Movement and equipositioning of plasmids by ParA
filament disassembly
The MIT Faculty has made this article openly available. Please share
how this access benefits you. Your story matters.
Citation
Ringaard, van Zon, Howard, and Gerdes (2009). Movement and
equipositioning of plasmids by ParA filament disassembly.
Proceedings of the National Academy of Sciences of the United
States of America 106:19369-19374. Copyright ©2009 by the
National Academy of Sciences
As Published
http://dx.doi.org/10.1073/pnas.0908347106
Publisher
National Academy of Sciences
Version
Final published version
Accessed
Thu May 26 04:00:51 EDT 2016
Citable Link
http://hdl.handle.net/1721.1/58578
Terms of Use
Article is made available in accordance with the publisher's policy
and may be subject to US copyright law. Please refer to the
publisher's site for terms of use.
Detailed Terms
Movement and equipositioning of plasmids
by ParA filament disassembly
Simon Ringgaarda,b, Jeroen van Zonc,d, Martin Howarde, and Kenn Gerdesa,1
aCentre
for Bacterial Cell Biology, Institute for Cell and Molecular Biosciences, Newcastle University, Newcastle upon Tyne NE2 4HH, United Kingdom;
of Biochemistry and Molecular Biology, University of Southern Denmark, Campusvej 55, 5230 Odense M, Denmark; cCentre for Integrative
Systems Biology, South Kensington Campus, Imperial College London, London SW7 2AZ, United Kingdom; dDepartment of Physics, Massachusetts
Institute of Technology, 77 Massachusetts Avenue, Cambridge, MA 02139; and eDepartment of Computational and Systems Biology, John Innes
Centre, Norwich NR4 7UH, United Kingdom
bDepartment
Edited by M. J. Osborn, University of Connecticut Health Center, Farmington, CT, and approved September 25, 2009 (received for review July 27, 2009)
cytoskeleton 兩 DNA segregation 兩 mathematical modeling 兩
ParA ParB 兩 pulling
I
n bacteria, it has been difficult to analyze how chromosomes are
segregated. To gain insight into the problem, partitioning (par)
loci encoded by plasmids have been used extensively as model
systems. Type I par loci encode 3 components: a Walker Box
ATPase (ParA), a DNA binding protein (ParB), and one or more
cis-acting DNA regions where the proteins act (parC). The ParB
proteins bind site-specifically to their cognate parC sites to form a
‘‘partition complex.’’ ParB also interacts with the cognate ParA
protein and thereby functions as an adaptor between ParA and
parC DNA. Thus, the parC region at which the segregation apparatus congregates is functionally equivalent of a eukaryotic centromere. Interestingly, ParA ATPases form helical structures that
dynamically relocate over the nucleoid (1–6). ParA relocation but
not the formation of filamentous structures depends on the presence of ParB bound to parC (1, 2, 4, 6). The presence of helical ParA
structures in living cells is consistent with the ability of the proteins
to polymerize in vitro (4, 6–13).
Purified ParAs of Thermus thermophilus and plasmid pSM19035
both dimerize in the presence of ATP (6, 14), whereas ParA of P1
dimerizes also without nucleotide (13). The ParA-ATP dimers bind
cooperatively and nonspecifically to DNA. Thus, the in vitro DNA
binding activity of ParA proteins is consistent with the nucleoid
association seen in vivo (1, 8). In all cases investigated, ParB
stimulates ParA ATPase activity, either on its own or in the
presence of its cognate centromere site (6, 9, 11, 15).
We showed previously that the type I par2 locus of pB171, on
average, distributes plasmids regularly over the bacterial nucleoid
(7). Our observations raised the possibility that the dynamic ParA
filaments generate the mechanical force that move and position
plasmids within the cell.
Here we analyze the relative movements of ParA and plasmids
in single cells. We find that ParA dynamics and plasmid movements
are intimately correlated in a pattern indicating that the partition
www.pnas.org兾cgi兾doi兾10.1073兾pnas.0908347106
complex stimulates disassembly of ParA structures. Strikingly,
plasmids consistently migrated in the wake of disassembling ParA
in manner suggesting that retracting ParA structures move plasmids
by a pulling mechanism. We used mathematical modeling of ParA
dynamics and plasmid movement to see if a simple pulling model
could yield the observed plasmid movements and distributions, and
ParA dynamics. The modeling reliably generated the observed
distributions, provided the rate of detachment of a plasmid from a
filament was filament-length dependent, a prediction that we
verified experimentally. In vivo data showed that perpetual cycles
of ParA assembly/disassembly continuously moved plasmids relative to each other, which results in a time-average equidistribution
of plasmids, as predicted by the mathematical model. Our observations and computations elucidate how the type I par locus of
Escherichia coli plasmid pB171 moves and positions plasmids.
Results and Discussion
Visualization of Plasmid, ParA, and Nucleoid by a Triple Labeling
System. We engineered a triple color labeling system to simulta-
neously analyze the subcellular dynamics of plasmids, ParA, and the
bacterial nucleoid in E. coli cells. A fully functional ParA-GFP
fusion was expressed at a level close to that of ParA expressed by
par2 (1). For brevity, ParA will be used interchangeably with
ParA-GFP in the following. Plasmids were visualized by a TetRmCherry fusion protein that binds to an array of 120 plasmidencoded tetO operators (16), and the nucleoid was stained with
Hoechst. When TetR-mCherry was donated in trans, the plasmid of
interest was visible as bright foci (Fig. 1). Plasmid stability assays
confirmed that the cytological data acquired using tetO/TetRmCherry to label plasmids were all obtained under conditions in
which par2 was functional (supporting information (SI) Fig. S1C),
consistent with the regular plasmid distributions shown in Fig. S1 A
and B.
