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Originally published In Press as doi:10.1074/jbc.M505126200 on July 13, 2005

J. Biol. Chem., Vol. 280, Issue 36, 31673-31678, September 9, 2005
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The Plastid Division Protein AtMinD1 Is a Ca2+-ATPase Stimulated by AtMinE1*{boxs}

Cassie Aldridge{ddagger} and Simon Geir Møller{ddagger}§

From the {ddagger}Department of Biology, University of Leicester, Leicester LE1 7RH, United Kingdom and the §Department of Science and Technology, University of Stavanger, N-4036 Stavanger, Norway

Received for publication, May 10, 2005 , and in revised form, July 7, 2005.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Bacteria and plastids divide symmetrically through binary fission by accurately placing the division site at midpoint, a process initiated by FtsZ polymerization, which forms a Z-ring. In Escherichia coli precise Z-ring placement at midcell depends on controlled oscillatory behavior of MinD and MinE: In the presence of ATP MinD interacts with the FtsZ inhibitor MinC and migrates to the membrane where the MinD-MinC complex recruits MinE, followed by MinD-mediated ATP hydrolysis and membrane release. Although correct Z-ring placement during Arabidopsis plastid division depends on the precise localization of the bacterial homologs AtMinD1 and AtMinE1, the underlying mechanism of this process remains unknown. Here we have shown that AtMinD1 is a Ca2+-dependent ATPase and through mutation analysis demonstrated the physiological importance of this activity where loss of ATP hydrolysis results in protein mislocalization within plastids. The observed mislocalization is not due to disrupted AtMinD1 dimerization, however; the active site AtMinD1(K72A) mutant is unable to interact with the topological specificity factor AtMinE1. We have shown that AtMinE1, but not E. coli MinE, stimulates AtMinD1-mediated ATP hydrolysis, but in contrast to prokaryotes stimulation occurs in the absence of membrane lipids. Although AtMinD1 appears highly evolutionarily conserved, we found that important biochemical and cell biological properties have diverged. We propose that correct intraplastidic AtMinD1 localization is dependent on AtMinE1-stimulated, Ca2+-dependent AtMinD1 ATP hydrolysis, ultimately ensuring precise Z-ring placement and symmetric plastid division.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Plastids are essential plant organelles vital for life on earth. They are not formed de novo but arise by binary fission from pre-existing plastids (1-3); plastid division is therefore essential for the maintenance and accumulation of plastid populations within plant cells. Many proteins involved in plastid division are derived from bacterial components conserved from the cyanobacterial origins of higher plant chloroplasts (4-7), including FtsZ, an ancient tubulin-like protein that forms a Z-ring to which other components of the division machinery are recruited (8, 9). The Z-ring is localized to the plastid midpoint (10-12), and correct Z-ring placement is mediated by the coordinated action of the prokaryotic-derived Min proteins. The Escherichia coli minB operon encodes MinC, MinD, and MinE, which together limit Z-ring placement to midcell (13-15): MinC is an antagonist of FtsZ polymerization (14, 15), and topological distribution of MinC is controlled by the ATPase MinD and the topological specificity factor MinE (16-18). ATP-bound MinD recruits MinC to the membrane where the MinD-MinC complex forms a stable inhibition structure at the polar zone of the cell (19-21). Topological specificity is conferred on this complex through interaction of MinE with membrane-bound MinD whereby MinE stimulates MinD ATPase activity, causing MinD to disassociate from the membrane and oscillation to the opposite side of the cell (22-24). The MinD dynamic behavior ensures the lowest MinD-MinC inhibitor complex concentration at midcell, resulting in FtsZ polymerization and appropriate placement of cell division (25).

Higher plants contain MinD and MinE homologs (5-7), and transgenic Arabidopsis plants with reduced AtMinD1 levels show Z-ring misplacement and asymmetric plastid division, whereas AtMinD1 overexpression leads to plastid division inhibition (5, 26). These phenotypes are reminiscent of minicelling and filamenting E. coli deficient in or overexpressing MinD (13), suggesting functional conservation between E. coli MinD and AtMinD1. In agreement with this, the polar localization of E. coli MinD reflects the distinct intraplastidic localization pattern of AtMinD1, which localizes to a single spot or two spots at opposite poles of chloroplasts (7, 27).

