Membrane-mediated Release of Nucleotide from an Initiator of Chromosomal Replication, Escherichia coli DnaA, Occurs with Insertion of a Distinct Region of the Protein into the Lipid Bilayer*

Jennifer GarnerDagger §, Peter Durrer, Jennifer KitchenDagger , Josef Brunner, and Elliott CrookeDagger par

From the Dagger  Department of Biochemistry and Molecular Biology, Georgetown University Medical Center, Washington, D. C. 20007 and the  Department of Biochemistry, Swiss Federal Institute of Technology Zürich, ETH-Zürich, CH-8092 Zürich, Switzerland

    ABSTRACT
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Abstract
Introduction
Procedures
Results
Discussion
References

DnaA protein, the initiator protein of E. coli chromosomal replication, can be rejuvenated from an inactive ADP form to active ATP-DnaA protein by acidic phospholipids in a fluid bilayer. Cross-linking studies with the photoactivable phospholipid analog 1-O-hexadecanoyl-2-O-[9-[[[2-[125I]iodo-4-(trifluoromethyl-3H-diazirin-3-yl)benzyl]oxy]carbonyl]nonanoyl]-sn-glycero-3-phosphocholine reveal insertion of DnaA protein into the hydrophobic region of the bilayer; this insertion is accompanied by membrane-mediated dissociation of the tightly bound allosteric nucleotides ADP and ATP. Photolabeling of DnaA protein occurred with membrane properties that resembled those needed for reactivation of ADP-DnaA protein; efficient labeling of DnaA protein was observed only when the lipid analog was incorporated into anionic vesicles and the temperature during treatment was above the gel to liquid crystalline phase transition. Predominant hydrophobic photolabeling was localized within a single region of DnaA protein, a region that contains putative amphipathic helices and has been shown to contain information essential for functional interaction with membranes.

    INTRODUCTION
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Abstract
Introduction
Procedures
Results
Discussion
References

Regulation of DnaA protein activity is thought to play a vital role in controlling initiation of chromosomal replication in Escherichia coli (1, 2). DnaA protein, an essential component for initiation, binds to four 9-mer DnaA boxes within the unique 245-base pair origin of chromosomal replication, oriC. After binding to oriC, DnaA protein promotes a local unwinding of the DNA, which leads to the formation of a prepriming complex and the subsequent reactions for DNA synthesis.

Initiation of chromosomal replication occurs at a precise time within the cell cycle (1, 2). Given that levels of DnaA protein remain constant throughout the cell cycle (3), other mechanisms must regulate the protein's activity. Prominent among these in vitro is the influence on the replicative action of DnaA protein by the tight binding of ATP and ADP (Kd of 30 and 100 nM, respectively) (4). Both the ADP and ATP forms of DnaA protein bind oriC (4-6), but only the ATP form is active for succeeding replication steps. The ADP form fails to promote strand opening (4, 7) and therefore is inert for replication (4, 8, 9).

Hydrolysis of bound ATP to ADP occurs gradually (4), is DNA-dependent (4), and may be modulated by a soluble factor (10). The resulting ADP remains tightly bound to DnaA protein, and exchange of bound ADP for ATP, even in the presence of high concentrations of ATP, is extremely slow (4). The initiation activity of ADP-DnaA protein, however, can be restored by treatment in vitro with acidic lipids in a fluid bilayer (11-14). In the presence of anionic lipid vesicles and oriC, DnaA-bound ADP can be exchanged for ATP, rendering DnaA protein active for initiating replication (11, 13).

Recent in vivo studies support the participation of acidic phospholipids in replication from oriC. Disrupted expression of phosphatidylglycerophosphate synthase (pgsA) results in cells that are depleted of acidic lipids and arrested for growth (15). The cells remain viable, and growth can be restored by induced expression of an intact copy of pgsA (16). Growth arrest also can be suppressed in the absence of pgsA expression if a mutation in rnhA is introduced into the cells (17). RNA in hybrid RNA-DNA duplexes persists in the absence of RNase H (rnhA gene product) at multiple sites in the genome and can serve as primers for DNA replication, thus bypassing the requirement for initiation at oriC and, thereby, DnaA protein (18). Such a means of suppressing arrested growth indicates that acidic phospholipids are necessary for normal, oriC-based initiations and implies that the primary cause of growth limitation in acidic phospholipid-deficient cells may be the inability to activate DnaA protein through phospholipid-mediated nucleotide exchange.

While accumulating in vitro and in vivo evidence suggests that lipid components of the cellular membrane play an active role in regulating DnaA protein activity, the interaction between DnaA protein and membranes remains poorly defined. It is known that other soluble proteins have their activities altered by interaction with membrane lipids. For example, protein kinase C is activated by the anionic phospholipid, phosphatidylserine, and has been shown to undergo a conformational change upon phosphatidylserine binding (19). Insertion of a domain of SecA protein into lipid vesicles promotes a conformational change in the soluble portion of the protein (20). This alteration is dependent upon temperature and shows specificity for acidic vesicles. A similar dependence on acidic lipids has been found for membrane binding and insertion of CTP:phosphocholine cytidylyltransferase, indicating that this protein has the same lipid requirements as does DnaA protein for membrane binding. Examination of the contributions of electrostatic and hydrophobic interactions in the activation of CTP:phosphocholine cytidylyltransferase by acidic vesicles indicate that its activation follows a two-step process of membrane binding followed by intercalation of a domain into the bilayer (21). Aspects of the DnaA protein's activation by fluid acidic membranes are similar to those for these other proteins and lead us to propose that functional interaction with phospholipids involves recruitment of DnaA protein to the membrane surface through electrostatic attraction, followed by insertion of a domain into the hydrophobic region of the lipid bilayer.

