-
BACTERIAL DIVISION
Mechanical crack propagation drivesmillisecond daughter cell
separationin Staphylococcus aureusXiaoxue Zhou,1,2,3* David K.
Halladin,2,3,4* Enrique R. Rojas,2,3,5* Elena F. Koslover,2,3
Timothy K. Lee,5 Kerwyn Casey Huang,4,5 Julie A.
Theriot2,3,4†
When Staphylococcus aureus undergoes cytokinesis, it builds a
septum, generating twohemispherical daughters whose cell walls are
only connected via a narrow peripheral ring.We found that
resolution of this ring occurred within milliseconds (“popping”),
withoutdetectable changes in cell volume. The likelihood of popping
depended on cell-wall stress,and the separating cells split open
asymmetrically, leaving the daughters connected bya hinge. An
elastostatic model of the wall indicated high circumferential
stress in theperipheral ring before popping. Last, we observed
small perforations in the peripheralring that are likely initial
points of mechanical failure. Thus, the ultrafast daughter
cellseparation in S. aureus appears to be driven by accumulation of
stress in the peripheralring and exhibits hallmarks of mechanical
crack propagation.
Most bacteria propagate through binaryfission, a process that is
highly coordi-nated and tightly controlled to pass ongenetic
material equally to the twodaughter cells and to regulate cell
size
and shape. Much of our knowledge of bacterialcell division comes
from rod-shaped bacteria,which double their cell length before
cytokinesis(1, 2); relatively less is known about cell divisionin
bacteria with other shapes. Staphylococcusaureus, a model system
for round bacteria, is aGram-positive pathogen well recognized for
itsvirulence and antibiotic resistance (3, 4). To di-vide, S.
aureus builds a septum, generating twohemispherical daughter cells
(5, 6). After construc-tion, the septal wall exists as two flat,
parallelplates, and the walls of the two daughter cellsare
connected only through a narrow periph-eral ring (Fig. 1A) (7).
Presumably, resolution ofthis peripheral wall ring leads to
daughter cellseparation, which is accompanied by a shapeconversion
of the daughter cells from hemispheresto spheres. This shape change
has previously beenassumed to occur through expansion of the
sep-tum to twice its original surface area, which woulddouble the
cell volume (5, 8). It remains unclearhow exactly the peripheral
ring is resolved toallow the daughter cells to separate,
particularlygiven that the S. aureus cell wall is quite thick (20to
30 nm) (9).Previous video microscopy–based observations
of S. aureus cell division have described daughter
cell separation as a dramatic “popping” eventwith no detectable
intermediate stages (10, 11). Toaddress the time scale and
mechanism of thepopping, we used phase contrastmicroscopywitha
temporal resolution of 1 ms. At this frame rate,we occasionally
observed intermediate stages ofpopping, whereas most separations
occurredwithin one or two frames (20% (fig. S1and movie S2),
indicating that the cell wall isnormally under substantial
mechanical stress. Ifcell-wall stress and consequent mechanical
fail-ure are contributing factors to the ultrafast cellseparation,
then altering turgor pressure shouldinfluence the likelihood of
separation. To test
this hypothesis, we exposed an unsynchronized,growing population
of cells to oscillatory changesin medium osmolarity over a range of
100 to500 mM in order to modulate turgor pressureand cell-wall
stress and recorded the time of pop-ping with respect to the phase
of the oscillatorycycle for hundreds of individual popping
events.We observed a large dose-dependent enrichmentof popping
events during the intervals when me-dium osmolarity was being
lowered (downshift),which corresponds to an increase in turgor
pres-sure and cell-wall stress, and a depletion of pop-ping events
during the intervals when mediumosmolarity was being raised
(upshift) (Fig. 1C andfig. S2). Thus, an externally induced
increase incell-wall stress promotes popping, whereas a de-crease
in wall stress delays popping, confirmingthe involvement of
cell-wall stress in determiningthe likelihood of popping.A further
prediction of the stress-driven crack
propagationmodel inwhich failure is initiated atone random point
along the periphery is thatafter splitting, when stress has been
released, thetwo daughter cells will remain connected at ahinge
point opposite the initial site of failure. Toprobe the relative
orientation of the two daugh-ter cells after popping, we tracked
the fate of theouter wall (Fig. 1A) relative to the septal wall
af-ter cell separation using fluorescent wheat germagglutinin (WGA)
and three-dimensional (3D)structured illumination microscopy (3D
SIM).WGA binds to N-acetylglucosamine residues inthe cell wall (19)
and does not penetrate into theseptum because of its size and can
therefore beused to selectively label the S. aureus outer wall(20).