AtMinD1 encodes a protein of 326 amino acids and, based on amino acid similarity, belongs to the ParA ATPase protein family containing a Walker A motif involved in the binding and hydrolysis of ATP. Like many ParA proteins, AtMinD1 can dimerize (27), and our studies on the accumulation and replication of chloroplasts 11 (arc11) mutant have demonstrated that the asymmetric plastid division observed in arc11 is due to a A296G missense mutation in AtMinD1 that leads to loss of dimerization and inappropriate intraplastidic localization (27). This suggests that AtMinD1 dimerization and correct intraplastidic localization is in part important for correct Z-ring placement during Arabidopsis plastid division.

Here we have expanded on our studies and demonstrated that AtMinD1 in Arabidopsis is an ATPase but, in contrast to its bacterial counterpart, is activated by Ca2+ rather than Mg2+. We further showed that the AtMinD1-mediated ATP hydrolysis is stimulated by AtMinE1 in the absence of envelope membrane lipids. Mutation analysis reveals that an active site AtMinD1(K72A) mutant leads to loss of ATPase activity and mislocalization within plastids and that, although AtMinD1(K72A) is able to dimerize, the interaction with AtMinE1 is abolished. Our findings reveal that appropriate AtMinD1 intraplastidic localization not only depends on AtMinD1 dimerization but on AtMinE1-stimulated ATP hydrolysis, which in turn governs correct Z-ring placement and symmetric plastid division.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Site-directed Mutagenesis of AtMinD1—A full-length 981-bp AtMinD1 cDNA was PCR amplified using primers MIND/5 (5'-ATCATATGGCGGTCTGAGATTGTTC-3'; NdeI is underlined) and MIND/7 (5'-ATGGATCCTTAGCCGCCAAAGAAAGAGAAGAAGCC-3'; BamH1 is underlined) and cloned into pCRScript (Stratagene) to generate pCRScript-AtMinD1. Two oligonucleotide primers, MIND/20 (5'-TTGCGGTGGTTGTCGTCGCTCCAACACCGCCTTTTC-3') and MIND/21 (5'-CGGAAAAGGCGGTGTTGGAGCGACGACAACCACCGC-3') were designed spanning the AtMinD1 Walker A motif containing single point mutations (underlined) changing the active site lysine (K) at position 72 to alanine (A). PCR amplification, using pCRScript-AtMinD1 as a template and the primer pairs MIND/5-MIND/20 and MIND/21-MIND/7 generated two AtMinD1 overlapping fragments that were joined together by Splicing by Overlap Extension using the flanking primers MIND/5 and MIND/7 to generate AtMinD1(K72A). The 981-bp full-length AtMinD1(K72A) cDNA was ligated into pCRScript (Stratagene) to generate pCRScript-AtMinD1(K72A) and subjected to DNA sequencing to verify the incorporated K72A mutation.

Protein Expression and Purification—Full-length wild-type AtMinD1 and AtMinD1(K72A) cDNAs were subcloned from pCRScript-AtMinD1 and pCRScript-AtMinD1(K72A) into pET14b (Novagen) to generate pET14b-AtMinD1 and pET14b-AtMinD1(K72A) and transformed into E. coli strain BL21 (DE3). 50-ml cultures were grown at 37 °C to a density of A600 = 0.6. Protein expression was induced with 1.5 mM isopropyl {beta}-D-thiogalactopyranoside for 2 h at 37 °C. Both AtMinD1 and AtMinD1(K72A) were insoluble and purified using TALON metal affinity resin (BD Biosciences) under denaturing conditions following the user manual. The purity of the proteins was verified by SDS-PAGE and refolded by dialysis against sodium phosphate buffers (50 mM sodium phosphate, 50 mM NaCl, 0.1 M EDTA, 1.5 mM dithiothreitol, 10% glycerol, pH 7.2) containing 8-0 M urea.

AtMinE1 was expressed as a translational fusion to the C terminus of glutathione S-transferase (GST)1 from pGEX-AtMinE1 (7) in E. coli BL21 (DE3). Protein expression was performed as described above and the soluble AtMinE1-GST fraction purified using glutathione resin (BD Biosciences) following the user manual. The purity was verified by SDS-PAGE. As a control, GST was purified from pGEX-6P (Amersham Biosciences) in E. coli BL21 (DE3).