A domain of DnaA protein necessary for functional interaction with acidic membranes was previously identified; tryptic disruption of this region blocked lipid-mediated release of nucleotide from DnaA protein (22). While the procedure of proteolytically mapping functional fragments permitted the identification of a segment important for membrane interaction, it did not reveal whether membrane-DnaA protein interaction is restricted to the polar head groups or if insertion into the bilayer occurs. Moreover, if insertion occurs, the question arises of whether it is restricted to a discrete region of the protein.

The photoreagent [125I]TID-PC/161 (see Fig. 1A) is an analog of the zwitterionic lipid phosphatidylcholine and can be incorporated into vesicles of desired composition (23). Upon photolysis, the diazirine group, located at the end of the sn-2 aliphatic chain, is converted to a carbene, which rapidly reacts with adjacent molecular bonds. The neighboring environment of the carbene consists largely of acyl chains, and to the extent of their presence, segments of protein that have inserted into the hydrophobic region of the lipid bilayer. This reagent has previously been employed to identify membrane-inserted domains of such proteins as the rabies and vesicular stomatitis viruses' envelope glycoproteins (24), influenza virus hemagglutinin (25), and the myristoylated alanine-rich protein kinase C substrate (MARCKS) and the MARCKS-related protein (26).

Here, [125I]TID-PC/16 was used to investigate the association between DnaA protein and membranes (Fig. 1B). Using vesicles of differing composition and fluidity, we compared the efficiency of [125I]TID-PC/16 photolabeling of DnaA protein with the efficiency of membrane-mediated nucleotide release. Following photolabeling under conditions that supported efficient incorporation of radioactivity into DnaA protein, the distribution of label along the polypeptide chain was examined to determine if DnaA protein was labeled randomly or if label was restricted to a distinct region, an indication that a specific domain inserts into the lipid bilayer.

    EXPERIMENTAL PROCEDURES
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Abstract
Introduction
Procedures
Results
Discussion
References

Materials-- Sources were as follows: HEPES, Tricine, CAPS, ATP, and CNBr (Sigma); [alpha -32P]ATP (3000 Ci/mmol), NEN Life Science Products; Coomassie Brilliant Blue R-250 and G-250, Bio-Rad; trans-paranaric acid, Molecular Probes; 2-(2-nitrophenylsulfenyl)-3-methyl-3'-bromoindolenine (BNPS-skatole), Pierce; 1-stearoyl-2-oleoyl-sn-glycero-3-[phospho-rac-[1-choline]] and 1-stearoyl-2-oleoyl-sn-glycero-3-[phospho-rac-[1-glycerol]], Avanti Polar Lipids, Inc.; trypsin, chymotrypsin, subtilisin, endoproteinase GluC, endoproteinase LysC, and endoproteinase ArgC, Boehringer Mannheim; type HA nitrocellulose filters, Millipore Corp.; Problott PVDF membrane, Applied Biosystems.

Nucleotide Binding to DnaA Protein and Phospholipid-mediated Release of Bound Nucleotide-- DnaA protein in buffer A (50 mM Tricine-KOH, pH 8.25, at 1 M; 2.5 mM magnesium acetate; 0.323 mM EDTA; 8 mM dithiothreitol; 0.0065% (v/v) Triton X-100; 20% (v/v) glycerol) was mixed with ATP (1 µM, radiolabeled where indicated) and incubated (10 min, 0 °C) to produce ATP-DnaA protein. To quantitate bound nucleotide, samples were filtered through nitrocellulose filters previously soaked in buffer G (50 mM Tricine-KOH, pH 8.25, at 1 M; 0.5 mM magnesium acetate; 0.3 mM EDTA; 5 mM dithiothreitol; 0.005% (v/v) Triton X-100; 10 mM ammonium sulfate; 17% (v/v) glycerol). Filters were washed with 5 ml of buffer G and dried, and retained nucleotide was measured by Cerenkov counting in a liquid scintillation counter. For phospholipid-mediated release, ATP-DnaA protein was incubated (10 min, temperature as indicated) with small unilamellar vesicles prior to the filtration through nitrocellulose.

Vesicle Preparation-- Total E. coli lipids, phosphatidylglycerol, phosphatidylcholine, and [125I]TID-PC/16 in CHCl3 were dried under a stream of nitrogen gas and suspended in water or buffer A (as indicated) by sonication (15 min, 0 °C) with 30-50% bursts from a microtip probe sonicator (Branson). Sonicated lipids were centrifuged (30 min, 140,000 × g, 4 °C), and the supernatant, which contained small unilamellar vesicles, was collected. Phospholipids were quantitated by a phosphopolymolybdate colorimetric assay (27).

Photolysis-- For [125I]TID-PC/16 photoactivation, reaction mixtures in Eppendorf tubes were irradiated for 30 s at room temperature (unless otherwise indicated) in a Pyrex vessel mounted approximately 10 cm from a Suss LH 1000 lamphouse (Karl Suss, Waterbury Center, VT) equipped with an Osram HBO 350-watt short arc high pressure mercury lamp (30 milliwatts·cm-2). Under these conditions, greater than 90% of the reagent was photolyzed.