ParA Recruits par2-Carrying Plasmids to the Nucleoid. Using our new
plasmid labeling system, we analyzed plasmid localization in cells
treated with nalidixic acid. This antibiotic, which inhibits DNA
gyrase, leads to the formation of filamentous cells with nonsegregated nucleoids. Foci of the R1 control plasmid without the par2
system localized almost exclusively to cytosolic regions (Fig. 1 A). By
striking contrast, foci of the par2-carrying plasmid colocalized with
the nucleoid (Fig. 1B). Moreover, a plasmid carrying an in-frame
deletion in parA had a localization pattern indistinguishable from
Author contributions: S.R., J.v.Z., M.H., and K.G. designed research; S.R., J.v.Z., and M.H.
performed research; S.R., J.v.Z., M.H., and K.G. analyzed data; and S.R., J.v.Z., M.H., and K.G.
wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Freely available online through the PNAS open access option.
1To
whom correspondence should be addressed. E-mail: kenn.gerdes@newcastle.ac.uk.
This article contains supporting information online at www.pnas.org/cgi/content/full/
0908347106/DCSupplemental.
PNAS 兩 November 17, 2009 兩 vol. 106 兩 no. 46 兩 19369 –19374
CELL BIOLOGY
Bacterial plasmids encode partitioning (par) loci that confer stable
plasmid inheritance. We showed previously that, in the presence of
ParB and parC encoded by the par2 locus of plasmid pB171, ParA
formed cytoskeletal-like structures that dynamically relocated over
the nucleoid. Simultaneously, the par2 locus distributed plasmids
regularly over the nucleoid. We show here that the dynamic ParA
patterns are not simple oscillations. Rather, ParA nucleates and
polymerizes in between plasmids. When a ParA assembly reaches a
plasmid, the assembly reaction reverses into disassembly. Strikingly,
plasmids consistently migrate behind disassembling ParA cytoskeletal
structures, suggesting that ParA filaments pull plasmids by depolymerization. The perpetual cycles of ParA assembly and disassembly
result in continuous relocation of plasmids, which, on time averaging,
results in equidistribution of the plasmids. Mathematical modeling of
ParA and plasmid dynamics support these interpretations. Mutational
analysis supports a molecular mechanism in which the ParB/parC
complex controls ParA filament depolymerization.
A
findings suggest that ParA recruits par2-carrying plasmids to the
nucleoid, consistent with previous observations (2).
B
Plasmid Movement and ParA Dynamics Are Intimately Connected.
d
5 µm
de
co
nv
Pa
ol
ve
Te rAd
tR GF
-m P
C h de
er co
Pa
r y nv
ol
Te rAve
tR GF
d/
-m P
C h de
er co
ry nv
/N o l
uc ve
Ky
le d/
m
oi
”p og
d
ea ra
k” ph
Pa o
rA f p
-G l a s
FP m
dy id /
na
m
ics
c
rA
-G
FP
b
Pa
a
Pa
Te rAtR GF
-m P
Ch /
er
ry
C
Ph
a
Te sec
t R on
-m o
Ch ntr
er ast
ry /
Pa
rA
-G
FP
5 µm
e
f
g
0’
0’
1’
1’
2’
2’
3’
3’
4’
4’
5’
5’
6’
6’
7’
7’
8’
8’
9’
9’
10’
10’
11’
11’
12’
12’
13’
13’
14’
14’
15’
15’
16’
16’
17’
17’
18’
18’
19’
19’
20’
20’
21’
21’
22’
22’
23’
23’
24’
24’
25’
25’
26’
26’
27’
27’
28’
28’
29’
29’
D
15’
15’
75o
29’
29’
Fig. 1. par2-carrying plasmids trail retracting ParA on the nucleoid. (A and B)
Intracellular localization of nucleoid (Hoechst stain, green) and plasmids (red) in
strain SR1 harboring (A) par⫺ plasmid pSR230 and (B) par2⫹ plasmid pSR233. (C)
Time lapse showing the subcellular localization of ParA-GFP and par2⫹ plasmid
pSR233 (R1 par2⫹ plac::parA::gfp tetO120) in strain SR1 (KG22⌬pcnB) also carrying the pBR322-based pSR124 plasmid (pBAD::tetR::mCherry). Numbers on the
side indicate minutes in the time lapse. (Ca) Overlay of phase-contrast images and
TetR-mCherry (red). (Cb) Intracellular localization of ParA-GFP. (Ac) Overlay of
ParA-GFP (green) and TetR-mCherry (red). (Cd) 2D deconvolved images of ParAGFP. (Ce) Overlay of deconvolved ParA-GFP (green) and TetR-mCherry (red). (Cf )
Overlay of deconvolved ParA-GFP (green), TetR-mCherry (red), and nucleoid
(blue). (Cg) Kymograph of plasmid (red) and maximum intensity of ParA-GFP
signal in (Cb). A movie of the time lapse is presented in Movie S1. (D) 3D surface
intensity plot of the dashed region of minutes 15–29 in (Cc).
that of the par⫺ plasmid, showing that ParA is required for plasmid
recruitment to the nucleoid. We also investigated if purified ParA
would bind DNA nonspecifically. Gel-shift analysis showed that
ParA indeed has nonspecific DNA binding activity in vitro (Fig. S2).