In nearly all of the daughter cell pairs ob-served (39 of 40), the
two sections of the pre-vious outer wall were still partially
connectedafter cell separation, and in all cases (40 of 40)they
appeared to have rotated around a hinge(Fig. 1D, 10 min, and movie
S3). In addition, wefollowed WGA-labeled live cells with
epifluo-rescence microscopy and observed two WGA la-beling patterns
after popping: the hinged patternas observed with 3D SIM (Fig. 1E,
left) and anonhinged pattern (Fig. 1E, right) that resemblesthe
labeling pattern reported previously (20).By correlating
epifluorescence microscopy toscanning electron microscopy (SEM), we
realizedthat the nonhinged pattern corresponds to cellswith their
hinge points oriented at the top orbottom surface of the cells
relative to the coverslip(Fig. 1F). Thus, daughter-cell separation
in S. aureusis achieved through mechanical crack propaga-tion that
initiates at some point around the pe-ripheral ring, connecting the
two daughter cells,and rapidly propagates circumferentially,
resultingin a hinge-like rotation.Given that popping occurs so
quickly, we ques-
tioned whether cell volume and surface areachange during this
process. It has been suggestedthat cell volume doubles at the
moment of cellseparation as a result of the septum expanding
tocover one-half of the new spherical cell (8, 21),whichwould
require substantial water influx overa very short time frame. To
address this question,we tracked growth of individual S. aureus
cells
574 1 MAY 2015 • VOL 348 ISSUE 6234 sciencemag.org SCIENCE
1Department of Chemistry, Stanford University, Stanford,
CA94305, USA. 2Department of Biochemistry, StanfordUniversity
School of Medicine, Stanford, CA 94305, USA.3Howard Hughes Medical
Institute (HHMI), StanfordUniversity School of Medicine, Stanford,
CA 94305, USA.4Department of Microbiology and Immunology,
StanfordUniversity School of Medicine, Stanford, CA 94305,
USA.5Department of Bioengineering, Stanford University,Stanford, CA
94305, USA.*These authors contributed equally to this work.
†Correspondingauthor. E-mail: [email protected]
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using the membrane dye FM 4-64 (Life Technol-ogies, Grand
Island, NY), and estimated cell vol-ume and surface area from the
2D cell outlinesby assuming a prolate cell shape (Fig. 2A).
Over-laying the 2D cell outlines from different stagesin the cell
cycle (Fig. 2B, inset) revealed that agrowing S. aureus cell
increases both its volumeand surface area throughout the cell
cycle, ac-companied by an overall increase in the cell as-pect
ratio after a small initial decrease (Fig. 2B).This small initial
decrease corresponded to a phasein which the two daughter cells
gradually (withinminutes) became more round and more sepa-rated
after popping. Following single cells andtheir progeny, we observed
a continuous increasein cell volume for each microcolony over
severalgenerations (Fig. 2C), which is consistentwith thecontinuous
exponential volume increase that has
been described for E. coli and other bacteria (22).With respect
to this continuous growth, the vol-ume change upon popping is
negligible on av-erage (Fig. 2D). Consistent with this, using
3Ddeconvolution fluorescence microscopy with aS. aureus strain
expressing cytoplasmic greenfluorescent protein (GFP), we observed
only min-imal changes in cell volume and GFP intensity(fig. S3)
after cell separation. Last, we estimatedchanges in cell surface
area from 2D cell outlinesby assuming a prolate cell shape, which
revealeda modest net decrease in surface area upon pop-ping (Fig.
2E and fig. S4), which is consistent witha geometric conversion
from hemi-ellipsoidal toellipsoidal shape, given constant volume. A
de-crease in surface area during popping indicatesthat the cell
wall must have been under tensilestress before popping, which is in
line with the
hypothesis that cell-wall stress contributes todaughter-cell
separation.We next questioned whether the septum ex-
pands to become one half of the new daughter’ssurface (8, 21),
given that the total surface areadecreases upon popping. To
determine the rel-ative contributions of the previous outer walland
septum to the surface of the new daughter,we used WGA pulse-chase
labeling and 3D SIMas described above. We found that ~73% of thenew
daughter’s surface was represented by theold wall regardless of
cell size (or stage in the cellcycle) (fig. S5), similar to the
ratio before cell sep-aration (Fig. 2F), indicating that the septum
con-stitutes only ~1/4 of the new daughter’s surfaceand does not
expand noticeably in surface areaupon popping, which is contrary to
the doublingof septal surface area assumed previously (8).
