our analyses, the most interesting step-arrest phenotypes are displayed by Mu A(G348D). Baker and Luo(1994) have not studied an analogous mutant.


EXPERIMENTAL PROCEDURES

Plasmids

Plasmid pRA158 contains the Mu A gene engineered to contain two new restriction sites within the coding region without altering the amino acid sequence: an EcoRV site was created by changing T A (third position of a Gly codon) at nucleotide 2137, and an HpaI site by changing G T (third position of a Val codon) at nucleotide 2812 on the Mu DNA sequence (Harshey et al., 1985; Priess et al., 1987). Both sites lie within the region encoding domain II of the A protein. The Mu DNA fragment from nucleotide 1309 to 3328 containing the engineered Mu A gene was cloned into vector pTZ19U (U. S. Biochemical Corp.) to give pRA158. In this construct, expression of the A gene is driven from an inducible T7 promoter. There was a basal level expression of A protein even in the absence of induction. Mutations obtained by directed mutagenesis were transferred to the Mu A gene within pRA158 by replacing a wild-type restriction fragment with the corresponding mutant fragment. pRA158 and its derivatives were used in excision assays (see below). It is known that high levels of Mu A are inhibitory to excision (Harshey and Cuneo, 1986). Hence, excision was monitored under non-induced conditions.

For efficient expression of Mu A, the Mu A gene from pRA158 was transferred to another T7 expression vector, pET11-a (Novagen), to generate pET158.

Plasmid pMK21, used in ``type 0'' and ``type I'' assays, was constructed as follows. The BamHI site on pRA167 (Leung et al., 1989) was changed to SalI, which upon SalI digestion and ligation eliminated a 7-kilobase SalI trp-lac fragment in the resulting plasmid pZW167. Restriction sites for BamHI and BglII were created on this plasmid by site-directed mutagenesis of nucleotide 58 (T C) and nucleotides 224, 228, and 229 (C A, A C, and A T), respectively. The resulting plasmid pMK21 has 1.8 kilobases of Mu sequences between attL and attR, which include the enhancer element.

``Me(2)SO assays'' were carried out using linear DNA fragments obtained by restriction enzyme digestion from a pUC19 derivative containing R1-R2 (Namgoong et al., 1994).

The plasmid engineered to be cut at the Mu right end by HindIII digestion was a gift from M. Surette and G. Chaconas (University of Western Ontario) and is described in Namgoong et al.(1994).

Mutagenesis and Cloning

Several of the point mutations were made by polymerase chain reaction mutagenesis of linearized plasmids by the mega-primer method (Barik, 1993). Others were obtained by the gapped duplex protocol for mutagenesis of appropriate Mu A fragments cloned into M13mp19. The latter procedure involved enrichment of mutants by selecting against the amber-containing parent templates (Kramer et al., 1984). The mutations were transferred to pRA158 or pET158 vectors by replacing a wild-type restriction fragment by the equivalent fragment harboring the mutation. Authenticity of mutations were verified by DNA sequencing.

Protein Purification

The wild-type Mu A protein and the variant proteins were expressed from pET158 and its derivatives by induction with isopropyl-1-thio-beta-D-galactopyranoside (0.4 mM) for 3 h at 30 °C. Four of the mutants, Mu A(D269V), Mu A(D294N), Mu A(G348D), and Mu A(E392A) were purified to near homogeneity (Kuo et al., 1991). These proteins were judged to be >95% pure from Coomassie Brilliant Blue staining of SDS-polyacrylamide gels in which they were fractionated. Concentration of Mu A or Mu A variants was derived from the extinction coefficient E = 1.83 (Kuo et al., 1991).

In Vivo Assay for Mu DNA Excision

The assay was done according to Bukhari(1975). Strain RH208 (F` prolac::mini-Mukan/Delta prolac his met Str^R; Harshey and Cuneo, 1986) was transformed with pRA158 plasmids bearing wild-type and mutant alleles of Mu A. The transformants were patched (1 cm^2) on McConkey lactose indicator plates and incubated at 30 °C for 4 days to allow precise excision of Mu DNA, which was scored by counting the number of Lac papillae.

In Vitro Assays for DNA-Protein Assembly and Transposition

Formation of the uncleaved Mu end synaptic complex (type 0) in the presence of Ca ions (Mizuuchi et al., 1992) was monitored by formation of a DSP (^2)(dithiobis(succinimidyl propionate))-cross-linked structure after restriction digestion as described by Surette et al.(1991). Alternatively, type 0 reaction mixtures were treated with 10 µg/ml heparin to remove all loosely bound Mu A protein (Kuo et al., 1991) before electrophoresis, which resulted in a distinct type 0 complex band, migrating just above the supercoiled donor plasmid. Me(2)SO assays were done as described earlier (Namgoong et al., 1994). Standard Mu end cleavage (type I) and strand transfer to target DNA in presence of Mu B and ATP (type II) assays were done as described by Surette et al.(1987). 10 µg/ml heparin was added to reaction mixtures when electrophoresed without SDS.