Thus, all our observations are consistent with the proposal that the
nucleoid functions as a scaffold for ParA assembly. Together, these
19370 兩 www.pnas.org兾cgi兾doi兾10.1073兾pnas.0908347106
Next, we analyzed plasmid and ParA dynamics in a cell with one
plasmid focus and one nucleoid (Fig. 1C and Movie S1). Plasmid
localization relative to the entire cell was obtained by phase contrast
microscopy (Fig. 1Ca), and relative to the nucleoid by labeling with
Hoechst (Fig. 1Cf ). We observed that the plasmid focus and ParA
oscillated back and forth over the nucleoid (Fig. 1C a, b, and f ).
Initially, the plasmid was located at midcell, with an assembly of
ParA to the right (Fig. 1Cc, 0⬘). The plasmid then moved rightward
while the ParA-GFP signal close to the focus disassembled. The
region behind the plasmid was left almost devoid of ParA-GFP
signal (Fig. 1Cc, 0⬘–8⬘). Simultaneously, ParA assembled in the left
half of the nucleoid. Eventually, the left assembly of ParA reached
the plasmid that was now located close to the right nucleoid pole
(Fig. 1Cc, 8⬘). At this stage (Fig. 1Cc, blue arrowheads), a new
dynamic event was triggered—the newly generated assembly of
ParA retracted leftward and again the plasmid focus followed the
retracting ParA-GFP signal. During this retraction, the region
behind the plasmid was again left devoid of ParA-GFP signal (Fig.
1Cc, 8⬘–17⬘). Once more, a ParA-GFP assembly was initiated on the
other (right) half, which eventually reached the plasmid focus (Fig.
1Cc, 18⬘–28⬘). The combined trajectories of plasmid and ParA
movements show that the plasmid consistently moved toward
regions of high ParA-GFP signal (Fig. 1Cg). Two-dimensional
deconvolution of the cytological recordings resolved the ParA-GFP
signal into filamentous structures colocalizing with the nucleoid
(Fig. 1C d–f ). It is obvious from these images that the filamentous
structures reversed from growth to shrinkage when they reached
the plasmid focus (Fig. 1C, yellow arrowheads), and that the focus
then started to move in the direction of the retracting ParA
structures, staying close to the filament end.
ParA assembly consistently initiated away from plasmid foci and
gradually continued to extend toward the nucleoid poles (Fig. 1Cf,
Figs. S3 and S4, and Movie S2). When the growing filaments
reached a focus, they reversed into disassembly. This pattern was
general for cells with one focus (see further examples in Figs. S3 and
S4). The oscillation frequency and amplitude of ParA-GFP and
plasmid foci varied considerably from one cell to another and also
in one cell over time: the plasmid foci shown in Fig. 1C and Figs.
S3 and S4 moved over most of the nucleoid. In other cases, the foci
tended to oscillate around midnucleoid whilst trailing retracting
ParA (Fig. S5). Importantly, the plasmid foci consistently followed
retracting ParA filaments. These observations unequivocally show
that ParA dynamics and plasmid segregation inherently go hand in
hand.
A focus often detached before the ParA assembly completely
disassembled. After detachment, ParA continued to disassemble,
leaving the focus behind until reached by a new ParA assembly (Fig.
1D and Fig. S5, 29⬘–34⬘). Fig. 1D is a 3D surface intensity plot of
the dashed region in Fig. 1Cc (15⬘–29⬘) showing that the focus
detached at 16⬘ but ParA-GFP continued to disassemble, leaving
the focus behind where it remained until at 27⬘ when it was moved
by a new ParA assembly. These data suggest that once ParA
disassembly has been initiated by ParB/parC, a continuous interaction with the ParB-bound plasmid is seemingly not required for
ParA filament depolymerization.
ParA Moves Plasmids by a Pulling Mechanism. When a ParA filament
reached a plasmid focus, it consistently reversed from growth
(interpreted as polymerization) to retraction (interpreted as depolymerization) (Fig. 1C). We find it reasonable to interpret retraction
of ParA filaments as filament depolymerization for 3 main reasons:
(i) ParA forms filaments in vitro (7), consistent with the filamentous
structures we see in vivo; (ii) the observation that ParA filaments
shrink from the focus/filament boundary leaving the space behind
Ringgaard et al.
a
Time
time (min)
time (min)
Ha
C0
time (min)
B0
A0
150
25
50
75 100
% of nucleoid length
E
time (min)
F
0
25
50
75 100
% of nucleoid length
0.4
frequency
25
50
75 100
% of nucleoid length
G 0.4
0.1
0.2
0
0
0
0.2
0
150
25
50
75 100
% of nucleoid length
0.3
frequency
D0
150
0
frequency
0
0
25
50
75 100
% of nucleoid length
25
50
75 100
% of nucleoid length
3
2
1
0
Hc
0
R2=0.84
0
0.2
0
25
50
75 100
% of nucleoid length
Ratio between focus
distance travelled and
initial filament length, Δc/a
150
Focus distance
travelled (μm), Δc
Hb
ParB
on plasmid
ParA
Δc = a- b
b
1.0
1
2
3
R2=0.51
0.8
0.6
0.4
0.2
0.0
0
1
2
3
Initial filament length (μm), a
the focus devoid of ParA-GFP signal; and (iii) the observation that
ParA in elongated cells accumulates over one nucleoid simultaneously with its disappearance from another when pulling a plasmid
(Fig. S6 and Movie S3). The consistent migration of plasmid foci at
the boundaries of retracting ParA filaments suggests that ParA
moves plasmids by a pulling mechanism in which depolymerization
of ParA generates the mechanical force for plasmid migration. The
assertion that ParA generates force on the plasmid foci was further
strengthened by the fact that ParA could transfer plasmids between
separate nucleoids, not only in cephalexin-treated cells (Fig. S6) but
also in dividing cells.
