A RT I C L E S

Reconstitution of the augmin complex provides insights into its architecture and function Kuo-Chiang Hsia1, Elizabeth M. Wilson-Kubalek2, Alejandro Dottore1, Qi Hao1, Kuang-Lei Tsai2, Scott Forth1, Yuta Shimamoto1, Ronald A. Milligan2 and Tarun M. Kapoor1,3 Proper microtubule nucleation during cell division requires augmin, a microtubule-associated hetero-octameric protein complex. In current models, augmin recruits γ-tubulin, through the carboxyl terminus of its hDgt6 subunit to nucleate microtubules within spindles. However, augmin’s biochemical complexity has restricted analysis of its structural organization and function. Here, we reconstitute human augmin and show that it is a Y-shaped complex that can adopt multiple conformations. Further, we find that a dimeric sub-complex retains in vitro microtubule-binding properties of octameric complexes, but not proper metaphase spindle localization. Addition of octameric augmin complexes to Xenopus egg extracts promotes microtubule aster formation, an activity enhanced by Ran–GTP. This activity requires microtubule binding, but not the characterized hDgt6 γ-tubulin-recruitment domain. Tetrameric sub-complexes induce asters, but activity and microtubule bundling within asters are reduced compared with octameric complexes. Together, our findings shed light on augmin’s structural organization and microtubule-binding properties, and define subunits required for its function in organizing microtubule-based structures. Error-free cell division depends on the regulated nucleation of microtubules, polar polymers of α/β-tubulin1–3. There have been important advances in our understanding of how microtubule nucleation at centrosomes is mediated by the recruitment of γ-tubulin and associated proteins (called the γ-tubulin ring complex, or γ-TuRC; refs 4–6). It has also been established that centrosomes are not required for the assembly of meiotic and mitotic spindles7–9. As a result, much attention has been focused on examining two centrosomeindependent microtubule formation pathways. The first is the chromosome-dependent microtubule formation pathway involving Ran GTPase and the Aurora B kinase complex10,11. The second pathway involves augmin, a recently discovered eight-protein complex needed to recruit γ-tubulin to microtubules within the spindle12–14. Several lines of evidence indicate that augmin is needed for proper centrosome-independent microtubule formation in dividing cells. First, the levels of spindle microtubules are reduced when augmin is knocked-down or mutant subunits are present12,13,15. Second, studies in Xenopus egg extracts depleted of augmin reveal that centrosomeindependent meiotic spindle assembly around chromatin-coated beads occurs at reduced rates16. Third, microtubule nucleation along the sides of other microtubules has been directly imaged in egg extracts and involves augmin17. Fourth, electron tomography-based analysis reveals that the number of minus-ends of microtubules distributed within the metaphase spindle are reduced in the absence of

augmin18. Further, these studies suggest that a rod-shaped (29 ± 14 nm long) structure, which could be augmin, crosslinks the minusend of newly formed microtubules to the lattice of pre-existing filaments18. Together, these data have led to a model in which augmin binds the sides of microtubules, recruits γ-tubulin, and promote the centrosome-independent nucleation of a new filament13. A recent study in Drosophila embryos indicates that augmin also contributes to centrosome-dependent astral microtubule assembly19. In metazoans augmin is comprised of eight subunits: Ccdc5 (HAUS1), Cep27 (HAUS2), hDgt3 (HAUS3), C14orf94 (HAUS4), hDgt5 (HAUS5), hDgt6 (HAUS6), UCHL5IP (HAUS7) and Hice1 (HAUS8) (Fig. 1a; refs 12–14). Recombinant Hice1 has been shown to bind microtubules in vitro20. It has been proposed that hDgt6 recruits γ-TuRC to spindle microtubules as a truncated construct cannot rescue the reduction of γ-tubulin signal in mitotic spindles following hDgt6 RNAi13. Furthermore, immunoprecipitation experiments suggest interactions between hDgt6 and NEDD1, a component of γ-TuRC (ref. 13). However, the functions of these different subunits and the overall organization of the augmin complex remain poorly understood. Here, we report the biochemical reconstitution of the augmin complex with recombinant proteins. Analyses of sub-complexes and electron microscopy reveal the subunit organization and overall architecture of this hetero-octameric complex. Comparisons between

1

Laboratory of Chemistry and Cell Biology, Rockefeller University, New York, New York 10065, USA. 2Department of Integrated Structure and Computational Biology, The Scripps Research Institute, La Jolla, California 92037, USA. 3 Correspondence should be addressed to T.M.K. (e-mail: [email protected]) Received 15 August 2013; accepted 16 July 2014; published online 31 August 2014; DOI: 10.1038/ncb3030

852

NATURE CELL BIOLOGY VOLUME 16 | NUMBER 9 | SEPTEMBER 2014 © 2014 Macmillan Publishers Limited. All rights reserved.

A RT I C L E S

1 hDgt6 (HAUS6)

b

410 432

955

NTD 1

368

UCHL5IP (HAUS7) 1

c

Hice1-MTBR

235

14

SP SP SP SP S P S P S P S P S P

7 Vo 6

1

9

12

15

18

Volume (ml)

363

C14orf94 (HAUS4)

8.5 9.5 10.5 11.5 12.5

ml

0.8 0.6 0.4 0.2

1

0 20 40 60 80 100 [Tubulin] (μM)

633

1

603

25 20 15 10

Hice1-MTBR

hDgt3 (HAUS3)

e Absorbance (a.u.)

3 PRC1-MTBD

1 0 –1

220

13.6 ml

50

Vo

–50 6 240

9

12

15

11.5

13.5

40 Vo 0 –20 6

18

15.5

9

h

Tetramer-II

12

15

10.5 11.5 12.5 13.5 14.5

ml

18 ml Mr (K)

Mr (K) 100 75 50 37 25

Hice1 hDgt6 (1–432) UCHL5IP Cep27

100 75 50 37 25

Hexamer

40

90 12.7 ml

Absorbance (a.u.)

Absorbance (a.u.)

13.0 ml

Volume (ml)

Hice1 hDgt6 (1–432)

20 Vo 0 –10

12.5 ml 60 30 Vo 0

6

9

12

15

18

6

9

12

15

18

Volume (ml)

Volume (ml) 10.5 11.5 12.5 13.5 14.5 ml

Hice1 hDgt6 (1–432) His–C14orf94 Ccdc5

80

Volume (ml)

260

Wavelength (nm)

g

Tetramer-I

150

Hice1-MTBR

–3 190 200

f

Hice1• hDgt6Δ (433–955) 250

Absorbance (a.u.)

d

–2

Kd = 9.3 (±1.4) μM

0

Mr (K)

hDgt5 (HAUS5)

θ × 10–4 (deg cm2 dmol–1)

1.0

278

Ccdc5 (HAUS1)

2

BSA Tubulin

Hice1-MTBR

0

Cep27 (HAUS2)

1

[Tubulin] (μM) 0 0.5 1.0 2.5 5 10 20 40 80

10.8 ml

Fraction bound

1 140 Hice1 (HAUS8) MTBR

Absorbance (a.u.)

a

Mr (K) 100 75 50 37 25

10

11

12

13

Hice1 hDgt6 (1–432) His–C14orf94 His–UCHL5IP Ccdc5 Cep27

Figure 1 Hice1·hDgt61(433–955) are components of distinct augmin sub-complexes. (a) Schematics for the 8 subunits of augmin: Hice1 (HAUS8), hDgt6 (HAUS6), UCHL5IP (HAUS7), Cep27 (HAUS2), C14orf94 (HAUS4), Ccdc5 (HAUS1), hDgt5 (HAUS5) and hDgt3 (HAUS3). The microtubule binding region (MTBR; a.a. 1–140) of Hice1 and the N-terminal domain (NTD; a.a. 1–432) of hDgt6 are highlighted (black). (b) Size-exclusion chromatography (Superdex 75) elution profile for Hice1-MTBR. (c) Microtubule co-sedimentation assays to analyse Hice1-MTBR binding. Microtubule-binding constants were determined by fitting to a hyperbola (Kd : 9.3 ± 1.4 µM, N = 3 independent experiments; error bars show s.d.). SDS–PAGE gel image, stained with Coomassie blue, is shown. BSA (final 0.25 mg ml−1 ), used to suppress nonspecific interactions, and tubulin are indicated. (d) Circular dichroism

14

ml Mr (K) 100 75 50 37 25

spectrum of Hice1-MTBR (10 µM, black) and PRC1-MTBD (10 µM, grey, dashed) (25 ◦ C). (e–h) Size-exclusion chromatography (Superose 6) elution profiles for Hice1·hDgt61(433–955) dimer (e), tetramer-I (Hice1·hDgt6 (1–432)· UCHL5IP·Cep27) (f), tetramer-II (Hice1·hDgt6 (1–432)·His–C14orf94·Ccdc5) (g) and hexamer (Hice1·hDgt6 (1–432)· His–UCHL5IP· Cep27·His-C14orf94·Ccdc5) (h). For all chromatography analyses peak fractions (volumes indicted) were analysed by SDS–PAGE (stained with Coomassie blue). Void volume (V o) is also indicated and absorbance (a.u.) at 280 nm is shown. SDS–PAGE resolved Hice1 to be slightly larger than hDgt6 (1–432), even though the calculated molecular mass of Hice1 is lower than that for hDgt6 (1–432). The tagged C14orf94 and UCHL5IP subunits in the hexamer could not be completely resolved by SDS–PAGE.

NATURE CELL BIOLOGY VOLUME 16 | NUMBER 9 | SEPTEMBER 2014 © 2014 Macmillan Publishers Limited. All rights reserved.

853

A RT I C L E S GFP–holo-complex

a

2

30

Absorbance (a.u.)

Absorbance (a.u.)

9.6 ml

4

GFP–octamer[hDgt6Δ(433–955)]

b

6

Vo

0 –2

10.3 ml 20 10

Vo

0 –10

–4 6

9

12

15

18

6

8 9 10 11 12 13

ml Mr (K) 250 150 100 75 50 37 25

GFP–hDgt6 His–hDgt3 and hDgt5 Hice1 and GFP–Cep27 C14orf94 UCHL5IP Ccdc5

c

12

15

18

7 8 9 10 11 12 13

d

Mr (K) Name

Coverage (%)

No. of peptides Unique/Total

hDgt3

62.2

34/43

20

hDgt5

47.2 42.3

24/27 12/18

10

Cep27 hDgt6 (1–432)

39.2 50.5

8/12 21/31

30 10.5 ml

Hice1 Vo

0 6

9

12

15

18

Volume (ml) 9 10 11 12 13 14

His–hDgt3 and hDgt5 GFP–Cep27 hDgt6 (1–432) C14orf94 UCHL5IP Hice1 (141–410) Ccdc5

ml Mr (K) 150 100 75 50 37 25

His–hDgt3 and hDgt5 Hice1 and GFP–Cep27 hDgt6 (1–432) C14orf94 UCHL5IP Ccdc5

GFP–octamer[hDgt6Δ(433–955), Hice1Δ(1–140)]

Absorbance (a.u.)