The 267-bp full-length E. coli minE gene was PCR amplified using primers EcE/3 (5'-ATCATATGGCATTACTCGATTTCTT-3'; NdeI is underlined) and EcE/4 (5'-ATGGATCCTTATTTCAGCTCTTCTGCTTCC-3'; BamH1 is underlined), ligated into pET14b to generate pET14b-EcMinE, and transformed into BL21 (DE3). Protein expression was performed as described above. EcMinE was soluble and was purified under native conditions using TALON metal affinity resin (BD Biosciences) following the user manual and the purity verified by SDS-PAGE.

ATPase Assays—For all assays, the reaction mixture contained 100 mM Tris-Cl (pH 7.4), 50 mM NaCl, 0.1 mM EDTA, 1.5 mM dithiothreitol, 10% glycerol, and 5 mM CaCl2, except for the cation effects assays where 5 mM CaCl2 was replaced with either 5 mM MgCl2, KCl, or MnCl2. In experiments testing the effects of the AtMinD1(K72A) mutation, different cation effects, and pH dependence, 10 µM [{gamma}-32P]ATP (specific activity 10 mCi/mmol) and 0.1 µM AtMinD1 or AtMinD1(K72A) were used, and reactions (20 µl) were incubated for 1 h at 35 °C and stopped with 1 µl of 1 M formic acid. In the time course assays for the double reciprocal plot 10-80 µM [{gamma}-32P]ATP and 0.1 µM AtMinD1 were used, and reactions were incubated at 35 °C and stopped at the specified time. To analyze the effect of AtMinE1 and EcMinE on AtMinD1 ATP hydrolysis, 0.1 µM AtMinE1 (1 pmol), 0.1 µM EcMinE (1 pmol), and 0.1 µM AtMinD1 (1 pmol) were used. Reactions were incubated at 35 °C and stopped after 10 min. In all assays a no-enzyme control was used to assess the background. Samples were spotted onto PEI-cellulose (POLYGRAM CEL 300 PEI; Macherey-Nagel) TLC plates and developed using 0.5 M LiCl and 0.5 M formic acid. Radioactive nucleotides were visualized by autoradiography using x-ray film (Kodak). For quantification purposes plates were scanned using phosphorimaging (Cyclone Storage Phosphor System; Packard).

AtMinD1/AtMinD1(K72A) Localization Analysis—Full-length AtMinD1 and AtMinD1(K72A) cDNAs were PCR amplified using primers MIND/1 (5'-TACTCGAGATGGCGTCTCTGAGATTGTTC-3'; XhoI underlined) and MIND/6 (5'-ATGGTACCGCCGCCAAAGAAAGAGAAGAAGCC-3'; KpnI underlined), removing the termination codon, and cloned into pWEN18 as N-terminal fusions to the YFP to generate pWEN18/AtMinD1 and pWEN18/AtMinD1(K72A). pWEN18/AtMinD1 and pWEN18/AtMinD1(K72A) were transiently expressed in tobacco leaves by particle bombardment (28) and visualized by fluorescence microscopy using a Nikon TE2000U inverted microscope, and image analysis was performed by using OPENLAB software (Improvision, Coventry, UK). The number of fluorescent spots in each chloroplast was recorded for each bombardment.



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  		FIG. 1.
AtMinD1 is an ATPase. A, SDS-PAGE of purified and refolded AtMinD1 and AtMinD1(K72A). B, autoradiography and quantification of AtMinD1 and AtMinD1(K72A) ATPase activity showing that AtMinD1 can hydrolyze ATP, whereas AtMinD1(K72A) shows no significant activity. A no-enzyme control was used to assess background.

 
Yeast Two-hybrid Analysis—Full-length AtMinD1(K72A) was PCR amplified using MIND/5 and MIND/7 and cloned into pGADT7 (GAL4 activation domain; AD) and pGBKT7 (GAL4 DNA-binding domain; BD) (Matchmaker two-hybrid system, version 3; Clontech). The resulting constructs, pGADT7/AtMinD1(K72A) and pGBKT7/AtMinD1(K72A) together with pGADT7/AtMinD1 and pGBKT7/AtMinD1 (27), pGADT7/ARC11 and pGBKT7/ARC11 (27), pGADT7/AtMinE1 and pGBKT7/AtMinE1 (29), and empty vector controls (pGADT7 and pGBKT7), were transformed into HF7c yeast cells in different combinations (Fig. 5). Double transformants were selected on minimal synthetic drop-out medium (S.D. medium) lacking tryptophan (T) (pGBKT7 vectors) and leucine (L) (pGADT7 vectors). Restoration of histidine (H) auxotrophy was used as a marker for protein-protein interactions. Quantitative data were obtained by growing 3-day-old double transformants in liquid S.D.-LT medium in a shaking incubator at 30 °C for 24 h. Cultures were grown to A600 of 1.0 before spotting 5 µl on to S.D.-LT and S.D.-LTH media plates. The plates were incubated for 4 days at 30 °C, and growth was assessed visually. For quantification each yeast spot was suspended in 1 ml of liquid S.D. medium and the A600 of the suspension recorded.