Recrystallization of BNPS-skatole-- BNPS-skatole was dissolved in hexanes (75 °C) and cooled to room temperature. The crystals were dried under nitrogen, dissolved in glacial acetic acid, and used immediately.

Preparation of DnaA Protein-Phospholipid Adducts for Proteolytic Digestion-- Lipid-DnaA protein adducts were electroeluted (recovered in 1-2 ml of 180 mM glycine, 25 mM Tris, 0.1% SDS) from excised bands and concentrated with a Centricon 10 microconcentrator (final volume of 30-40 µl). To remove salts, the samples were diluted with water (2 ml) and concentrated again. Where indicated, additional DnaA protein was added so that the level of protein was sufficient for subsequent resolution and detection of fragments. For samples that were treated with BNPS-skatole, SDS was removed by ion pair extraction (28) prior to cleavage.

Other Methods-- [125I]TID-PC/16 was prepared according to Weber and Brunner (23) at a specific activity of approximately 200 Ci/mmol. DnaA protein was purified (29) from BL21(DE3)pLysS/pKA211, a transformed strain in which high level inducible expression of the wild-type dnaA gene is under the control of a bacteriophage T7 RNA polymerase promoter.2 Transfer of fragments from SDS-polyacrylamide gel to PVDF membrane was performed as described by Matsudaira (30). E. coli phospholipid extraction was performed as described by Kagawa and Racker (31) from strain W3110 (F-, lambda -, IN(rrnD-rrnE)). Protein quantitation was performed as described by Bradford (32). Polyacrylamide gel electrophoresis (19% acrylamide Tris-glycine SDS-PAGE and 16.5% acrylamide Tris-Tricine step electrophoresis were performed as described respectively (33, 34). Amino-terminal sequencing was performed by Dr. A. Fowler (UCLA Protein Microsequencing Facility).

    RESULTS
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Abstract
Introduction
Procedures
Results
Discussion
References

Use of a Radiolabeled Photoactive Lipid Analog to Assess Insertion of DnaA Protein into Membranes-- Neutral vesicles composed of 1-stearoyl-2-oleoyl phosphatidylcholine (PC) or acidic vesicles composed of the 1-stearoyl-2-oleolyl forms of phosphatidylglycerol and phosphatidylcholine (4:1) (PG:PC) were prepared in the presence of [125I]TID-PC/16 (Fig. 1A). Nucleotide-bound DnaA protein was incubated with the vesicles and exposed to light to cross-link the lipid analog with regions of the protein that had inserted into the hydrophobic portion of the membrane bilayer. Lipid-protein adducts were precipitated with a mixture of chloroform and methanol to remove the bulk of unincorporated lipid and solubilized in SDS for further resolution by SDS-PAGE. DnaA protein was identified by Coomassie staining. The band was excised from the gel and analyzed by gamma -counting to determine the efficiency with which neutral versus acidic vesicles were able to photolabel DnaA protein. Alternatively, the cross-linked protein was electroeluted from the gel, and SDS was removed by ion-pair extraction (28). DnaA protein was proteolytically or chemically cleaved, and the resulting fragments were separated electrophoretically. Peptides were visualized by Coomassie staining, and the amount of lipid linked to individual peptides was determined by gamma -counting bands excised from the gel. The peptides were mapped within DnaA protein by N-terminal sequencing (Fig. 1B).


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Fig. 1.   Photolabeling DnaA protein with a radiolabeled phospholipid analog. A, [125I]TID-PC/16 is an analog of phosphatidylcholine. Upon photolysis, the trifluoromethyl diazirine group forms a reactive carbene (23). B, ATP-DnaA protein (in buffer A, 1 µM ATP) was incubated (10 min, 38 °C) with small unilamellar vesicles prepared from phosphatidylcholine or phosphatidylglycerol:phosphatidylcholine (4:1 weight ratio), which also contained [125I]TID-PC/16. The samples were subjected to photolysis, chloroform:methanol (1:2 (v/v); 3 times sample volume) was added, and after a period of time (1 h, 23 °C) samples were centrifuged in a microcentrifuge (15 min, 14,000 × g, 23 °C). The supernatants were decanted to remove unincorporated lipid, and the pellets were dried in vacuo and dissolved (3 min, 95 °C) in SDS-PAGE sample buffer. DnaA protein was further isolated electrophoretically (12% SDS-PAGE) and visualized by staining with Coomassie Brilliant Blue R-250. The bands containing DnaA protein were excised, and the quantity of incorporated lipid was determined by gamma -counting. Alternatively, the samples were prepared (see "Experimental Procedures") for chemical or protease digestion. Proteolytic products were separated by appropriate SDS-PAGE systems (see "Experimental Procedures"). Polypeptides were visualized by staining with Coomassie Brilliant Blue, and covalently attached lipid was detected in excised bands in a gamma -counter. Certain digestion fragments were identified by amino-terminal sequence analysis after transfer of peptides from parallel reactions to PVDF membrane.

Membrane Composition for Efficient Photolabeling of DnaA Protein Parallels That Required for Nucleotide Release-- Zwitterionic phosphatidylcholine vesicles containing [125I]TID-PC/16 were feeble at photolabeling nucleotide-bound DnaA protein. However, cross-linking was 23-fold higher with anionic PG:PC vesicles (Table I). The observed cross-linking was completely dependent upon generation of the reactive carbene, as shown by the low level of lipid cross-linking in the absence of photolysis, even with the acidic PG:PC vesicles. The neutral and acidic vesicles used to photolabel DnaA protein were also assessed for their ability to induce the release of bound nucleotide from DnaA protein. The acidic vesicles, which were necessary for effective photolabeling of DnaA protein, promoted the release of bound ATP. In contrast, a poor release response was observed with the neutral vesicles, which were ineffective in photolabeling DnaA protein (Table I). It is worth noting that the difference in the efficiency for photolabeling DnaA protein was dependent on the surface charge of the vesicles and not the head group charge of the photoactive lipid analog. The extent of photolabeling and specificity for acidic vesicles was the same for ADP-DnaA protein and the ATP form (data not shown).

