Transposase–Transposase Interactions in MOS1 Complexes: A Biochemical Approach

doi:10.1016/j.jmb.2010.11.032

J. Mol. Biol. (2011) 405, 892–908 Contents lists available at www.sciencedirect.com

Journal of Molecular Biology j o u r n a l h o m e p a g e : h t t p : / / e e s . e l s e v i e r. c o m . j m b

Transposase–Transposase Interactions in MOS1 Complexes: A Biochemical Approach Guillaume Carpentier†, Jérome Jaillet†, Aude Pflieger†, Jérémy Adet, Sylvaine Renault and Corinne Augé-Gouillou⁎ Université François Rabelais de Tours, GICC, CNRS, UMR 6239, UFR Sciences & Techniques, Parc Grandmont, 37200 Tours, France Received 15 July 2010; received in revised form 3 November 2010; accepted 16 November 2010 Available online 24 November 2010 Edited by J. Karn Keywords: mariner transposition; transposition regulation; transposase dimerization; transposase/DNA complexes

Transposases are proteins that have assumed the mobility of class II transposable elements. In order to map the interfaces involved in transposase–transposase interactions, we have taken advantage of 12 transposase mutants that impair mariner transposase–transposase interactions taking place during transposition. Our data indicate that transposase– transposase interactions regulating Mos1 transposition are sophisticated and result from (i) active MOS1 dimerization through the first HTH of the N-terminal domain, which leads to inverted terminal repeat (ITR) binding; (ii) inactive dimerization carried by part of the C-terminal domain, which prevents ITR binding; and (iii) oligomerization. Inactive dimers are nonpermissive in organizing complexes that produce ITR binding, but the interfaces (or interactions) supplied in this state could play a role in the various rearrangements needed during transposition. Oligomerization is probably not due to a specific MOS1 domain, but rather the result of nonspecific interactions resulting from incorrect folding of the protein. Our data also suggest that the MOS1 catalytic domain is a main actor in the overall organization of MOS1, thus playing a role in MOS1 oligomerization. Finally, we propose that MOS1 behaves as predicted by the pre-equilibrium existing model, whereby proteins are found to exist simultaneously in populations with diverse conformations, monomers and active and inactive dimers for MOS1. We were able to identify several MOS1 mutants that modify this pre-existing equilibrium. According to their properties, some of these mutants will be useful tools to break down the remaining gaps in our understanding of mariner transposition. © 2010 Elsevier Ltd. All rights reserved.

Introduction Transposases are proteins that have assumed the mobility of class II transposable elements, which *Corresponding author. E-mail address: [email protected]. † G.C., J.J., and A.P. contributed equally to this work. Abbreviations: MBP, maltose binding protein; ITR, inverted terminal repeats; LZ, leucine zipper; SEC, single-end complex; PEC, paired-end complex; OPI, overproduction inhibition; GF, gel filtration; EMSA, electrophoretic mobility shift assay; BSA, bovine serum albumin.

are DNA elements that move around within their host genome via a DNA intermediate. Among them, the mariner group consists of well-characterized transposons belonging to the large ITm superfamily.1 They transpose using a cut-andpaste mechanism, involving several steps: (i) sequence-specific binding of transposase to the inverted terminal repeats (ITRs) present at each end of the transposon; (ii) pairing of the transposon ITRs in a paired-end complex (PEC); (iii) cleavage of both DNA strands at each transposon end; (iv) capture of target DNA; and (v) strand transfer to insert the transposon at a new locus. Two recent

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Transposase–Transposase Interactions

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Fig. 1. Current models for the early stages of mariner transposition. The 1.3-kb Mos1 transposon (light gray) has 28-bp imperfect ITRs at both ends (IRR and IRL; blue triangles) and encodes a transposase (MOS1; pink circle), the sole requirement for Mos1 transposition. Depending on the authors, MOS1 binds to ITR as a monomer (SEC1)3 or dimer (SEC2).2,4 The ends are brought together to form a PEC that could contain either a MOS1 dimer2 or a MOS1 tetramer.5–7

publications have provided information that helps to elucidate the molecular mechanisms underlying mariner transposition. One provides the first report of the crystal structure of a full-length eukaryotic transposase in a PEC with transposon-end DNAs, revealing that the ends are held in a trans arrangement by a dimeric transposase.2 This was done using a soluble variant of the mariner Mos1 transposase, T216A. The authors have demonstrated that T216A is produced and purified as a dimer that binds to one ITR to form a single-end complex (designated SEC2; Fig. 1). In this study, the first-strand nicking takes place in the SEC2. The molecular rearrangements that follow allow the PEC to be assembled by the simple addition of a second ITR within the rearranged SEC2. Secondstrand cleavages take place within the PEC, leading to the excision of Mos1. The second study5 focused on Hsmar1, a reconstructed human mariner. Once again, the authors demonstrated that the Hsmar1 transposase is both produced and purified as a dimer that binds to the ITRs to form SEC2, possibly at each transposon end (Fig. 1), suggesting that the subsequent PEC may contain four transposase molecules. Molecular rearrangements follow the first-strand cleavages, and then second-strand cleavages take place within the PEC, leading to the excision of Hsmar1. According to these authors, we await definite proof of whether single-strand nicks take

place in the SEC2 or in the PEC. A third model had been proposed, in which the mariner transposase binds as a monomer to each ITR to form single-end complexes, designated SEC13 (Fig. 1). Since it is supposed that the transposase dimerizes prior to ITR binding,4 this model (involving SEC1) is gradually abandoned. A simplified view of the various proposed mariner complexes assembly at the early stages of mariner transposition is given in Fig. 1. Comparison of these publications reveals that the two first models have much in common but have several differences. It indicates that mariner transposases interact with their cognate ITR as dimers, as proposed,4,6 thus forming SEC2, which is a complex of primary importance during the course of transposition. It also indicates that major molecular rearrangements are required to progress from SEC2 to the PEC. It seems clear that single-strand cleavages contribute to the reorganization of complexes. The main differences between the studies reported here concern (i) the complex supporting the single-strand cleavages (SEC2 or PEC) and (ii) the stoichiometry of the PEC, which contains a transposase dimer in the first case and a possible transposase tetramer in the second case, suggesting that our understanding of mariner transposition mechanism is far from complete. In addition to the transposase complexes described above (free dimers, SEC2, and PEC), genetic studies 9 have suggested that

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Transposase–Transposase Interactions