Bimolecular Fluorescence Complementation—Full-length AtMinD1(K72A) cDNA was PCR amplified using primers MIND/1 and MIND/6 and cloned into pWEN-NY (29) as a fusion to the N-terminal fragment of YFP (containing amino acids 1-154 of YFP). The resulting construct pWEN-NY/AtMinD1(K72A) was co-bombarded into tobacco along with pWEN-CY/AtMinD1 (29) (containing amino acids 155-238 of YFP), and bimolecular fluorescence complementation was visualized by fluorescence microscopy. As a positive control tobacco was also co-bombarded with pWEN-NY/AtMinD1 and pWEN-CY/AtMinD1 (29). As a negative control pWEN-CY/AtMinD1, pWEN-NY/AtMinD1, and pWEN-NY/AtMinD1(K72A) were bombarded separately into tobacco cells.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
AtMinD1 Is a Ca2+-dependent ATPase—AtMinD1 contains a putative Walker A motif, and to test whether AtMinD1 is an ATPase a His6-AtMinD1 fusion protein was expressed in E. coli followed by denaturing Co2+ affinity chromatography purification and refolding by dialysis. The purity of refolded AtMinD1 was verified by SDS-PAGE (Fig. 1A) followed by incubation with radiolabeled [{gamma}-32P]ATP at pH 7.4 in the presence of 5 mM CaCl2. TLC analysis and autoradiography showed clear AtMinD1-induced radiolabeled inorganic phosphate (Pi) release compared with a no-enzyme control, revealing that AtMinD1 is an ATPase (Fig. 1B). To ensure the measured ATP hydrolysis was due to AtMinD1 and not a contaminating E. coli ATPase, we generated an active site AtMinD1 mutant by substituting the conserved Walker A lysine for alanine, creating AtMinD1(K72A). AtMinD1(K72A) was expressed and purified as a His6 fusion protein as for wild-type AtMinD1 (Fig. 1A), incubated with [{gamma}-32P]ATP in the presence of 5 mM CaCl2, and analyzed by TLC. Autoradiography revealed no significant Pi release above the no-enzyme control reaction (Fig. 1B), confirming that the K72A mutation inactivates AtMinD1 ATPase activity and that AtMinD1 ATP hydrolysis is mediated through the Walker A domain.



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  		FIG. 2.
Characterization of AtMinD1 ATP hydrolysis. A, increasing amounts of AtMinD1 lead to a linear increase in ATP hydrolysis. B, steady-state kinetics of AtMinD1 ATP hydrolysis. A double reciprocal plot of Pi release versus ATP concentration was used to calculate the Km and Vmax of AtMinD1. C, cation dependence of AtMinD1 ATPase activity. D, the effect of altering pH on the ATPase activity of AtMinD1.

 
To more fully characterize the catalytic activity of AtMinD1, we determined key kinetic parameters of AtMinD1-mediated ATP hydrolysis. The extent of ATP hydrolysis as a function of protein concentration was calculated by measuring Pi release in response to increasing AtMinD1 amounts (Fig. 2A). As seen from Fig. 2A, the extent of ATP hydrolysis was proportional to the amount of input AtMinD1, and in the linear range of enzyme dependence 7 pmol of ATP was hydrolyzed per pmol AtMinD1, which translates to a turnover number of ~2 fmol s-1. Using input [{gamma}-32P]ATP concentrations from 10-80 µM, we quantified Pi release as a function of time in separate time course assays (supplemental Fig. 1). From initial reaction rates a reciprocal plot revealed that AtMinD1 has a Km of 500 µM ATP and a Vmax of 0.2 pmol ATP s-1 (Fig. 2B). These values show that AtMinD1 is a weak ATPase.