                              
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Table I
Specificity of lipid headgroups for efficient phospholipid cross-linking matches that needed for nucleotide release
[alpha 32P]ATP-DnaA protein (24.4 µg, in buffer A containing 1 µM ATP) was incubated (10 min, 38°C) with small unilamellar vesicles (137 nmol of phospholipid, in buffer A) composed of PC and PG:PC (4:1 w/w), which also contained [125I]TID-PC/16 (71 cpm/pmol phospholipid and 77 cpm/pmol phospholipid for the PC and PG:PC vesicles, respectively). Release of bound nucleotide was assessed for a portion (41 µl) of each reaction (top three rows), and the remainder (203 µl) was subjected to photolysis (30 s; top two rows), and covalently attached lipid was measured (top three rows). The dependence on DnaA protein for measurable cross-linked lipid was confirmed by gamma -counting a region of the gel that did not stain for protein (blank gel slice). The value of 0% nucleotide release (i.e. 100% nucleotide bound) corresponds to an ATP:DnaA protein molar ratio of 0.46.

E. coli Phospholipid Vesicles Efficiently Photolabel DnaA Protein-- The major lipid components of the inner membrane of E. coli are phosphatidylethanolamine (74%), phosphatidylglycerol (19%), and cardiolipin (4%) (35). E. coli phospholipid liposomes can promote the release of nucleotide from DnaA protein (11-14). Total E. coli lipid vesicles that contained [125I]TID-PC/16 photolabeled DnaA protein at protein:lipid ratios comparable with those needed for membrane-mediated release of bound nucleotide from DnaA protein (Fig. 2). The amount of DnaA protein photolabeled was similar (3.3% of added DnaA protein) to that photolabeled (2.7%) by an equivalent amount of the vesicles composed of the 1-stearoyl-2-oleolyl forms of PG and PC (Table I). As the concentration of E. coli phospholipids was increased, up to 19% of the added DnaA protein was photolabeled (Fig. 2).


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Fig. 2.   DnaA protein is efficiently photolabeled by vesicles composed of E. coli phospholipids. [alpha -32P]ATP-DnaA protein (1.26 µg in buffer A containing 1 µM ATP) was incubated (10 min, 38 °C) with small unilamellar vesicles (in buffer A) prepared with phospholipids extracted from E. coli (strain W3110) and [125I]TID-PC/16 (420 cpm/pmol phospholipid). A portion (<FR><NU>1</NU><DE>6</DE></FR> volume) of each sample was assayed for retention of bound nucleotide (bullet ), and the remainder was subjected to photolysis (30 s) and analyzed for covalently attached lipid (black-square).

Maximal Photolabeling of DnaA Protein Occurs with Membranes in the Fluid Phase-- Although the necessity for anionic polar head groups for efficient cross-linking argues that the photolabeling of DnaA protein is specific, another membrane parameter, fluidity, was also examined. Membrane fluidity is essential for the release of nucleotide bound to DnaA protein (12, 14). The ATP form of DnaA protein was treated at different temperatures with vesicles composed of E. coli phospholipids. The ability of the membranes to dissociate nucleotide from DnaA protein decreased at lower temperatures and occurred only at very low levels when membranes were below their phase transition temperature (see Ref. 14 and Fig. 3, A and B). Correspondingly, photolabeling of DnaA protein, an indication of insertion into the hydrophobic portion of the lipid bilayer, increased as the membranes underwent a transition from a mainly gel-like state to the liquid crystalline phase (Fig. 3C).


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Fig. 3.   Effects of temperature on membrane-DnaA protein interaction. A, [alpha -32P]ATP-DnaA (1.26 µg in buffer A containing 1 µM ATP) was treated (10 min) with total E. coli phospholipid small unilamellar vesicles (33 nmol of phospholipid in buffer A) that contained [125I]TID-PC/16 (520 cpm/pmol of phospholipid). Nucleotide remaining bound to DnaA protein was assessed for a portion (22.6 µl) of each reaction (bullet ). Values are calculated as a percentage of the ATP retained by DnaA protein incubated at the indicated temperature in the absence of phospholipid. B, total E. coli phospholipid small unilamellar vesicles (100 µM in phospholipid) were labeled with trans-paranaric acid (phospholipid:fluorescent probe, 100:1 molar ratio). Fluorescence anisotropy (black-square) at different temperatures was measured as the sample was heated at a rate of 30 °C/h. C, the remainder (113 µl) of the reactions from A were subjected to photolysis (30 s), and photolabeling was measured as described in Fig. 1B.

