DOI: 10.1126/science.1221886, 355 (2012);337 Science

et al.Sophie DumontActive and Passive Force GenerationDeformations Within Moving Kinetochores Reveal Different Sites of

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Deformations Within MovingKinetochores Reveal Different Sitesof Active and Passive Force GenerationSophie Dumont,1*† E. D. Salmon,2 Timothy J. Mitchison1

Kinetochores mediate chromosome segregation at mitosis. They are thought to contain bothactive, force-producing and passive, frictional interfaces with microtubules whose relativelocations have been unclear. We inferred mechanical deformation within single kinetochoresduring metaphase oscillations by measuring average separations between fluorescently labeledkinetochore subunits in living cells undergoing mitosis. Inter-subunit distances were shorter inkinetochores moving toward poles than in those moving away. Inter-subunit separation decreasedabruptly when kinetochores switched to poleward movement and decreased further whenpulling force increased, suggesting that active force generation during poleward movement compresseskinetochores. The data revealed an active force-generating interface within kinetochores and aseparate passive frictional interface located at least 20 nanometers away poleward. Together,these interfaces allow persistent attachment with intermittent active force generation.

Kinetochores link chromosomes to micro-tubules in the mitotic spindle and gen-erate forces for chromosome movement

(1). Mammalian kinetochores consist of morethan 80 proteins (2), many of whose positionshave been mapped with nanometer-level pre-cision (3–9). Kinetochores are thought to inter-act both actively and passively with microtubules,but the molecular identity and physical nature

of the relevant interfaces are unclear (10). Ac-tive interfaces generate pulling force by trans-ducing the free energy of microtubule plus-enddepolymerization into mechanical work (11, 12)at kinetochores moving poleward (P). Passiveinterfaces generate molecular friction when ki-netochores are forced to slide over the micro-tubule lattice toward plus-ends (13). This occurswhen kinetochores moving away from poles (AP)

are pulled by P sisters and also when polewardflux pulls microtubules away from stationarykinetochores in some systems (14, 15).

Mechanical compliance (deformation in re-sponse to force) should report on the positionand direction of forces acting on kinetochores (6).We used the separation between red and greenprobes to measure kinetochore deformation inliving Ptk2 cells. The CenpC probe reported onkinetochore subunits near the chromosome, andCdc20 and Hec1 probes reported on subunits nearmicrotubules. CenpC binds DNA (16), Cdc20reports on microtubule-binding protein KNL1,and Hec1 binds microtubules and is part of theload-bearing Ndc80 complex (Fig. 1A) (17). Meta-phase oscillations, in which kinetochores switchfairly regularly between P and AP movement,provided natural force fluctuations (18, 19).

A one-dimensional map of kinetochore sub-units (Fig. 1A) was generated from light-levelmeasurements in fixed cells and in vitro struc-tural work (6). We coexpressed two fluorescentkinetochore protein pairs in Ptk2 cells, either

Fig. 1. Tracking kinetochore probe pairs. (A) Simplified one-dimensional representation of kinetochore organization (6),with probe positions indicated by stars. (B) Experimental sche-matic. Kinetochore positions oscillate during metaphase. Pdenotes movement toward a pole, and AP denotes movementaway. Probes are indicated as colored circles. (C) Representa-tive images of EYFP-Cdc20 and mCherry-CenpC at one timepoint. Scale bar, 5 mm. (D) Enlarged image of the single kine-tochore pair indentified in (C). Scale bar, 500 nm. (E) Repre-sentative tracks of a single kinetochore [circle-identified in(C)], with direction indicated. Dashed lines indicate direction reversals. (F)Inter-probe distance versus time from the tracks in (E). Dotted lines indicatedirection reversals. (G and H) Histograms of all CenpC(N)–Cdc20 distancemeasurements during metaphase (G) and anaphase (H). The average of themeans of P and AP distributions is consistent with the reported CenpC–KNL1

N-terminal distance in fixed cells (53 to 54 nm) (6, 17). (I) Histograms of allCenpC(N)–Hec1(C) distance measurements during metaphase. The noise insingle traces and histogram widths [(F) to (I)] stemmed from both experimentalnoise (such as centroid determination and two-color registration) andbiological variation. Stars indicate histogram distribution means.