Divalent cations are known to influence the activity of ATPases (30), and we investigated whether different cations affected AtMinD1 activity. Surprisingly, Mg2+ had no significant effect on AtMinD1 activity (Fig. 2C) in contrast to the Mg2+-dependent activity of E. coli MinD (31). Similarly, K+ and Mn2+ did not significantly stimulate AtMinD1-mediated ATP hydrolysis (Fig. 2C). However, addition of Ca2+ ions had a dramatic effect leading to an ~5-fold increase in ATP hydrolysis compared with reactions containing Mg2+ (Fig. 2C). Further analysis showed maximum ATP hydrolysis between pH 7.5-8 (Fig. 2D). Combined, these data demonstrate that AtMinD1 is a Ca2+-dependent ATPase.

AtMinE1 Stimulates AtMinD1 ATPase Activity—Because of the low basal ATPase activity of AtMinD1, we analyzed whether AtMinE1 can stimulate AtMinD1-mediated ATP hydrolysis. Equimolar amounts of purified AtMinE1-GST and AtMinD1 were incubated with CaCl2 and [{gamma}-32P]ATP for 10 min. The inclusion of AtMinE1 had a marked effect on the amount of Pi release, showing an ~3-fold increase in ATP hydrolysis compared with AtMinD1 alone and taking into account background Pi release in assays only containing AtMinE1 (Fig. 3), demonstrating that AtMinE1 can stimulate AtMinD1 activity. To ensure the increase in ATP hydrolysis was not due to either inherent AtMinE1 ATPase activity or a contaminating E. coli ATPase, we performed ATPase assays using only purified AtMinE1-GST. Although AtMinE1-GST assays did result in low background ATP hydrolysis (Fig. 3), probably due to contaminating E. coli protein(s), it is clear that the measured increase in ATP hydrolysis in reactions containing both AtMinD1 and AtMinE1 is because of AtMinD1 activity. To further verify this, we performed assays with AtMinD1(K72A) and AtMinE1 that resulted in similar ATP hydrolysis levels as observed for AtMinE1-GST alone (data not shown). As a control for AtMinE1-GST, we performed assays using purified GST that resulted in no Pi release (Fig. 3).



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  		FIG. 3.
The effect of AtMinE1 on AtMinD1 activity. AtMinE1 and E. coli MinE (EcMinE) were added to AtMinD1 ATPase assays in equimolar amounts (1 pmol). The activity of AtMinD1 was analyzed by autoradiography and quantified using phosphorimaging. GST is provided as a control for GST-tagged AtMinE1.

 
To investigate the evolutionary conservation of AtMinE1-stimulated AtMinD1 activity, E. coli MinE (EcMinE) was purified and substituted for AtMinE1 in the ATPase assays. Although we have previously demonstrated that AtMinE1 can function as a topological specificity factor in E. coli (7), experiments revealed EcMinE was unable to stimulate AtMinD1 activity (Fig. 3).

Loss of ATP Hydrolysis Results in Abnormal AtMinD1 Localization—AtMinD1 shows distinct intraplastidic localization patterns localizing into one or two discrete spots at polar regions of ellipsoidal chloroplasts (7, 27). In contrast, the A296G mutation in AtMinD1/ARC11 results in mislocalization, forming large and distorted fluorescent aggregates and/or multiple speckles (27). To test whether an active site mutation affects AtMinD1 localization, translational fusions of AtMinD1 and AtMinD1(K72A) to YFP were created and transiently expressed in tobacco leaves by particle bombardment. As expected, the majority of AtMinD1-YFP (75%) localized into one or two discrete spots (Fig. 4A). In contrast, AtMinD1(K72A)-YFP forms multiple speckles often up to six or more speckles within chloroplasts (Fig. 4). Unlike the mislocalization of AtMinD1(A296G), protein aggregation is rarely observed and AtMinD1(K72A)-YFP frequently forms more speckles per chloroplast than reported for AtMinD1(A296G)-GFP (27), suggesting a different mechanism is responsible for the mislocalization. It is clear from these results that ATP binding/hydrolysis plays an important role in ensuring correct intraplastidic localization of AtMinD1.