Fig. 2. Gel filtration analyses. (a) Analytical gel filtration chromatogram of wild-type MBP-MOS1 at various concentrations, 3 μM (blue), 300 nM (red), and 80 nM (yellow). Two hundred microliters of each protein was applied to a Bio-Silect SEC 250 (Bio-Rad) column at various concentrations in buffer solution [20 mM Tris, 100 mM NaCl, 5 mM MgCl2 (pH 9)]. Two peaks are labeled: a monomer of MBP-MOS1 (83 kDa, eluting at 8.6 min) and a dimer (166 kDa, eluting at 6.5 min). A shoulder is also present (40 kDa, eluting at 10–11 min), which contains degradation products that co-purified with MBP-MOS1. The molecular weight standards (Bio-Rad) used to calibrate the column are positioned to indicate their elution retention time at the top of the chromatogram. The chromatogram presented here is representative of the various tests. The x-axis indicates the retention times in minutes, and the y-axis indicates the absorbance (at 220 nm) in milliabsorbance units. The monomer usually eluted between 8.4 and 8.9 min, whereas the dimer usually eluted between 6.4 and 6.8 min (Supplementary data2). (b) Western blot analysis of the gel filtration fractions. Each fraction was collected, and 20 μL was used to perform SDS-PAGE and Western blot analyses with an anti-MBP monoclonal antibody (New England Biolabs). Relative amounts (in percentage) of monomer (M) versus dimer (D) were extrapolated from this quantification according to the area of the corresponding peaks. Molecular weight standards (Bio-Rad) are indicated on the left.

overexpression of the mariner transposase leads to the formation of inactive oligomers, a phenomenon known as overproduction inhibition (OPI). In this study, we used MOS1 mutants to reach a better understanding of the Mos1 transposition mechanism, especially at the level of MOS1–MOS1 interactions that take place during the process. Twelve mutants had been identified in the previous decade using the yeast two-hybrid system10 in the course of a search for MOS1 variants with impaired transposase–transposase interactions. These mutants allow us to demonstrate that MOS1 obeys the preexisting equilibrium model, with a state of equilibrium existing between monomers and dimers, and only the latter being able to bind to the ITR. Furthermore, we identified mutants that were unable

to promote the PEC assembly, which made them useful for finding out whether first-strand nicking takes place within the SEC2 or the PEC. Finally, we mapped the MOS1 domain involved in the formation of active dimers, and our data suggest that the first helix–turn–helix motif (HTH1) drives two separable activities, dimerization and ITR binding.

Results MOS1 and the equilibrium model Data accumulated to date indicate that MOS1 exists under various conformations (multimers, dimers, monomers), which implies the existence of

895

Transposase–Transposase Interactions Table 1. Transposition activity of the two-hybrid mutants Prot T. rate

WT

F53Y

Q91R

L92H

A93V

S104P

V120A

L124S

W159R

I160T

Y237C

D279G

S302P

1

1

10

0.1

0

0

0.2

0

0

0

1

0

0

11

Transposition rates were established using a bacterial assay, which is the most sensitive method available. Data are expressed relative to the wild-type transposition rate. Prot, protein; WT, wild type; T. rate, transposition rate. Zero means undetectable events.

equilibrium between these various forms. Therefore, we first wanted to confirm the multimeric status of the Mos1 transposase, when the ITR is not present. This was done using gel filtration (GF) analyses, performed with various amounts of MOS1 (50– 1.5 μg, equivalent to concentrations ranging from 3 μM to 80 nM) (Fig. 2a). GF analyses confirmed that MOS1 was purified as a mixture of monomers and dimers. The elution profiles at three different concentrations looked similar, suggesting that the concentration of MOS1 does not affect the equilibrium between monomers and dimers, at least in the range of concentrations used here. The content of each peak was analyzed by SDS-PAGE and Western blotting (Fig. 2b). The dimer-containing fractions were highly enriched with MOS1 transposase, whereas in the monomer-containing fractions, MOS1 transposase was contaminated by MOS1 degradation products (Supplementary data 1). This striking finding was unexpected, as the sizes of the degradation products were not compatible with such a profile, which suggested that these degradation products must be able to dimerize. We will return to this question later. Relative amounts (in percentage) of monomer (M) versus dimer (D) were extrapolated from SDS-PAGE and Western blot quantification taking into account the area of the corresponding peaks. It indicated that the wild-type

protein was present mainly in the dimeric conformation (84%) (versus 16% monomers). Coupled GF and Western blot analyses were used to define the multimerization state of various MOS1 mutants (as done for the wild-type protein) and the changes in MOS1–MOS1 interactions in these mutants. Surprisingly, we verified that all these mutants were still able to form dimers, albeit with a lower efficiency for some of them (see thereafter). The two-hybrid detected mutants were also tested for their ability to promote transposition (Table 1). Two kinds of mutants were then identified: three (F53Y, Q91R, and Y327C) were still able to promote the transposition of a pseudo-Mos1 element, whereas the remaining nine were either partially or totally inactive in transposition assays (L92H, A93V, S104P, V120A, L124S, W159R, I160T, D279G, and S302P). From these data, we assume that MOS1 monomers are able to form two kinds of dimers: (1) active dimers that permit transposition to occur, and (2) inactive dimers that do not support transposition due to inappropriate conformations (Fig. 3). In this model, the two-hybrid mutants can shift the equilibrium in various ways. Mutants that prevent (either directly or indirectly) the formation of active dimers and/or the assembly of the synaptic complex are mutants of the transposition pathway. From the

Fig. 3. MOS1 equilibrium model. Several MOS1 conformations can co-exist in solution in the absence of ITRs. MOS1 monomers may be able to engage in two ways: one that allows MOS1 dimerization for ITR binding and transposition to occur (active dimer; the irregular shape with pink border), and the other allowing MOS1–MOS1 interactions to occur, which prevent transposition (regular ellipse with blue border), leading to inactive conformations. The Mos1 ITR is represented as a black arrow. The formation of inactive complexes involving mariner ITR has been described in the case of Himar1.8 As nothing is known about this possibility with regard to Mos1, and as our aim was to propose an as widely relevant model as possible, this kind of inactive complex is proposed here.

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Transposase–Transposase Interactions

standpoint of transposition, they correspond to lossof-function mutations. On the other hand, mutants that prevent the formation of inactive dimers (or oligomers) are mutants of the inhibition pathway. The latter are not expected to impair the transposition efficiency and may, in some cases, actually improve it, as was observed for F53Y, Q91R, and Y237C. Before we continue, we must define what “active dimer” means. MOS1 active dimers are those allowing SEC2 assembly. Keeping this definition in mind, and because we are mainly interested in Mos1 early transposition steps (i.e., SEC2 and PEC assembly), we searched within the nine mutants unable to promote transposition the ones still able to promote SEC2 assembly. Mutants allowing the assembly of active dimers The ability of the nine loss-of-function MOS1 mutants (derived from yeast two-hybrid screening) to interact with Mos1 ITR was compared to that of the wild-type transposase (Fig. 4), under conditions favorable for SEC2 assembly.4,6 SEC2 was in fact observed with four mutants: V120A (giving a weaker signal than the wild-type transposase), L124S, W159R, and D279G. These mutants displayed differing profiles in GF analyses (Supplementary data 2). V120A, L124S, and W159R gave elution profiles indicating that the equilibrium had been shifted towards the monomer (present at levels of 33%, 52%, and 50% for V120A, L124S, and W159R, respectively), in contrast to that recovered for D279G (7% monomer) (Fig. 4). Thus, these mutants are still able to form dimers, but in less proportion than the wild type. In order to check how the corresponding positions were actually involved in MOS1 activity, all four mutants (V120A, L124S, W159R, and D279G) were checked for their ability to promote first- and secondstrand cleavages, using excision assays12 performed with two plasmids: pBC-3T3 to detect first strandand second strand-specific cleavages, and pBC (native) to detect nonspecific DNA nicking activity. Time-course analyses were first performed using both plasmids and with the MOS1 wild-type transposase as a reference. Our data show that with pBC-3T3, the first-strand specific cleavage occurred quickly, with about 100% of the supercoiled form being converted into the open circle form in less than 30 min (Fig. 5a). Similar assays performed with pBC identified MOS1 DNA nonspecific nicking activity that converted 80% of the supercoiled DNA into open-circle DNA within 2 to 4 h (Fig. 5a and Supplementary data 3). Finally, PEC assembly soon resulted in the detection of the plasmid backbone (a dead-end product of the excision reaction) within 15 to 30 min with pBC-3T3, together with a decrease in the level of open-circle DNA, whereas this latter form accumulated when pBC was used. These different