Single Amino Acid Change in the ParB N Terminus Abolishes ParA
Dynamics. The components encoded by type Ib parFGH locus of
TP228 has been extensively analyzed biochemically. Importantly,
the N terminus of ParG (ParB homolog) stimulates the ATPase
activity of ParF (ParA homolog) (11). Arg-19 in the N terminus of
ParG is essential for this stimulation. Alignments of ParG and ParB
sequences showed that Arg-19 of ParG corresponds to Arg-26 of
ParB (Fig. S7). We mutated 2 arginines in the N terminus of ParB
(Arg-12 and Arg-26) to either lysine or alanine (Fig. S7A). Plasmid
segregation assays showed that par2 carrying parBR26K exhibited a
3-fold reduced activity, whereas par2 parBR26A exhibited an almost
complete par-deficient phenotype (Fig. S7B). By contrast, mutational changes of Arg-12 had no measurable effect on plasmid
segregation by par2.
We then performed time-lapse microscopy of ParA in the
context of parBR26A (Fig. S7C). In this case, ParA still localized to
the nucleoid. However, ParA dynamics occurred in 15% of the cells
Ringgaard et al.
only, in contrasted to the WT case in which virtually all cells showed
ParA movement. Thus, the N terminus (Arg-26) of ParB is essential
for both ParA dynamics and par2 activity. Because the corresponding arginine (Arg-19) of ParG is responsible for stimulation of the
ATPase activity of ParF in vitro, and the fact that Arg-26 of ParB
is essential for ParA oscillation and plasmid stability, our observations suggest that ParB similarly stimulates the ATPase activity of
ParA and that this stimulation is a prerequisite for ParA dynamics
by means of regulating ParA depolymerization (see text following).
To further substantiate that the par loci of plasmids pB171 and
TP228 function by similar mechanisms, we tagged ParF with a
fluorescent protein and analyzed the subcellular localization and
dynamics of the fusion protein. As for ParA, ParF also localized to
the nucleoid (Fig. S7D). Most importantly, time-lapse experiments
showed that, in the presence of the entire par system (including
ParG and parH centromere), ParF oscillated over the nucleoid (Fig.
S7E). In the absence of ParG/parH, no oscillation of ParF was
observed (Fig. S7F). These observations further indicate that the 2
par loci function by similar, if not identical, molecular mechanisms.
Mathematical Model Describing the par Mechanism. Although it is
intuitively clear that a pulling-like mechanism will generate singlecell kymographs similar to those seen in Fig. 1C, it is far from
obvious that such a mechanism can generate the regular focus
distributions obtained from many cells (Fig. S1 A and B). We
therefore constructed a simple, stochastic computational model
that could predict plasmid focus distributions and ParA dynamics.
The model consisted of randomly nucleated ParA filaments that
grew stochastically until they reached a plasmid, after which they
PNAS 兩 November 17, 2009 兩 vol. 106 兩 no. 46 兩 19371
CELL BIOLOGY
Fig. 2. Simulated kymographs and plasmid foci distributions generated by mathematical modeling. (A–C) Simulated kymographs of ParA/focus dynamics for (A) 1,
(B) 2, and (C) 3 foci cases, a movie of (C) is presented in Movie S7. (D) Simulated kymograph of one focus splitting into 2 and being segregated. (E-G) Simulated foci
distributions for (E) 1, (F) 2, and (G) 3 foci cases. For all simulated foci distributions, the distributions were built up over 7,500 min of simulated time, with sampling every
7.5 min, and means and error bars constructed from 40 independent runs. (H) Plasmid travel distance is ParA filament length dependent. (Ha) Schematics showing how
the focus travel distance was measured relative to initial ParA filament length. (Hb) The plot shows the focus travel distance, ⌬c, as function of initial ParA filament
length, a. (Hc) The plot shows the ratio between focus travel distance and initial ParA filament length, ⌬c/a, as function of initial ParA filament length, a.
