SUPPLEMENTARY INFORMATION

Transcription

1 SUPPLEMENTARY INFORMATION Figure S1 Examples of spindles assembled on small chromatin bead structures. (a) Spindle assembled on a short string of aligned chromatinbeads. (b-d) Spindles assembled on chromatin-bead clusters of various sizes. Fixed after 70 minutes of spindle assembly. Tubulin, red; chromatinbeads, green. Scale bars, 10 µm. Figure S2 Analysis of chromatin-bead structure bending during spindle assembly. (a, b) Two images from a time-lapse recording of fluorescent tubulin during spindle assembly on a single chromatin-bead string. Bending of a chromatin-bead structure can be seen over time. Beads outline drawn in yellow. Timestamps, min:sec. Scale bars, 10 µm. (c, d) Higher magnification images of the region of chromatin structure bending from the time-lapse recording shown in a and b. (e) Schematic illustration of bending of chromatin linkages between adjacent beads. 1

2 SUPPLEMENTARY INFORMATION Figure S3 Incorporation of non-chromatin ( dud ) beads into strings of chromatin beads to perturb the chromatin signal. Fluorophore-labeled dud beads (green) were mixed with DNA-coated beads (blue) before alignment in the magnetic field. (a, b) Many short strings were observed that had nonchromatin beads at one end, suggesting that these shorter strings result from breakage of longer strings at sites of incorporated dud beads. (c, d) In samples in which spindles were assembled on arrays containing both dud and chromatin-beads, dud beads were found predominantly at ends of chromatin-bead structures and sites of bead structure kinks. Tubulin, red. Scale bars, 10 µm. Supplemental Movie Movie corresponding to data shown in Fig. 5. Dynamic microtubules explore the region around the entire chromatin-bead structure. Fluorescent EB1 was imaged in fields surrounding bipolar spindles assembled on a chromatin-bead string by spinning-disc confocal microscopy. 2

3 Supplementary Information Estimation of the force required to bend chromatin-bead strings The stiffness and flexibility of the chromatin beads alignment comes from the contribution of each chromatin polymer, assembled during the interphase, that take part in the cross-linking of adjacent beads (Supplementary Figure S2). We assume for simplicity that the linkers of two adjacent beads, made of chromatin, have a rod-like shape (length L and cross section S). We estimate the surface of contact S to be ~ 1 µm 2 (fluorescent microscopy observations and electron microscopy of chromatin beads 1 ). During spindle pole formation the stress is applied on a few beads (Supplementary Figure S2). The bending modulus B of chromatin linkers is estimated by two ways: a. Estimation of B is obtained by assuming that the chromatin linkers form a continuous rod-like shape having a Young modulus Y (measured for the chromatin to be Pa 2 and chosen to be 500 Pa), and of radius r (~ 0.5 µm) 2. Linear elasticity allows us to calculate the bending modulus B of a continuous rod shape of radius r and Young modulus Y: B = YI = π 4 Yr4 ~ Nm 2 (I is the moment of inertia of a cylinder) 3. b. Estimation of B from the individual contributions of chromatin-molecules grafted on the beads. These polymers contribute equally to the stiffness between beads. Each chromatin polymer has a cross-section area of ~ 200 nm 2 (assumed to be ~ Rg 2 (where Rg is the radius of gyration of the DNA polymer)). We estimate the chromatin polymers number forming the crosslinker to be The chromatin linker bending rigidity is chosen to be the one measured for a single chromatid fiber assembled in vitro, in single chromatin stretching experiments 4. This

4 bending rigidity is b ~ Nm 2. Each linker contribute to the total bending rigidity B, thus B ~ b ~ Nm 2. Both calculations give results of the same order of magnitude. We choose B ~ Nm 2. The force required to bend a rod shape of length L is F = α YI L 2 = π2 B L 2 We find F ~ 2 nn, for a length L ~ 0.3 µm.

5 Methods Reagents Cytostatic factor-arrested Xenopus laevis egg extracts were prepared as described, and driven into interphase by the addition of 0.3 mm CaCl 5 2. Tubulins labeled with fluorophores 6, monastrol 7, p50/dynamitin 8, and p150-cc1 9 were prepared as previously described. DNAcoated beads were prepared as described 10, except that 2-3 pg DNA was bound to each bead. Only mono-disperse preparations of DNA-coated beads with uniform DNA loading (by Hoechst (Sigma-Aldrich) staining) were used. To make labeled dud beads (not coated with DNA), streptavidin Dynabeads (Dynal) were labeled with Oregon Green-succinimidyl ester (Molecular Probes). EB1 labeled with Alexa-594 (Molecular Probes) was prepared and used as previously described 11. Imaging and Data analysis Fluorescence imaging of spindles for quantitation was done using an Axioplan 2 (Carl Zeiss MicroImaging, Inc.) and a 20X (Plan Neo, NA 0.5) or 40x (Plan Neo, NA 0.75) objective, equipped with an Axiocam MRm camera (Carl Zeiss MicroImaging, Inc.). For high-resolution imaging of microtubule organization and EB1 dynamics, images were acquired using an Axiovert 200M (Carl Zeiss MicroImaging, Inc.) and a 100X (Plan Apo, NA 1.4) objective, equipped with a piezo z-motor (Applied Scientific Instrumentation), an Orca ER CCD camera (Hamamatsu), a spinning-disk confocal head, 488- and 568-nm excitation filters, and krypton and argon ion lasers (Solamere Inc., Utah). For microtubule organization, z-sections (separated by 1.0 µm) were acquired, and maximal intensity projections of entire Z-stacks are displayed.