9

Volume (ml)

Volume (ml)

C14orf94

62.5

29/33

UCHL5IP

40.5

12/13

Ccdc5

78.4

10/33

100 75 50 37 25 20 15

ml Mr (K) 150 100 75 50 37 25

Figure 2 Reconstitution of the GFP-tagged hetero-octameric human augmin complex with recombinant proteins. (a–c) Size-exclusion chromatography (Superose 6) elution profiles for GFP-labelled holocomplex (Hice1·GFP–hDgt6·UCHL5IP·GFP–Cep27·C14orf94·Ccdc5·HishDgt3·hDgt5) (a), octamer[hDgt61(433–955)] (Hice1·hDgt6(1– 432)·UCHL5IP·GFP–Cep27·C14orf94·Ccdc5·His–hDgt3·hDgt5) (b) and octamer[hDgt61(433–955), Hice11(1–140)] (Hice1 (141–410)·hDgt6

(1–432)·UCHL5IP·GFP–Cep27· C14orf94·Ccdc5·His–hDgt3·hDgt5) (c) with peak fractions (volumes indicated) analysed by SDS–PAGE (stained with Coomassie blue). The void volume (V o) is indicated and absorbance (a.u.) at 280 nm is shown. (d) Analysis of the subunit composition of the augmin octamer[hDgt61(433–955)] by mass spectrometry. Identities, percentage of sequence coverage, and number of peptides (unique and total) detected are shown.

direct microtubule binding and metaphase spindle localization of the augmin complexes reveal how different subunits contribute to these properties. We also analyse the activity of the recombinant augmin holo-complex and sub-complexes using a microtubule aster assembly assay. Together, these studies shed light on how augmin contributes to the formation of microtubule-based structures.

we focused on characterizing individual subunits and generating sub-complexes. We first examined Hice1, the subunit that has been shown in vitro to bind microtubules through a region at its amino terminus (hereafter, Hice1-MTBR, for Hice1-microtubule-binding region; amino acids 1–140; ref. 20). We expressed and purified Hice1-MTBR and find that it is monodisperse by size-exclusion chromatography (Fig. 1b). Co-sedimentation assays revealed that Hice1-MTBR binds microtubules (Kd : ∼9 µM, Fig. 1c). Circular dichroism spectrometry indicated that Hice1-MTBR is mostly a random coil in solution (Fig. 1d). As a control, we examined the microtubule-binding domain (MTBD; amino acids 341–466) of PRC1 under similar conditions and found that it was mainly helical in solution (Fig. 1d), consistent with structural data22. These data suggest

RESULTS Hice1·hDgt6 are core components of distinct augmin sub-complexes To biochemically characterize and reconstitute this multi-protein complex we first tried co-expressing all eight subunits in bacteria using polycistronic systems21. As these attempts were unsuccessful,

854

NATURE CELL BIOLOGY VOLUME 16 | NUMBER 9 | SEPTEMBER 2014 © 2014 Macmillan Publishers Limited. All rights reserved.

A RT I C L E S b

GFP–Hice1•hDgt6Δ(433–955)

c

MT

a

GFP GFP

GFP

Augmin

d

1 〈t1〉 = 0.52 ± 0.02 s

X-rhodamine microtubule Glass surface

1-CDF

Time (s)

Augmin

0.1 0.01 0.001 0

1

2

3

4

Dwell time (s)

GFP–tetramer-I

f

h GFP–octamer[hDgt6Δ(433–955)] i

MT

MT

GFP

GFP 1

j

〈t1〉 = 0.59 ± 0.05 s

0.1 Time (s)

1-CDF

Time (s)

g

〈t2〉 = 4.2 ± 0.2 s 0.01

(9.9 ± 0.7%)

1

1-CDF

e

0

GFP–holo-complex

0.1 〈t2〉 = 13 ± 1 s (1.9 ± 0.2%)

0.01 0.001

0.001

k

〈t1〉 = 0.77 ± 0.05 s

5 10 15 20 Dwell time (s)

0

25

10 20 30 40 Dwell time (s)

50

n

l

MT

GFP

MT GFP

1 1-CDF

Time (s)

m

〈t1〉 = 0.30 ± 0.04 s

0.1

GFP–octamer[hDgt6Δ(433–955)]: 60 nM 〈t2〉 = 3.2 ± 0.8 s (16.9 ± 1.0%)

0.01

MT

GFP

0.001 0

5

10

15

20

25

Dwell time (s)

GFP–octamer[hDgt6Δ(433–955), Hice1Δ(1–140)]: 60 nM

Figure 3 TIRF microscopy-based analysis of augmin-microtubule interactions. (a) A schematic of the assay. Microtubules immobilized on the glass surface and GFP-tagged complexes were imaged using TIRF microscopy. (b–m) Single-molecule fluorescence analysis. Image of GMPCPP-stabilized microtubules (X-rhodamine- and biotin-labelled) (top), GFP-tagged complexes (maximum intensity projections, 300 images) and corresponding kymographs (below) for GFP-tagged Hice1·hDgt61(433–955) dimer (b), tetramer-I (GFP-Cep27) (e), octamer[hDgt61(433–955)] (h) and holo-complex (k). Scale bars, horizontal, 2 µm; vertical, 2 s. (c,f,i,l) The region highlighted (rectangle) in each kymograph is also shown in greater detail as a montage. Scale bars, 2 µm. (d,g,j,m) Binding events of individual GFP-tagged augmin complexes on microtubules were tracked to compute the cumulative distribution function (CDF;

right panels). Mean dwell times hti and relative amplitudes (in parentheses) were obtained by fitting to bi-exponential functions (grey curve) for each case, whereas Hice1·hDgt61(433–955) was fitted to a mono-exponential function. Three or more independent experiments were analysed in each condition (n = 5, d,g,m; n = 4, j). Error bars: horizontal, binning intervals; vertical, 1/(total events). (n) GMPCPPstabilized microtubules (X-rhodamine- and biotin-labelled), immobilized on a glass surface, were incubated separately with equal concentrations (60 nM) of GFP-tagged octamer[hDgt61(433–955)] and octamer[hDgt61 (433–955), Hice11(1–140)]. Under identical imaging conditions, octamer[hDgt61(433–955)] decorated microtubules (upper-left panel) but octamer[hDgt61(433–955), Hice11(1–140)] did not target to microtubules (lower-left panel).

that the Hice1 microtubule-binding region is unlikely to have a welldefined secondary structure in solution.

We were unable to purify monodispersed full-length recombinant Hice1 and therefore examined co-expression with

NATURE CELL BIOLOGY VOLUME 16 | NUMBER 9 | SEPTEMBER 2014 © 2014 Macmillan Publishers Limited. All rights reserved.

855

A RT I C L E S a [Tubulin] (μM) His–Dgt3 and Dgt5 Hice1 and GFP–Cep27 hDgt6 (1–432) C14orf94 and UCHL5IP Ccdc5

c [Tubulin] (μM)

b

GFP–octamer[hDgt6Δ(433–955)] 0.0 1.0 2.5 5.0 Mr (K) S P S P S P S P 75 BSA Tubulin

[Tubulin] (μM)

[Tubulin] (μM) His–Dgt3 and Dgt5 GFP–Cep27 hDgt6 (1–432) C14orf94 and UCHL5IP Hice1 (141–410) Ccdc5

BSA Tubulin

Hice1 hDgt6 (1–432) UCHL5IP

25

Cep27

Tetramer-II 0.0 0.1 0.2 0.5 S P S P S P S P

d [Tubulin] (μM)

1 2 3 5 7 S P S PS P S P S P BSA Tubulin

Hice1 hDgt6 (1–432) C14orf94 Ccdc5

e

Tetramer-I 0.0 0.1 0.2 0.5 1 2 3 5 10 S P S P S P S P S P S P S P S P SP

[Tubulin] (μM)

75 BSA Tubulin

Tetramer-I[ΔHice1Δ(1–142)] 1 2 3 5 10 0.0 0.1 0.2 0.5 S P S PS P S P S P S P S P S P S P

hDgt6-NTD UCHL5IP Hice1(143–410) Cep27

25

g

BSA Tubulin

Hice1 hDgt6 (1–432)

f

GFP–octamer[hDgt6Δ(433–955), Hice1Δ(1–140)] 0 1 2.5 5 Mr (K) S P S P S P S P

Hice1•hDgt6Δ(433–955) 0.0 0.1 0.2 0.5 1 2 3 5 10 S P S P S P S P S P S P S P S P S P

BSA Tubulin

1.0

Fraction bound

0.8

0.6

0.4 Octamer Tetramer-I

0.2

Tetramer-II Hice1•hDgt6Δ(433–955)

0 0

2

4 6 [Tubulin] (μM)

8

10

Figure 4 Analyses of microtubule binding by augmin complexes. (a–f) SDS–PAGE analysis of microtubule co-sedimentation assays for GFP–octamer[hDgt61(433–955)] (a), tetramer-I (b), tetramer-II (c), Hice1·hDgt61(433–955) (d), GFP–octamer[hDgt61(433–955), Hice11(1–140)] (e) and tetramer-I[Hice11(1–142)] (f). BSA (final 0.25 mg ml−1 ), used to suppress nonspecific interactions, and tubulin are indicated. (g) Analysis of microtubule binding by GFP–octamer[hDgt61(433– 955)] (diamond), tetramer-I (square), tetramer-II (triangle) and Hice1·hDgt61(433–955) (circle). For the octamer, band intensities of His– Dgt3&Dgt5, C14orf94, UCHL5IP and Ccdc5 from the SDS–PAGE gels were used to determine the average fraction of protein bound. For tetramer-I, band

intensities of hDgt6 (1–432), UCHL5IP and Cep27 were used to determine the average fraction of protein bound. For tetramer-II, band intensities of C14orf94, and Ccdc5 were used to determine the average fraction bound. For the dimer, only the hDgt6 (1–432) band intensity was used to determine the fraction of protein bound. Microtubule-binding constants Kd were determined by fitting to a hyperbola (octamer: ∼1.1 ± 0.2 µM; tetramer-I: 0.6 ± 0.1 µM; tetramer-II: ∼0.5 ± 0.1 µM; dimer: 1.0 ± 0.1 µM). Approximate Kd values are provided for octamer and tetramer-II, as more than 50% of these complexes did not bind microtubules at the highest protein concentration we could test. n = 3 independent experiments were analysed. Error bars show s.d.

the hDgt6 N-terminal domain (amino acids 1–432) (hereafter, Hice1·hDgt61(433–955), as an interaction between these has been shown by yeast two-hybrid results13. We found a soluble and stable hetero-dimer formed by Hice1 and hDgt61(433–955) (Fig. 1e). Light-scattering analysis indicated that these proteins form a heterodimeric complex with a relative molecular mass consistent with that calculated (Mr 91K; Supplementary Fig. 1a,d). We next examined whether other augmin subunits could be co-purified with the Hice1·hDgt61(433–955) hetero-dimer. After trying different combinations, we obtained two different stable tetrameric complexes. One tetrameric complex (hereafter, tetramer-I) was comprised of Hice1, hDgt6 (1–432), UCHL5IP

and Cep27 (Fig. 1f). The other tetramer (hereafter, tetramer-II) was comprised of Hice1, hDgt6 (1–432), C14orf94 and Ccdc5 (Fig. 1g). Light-scattering analysis indicated that these complexes adopt an extended conformation, and the relative molecular masses of tetramer-I and -II were 149K and 159K, respectively (Supplementary Fig. 1b–d), consistent with the subunits associating with equal stoichiometry. Next, to determine whether the proteins represented in tetramer-I and -II assemble into a larger complex, we co-expressed all six proteins in bacteria. We were able to purify a hetero-hexameric complex, with either His-tags on both C14orf94 and UCHL5IP (Fig. 1h), or with only His-tagged C14orf94 (Supplementary Fig. 1e). Tagged C14orf94 and

856

NATURE CELL BIOLOGY VOLUME 16 | NUMBER 9 | SEPTEMBER 2014 © 2014 Macmillan Publishers Limited. All rights reserved.