3D
su
in t rfac
en
e
Pa sity
pl o
rA
to
Te -GF
f
P/
tR
-m
Ch
er
ry
cle
e
3
Ky
m
p l a og r
a
”p smi ph o
ea d /
f
av k” P dyn
e
a
po rage arA- mic
s
G
sit
ion focu FP /
s
0’
Nu
d
12
oid
ry
c
Pa
rA
Te -GF
tR
-m P/
Ch
er
b
Pa
rA
-G
FP
a
Ph
as
Te eco
tR
- m nont
Ch ras
er
ry t/
A
f
0’
1’
1’
2’
2’
3’
3’
4’
4’
5’
5’
6’
6’
7’
7’
8’
8’
9’
9’
10’
10’
11’
11’
12’
12’
13’
13’
14’
14’
15’
15’
16’
16’
17’
17’
18’
18’
19’
19’
20’
20’
21’
21’
22’
22’
23’
23’
24’
24’
25’
25’
26’
26’
27’
27’
28’
28’
29’
29’
30’
30’
31’
Kymograph of plasmid/
“peak” ParA-GFP dynamics
C
1’
a
a
b
c
2’
3’
4’
5’
6’
1 2
7’
1
8’
9’
1
10’
1
12’
1
13’
1
14’
1
15’
1
16’
1
17’
14
2
2
1
1
25’
4
2
3
2
3
1 4
22’
2
3
1 4
1
2
3
19’
21’
2
3
1
24’
2
2
18’
23’
2
2
1
11’
20’
2
2
3
3
2
4
3
2
4
3
4
1
4
1
4
2
3
2
3
2
d
0’
1’
2’
3’
4’
5’
6’
7’
8’
9’
10’
11’
12’
13’
14’
15’
16’
17’
18’
19’
20’
21’
22’
23’
24’
25’
26’
27’
28’
29’
b
c
nv
olv
Pa
ed
r
Te A-G
tR
FP
-m
C h dec
o
er
ry n v o
lve
Pa
rA
d /
-G
Te
F
tR
-m P d e
Ch
co
nv
er
ry
olv
/N
ed
uc
/
Ky
le o
mo
id
gr
”p
a
ea
ph
k”
P a of pl
as
rA
mi
-G
d /
FP
ParA-GFP/
TetR-mCherry
ParA-GFP
0’
Pa
r
Te A-G
tR
F
-m P/
Ch
er
ry
Pa
rA
-G
FP
de
co
Phasecontrast/
TetR-mCherry
Ph
as
Te eco
tR
n
-m ont
Ch ras
t
er
ry /
Pa
rA
-G
FP
B
31’
d
e
f
g
0’
1’
2’
3’
4’
5’
6’
7’
8’
9’
10’
11’
12’
13’
14’
15’
16’
17’
18’
19’
20’
21’
22’
23’
24’
25’
26’
27’
28’
29’
Fig. 3. Perpetual cycles of ParA assembly/disassembly move and position plasmids. (A–C) Time lapses showing the subcellular localization of ParA-GFP and par2⫹
plasmids (A and B) in a cephalexin-treated cell. Numbers on the side indicate minutes in the time lapse. Experimental setup as in Fig. 1C. Videos of the time lapse in (A–C)
are presented in Movies S4, S5, and S6, respectively. (Aa, Ba, and Cc) Overlay of phase-contrast images and TetR-mCherry (red). (Ab, Bb, and Cc) Intracellular localization
of ParA-GFP. (Ac, Bc, and Cc) Overlay of ParA-GFP (green) and TetR-mCherry (red). (Ad) Nucleoid stained with Hoechst. (Ae) 3D surface intensity plot of (Ac). (Af )
Kymograph of plasmid (red) and maximum intensity of ParA-GFP signal (green) in (Ac). Dotted blue lines indicate the average foci positions during the time lapse. (Bd
and Cg) Kymograph of plasmid (red) and highest-intensity ParA-GFP signal in (Bb) and (Cb). (C) Focus splitting and segregation by ParA filaments. (Cd) Deconvolved
images of ParA-GFP. (Ce) Overlay of deconvolved ParA-GFP (green) and TetR-mCherry (red). (Cf ) Overlay of deconvolved ParA-GFP (green) and TetR-mCherry (red) and
nucleoid (blue).
19372 兩 www.pnas.org兾cgi兾doi兾10.1073兾pnas.0908347106
Ringgaard et al.
ADP
1
ATP
ATP ATP ATP
AD
ATP
P
5
2
ATP
ATP
P
AD
ATP
ATP
ATP
ATP
ATP ATP ATP
3
4
4’
ATP
ATP
P
AD
P
AD
ATP ATP ATP
ADP
ParA dimer
ParB
ParB bound on
plasmid parC site
Chromosomal DNA
Direction of
polymerization
Direction of
depolymerization
Fig. 4.
Molecular model showing how plasmid movement is generated by dynamic ParA
filaments. See Discussion for a detailed description of the molecular model.
switched to depolymerization, during which the plasmid was
‘‘dragged’’ along with the shrinking filament before being released.
Full details of the model are given in SI Text.
In our initial modeling, a plasmid focus detached from a ParA
filament only after the filament had completely disassembled. This
assumption predicted a roughly flat plasmid focus distribution
across the nucleoid with peaks toward the poles (Fig. S8). When
compared with our experimental data (Fig. S1 A and B, second
panels), this distribution clearly had qualitatively the wrong form,
a problem that persisted even with a constant nonzero probability
of detachment per depolymerization step (see SI Text). To correct
this problem, we implemented a revised rule that a focus would
detach from a filament during depolymerization with a probability
that depended on the current filament length, with longer filaments
having a reduced probability of focus detachment. Simulations
generated by this revised model produced kymograph and plasmid
distributions that agreed well with our observations in the single
focus case (cf. Fig. 2A with Fig. 1C, and Fig. 2E with Fig. S1B). In
Fig. 2D we show simulated kymographs for the case where one
focus separates into 2 foci, which are then rapidly pulled apart and
subsequently move over separate nucleoid halves.