Miscellaneous Methods

Bacterial transformation, isolation of plasmid DNA, restriction enzyme digestion, DNA ligation, and other miscellaneous procedures were done as described by Maniatis et al.(1982).


RESULTS

Mutations That Block Mu Excision in Vivo

It is known that mutations in Mu A that affect transposition in vitro also affect Mu DNA excision in vivo (Harshey and Cuneo, 1986).(^3)Hence precise excision of Mu DNA, and consequent restoration of an otherwise interrupted lacZ gene (Table 1), provides a quick in vivo functional test for Mu A variants obtained by directed mutagenesis. Several lines of reasoning determined which residues were to be targeted for mutagenesis. Based on the mechanistic similarity of Mu transposition to retroviral integration and the conservation of the Asp-Asp-35-Glu motif among retroviral integrases, we initially scanned for such motifs within the presumed catalytic domain (domain II) of Mu A protein. Asp/Glu and Asp/Glu of Mu A (see Fig. 1A) have the right spacing expected for a canonical Asp-Asp-Glu motif. The in vivo excision activities of Mu A substituents altered at these positions are shown in Table 1. While mutations at three of these four positions (D294V or D294N, D405V, and E442L) markedly reduced excision, a change from Glu to leucine did not affect excision. It is unlikely therefore that Asp/Glu is part of a typical Asp-Asp-35-Glu motif relevant to transposition. The in vivo phenotype of mutations at Asp and Glu suggest that this pair might well contribute to a functional Asp-Asp-35-Glu motif. However, further in vitro analysis of the mutants was impeded by difficulties in their purification. Effects of other substitutions at these two positions are under investigation. The sequence alignment shown in Fig. 1B reveals that Asp, Asp, and Glu of Mu A correspond to highly conserved residues within the transposase/retroviral integrase family. Mutations at Asp and Glu (as was also the case with those at Asp) resulted in a large drop in in vivo excision (Table 1). The alignment in Fig. 1B is also significant by the conspicuous absence of a conserved aspartic acid at position 348 in Mu A and the presence of a glycine instead. We therefore wanted to test the effect of changing this position to aspartic or glutamic acid. Both changes led to nearly complete loss of in vivo excision (Table 1). A number of other aspartic residues that do not show a strict 35 amino acid spacing with a downstream glutamic residue were also mutated to provide a frame of reference. None of these mutations had a significant effect on excision (Table 1).



In this paper we provide a detailed in vitro analysis of three mutant A proteins that were functionally defective as judged by the excision assay: Mu A(D269V), Mu A(G348D), and Mu A(E392A). Particularly interesting was the characteristic ``step-arrest'' behavior of the variant proteins in the transposition pathway and the differential response of two of these proteins to different divalent cations in the reaction (see below). Mu A(D294N) was uniformly defective in all in vitro assays detailed below, retaining approximately 5-10% of the wild-type activity (data not shown). Hence this variant is not revealing in regard to the mechanism of transposition and is not discussed in detail.

Assembly of the Uncleaved Mu Synaptic Complex (Type 0) in Ca

The ability of the mutants to assemble Mu ends into an uncleaved high-order nucleoprotein complex (type 0) in the presence of Ca ions was examined by the indirect assay (Fig. 2; Surette et al., 1991). The substrate in these reactions is a negatively supercoiled plasmid that contains attL, attR, and the enhancer sequences. After protein cross-linking with DSP to stabilize the synapse, HindIII plus XbaI digestion was carried out to generate the ``[chi]'' structure. The uncut closed and open circular forms of the substrate plasmid (CCP and OCP, lane 3, Fig. 3A), the linear form (LP, lane 1, Fig. 3A), linear attL and attR fragments resulting from HindIII plus XbaI digestion (L and R, lane 2, Fig. 3A) and the form (migrating between the linear and open circular plasmid forms; lane 4, Fig. 3A) can be resolved by electrophoresis in 1% agarose gels. Mu A(D269V) was defective in assembly of the synpased structure (note the absence of in lane 5, Fig. 3A). By contrast, Mu A(G348D) showed roughly 20-25% of wild-type activity (lane 6, Fig. 3A), whereas Mu A(E392A) showed approximately 50% activity (lane 7, Fig. 3A). Prolonged incubations with Mu A(E392A) resulted in nearly 80% of the donor substrate being converted into type 0 (data not shown; see also lane 5, Fig. 3B). Treatment with SDS prior to electrophoresis resulted in dissociation of the structure to yield linear products (lanes 8-11, Fig. 3A).