A Distinct Region of DnaA Protein Is Preferentially Photolabeled-- DnaA protein-lipid adducts were chemically and proteolytically digested and resulting fragments analyzed to determine whether incorporated lipid was found throughout the protein or localized to a single region (Fig. 1B). A discrete region of DnaA protein centered on a possible amphipathic helix of residues Asp357 to Val374 has been shown to be indispensable for membrane-mediated release of nucleotide from proteolytic fragments of DnaA (22). To determine if this essential domain inserts into the bilayer, DnaA protein-lipid adducts were cleaved with BNPS-skatole. This selective hydrolysis at the rare five tryptophans permitted a gross dissection of DnaA protein. BNPS-skatole cleavage of DnaA protein results in two large fragments: a 19-kDa fragment between tryptophans 117 and 288 and a 20-kDa fragment between tryptophan 288 and the carboxyl terminus. Resolution and visualization of the generated polypeptides by SDS-PAGE and Coomassie staining shows that the fragment patterns are the same, whether the protein was treated with neutral or acidic vesicles (Fig. 4A, lanes 1 and 2, respectively). However, reflective of the need for anionic phospholipids for efficient photolabeling of DnaA protein (Table I), only the reaction with acidic vesicles contained radiolabel. BNPS-skatole digestion of the DnaA protein-lipid adducts showed that label was found almost exclusively on a 20-kDa fragment (Fig. 4B, lane 4). Amino-terminal sequence analysis identified this as the carboxyl-terminal BNPS-skatole fragment of DnaA protein (data not shown).


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Fig. 4.   Photolabel resides on the carboxyl-terminal 20-kDa BNPS-skatole fragment of DnaA protein. ATP-DnaA protein (20 µg, in 100 µl of buffer A containing 1 µM ATP) was incubated (10 min, 38 °C) with PC vesicles (22.5 nmol of PC, 1.9 × 107 cpm [125I]TID-PC/16, in 160 µl of buffer A; lanes 1 and 3) and PG:PC vesicles (18 nmol of PG, 4.5 nmol of PC, 1.9 × 107 cpm [125I]TID-PC/16, in 160 µl of buffer A; lanes 2 and 4) prior to photolysis (30 s). DnaA protein, with any covalently attached phospholipid, was precipitated, further isolated by SDS-PAGE, visualized by staining, and eluted from the gel, and concentrated and SDS was removed (see Fig. 1B and "Experimental Procedures"). Following ion pair extraction, DnaA protein was dissolved (final concentration of 0.25 mg/ml) in 60% acetic acid and cleaved (23 °C, 48 h) with freshly recrystallized BNPS-skatole (200 µg). The resulting fragments were separated by 15% SDS-PAGE. Polypeptides were visualized by Coomassie staining (A), and radiolabel was detected with a PhosphorImager (Molecular Dynamics) (B). The band numbered 1 indicates the fragment subjected to N-terminal sequence analysis of peptides transferred to PVDF from a similar reaction. Lane M contains molecular weight standards.

Determining the specific attachment sites of the radiolabeled lipid on the 20-kDa fragment is impractical due to the limited quantity of DnaA protein-lipid adducts generated. Thus, to better define the regions of DnaA protein that were accessible to the hydrophobic photoreagent, full-length DnaA protein-lipid adducts were subjected to other hydrolytic treatments, including cleavage by cyanogen bromide (Fig. 5, A and B) and digestion with a variety of proteases (Fig. 5, C and D). Peptides that could be clearly resolved by electrophoresis were examined for covalently attached radiolabeled lipid and mapped within DnaA protein by amino-terminal sequencing (Table II). Peptides that did not contain radiolabel were also sought, such that the complete sequence of DnaA protein was represented. Therefore, if radiolabel was found on peptides arising from only one or two regions of DnaA protein, its absence in other regions was not due to the inability to generate and obtain peptides for those portions of DnaA protein.


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Fig. 5.   Proteolytic fragments of DnaA protein-phospholipid adducts. A and B, ATP-DnaA protein (18.2 µg in 109 µl of buffer A containing 1 µM ATP) was incubated (10 min, 38 °C) with PG/PC vesicles (71 nmol of PG, 17.6 nmol of PC, in 420 µl of buffer A, 1.5 × 108 cpm [125I]TID-PC/16), subjected to photolysis (30 s) and prepared for digestion (see "Experimental Procedures"). The sample (60 µg) was solubilized in formic acid (70%, 170 µl) and digested (16 h, 23 °C) with CNBr (300 µg dissolved in 70% formic acid). The reaction was diluted with water, and CNBr and formic acid were removed by drying under a stream of N2 gas. Digestion products were dissolved in SDS-PAGE sample buffer. Half of the sample was resolved by 16.5% acrylamide Tris-Tricine step electrophoresis (34) (A), and the remainder was resolved by 19% SDS-PAGE (B). Fragments were visualized by staining with Coomassie Brilliant Blue. Numbered bands indicate fragments subjected to N-terminal sequence analysis of peptides transferred to PVDF in parallel reactions. Lanes 1 and 4 contain 2 µg of undigested DnaA protein; lanes 3 and 6 are molecular weight standards. C and D, ATP-DnaA protein (20.2 µg, in 100 µl of buffer A containing 1 µM ATP) was incubated (10 min, 38 °C) with PG:PC vesicles (18 nmol of PG, 4.5 nmol of PC, 1.6 × 107 cpm of [125I]TID-PC/16) subjected to photolysis (30 s), and prepared for digestion (see "Experimental Procedures"). DnaA protein-lipid adducts (15 µg/digestion) were denatured by heating in SDS (0.1%, 5 min, 100 °C, 50-µl final volume), cooled to room temperature, and digested (23 °C) with the following proteases: subtilisin (0.1 µg) (lanes 7 and 8), chymotrypsin (0.4 µg) (lanes 9 and 10), trypsin (0.48 µg) (lanes 11 and 12), endoproteinase GluC (0.5 µg) (lanes 13 and 14), endoproteinase LysC (0.5 µg) (lanes 15 and 16), endoproteinase ArgC (0.5 µg) (lanes 17 and 18). After 12 h, an equal portion of protease was added, and the digestions continued for an additional 12 h. Fragments were separated on 16.5% acrylamide Tris-Tricine gels, and peptides were visualized by staining with Coomassie Brilliant Blue. Numbered bands indicate fragments subjected to N-terminal sequence analysis of peptides transferred to PVDF in parallel reactions.