Testing the Model: Plasmid Detachment Rates. In cells with one focus,
we measured experimentally the absolute distance traveled by the
focus relative to the initial length of the ParA filament pulling the
focus (Fig. 2H). The focus typically detached before the filament
completely depolymerized (Fig. 1D), regardless of filament length.
From the measurements it is evident that long ParA filaments pull
the plasmid a longer distance than shorter filaments (Fig. 2Hb).
Furthermore, a long filament moved a plasmid longer relative to its
initial length compared to short filaments (Fig. 2Hc). Hence, the
longer distances traversed by a focus attached to longer filaments
was not only due to the filament itself being longer, but is also a
consequence of the focus detachment rate being lower for longer
filaments. Therefore, the data in Fig. 2H supported the revised rule
of the mathematical model.
It is not known why longer filaments have lower detachment rates
than shorter filaments. One possibility is that ParB/parC-carrying
plasmids contact bundles of ParA protofilaments rather than a
single filament, and that long bundles consist of a larger number of
protofilaments than short ones. The parC1 and parC2 regions of
par2 consist of 17 and 18 ParB dimer binding sites, respectively, and
deletion of parC2 reduced the efficiency of par2 (1, 17). These
observations are consistent with the proposal that multiple ParA
protofilaments can simultaneously contact the partition complex of
a par2-carrying plasmid, and that filament bundling reduces the
detachment rate.
Ringgaard et al.
foci cases (Fig. 2 B and C). Here, the model predicted much more
complex ParA dynamics than in the single focus case, with cycles of
ParA polymerization/depolymerization in between plasmids moving relative to each other (Fig. 2 B and C), which on time averaging
resulted in equipositioning of plasmids (Fig. 2 F and G) similar to
our experimental data (7) (Fig. S1 A and B).
Analysis of ParA and plasmid dynamics was difficult in cells with
multiple foci. Thus, to analyze the multiple foci case, we treated cells
with cephalexin to obtain elongated, nondividing cells (Fig. 3A and
Movie S4). Strikingly, as predicted by the mathematical modeling,
ParA dynamics was much more complex in the multifoci cases (Fig.
3Ab) than the simple pole-to-pole oscillation seen in cells with a
single focus. Instead, ParA exhibited continuous rounds of assembly/disassembly in between plasmid foci (Fig. 3A c and e). Similar
to cells with one focus, when assembling ParA structures reached
a focus, assembly was reversed to disassembly and the focus
followed behind the retracting ParA-GFP signal (Fig. 3A a–d, focus
3, 0⬘–10⬘). Concomitantly, a new ParA assembly was initiated
elsewhere and the cycle was repeated. Often, a ParA assembly
contacted foci at both ends. This was consistently accompanied by
bipolar disassembly of ParA and movement of the 2 foci toward
each other (Fig. 3A, 14⬘–19⬘, and Fig. S6A). The fact that disassembling ParA can move 2 plasmids toward each other is in further
support of a system where ParA moves plasmids by a pulling
mechanism. This perpetual cycle of ParA assembly/disassembly
between foci moved and positioned plasmids relative to each other,
resulting in a time-averaged equidistribution of the foci in single
cells (Fig. 3A e and f ) and at the level of the cell population (Fig.
S1 A and B). Fig. 3Af shows kymographs of plasmid foci and peak
ParA-GFP dynamics. Dotted blue lines indicate the average foci
positions during the time lapse. It is evident that the foci continuously move around these average position. As a further characteristic example, we include a case in which one plasmid focus was
separated by cycles of ParA-GFP assembly/disassembly into 4 foci
that eventually became distributed throughout the cell (Fig. 3B and
Movie S5).
We also analyzed a focus splitting event. When separation of
plasmids occurred, in some instances both plasmids were pulled in
the same direction (Fig. 3C, 12⬘–19, and Movie S6). In other
examples, after splitting, only one focus moved with ParA, and the
other focus stayed behind, resulting in focus separation (Fig. 3C,
26⬘–29, and Figs. S3 and S4). ParA filaments then contacted the
plasmid which had been left behind (Fig. 3C, blue arrowheads, and
Figs. S3 and S4), seemingly triggering movement of the plasmid
toward the other half of the cell, thus resulting in the 2 plasmids
being localized away from each other with the region in between
devoid of ParA-GFP signal (Fig. 3C, green arrowheads). Again we
see good general agreement between our mathematical simulations
and experiments (cf. Figs. 2D and 3C, and Figs. S3 and S4).
Molecular Model Explaining Plasmid Movement and ParA Relocation.
The cytological observations presented here, together with previous biochemical data obtained with other type I par loci components (6, 9, 11, 15, 17–19) and our mathematical modeling, all
support a simple picture of how the perpetual cycles of ParA
filament growth/shrinkage may generate the force that moves and
positions plasmids over the nucleoid (Fig. 4). In step 1, ParA2-ATP
dimers bind cooperatively to nucleoid DNA, leading to large ParA
filaments. This contention is supported by in vitro data obtained
with 2 type I ParAs (Soj and ParA of pSM19035) (6, 14). Formation
of filaments begins with a nucleating core from which rapid
polymerization proceeds. In step 2, a growing filament contacts a
plasmid via ParB bound to parC. In step 3, ParBs bound to parC on
the plasmid stimulate the ATPase activity of ParA2-ATP at the end
of the filament. This step is supported by the fact that ParG of
TP228 stimulates the ATPase activity of ParF, that ParF of TP228
PNAS 兩 November 17, 2009 兩 vol. 106 兩 no. 46 兩 19373
CELL BIOLOGY
Equidistribution of Plasmids by Perpetual Cycles of ParA Assembly/
Disassembly. Next we generated simulated kymographs for multi-
ATP
6
P
AD
oscillates over the nucleoid (Fig. S7 D and E), and our finding that
the N terminus of ParB was required for ParA dynamics (Fig. S7
A–C). By this reaction, ParA2-ATP is converted to its ADP form
and released from the DNA (6, 14), leaving a new ParA2-ATP
filament end accessible for interaction with the partition complex.