1Department of Systems Biology, Harvard Medical School,Boston, MA 02115, USA. 2Department of Biology, Universityof North Carolina at Chapel Hill, Chapel Hill, NC 27599, USA.

*Present address: Department of Cell and Tissue Biology, Uni-versity of California, San Francisco, San Francisco, CA 94143,USA.†To whom correspondence should be addressed. E-mail:sophie.dumont@ucsf.edu

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mCherry-CenpC [N-terminal label, CenpC(N)]and EYFP-Cdc20, or mCherry-CenpC and Hec1-EGFP [C-terminal label, Hec1(C)]. These probesdid not perturb metaphase oscillations (17). TheCdc20 probe was brighter than the Hec1 probeand was used for most experiments. We imagedred and green probes simultaneously by meansof confocal fluorescence microscopy with a di-chroic beam splitter and a single camera. Cellswere compressed with an agarose pad so as tokeep kinetochores in focus (Fig. 1B), and com-pression did not perturb oscillations (table S1)(20). Occasionally, drastic compression was usedto induce unusually large forces (17). Using avariant (3, 6) of SHREC (single-molecule high-resolution colocalization) (21) in vivo (Fig. 1C), wemeasured the distance between centroids of theprobes (Fig. 1D) every 10 s during several oscil-lation cycles (Fig. 1E and movies S1 and S2) (17).

We first asked whether P and AP kinetochoreswere on average different. Graphs of interprobedistance over time for a single kinetochore (Fig.1F), and a histogram of many kinetochores (Fig.1G), revealed CenpC(N)–Cdc20 distances of47 T 20 nm in P kinetochores (n = 525 measure-ments), and 55 T 19 nm in AP kinetochores (n =569 measurements). These values differ with highsignificance (P=10−10) (table S2). Of kinetochoresimaged, 93% displayed a greater mean inter-probedistance in AP than in P state (fig. S1A). The 15%shorter CenpC(N)–Cdc20 distance in P as com-paredwithAP kinetochores could stem from eithera mechanically compliant CenpC(N)–Cdc20 link-age that responds to force or from biochemicalchanges that relocalize a probe molecule.

To distinguish mechanical from biochemi-cal causes of inter-probe distance change, wefirst asked whether it also occurred during ana-phase, which drastically changes kinetochore bio-chemistry (22). Anaphase kinetochores are biasedtoward P motion to segregate chromosomes, butAP transients still occur in Ptk2 cells with sim-ilar velocities to metaphase (18, 23), probablybecause of polar ejection forces (17). Inter-probedistances in anaphase were statistically indistin-guishable from those at metaphase (P = 0.2 forP, and P = 0.4 for AP). The mean CenpC(N)–Cdc20 distance in anaphase was 49 T 22 nmin P kinetochores (n = 204 measurements) and57 T 20 nm in AP kinetochores (n = 89 measure-ments) (Fig. 1H and table S2). Just as for meta-phase, anaphase P and AP inter-probe distanceswere statistically different from each other (P =0.004), and all but one kinetochore displayed agreater mean inter-probe distance in AP than inP state.

Next, we measured the inter-probe distanceat metaphase between CenpC(N) and Hec1(C),which is part of the main load-bearing complex,Ndc80. The mean CenpC(N)–Hec1(C) distancewas 38 T 15 nm in P kinetochores (n = 564 mea-surements) and 43 T 17 nm in AP kinetochores(n = 487 measurements) (Fig. 1I and table S2).These values differwith high significance (P=10−5),and their average was consistent with the localiza-

tion of Hec1 in fixed cells (6). Here too, the meanbehavior was representative of individual kineto-chores (fig. S1B). Observing similar changes indistance fromCenpC to twodifferent probes report-ing on the microtubule-binding KMN network(2, 17) strongly argues that length changes be-tween P andAP are due tomechanical compliance.