SCIENCE sciencemag.org 1 MAY 2015 • VOL 348 ISSUE 6234 575
FM WGAPhase
0 ms 1 ms
1 µm
FM WGAPhase
0 minchase
10 minchase
NHS-568 WGA-488 Overlay
0 20 40 60 80 100 1200
10
20
30
40
50
60
70
80
90
# of
Pop
ping
s
0
20
40
60
80
100
120
140
160
180
200
Time since start of cycle (s)
Con
c. o
f Sor
bito
l (re
d lin
e) (
mM
)0
min
3 m
in
FM WGAPhase Overlay FM WGAPhase Overlay
~20% frequency~80% frequency
0 2 4 6 8 10 120
1
2
3
4
5
6
7
8
Cou
nts
Separation duration (ms)
outer wall
septum
peripheral ring
Downshift:high wall stress
Upshift:low wall stress
Fig. 1. Daughter cell separation in S. aureus occurs within
millisecondswith characteristics of mechanical crack propagation.
(A) A schematicdiagram of the cell wall before daughter cell
separation. (B) Snapshots ofS. aureus strain Newman “popping”
(inset) and histogram of daughter cellseparation duration captured
by means of phase contrast microscopy at1000 frames/s (n = 16
popping events). (C) Distribution of cumulative countsof popping
events plotted over the 2-min oscillatory period for 200 mM
os-motic shocks. The red line denotes the concentration of sorbitol
in the medium,and the dashed line denotes average popping counts,
assuming a uniformdistribution (n = 400 popping events). (D) 3D SIM
images of fixed Newman cells
labeled with fluorescent WGA (WGA-488, green), which marks the
outer wall andfollowed by 0 or 10min of growth in the absence of
WGA. Cell surfaces and septawere stained with an amine reactive dye
(NHS-568, red). (E) Time-lapse epi-fluorescence images of Newman
cells labeled with WGA (green) before (0 min)and after (3 min)
popping. Corresponding phase-contrast (gray), membranestaining with
FM 4-64 (red), and overlay of WGA and FM signals are alsodisplayed.
Two types of old wall geometry after popping were observed:hinged
(left, ~80%) and nonhinged (right, ~20%). (F) Correlative light
andSEM on Newman cells labeled with WGA followed by 10-min chase
showingthe two types of WGA labeling patterns as in (E). Scale
bars, 1 mm.
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Additionally, the finding that the previous outerwall makes up
~3/4 of the new daughter’s cellwall (as opposed to half) suggests
that there mustbe new wall synthesis in the outer wall as well asat
the septum to sustain the continuous surfaceexpansion required for
cell size homeostasis gen-eration after generation.Because our data
suggest that accumulation of
stress in the cell wall plays an important role inthe ultrafast
daughter cell separation, we soughtto model the stress distribution
in the cell wallbefore popping using a continuum
elastostaticapproach. On the basis of previous cryogenicelectron
microscopy data (7) and constraints de-termined by our experimental
measurements oncell volume and surface area, we built a 3D
finiteelement model of a “ready-to-pop” S. aureus cellwall as a
prolate ellipsoidal shell with two sep-arated septal plates that
are connected with aperipheral ring (fig. S6). We assumed that
theperipheral ring does not grow asmuch as the restof the cell once
septation begins because it is notin direct contact with the
cytoplasmicmembranewhere new wall material is incorporated
(sup-
plementary text 1 and fig. S7). After inflating themodeled cell
wall with a uniform turgor pressurein both compartments, we
calculated the vonMises stress (a criterion for material failure)
for
the entire surface. Indeed, the vonMises stress atthe peripheral
ring was found to be higher thanelsewhere in the outer wall (Fig.