A RT I C L E S a

Tetramer-I Overview

b

51

c

Mr (K)

MBP–N-Hice1

d

45

hDgt6 (1–432) UCHL5IP CEP27

100 75 50 37 25

81

156

e

f

GFP–octamer[hDgt6Δ(433–955)] Overview

68

72

70

Short arm

Stem Long arm

Figure 5 2D class averages of augmin complexes examined by electron microscopy. (a) Negative-stain electron micrograph of tetramer-I (Hice1·hDgt6 (1–432)·UCHL5IP·Cep27) particles. A field of particles adsorbed onto a glow-discharged carbon grid was stained with 2% (w/v) uranyl acetate and processed for imaging. Scale bar, 50 nm. (b) A representative 2D class average image of tetramer-I particles (N = 51). Scale bar, 10 nm (c) SDS–PAGE analysis (stained with Coomassie blue) of purified tetramer-I with MBP-tagged Hice1 N-terminus. (d) Three representative class averages of the MBP-tagged tetramer-I (MBP–Hice1·hDgt6 (1–432)·UCHL5IP·Cep27) particles (N = 45, 81 and 156). The position of the MBP-tag is indicated (arrow). Scale bar, 10 nm (e) Negative-stain electron micrograph of GFP–octamer[hDgt61(433–955)] particles. A field of GFP–octamer[hDgt61(433–955)] particles adsorbed onto a glow-discharged carbon grid was stained with 2% (w/v) uranyl acetate and processed for imaging. Scale bar, 50 nm. (f) Representative class averages of the GFP–octamer[hDgt61(433–955)] particles (N = 68, 70 and 72). Prominent features in the structure are indicated. Scale bar, 10 nm.

UCHL5IP could not be resolved by SDS–PAGE, and their presence was independently confirmed (Supplementary Fig. 1f). Although we were unable to reconstitute octameric augmin complexes in bacteria, these analyses revealed a connectivity model for the different subunits in augmin (see below).

Biochemical reconstitution of GFP-tagged hetero-octameric human augmin complexes To reconstitute the augmin holo-complex, we used the MultiBac system for poylcistronic gene expression in insect cells23. Remarkably, the entire hetero-octameric complex (hereafter, holo-complex) could be isolated, albeit with low yields. We confirmed the presence of the subunits by western blot analyses (Supplementary Fig. 2) and using SDS–PAGE-based comparisons with another augmin octameric complex characterized by mass spectrometry (see below). The sizeexclusion chromatography elution profile and the intensities of bands resolved by SDS–PAGE suggest that subunits in the holo-complex are present at equal stoichiometry (Fig. 2a). As we could obtain only small amounts of the holo-complex (∼20 µg from 1 l of insect cell culture), we generated an octameric complex with a truncated untagged hDgt6 (1–432). This octameric complex (hereafter, octamer[hDgt61(433–955)]) eluted as a single peak during size-exclusion chromatography (Fig. 2b). The presence of the different subunits in the octamer[hDgt61(433–955)] was validated by mass spectrometry (Fig. 2d). We also purified a related octameric complex lacking the Hice1 N terminus (hereafter, octamer[hDgt61(433–955), Hice11(1–140)]), thereby removing a known microtubule-binding region (Fig. 2c). Together, these data indicate that augmin can be reconstituted by coexpressing eight subunits as recombinant proteins, and the C-terminal residues of hDgt6 and the N-terminal residues of Hice1 are dispensable for forming an octameric complex. Characterizing the microtubule interaction of different purified augmin complexes in vitro We next used total internal reflection fluorescence (TIRF) microscopy to analyse the interaction of GFP-tagged augmin complexes with microtubules. For these analyses we generated GFP-labelled dimer and tetramer-I complexes in addition to the GFP-labelled octameric complexes. Fluorescence intensity measurements of single molecules indicated that Hice1·hDgt61(433–955) dimer, tetramer-I, and octamer[hDgt61(433–955)] particles have a single GFP tag and the holo-complex has two GFP tags, as expected (Supplementary Fig. 3a–f). All GFP-tagged augmin complexes bound to microtubules immobilized on coverslips (Fig. 3a–c,e,f,h,i,k,l and Supplementary Fig. 3l,m). Analyses of time-lapse sequences using kymographs showed that the different augmin complexes diffused in one dimension along microtubules without detectable directional bias (Supplementary Fig. 3g–k). In addition, all of these complexes had similar microtubule-association times (t1/2 less than 1 s, with a subset (∼2–16%) exhibiting longer (∼3–13 s) lifetimes; Fig. 3d,g,j,m and Supplementary Fig. 3n). Although high-intensity spots, most likely due to aggregation under the assay conditions, could be observed for the GFP-tagged holocomplex (Supplementary Fig. 3e), several microtubule-bound particles with the expected intensities were observed and analysed. At equal concentrations, GFP-tagged octamer[hDgt61(433–955)] bound microtubules, whereas octamer[hDgt61(433–955), Hice11(1–140)] did not (Fig. 3n). Together, these analyses reveal that the interaction of augmin with microtubules depends on Hice1-MTBR, results in onedimensional (1D) diffusion along the filaments and is not biased to ends of stabilized filaments.

NATURE CELL BIOLOGY VOLUME 16 | NUMBER 9 | SEPTEMBER 2014 © 2014 Macmillan Publishers Limited. All rights reserved.

857

A RT I C L E S i

58

150 100 50 0 10

GFP intensity

300 200 100 0 20 30 40 Distance (μm)

50

60 50 40 30 20 10 0

400 300 200 100 0 0

10

20 30 40 50 Distance (μm)

Rhodamine intensity

GFP intensity

h

10

Rhodamine intensity

400

60 50 40 30 20 10 0 0

g GFP-tagged octamer[hDgt6Δ(433–955), Hice1Δ(1–140)]

50 55

0

[h

f

e GFP-tagged tetramer-I

20 30 40 Distance (μm)

1

Dg t6 Δ

0

Rhodamine intensity

60 50 40 30 20 10 0

Oc 33 tam –9 er 55 )]

20 30 40 50 Distance (μm)

d GFP intensity

c GFP-tagged tetramer-II

10

(4

0 0

2

m er -II

100

ra

200

MT GFP

m er -I

300

3

Te t

400

Pole/midzone intensity

500

60 50 40 30 20 10 0

ra

b Overlay GFP intensity

MT

Rhodamine intensity

GFP

Te t

a GFP-tagged octamer[hDgt6Δ(433–955)]

56

Figure 6 Analysis of augmin’s localization in metaphase spindles assembled in Xenopus egg extracts. (a–h) Fluorescence microscopy-based analysis. Recombinant GFP-tagged octamer[hDgt61(433–955)] (a), tetramer-II (c), tetramer-I (e) and octamer[hDgt61(433–955), Hice11(1–140)] (g) were added (15 nM) and imaged without fixation (left panels); tubulin (X-rhodamine-labelled, middle panels); and overlays (right panels; tubulin, red; GFP, green; and DNA, blue). (b,d,f,h) Corresponding linescans, along the long axis of the spindles, for the GFP (green) and X-rhodamine (red) signals. (i) Analysis of augmin (green) and

microtubule (red) levels in spindles. Ratios of average fluorescence signal at spindle poles versus signal in the middle of the spindle for tetramer-I, tetramer-II and octamer[hDgt61(433–955)] are shown. Tetramer-I: 0.99 ± 0.18 s.d. (N = 13 spindles), microtubule, 1.06 ± 0.14 s.d. (N = 13 spindles); Tetramer-II: 0.91 ± 0.22 s.d. (N = 14 spindles), microtubule, 0.9 ± 0.2 s.d. (N = 14 spindles); octamer[hDgt61(433–955)], 1.91 ± 0.53 s.d. (N = 14 spindles), microtubule, 1.11 ± 0.21 s.d. (N = 14 spindles). S.d. was determined from data pooled from 3 or more independent experiments.

We next measured the microtubule-binding affinities of the Hice1·hDgt61(433–955) dimer, tetramer-I, tetramer-II and GFP– octamer[hDgt61(433–955)] using co-sedimentation assays. We found that these multi-protein complexes bound microtubules with more than tenfold higher affinity (Kd : 0.5–1.1 µM) than Hice1-MTBR alone (Figs 1c and 4). GFP–octamer[hDgt61(433–955), Hice11(1–140)] and tetramer-I[Hice11(1-142)] did not bind microtubules under these conditions at concentrations up to 10 µM (Fig. 4e,f). We also visualized microtubule binding of the dimeric and tetrameric complexes by negative-stain electron microscopy (Supplementary Fig. 4a,b). Together, these data indicate that Hice1MTBR alone is not sufficient to achieve microtubule-binding affinities comparable to the multi-protein complexes, but this region is needed for strong augmin–microtubule interactions.

Single-particle electron microscopy analysis of augmin complexes To examine the overall organization of the augmin complex, we used negative-stain single-particle electron microscopy. Images of tetramer-I (Hice1·hDgt6 (1–432)·UCHL5IP·Cep27) and tetramer-II (Hice1·hDgt6 (1–432)·His-C14orf94·Ccdc5) complexes seemed to be similar (Fig. 5a and Supplementary Fig. 4e,f), and therefore we carried out standard single-particle analysis only of tetramer-I. We generated 2D class averages of tetramer-I and found it to have a ‘drumstick’like structure with a length of 16–20 nm (Fig. 5b). To localize the position of the microtubule-binding Hice1 subunit, we generated two tetramer-I complexes containing Hice1 tagged with a maltose-binding protein (MBP) at either the N or C terminus. Coexpression, followed by multi-step chromatography, led to homogeneous tetrameric protein

858

NATURE CELL BIOLOGY VOLUME 16 | NUMBER 9 | SEPTEMBER 2014 © 2014 Macmillan Publishers Limited. All rights reserved.

A RT I C L E S a

Aster number

1,000 800

A Buffer B GFP–octamer[hDgt6Δ(433–955)] Ran+ C GFP–holo-complex 6 nM augmin complexes

b

GFP-tagged holo-complex (15 nM) MT GFP

600 400 A BC

A B C

A B C

200

Ran+

0 15'

d

GFP-tagged octamer[hDgt6Δ(433–955)] (15 nM) MT

GFP

Ran+

Aster number

3,000 2,500

f

Ran+ 60 nM augmin complexes 10'

2,000 1,500 500

A

B

C

D

E

3,000 Aster number

e

2,500

Rhodamine signal GFP signal

6 4 2 0

A B A GFP–holo-complex B GFP–octamer[hDgt6Δ(433–955)]

g

Ran+ 10'

2,000 1,500 500

AB

AB

A B

AB

Aster number

10'

Intensity ratio (1 μm versus 5 μm radius)

5'

c

0 0 7.5 15 30 60 (nM) A GFP–octamer[hDgt6Δ(433–955), C Tetramer-I A Tetramer-II Hice1Δ(1–140)] D Tetramer-II B GFP–octamer[hDgt6Δ(433–955)] E GFP–octamer[hDgt6Δ(433–955)] B Hice1•hDgt6Δ(433–955)

3,000 2,500

Ran – 60 nM augmin complexes 10'

2,000 1,500 500

A

B

0 A Tetramer-II B GFP–octamer[hDgt6Δ(433–955)]

Figure 7 Analysis of augmin-induced microtubule aster formation. Augmin complexes and Ran(Q69L) (15 µM) were added to extracts, incubated for 10 min (or as noted), fixed and processed to determine the number of asters formed. (a) Aster assembly in the presence of buffer (A, light grey), GFP-tagged octamer[hDgt61 (433–955)] (6 nM) (B, grey), and holo-complex (6 nM) (C, black). Mean and s.d. were calculated from 3 independent experiments with separate extract preparations. For each experiment, data were pooled from two measurements. (b,c) Asters formed in the presence of Ran(Q69L) and GFP-tagged holo-complex (b) or octamer[hDgt61(433–955)] at 15 nM (c). Tubulin (X-rhodamine-labelled), left panels; GFP fluorescence, right panels, are shown. Scale bar, 10 µm. (d) Analysis of microtubule (rhodamine signal) and augmin (GFP signal) levels in asters. Ratios of average fluorescence at 1 µm versus 5 µm radius for holo-complex (A) and octamer[hDgt61(433–955)] (B) induced asters are shown. Mean and s.d. were calculated from: Holo-complex, N = 23 asters; octamer, N = 22 asters; analysed asters were from 3 separate extract preparations.