6 For EB1 imaging, single confocal sections were acquired every 2 seconds. X-Rhodamine or Alexa-488-conjugated tubulin was used at ~300 nm for imaging microtubule distribution. Chromatin-beads were imaged by autofluorescence of the paramagnetic beads. Data analysis was performed using Metamorph software (Universal Imaging Corp.) and Axiovision software (Carl Zeiss MicroImaging, Inc.). For purposes of quantitation, counting was limited to structures containing greater than 6, but fewer than 50 beads, and structures were considered strings if they possessed no more than one defect in the linearity of the structure (Fig. 2e, f; 6c; 7c). DNA-Bead Alignment and Spindle Assembly Chambers for DNA-bead alignment and spindle assembly were constructed by sealing two rubber o-rings (9 mm I.D./12 mm O.D., and 20 mm I.D./27 mm O.D., respectively) in an approximately concentric arrangement to a #1.5 coverglass (Fisher Scientific) using VALAP (1:1:1 mixture of Vaseline, lanolin and paraffin). To regulate temperature and humidity in the chambers, distilled water was added to the space between the two o-rings, and each chamber was placed on a homemade metal block, water-cooled to 16 C using a refrigerated water bath circulator (RTE7, Neslab). A solenoid electromagnet (EM or EM , AZeer Enterprises, Inc.) operating at 6V DC was positioned ~2 mm from the edge of each chamber, such that the chamber was near the center of the magnetic field. 60µl fresh egg extract containing 0.3 mm CaCl 2 was gently added to each chamber, and 5 µl DNA-coated beads (at ~6x10 8 beads/ml in 10mM Hepes (ph 7.7), 1 mm MgCl 2, 100 mm KCl, 150 mm sucrose) were gently dispersed through the extract in the chamber using an 130 µm bore beveled-tip syringe (#701, Hamilton Company). The top of the chamber was then

7 sealed with a second coverglass and silicon grease (Beckman). The sealed chambers containing extract and DNA-beads were incubated for two hours at 16 C to assemble chromatin on the beads. 50 µl fresh cytostatic factor-arrested egg extract was then gently mixed into each sample using a large-bore pipette. The chambers were re-sealed and incubated at 16 C for an additional 30 min to allow the extract to enter mitosis. Chambers were then removed from the magnetic field and cooled on ice for at least 15 min to disassemble microtubule structures, and warmed to 16 C to initiate spindle assembly. Indistinguishable results were obtained with or without depolymerization of microtubules by cold treatment. For fixed timepoint analysis, small sample volumes were removed and fixed as described 5. For real-time observations of spindle assembly, µl samples were removed from chambers on ice, and placed in ~75 µm thick flow cells consisting of two thin strips of doublestick tape separating a coverslip and microscope slide. Each flow cell was sealed with VALAP, and imaged in a temperature-controlled room at ~18 C, as described 5.

8 Supplemental References 1. Reinsch, S. & Karsenti, E. Movement of nuclei along microtubules in Xenopus egg extracts. Curr Biol 7, (1997). 2. Poirier, M. G., Eroglu, S. & Marko, J. F. The bending rigidity of mitotic chromosomes. Mol Biol Cell 13, (2002). 3. Kosevich, A. M., Lifshitz, E. M., Landau, L. D. & Pitaevskii, L. P. Theory of Elasticity (ed. 3) (Butterworth-Heinemann, 1986). 4. Houchmandzadeh, B., Marko, J. F., Chatenay, D. & Libchaber, A. Elasticity and structure of eukaryote chromosomes studied by micromanipulation and micropipette aspiration. J Cell Biol 139, 1-12 (1997). 5. Desai, A., Murray, A., Mitchison, T. J. & Walczak, C. E. The use of Xenopus egg extracts to study mitotic spindle assembly and function in vitro. Methods Cell Biol 61, (1999). 6. Hyman, A. A. Preparation of marked microtubules for the assay of the polarity of microtubule-based motors by fluorescence. J Cell Sci Suppl 14, (1991). 7. Mayer, T. U. et al. Small molecule inhibitor of mitotic spindle bipolarity identified in a phenotype-based screen. Science 286, (1999). 8. Wittmann, T. & Hyman, T. Recombinant p50/dynamitin as a tool to examine the role of dynactin in intracellular processes. Methods Cell Biol 61, (1999). 9. Gaetz, J. & Kapoor, T. M. Dynein/dynactin regulate metaphase spindle length by targeting depolymerizing activities to spindle poles. J Cell Biol 166, (2004). 10. Heald, R. et al. in Cell Biology: A Laboratory Handbook (ed. Celis, J.) (Academic Press, San Diego, CA, 1998). 11. Tirnauer, J. S., Grego, S., Salmon, E. D. & Mitchison, T. J. EB1-microtubule interactions in Xenopus egg extracts: role of EB1 in microtubule stabilization and mechanisms of targeting to microtubules. Mol Biol Cell 13, (2002).