Figure 2: The rationale for the assay for synapsis. Synapsis between attL and attR within the circular plasmid in presence of Ca leads to the type 0 complex within which Mu A is shown to exist as a tetramer (Lavoie et al., 1991; Mizuuchi et al., 1992). The enhancer element is located across the HindIII (H) site (not shown). HindIII plus XbaI (X) digestion would give rise to the form (Surette et al., 1991). Digestion with either HindIII or XbaI alone would yield one of two alpha forms. When the chi is treated with SDS, two linear DNA fragments (one harboring attL; the other harboring attR) will be generated.




Figure 3: Assembly of the type 0 complex in presence of Ca. A, after incubation of the substrate plasmid with Mu A or Mu A variants (approximately 2 pmol of protein/pmol of att subsite) in the presence of HU and Ca followed by DSP treatment, extensive digestion with HindIII plus XbaI was carried out prior to electrophoretic fractionation in agarose. Lane designations are as follows: lane 1, linearized plasmid; lane 2, plasmid cut with HindIII plus XbaI; lane 3, uncut plasmid; lanes 4-7, reactions with Mu A, Mu A(D269V), MuA(G348D), and Mu A(E392A), respectively. Lanes 8-11, reactions as in lanes 4-7 treated with SDS. B, unlike the assays shown in A, reactions here were not digested with restriction enzymes, nor treated with SDS. 10 µg/ml of heparin was added to the samples prior to electrophoresis. CCP, closed circular plasmid; OCP, open circular plasmid; LP, linear plasmid; R, attR DNA fragment; L, attL DNA fragment. The species above the open circular form of the substrate plasmid in B is probably a trace of the supercoiled dimer form of the plasmid.



When samples were fractionated without restriction digestion but with addition of heparin prior to electrophoresis, the pattern shown in Fig. 3B was obtained. The band that migrates just above the covalently closed circular form (CCP) of the plasmid is the type 0 complex, the precursor of the band seen in Fig. 3A. When these reaction mixtures were treated with HindIII and XbaI under partial digestion conditions, there was a strong correspondence between the disappearance of type 0 and the appearance of together with ``alpha'' bands (alpha arises from a single restriction cut within the synapsed complex; see Fig. 2) (data not shown). The type 0 band was present in the Mu A (lane 2, Fig. 3B), Mu A(G348D) (lane 4, Fig. 3B), and the Mu A(E392A) (lane 5, Fig. 3B) reactions and absent in the Mu A(D269V) reaction (lane 3, Fig. 3B). The level of type 0 complex yielded by Mu A(G348D) was lower than that obtained with Mu A or Mu A(E392A), possibly because the synapsed complex produced by Mu A(G348D) in presence of Ca was less stable than the normal complex.

Thus, while Mu A(G348D) and Mu A(E392A) can assemble the type 0 complex, MuA(D269V) is defective in assembly.

Cleavage of Mu Ends in Presence of Mg: Type I Assays

Under standard assay conditions with a supercoiled substrate in presence of Mg, Mu A can nick attL and attR to yield the cleaved donor complex (type I complex). The ability of the mutant proteins to cleave Mu ends in this assay is shown in Fig. 4A. After incubation of the substrate plasmid with Mu A or Mu A variants in presence of Mg, samples were treated with SDS prior to agarose gel electrophoresis. The extent of type I complex formed was indirectly measured as the increase in the relaxed form of the substrate (labeled OCP). All three Mu A variants tested negative (lanes 5-13, Fig. 4A) in formation of the type I complex. Wild-type Mu A showed normal reaction as expected (lanes 2-4, Fig. 4A). In reactions not treated with SDS, the type I complex (migrating in between the supercoiled and relaxed forms of the plasmid) was seen only with wild type Mu A and not with any of the mutant proteins (data not shown). As in the Ca reaction (see Fig. 3B, lane 5), Mu A(E392A) yielded the type 0 complex. More importantly, in contrast to the Ca incubations (see Fig. 3B, lane 4), no band corresponding to the type 0 form was detected with Mu A(G348D) under these type I (Mg) assay conditions (data not shown).


Figure 4: Reactions of Mu A variants in Mg, in Ca followed by Mg and in Mn. A, standard type I assays were done in presence of Mg. Samples were treated with SDS prior to electrophoresis. Lane 1 is a control reaction without addition of Mu A protein. For each reaction set, reactions from left to right contained 1, 2, and 4 pmol of Mu A or A variant/att subsite. B, reaction mixtures were first incubated with Ca under type 0 assay conditions. Incubations were then continued in presence of added Mg (10 mM). Samples were fractionated without SDS treatment. 1. C, reactions were done as in A, except that Mg was replaced by Mn, and samples were fractionated without SDS. Lane 1 is a reaction without added Mu A protein. Lane 2 is a wild-type Mu A reaction in 2 mM Mn. Lanes 3-14 represent reactions with Mu A variants. For each variant, reactions from left to right contain 5, 10, 20, and 40 mM Mn, respectively. All other symbols as in Fig. 3. The bands migrating below the open circular plasmid (OCP) forms in some lanes are likely to be a trace amounts of the linear form of the substrate plasmid. CCP, closed circular plasmid.