(e) Aster formation in the presence of GFP-tagged octamer[hDgt61 (433–955), Hice11(1–140)] (A), Hice1·hDgt61(433–955) dimer (B), tetramer-I (C), tetramer-II (D), and GFP-tagged octamer[hDgt61 (433–955)] (E) at 60 nM. Mean and s.d. were calculated from either n = 3 (dimer, tetramer-I and tetramer-II) or n = 4 (octamer[hDgt61 (433–955), Hice11(1–140)] and octamer[hDgt61(433–955)) independent experiments with separate extract preparations. For each experiment, data were pooled from two measurements. (f) Dose-dependent analysis of aster formation in the presence of tetramer-II (A) and GFP-tagged octamer[hDgt61(433–955)] (B). Mean and s.d. were calculated from either n = 3 (tetramer-II) or n = 5 (octamer[hDgt61(433–955)]) independent experiments with separate extract preparations. For each experiment, data were pooled from two measurements. (g) Aster assembly in the absence of Ran(Q69L). Aster number for tetramer-II (A, 60 nM) and GFP-tagged octamer[hDgt61(433–955)] (B, 60 nM). Mean and s.d. were calculated from n = 3 independent experiments with separate extract preparations. For each experiment, data were pooled from two measurements.

complexes (Fig. 5c and Supplementary Fig. 4c). 2D averages of these N- and C-terminally labelled tetramer-I complexes revealed additional density at similar positions at the narrow end of the ‘drumstick’ structure (Fig. 5d and Supplementary Fig. 4d). Together, these data indicate that Hice1 localizes at one end of tetramer-I formed by Hice1, hDgt6 (1–432), UCHL5IP and Cep27. We next examined the structure of the purified recombinant hetero-octameric complex lacking the C-terminal portion of hDgt6 (octamer[hDgt61(433–955)]). In this complex, Cep27 is tagged with GFP. Analysis of ∼8,000 augmin particle images revealed classes exhibiting general structural features (representative class averages shown in Fig. 5e,f). The complex is 38–44 nm long with a splayed

Y-shape at one end. The angle between the long and short arms of the Y is variable, underscoring the flexibility of the splayed end of the complex seen in the raw images. Attempts to localize tetramer-I in the octamer averages did not yield an unambiguous result. Overall, our findings indicate that the augmin octamer adopts a ∼40 nm extended Y-shaped complex, the splayed end of which can access a large number of conformations. Analysing augmin targeting to metaphase spindles To examine the contributions of the different subunits to augmin’s localization in metaphase spindles we used Xenopus egg extracts24. Using an antibody against Xenopus Ccdc5 (Supplementary Fig. 5a)

NATURE CELL BIOLOGY VOLUME 16 | NUMBER 9 | SEPTEMBER 2014 © 2014 Macmillan Publishers Limited. All rights reserved.

859

A RT I C L E S c

b GFP-tagged octamer

GFP-tagged tetramer-II (60 nM)

MT

Ran+ 45º

45º MT

le

An g

le

(º)

(º)

0º 0º

0º Radial distance (μm)

Ran+

60 nM

0.6

f CV for microtubule intensity

Normalized intensity (a.u.)

Ran+

Tetramer-II Octamer [hDgt6Δ(433–955)]

0.2

90

180

270

360

g

60 nM

A B

A Tetramer-II B Octamer A B [hDgt6Δ(433–955)]

4 8 Radius (μm)

15 nM

i Ran alone Holo-complex Octamer [hDgt6Δ(433–955)]

0

90

180

270

0.6 CV for microtubule intensity

Normalized intensity (a.u.)

0º Radial distance (μm)

0

0

j

15 nM A Ran alone B Holo-complex C Octamer [hDgt6Δ(433–955)]

0.4

0.2

ABC

ABC

0

360

4 8 Radius (μm)

Angle (º)

k

60 nM 6 5 4 3 2 1 0

6 5 4 3 2 1 0

Tetramer-II Octamer [hDgt6Δ(433–955)]

0

5

10 15 Radius (μm)

20

15 nM

Ran alone Holo-complex Octamer [hDgt6Δ(433–955)]

0

5

10 15 Radius (μm)

20

Direct binding partners Cep27

Tetramer-I

hDgt5

Microtubule binding

UCHL5IP hDgt3

Hice1 hDgt6 (1−432)

Hexamer Ccdc5

Tetramer-II

Figure 8 Morphological analysis of asters induced by GFP-tagged augmin complexes and schematic for augmin organization and function. Augmin complexes and Ran(Q69L)(15 µM) were added to extracts, and aliquots were fixed and processed for imaging. (a,b) Two examples of microtubule asters induced by GFP-tagged tetramer-II (a) and GFP-tagged octamer[hDgt61(433–955)] (b) at 60 nM. Defined angular coordinates are indicated (blue and yellow lines). (c,d) Representative images after polar transformation of asters induced by tetramer-II (c) and octamer[hDgt61(433–955)] (d). The microtubule fluorescence intensity along the radial direction (x axis, where radius = 0 corresponds to the centre of the aster), is shown for all angles (y axis, from 0 to 360 degrees). Labelled radial profiles (blue and yellow lines) correspond to those shown in a and b. (e) Normalized angular intensity values at radius = 8 µm in asters induced by teramer-II and octamer[hDgt61(433–955)]. (f) Coefficient of variation (CV, equal to the standard deviation divided by the mean of the intensity) of tetramer-II- and octamer[hDgt61(433–955)]-induced asters at 4 and 8 µm radii. CV was calculated for half-circle (180 degree) regions containing the highest detected microtubule signal. (Tetramer-II, N = 27 asters; octamer,

Microtubule aster formation

Hice•1hDgt6Δ(433–955) × Tetramer-I × √ Tetramer-II √ √ Octamer √ √ Octamer (MTBR deletion) × × √

C14orf94

860

45º

0.4

Angle (º)

h

45º

Normalized intensity (a.u.)

An g

Normalized intensity (a.u.)

MT

Angle (º)

Ran+

Octamer[hDgt6Δ(433–955)]

Tetramer-II

MT

e

d

[hDgt6Δ(433–955)] (60 nM)

Angle (º)

a

Octamer

N = 37 asters; s.d. was determined from data pooled from 3 independent experiments.) (g) Normalized intensity averaged across all angles plotted as a function of radial distance from the aster centre. (h) Normalized angular intensity values at radius = 8 µm in asters induced by Ran-alone, holocomplex (15 nM) and octamer[hDgt61(433–955)] (15 nM). (i) Coefficient of variation calculated from angular intensity values (Ran(Q69L) alone, N = 28 asters; holo-complex, N = 31 asters, octamer[hDgt61(433–955)], N = 35 asters; s.d. was determined from data pooled from 3 or more independent experiments.) (j) Normalized intensity averaged across all angles plotted as a function of radial distance from the aster centre. (k) Schematic for augmin’s subunit organization and function. Direct interaction between subunits is indicated as a red bar. Hice1 and hDgt6 (1–432) form heterodimers. Distinct tetrameric complexes have a common Hice1·hDgt61 (433–955) core. The six proteins in these two tetramers form a stable hexamer. Together with hDgt5 and hDgt3, these proteins form an octameric complex. The table provides a summary of the results for microtubule interaction in vitro and microtubule aster formation induced by augmin complexes.

NATURE CELL BIOLOGY VOLUME 16 | NUMBER 9 | SEPTEMBER 2014 © 2014 Macmillan Publishers Limited. All rights reserved.

A RT I C L E S we find that this subunit localizes along spindle microtubules and accumulates at spindle poles (Supplementary Fig. 5b), consistent with the reported augmin localization in Drosophila meiotic spindles15. Next, we analysed the localization of recombinant GFP-tagged octamer[hDgt61(433–955)], tetramer-II and tetramer-I (15 nM). Although all of these complexes targeted to spindle microtubules (Fig. 6a–f and Supplementary Fig. 5c), octamer[hDgt61(433–955)] had a twofold higher signal at spindle poles (Fig. 6i). The GFPtagged Hice1·hDgt61(433–955) dimer (15 nM) localized to the spindle, but also targeted to chromosomes, possibly owing to nonspecific interactions (Supplementary Fig. 5d,e). These results suggest endogenous augmin and octamer[hDgt61(433–955)], but not the other sub-complexes, have a similar protein localization in Xenopus metaphase spindles. As the recombinant GFP-tagged holocomplex could be obtained only at concentrations that were ∼10-fold lower than other complexes, we were unable to achieve sufficient levels in extracts for detection of its spindle association without perturbing organization owing to extract dilution (Supplementary Fig. 5i). We next examined GFP-tagged octamer[hDgt61(433–955), Hice11(1–140)] (15 nM) localization in spindles. The GFP intensity in the middle of the spindle is similar to that of recombinant GFP alone (Fig. 6h and Supplementary Fig. 5f,g). Interestingly, a weak GFP signal could be consistently detected at spindle poles (Fig. 6g,h and Supplementary Fig. 5h), suggesting the presence of low-affinity microtubule-independent interactions between pole-associated proteins and augmin. Together, these results suggest that augmin sub-complexes with similar microtubule-binding properties in vitro target to metaphase spindles. However, accumulation at spindle poles requires subunits in addition to those present in tetramer-I or -II alone. Recombinant augmin promotes microtubule aster formation in Xenopus egg extracts To test the function of the recombinant augmin complexes, we used a microtubule aster formation assay in Xenopus egg extracts16,17. It has recently been shown that augmin depletion delays microtubule aster formation in the presence of Ran(Q69L) (ref. 16), a mutant form of Ran locked in the GTP-bound state. We find that addition of augmin holo-complex (6 nM) leads to substantially more asters, formed within minutes, in the presence of Ran(Q69L) (15 µM), compared with controls (buffer; Fig. 7a and Supplementary Figs 6b and 7a). Aster formation was also promoted by octamer[hDgt61(433–955)] (6 nM), (Fig. 7a and Supplementary Figs 6a and 7a), albeit with reduced efficiency (15 min, ∼32% lower number) compared with the holocomplex (Fig. 7a). We examined the localization of the GFP-tagged augmin holo-complex and octamer[hDgt61(433–955)] in microtubule asters at 15 nM, the highest concentration of the holo-complex we could reliably achieve without loss of aster stability. Both holo-complex and octamer[hDgt61(433–955)] associated with microtubules in the asters (Fig. 7b–d). We were unable to sufficiently deplete endogenous augmin from the egg extracts using our xCcdc5 antibody (Supplementary Fig. 6c), and therefore could not properly examine whether the recruitment of γ-tubulin in the asters generated by octamer[hDgt61(433–955)] was reduced compared with that in asters induced by the holo-complex.