3, A and B, andmovie S6). The high von Mises stress in the
576 1 MAY 2015 • VOL 348 ISSUE 6234 sciencemag.org SCIENCE
0 10 20 30 40 50 60 700
0.5
1
1.5
2
2.5
3
Time (min)
Vol
ume
(µm
3 )
Total Volume
Pop Pop
Corrected relative surface area change
Cou
nts
0.5
0.6
0.7
0.8
0.9
1
After popBefore pop
Fra
ctio
n of
“ol
d” s
urfa
ce
0 0.2 0.4 0.6 0.8 11.2
1.25
1.3
1.35
Fraction into cell cycle
Asp
ect R
atio
FM 4-64 Outlined
20 min
30 min
80 min
Corrected relative volume change
Cou
nts
0
5
10
15
20
0 0.1 0.2 0.3 0.4 0.5 0 0.1 0.2 0.30
5
10
15
Cell Outline Overlay
NHS-568 WGA Overlay Extracted
Pop
Fig. 2. Cell volume increases continuously throughout the cell
cycle.(A) Time-lapse images of S. aureus cells stained with FM 4-64
(left) and out-lined by fitting with ellipses (right). (B) Average
aspect ratio of S. aureus cellsthroughout the cell cycle (from
immediately after previous popping to ready-to-pop) and overlay of
the cell outlines (inset) from a typical cell at different pointsof
the cell cycle colored from blue (early) to red (late). Error bars
denotestandard errors (n = 27 cells). Red bars on top indicate the
time fraction intothe cell cycle when septation starts (left, 0.35
T 0.03 SEM) and completes(right, 0.77 T 0.02 SEM), respectively (n
= 26 cells). (C) Representative tracesof cell volume as a function
of time following a microcolony starting from asingle cell; solid
blue traces indicate cell volumes of individual cells
beforepopping, and the dashed black line denotes the total cell
volume of all the cells
present at a given time. Cell volume and surface area were
estimated from the2D cell outlines by fitting to ellipses and
assuming prolate cell shapes (that eachcell was rotationally
symmetric around the long axis). (D and E) Distribution ofrelative
changes in (D) volume and (E) surface area during popping, after
cor-recting for baseline growth rate. The black solid line
represents kernel densityestimate of the distribution, and the red
dashed line denotes the average (2 T10% SD for volume, –11 T 6.5%
SD for surface area; n = 69 division events).(F) (Top) 3D SIM
images and corresponding extracted data (fig. S5); (bottom)fraction
of old surface before (0.71 T 0.01 SD; n = 15 cells) and after
(0.73 T 0.03SD; n = 36 cells) popping. Cells were modeled as
ellipsoids, and the contributionof the old, WGA-labeled wall to the
daughter cells’ total surface area was mea-sured by fitting a plane
to the old/new boundary (fig. S5A). Scale bars, 1 mm.
0.5
0.4
0.3
0.2
0.1
0
von Mises Circumferential Axial
Fig. 3. High stress in the peripheral ring prepares the cell for
popping. (A) von Mises stressdistribution in the “ready-to-pop” S.
aureus cell wall (fig. S6, state 3) modeled as a linear elastic
material(details of model construction are provided in the
supplementary materials). Color represents therelative magnitude of
stress. The stress at the peripheral ring (red arrow), where the
cell wall splits openduring popping, is higher than elsewhere in
the outer wall. (B) Enlarged views of a cut-through slice ofthe
cell in (A) shows high von Mises stress at the peripheral ring (red
arrow) as well as the stressdistribution in the circumferential and
axial directions, respectively.
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peripheral ring was mostly due to the high cir-cumferential
stress (Fig. 3B, supplementary text2, and fig. S8) resulting from
differential growthof the peripheral ring compared with the rest
ofthe cell wall. The axial stress, which resultsmain-ly from turgor
pressure (fig. S9), is actually lowerin the peripheral ring than
elsewhere (Fig. 3B).Modeling a series of intermediate stages
through-out the growth and division cycle that includesthe growth
of the septum resulted in an increasein the overall cell aspect
ratio during growth(movie S6), which is similar to our
experimentalobservations (Fig. 2B).Last, we examined the cell
surface closely with
high-resolution SEM in order to search for po-tential features
of mechanical failure or fracture,
and we noticed structures in a subpopulation ofcells that
appeared to be perforation-like holesand cracks along the
peripheral ring. Similarstructures have been observed with atomic
forcemicroscopy on hydrated live cells (23). To deter-mine whether
the appearance of these structureswas cell-cycle dependent, we
correlated SEMwithfluorescence microscopy on FM 4-64–stainedcells.