In a mechanical model, force changes anddeformation changes are expected to be closelycorrelated in time. To test whether this require-ment is met, we tracked sister kinetochores dur-ing direction reversals (Fig. 2A) in which forceschange abruptly. We measured the distance with-in one kinetochore and distance between that ki-

netochore and its sister, where the latter reportson force between kinetochores from centromerestretch, during 40 s of observation (five sequen-tial images) centered on reversals, averaged overall reversals in the data set (Fig. 2, B to D).Kinetochore direction (Fig. 2B) (19) and inter-kinetochore distance (Fig. 2C) both changedabruptly at reversals. P-to-AP andAP-to-P reversalsoccurred at markedly different inter-kinetochoredistances (P = 10−13) (Fig. 2C and table S3) andtimes. The leading P kinetochore reversed on av-erage 6 T 11 s (n = 133 reversals) before thetrailing AP one (and in 15% of the cases afterthe AP one), whichwas illustrated by a shift on the

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Fig. 2. Distance between sister kinetochores andbetween probes in one kinetochore during directionreversals. (A) Single kinetochore position (top) andinter-kinetochore distance (bottom) over time for asister kinetochore pair (that of Fig. 1C). Black starsindicate all direction reversals, and dotted lines indicatedirection reversals for the circle-identified kinetochore.(B) Mean kinetochore position, (C) inter-kinetochore dis-tance, and (D) CenpC(N)–Cdc20 distance over time formetaphase P-to-AP (n = 104 traces) and AP-to-P (n =98 traces) reversals (P in red, AP in blue). AP-to-P rever-sals are positioned 6 s later than P-to-AP reversals onthe time axis so as to reflect the average time betweenthem. Bars indicate SEM. Direction reversals cause abrupt changes in inter-probe distance within akinetochore, which is consistent with a mechanical response to a change in force.

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Fig. 3. Kinetochore velocity and CenpC(N)–Cdc20 distance at different inter-kinetochore distances. (A)Kinetochore velocity as a function of inter-kinetochore distance during metaphase P (n = 104 traces)and AP (n = 104 traces) movement before and after P-to-AP reversals, respectively, when the highestinter-kinetochore distances are visited. P velocity decreases and AP velocity increases with inter-kinetochoredistance, as expected if P kinetochores generate force and inter-kinetochore distance is a metric for force.We include data from drastic spindle compression to probe even higher forces (17). (B) CenpC(N)–Cdc20distance as a function of inter-kinetochore distance during metaphase P (n = 547 measurements) and AP(n = 529 measurements) movement. Inter-probe distance decreases with inter-kinetochore distance in Pkinetochores, which is consistent with a force-deformation relationship. Lines are best linear fits.

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time axis (Fig. 2, B to D). Inter-kinetochore dis-tance tended to increase during coordinated move-ment because the leading P kinetochore movedslightly faster (on average) than did its trailingAP sister (fig. S2, A to D). P-to-AP reversal oc-curred on average 7 T 15 s (n = 151 reversals) afterthe maximum inter-kinetochore distance had beenreached; inter-kinetochore distance was 2.7 T1.0 mm at this reversal and decreased abruptly af-terward as both sisters transiently moved towardeach other (Fig. 2C). AP-to-P reversal occurredafter this decrease in inter-kinetochore distance at1.9 T 0.6 mm,which is close to the globalminimum(Fig. 2C and fig. S2D). The above data areconsistent with a mechanical model (24) in whichhigh centromere stretch favors P-to-AP reversaland low stretch favors AP-to-P reversal (14).

Within our time resolution, changes in move-ment direction and CenpC(N)–Cdc20 distances co-incided (17). CenpC(N)–Cdc20 distances increasedabruptly after P-to-AP reversals and decreasedabruptly after AP-to-P reversals (Fig. 2D; fig. S2,E and F; and tables S3 and S4), which was alsotrue at anaphase (table S3). That change in forcesexerted by P and on AP kinetochores—measuredthrough inter-kinetochore stretch—coincides closelywith changes in inter-probe distances within ki-netochores supports the mechanical interpretationof inter-probe distances and constrains time scalesassociated with force transitions.