Thus, in presence of Mg, Mu A(E392A) is normal and Mu A(G348D) is defective in the assembly of a stable precleavage complex (equivalent to type 0).

Conversion of Type 0 Complex Assembled in the Presence of Ca to Type I Complex by Added Mg

It is known that the type 0 complex assembled by Mu A in the presence of Ca can be converted to type I by the addition of Mg (Mizuuchi et al., 1992). Can any of the Mu A variants mediate the type 0 to type I conversion? In these reactions, the substrate was preincubated in presence of Ca and then made 10 mM in Mg. With wild type Mu A, a prominent type I band was detected (lane 1, Fig. 4B). There was no evidence for the formation of the type 0 or the type I complex with Mu A(D269V) (lane 2, Fig. 4B). In the Mu A(G348D) reaction, the type 0 band disappeared with a concomitant increase in the relaxed plasmid band (OCP, lane 3, Fig. 4B). Primer extension reactions showed that the relaxed species did not have nicks at Mu ends (data not shown). Although the nature of the relaxation step remains obscure at this time, the reaction resulted in complete dissociation of the type 0 complex. With Mu A(E392A) the type 0 complex was observed (lane 4, Fig. 4B), but no att cleavage within this complex (conversion to the type I complex) occurred. When the Mu A(E392A) reaction was treated with SDS prior to fractionation, the type 0 band was quantitatively converted to the supercoiled form of the plasmid (data not shown).

Thus, neither Mu A(G348D) nor Mu A(E392A) can mediate att-specific DNA cleavage within the pre-organized type 0 complex.

Effect of Mn on the Steps of the Transposition Pathway

Mn at 2-5 mM concentration can substitute for Mg (10 mM) in the formation of the type I complex with wild type Mu A (lane 2, Fig. 4C). At higher concentrations of Mn, reaction with Mu A was inhibited (data not shown). Among the mutants, Mu A(D269V) showed no reactivity at Mn concentrations ranging from 5 to 40 mM (lanes 3-6, Fig. 4C). At 5-10 mM Mn, the type I complex was detectable with Mu A(G348D) (lanes 7 and 8, Fig. 4C). Similarly, the type 0 band was seen with Mu A(E392A) at 5-20 mM Mn (lanes 11 and 12, Fig. 4C). However, type I and type 0 complex formation by the latter two mutants was inhibited at Mn concentrations above 5 and 10 mM, respectively (lanes 8-10, 13, and 14, Fig. 4C). An increase in the nicked substrate population with increasing Mn concentration was also noted in reactions with both these A variants (lanes 9, 10, 13, and 14, Fig. 4C). A primer extension reaction verified that this increase did not result from specific cleavage at Mu ends (data not shown). When the reactions included ATP, Mu B, and a target substrate (type II assay), a trace of strand transfer activity was observed with Mu A(E392A) at 10-20 mM Mn; higher levels of strand transfer were obtained with Mu A(G348D) at 5-10 mM Mn (results not shown). The reactivity of Mu A(G348D) correlated well with the extent of the type I complex yielded by it (lanes 7 and 8, Fig. 4C).

Thus, Mn but not Mg ions can support att cleavage by Mu A(G348D) at concentrations comparable to those in the wild-type reaction. The cleaved ends can be used for strand transfer. Mn-supported att cleavage by Mu A(E392A) must be quite low as inferred from the trace amounts of strand transfer yielded by this mutant.

Cleavage and Strand Transfer in the Me(2)SO Assay

Me(2)SO has been found to relax the topological constraints and the enhancer requirement of Mu transposition without sacrificing many of the physico-chemical aspects of the reaction (Mizuuchi and Mizuuchi, 1989). We have recently used the Me(2)SO reaction to unmask features of the transposition reaction that could not be revealed in the standard type I assay (Namgoong et al., 1994). In this assay, linear DNA containing R1 and R2 subsites from attR of Mu can efficiently partake in the assembly of high-order DNA-protein complexes capable of cleavage and strand transfer. Results of the Me(2)SO reaction (in presence of Mg) with and without SDS treatment are displayed in Fig. 5, A and B, respectively. Wild-type Mu A formed a ladder of high-order complexes (HC, lane 2, Fig. 5A), which upon SDS treatment dissociated to reveal products of strand transfer (STP, lane 2, Fig. 5B). Mu A(E392A) assembled a distinct complex (lane 5, Fig. 5A), but not Mu A(D269V) (lane 3, Fig. 5A) nor Mu A(G348D) (lane 4, Fig. 5A). The presence of Mu A protein within the complexes was confirmed by probing with Mu A-specific antiserum as described by Namgoong et al.(1994). The SDS-treated reaction revealed no strand transfer products with any of the mutant proteins (lanes 3-5, Fig. 5B).