Fig. 4. Analyses of MOS1 two-hybrid mutants by EMSA. The ability of MOS1 mutants to form SEC2 was evaluated by EMSA using conditions that promote SEC2 formation (30 min at 4 °C, in the presence of competitor DNA and 5 mM MgCl2). Complexes (SEC1, SEC2) and free probe (ITR30) are indicated on the left margin. The protein used is indicated at the top of each lane. For each protein, the percentage of monomer versus dimer (extrapolated from GF and Western blot analyses) is indicated under the corresponding lane.

products of the excision reaction are used as landmarks in analyzing the activity of the four mutants (relaxation of the supercoiled plasmids for firststrand nicking and the appearance of the backbone for full excision, i.e., following PEC assembly). V120A First-strand cleavages give rise to a quite normal pattern for this mutant (Fig. 5b), this step being simply slightly slower than that carried out by the wild protein. Consequently, V120A performed second-strand cleavages and, indeed, the whole excision process less efficiently (slower), which was demonstrated by the later detection of the released backbone (after 2 h, instead of 30 min as for the wild type). This is consistent with the fact that V120A is able to promote transposition, albeit less efficiently than the wild-type protein, indicating that this mutation did not prevent PEC assembly. Together with GF data, this indicates that V120A is in fact a weak dimerization mutant.

Transposase–Transposase Interactions

897

Fig. 5. First-strand nicking analyses. (a) Left panel: Time-course analyses were done using two supercoiled plasmids (pBC-3T3, and pBC) and purified MBP-MOS1. The assays were performed at 30 °C, and the resulting products were loaded onto agarose gel. Only the gel for pBC-3T3 is shown. Molecular weight markers are indicated on the left. The various products are depicted on the right, and their positions on the gel are indicated. First-strand nicking at one transposon end generates an open circular product (OC). Second-strand nicking linearizes the donor (L), yielding the single-end break product. A similar sequence of nicks at the other transposon end yields the double-end break products, which consist of the plasmid backbone (B) plus the excised transposon fragment (Tpson = 3T3). SC, supercoiled donor. Right panel: DNA products detected in the gel are quantified, and the disappearance of the SC donor is used as a measure of the first-strand nicking activity. It is plotted against time, yielding the following curves: first-strand nicking activity of MOS1 on pBC-3T3 (red squares) and on pBC (black triangles). (b–e) Similar analyses were done using the MBP-V120A, MBP-L124S, MBP-W159R, and MBP-D279G mutants as indicated. Color coding and abbreviations are as in (a).

898 L124S and W159R First-strand cleavages were markedly slowed down for both these mutants, since supercoiled DNA was still detected after incubating for 8 and 24 h for L124S and W159R, respectively, reaching the nonspecific cleavage level observed with pBC (Fig. 5c and d). In addition, both were unable to promote full excision, as demonstrated by the lack of backbone release, even at the end of the reaction time course. This is consistent with the fact that L124S and W159R are unable to promote transposition. Taken together, these data indicate that L124S and W159R mutants are able to bind Mos1 ITR, thus forming a SEC2 complex, but that they are unable to promote first-strand cleavage efficiently. This might be due either to a wrong SEC2 organization (with the catalytic triad situated too far from the DNA to be cleaved) or to direct impairment of the catalytic activity. Alternatively, this could indicate that the cleavages (of both the first and second strands) do not occur in the SEC2, but rather in the PEC, the assembly of which is prevented when using L124S and W159R. This latter hypothesis was tested and data are presented thereafter. D279G First-strand cleavages were completely prevented when this mutant was used, since supercoiled DNA was still present after 24 h of incubation (Fig. 5e). Consequently, this mutant was unable to promote full excision, which is consistent with the fact that it is unable to promote transposition. Considering the position of the mutated residue relative to the catalytic triad (D156-D249-D284),2,13 and the close vicinity between the D279 residue and the end of one ITR (in the PEC structure2) on the one hand and the GF filtration data on the other, we surmised that the D279G mutation only impairs catalysis, regardless of PEC assembly. This is supported by the fact that the D279G mutant did not support first-strand cleavage in pBC3T3 (specific cleavages) or in pBC (nonspecific cleavages). This finding is in fact the first evidence to attribute an activity to the YSPD279LAP motif, which is highly conserved among mariner transposases. Finally, our data are consistent with the most recently elucidated structures of MOS1,2,13 according to which the V120 and L124 residues belong to the [113–125] linker region, a subdomain that links together the MOS1 N-terminal ITR binding domain to the C-terminal catalytic domain. This linker is involved in various MOS1–MOS1 interfaces. It was first described as playing a role in the interaction between transposase monomers as a linker belonging to one monomer interacts with the homologous sequence of the second monomer across the dimerization interface found in the crystal of the

Transposase–Transposase Interactions

catalytic domain.13 This role is consistent with the activity of the V120A mutant described here. It has also been shown that in order to play a role in the interaction between transposase monomers, the [113–125] linker interacts with the clamp loop (amino acids 162–189), thus locking the PEC,2 and this is probably prevented by the L124S mutation. Taken together, these data account for the involvement of the [113–125] linker in the molecular rearrangements that allow progressing from SEC2 to the PEC, as proposed.4 Furthermore, the W159 residue is near the ends of the clamp loop (residues F161 and K190), but the distances involved (more than 6 Å) and the respective charges prevent strong interactions.2 In contrast, the W159R substitution brings the modified amino acid (now an Arginine) nearer to the R186 residue (about 5 Å), thus allowing hydrogen bonds to be formed. This, in turn, may disrupt the organization of the clamp loop (amino acids 162– 189) and therefore prevent the PEC assembly. Due to their properties, the four mutants, and particularly L124S, W159R, and D279G, are of main interest in the biochemical and structural studies of mariner transposition. Indeed, they will permit the study of the early steps of mariner transposition, allowing the accumulation of intermediate complexes (SEC2 and/or PEC) that are difficult to stabilize with the wild-type protein (because they are active complexes). MOS1 C-terminal part drives the assembly of dimers that do not support transposition To complete our analyses of two-hybrid loss-offunction mutants, we next focused on the remaining five: L92 H, A93V, S104P, I160T, and S302P. Their inability to form SEC2 with the Mos1 ITR (Fig. 4) is consistent with their inability to promote transposition (Table 1). Using GF analyses, we checked the equilibrium state of these five mutants (Supplementary data 2). They all gave elution profiles indicating that the equilibrium had been shifted further towards the monomer than in the wild type (present at 31%, 38%, 34%, 63%, and 24% for L92H, A93V, S104P, I160T, and S302P, respectively, versus 16% for the wild-type protein) (Fig. 4). Compared to our previous GF data, and particularly to that obtained for L124S (52% monomers), the GF data recovered for L92H, A93V, and S104P, at least, are not sufficient to account for the DNA binding data (no detectable trace of the complex). In fact, these mutants have mutations in one of the helix–turn– helix (HTH2) motifs, which is responsible for the specific recognition of the ITR. On the basis of the known structure of the PEC,2 we searched for data that could account for the biochemical behavior of these mutants. First, we found that only S104 belongs to the sixth α-helix, which is responsible