Extent of deformation is expected to correlatewith magnitude of force in a mechanical model.To test this, we plotted inter-probe distance againstinter-kinetochore distance. Here too, chromosome

oscillations provided natural changes in kineto-chore forces (fig. S2); to extend the range of forces,we included measurements from drastically com-pressed cells in which inter-kinetochore stretchwas up to 6 mm. Force between kinetochores isdue to active force from the P kinetochore op-posed largely by friction from the AP kinetochore(14, 15, 24). Velocity of AP kinetochores increasedwith inter-kinetochore distance, as expected ifAP movement is due to pulling by the P sisteropposed by frictional drag at the AP kinetochore(P = 10−4) (Fig. 3A and table S5). Consistent withthis view, velocity of P kinetochores decreasedwith inter-kinetochore distance (P = 0.08) (Fig.3A and table S5) (13). Inter-probe distances in Pkinetochores decreased with inter-kinetochoredistance (P = 10−8) (Fig. 3B and table S5). Thissuggests that P kinetochores that exert more forceare more compressed. No correlation betweeninter-probe and inter-kinetochore distances wasdetected in AP kinetochores (P = 0.6) (Fig. 3Band table S5). When little active force was gen-erated at low inter-kinetochore distances, both Pand AP kinetochores displayed similar inter-probe distances (Fig. 3B).

Because P kinetochores, where active forceis generated, are internally compressed relativeto AP kinetochores and because the larger theforce generated at P kinetochores the more com-pressed they are, we developed a simple me-chanical model in which frictional forces aregenerated at a more outward position than thatof active forces in P kinetochores, leading tointernal compression (Fig. 4A, figs. S3 and S4,

and tables S6 and S7) (17). AP kinetochores,which lack active force generation, are extendedby pulling from centromeric chromatin balancedby friction at the kinetochore-microtubule interface.Kinetochore deformation need not vary linearlywith force and may either represent changes inlength of protein linkages or their reorientationand reorganization. We extended this mechan-ical model in light of previous mapping data(Fig. 4B) (6). The active, force-generating inter-face for P movement lies internal to the meanposition of the Hec1(C) probe, and the passive,frictional interface is at least 20 nm outward ofthe active interface (Fig. 3B). This makes themicrotubule-binding site at the outer end of Hec1(Fig. 4A, 0 nm mean position) a good candidatefor passive force generation (17).

Spatially separated passive and active inter-faces at kinetochores, whether comprising dif-ferent molecules or different interactions of thesame molecule with microtubules in differentlocations (17), may represent a design principlewith important advantages. The passive friction-al interface binds persistently to microtubulesindependently of the microtubule dynamics stateor movement direction, ensuring segregation ac-curacy. The active interface consumes energyto efficiently move kinetochores poleward butcan evolve without the constraint of requiringpersistent attachment. Together, both interfacesallow the kinetochore to harness force from de-polymerizing microtubules without losing grip.That said, kinetochores may be able to functionusing only the passive interface—for example, insystems without anaphase A (25) or where mi-crotubules polymerize continuously at kinetochores,even during anaphase (26). In these systems, seg-regation forces will be generated elsewhere inthe spindle and presumably transmitted to chro-matin via molecular friction.

References and Notes1. S. Santaguida, A. Musacchio, EMBO J. 28, 2511 (2009).2. I. M. Cheeseman, A. Desai, Nat. Rev. Mol. Cell Biol. 9, 33

(2008).3. A. P. Joglekar, K. Bloom, E. D. Salmon, Curr. Biol. 19,

694 (2009).4. S. A. Ribeiro et al., Proc. Natl. Acad. Sci. U.S.A. 107,

10484 (2010).5. R. B. Schittenhelm et al., Chromosoma 116, 385 (2007).6. X. Wan et al., Cell 137, 672 (2009).7. T. J. Maresca, E. D. Salmon, J. Cell Biol. 184, 373

(2009).8. K. S. Uchida et al., J. Cell Biol. 184, 383 (2009).9. Z. Venkei et al., Open Biol. 2, 110032 (2012).