For each depolymerization event, the plasmid can either drop off
(with a probability dependent on the filament length, step 4⬘; Fig.
2H) or continue (step 4) to be attached to the end of the
depolymerizing ParA filament. In steps 4 and 4⬘, the ParA filament
continues to depolymerize until it completely disappears (Fig. 1 C
and D). In step 5, the plasmid has been released and ParA2-ATP
subunits have assembled into a new filamentous structure away
from the plasmid. Eventually, the released plasmid interacts with a
new ParA filament approaching from the opposite side. When
contact is made, this filament will move the plasmid in the opposite
direction. Finally, in step 6, free ParA2-ADP is rejuvenated to
ParA2-ATP (6). This perpetual cycle of ParA relocation converts
energy in the form of ATP to mechanical force that powers plasmid
movement.
Comparison with Related Systems. The type I par locus of plasmid
F (sopABC) has also been investigated with respect to SopA and
plasmid dynamics in living cells (3). The observed patterns of
plasmid movement and localization were similar to those described here. However, SopA-GFP dynamics differed from that
of ParA-GFP. First, SopA-GFP formed a big focus that oscillated from one end of the nucleoid to the other. While oscillating, the F plasmid focus followed the SopA focus, but there was
no apparent contact between the SopA and plasmid foci. Second, SopA-GFP also formed a static filamentous structure that
spanned the entire length of the nucleoid without being shortened as the plasmid migrated. Based on these observations,
Hatano et al. (3) suggested that the SopA focus redistributes
within the stationary filament while maintaining an overall
filament structure. Importantly, it was not explained how the sop
system positioned plasmids relative to each other. Furthermore,
the nucleoid did not appear to be essential for SopA oscillation.
In contrast, our data clearly show that the nucleoid plays an
important role for par2-mediated plasmid partitioning.
Most bacterial chromosomes encode type I par loci (20) that have
1. Ebersbach G, Gerdes K (2001) The double par locus of virulence factor pB171: DNA
segregation is correlated with oscillation of ParA. Proc Natl Acad Sci USA 98:15078 –15083.
2. Ebersbach G, Gerdes K (2004) Bacterial mitosis: Partitioning protein ParA oscillates in
spiral-shaped structures and positions plasmids at mid-cell. Mol Microbiol 52:385–398.
3. Hatano T, Yamaichi Y, Niki H (2007) Oscillating focus of SopA associated with filamentous structure guides partitioning of F plasmid. Mol Microbiol 64:1198 –1213.
4. Lim GE, Derman AI, Pogliano J (2005) Bacterial DNA segregation by dynamic SopA
polymers. Proc Natl Acad Sci USA 102:17658 –17663.
5. Adachi S, Hori K, Hiraga S (2006) Subcellular positioning of F plasmid mediated by
dynamic localization of SopA and SopB. J Mol Biol 356:850 – 863.
6. Pratto F, et al. (2008) Streptococcus pyogenes pSM19035 requires dynamic assembly of
ATP-bound ParA and ParB on parS DNA during plasmid segregation. Nucleic Acids Res
36:3676 –3689.
7. Ebersbach G, et al. (2006) Regular cellular distribution of plasmids by oscillating and
filament-forming ParA ATPase of plasmid pB171. Mol Microbiol 61:1428 –1442.
8. Hester CM, Lutkenhaus J (2007) Soj (ParA) DNA binding is mediated by conserved
arginines and is essential for plasmid segregation. Proc Natl Acad Sci USA 104:20326 –
20331.
9. Bouet JY, Ah-Seng Y, Benmeradi N, Lane D (2007) Polymerization of SopA partition
ATPase: Regulation by DNA binding and SopB. Mol Microbiol 63:468 – 481.
10. Barilla D, Rosenberg MF, Nobbmann U, Hayes F (2005) Bacterial DNA segregation
dynamics mediated by the polymerizing protein ParF. EMBO J 24:1453–1464.
11. Barilla D, Carmelo E, Hayes F (2007) The tail of the ParG DNA segregation protein
remodels ParF polymers and enhances ATP hydrolysis via an arginine finger-like motif.
Proc Natl Acad Sci USA 104:1811–1816.
12. Batt SM, Bingle LE, Dafforn TR, Thomas CM (2009) Bacterial genome partitioning:
N-terminal domain of IncC protein encoded by broad-host-range plasmid RK2 modulates oligomerisation and DNA binding. J Mol Biol 385:1361–1374.
13. Dunham TD, Xu W, Funnell BE, Schumacher MA (2009) Structural basis for ADPmediated transcriptional regulation by P1 and P7 ParA. EMBO J 28:1792–1802.
14. Leonard TA, Butler PJ, Lowe J (2005) Bacterial chromosome segregation: Structure and
DNA binding of the Soj dimer—a conserved biological switch. EMBO J 24:270 –282.