Transposase–Transposase Interactions

for recognition; both L92 and A93 belong to the fifth α-helix. Moreover, all three residues are (i) too far from the DNA to be involved in the specific recognition of the ITR2 and (ii) do not display properties that could account for DNA strong interactions; these two considerations rule out the possibility that these mutations could directly affect specific ITR recognition. Instead, we propose that any point mutation in the L92H, A93V, and S104P mutants can prevent ITR binding either directly by locally disrupting the folding of the HTH or indirectly, for instance, by steric hindrance of DNA binding. This effect could be strong enough to prevent the detection of traces of the complex by electrophoretic mobility shift assay (EMSA), even after overexposure (not shown). It is still not clear why the two-hybrid screening revealed these positions; we assume that the overall organization of HTH2 must be important for the overall organization of the protein. In contrast, with I160T and S302P, traces of SEC1 were detected. I160T was mainly detected as monomers in GF analyses (63%), suggesting that (i) the amount of dimers was too low to sustain direct SEC2 assembly, and (ii) although binding of the monomers may occur under certain biochemical conditions, this does not support the recruitment of a second transposase subunit, thus preventing SEC2 formation. In contrast, S302P was mainly detected in the form of dimers in GF analyses (76%), but these dimers were unable to interact with the Mos1 ITR to form SEC2. Notably, detection of traces of SEC1 with S302P supports the idea that the binding of the monomers may occur under certain biochemical conditions, without supporting the recruitment of a second transposase subunit, thus preventing SEC2 formation. Given the equilibrium model, S302P must therefore be able to form inactive dimers. Two hypotheses could account for this finding. First, the MOS1 C-terminal region (via the S302 position and/or residues around it) could be directly involved in the assembly of inactive dimers, and the S302P mutation strongly potentiates this function. Second, the S302P mutation could allow new interactions to take place, thus preventing the establishment of active dimers. These hypotheses are not mutually exclusive. The literature contains fragmentary information suggesting that the MOS1 C-terminal region may play a role in the assembly of MOS1 complexes. This was also recently illustrated for HIMAR1, a related mariner transposase.7 We therefore first analyzed the role of the MOS1 C-terminal region in Mos1 ITR binding, using various MOS1 forms truncated at the carboxyl-terminal end of the protein. In the first construct [1–283], the C terminus, including S302 and the third residue (D284) involved in the catalytic triad, was removed while maintaining the YSPDLAP motif; in the second construct [1–253], the YSPDLAP motif and the fifth β-sheet of the RNaseH

899 fold were removed; in the others, the catalytic core was either totally (fragment [1–143]) or partially (fragment [1–169]) removed. The truncated transposases were analyzed using GF assays (to check their monomer/dimer equilibrium) and EMSA (to check their ability to form active dimers via Mos1 ITR binding). MOS1, [1–143], and M[1–169] were found to be able to bind efficiently to the labeled Mos1 ITR, forming the expected SEC2 (Fig. 6a), as previously shown.4 A similar pattern was obtained for the [1–283] truncated transposase. In contrast, complex formation was prevented when the [1–253] transposase fragment was used (Fig. 6a). These data are close to that obtained with HIMAR1, for which eliminating amino acids between the C terminus and position 242 reduced the intensity of HIMAR1/ ITR complexes, while eliminating amino acids between the C terminus and position 180 enhanced the formation of SEC2.7 In an attempt to correlate these striking DNA binding activities to the oligomerization status of the various proteins, the truncated MOS1 mutants were analyzed using GF assays. Despite the fact that all the truncated proteins are shorter than the fulllength protein (MBP)-MOS1, they all displayed a first elution peak with a retention time of ∼ 6.3 min (i.e., eluted faster than the full-length protein; Supplementary data 2), suggesting the formation of oligomers containing more than two protein molecules (Fig. 6b). In addition, Western blot analyses revealed that degradation products were eluted together with the oligomer peak (Fig. 6b and Supplementary data 4), which was not true for the peaks of the full-length dimer (Fig. 2b), thus confirming the formation of aggregates when truncated proteins were used. Such aggregates were also observed when using the truncated HIMAR1 mentioned earlier.7 According to the preexisting equilibrium model and to the EMSA data, some of the protein fragments used here may exist in at least two conformations (oligomers and active dimers for ITR binding, as in the [1–143], [1–169], and [1–283] fragments), whereas the [1–253] fragment only exists in an inactive conformation. These findings are in agreement with previously published data showing that the minimal domain for ITR binding (i.e., active dimer formation) involves the MOS1 first 120 amino acids.4,10,14 The C-terminal part of MOS1 transposase [279–345] is indeed not directly involved in the formation of active dimers, and correct folding of the DDD domain should prevent MOS1 oligomerization. According to data obtained with the S302P mutant, and in order to complete our investigation of the properties of the MOS1 C-terminal domain, we next analyzed the ability of the [279–345] fragment (containing only the last 65 amino acids) to interact with itself in glutaraldehyde fixation assays. Proteins were incubated for 30 min with or without glutaraldehyde, a

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Transposase–Transposase Interactions

Fig. 6. C-terminal domain oligomerization activity. (a) Truncated MOS1 transposases were analyzed by EMSA using Mos1 ITR as a probe and compared to the full-length transposase. Complexes (SEC1, SEC2) and free probe (ITR30) are indicated on the left margin. The protein used is indicated at the top of each lane. (b) Analytical gel filtration chromatogram of wild-type MBP-MOS1 (in black) and truncated MBP-[1–253] (in red) at 3 μM. Axes are as in Fig. 2b. Several peaks were collected (three for MOS1 and four for the [1–253] fragment), and 20 μL was used for SDS-PAGE and Western blot analyses (inserted). The positions of the expected proteins (MBP-MOS1 and MBP-[1–253]) are indicated, as well as those of degradation products (DP1 and DP2). (c) MBP5 and the [279–345] MOS1 fragment were exposed (+) or not exposed (−) to 0.005% glutaraldehyde and then loaded onto SDS-PAGE. After electrophoresis, the gel was subjected to Coomassie blue staining. Molecular weight standards are shown on the left. The proteins used in each lane are indicated at the top of the figure. [279-S302P-345] corresponds to [279–345] containing the S302P mutation. Asterisks indicate the positions of monomer (*) and dimer (**).

