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

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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: theriot@stanford.edu

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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

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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

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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.

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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

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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: j.r.lewandowski@warwick.ac.uk(J.R.L.); martin.blackledge@ibs.fr (M.B.); lyndon.emsley@epfl.ch(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+

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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 http://science.sciencemag.org/content/348/6234/574

MATERIALSSUPPLEMENTARY http://science.sciencemag.org/content/suppl/2015/04/29/348.6234.574.DC1

REFERENCES

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