We next analysed aster formation by the recombinant augmin sub-complexes and the octameric complex lacking Hice1-MTBR. We estimated that the concentration of Ccdc5 in Xenopus extracts is 60 nM (Supplementary Fig. 5a). We were unable to obtain these high concentrations of the holo-complex without diluting the extract and disrupting aster assembly, but could with octamer[hDgt61 (433–955)]. Within 10 min, this octameric complex (60 nM) induced aster assembly. In contrast, essentially no asters formed in the presence of octamer[hDgt61(433–955), 1Hice1(1–140)] or Hice1·hDgt61(433–955) dimer (Fig. 7e and Supplementary Fig. 7b, 60 nM). Notably, whereas some asters were induced by tetramer-I, the activity of tetramer-II was comparable to that of octamer[hDgt61(433–955)] in this assay (Fig. 7e (60 nM)). The localization of tetramer-II in asters was similar to that of the holocomplex and octamer[hDgt61(433–955)] (Supplementary Fig. 6d,e (15 nM)). Addition of tetramer-II induced aster formation in a dose-dependent manner, with efficiency only slightly lower than that of octamer[hDgt61(433–955)] (Fig. 7f and Supplementary Fig. 7c). Interestingly, octamer[hDgt61(433–955)] and tetramer-II promoted microtubule aster formation even in the absence of Ran(Q69L), but overall efficiency was lower (Fig. 7g and Supplementary Fig. 7d). Together, our data indicate that recombinant augmin promotes microtubule aster formation in Xenopus egg extracts and this Ran GTPase-regulated activity depends on Hice1-MTBR, but not the hDgt6 C terminus. Morphological differences between microtubule asters promoted by octameric and tetrameric augmin complexes We next compared the morphology of the asters induced by the recombinant GFP-tagged tetramer-II and octamer[hDgt61(433– 955)] in the presence of Ran(Q69L). We used recombinant complexes at concentrations similar to that of the native protein (60 nM). Tetramer-II-induced asters were symmetric with a more uniform microtubule density, whereas octamer[hDgt61(433–955)]-induced asters were more asymmetric and exhibited more regions of dense microtubule ‘bundles’ adjacent to regions void of microtubules (Fig. 8a,b and Supplementary Fig. 6l,m). To quantitatively compare the morphology of the asters induced by these complexes we first performed a coordinate transformation on images (bottom panel, Fig. 8a,b). Here, the fluorescence intensity along the radial direction is plotted for all angles (Fig. 8c,d). The angular intensity values at a specified radius were then determined (Fig. 8e, radius = 8 µm), and the coefficient of variation was calculated (Fig. 8f). For asters induced with tetramer-II, and at radii near the centre of the aster, the coefficient of variation was relatively small. In contrast, for octamer[hDgt61(433–955)]-induced asters, the coefficient of variation was much larger. These fluctuations in microtubule intensity increased with distance from the aster centre (Fig. 8f). Further, determined under a comparable total intensity within the asters, the normalized intensity of all microtubule signal reveals that octamer[hDgt61(433–955)]-induced asters have a sharper reduction of microtubule density along the radial direction compared with that of tetramer-II-induced asters (Fig. 8g and Supplementary Fig. 7e). We next analysed asters induced by Ran(Q69L) alone to those generated by Ran(Q69L) and holo-complex or octamer[hDgt61 (433–955)], at the highest holo-complex concentration we could

NATURE CELL BIOLOGY VOLUME 16 | NUMBER 9 | SEPTEMBER 2014 © 2014 Macmillan Publishers Limited. All rights reserved.

861

A RT I C L E S reliably achieve in the assay (15 nM, Fig. 7b,c and Supplementary Fig. 6i–k). Our analysis revealed that the coefficients of variation at different radii and the normalized intensity measured as a function of radius were similar for each of these conditions. (Fig. 8h–j and Supplementary Figs 6f–h and 7f). Together, these data show that different augmin complexes can generate asters with distinct morphological features, and that the addition of holo-complex and octamer[hDgt61(433–955)] induce structures with morphologies similar to the Ran-induced asters. DISCUSSION In summary, the biochemical reconstitution of the hetero-octameric augmin complex has revealed its overall architecture and allowed functional tests. Our electron microscopy analyses indicate that augmin forms an elongated Y-shaped structure. However, owing to the resolution of this structure and conformational flexibility of the holo-complex, we are unable to position sub-complexes within the holo-complex with high confidence. On the basis of our analyses thus far, we propose that Hice1·hDgt61(433–955) forms a core scaffold within augmin. Within the holo-complex, C14orf94 and Ccdc5 are likely to be directly connected (Supplementary Fig. 1g), and together interact with Hice1·hDgt61(433–955) independent of other subunits. At present, we are unable to position hDgt3 and hDgt5 subunits within augmin. Together, our findings provide valuable constraints on how the different subunits interact within the augmin holocomplex (Fig. 8k). Our findings also help dissect the contributions of the different augmin subunits to its microtubule-organizing function. Octamer[hDgt61(433–955)], Hice1·hDgt61(433–955) dimer and tetramer-I have similar microtubule-binding affinities, filamentassociation lifetimes and distributions along stabilized microtubules. As these properties do not differ significantly for the octameric and dimeric complexes, and Hice1·hDgt61(433–955) is present in tetramer-II, we infer that these properties are probably comparable for all of these complexes. Further, our data indicate that Hice1-MTBR mediates direct microtubule binding in the augmin complex in vitro and in spindles. Although different sub-complexes can bind spindle microtubules, enrichment of these proteins at spindle poles, similar to that of endogenous augmin, requires the additional subunits present in the octameric complex. Our data also indicate that this localization does not depend on the hDgt6 C terminus that has been shown to interact with γ-tubulin13. Current models predict that augmin should localize at the branch point between the existing ‘mother’ filament and the new ‘daughter’ filament13. However, such localization has not been shown and our own efforts have not been successful thus far. This could be due to limited signal over background in our imaging assays or augmin not preferentially localizing to the filament branch points relative to the sides of the filaments. We find that microtubule asters with similar size and overall microtubule levels are promoted by tetramer-II and octameric complexes, including one that lacks the hDgt6 C terminus. The main morphological difference between asters promoted by tetramer-II and octameric complexes is the amount of microtubule bundling. This difference could result from variations in the branching angles between the ‘mother’ and ‘daughter’ filaments, or the efficiency in recruiting γ-tubulin. This aster-promoting activity is not observed for

862

other sub-complexes that have similar microtubule-binding properties in vitro. C14orf94 and Ccdc5, the two subunits in tetramer-II not present in tetramer-I or the dimeric complexes, are unlikely to substantially modulate the microtubule interaction in vitro. These two subunits may instead mediate regulation of aster assembly by Ran– GTP, or contribute to γ-tubulin recruitment independent of hDgt6. These data, along with our observation that this aster-promoting activity requires Hice1-MTBR, raise the possibility that augmin functions in two ways, by recruiting γ-tubulin and through directly stabilizing microtubules. In vitro assays with dynamic microtubules and the augmin complexes we have reconstituted will help test this model and reveal how this hetero-octameric complex contributes to microtubule-formation pathways needed to ensure successful cell division.  METHODS Methods and any associated references are available in the online version of the paper. Note: Supplementary Information is available in the online version of the paper ACKNOWLEDGEMENTS We thank G. Goshima (Nagoya University, Japan) for the gift of UCHL5IP antibody; and L. Pelletier (University of Toronto, Canada) for the antibodies against C14orf94 and Cep27. The SEC-LS/UV/RI instrumentation used for light-scattering analysis was supported by a National Institutes of Health (NIH) award (1S10RR023748-01). TEM studies were conducted at the National Resource for Automated Molecular Microscopy, which is supported by the National Institute of General Medical Sciences (9 P41 GM103310). K-C.H. was supported by the Kimberly LawrenceNetter Cancer Research Discovery Fund at The Rockefeller University and is supported by a Special Fellow Award from The Leukemia and Lymphoma Society. S.F. acknowledges postdoctoral support from an NIH National Research Service Award Fellowship (F32GM099380). R.A.M. acknowledges support from the NIH (GM-052468). T.M.K. acknowledges support from the NIH (GM-65933). AUTHOR CONTRIBUTIONS K-C.H. carried out experiments in Figs 1–4 and assembled all figures. Q.H. helped purify and characterize proteins. Acquisition and interpretation of the electron microscopy and image analysis data shown in Fig. 5 were carried out by E.M.W-K., K-L.T. and R.A.M. Single-molecule assays in Fig. 3 were carried out by A.D. K-C.H. and Y.S. carried out experiments involving Xenopus egg extracts (Figs 6–8). S.F. quantitatively analysed aster morphology (Fig. 8). All authors helped write the manuscript. T.M.K. directed the project and helped design experiments and prepare the manuscript. COMPETING FINANCIAL INTERESTS The authors declare no competing financial interests. Published online at www.nature.com/doifinder/10.1038/ncb3030 Reprints and permissions information is available online at www.nature.com/reprints 1. Walczak, C. E. & Heald, R. Mechanisms of mitotic spindle assembly and function. Int. Rev. Cytol. 265, 111–158 (2008). 2. Meunier, S. & Vernos, I. Microtubule assembly during mitosis—from distinct origins to distinct functions? J. Cell Sci. 15, 2805–2814 (2012). 3. Teixidó-Travesa, N., Roig, J. & Lüders, J. The where, when and how of microtubule nucleation—one ring to rule them all. J. Cell Sci. 125, 4445–4456 (2012). 4. Lüders, J. & Stearns, T. Microtubule-organizing centres: a re-evaluation. Nat. Rev. Mol. Cell Biol. 8, 161–167 (2007). 5. Bettencourt-Dias, M. & Glover, D. M. Centrosome biogenesis and function: centrosomics brings new understanding. Nat. Rev. Mol. Cell Biol. 8, 451–463 (2007). 6. Kollman, J. M., Merdes, A., Mourey, L. & Agard, D. A. Microtubule nucleation by γ-tubulin complexes. Nat. Rev. Mol. Cell Biol. 12, 709–721 (2011). 7. Wadsworth, P. & Khodjakov, A. E pluribus unum: towards a universal mechanism for spindle assembly. Trends Cell Biol. 14, 413–419 (2004). 8. Walczak, C. E., Cai, S. & Khodjakov, A. Mechanisms of chromosome behaviour during mitosis. Nat. Rev. Mol. Cell Biol. 11, 91–102 (2010). 9. Dumont, J. & Desai, A. Acentrosomal spindle assembly and chromosome segregation during oocyte meiosis. Trends Cell Biol. 22, 241–249 (2012). 10. Karsenti, E. & Vernos, I. The mitotic spindle: a self-made machine. Science 294, 543–547 (2001).

NATURE CELL BIOLOGY VOLUME 16 | NUMBER 9 | SEPTEMBER 2014 © 2014 Macmillan Publishers Limited. All rights reserved.

A RT I C L E S 11. Kelly, A. E. & Funabiki, H. Correcting aberrant kinetochore microtubule attachments: an Aurora B-centric view. Curr. Opin. Cell Biol. 21, 51–58 (2009). 12. Goshima, G., Mayer, M., Zhang, N., Stuurman, N. & Vale, R. D. Augmin: a protein complex required for centrosome-independent microtubule generation within the spindle. J. Cell Biol. 181, 421–429 (2008). 13. Uehara, R. et al. The augmin complex plays a critical role in spindle microtubule generation for mitotic progression and cytokinesis in human cells. Proc. Natl Acad. Sci. USA 106, 6998–7003 (2009). 14. Lawo, S. et al. HAUS, the 8-subunit human Augmin complex, regulates centrosome and spindle integrity. Curr. Biol. 19, 816–826 (2009). 15. Colombié, N., Głuszek, A. A., Meireles, A. M. & Ohkura, H. Meiosis-specific stable binding of augmin to acentrosomal spindle poles promotes biased microtubule assembly in oocytes. PLoS Genet. 9, e1003562 (2013). 16. Petry, S., Pugieux, C., Nédélec, F. J. & Vale, R. D. Augmin promotes meiotic spindle formation and bipolarity in Xenopus egg extracts. Proc. Natl Acad. Sci. USA 108, 14473–14478 (2011). 17. Petry, S., Groen, A. C., Ishihara, K., Mitchison, T. J. & Vale, R. D. Branching microtubule nucleation in Xenopus egg extracts mediated by augmin and TPX2. Cell 152, 768–777 (2013).