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Transposase–Transposase Interactions

Fig. 7 (legend on next page)

902 chemical that covalently cross-links proteins. We confirmed that this fragment is indeed able to dimerize by itself in glutaraldehyde fixation assays (Fig. 6c). In addition, the same fragment, with the S302P mutation, retains its ability to dimerize, supporting the idea that the inactive dimer probably directly involves the last 65 amino acids of MOS1. Taken together, these data revealed a complex pattern of MOS1–MOS1 interactions relying on the C-terminal domain of the protein. All these activities (oligomerization and active or inactive dimer formation) probably compete with MOS1/ITR assembly, thus reinforcing the pre-equilibrium model hypothesis. The MOS1 N-terminal part drives the assembly of active dimers Two-hybrid screening did not detect any mutants in the first 50 amino acids, a region that is known to drive strong protein–protein interactions, in addition to ITR binding.2,7 Data presented here, together with previous published data, suggest that MOS1 active dimers (i.e., those allowing Mos1 ITR binding to occur) are formed through this domain.7,10,14 This could explain why the corresponding mutants were not detected in two-hybrid assays: mutation within the first 50 amino acids makes the active dimers unstable, thus favoring inactive ones formed through the C-terminal part of MOS1. As twohybrid screening is based on the abolition of MOS1–MOS1 interactions, it was not suitable in this situation. The first 116-amino-acid region is now depicted as containing two HTHs involved in Mos1 ITR specific binding.2 One of these HTHs (HTH1, spanning residues 8–53) is described as having two activities, MOS1 dimerization and Mos1 ITR binding, which were also demonstrated by biochemical approaches for HIMAR1.7 We propose that the MOS1 dimerization activity associated with HTH14 (Supplementary data 5) in fact controls MOS1 active dimerization (i.e., the type of interaction that permits ITR binding to occur). According to this

Transposase–Transposase Interactions

hypothesis, we assume that these two functions (dimerization and ITR binding) may be due to distinct parts of the HTH1. This is supported by the recently solved structure of the MOS1 PEC,2 which reveals that the first helix [8–21] is mainly involved in MOS1–MOS1 dimerization, the second [25–36] in MOS1–MOS1 interactions (intra and intermolecular), whereas the third helix [42–53] is involved in ITR binding through the DNA minor groove. This is supported by recent data on HIMAR1, in which a mutation within the corresponding HTH1 (Y12A) results in a transposase able to interact strongly with the ITR, but gives a band in EMSA that migrates faster than the wild-type, therefore indicating that a monomer of Y12A transposase was bound to the ITR DNA instead of a dimer.7 However, the MOS1– MOS1 dimerization we wanted to characterize here drives dimerization prior to ITR binding, but the lack of a solved crystallographic structure corresponding to the MOS1 N-terminal domain (without the ITR) prevents direct observation of MOS1 active dimers. We used functional and biochemical approaches to address this question, with the assumption that the first two helixes are involved in MOS1 active dimerization, whereas the third drives specific interactions between MOS1 and its ITR. If this is true, we postulate that an unrelated but wellcharacterized dimerization domain could replace the first two helixes to drive MOS1 active dimerization. The dimerization domain of the yeast regulatory protein Gal4 (amino acids 41– 10015) and the leucine zipper (LZ) motif of the yeast control general protein GCN4 (amino acids 253–27816) were substituted for the first 34 amino acids of MOS1 to yield the LZ[35–345] and Gal4 [35–345] transposases, respectively. The modified transposases were tested for their ability to trigger Mos1 transposition in bacteria (Fig. 7a). No transposition event was detected using a Mos1 transposase in which the [1–34] region was either truncated ([35–345] MOS1 fragment) or had been mutated by point substitutions (V14P or S28P) that disrupt the local structure of this region (Table 2).18

Fig. 7. In vivo transposition analyses. (a) In vivo transposition assays were performed using pBC-3T3 as the donor and target plasmid. Together with a plasmid encoding the transposase under investigation (pMal-Tpase), pBC-3T3 was cotransformed in bacteria. Transposition products were detected and recovered by promoter targeting. (b) Analyses of MBP-Gal4[35–345] transposition products. After purification, plasmids were digested by HindIII and analyzed by 1% agarose gel electrophoresis. MW, 1-kb DNA step ladder (Promega). T, pBC-3T3 (donor plasmid). Lanes 1 to 5: five independent clones from transposition assay. (c) Chart of transposases under investigation; the MBP N-terminal part is not represented. The full-length MOS1 transposase is shown with both HTH (HTH1 spanning residues 1–55 and HTH2 spanning residues 72–109) and the catalytic domain (green box with a blue border, labeled with DD34D). Truncated and substituted transposases are as follows: [50–345] is a transposase lacking the first HTH, whereas Gal4[50–345] and LZ[50– 345] are transposases in which the first HTH was replaced by the Gal4 dimerization domain (in black) or a leucine zipper (in mauve), respectively; [35–345] is a transposase lacking the first two α-helixes of HTH1, whereas Gal4[35–345] and LZ [35–345] are transposases in which the lacking part of MOS1 was replaced by the Gal4 dimerization domain or a leucine zipper, respectively. (d) Insertion site sequences. Target plasmids recovered in transposition assays were sequenced. Insertion sites are reported, with the duplicated TA (underlined) flanking the new inserted copy. All of them were located in the integration hot spot, the cat gene.17 Only the first four bases of Mos1 are indicated.

903

Transposase–Transposase Interactions Table 2. Transposition rates for N-terminally substituted mutants Transposase No MOS1 V14P S28P R48A R48Q [35–345] [50–345] Gal4[35–345] LZ[35–345] Gal4[50–345] LZ[50–345]

Transposition frequency 0 10− 4 [3 × 10− 5] Undetecteda Undetecteda Undetecteda Undetecteda Undetecteda Undetecteda 6.8 × 10− 9 [7 × 10− 9] 2 × 10− 8 [4 × 10− 9] Undetecteda Undetecteda

Transposition rates were established using a bacterial assay.11 Medians [quartiles] are indicated. a Sensitivity threshold: 2 × 10− 10.