Figure 5: High-order DNA-protein assembly and strand transfer under Me(2)SO assay conditions. A and B, reactions were carried out in Me(2)SO with a linearized plasmid containing attR subsites R1-R2 (Namgoong et al., 1994). C and D, the substrate was a linear DNA fragment containing precleaved attR (R1-R2-R3) (Namgoong et al., 1994). Samples were fractionated by electrophoresis without SDS treatment (A and C) or after SDS treatment (B and D). R, linear DNA fragment containing R1-R2; RC; linear DNA containing precleaved attR; HC, high-order DNA-protein complex; STP, strand transfer products.



The results when linear DNA precleaved at attR was used as the substrate in the Me(2)SO reaction are shown in Fig. 5, C and D. Surprisingly, Mu A(D269V), which tested negative in all assays so far, was seen to assemble a complex (lane 3, Fig. 5C) in amounts comparable with that formed by Mu A(E392A) (lane 5, Fig. 5C). Furthermore, Mu A(G348D) also yielded a similar complex with the precleaved DNA (lane 4, Fig. 5C). The formation of this complex with all three mutants as well as wild-type Mu A was independent of the addition of metal ions (not shown). Among the mutants, only Mu A(G348D) (lane 4, Fig. 5D) and not the other two mutants (lanes 3 and 5, Fig. 5D) could yield strand transfer products from the prenicked substrate. This strand transfer activity was absolutely dependent on added metal ions (Mg or Mn) in contrast to the strand transfer activity of wild-type protein which did not require an exogenous supply of either of these two metals under the Me(2)SO assay conditions (data not shown).

Metal Ion Specificity and Step-arrest Phenotypes of Mu A Mutants

We may now summarize the step-arrest phenotypes of the mutants as deduced from the various assays described above ( Table 2and Table 3). Mu A(D269V) cannot assemble the type 0 complex with Ca from a supercoiled substrate, nor can it assemble the type 0, type I, or type II complex in presence of Mg or Mn. It cannot utilize a linear substrate to organize a high-order complex under Me(2)SO assay conditions (in the presence of 10 mM Mg). However, when provided with a precleaved attR DNA fragment, it assembles a high-order complex that shows no strand transfer potential. Mu A(E392A) forms the type 0 complex with Ca nearly as efficiently as wild-type Mu A, but cannot cleave the Mu ends within this complex and convert it to type I when Mg is provided. It can also form a high-order complex with a linear attR fragment in Me(2)SO whether or not the Mu end is precleaved. However, no strand transfer is mediated by this protein. In presence of 10-20 mM Mn, this mutant executes a very low level of att cleavage and strand transfer when provided with Mu B, ATP, and a DNA target. Mu A(G348D) assembles the type 0 complex (albeit not very efficiently) in the presence of Ca, but no such complex is formed with Mg, nor can it cleave a pre-assembled type 0 complex in the presence of Mg. At 5-10 mM Mn, Mu A(G348D) can form modest levels of a type I-like complex which can undergo strand transfer to a target substrate in the presence of Mu B and ATP. In the Me(2)SO assay, Mu A(G348D) can build a high-order DNA-protein complex from a precleaved attR. The assembled complex is competent in strand transfer in the presence of Mg or Mn.





The properties of Mu A(E392A) described here are consistent with those of Mu A(E392Q) reported by Baker and Luo(1994). However, Mu A(D269V) is distinct from Mu A(D269N) studied by Baker and Luo(1994). In our assays, Mu A(D269V) shows a severe defect in DNA-protein assembly and does not show att cleavage or strand transfer activity in presence of Mn.


DISCUSSION

Divalent metal ions play a central role in phosphoryl transfer reactions in nucleic acids. Examples include reactions mediated by polymerases, nucleases, restriction enzymes, DNA-dependent ATPases, and ribozymes, among others. It is reasonable to expect that key acidic residues within the ``phosphoryl transferase'' family might be involved in coordinating the metal and orienting it within the active site. The x-ray structure of the Klenow fragment of E. coli DNA polymerase I, together with directed mutational analysis, provides insights into how two metal ions can be positioned within the transition state for the 3`-5` exonuclease activity of this enzyme (Beese and Steitz, 1991; Derbyshire et al., 1991). Properties of residue-specific mutants of retroviral integrases have led to the hypothesis that key acidic amino acids (forming a Asp-Asp-35-Glu motif), critical for catalysis, may function by coordination of the metal cofactor (Kulkosky et al., 1992; Engelman and Craigie, 1992; van Gent et al., 1993; Leavitt et al., 1993). In this study we describe variants of Mu A protein that are arrested at specific steps of the transposition pathway in a metal-specific manner.