18. Kamasaki, T. et al. Augmin-dependent microtubule nucleation at microtubule walls in the spindle. J. Cell Biol. 202, 25–33 (2013). 19. Hayward, D., Metz, J., Pellacani, C. & Wakefield, J. G. Synergy between multiple microtubule-generating pathways confers robustness to centrosome-driven mitotic spindle formation. Dev. Cell 28, 81–93 (2014). 20. Wu, G. et al. Hice1, a novel microtubule-associated protein required for maintenance of spindle integrity and chromosomal stability in human cells. Mol. Cell. Biol. 28, 3652–3662 (2008). 21. Tan, S., Kern, R. C. & Selleck, W. The pST44 polycistronic expression system for producing protein complexes in Escherichia coli. Protein Expr. Purif. 40, 385–95 (2005). 22. Subramanian, R. et al. Insights into antiparallel microtubule crosslinking by PRC1, a conserved nonmotor microtubule binding protein. Cell 142, 433–443 (2010). 23. Trowitzsch, S., Bieniossek, C., Nie, Y., Garzoni, F. & Berger, I. New baculovirus expression tools for recombinant protein complex production. J. Struct. Biol. 172, 45–54 (2010). 24. 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, 385–412 (1999).

NATURE CELL BIOLOGY VOLUME 16 | NUMBER 9 | SEPTEMBER 2014 © 2014 Macmillan Publishers Limited. All rights reserved.

863

METHODS

DOI: 10.1038/ncb3030

METHODS Construction of augmin subunit genes in Escherichia coli expression plasmids. DNA fragments of the Hice1 microtubule-binding region (MTBR; amino acids (a.a.) 1–140) were amplified by polymerase chain reaction (PCR) and cloned into NdeI/XhoI sites of a modified pET28a (Novagen) vector with a PreScission protease site directly after the N-terminal 6-His tag. The amplified PCR fragments of hDgt6 (1–432) and Hice1 (a.a. 1–410) were cloned into a modified pGEX-6P vector (GE Healthcare) that contains two multiple cloning sites (MCSs) and an ampicillin resistance marker. hDgt6 (1–432) and Hice1 were inserted into the vector through BamHI/SalI and BglII/XhoI sites, respectively. hDgt6 (1–432) was fused to a cleavable GST and Hice1 was untagged. Hice1 (a.a. 143–410) was cloned into identical restriction sites in pGEX-6P as the full-length Hice1. Full-length UCHL5IP (a.a. 1–368) and Cep27 (a.a. 1–235) were cloned into a modified bi-cistronic pCDFDuet-1 vector (Novagen) that contains a streptomycin resistance marker and a PreScission protease site directly after the N-terminal 6-His tag. UCHL5IP was inserted into MCS1 through BamHI/NotI sites, and Cep27 was cloned into MCS2 through NdeI/XhoI sites. UCHL5IP was fused with a cleavable N-terminal His tag and Cep27 was expressed as an untagged protein. Full-length C14orf94 (a.a. 1–363) and Ccdc5 (a.a. 1–278) were cloned into another modified bi-cistronic pRSFDuet-1 vector (kanamycin resistance) (Novagen) with either a cleavable or an un-cleavable N-terminal 6-His tag in C14orf94, and Ccdc5 was expressed as an untagged protein. C14orf94 was inserted into MCS1 through BamHI/NotI sites, and Ccdc5 was cloned into MCS2 through NdeI/XhoI sites. To generate the N-terminal MBP-tagged Hice1 construct, DNA fragments of MBP amplified by PCR were inserted in-frame through a BglII site at the 50 end of Hice1 in pGEX-6P. For C-terminal MBP-tagged Hice1, the stop codon in pGEX-6P-Hice1 was removed by site-directed mutagenesis and PCR-amplified MBP DNA fragments including its stop codon were fused in-frame through a XhoI site at the 30 end of the Hice1 gene. Cloning of GFP-tagged hDgt6 (1–432) and Cep27 constructs used the same strategy as was employed to generate MBP-tagged constructs. For expression of C14orf94 (a.a. 1–188) and full-length Ccdc5, DNA fragments containing individual genes were cloned into the modified pGEX-6P vector through BamHI/SalI and BglII/XhoI sites, respectively. Ccdc5 is fused to a cleavable GST and C14orf94 (a.a. 1–188) is untagged.

Purification of recombinant augmin sub-complexes from bacteria. For the expression of Hice1-MTBR, E. coli Rosetta cells (Stratagene) were employed and protein expression was carried out in LB medium and induced by the addition of 0.5 mM IPTG at 18 ◦ C for 16 h. Cells were collected by centrifugation and resuspended in a buffer containing 50 mM potassium phosphate, pH 7.4, 150 mM NaCl, 5 mM β-mercaptoethanol and complete EDTA-free protease inhibitors (Roche). The cells were lysed with a cell disrupter (Avestin) and the lysate was centrifuged for 30 min at 40, 000g . The supernatant was then applied to a His-Select nickel column (Sigma) and eluted using an imidazole gradient. Fractions containing Hice1-MTBR were pooled and digested by PreScission protease for 24 h in a buffer containing 20 mM HEPES, pH 7.4, 100 mM NaCl and 3 mM dithiothreitol. The protein was purified over a HiTrap S HP column (GE Healthcare) using a NaCl gradient and further purified over a 16/60 Superdex 75 column (GE Healthcare). Selected fractions containing Hice1-MTBR were pooled, concentrated and stored at −80 ◦ C. Full-length Hice1-containing sub-complexes, including Hice1·hDgt61 (433– 955) dimer, tetramer-I, tetramer-II and hexamer, were expressed in E. coli BL21 StarRIL strain (Invitrogen) and proteins were induced by the addition of 0.5 mM IPTG at 18 ◦ C for 16 h. Cells were collected by centrifugation and resuspended in a buffer containing 20 mM HEPES, pH 7.4, 300 mM NaCl, 3 mM dithiothreitol and complete EDTA-free protease inhibitors (Roche). The lysate was then applied to a glutathione S-transferase column (GE Healthcare) and eluted using a glutathione gradient. Fractions containing the proteins were pooled and GST was removed by PreScission protease cleavage for 24 h in a buffer containing 50 mM K-phosphate, pH 7.4, 300 mM NaCl and 5 mM β-mercaptoethanol. The proteins were then purified using a HisTrap column (GE Healthcare) through imidazole elution and further purified over either a 16/60 Superdex 200 or a 10/300 Superose 6 column (GE Healthcare). Fractions from size-exclusion columns were applied to and concentrated by a Hitrap S HP column (GE Healthcare). Selected fractions were dialysed against buffer containing 20 mM HEPES, pH 7.4, 300 mM NaCl, 3 mM dithiothreitol and 10% sucrose and stored at −80 ◦ C. Purifications of sub-complexes that contain truncated Hice1 were essentially the same as for the isolation of the sub-complexes containing full-length Hice1, except that the Hitrap S column was replaced by a Hitrap Q column in the final step (GE Healthcare). Various MBP-tagged and GFP-tagged protein complexes were isolated using a similar expression and purification protocol.

Construction of multi-gene baculovirus expression vectors. To generate baculovirus strains expressing the augmin octameric complex, we used the Multibac system. Augmin subunits were PCR-amplified using high-fidelity hot-start DNA polymerase (Thermo Scientific) and individually cloned into either pACEB1 or

pACEB2 transfer vector (ATG:biosynthetics GmbH). Restriction sites needed for cloning into the MultiBac vectors were included in the PCR primers. To facilitate cloning, internal BstXI restriction sites in augmin subunits hDgt6 and Ccdc5 were removed by introducing silent mutations using site-directed mutagenesis. GST, 6xHis and GFP tags were engineered in-frame at the 50 end of the augmin subunits. All augmin gene constructs were sequence verified. Full-length Hice1, truncated Hice1 (a.a. 141–410), UCHL5IP and C14orf94 were cloned into BamHI/NotI sites within the MCS of the pACEB1 vector. Ccdc5 was cloned through BamHI/SalI sites for insertion into pACEB1 vector. hDgt5 was cloned through EcoRI/SalI sites for assembly into the pACEB1 vector. A TEVcleavable 6-His-tagged hDgt3 was cloned through XhoI/KpnI into the pACEB2 vector. GFP–Cep27 and GST–hDgt6 (1–432) were amplified by PCR using bacterial expression constructs as templates and cloned through XhoI/KpnI sites in the pACEB2 vector. PCR-amplified DNA fragments of GST-tagged full-length hDgt6 (a.a. 1–955) were cloned through XhoI/KpnI sites for insertion into the pACEB2 vector and subsequently GFP was inserted between GST and hDgt6. Individual gene expression cassettes that included the sequence-verified augmin gene, promoter and terminator were excised by I-CeuI and BstXI. Stepwise assembly of individual gene expression cassettes into the pACEB1-full-length Hice1 and pACEB1-truncated Hice1 (a.a. 141–410) resulted in the following final expression constructs: (A) pACEB1-Hice1/C14orf94/UCHL5IP/Ccdc5/GFP– Cep27/hDgt5/His–hDgt3/GST–GFP–hDgt6; (B) pACEB1-Hice1(a.a. 141–410)/ C14orf94/UCHL5IP/Ccdc5/ GFP–Cep27/hDgt5/His–hDgt3/GST–hDgt6 (1–432); (C) pACEB1–Hice1/C14orf94/UCHL5IP/Ccdc5/ GFP–Cep27/hDgt5/His–hDgt3/ GST–hDgt6 (1–432). Subsequently, the expression constructs containing eight subunits of augmin were transformed into DH10MultiBacTurbo cells (ATG:biosynthetics GmbH) to generate bacmids.

Purification of recombinant octameric augmin complexes from insect cells. High Five insect cells (Invitrogen) were infected with baculovirus that contained eight subunits of the augmin complex. Cells were collected at 1, 000g for 15 min and resuspended in lysis buffer that contained 50 mM K-phosphate, pH 7.4, 300 mM NaCl, 5% sucrose, 0.1% Tween-20, 5 mM β-mercaptoethanol, 0.1 mM phenylmethylsulphonyl fluoride and complete EDTA-free protease inhibitors. After sonication, cell lysate was centrifuged for 30 min at 40, 000g . The supernatant was then applied to a glutathione S-transferase column. Subsequently, the column was equilibrated with washing buffer (50 mM K-phosphate, pH 7.4, 300 mM NaCl and 5 mM β-mercaptoethanol) and recombinant augmin was eluted with 50 mM glutathione. Fractions containing the proteins were pooled and GST was removed by PreScission protease cleavage for 24 h at 4 ◦ C in a buffer containing 50 mM K-phosphate, pH 7.4, 300 mM NaCl and 5 mM β-mercaptoethanol. The proteins were then purified using a HisTrap column through imidazole elution and further purified over a 10/300 Superose 6 column. Fractions from size-exclusion columns were applied to and concentrated by a HisTrap column. Pure proteins were dialysed against buffer containing 1×BRB80 (80 mM PIPES, 1 mM EGTA and 1 mM MgCl2 , pH 6.8), 100 mM KCl, 3 mM dithiothreitol and 10% sucrose and stored at −80 ◦ C.

Circular dichroism spectroscopy. Circular dichroism spectra were recorded using an Aviv Circular Dichroism Spectrometer, Model 420. A 10 µM solution of each protein in 10 mM potassium phosphate, pH 7.0 and 100 mM NaCl was analysed at 20 ◦ C. Data were collected from 190–260 nm at 0.1 nm intervals recorded at 50 nm min−1 ; measurements were averages of 5 scans. The mean residue molar ellipticity [θ] (deg × cm2 × dmol−1 ) was determined using ellipticity in degrees, protein concentration, cell path length and molecular weight of protein.