In contrast, transposition events were detected when the LZ[35–345] and Gal4[35–345] transposases were used (Table 2), thus demonstrating that these proteins were indeed able to trigger Mos1 transposition, albeit with reduced efficiency. Synaptic complexes are highly organized structures, and thus it is not surprising to find that their transposition efficiency is altered by changing the way by which the complexes are assembled (dimerization equilibrium, DNA binding domain accessibility to ITR, transposase–transposase interactions in the complex, etc.). Finally, when the whole HTH1 was deleted and replaced by wellcharacterized dimerization domains, the resulting proteins ([50–345] MOS1 fragment, LZ[50–345], and Gal4[50–345]) were unable to promote transposition, even with very low efficiency (Table 2). This therefore supported the idea that HTH1 in fact drives two distinct and separable activities: dimerization and ITR binding. Due to the low transposition efficiency of the LZ[35–345] and Gal4[35–345] transposases, we checked that what we had detected was in fact true transposition. Five independent in vivo transposition events for each recombinant transposase were analyzed by restriction mapping and sequencing. A single fragment of about 4600 bp was expected from the pBC-3T3 donor, which contained a single restriction site for HindIII (in the pseudotransposon). When a pBC-3T3 molecule was used as the target, a pBC-3T3 with an additional 3T3 was recovered. These new plasmids gave rise to two bands after HindIII digestion, the sizes of which depended on the location of the 3T3 insertion (Fig. 7b). Whichever substituted transposase was used, all insertions were due to complete elements and were flanked by a TA duplication (Fig. 7c), which are both expected landmarks for accurate transposition.19

On the other hand, the fact that the third helix is mainly involved in ITR binding was confirmed using MOS1 transposases with point substitutions in this helix (R48A or R48Q). Both were analyzed by transposition assays and EMSA. Transposition assays in bacteria did not allow to detect transposition events, and EMSA failed to show complexes formed between the Mos1 ITR and either the R48A or R48Q mutants (Supplementary data 6). The resulting patterns are similar to those found for the HTH2 mutants (L92H, A93V, and S104A; Fig. 4). These data therefore suggest that the activities (dimerization and ITR binding) of the HTH1 can be separated: the first two helixes being involved in dimerization, and the third in DNA binding. However, this needs further investigation to provide conclusive evidence. First-strand nicking requires PEC assembly As previously mentioned, some of the mutants studied here might be useful for biochemical and structural studies of mariner transposition. Although it was not the main topic of this study, it was tempting to benefit from the L124S properties in order to check whether first-strand nicking takes place in SEC2 or in PEC. Indeed, our data indicated that L124S allows SEC2 assembly but does not support first-strand nicking, suggesting that it requires PEC assembly. This hypothesis was investigated in excision assays using plasmids that contained only one end (pBS-3) and the wild-type transposase. We assume that synaptic complex assembly would be more difficult to achieve under these conditions. If this is true (i.e., cleavages require PEC assembly), then first-strand nicking would not be expected to occur. Data obtained using this plasmid were similar to those obtained with pBC and to those obtained with the L124S mutant, supporting the idea that first-strand cleavage requires the presence of two Mos1 ITRs on the same DNA molecule, with an organization that allows the PEC to assemble (Fig. 8). These data therefore confirm that first-strand cleavages do occur within the PEC, resolving one of the two ambiguities between the two models2,5 presented at the beginning of the study.

Discussion Several models have attempted to explain proteinbinding mechanisms.20 However, there is increasing support for the pre-existing equilibrium model, whereby proteins are found to exist simultaneously in populations with diverse conformations. This model assumes that in its native state, a protein exhibits a series of conformations, and that equilibrium exists between these conformational states.20 This model proposes that MOS1 monomers are able

904

Transposase–Transposase Interactions

Fig. 8. First-strand nicking requires PEC assembly. Timecourse analyses were done using various supercoiled (SC) donor plasmids (pBC-3T3, pBC, or pBS3) and purified MBP-MOS1 or MBP-L124S, as indicated on the left. The assays were performed at 30 °C, and the resulting products were loaded onto agarose gels. Only the part of the gel corresponding to the SC donor is shown. The disappearance of the SC donor is used as a measure of the first-strand nicking activity. For the wild-type transposase acting on the pBC-3T3, the backbone is also seen in this zone, as presented in Fig. 5.

to form two kinds of dimers: (1) active dimers that permit ITR binding and transposition to occur, and/ or (2) inactive dimers that do not support transposition by having inappropriate conformations. In the present work, we have demonstrated that MOS1 behaves as predicted by the pre-equilibrium existing model. There is equilibrium between MOS1 monomers and active and inactive dimers. The work reported here has allowed us to identify several MOS1 mutants that modify this pre-existing equilibrium. For instance, V120A is a mutant that weakly inhibits active dimers formation, whereas I160T is a mutant that strongly inhibits active dimers formation. Alternatively, S302P is a mutant that strongly promotes inactive dimer assembly. In the course of the study, we have also shown that L124S and W159R prevent PEC assembly, whereas D279G mutation results in a MOS1 variant that is unable to promote catalysis. This latter mutant is an interesting one, since it can advantageously be used in biochemical studies to replace the usual mutants of the catalytic triad (D156-D249-D284), giving a protein with unaltered metal ion coordination properties. Taken together, these studies allow us to propose that there is a threshold above which the formation of SEC2 is allowed, as this assembly requires the presence of at least 50% of active dimers. Our data agree with an old hypothesis formulated by Dan Hartl and co-workers in 199621: “overproduction of wild-type subunits shifts a dynamic equilibrium toward the formation of catalytically less active or inactive oligomers.” Indeed, such inactive conformations were first suspected by the way of genetic studies, where some mutants antagonize the activity of the wild-type MOS1 by negative complementation.9,21 Dominant-negative complementation may have a number of possible molecular mechanisms, but the most likely is the formation of oligomers (dimers or higher order multimers) of reduced activity that contain both wild-type and defective subunits. Interestingly, Dan Hartl and co-workers have identified several MOS1

mutants that are inactive in Mos1 germ-line excision and at the same time very efficient in germ-line dominant-negative complementation (R12Q, S99N, A102V, D284E, G292R),22 indicating that they are still able to promote MOS1–MOS1 interactions. Three of them are obviously mutants of the transposition pathway, affecting ITR binding (S99N and A102V) and catalysis (D284E). The two others could be explained in the light of our actual work, asking whether they modify the pre-existing equilibrium or not. This is probably the case for R12Q, which is unable to make dimers through HTH1, thus favoring the assembly of inactive dimers. It is more difficult to conclude about G292R, without the help of the crystal structure,2 in which the G292 residue is close to the end of an ITR, similarly to D279. It is therefore tempting to conclude that G292R prevents catalysis, as does D279G. According to published data,2,5,7 the pre-existing equilibrium model can probably be extended to all the members of the mariner family, suggesting that there is a pathway regulating mariner transposition through the transposase state. We do not yet know which factors regulate the monomer/dimer equilibrium. Interestingly, the transposase concentration (at least in the conditions assayed here) does not appear to affect the equilibrium, but GF analyses failed to distinguish between active and inactive dimers. Under this point of view, it could be proposed that increased transposase concentrations might shift the equilibrium towards inactive dimers, accounting for OPI. Alternatively, the mechanism of action of OPI might depend on the stoichiometry of the PEC. In the case where the PEC contains two transposase molecules forming at first a SEC2 and then recruiting a free ITR, OPI could be the result of ITR saturation. Further investigations will be required to explain this and particularly the question of the impact of mariner ITR on the transposase equilibrium. According to recent findings concerning mariner transposition biochemistry, one question still awaiting incontrovertible proof concerns the kind of