The holes and cracks were observed mostlyon cells at later stages
in the cell cycle withcompleted septa (Fig. 4, A to D).
Specifically, 53out of 54 cells with visible holes had
completedsepta (98%), whereas only 53 of 108 cells withcompleted
septa had holes (49%), suggesting thattheholes are formedafter the
septum is completed.Enrichment of WGA binding at the peripheralring
region was also observed when holes werepresent (fig. S10),
suggesting that these holes aretrue structural changes permitting
access of largeproteins through the wall that are excluded
atearlier stages. These perforations are likely pointsof mechanical
failure that could initiate a prop-agating crack. Although the
axial stress necessaryfor circumferential crack propagation is
relativelylow at the peripheral ring, the presence of
per-forationswill lead to locally high stresses at the tipsof the
cracks (24) that could be sufficient to drivepropagation.
Consistent with this hypothesis, thedistribution of the perforation
lengths observedwith SEM featured a cutoff length so that
mostperforations were
-
ACKNOWLEDGMENTS
We thank P. A. Levin for strain Newman; A. Cheung for
RN6390∆spa;W. M. Nauseef for plasmid pCM29; J. Bose and K. W.
Bayles forUAMS-1 and UAMS-1 Datl; J. Mulholland and L. Joubert in
the CellSciences Imaging Facility at Stanford for technical
assistance with 3DSIM (funded by National Center for Research
Resources awardnumber 1S10OD01227601) and SEM; S. Lou and L. Harris
for helpwith the blind SEM analysis; M. Pinho, F. Chang, and L.
Cegelski fordiscussions; and J. Shaevitz, D. Fisher, L. Harris, and
the anonymousreviewers for helpful comments on the manuscript. X.Z.
wassupported by a Stanford Interdisciplinary Graduate
Fellowship;
D.K.H. was supported by the Stanford Cell and Molecular
BiologyTraining Grant (T32-GM007276); E.R.R. was supported by
aDistinguished Postdoctoral Fellowship from the Simbios NIH
Centerfor Biomedical Computation (U54-GM072970); E.F.K. was
supportedby the James S. McDonnell Foundation Postdoctoral
Fellowship Awardin Studying Complex Systems; T.K.L. was supported
by a SiebelScholars Graduate Fellowship and an NIH Biotechnology
TrainingGrant; K.C.H. was funded by a NIH Director’s New Innovator
Award(DP2OD006466); and J.A.T. was funded by HHMI, the
NationalInstitute of Allergy and Infectious Diseases (R01-AI36929)
and theStanford Center for Systems Biology (P50-GM107615).
SUPPLEMENTARY MATERIALS
www.sciencemag.org/content/348/6234/574/suppl/DC1Materials and
MethodsSupplementary TextFigs. S1 to S14Table S1References
(32–36)Movies S1 to S7
27 October 2014; accepted 25 March
201510.1126/science.aaa1511
PROTEIN DYNAMICS
Direct observation of hierarchicalprotein dynamicsJózef R.
Lewandowski,1*† Meghan E. Halse,1‡ Martin Blackledge,2* Lyndon
Emsley1,3*
One of the fundamental challenges of physical biology is to
understand the relationshipbetween protein dynamics and function.
At physiological temperatures, functional motionsarise from the
complex interplay of thermal motions of proteins and their
environments.Here, we determine the hierarchy in the protein
conformational energy landscape thatunderlies these motions, based
on a series of temperature-dependent magic-anglespinning
multinuclear nuclear-magnetic-resonance relaxation measurements in
a hydratednanocrystalline protein. The results support strong
coupling between protein and solventdynamics above 160 kelvin, with
fast solvent motions, slow protein side-chain motions,and fast
protein backbone motions being activated consecutively. Low
activation energy,small-amplitude local motions dominate at low
temperatures, with larger-amplitude,anisotropic, and functionally
relevant motions involving entire peptide units becomingdominant at
temperatures above 220 kelvin.