Electron microscopy and image processing. The tetramer and octamer protein samples were diluted to ∼60 µg ml−1 in buffer (20 mM HEPES pH 7.4, 100 mM NaCl). Three microlitres of protein solution was applied to a glow-discharged, carbon-coated Maxtaform, 400-mesh Cu/Rh EM specimen grid (Ted Pella) or a CFlat 2/2 400-mesh EM grid (Electron Microscopy Sciences), with a thin layer of carbon over the holes and preserved with 2% (w/v) uranyl acetate or uranyl formate. Images were recorded with a Tecnai Spirit (Philips/FEI) electron microscope (FEI), equipped with an LaB6 filament and operating at an acceleration voltage of 120 kV at a magnification of ×52,000 (2.05 Å per pixel) on a 4k × 4k Teitz CMOS camera, at 1 ∼ 2 µm underfocus. The automatic data collections were carried out using Leginon25. Tetramer-I (2,831 particles), MBP-labelled tetramer-I (N-terminal MBPtagged Hice1, 1,539 particles; C-terminal MBP-tagged Hice1, 2,967 particles) and octamer (8,000 particles) images were manually selected and boxed using e2boxer26. Twofold pixel binning of the original particle images resulted in a final pixel size of 4.1 Å. We carried out image alignment and classification using the Iterative Stable Alignment and Clustering (ISAC) program in the SPARX software package27,28.

Microtubule co-sedimentation assay. Taxol-stabilized microtubules were polymerized from purified pre-cleared bovine tubulin. Microtubules were

NATURE CELL BIOLOGY © 2014 Macmillan Publishers Limited. All rights reserved.

METHODS

DOI: 10.1038/ncb3030 incubated in ∼1 µM proteins for 20 min at room temperature in the buffer containing 0.5× BRB80, 100 mM KCl, 10% sucrose and 0.25 mg ml−1 BSA. Subsequently, reaction solutions were subjected to sedimentation in a TLA 120.1 rotor (Beckman Coulter) at 75,000 rpm for 10 min at 27 ◦ C. The proteins in the pellet and supernatant were analysed by SDS–PAGE and the Coomassie-stained bands were quantified (LI-COR Odyssey). Band intensities from the gels were used to determine the fraction of protein bound, which was plotted against microtubule concentration. The data were fitted to a modified Hill equation to determine Kd .

Spin-down and fixation of Xenopus metaphase spindles and microtubule asters onto coverslips. Xenopus egg extracts containing either spindles or

Single-molecule TIRF analysis of GFP-tagged augmin complexes. TIRF data

of asters were recorded and stored as 16-bit TIFF files. Asters were first identified visually, and regions that fully encompassed single isolated asters were cropped. Polar transforms were performed by first locating the centre of the aster, and running the Image J plug-in ‘Polar Transformer’, resulting in the mapping of the image from Cartesian coordinates to radial and angular coordinates. The average intensity across all angles at each radius was then calculated using the Plot Profile tool in ImageJ. These data were then normalized against the integrated intensity within each aster, such that the percentage of the aster’s total microtubule intensity at a given radius could be determined. Radial intensity data from multiple asters formed under a given set of conditions were then averaged to determine a characteristic radial profile (Ran control asters, N = 28; Holo-complex, N = 31; octamer[hDgt61 (433–955)] (15 nM), N = 35; octamer[hDgt61(433–955)] (60 nM), N = 37; Tetramer-II (60 nM), N = 27). Angular profiles were determined by drawing a circular path at a fixed radius from the aster’s centre. The ImageJ plug-in ‘Oval Profile’ was used to determine the intensity at all angles along the circular path. For each aster, paths were drawn at radii = 4, 8 and 12 µm. From these angular intensity profiles, the centre of the region containing the most microtubule density was determined by a centre-of-mass analysis. Intensity data contained within a half circle (or, ±90 degrees on either side of the region’s centre) were selected, and the coefficient of variation (the standard deviation divided by the mean) was calculated. The mean and standard deviation of data pooled for each aster-forming condition were then measured (experimental statistics identical to above).

were collected using a Nikon Ti Eclipse inverted microscope with a custom twocolour TIRF illumination system. GFP was excited using a 488 nm laser (Spectra Physics) and X-rhodamine with a 561 nm laser (Cobolt Jive). Microtubules were polymerized from a solution that contained unlabelled, X-rhodamine-labelled, and biotin-labelled tubulin in a ratio of 10:1:0.7 at 37 ◦ C in the presence of guanalyl-(α,β)-methylene diphosphate (GMPCPP). Flow chambers assembled with biotin–polyethylene glycol (bioPEG)-coated coverslips were sequentially filled with 0.5 mg ml−1 α-casein, 0.2 mg ml−1 NeutrAvidin, and biotin-labelled microtubules in the presence of 20 µM taxol and BRB80. Finally, GFP-labelled protein was added to the chamber in BRB80 supplemented with 20 µM taxol, 0.5 mg ml−1 α-casein, 10% sucrose, 2 mM dithiothreitol, 200 µg ml−1 glucose oxidase, 35 µg ml−1 catalase and 4.5 µg ml−1 glucose. A single image of the microtubules was taken and time-lapse sequences were recorded in the GFP channel at 4.4 frames per second with 100 ms exposure. Spot detection and track assembly were performed using the Speckle Tracker J plugin for ImageJ (ref. 29). The dwell time of each track was determined from the duration of the track using a custom routine in Excel. The dwell time frequency (d.t.f.) was calculated using the equation N (ti ) d.t.f(ti ) = P N (tj ) j=1

where N (ti ) is the number of tracks that have a given dwell time (ti ). The empirical Cumulative Distribution Function (CDF) was calculated using the equations i−1 X

CDF(ti ) =

d.t.f.(tj )

For i > 1

CDF(ti ) = 0

For i = 1

j=1

The paired data (1 − CDF, ti ) was fitted to the exponential sum 1 − CDF(ti ) = A1 · e−ti /τ1 + A2 · e−ti /τ2 , using a ‘least-squares’ approach in MATLAB. The second part of the sum was excluded in the case of the mono-exponential fit on Hice1·GFP-hDgt6 (1–432) data. The parameters A and τ , the relative amplitude and mean dwell time respectively, are expressed as the mean value for the estimator ± the 95% confidence intervals extracted from the fit. For the GFP intensity analyses, GFP-labelled augmin complexes were nonspecifically bound to the glass surface of flow chambers that were assembled from untreated coverslips to obtain a sparse decoration. The same buffer solution used in the single-molecule microtubule-binding assay was used in these assays. Images were acquired in the GFP channel at 4.4 frames per second with a 100 ms exposure. The first three frames were averaged and used for the analysis. Spots were detected using Speckle Tracker J, and their intensities were determined using a custom routine in MATLAB and Microsoft Excel.

Spindle and microtubule aster assembly in Xenopus egg extracts. Cytoplasmic extracts from the Xenopus laevis eggs, arrested in metaphase of meiosis II, were prepared as previously described24. Spindles were assembled in the presence of Xrhodamine-labelled tubulin and sperm DNA. Imaging analysis was carried out to examine the spindle localization of augmin complex without fixation. Experiments were repeated three times or more for each condition. Linescans of GFP fluorescence in each spindle were carried out with a 10-pixel-wide stripe along the spindle axis. Pole microtubule and GFP intensities were measured by averaging over a 40-pixel region within the spindle. Midzone microtubule and GFP intensities were measured by averaging over a 40-pixel region near the spindle centre. For microtubule aster assembly, X-rhodamine-labelled tubulin and Ran(Q69L) were added to a final concentration of 0.1 mg ml−1 and 15 µM, respectively. Ten microlitres of egg extracts was mixed with 1 µl augmin protein solution. Subsequently, a total of 2 µl of extract for each condition was fixed onto two different coverslips, imaged and analysed. To quantify aster number, fixed samples were scanned automatically using a motorized XY stage and captured images were stitched into one composite image (NIS-Elements). Fluorescence images of spindles and microtubule asters were acquired with a Cooled-CCD (charge-coupled device) camera (CoolSNAP, Photometrics), which was mounted on the microscope base-port. Image acquisition was performed with either Metamorph (Molecular Devices) or NIS-Elements (Nikon).

microtubule asters were diluted in 1 ml of dilution buffer (1×BRB80, 30% glycerol, and 0.5% Triton X-100) and subsequently an equal volume (1 ml) of fixation solution (1×BRB80, 30% glycerol, 0.5% Triton X-100 and 4% formaldehyde) was added. Fixed samples were layered onto cushion (1×BRB80 and 40% glycerol) and then spun down spindles or asters onto coverslips.

Polar and radial intensity analyses of microtubule asters. Fluorescence images

Generation of antibody against Xenopus laevis Ccdc5. The recombinant GSTtagged Xenopus laevis Ccdc5 protein (a.a. 1–204) was purified by GST affinity and size-exclusion chromatography. GST tag was removed by PreScission protease and non-tagged recombinant xCcdc5 was used to produce rabbit polyclonal anti-xCcdc5 antibody serum (Cocalico). For affinity purification of antibody, recombinant Ccdc5 protein was coupled to a HiTrap NHS column (GE Healthcare) and subsequently sera containing the anti-Ccdc5 antibody were passed over the antigen-coated column. The antibody was eluted with low-pH buffer and dialysed check aganist storage buffer (50 mM PBS and 50% glycerol).

Electron microscopy of augmin-decorated microtubules. Microtubules (∼150 µg ml−1 in 1×BRB80) were applied to glow-discharged carbon grids, then incubated with either GFP–teramer-I or dimer (∼200 µg ml−1 in 10 mM HEPES, pH 7.0, 2% sucrose and 60 mM NaCl) for 2 min, grids were washed with buffer without sucrose and stained with 2% (w/v) uranyl acetate. Images were recorded with a Tecnai Spirit electron microscope (FEI). Images were recorded at 29,000× (3.75 ångström per pixel on a 2K × 2K Tietz camera).

Size-exclusion chromatography coupled light scattering. The light-scattering data were collected using a Superose-6 chromatography column (GE Healthcare), connected to a high-performance liquid chromatography system (HPLC), Agilent 1200 (Agilent Technologies), equipped with an autosampler. The elution from SEC was monitored by a photodiode array (PDA) UV/VIS detector (Agilent Technologies), a differential refractometer (OPTI-Lab rEx Wyatt), and a static and dynamic, multi-angle laser light scattering (LS) detector (HELEOS II with QELS capability, Wyatt). The weight average molecular masses were determined across the entire elution profile in 1 s intervals from static LS measurements using ASTRA software as previously described30. 25. Suloway, C. et al. Automated molecular microscopy: the new Leginon system. J. Struct. Biol. 151, 41–60 (2005). 26. Tang, G. et al. EMAN2: an extensible image processing suite for electron microscopy. J. Struct. Biol. 157, 38–46 (2007). 27. Hohn, M. et al. SPARX, a new environment for cryo-EM image processing. J. Struct. Biol. 157, 47–55 (2007). 28. Yang, Z., Fang, J., Chittuluru, J., Asturias, F. J. & Penczek, P. A. Iterative stable alignment and clustering of 2D transmission electron microscope images. Structure 20, 237–247 (2012). 29. Smith, M. B. et al. Interactive, computer-assisted tracking of speckle trajectories in fluorescence microscopy: application to actin polymerization and membrane fusion. Biophys. J. 101, 1794–1804 (2011). 30. Folta-Stogniew, E. & Williams, K. R. Determination of molecular masses of proteins in solution: implementation of an HPLC size exclusion chromatography and laser light scattering service in a core laboratory. J. Biomol. Tech. 10, 51–63 (1999).

NATURE CELL BIOLOGY © 2014 Macmillan Publishers Limited. All rights reserved.