Transposase–Transposase Interactions

905

Fig. 9. Overview of the mapped domains. Two transposases are shown, with domains involved in various MOS1– MOS1 interactions. The first 35 amino acids (part of HTH1; green) are proposed to promote active dimer assembly, whereas the last 65 amino acids (in orange) are proposed to promote inactive dimer assembly. The linker region (in brown) and the clamp loop (in purple) are mentioned, since they are involved in the PEC assembly. The nine mutants (known to prevent or to weaken MOS1–MOS1 interactions) unable to promote transposition are indicated at the top of the figure, together with their related specific motifs. The catalytic triad (D156-D249-D284) is also indicated.

complex (SEC2 or PEC) that allows first-strand nicking. Here, we demonstrate that the complex that supports this activity is in fact the PEC. Richardson et al.2 did not assume this, but our data are in total agreement with theirs. In their elegant study, the authors assayed mixtures of various MOS1 mutants to check whether cleavages occurred in cis or trans within SEC2, and found at first that a mixture of DNA-binding and catalytic mutants was unable to cleave Mos1 ITR (see Ref. 2; Fig. 7). In the light of our findings, this simply reflects the fact that this mixture does not allow PEC assembly to occur, thereby preventing the occurrence of first-strand cleavage. Their other data can also be interpreted with this assumption, giving best-matching values. Completing the schema previously proposed,5 we therefore conclude that the early steps in the mariner transposition pathway are as follows: transposase dimerization→SEC2 assembly→PEC assembly→ first-strand nicking→complex reorganization→ second-strand nicking→excision. What remains unclear is whether the cleavages at both ends are concerted. Evidence has been published indicating that second-strand cleavage does not occur at the same time at both ITRs, leading to the accumulation of linear (single-end break) products in excision assays.5,8,10,12 The mechanism of second-strand cleavage (i.e., at each ITR) in mariner transposons is one of the most significant gaps in our understanding of transposition. Indeed, almost all DDE family cut-and-paste transposons thus far examined use a DNA hairpin mechanism,23–25 allowing one subunit of the homomultimeric protein complex to perform double-strand cleavage at one transposon end. One well-known exception is Tn7, where two different subunits of the heteromultimeric protein complex perform double-strand cleavage at one transposon end.26 Mariner transposons are unusual because they use a homomeric transposase but lack

the DNA hairpin intermediate.3,13 The complex reorganization necessary to promote second-strand cleavage suggests that transposase subunits, which have already undergone the first single-strand cleavages at both ends, subsequently move to initiate the second-strand cleavages. Overall, this pathway is consistent with the paradigm established for other well-characterized members of the DDE family of transposases, where synapsis is a prerequisite for catalysis.27–29 Synapsis is believed to prevent single-strand genotoxic cleavages, thus protecting the host genome. However, MOS1 is able to cleave a plasmid that does not contain an ITR, and one can imagine that in a genomic context, the amount of ITR is negligible, thus tending to promote DNA damage. This nonspecific activity is not shared by all the transposases of the mariner family; for instance, the Hsmar1-ra transposase5 displays only specific DNA cleavages. This raises the question of which MOS1 conformer is able to promote nonspecific DNA cleavage. Preliminary data obtained using the I160T mutant (mainly a monomer) suggested that the MOS1 monomer should promote nonspecific DNA cleavages (our unpublished data). These data are supported by the fact that Hsmar1-ra transposase has only been detected in dimer form, but this hypothesis remains to be proved. Our data also confirm that interactions within MOS1 dimers are sophisticated and result from (i) active dimerization due to the N-terminal domain, which leads to ITR binding; (ii) inactive dimerization carried by part of the C-terminal domain, which prevents ITR binding; and (iii) oligomerization (Fig. 9). Active dimer assembly requires part of the HTH1 (the first 35 amino acids), which should be considered as the MOS1 active dimerization domain. Replacement experiments (that allowed demonstrating this point) deal with the idea that proteins are modular molecules in which each module drives

906 a specific activity. The fine-mapping of the modular organization of a particular protein is of main interest for people who engineer proteins for biotechnical purposes. On the other hand, inactive dimers are nonpermissive in organizing complexes that produce ITR binding, but the interfaces (or interactions) supplied in this state could play a role in the various rearrangements needed during transposition. Finally, oligomerization is probably not due to a specific MOS1 domain, but rather the result of nonspecific interactions resulting from incorrect folding of the protein. These interactions take place when the catalytic core is partially deleted or absent, suggesting that the MOS1 catalytic domain is a main actor in the overall organization of MOS1, including that of the N-terminal domain. Eukaryotic and prokaryotic transposases display very different properties regarding their ability to interact with their ITRs. When produced and purified as a recombinant protein, the wild-type Tn5 transposase is monomeric and unable to interact with its ITR in vitro. To allow this binding to occur, either the C-terminal domain must be removed or the whole protein must be mutated (which leads to an active system in vitro).30 In contrast, mariner transposases can be produced and purified as active dimeric recombinant proteins. One of the main differences between the two systems is that in prokaryotes (e.g., Mu, Tn10, and Tn5), transcription, translation, and, in some cases, DNA binding are linked phenomena. In Tn5, the transposase interacts with its ITR upon translation and before the synthesis of the C-terminal domain, thus promoting in situ SEC and PEC assembly. In addition, SECs with full-length Tn5 transposase are never detected in vitro,27 suggesting that Tn5 transposase spontaneously dimerizes after ITR binding. In contrast, in eukaryotes, mariner transposases are translated in the cytoplasm and need to return to the nucleus before they can undergo ITR binding, synapsis, and transposition. ITR binding also requires transposase dimerization. Together with our previous remarks about the activities of monomers versus dimers, mariner transposase dimerization could take place in the cytoplasm, just after synthesis. In this case, the dimers would have to return to the nucleus. If the nuclear translocation of dimers and/or monomers is a controlled mechanism, this would provide a way of reducing monomer levels in the nucleus and, thus, reducing the genotoxicity of the transposase.

Materials and Methods DNA constructs The 12 MOS1 mutants used, F53Y, Q91R, L92H, A93V, S104P, V120A, L124S, W159R, I160T, Y237C, D279G, and S302P, have been described elsewhere.10,11 The only