Conservation of Catalytic Motifs among Transposases and Retroviral Integrases?

Can the identical chemical mechanisms of strand breakage and transfer by Mu A and retroviral integrases (Mizuuchi, 1992) be correlated with a conserved Asp-Asp-35-Glu active site cluster (Kulkosky et al., 1992; Radstrom et al., 1994)? Our mutational studies have not revealed a typical Asp-Asp-35-Glu motif within Mu A. Nevertheless, the functional defects observed with Mu A variants altered at specific aspartic and glutamic acid residues (for example Asp and Glu) within the presumed catalytic domain lends credence to the notion that these amino acids are either active site residues or contribute significantly to organization of the reaction pocket. Baker and Luo (1994) demonstrate that such mutants (for example Mu A(D269N) and Mu A(E392A)) can be rescued, albeit inefficiently by replacement of Mg by Mn at higher levels. It is possible that the presumed catalytic function of these residues might involve interactions with the metal. Alternately, they may affect catalysis by interacting with residues directly involved in metal coordination.

Our studies reveal specific effects of these mutations in individual steps of the transposition pathway and their distinct responsiveness to a set of divalent metal ions, Ca, Mg or Mn. Quite striking is the observation that changing Gly (which aligns with the second Asp of the retroviral Asp-Asp-35-Glu motif (see Fig. 1B)) to aspartic acid results in bi-specificity of metal ions: Ca but not Mg can be utilized in the assembly step, only Mn can function in cleavage, and Mg or Mn can function in the strand transfer steps. This mutation, in conjunction with Glu, artificially creates an analog of the Asp-35-Glu motif that is absent in wild-type Mu A. The novel assembly and catalytic potentials revealed by the mutation strongly implies its location within the active site pocket or its influence on the active site configuration. Overall, the properties of the mutants suggest that the transposase and the retroviral integrases may utilize analogous metal coordinating mechanisms although the amino acid residues that perform this function may not conform to an obvious primary sequence motif.

Role of Precleaved Ends in Assembly

A number of experimental observations link the cleavage of Mu ends to the formation of a stable, strand transfer-competent DNA-protein complex. In the normal transposition assay using negatively supercoiled substrates, formation of the stable type I complex is dependent on the nicking of the Mu ends (Surette et al., 1991). Mutations of nucleotides at Mu ends that block cleavage by Mu A effectively prevent the assembly of the type I complex. Furthermore, in the Me(2)SO reaction, the defect in high-order DNA-protein assembly of linear, Mu A-noncleavable att DNA fragments can be overcome or ameliorated by precleavage of Mu ends with restriction enzymes (Namgoong et al., 1994). However, att cleavage is not a prerequisite for stable assembly. Noncleaved complexes (type 0) can be assembled by wild-type Mu A (Mizuuchi et al., 1992) and by Mu A variants described here. However, in the Me(2)SO assays, such complexes appear to be less stable than type I complexes (Namgoong et al., 1994). We propose that the DNA-protein interactions that stabilize the type 0 and type I complexes are distinct. Recognition of the terminal bases at the two ends of Mu is critical to type 0 assembly. Indeed, point mutations of these nucleotides block type 0 complex formation. (^4)Cleavage of Mu ends likely promotes a new set of interactions (perhaps involving the exposed 3`-hydroxyl and/or the 5`-phosphate) that contribute to stable type I assembly. The rescue of Mu A(D269V), which cannot assemble type 0, in Me(2)SO-aided type I-like assembly by att cleavage further demonstrates the functional significance of the cleaved ends in organizing a stable DNA-protein complex. A strong correlation between DNA cleavage and stabilization of high-order DNA-protein associations has been observed with the Flp recombinase as well (Qian et al., 1991).

Metal-specific Step-arrest Phenotypes

The Mu A variants described here together with the previously characterized strand transfer-defective variant Mu Ats5045 (Leung and Harshey, 1991) now allow us to arrest the transposition reaction at the assembly/synapsis step, the strand cleavage step, or the strand transfer step under appropriate reaction conditions. These step-arrest mutants therefore provide a tool for mechanistic analysis of the partial reactions of a complex biochemical pathway. Such mutants have proven seminal to understanding the mechanism of another phosphoryl transfer reaction: site-specific recombination by the Flp recombinase. Pairwise combinations of Flp step-arrest mutants helped reveal the mode of active site assembly and the mechanism of strand cleavage and strand transfer in this recombination system (Parsons et al. 1988, 1990; Chen et al., 1992; Lee and Jayaram, 1993). Whereas Flp uses a tetrameric protein assembly to mediate four strand cleavages and four strand joinings during one recombination event, Mu A uses a tetrameric assembly to bring about four strand breakages and two strand unions during one round of transposition. The prospect of catalytic complementation between pairs of step-arrest mutants holds the promise for deciphering the role of individual A monomers within the transposition complex.