S U P P L E M E N TA R Y I N F O R M AT I O N DOI: 10.1038/ncb3030

Supplementary Figure 1 Biochemical characterization of augmin subcomplexes. (a-c) Oligomeric states of Hice1· hDgt6D(433-955) (a), tetramer-I (Hice1·hDgt6 (1-432)·UCHL5IP·Cep27) (b), and tetramer-II (Hice1·hDgt6 (1-432)·His-C14orf94·Ccdc5) (c) were analyzed by size exclusion chromatography coupled with light scattering. Recombinant subcomplexes elute earlier than expected for a globular protein of equivalent molecular mass. Elution volumes of molecular weight standards used in the size exclusion chromatography are indicated. The average molecular mass for sub-complexes analyzed by light scattering are indicated (lines across the respective elution peaks). (d) Observed results from light

scattering experiments (Obs) are presented beside molecular weights calculated using standards (Cal). (e) Purified hexamer (Hice1·hDgt6 (1-432)·UCHL5IP·Cep27·His-C14orf94·Ccdc5) was analyzed by SDSPAGE, followed by staining with Coomassie blue. (f) Western blot analysis of the hexameric complex with the indicated antibodies. (g) C14orf94 (a.a. 1-188) · full length Ccdc5 hetero-dimer was examined by size exclusion chromatography (Superdex 200 16/60). Peak fractions (between 60 and 80 ml; volumes indicted) were analyzed by SDS-PAGE (staining with Coomassie blue). Void volume (Vo) is indicated. The star indicates a GST-containing peak. Absorbance (A.U.) is 280 nm.

WWW.NATURE.COM/NATURECELLBIOLOGY

1 © 2014 Macmillan Publishers Limited. All rights reserved.

S U P P L E M E N TA R Y I N F O R M AT I O N

Supplementary Figure 2 Analysis of subunits in the augmin holo-complex. (a) Recombinant holo-complex (Hice1·GFPhDgt6·UCHL5IP·GFP-Cep27·C14orf94·Ccdc5·His-hDgt3·hDgt5) was analyzed by SDS-PAGE, followed by staining with Coomassie blue.

(b-d) Western blots were carried out using the indicated antibodies. GFP-tagged Cep27 within purified tetramer-I was also detected using anti-GFP antibody (b). The star indicates a GFP-containing degradation product of GFP-Cep27.

2

WWW.NATURE.COM/NATURECELLBIOLOGY © 2014 Macmillan Publishers Limited. All rights reserved.

S U P P L E M E N TA R Y I N F O R M AT I O N

Supplementary Figure 3 Single molecule analysis of augmin complexes. (a-f) Fluorescence intensity average and distribution analysis of single particles for each complex adhered to a glass surface: (a) Hice1· hDgt6D(433-955) (intensity average 5400 ± 1800, N = 593 particles), (b) tetramer-I (GFPCep27) (intensity average 4800 ± 1800, N = 600 particles), (c) tetramer-I (GFP-hDgt6 (1-432)) (intensity average 5400 ± 2500, N = 631 particles), (d) octamer[hDgt6D(433-955)] (intensity average 4500 ± 2000, N = 626 particles), (e) holo-complex (8300 ± 7000, N = 662 particles). (f) Monomeric GFP (intensity average 4800 ± 1700, N = 577 particles) was used as reference. Intensity averages are reported as mean ± SD. Three or more independent experiments were analyzed. (g-k) The mean square displacements for Hice1· hDgt6D(433-955) (g), tetramer-I (GFP-Cep27) (h), tetramer-I (GFP-hDgt6 (1-432)) (i), octamer [hDgt6D(433-955)] (j),

holo-complex (k) are shown. Diffusion coefficients (D) are calculated as half the slope of the fit line. Three or more independent experiments were analyzed. (l) Image of GMPCPP-stabilized microtubules (X-rhodamine- and biotin-labeled) (top), GFP-tagged complexes (middle, maximum intensity projections from 300 images in the time-lapse sequence) and corresponding kymographs (below) are shown for GFP-tagged tetramer-I (GFP-hDgt6 (1432)) that also has a GFP on hDgt6, as is the case for the dimer. Scale bars: horizontal, 2 µm; vertical, 2 seconds. (m) The region highlighted (box) in each kymograph is also shown in greater detail as a montage. Scale bar: 2 µm. (n) Binding of individual molecules to a microtubule was tracked to compute the CDF of the dwell time for the complex. Mean dwell time < t > and relative amplitude (in parentheses) were obtained by fitting to bi-exponential functions (gray curve). Data from three or more independent experiments were analyzed.

WWW.NATURE.COM/NATURECELLBIOLOGY

3 © 2014 Macmillan Publishers Limited. All rights reserved.

S U P P L E M E N TA R Y I N F O R M AT I O N

Supplementary Figure 4 Electron microscopy analyses of augmin subcomplexes. (a,b) Negative stain electron micrographs of taxol stabilized microtubules in the presence of (a) Hice1·hDgt6D(433-955) and (b) GFPtagged tetramer-I. Scale bar, 40 nm. Microtubule without tetramer-I binding is indicated by arrow. (c) SDS-PAGE analysis (stained with Coomassie blue) of purified tetramer-I with MBP-tagged Hice1 C-terminus. (d) Negative stain electron micrograph of the MBP-tagged tetramer-I (Hice1-MBP·hDgt6 (1-432)

·UCHL5IP·Cep27) particles. Three representative class averages of individual particles (N=82, 52 and 49) are shown. The position of the MBP-tag is indicated (arrow). Scale bar, 10 nm (e,f) Negative stain electron micrograph of tetramer-I (Hice1·hDgt6 (1-432)·UCHL5IP·Cep27) (e) and tetramer-II (Hice1·hDgt6 (1-432)·C14orf94·Ccdc5) (f) particles. A field of either tetramer-I or -II particles adsorbed onto a glow-discharged carbon grid was stained with 2% (w/v) uranyl acetate and processed for imaging. Scale bar, 100 nm.

4

WWW.NATURE.COM/NATURECELLBIOLOGY © 2014 Macmillan Publishers Limited. All rights reserved.

S U P P L E M E N TA R Y I N F O R M AT I O N

Supplementary Figure 5 Analysis of metaphase spindle localization of augmin complexes. (a) Western blot analysis of crude Xenopus egg extracts (10-fold dilution in CSF-XB buffer, 7ml) with anti-xCcdc5 antibody (lane 6). Equal volumes (7 ml) of serial dilutions of recombinant xCcdc5 (a.a. 1-204) were analyzed SDS-PAGE to determine endogenous xCccdc5 concentration (lane 1 to 5). Concentrations of recombinant xCcdc5 protein are indicated. (b) Localization of endogenous augmin in the metaphase spindles assembled in Xenopus egg extracts, fixed, spun down on coverslips and stained with antixCcdc5 and anti-a-tubulin antibodies. xCcdc5 antibody staining, left panels; and a-tubulin antibody staining (middle panels); and overlays (right panels; microtubules, red; xCcdc5, green; and DNA, blue), are shown. Scale bar, 10 μm. (c) Additional fluorescence images of metaphase spindles in the presence GFP-tagged octamer[hDgt6D(433-955). GFP (left panels); X-rhodamine (middle

panels); and overlays (right panels; microtubules, red; GFP, green; and DNA, blue), are shown. Scale bar, 10 μm. (d,f) Fluorescence images of metaphase spindles in the presence of GFP-tagged Hice1·hDgt6D(433-955) (d) and GFP protein (f) (left panels, 15 nM); X-rhodamine (middle panels); and overlays (right panels; microtubules, red; GFP, green; and DNA, blue), are shown for spindles in d, f. Scale bar, 10 μm. (e,g) Linescans for GFP fluorescence (green) and X-rhodamine (red) along the long axis of the spindle are shown. (h) The intensity of the GFP image in the presence of GFP-tagged octamer[hDgt6D(433-955), Hice1D(1-140)] was adjusted to visualize signal at the spindle poles. (i) Localization analysis of augmin holo-complex in metaphase spindles using Xenopus egg extracts. Recombinant GFP-tagged holo-complex (6 nM) (left panels); tubulin (X-rhodamine-labeled, middle panels); and overlays (right panels; tubulin, red; GFP, green; and DNA, blue), are shown. Scale bar, 10 μm

WWW.NATURE.COM/NATURECELLBIOLOGY

5 © 2014 Macmillan Publishers Limited. All rights reserved.

S U P P L E M E N TA R Y I N F O R M AT I O N

Supplementary Figure 6 Microtubule aster formation in the presence of augmin complexes. (a, b)Aster assembly in the presence of Ran(Q69L) and GFP-tagged octamer[hDgt6∆(433-955)] (a) or holo-complex (b) at 6 nM. (c) Immunodepletion of xCcdc5 from Xenopus egg extracts using anti-xCcdc5 antibody. xCcdc5 bands in crude (A), IgG-depleted (B) and xCcdc5-depleted (C) egg extracts were analyzed by western blots. Equal volumes of egg extract samples with the same dilution were analyzed by SDS-PAGE. Supernatant fractions (S) and bead bound fractions (P) are indicated. IgG and xCcdc5 bands are indicated. Three independent experiments were performed and depletion was found to be ~60%. (d) A representative image of microtubule aster formed in the presence of Ran(Q69L) and GFP-tagged tetramer-II at 15 nM. Tubulin (X-rhodaminelabeled), left panel; and GFP fluorescence, right panel, are shown. Scale

bar, 10 μm. (e) Analysis of microtubule (rhodamine signal) and augmin (GFP signal) levels in asters. Ratios of average fluorescence at 1mm versus 5 mm radius for tetramer-II (15 nM) induced asters are shown. (N=19 asters; S.D. was determined from data pooled from 3 independent experiments.) (f-h) Representative images after polar transformation of asters induced by RanQ69L alone (f) and in the presence of Ran(Q69L) and holo-complex (15 nM) (g) and octamer[hDgt6∆(433-955)] (15 nM) (h). (i-m) Representative images of microtubule asters induced by Ran(Q69L) alone (i) and in the presence of Ran(Q69L) and GFP-tagged holo-complex (15 nM) (j), octamer[hDgt6∆(433-955)] (15 nM) (k), octamer[hDgt6∆(433-955)] (60 nM) (l) or tetramer-II (60 nM) (m). Three half-circle (180 degree) regions (at 4 and 8, and 12 mm radii) used for determination of coefficient of variation are indicated in first image for each panel (dashed white lines).

6

WWW.NATURE.COM/NATURECELLBIOLOGY © 2014 Macmillan Publishers Limited. All rights reserved.

S U P P L E M E N TA R Y I N F O R M AT I O N

Supplementary Figure 7 Analysis of aster number and microtubule intensity in asters promoted by addition of recombinant augmin complexes to egg extracts. Individual fluorescence images of X-rhodamine-labeled microtubules were taken automatically using a motorized XY stage and then 400 captured images were stitched into one composite image using NIS-Element software. A representative single image from each composite image is shown in the bottom inset. Augmin complexes and Ran(Q69L) were added to extracts, incubated for 10 min (or as noted), fixed and processed. (a) Representative composite images of microtubule asters in the presence of buffer control, GFP-tagged octamer[hDgt6D(433-955)] and holo-complex. (b) Representative composite images of microtubule asters in the presence of GFP-tagged octamer[hDgt6D(433-955),

Hice1D(1-140)] and octamer[hDgt6D(433-955)] with Ran(Q69L). (c) Representative composite images of microtubule asters in the presence of indicated GFP-tagged octamer[hDgt6D(433-955)] concentrations. (d) Representative composite images of microtubule asters in the presence of GFP-tagged octamer[hDgt6D(433-955)] without Ran(Q69L). Scale bars, 1 mm (composite image); 100 mm (single image). (e) Total microtubule intensity in asters induced in the presence of Ran(Q69L) and GFP-tagged tetramer-II or octamer[hDgt6D(433-955)] at 60 nM. (f) Total microtubule intensity in asters induced by of Ran(Q69L) alone and in the presence of Ran(Q69L) and GFP-tagged holo-complex or octamer[hDgt6D(433-955)] at 15 nM. S.D. was determined from data pooled from at least 3 independent experiments.

WWW.NATURE.COM/NATURECELLBIOLOGY

7 © 2014 Macmillan Publishers Limited. All rights reserved.

Reconstitution of the augmin complex provides insights into its architecture and function.

Proper microtubule nucleation during cell division requires augmin, a microtubule-associated hetero-octameric protein complex. In current models, augm...
3MB Sizes 0 Downloads 4 Views