                              
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Table II
Lipid cross-linked to DnaA proteolytic fragments
Proteolytic reactions, as in Figs. 4 and 5, were resolved by SDS-PAGE and transferred to PVDF membranes, and indicated fragments were subjected to Edman degradation to identify their amino-terminal sequences. Molecular masses and putative carboxyl-terminal residues were predicted by the polypeptides' migration during SDS-PAGE in conjunction with potential chemical and protease cleavage sites. Covalently attached lipid was measured by gamma -counting of excised bands, and the results presented were normalized to 2500 cpm of digestion reaction loaded onto the gels.

The distribution of radiolabel is presented graphically in Fig. 6. As with the 20-kDa BNPS-skatole fragment, it was not possible to localize within a given fragment where lipid was attached. However, the peptides that contained radiolabel where found to overlap, being derived from a single region of DnaA protein; substantial levels of lipid were found on fragments encompassed by Leu282 and Ser467 (Fig. 6A and Table II). The region of DnaA protein that inserts into the membrane can be further delineated in that fragments 12 (Val211-Met307) and 9 (Ser400-Arg463) both lacked appreciable attached lipid. Thus, the domain of DnaA protein that inserts into the hydrophobic environment of the bilayer is likely to be limited to a sequence that lies between Lys309 and Arg399 (Table II and Fig. 6). In contrast, only low levels of covalently attached lipid were detected throughout the remainder of DnaA protein.


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Fig. 6.   Distribution of photolabel within DnaA protein. Each fragment listed in Table II was assigned a rectangular area proportional to the amount of associated photolabel (cpm from Table II); the horizontal dimension corresponds to the length of the fragment, and the vertical dimension is dependent on the magnitude (cpm) of photolabeling (for this analysis, photolabel associated with a DnaA fragment was assumed to be evenly distributed on the fragment, since its precise location cannot be defined). A, the rectangular areas of the fragments were aligned, additively where applicable, to the linear sequence of DnaA protein (residues 1-467). B, the amount of photolabel associated with each residue was normalized to the number of fragments that contain that residue. The arbitrary units for the magnitude of photolabeling in panels A and B are the same.

    DISCUSSION
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Abstract
Introduction
Procedures
Results
Discussion
References

These studies, which employed a lipophilic photoreagent, indicate that an essential step in the membrane activation of DnaA protein is the insertion of a distinct domain of the protein into the hydrophobic region of the lipid bilayer. As has been observed for the rejuvenation of ADP-DnaA protein (see Refs. 11 and 14 and Table I), insertion requires membranes that contain acidic phospholipids (Table I). Efficient photolabeling was observed with [125I]TID-PC/16 incorporated into acidic vesicles, although the lipid analog possesses a zwitterionic head group (Table I). Interestingly, using [125I]TID-PG/16 in PG:PC (4:1) vesicles further increased the labeling of DnaA (data not shown), suggesting that in addition to creating an acidic surface on the lipid bilayer, the anionic head groups may be involved in direct contact with DnaA protein. This also may be reflected in the observation that clustering of acidic phospholipids in bilayers of mixed head group composition can have a strong influence on the affinity DnaA protein has for nucleotides (36).

The covalently attached lipid was found within a single, distinct region of DnaA protein (Table II and Fig. 6). This segment of DnaA protein includes a portion previously shown to be critical for membrane-mediated release of nucleotide from DnaA protein (22). A recent prediction on the structure of DnaA protein (37) suggests that the photolabeled region of DnaA is largely alpha -helical and lies within the structures termed helices H9 through H12. Helix H9 is thought to be part of the open twisted alpha /beta core, and H12 lies within the known DNA binding domain located at the carboxyl terminus of DnaA protein. Thus, these structures may not participate in the interaction with membranes. Helix H10 is thought to be long and amphipathic, and it has been suggested to be involved in a helix-loop-helix DNA binding motif (37), but H10 is dispensable for specific DNA binding (38). H11 corresponds to the segment identified as important for membrane association (22). Therefore, either or both of these elements and their intervening sequence are strong candidates for the structures that interact with membranes.

A significant level of photolabeling was observed even when the E. coli phospholipid vesicles were below their phase transition temperature (Fig. 3). Such photolabeling of DnaA protein did not emanate from hydrophilic reactive derivatives of [125I]TID-PC/16 generated during vesicle formation, as is evident by the lack of similar labeling when DnaA protein was treated with phosphatidylcholine vesicles prepared with [125I]TID-PC/16 (Table I). Further support for the possibility that the low temperature photolabeling is due to a DnaA protein-membrane association is the ability of acidic vesicles in the gel phase to protect a region of DnaA protein from digestion by trypsin (22). Thus, the photolabeling reflects the capacity of DnaA protein to insert into the hydrophobic region of the bilayer at relatively low temperatures. The fluorescent anisotropy data reveal that at 0 °C, the E. coli total phospholipid vesicles were below their phase transition temperature (Fig. 3B). As such, DnaA protein is either able to intercalate into lipid bilayers that are in the gel phase, or vesicles composed of heterogeneous phospholipids have fluid microdomains into which DnaA protein can insert.