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  		FIG. 4.
Intraplastidic localization patterns of AtMinD1 and AtMinD1(K72A) YFP fusion proteins in living chloroplasts. A, AtMinD1 forms one or two spots within the chloroplasts, whereas AtMinD1(K72A) forms multiple speckles. B, quantitative data from a typical analysis of a single particle bombardment of a tobacco leaf show significant difference in the number of spots/chloroplast between AtMinD1 and AtMinD1(K72A). Only chloroplasts in which the number of spots/speckles could be accurately counted are included, and analysis for each bombardment used a range of cells. Scale bars, 5 µm.

 
AtMinD1(K72A) Can Dimerize—Yeast two-hybrid and fluorescence resonance energy transfer analyses have demonstrated AtMinD1 is able to form homodimers (27). In line with this, E. coli MinD undergoes self assembly on phospholipid vesicles forming filamentous polymeric structures (24). Both ARC11/AtMinD1(A296G) and AtMinD1(K72A) exhibit mislocalization within plastids, and we have previously shown that ARC11/AtMinD1(A296G) mislocalization is due to loss of dimerization (27). To investigate whether AtMinD1(K72A) mislocalization was due to loss of dimerization, we performed yeast two-hybrid studies using restoration of histidine auxotrophy as a marker for interaction. In agreement with our previous data (27) we found His auxotrophy restoration in cells expressing AD-AtMinD1 and BD-AtMinD1 (Fig. 5A). Similarly, HF7c expressing AD-AtMinD1(K72A) and BD-AtMinD1 (Fig. 5A) or AD-AtMinD1 and BD-AtMinD1(K72A) (data not shown) showed restoration of His auxotrophy, revealing the K72A mutation does not affect AtMinD1 dimerization. As a negative control we expressed AD-ARC11/AtMinD1(A296G) and BD-AtMinD1 (Fig. 5A) and AD-AtMinD1 and BD-ARC11/AtMinD1(A296G) (data not shown), which showed no growth without His. BD-ARC11/AtMinD1(A296G) and AD-AtMinD1(K72A) (Fig. 5A) and AD-ARC11/AtMinD1(A296G) and BD-AtMinD1(K72A) (data not shown) were also co-expressed, showing no growth without His. To ensure the interactions detected were not because of autoactivation, each construct was co-expressed with the empty vector controls and showed no restoration of His auxotrophy.

To verify that AtMinD1(K72A) can dimerize inside living chloroplasts, bimolecular fluorescence complementation assays were carried out. Separate, non-fluorescent N-terminal (NY) and C-terminal (CY) YFP protein domains can associate to form a functional fluorescent bimolecular complex when brought into proximity by interacting proteins. As previously observed (29), tobacco co-bombarded with pWEN-NY/AtMinD1 and pWEN-CY/AtMinD1 showed clear fluorescence (Fig. 5B). Tobacco co-bombarded with pWEN-NY/AtMinD1(K72A) and pWEN-CY/AtMinD1 also showed clear fluorescence (Fig. 5B), demonstrating that AtMinD1(K72A) is able to dimerize in planta and showing that AtMinD1(K72A) mislocalization is not because of loss of dimerization as observed for ARC11/AtMinD1 (A296G). Tobacco cells bombarded with single vectors (negative controls) showed no fluorescence, as expected.