Role of Metal Ions in Transposition by MuA

The central role played by divalent metal ions such as Mg and Mn in phosphoryl transfer in nucleic acids has been well documented. However, revealing the precise details of how metal facilitates catalysis has not been easy. Structural analysis of E. coli alkaline phosphatase (Kim and Wyckoff, 1991) and of the Klenow polymerase (Beese and Steitz, 1991) complexed with a deoxynucleoside monophosphate and a single-stranded DNA substrate provides a rational explanation for the role of metal coordination during phosphoryl transfer. A common feature of the two active site configurations is two divalent metal ions held 3.9 Å apart. The polymerase structure strongly suggests that the two metal ions liganded in part to specific aspartic and glutamic acid residues of the protein contribute to the stabilization of the transition state. Although one metal facilitates the production of the attacking nucleophile (the hydroxide ion), the other is thought to facilitate the departure of the leaving group (3`-hydroxyl). The involvement of a metal ion in stabilizing the leaving group has been inferred for the Tetrahymena ribozyme as well (Piccirilli et al., 1993). The restoration of cleavage activity by Mn and Zn to a ribozyme variant in which 3`-bridging oxygen is replaced by sulfur is consistent with the ability of these metals to coordinate sulfur more strongly than Mg. A multiple metal ion mechanism has been proposed for the ribonuclease P reaction (Smith and Pace, 1993). In this case, a catalytically important Mg appears to be coordinated by the 2`-hydroxyl at the cleavage site within the tRNA substrate.

The differential effects of various divalent cations on Mu A and Mu A variants in the steps of transposition would be consistent with a two metal or multiple metal ion mechanism, although a mechanism involving a single metal ion (Suck, 1992) cannot be ruled out. The requirement for Ca (and not Mg or Mn) for type 0 assembly and that of Mn (rather than Mg) for strand cleavage and stransfer by Mu A(G348D) is suggesstive of the involvement of more than one metal ion during transposition. The extreme simplicity of a two metal ion mechanism in phosphoryl transfer (one to generate the oriented nucleophile and the other to stabilize the leaving group) suggests that this catalytic strategy may be utilized by a number of enzyme systems. The evolutionary design of their active sites would be guided by the need to achieve precise orientation of the metal ions with respect to the substrate. Structural analyses of the active site configurations of Mu A and its step-arrest variants should be insightful in this regard.


FOOTNOTES

*
This work was supported by funds from National Institutes of Health. 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.

§
To whom correspondence should be addressed. Tel.: 512-471-6881; Fax: 512-471-5546; rasika{at}uts.cc.utexas.edu.

(^1)
L. Sundstrom, personal communication.

(^2)
The abbreviation used is: DSP, dithiobis(succinimidyl propionate).

(^3)
R. M. Harshey, unpublished data.

(^4)
S.-Y. Namgoong and R. M. Harshey, unpublished results.


ACKNOWLEDGEMENTS

We thank Jehee Lee for help in preparing Fig. 2.