In contrast to the photolabeling that occurred at 0 °C, release of nucleotide from DnaA protein was virtually nonexistent (Fig. 3A). In light of these results, it appears that insertion of DnaA protein into the anionic membrane bilayer, while essential, is not sufficient to catalyze nucleotide release. Previously, it was speculated that the need for elevated temperatures was to provide the membranes with adequate fluidity (11, 12, 14). While this contribution may be necessary, DnaA protein itself seems to have a thermal requirement for dissociation of nucleotide. In support of this hypothesis, additional insertion of DnaA protein (as detected by photolabeling) did not occur at temperatures above the vesicles' phase transition temperature, although the efficiency of nucleotide release continued to improve (Fig. 3). Similarly, a thermal requirement beyond membrane fluidity has been observed for the membrane-induced conformational change in SecA protein (20).

We propose that in the process of membrane-mediated rejuvenation of ADP-DnaA protein, the charge of anionic phospholipids invites close contact between DnaA protein, which is a basic protein, and the surface of the lipid bilayer. Upon association with the membrane, a discrete region of DnaA protein inserts into the hydrophobic portion of the bilayer. The introduction of part of the protein into an aliphatic environment, along with sufficient thermal energy, produces a conformational change in DnaA such that the affinity for bound nucleotide is decreased.

It is not clear at this time if the function of acidic phospholipids is solely for the recruitment of DnaA protein to the membrane or if they play additional roles. DnaA protein does not need to recognize a specific anionic head group, since the requirement for acidic lipids can be met not only by a variety of glycerophospholipids (phosphatidylglycerol, phosphatidylserine, phosphatidylinositol, phosphatidic acid, and cardiolipin) (11, 14) but also by gangliosides (22).

A concerted interaction of DnaA protein, oriC, and acidic phospholipids is necessary for membrane rejuvenation of ADP-DnaA protein (11, 13). However, the level of photolabeling and the distribution of covalently attached lipid on DnaA protein bound to oriC was indistinguishable from that seen for DnaA protein in the absence of oriC (data not shown). Thus, oriC appears not to influence protein-lipid association during reactivation of DnaA. Instead, oriC may stabilize a specific conformation of DnaA protein such that following the release of ADP, DnaA protein is able to bind ATP with high affinity.

It is still unclear whether the DnaA-membrane association is transient or DnaA protein remains bound to the membrane. Further biochemical studies on the interaction of DnaA protein and cellular components, in conjunction with examining the cytolocation of DnaA throughout the cell cycle should help to answer these questions.

    ACKNOWLEDGEMENT

We thank Hiroshi Nakai and Sarah Spiegel for critical reading of this manuscript and Tsutomu Katayama for providing us with the DnaA-overproducing plasmid pKA211.2 We also acknowledge Dr. Audree Fowler (UCLA Protein Microsequencing Facility).

    FOOTNOTES

* This work was supported in part by National Institutes of Health Grant R01-GM49700 (to E. C.), National Science Foundation Grant MCB 9408830 (to E. C.), and Swiss National Science Foundation Grant 31-45951.95 (to J. B.). The UCLA Protein Microsequencing Facility is partially supported by Cancer Center Support Grant CA 16042-20 from NCI, National Institutes of Health (to the Jonsson Comprehensive Cancer Center).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§ Present Address: Dept. of Biochemistry, New York University Medical Center, 550 First Ave., New York, NY 10016.

par To whom correspondence should be addressed: Dept. of Biochemistry and Molecular Biology, Georgetown University Medical Center, 3900 Reservoir Rd. NW, Washington, DC 20007. Tel.: 202-687-1644; Fax: 202-687-7186.

1 The abbreviations used are: [125I]TID-PC/16, 1-O-hexadecanoyl-2-O-[9-[[[2-[125I]iodo-4-(trifluoromethyl-3H-diazirin-3-yl)benzyl]oxy]carbonyl]nonanoyl]-sn-glycero-3-phosphocholine; PC, phosphatidylcholine; PG, phosphatidylglycerol; MARCKS, myristoylated alanine-rich protein kinase C substrate; CAPS, 3-(cyclohexylamino)propanesulfonic acid; BNPS-skatole, 2-(2-nitrophenylsulfenyl)-3-methyl-3'-bromoindolenine; PVDF, polyvinylidene difluoride; Tricine, N-[2-hydroxy-1,1-bis(hydroxymethyl)ethyl]glycine; PAGE, polyacrylamide gel electrophoresis.