AtMinD1(K72A) Is Unable to Interact with AtMinE1—The MinE binding site in E. coli MinD is in close proximity to the ATP binding site on {alpha}-helix 7 (32) and lysine 11 within the Walker A motif (equivalent to AtMinD1 lysine 67) interacts with residues within {alpha}-helix 7 competing with MinE (32), suggesting the Walker A motif is involved in mediating MinD-MinE interaction. To test whether the AtMinD1 K72A mutation affects interaction with AtMinE1 we expressed AD-AtMinE1 and BD-AtMinE1 with BD-AtMinD1(K72A) and AD-AtMinD1(K72A), respectively, in HF7c. In contrast to yeast cells expressing AD-AtMinE1 and BD-AtMinD1 (Fig. 5A) or AD-AtMinD1 and BD-AtMinE1 (data not shown) showing growth on His-free media, cells containing AD-AtMinE1 and BD-AtMinD1(K72A) or AD-AtMinD1(K72A) and BD-AtMinE1 (Fig. 5A) showed no restoration of His auxotrophy, demonstrating that the K72A mutation in AtMinD1 abolishes the interaction with AtMinE1. AD-AtMinE1 and BD-ARC11/AtMinD1(A296G) were also co-expressed in HF7c and demonstrated the AtMinD1 A296G mutation has no effect on AtMinD1-AtMinE1 interaction (Fig. 5A). To ensure the interactions detected were not due to AtMinE1 autoactivation, empty BD and AD vector were expressed with AD-AtMinE1 and BD-AtMinE1, revealing no His auxotrophy restoration (Fig. 5A).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The importance of AtMinD1 in plastid division has been shown by studies demonstrating that a disequilibrium of AtMinD1 levels in transgenic plants results in Z-ring misplacement and inappropriate plastid division (5, 26). Although little is known about how AtMinD1 ensures correct Z-ring placement, we recently reported, through the cloning of the disrupted locus in arc11, that AtMinD1 dimerization is important for correct interplastidic localization and central Z-ring positioning during plastid division (27). In this report we have revealed the importance of the biochemical activity of AtMinD1 and shown how the topological specificity factor AtMinE1 modulates both AtMinD1 activity and AtMinD1 localization dynamics. We showed that AtMinD1, in contrast to its bacterial counterpart, is a Ca2+-dependent ATPase and that the Walker A motif is important for both ATPase activity and correct intraplastidic localization. Although an active site AtMinD1 mutant can still dimerize, loss of ATPase activity abolishes its interaction with AtMinE1. Together with the fact that AtMinE1 can stimulate the ATPase activity of AtMinD1, our data suggest that AtMinE1 not only modulates ATP hydrolysis but also ensures correct AtMinD1 localization within plastids during division.

AtMinD1 ATP hydrolysis is mediated through the Walker A domain, as a K72A mutation within this motif leads to a complete loss of ATPase activity (Fig. 1B). The high Km and low Vmax values show that AtMinD1 is a weak ATPase, like the MinD-related protein ParA (33). The weak activity exhibited by AtMinD1 may be because of electrostatic properties of the ATP binding site. In the F1 ATPase (34) and in hydrolases (35), basic amino acids near the ATP {gamma}-phosphate are responsible for stabilization of the transition state negative charge (36); both E. coli MinD and AtMinD1 lack these basic amino acids, possibly explaining the weak ATP turnover. The low basal activity may, however, be an important feature of the AtMinD1 mode of action, as in E. coli MinD membrane dissociation only occurs after stimulation by MinE and this plays an essential role in the MinCDE oscillatory cycle (22, 24). Although AtMinD1 and AtMinE1 oscillatory behavior has not been reported, based on the evolutionary conservation of the division machinery it is probable that a similar mechanism occurs during plastid division.



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  		FIG. 5.
Interactions of AtMinD1(K72A). A, yeast two-hybrid analysis of AtMinD1(K72A), AtMinD1, and AtMinE1 interactions. HF7c yeast cells were co-transformed with different vector combinations. Co-transformed HF7c colonies were grown overnight and spotted onto plates containing S.D.-tryptophan and leucine (S.D.-TL) and plates containing S.D.-tryptophan, leucine, and histidine (S.D.-HTL). Interaction was determined by restoration of histidine (H) auxotrophy and quantative analysis based on the ratio of growth between growth on S.D.-TL and S.D.-HTL, which gives relative strength of interaction. B, biomolecular fluorescence complementation in living chloroplasts showing that both AtMinD1 and AtMinD1(K72A) can dimerize. Scale bar, 5 µm.

 
Although E. coli MinD and AtMinD1 show a high degree of similarity at the amino acid level, we have found significant differences in the functioning of the two proteins, First, a difference in cation dependence of ATPase activity between Ca2+-dependent AtMinD1 and Mg2+-dependent E. coli MinD implies an important functional difference, probably signifying evolutionary adaptation as many plant processes are regulated by calcium. Indeed, studies have suggested a regulatory role for plastidic Ca2+ fluxes (37), and our findings suggest plastidic Ca2+ levels regulate AtMinD1 activity during plastid division. Second, although we have demonstrated that AtMinE1 stimulates the activity of AtMinD1, this stimulation can occur independently of membrane binding (Fig. 3) in contrast to the phospholipid-dependent MinE stimulation of E. coli MinD (31), suggesting functional divergence in the mechanism of AtMinE1-stimulated AtMinD1 ATP hydrolysis during plastid division in Arabidopsis. In addition, purified MinE from E. coli is unable to stimulate AtMinD1 ATPase activity in vitro (Fig. 3), further indicating that at least in part the mechanism of AtMinE1-stimulated AtMinD1 activity is different from that in prokaryotes.