REFERENCES

  1. Baker, T. A., and Luo, L. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 6645-6658
  2. Baker, T. A., Mizuuchi, M., and Mizuuchi, K. (1991) Cell 65, 1003-1013 [Medline] [Order article via Infotrieve]
  3. Baker, T. A., Mizuuchi, M., Savilahti, H., and Mizuuchi, K. (1993) Cell 74, 723-733 [Medline] [Order article via Infotrieve]
  4. Barik, S. (1993) Methods Mol. Biol. 15, 277-286
  5. Beese, L. S., and Steitz, T. A. (1991) EMBO J. 10, 25-33 [Abstract]
  6. Betermier, M., Alazard, R., Lefrere, V., and Chandler, M. (1989) Mol. Microbiol 3, 1159-1171 [Medline] [Order article via Infotrieve]
  7. Bremer, E., Silhavy, T. J., and Weinstock, G. M. (1988) Gene (Amst.) 71, 177-186 [CrossRef][Medline] [Order article via Infotrieve]
  8. Bukhari, A. I. (1975) J. Mol. Biol. 96, 87-99 [Medline] [Order article via Infotrieve]
  9. Chen, J. W., Lee, J., and Jayaram, M. (1992) Cell 69, 647-658 [Medline] [Order article via Infotrieve]
  10. Derbyshire, V., Grindley, N. D. F., and Joyce, C. M. (1991) EMBO J. 10, 17-24 [Abstract]
  11. Engelman, A., and Craigie, R. (1992) J. Virol. 66, 6361-6369 [Abstract]
  12. Fayet, O., Raymond, P. P., Prere, M. F., and Chandler, M. (1990) Mol. Microbiol. 4, 1771-1777 [Medline] [Order article via Infotrieve]
  13. Harshey, R. M., and Cuneo, S. (1986) J. Genet. 65, 159-174
  14. Harshey, R. M., Getzoff, E. D., Baldwin, D. L., Miller, J. L., and Chaconas, G. (1985) Proc. Natl. Acad. Sci. U. S. A. 82, 7676-7680 [Abstract]
  15. Kim, E. E., and Wyckoff, H. W. (1991) J. Mol. Biol. 218, 449-464 [Medline] [Order article via Infotrieve]
  16. Kramer, W., Drusta, V., Jansen, H. W., Kramer, B., Pflugfelder, M., and Fritz, H.-J. (1993) Nucleic Acids Res. 12, 9441-9456 [Abstract]
  17. Kulkosky, J., Jones, K. S., Katz, R. A., Mack, J. P. G., and Skalka, A. M. (1992) Mol. Cell. Biol. 12, 2331-2338 [Abstract]
  18. Kuo, C.-F., Zou, A., Jayaram, M., Getzoff, E. D., and Harshey, R. M. (1991) EMBO J. 10, 1585-1591 [Abstract]
  19. Lavoie, B. D., Chan, B. S., Allison, R. G., and Chaconas, G. (1991) EMBO J. 10, 3051-3059 [Abstract]
  20. Leavitt, A. D., Shiue, L., and Varmus, H. E. (1993) J. Biol. Chem. 268, 2113-2119 [Abstract/Free Full Text]
  21. Lee, J., and Jayaram, M. (1993) J. Biol. Chem. 268, 17564-17570 [Abstract/Free Full Text]
  22. Leung, P. C., and Harshey, R. M. (1991) J. Mol. Biol. 219, 189-199 [Medline] [Order article via Infotrieve]
  23. Leung, P. C., Teplow, D. B., and Harshey, R. M. (1989) Nature 338, 656-658 [CrossRef][Medline] [Order article via Infotrieve]
  24. Maniatis, T., Fritsch, E. F., and Sambrook, J. (1982) Molecular Cloning : A Laboratory Manual , Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
  25. Mizuuchi, K. (1992) Annu. Rev. Biochem. 61, 1011-1051 [CrossRef][Medline] [Order article via Infotrieve]
  26. Mizuuchi, M., and Mizuuchi, K. (1989) Cell 58, 399-408 [Medline] [Order article via Infotrieve]
  27. Mizuuchi, M., Baker, T., and Mizuuchi, K. (1992) Cell 70, 303-311 [Medline] [Order article via Infotrieve]
  28. Nakayama, C., Teplow, D., and Harshey, R. M. (1987) Proc. Natl. Acad. Sci. U. S. A. 84, 1809-1813 [Abstract]
  29. Namgoong, S.-Y., Jayaram, M., Kim, K., and Harshey, R. M. (1994) J. Mol. Biol. 238, 514-527 [CrossRef][Medline] [Order article via Infotrieve]
  30. Piccirilli, J. A., Vyle, J. S., Caruthers, M. H., and Cech, T. R. (1993) Nature 361, 85-88 [CrossRef][Medline] [Order article via Infotrieve]
  31. Parsons, R. L., Prasad, P. V., Harshey, R. M., and Jayaram, M. (1988) Mol. Cell. Biol. 8, 3303-3310 [Medline] [Order article via Infotrieve]
  32. Parsons, R. L., Evans, B. R., Zheng, L., and Jayaram, M. (1990) J. Biol. Chem. 265, 4527-4533 [Abstract/Free Full Text]
  33. Priess, H., Schmidt, C., and Kamp, D. (1987) Phage Mu (Symonds, N., Toussaint, A., van de Putte, P., and Howe, M. M., eds) pp. 277-296, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
  34. Qian, X.-H., Inman, R. B., and Cox, M. M. (1990) J. Biol. Chem. 265, 21779-21788 [Abstract/Free Full Text]
  35. Radstrom, P., Skold, O., Swedberg, G., Flensburg, J., Roy, P. H., and Sundstrom, L. (1994) J. Bacteriol. 176, 3257-3268 [Abstract]
  36. Rowland, S.-J., and Dyke, K. G. H. (1990) Mol. Microbiol. 4, 961-975 [Medline] [Order article via Infotrieve]
  37. Suck, D. (1992) Curr. Opin. Struct. Biol. 2, 84-92
  38. Smith, D., and Pace, N. R. (1993) Biochemistry 32, 5273-5281 [Medline] [Order article via Infotrieve]
  39. Surette, M. G., Buch, S. J., and Chaconas, G. (1987) Cell 49, 253-262 [Medline] [Order article via Infotrieve]
  40. Surette, M. G., Harkness, T., and Chaconas, G. (1991) J. Biol. Chem. 266, 3118-3124 [Abstract/Free Full Text]
  41. van Gent, D. C., Oude Groeneger, A. A. M., and Plasterk, R. H. A. (1993) Nucleic Acids Res. 21, 3373-3377 [Abstract]

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