Proteins must traverse complex conforma-tional energy landscapes
to perform theirphysiological function. This is achievedthrough
thermally activated fluctuations(1), and when specific molecular
motions
cease at low temperatures, function also ceases orbecomes
reduced (2, 3). Understanding the hi-erarchy of these motions thus
holds the key tounderstanding how proteins function on a mo-lecular
level at physiological temperatures. Contraryto expectations, and
despite large differences instructure and function between proteins
of dif-ferent families, dynamic properties appear toexhibit common
general features, in particularapparent transitions between
different dynamicregimes as a function of temperature. These
tran-sitions are thought to occur as a result of cou-pling between
proteins and the surroundingsolvent (4).
Observing and understanding protein dynam-ic transitions has
been a focus of many fields ofresearch over the past 40 years,
including neu-
tron scattering (5),Mössbauer spectroscopy (4, 6),Terahertz
spectroscopy (7), dielectric spectroscopy(4), differential scanning
calorimetry (8), x-raycrystallography (2, 9), and molecular
dynamicssimulation (10, 11). This relative wealth of infor-mation
has not, however, led to a consensus pic-ture of dynamical
transitions and their origins,with different techniques detecting
distinct pro-cesses, leading to apparently contradictory
de-scriptions (12, 13). Thismay be due to the widelyvarying
conditions required for the diverse tech-niques or to the
sensitivity of the different phys-ical measurements to dynamics
occurring ondifferent time scales.Herewemeasure, in a single
sample, a set of 13
different nuclear magnetic resonance (NMR) ob-servables that are
sensitive to dynamics occur-ring on different time scales and in
different partsof the system over temperatures from 105 to 280 Kin
the fully hydrated crystalline protein GB1, asmall globular protein
specifically binding to an-tibodies. The analysis of multiple
probes that re-port on the different structural components ofthis
complex system allows us to develop a com-plete and coherent
picture of the dynamic pro-cesses across the whole temperature
range, as
578 1 MAY 2015 • VOL 348 ISSUE 6234 sciencemag.org SCIENCE
1Université de Lyon, Institut de Sciences
Analytiques(CNRS/ENS-Lyon/UCB-Lyon 1), Centre de
RésonanceMagnétique Nucléaire à Très Hauts Champs,
69100Villeurbanne, France. 2Université Grenoble Alpes, Institut
deBiologie Structurale (IBS), F-38044 Grenoble, France; CNRS,IBS,
F-38044 Grenoble, France; CEA, IBS, F-38044 Grenoble,France.
3Institut des Sciences et Ingénierie Chimiques, EcolePolytechnique
Fédérale de Lausanne (EPFL), CH-1015Lausanne,
Switzerland.*Corresponding author. E-mail:
[email protected](J.R.L.); [email protected]
(M.B.); [email protected](L.E.) †Present address: Department of
Chemistry, University ofWarwick, Coventry CV4 7AL, UK. ‡Present
address: Department ofChemistry, University of York, York YO10 5DD,
UK.
hydration water+
side chain
side chain
R1,CP : 1H
R1:13CH3,
15Nζ :
13CH3, 15Nζ
bulk water
R1, 1H: 1H
hydrated protein crystals
fast (ps-ns): R1slow (ns-ms): R1ρ & R2’
backboneR1:
15N, 13C’R1ρ & R2’:
15N, 13C’, 13CαR1ρ & R2’:
Fig. 1. Cartoon representation of the loca-tion of motions and
the relaxation ratesthat are most sensitive to those motions.The
rates in green, purple, and red report onbackbone, side-chain, and
solvent motions,respectively.
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aureusStaphylococcusMechanical crack propagation drives
millisecond daughter cell separation in
TheriotXiaoxue Zhou, David K. Halladin, Enrique R. Rojas, Elena
F. Koslover, Timothy K. Lee, Kerwyn Casey Huang and Julie A.
DOI: 10.1126/science.aaa1511 (6234), 574-578.348Science
, this issue p. 574Sciencegradually, tiny imperfections in the
mother cell wall were seen to crack open, leaving two daughter
cells linked by a hinge.who examined dividing cells with
millisecond precision using high-speed videomicroscopy. Rather than
proceeding
,et al. proceeds much like the cracking of an egg. So say Zhou
Staphylococcus aureusDaughter cell separation in Pop goes the
coccus
ARTICLE TOOLS
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MATERIALSSUPPLEMENTARY
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REFERENCES
http://science.sciencemag.org/content/348/6234/574#BIBLThis
article cites 33 articles, 9 of which you can access for free
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