In E. coli, wild-type MinD localizes to the cell periphery, whereas the active site MinD(K16Q) mutant is distributed throughout the cell (24). MinD(K16Q) is, however, still able to bind ATP but cannot bind phospholipids (24). In agreement with this we have shown that the active site AtMinD1(K72A) mutant exhibits aberrant localization patterns, distributed as speckles throughout plastids (Fig. 4). This mislocalization is not because of loss of AtMinD1 dimerization (Fig. 5) as observed for ARC11 (27) but most probably because of lack of interaction with AtMinE1 (Fig. 5A). The loss of interaction between AtMinD1(K72A) and AtMinE1 may either be because lysine 72 is directly involved in the AtMinE1 interaction or because AtMinD1(K72A) is unable to adopt the correct conformation upon binding ATP necessary for AtMinE1 interaction. In E. coli, the {alpha}-helical region of the MinE anti-MinCD domain interacts with MinD {alpha}-helix 7 forming a coiled-coil structure (38, 32), and lysine 11 within the Walker A region (P-loop) competes with MinE for residues within {alpha}-helix 7 (32). MinE-mediated disruption of the non-covalent interaction between lysine 11 and {alpha}-helix 7 changes the lysine 11 side-chain orientation and the P-loop conformation. This transmits an activation signal to the neighboring catalytic domain or to the bound ATP, bringing about ATP hydrolysis (32). This model suggests MinE stimulation is through conformational change in the Walker A motif rather than through direct interaction between MinE and Walker A residues. Based on this model it is unlikely that AtMinE1 interacts directly with the Walker A motif; we favor the theory that a Walker A mutation in AtMinD1 changes the overall conformation, ultimately disabling its interaction with AtMinE1.



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  		FIG. 6.
A working model for AtMinD1 mode of action during plastid division. AtMinD1 dimerizes and binds ATP followed by recruitment of AtMinE1, which stimulates AtMinD1-mediated ATP hydrolysis. ATP hydrolysis by AtMinD1 is dependent on Ca2+.

 
Based on our data we propose a working model for the AtMinD1 mode of action during plastid division in Arabidopsis (Fig. 6). We suggest AtMinD1 undergoes dimerization and binds ATP and that this AtMinD1 dimer complex exhibits low basal Ca2+-dependent ATPase activity. AtMinD1 then interacts with AtMinE1, stimulating ATP hydrolysis. How does this impinge on the function of AtMinD1 in ensuring the correct placement of the Z-ring? In line with the prokaryotic model, dimerized ATP-bound AtMinD1 may bind to the chloroplast envelope before AtMinE1 interaction, which stimulates ATP hydrolysis and membrane release followed by protein relocation. However, in contrast to E. coli, AtMinE1 can stimulate ATP hydrolysis in the absence of envelope lipids; therefore, AtMinE1 can enhance AtMinD1 activity prior to envelope binding. In addition, plants do not harbor MinC, suggesting that AtMinD1 and AtMinE1 modes of action differ from those in prokaryotes. Together with the fact that AtMinD1 is dependent on Ca2+ and not Mg2+, it is clear that AtMinD1 has evolved at the biochemical and cell biological level, presumably to adapt from being a part of cell division machinery in free living prokaryotes to becoming an integral component of the plastid division machinery in higher plants.


   FOOTNOTES
 
* This work was supported by grants from the Biotechnology and Biological Sciences Research Council (SO3/P002, 91/REI18421), The Royal Society, and the European Molecular Biology Organization Young Investigator Program (to S. G. M.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{boxs} The on-line version of this article (available at http://www.jbc.org) contains a supplemental figure. Back

To whom correspondence should be addressed. Tel.: 44-116-252-5302; Fax: 44-116-252-3330; E-mail: sgm5{at}le.ac.uk.

1 The abbreviations used are: GST, glutathione S-transferase; S.D., synthetic dropout medium; AD, activation domain; BD, binding domain; YFP, yellow fluorescence protein. Back


   ACKNOWLEDGMENTS
 
We thank Jodi Maple and Xiang Ming Xu for helpful discussions.



   REFERENCES
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 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
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