2 T. Katayama, unpublished data.

    REFERENCES
Top
Abstract
Introduction
Procedures
Results
Discussion
References

  1. Skarstad, K., and Boye, E. (1994) Biochim. Biophys. Acta 1217, 111-130[Medline] [Order article via Infotrieve]
  2. Messer, W., and Weigel, C. (1996) in Initiation of Chromosome Replication in Escherichia coli and Salmonella (Neidhardt, F. C., Curtiss, R., III, Ingraham, J., Lin, E. C. C., Low, K. B., Magasanik, B., Reznikoff, W. S., Riley, M., Schaechter, M., and Umbarger, H., eds), 2nd Ed., pp. 1579-1601, American Society of Microbiology, Washington, D. C.
  3. Sakakibara, Y., and Yuasa, S. (1982) Mol. Gen. Genet. 186, 87-94[CrossRef][Medline] [Order article via Infotrieve]
  4. Sekimizu, K., Bramhill, D., and Kornberg, A. (1987) Cell 50, 259-265[Medline] [Order article via Infotrieve]
  5. Yung, B. Y. M., and Kornberg, A (1989) J. Biol. Chem. 264, 6146-6150[Abstract/Free Full Text]
  6. Crooke, E., Thresher, R., Hwang, D. S., Griffith, J., Kornberg, A. (1993) J. Mol. Biol. 233, 16-24[CrossRef][Medline] [Order article via Infotrieve]
  7. Bramhill, D., and Kornberg, A. (1988) Cell 52, 743-755[Medline] [Order article via Infotrieve]
  8. Yung, B. Y. M., Crooke, E., and Kornberg, A. (1990) J. Biol. Chem. 265, 1282-1285[Abstract/Free Full Text]
  9. Sekimizu, K., Bramhill, D., and Kornberg, K. (1988) J. Biol. Chem. 263, 7124-7130[Abstract/Free Full Text]
  10. Katayama, T., and Crooke, E. (1995) J. Biol. Chem. 270, 9265-9271[Abstract/Free Full Text]
  11. Sekimizu, K., and Kornberg, A. (1988) J. Biol. Chem. 263, 7131-7135[Abstract/Free Full Text]
  12. Yung, B. Y. M., and Kornberg, A. (1988) Proc. Natl. Acad. Sci. U. S. A. 85, 7202-7205[Abstract]
  13. Crooke, E., Castuma, C. E., and Kornberg, A. (1992) J. Biol. Chem. 267, 16779-16782[Abstract/Free Full Text]
  14. Castuma, C. E., Crooke, E., and Kornberg, A. (1993) J. Biol. Chem. 268, 24665-24668[Abstract/Free Full Text]
  15. Heacock, P. N., and Dowhan, W. (1987) J. Biol. Chem. 262, 13044-13049[Abstract/Free Full Text]
  16. Heacock, P. N., and Dowhan, W. (1989) J. Biol. Chem. 264, 14972-14977[Abstract/Free Full Text]
  17. Xia, W., and Dowhan, W. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 783-787[Abstract]
  18. Kogoma, T., and von Meyenburg, K. (1983) EMBO J. 2, 463-468[Medline] [Order article via Infotrieve]
  19. Orr, J. W., Keranen, L. M., and Newton, A. C. (1992) J. Biol. Chem. 267, 15263-15266[Abstract/Free Full Text]
  20. Ulbrandt, N. D., London, E., and Oliver, D. B. (1992) J. Biol. Chem. 267, 15184-15192[Abstract/Free Full Text]
  21. Arnold, R. S., and Cornell, R. B. (1996) Biochemistry 35, 9917-9924[CrossRef][Medline] [Order article via Infotrieve]
  22. Garner, J., and Crooke, E. (1996) EMBO J. 15, 3477-3485[Abstract]
  23. Weber, T., and Brunner, J. (1995) J. Am. Chem. Soc. 117, 3084-3089
  24. Durrer, P., Gaudin, Y., Ruigrok, R. W. H., Graf, R., Brunner, J. (1995) J. Biol. Chem. 270, 17575-17581[Abstract/Free Full Text]
  25. Durrer, P., Galli, C., Hoenke, S., Corti, C., Gluck, R., Vorherr, T., and Brunner, J. (1996) J. Biol. Chem. 271, 13417-13421[Abstract/Free Full Text]
  26. Vergeres, G., Manenti, S., Weber, T., and Sturzinger, C. (1995) J. Biol. Chem. 270, 19879-19887[Abstract/Free Full Text]
  27. Ames, B. V., and Dubin, D. T. (1960) J. Biol. Chem. 235, 769-775[Medline] [Order article via Infotrieve]
  28. Konigsberg, W. H., and Henderson, L. (1983) Methods Enzymol. 91, 254-259[Medline] [Order article via Infotrieve]
  29. Sekimizu, K., Yung, B. Y. M., and Kornberg, A. (1988) J. Biol. Chem. 263, 7136-7140[Abstract/Free Full Text]
  30. Matsudaira, P. (1987) J. Biol. Chem. 262, 10035-10038[Abstract/Free Full Text]
  31. Kagawa, Y., and Racker, E. (1971) J. Biol. Chem. 246, 5477-5487[Abstract/Free Full Text]
  32. Bradford, M. M. (1976) Anal. Biochem. 72, 248-254[CrossRef][Medline] [Order article via Infotrieve]
  33. Crooke, E., Guthrie, B., Lecker, S., Lill, R., and Wickner, W. (1988) Cell 54, 1003-1011[Medline] [Order article via Infotrieve]
  34. Schagger, H., and von Jagow, G. (1987) Anal. Biochem. 166, 368-379[Medline] [Order article via Infotrieve]
  35. Gennis, R. B. (1989) Biomembranes: Molecular Structure and Function, Springer-Verlag New York, Inc., New York
  36. Mizushima, T., Ishikawa, Y., Obana, E., Hase, M., Kubota, T., Katayama, T., Kunitake, T., Watanabe, E., and Sekimizu, K. (1996) J. Biol. Chem. 271, 3633-3638[Abstract/Free Full Text]
  37. Schaper, S., and Messer, W. (1997) Proteins 28, 1-9[CrossRef][Medline] [Order article via Infotrieve]
  38. Roth, A., and Messer, W. (1995) EMBO J. 14, 2106-2111[Abstract]


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