19374 兩 www.pnas.org兾cgi兾doi兾10.1073兾pnas.0908347106
been proposed to be involved in chromosome segregation (21–24).
More direct evidence for this notion has come from Caulobacter
crescentus (23) and Vibrio cholerae (24). In particular, ParAI
encoded by ChrI of V. cholerae exhibited a disassembly pattern
suggesting that it generates force and segregates ChrI origins by a
pulling mechanism (24).
The minCDE locus of E. coli encodes oscillating and filamentforming MinD that, in combination with MinC and MinE, prevents
FtsZ-ring formation at the cell poles (25). The similarities between
the E. coli minCDE system and type I par loci are striking. ParA and
MinD are both ‘‘deviant’’ Walker A Box ATPases that form
dynamic patterns on a cellular surface (nucleoid and cell membrane, respectively) (1, 25). Both ParA and MinD form cytoskeletal
filaments that interact with their surfaces as ATP-bound dimers (6,
14, 26). The nucleotide state determines the cellular location of
ParA and MinD, where ATP hydrolysis releases the proteins from
their surface-bound states (6, 14, 27). The ATPase activities of the
proteins are stimulated by the N termini of their dimeric partner
proteins (ParB and MinE, respectively) (6, 11, 14, 15, 27). Furthermore, filament dynamics functions to position another cellular
structure (FtsZ ring or plasmid, respectively) (ref. 25; this work).
These observations reveal that evolution has solved 2 very different
spatial problems related to cell division (DNA segregation and
septum placement) by related molecular mechanisms.
Materials and Methods
A functional ParA-GFP fusion was constructed as described previously (1).
Plasmids were visualized by binding of TetR-mCherry fusion protein to an
array of 120 plasmid-encoded tetO operators on the plasmid of interest (16).
Microscopy was performed essentially as described previously (2, 16) with a
few changes described in SI Text. Microscopy was performed in a ⌬pcnB strain
[lacking poly(A) polymerase]. Mathematical modeling, additional materials
and methods, tables, and movie legends are presented as SI Text. Strains and
plasmids are listed in Table S1.
ACKNOWLEDGMENTS. We thank Kathrin Schirner, Jeff Errington, and Heath
Murray for comments on the manuscript; François X. Barre (Gif-sur-Yvette Cedex,
France) for plasmid pFX240; and Gitte Ebersbach (Odense, Denmark), Qing Wang
(Cambridge, United Kingdom), and Simon Syvertssen (Newcastle, United Kingdom) for the construction plasmids. This work was supported by grants from the
U.K. Biotechnology and Biological Sciences Research Council (to M.H. and K.G.),
the Danish Natural Science Research Council (FNU) (to K.G.), and from the Royal
Society (to M.H.).
15. Radnedge L, Youngren B, Davis M, Austin S (1998) Probing the structure of complex
macromolecular interactions by homolog specificity scanning: The P1 and P7 plasmid
partition systems. EMBO J 17:6076 – 6085.
16. Lau IF, et al. (2003) Spatial and temporal organization of replicating Escherichia coli
chromosomes. Mol Microbiol 49:731–743.
17. Ringgaard S, Lowe J, Gerdes K (2007) Centromere pairing by a plasmid-encoded type
I ParB protein. J Biol Chem 282:28216 –28225.
18. Leonard TA, Butler PJ, Lowe J (2004) Structural analysis of the chromosome segregation
protein Spo0J from Thermus thermophilus. Mol Microbiol 53:419 – 432.
19. Ringgaard S, Ebersbach G, Borch J, Gerdes K (2007) Regulatory cross-talk in the double
par locus of plasmid pB171. J Biol Chem 282:3134 –3145.
20. Gerdes K, Moller-Jensen J, Bugge JR (2000) Plasmid and chromosome partitioning:
Surprises from phylogeny. Mol Microbiol 37:455– 466.
21. Ireton K, Gunther NW, Grossman AD (1994) spo0J is required for normal chromosome
segregation as well as the initiation of sporulation in Bacillus subtilis. J Bacteriol
176:5320 –5329.
22. Jakimowicz D, et al. (2007) Alignment of multiple chromosomes along helical ParA
scaffolding in sporulating Streptomyces hyphae. Mol Microbiol 65:625– 641.
23. Toro E, Hong SH, McAdams HH, Shapiro L (2008) Caulobacter requires a dedicated
mechanism to initiate chromosome segregation. Proc Natl Acad Sci USA 105:15435–15440.
24. Fogel MA, Waldor MK (2006) A dynamic, mitotic-like mechanism for bacterial chromosome segregation. Genes Dev 20:3269 –3282.
25. Raskin DM, de Boer PA (1999) Rapid pole-to-pole oscillation of a protein required for
directing division to the middle of Escherichia coli. Proc Natl Acad Sci USA 96:4971–
4976.
26. Hu Z, Saez C, Lutkenhaus J (2003) Recruitment of MinC, an inhibitor of Z-ring formation,
to the membrane in Escherichia coli: Role of MinD and MinE. J Bacteriol 185:196 –203.
27. Ma L, King GF, Rothfield L (2004) Positioning of the MinE binding site on the MinD
surface suggests a plausible mechanism for activation of the Escherichia coli MinD
ATPase during division site selection. Mol Microbiol 54:99 –108.
Ringgaard et al.