10. A. P. Joglekar, K. S. Bloom, E. D. Salmon, Curr. Opin.Cell Biol. 22, 57 (2010).

11. E. L. Grishchuk, M. I. Molodtsov, F. I. Ataullakhanov,J. R. McIntosh, Nature 438, 384 (2005).

12. D. E. Koshland, T. J. Mitchison, M. W. Kirschner, Nature331, 499 (1988).

13. R. B. Nicklas, J. Cell Biol. 97, 542 (1983).14. A. Khodjakov, C. L. Rieder, J. Cell Biol. 135, 315 (1996).15. P. Maddox, A. Straight, P. Coughlin, T. J. Mitchison,

E. D. Salmon, J. Cell Biol. 162, 377 (2003).16. T. Hori et al., Cell 135, 1039 (2008).17. Materials and methods are available as supplementary

materials on Science Online.18. R. V. Skibbens, V. P. Skeen, E. D. Salmon, J. Cell Biol.

122, 859 (1993).19. B. Akiyoshi et al., Nature 468, 576 (2010).

Fig. 4. Mechanical modelfor kinetochore compliance.(A) Kinetochores and inter-kinetochore chromatin viewedas three springs in series, withprobes indicated by stars. Pkinetochores are compressed,on average, relative to AP ki-netochores. We interpret thisas indicating that an activeforce-generating interface thatis only engaged in P kineto-chores (orange) lies inward ofthe mean positions of Cdc20and Hec1(C), whereas a pas-sive frictional interface thatis engaged in all kinetochores(blue) lies outward of thesemarkers. (B) Preliminary struc-tural interpretation, with inter-probe distances indicated forCenpC(N)–Cdc20 (meanT SEM).The active and passive inter-faces could differ because theycomprise different moleculesor because force generationis restricted to the end of themicrotubule, whereas frictionoccurs all along the embeddedlattice.

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20. S. Dumont, T. J. Mitchison, Curr. Biol. 19, 1086 (2009).21. L. S. Churchman, Z. Okten, R. S. Rock, J. F. Dawson,

J. A. Spudich, Proc. Natl. Acad. Sci. U.S.A. 102, 1419 (2005).22. B. J. Howell et al., Curr. Biol. 14, 953 (2004).23. J. S. Tirnauer, J. C. Canman, E. D. Salmon, T. J. Mitchison,

Mol. Biol. Cell 13, 4308 (2002).24. R. V. Skibbens, C. L. Rieder, E. D. Salmon, J. Cell Sci.

108, 2537 (1995).25. K. Oegema, A. Desai, S. Rybina, M. Kirkham, A. A. Hyman,

J. Cell Biol. 153, 1209 (2001).26. J. R. J. LaFountain Jr., C. S. Cohan, A. J. Siegel,

D. J. LaFountain, Mol. Biol. Cell 15, 5724 (2004).

Acknowledgments: We thank J. Shah for the stable Ptk2EYFP-Cdc20 line, C. Carroll and A. Straight for themCherry-CenpC construct, J. DeLuca for Hec1 constructs,X. Wan for sharing SpeckleTracker, and M. Kirschnerfor equipment loan. We thank I. Cheeseman, P. Choi,S. Churchman, J. DeLuca, M. Ginzberg, Q. Justman,J. Shah, X. Wan, and J. Waters (HMS Nikon ImagingCenter) for discussions. S.D. was supported by the CharlesA. King Trust, Bank of America and by NIH K99GM094335;E.D.S. was supported by NIH R37GM024364; and T.J.M.was supported by NIH R01GM039565. Data are presented inthe supplementary materials.