Transposase–Transposase Interactions difference was that they were cloned in the pMal-c2 system (New England Biolabs) at the SnaBI and HindIII restriction sites, as had previously been done for the wild-type MOS1.6 The resulting plasmids encoded MBP-MOS1 variants. The truncated transposases, corresponding to MOS1 fragments [1–143], [1–169], [1–253], [1–283], [279–345], and [35–345], used in this study have also been described previously.14 Genes encoding the yeast Gal4 dimerization domain (SPKTKRSPLTRAHLTEVESRLERLEQLFLLIFPREDLDMILKMDFLQDIKALLTGLFVQD; accession no. P04386) and the yeast GCN4 leucine zipper (ILPILEDKVEELLSKNYHLENEVASLKKLVGES; accession no. P03069) were synthesized by Entelechon (Germany). They are flanked by classic flexible linkers (MPGGGGSGGGGS// GGGGSGGGGSPG) and SmaI restriction sites. These domains were cloned into the pMal-c2 (New England Biolabs) vector at the SnaBI restriction site, giving pMalGal4 and pMal-LZ. These vectors were then used to insert either the [35–345] or [50–345] part of MOS1 at the EcoRI and HindIII restriction sites. The final constructs were designated pMal-Gal4[35–345], pMal-Gal4[50–345], pMalLZ[35–345], and pMal-LZ[50–345], and they encode MBPGal4[35–345], MBP-Gal4[50–345], MBP-LZ[35–345], and MBP-LZ[50–345], respectively. Transposition assays in bacteria were performed as described,11 with a sensitivity threshold of about 2 × 10− 10 events per assay. Production and purification of proteins The wild-type Mos1 transposase (MOS1), the MOS1 mutants, and the MOS1 fragments were produced and purified as fusion proteins linked to MBP (387 amino acids), using the pMal-c2 system (New England Biolabs) and following the manufacturer's instructions. Protein quantification was done by the Bradford method, versus a bovine serum albumin (BSA) standard. MBP5 (43 kDa) was purchased from Promega. Here, we use the terms “oligomers” or “oligomerization” to refer to inactive complexes containing several (at least two) transposase molecules, whereas the terms “multimers” or “multimerization” are used in reference to active complexes containing several (at least two) transposase molecules. Excision assays Basic excision reaction mixtures contained 10 mM Tris– HCl (pH 9), 50 mM NaCl, 20 mM MgCl2, 0.5 mM EDTA (ethylenediaminetetraacetic acid), 0.5 mM DTT, 100 ng of BSA, 600 ng of supercoiled pBC-3T3, and 80 nM MBPMOS1, in a volume of 20 μL. The reactions were allowed to proceed at 30 °C for various periods, as indicated, before 2 μL of stop solution (loading buffer) was added. The reaction mixtures were then incubated at 65 °C for 10 min. Each reaction was analyzed by overnight electrophoresis at 2.7 V/cm on a TAE-buffered 1% agarose gel. After electrophoresis, the gel was stained with ethidium bromide (0.3 mg/mL) and photographed on a transilluminator. ImageGauge 4.22 software (Fujifilm) was used to quantify the bands. Three kinds of plasmids were used: pBC-3T3 (containing a whole pseudo-Mos1) to monitor the usual excision reaction, pBS-3 (containing only a single 3′ ITR, without the other sequences that constitute pseudoMos1) to monitor cleavages occurring at a single end, and pBC (native) to monitor the nonspecific cleavages.

907

Transposase–Transposase Interactions Electrophoretic mobility shift assays

Insertion site analysis

Binding reactions were carried out in 50 mM NaCl, 0.5 mM DTT, 10 mM Tris (pH 9), 5% glycerol, 1 μg of sonicated salmon sperm DNA, and 100 ng of BSA. Each 20-μL reaction mixture contained 0.2 pmol of 32P-labeled, double-stranded oligonucleotide containing the 3′ITR sequence [ITR30]6 and 10 pmol of purified wild-type MOS1, mutants, or fragments (in fusion with MBP). Reactions were carried out in 5 mM MgCl2 for 30 min at 4 °C. Reaction products were separated using 6% native polyacrylamide/0.25× TBE gels (30:0.93 acrylamide/ bisacrylamide) containing 5% glycerol. Gels were run at 200 V for 3 h and then autoradiographed.

Five tetracycline-resistant colonies from independent transposition assays were taken, and plasmids were extracted (Promega DNA purification system), diluted, and used to transform DH5α-competent cells. This step was needed to purify the plasmids conferring the tetracycline-resistant phenotype. For each transformation, one tetracycline-resistant colony was taken, and plasmids were extracted and analyzed using HindIII digest. Transposition signatures were verified by DNA sequencing, using primers anchored in the cat-coding gene of the pBC.17

Glutaraldehyde fixation assays Proteins were prepared in 20 mM phosphate buffer (pH 8), as previously described.6 Reaction mixtures (16 μL) containing 10 to 20 pmol of truncated transposase, either alone or with 0.005% (v/v) glutaraldehyde, were incubated for 30 min at 25 °C. The reactions were stopped by adding 3 μL of 6× SDS-PAGE loading buffer, before boiling for 10 min and loading onto discontinuous 4–8% SDS-PAGE, together with the protein standards (Promega). Gels were run at 30 mA for 1 h and then stained with Coomassie blue. Size-exclusion chromatography experiments The Bio-Silect SEC 250 column (range, 10–300 kDa; BioRad) equilibrated with Tris–HCl 20 mM (pH 9), NaCl 100 mM, MgCl2 5 mM was used at a flow rate of 0.8 mL/ min. Data acquisition and analyses were performed with the ChromQuest 4.2 software. The molecular weight calibration curve was obtained by loading a mix of known proteins (Gel Filtration Standard #151-1901, BioRad) containing 50 μg of thyroglobulin (bovine, 670 kDa), 50 μg of γ-globulin (bovine, 158 kDa), 50 μg of ovalbumin (chicken, 44 kDa), 25 μg of myoglobin (horse, 17 kDa), 5 μg of vitamin B12 (1.35 kDa), and 50 μg of BSA (66 kDa, Promega) in 200 μL of 20 mM Tris–HCl (pH 9), 100 mM NaCl, 5 mM MgCl2. All proteins except thyroglobulin (which was in the void volume) were detected at the end of runs by UV light (220 nm). As expected, the curve fitted a class III polynomial equation: Y = aX3 + bX2 + cX + d, with X = log(molecular weight; Da) and Y = retention time (min). For protein analyses, 200 μL of buffer [20 mM Tris–HCl (pH 9), 100 mM NaCl, 5 mM MgCl2] containing, unless otherwise specified, 50 μg of each purified recombinant protein was applied to the column, and proteins were detected after the runs by UV light (220 nm). Each peak was collected, and 20 μL was analyzed by SDSPAGE and Western blot using an anti-MBP monoclonal antibody (#E8032, New England Biolabs). The HRP luminescence of the secondary antibody (HRP goat antimouse, #UP446330, Interchim) was measured using the LAS-4000 machine. The intensity of each band was calculated from a nonsaturated picture with the Multi Gauge V3.0 Software (Fuji). For each protein, relative amounts (in percentage) of monomer versus dimer were extrapolated from this quantification based on the area of the corresponding peaks.

Acknowledgements We would like to thank Dr. David Finnegan who kindly gave us 12 MOS1 mutants obtained using the yeast two-hybrid system. Many thanks to M.V. Demattei and M. Genty for their excellent technical assistance. Dr. M. Ghosh revised the English text. Funding was provided by the Université François Rabelais (Tours); the Centre National de la Recherche Scientifique (CNRS); the Ministère de l'Education Nationale, de la Recherche et de la Technologie; the European Commission (Project SyntheGeneDelivery, no. 018716); the Association Française contre les Myopathies (AFM grant no. 11468); the Région Centre (InhDDE project); and the Groupement de Recherche CNRS 2157.

Supplementary Data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/ j.jmb.2010.11.032

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