Supplementary Materialswww.sciencemag.org/cgi/content/full/science.1221886/DC1Materials and MethodsSupplementary TextFigs. S1 to S4Tables S1 to S7References (27–76)Movies S1 and S2

13 March 2012; accepted 14 May 2012Published online 21 June 2012;10.1126/science.1221886

Regional Astrocyte Allocation RegulatesCNS Synaptogenesis and RepairHui-Hsin Tsai,1,2,5 Huiliang Li,6* Luis C. Fuentealba,4,5* Anna V. Molofsky,1,3,5*Raquel Taveira-Marques,6* Helin Zhuang,6 April Tenney,1,2 Alice T. Murnen,1,2,5

Stephen P. J. Fancy,1,2,5 Florian Merkle,4† Nicoletta Kessaris,6 Arturo Alvarez-Buylla,4,5‡William D. Richardson,6‡ David H. Rowitch1,2,4,5‡

Astrocytes, the most abundant cell population in the central nervous system (CNS), are essential fornormal neurological function. We show that astrocytes are allocated to spatial domains in mousespinal cord and brain in accordance with their embryonic sites of origin in the ventricular zone. Thesedomains remain stable throughout life without evidence of secondary tangential migration, evenafter acute CNS injury. Domain-specific depletion of astrocytes in ventral spinal cord resulted inabnormal motor neuron synaptogenesis, which was not rescued by immigration of astrocytesfrom adjoining regions. Our findings demonstrate that region-restricted astrocyte allocation is ageneral CNS phenomenon and reveal intrinsic limitations of the astroglial response to injury.

Astrocytes serve roles essential for normalneurological function such as regulationof synapse formation, maintenance of

the blood-brain barrier (BBB), and neuronal ho-meostasis (1, 2). Although astroglia are region-

ally heterogeneous in terms of gene expressionand their electrical and functional properties(3–5), astrocyte diversification and migrationremain poorly understood. Two generally rec-ognized types of astrocytes are fibrous astrocytes

(FAs) of white matter that express glial fibrillaryacidic protein (GFAP), and protoplasmic astro-cytes (PAs) of gray matter that normally expresslittle or no GFAP. Aldh1L1-GFP and AldoC aremore recently described markers of both PAs

1Howard Hughes Medical Institute, University of CaliforniaSan Francisco, San Francisco, CA 94143, USA. 2Department ofPediatrics, University of California San Francisco, San Francisco,CA 94143, USA. 3Department of Psychiatry/Langley-PorterInstitute, University of California San Francisco, San Francisco,CA 94143, USA. 4Department of Neurosurgery, University ofCalifornia San Francisco, San Francisco, CA 94143, USA. 5Eliand Edythe Broad Center of Regeneration Medicine andStem Cell Research, University of California San Francisco,San Francisco, CA 94143, USA. 6Wolfson Institute for Bio-medical Research and Research Department of Cell andDevelopmental Biology, University College London, LondonWC1E 6BT, UK.

*These authors contributed equally to this work.†Present address: Departments of Stem Cell and Regener-ative Biology and Molecular and Cellular Biology, HarvardUniversity, Cambridge, MA 02138, USA.‡To whom correspondence should be addressed. E-mail:rowitchd@peds.ucsf.edu (D.H.R.); abuylla@stemcell.ucsf.edu (A.A.-B.); w.richardson@ucl.ac.uk (W.D.R.)

Fig. 1. Segmental distribution of fibrous and proto-plasmic astrocytes in spinal cord. (A)Nkx2.2-creERT2(tamoxifen induction E10.5 to E12.5):Rosa26-YFPfate map shows YFP+, GFAP+ cells at the ventral mid-line at P0. YFP, yellow fluorescent protein. (B) In P2Olig2-tva-cre:CAG-GFPmice, astrocytes remain in reg-ister with pMN, whereas Olig2+ OPs distribute wide-ly. (C) Ngn3-cre:Z/EG P1 cord shows intermediatewedge of astrocytes. (D, G, G′) At P2, FAs and PAsin Pax3-cre animals remain dorsally restricted. DAPI,4´,6-diamidino-2-phenylindole. (E and F) Aldh1L1-GFP coexpression with GFAP+ (FAs) and AldoC+ (FAsand PAs) cells. (H to N) Astrocytes from Olig2cre/+spinal cord have a restricted ventral distribution. InOlig2cre/cre nulls, we observe significantly (*P <0.0001) increased p2-type (GFAP+, Pax6+, AldoC+)astrocytes (fig. S1I), which fail to migrate from theventral domain. Scale bars, 200 mm.

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