1
SUPPLEMENTARY INFORMATION
Supplementary Methods
Assembly and visualization of GDP-tubulin helical tubes. Preparation of subtilisincleaved GDP-tubulin was done as follows. Bovine brain tubulin from Cytoskeleton
(Denver, CO) at a concentration of 6 mg/ml was incubated at 37 ˚C for 30 minutes in
CBII buffer (80 mM PIPES, 1 mM EGTA, 1 mM MgCl2, and 10% glycerol at pH 6.8)
in the presence of 2.5 mM GTP to form microtubules. 25 g/ml subtilisin was added,
and then quenched after 10 minutes by adding 3.0 mM PMSF. The subtilisin-cleaved
microtubules were centrifuged down at 13.5K RPM for 15 minutes, washed with
prewarmed CBII buffer and pelleted by an additional 8 minutes’ centrifugation. The
pellet was resuspended in cold CBI buffer (as CBII buffer minus glycerol) to a
concentration of about 10 mg/ml. After 20 minutes in ice all the microtubules were fully
depolymerized. The solution was centrifuged at 50K RPM for 15 minutes at 4oC to
remove any insoluble aggregates. The supernant was characterized by SDS-PAGE and
Western blotting with antibodies against -tubulin and -tubulin. Following this
procedure about half of the -tubulins’ C-termini were cleaved, while practically all tubulin molecules were intact. Although partial removal of the C-terminus of tubulin by
subtilisin treatment is not necessary for the formation of the tubes, it has a positive
effect on their crystalline order. More extensive subtilisin digestion or higher
concentration of manganese resulted in more triple-layered tubes and were avoided.
The GDP-tubulin prepared as above was diluted to a final concentration of 2~3
mg/ml in CBI buffer. GDP and MnCl2 were added to the tubulin solution to give final
concentrations of 2 mM and 20~50 mM, respectively. The solution was incubated at
37oC to promote GDP-tubulin assembly (although tubes can form at any temperature
between 4 and 37 ˚C). After 3~5 hours incubation, the solution became cloudy. 3.5 l of
2
this solution was taken and applied onto a chloroform-pre-treated Quantifoil electron
microscope grid (R1.2/1.3 Cu 400 mesh, SPI, PA) mounted in a Vitrobot (FEI,
Netherlands). The grid was blotted for 1.8 seconds at room temperature with 100%
humidity and plunged into liquid ethane slush cooled by liquid nitrogen at about 172oC. The vitrified grids were stored in liquid nitrogen until examination. Cryo-EM
grids were transferred to a Gating 626 cry-holder (Gating Inc., PA) and inserted into a
CM200 electron microscope with a field emission gun (FEY, Netherlands). The
temperature of the grid in the microscope was maintained below -172oC during the
whole process of examination and image recording. Using low-dose protocols, images
of the tubular crystals of GDP-tubulin were recorded on Kodak SO-163 films at a
magnification of 50 K and acceleration voltage of 200 KV with a dose lower than 15 e/Å2 per picture. The defocus of the images ranged between 0.5 m and 2.0 m. In some
sessions, focal pairs were taken of the same area to facilitate the classification of the
tubes into different families (see below) using the higher defocus image. The films were
developed and screened both by eye and optical diffraction to remove those with
astigmatism or drift. Selected films were digitized on a Nikon Super Coolscan 8000 ED
scanner (Nikon, Japan) at 2000 dpi, resulting in a final pixel size of 2.54 Å. The images
were transferred to a SGI workstation and converted from transmission to optical
density using the EMAN image processing package1.
Reconstruction of the helical tubular crystals of GDP-tubulin. In the presence of
high concentration of manganese in solution, GDP-tubulin formed well-ordered, multilayered tubular crystals. Most of them are double-layered tubes while some are triplelayered (Fig. S1a). Analysis of the diffraction patterns from images of these tubes
indicated that each layer is a one-start helix corresponding to a curved tubulin
protofilament (Fig. S1b and S2). Depending on the number of tubulin subunits per turn
3
of these helices, the tubular crystals can be classified into different families. Each tube
was assigned to a certain family based on its number of layers, the diameters of each
layer in the tube (and thus the number of tubulin subunits per turn) and the parallel/antiparallel relationship among the protofilaments of different layers (whether they were
parallel or anti-parallel was determined by examining the directions of tubulins’ end-onview projections at the edge walls of the two layers in filtered images). Table S1
summarizes the families of tubes observed under our experimental conditions. For the
structure analysis described in this study, we focused on the most abundant family of
double-layered tubes with 24/32 subunits in anti-parallel protofilaments. The double
layer character of the tubes results in systematic overlap of the Bessel terms from inner
and outer layers on all the layer lines2 (supplementary Fig S2) and makes it impossible
to use traditional helical reconstruction methods. Here we have implemented an iterative
Fourier Bessel method developed in our laboratory2 by which the relative orientation of
different tube images can be determined and used to produce independent 3-D
reconstructions of the inner and outer layers of the tube.
Images of 19 tubes from the anti-parallel 32/24 family of double-layered tubes
were chosen for alignment. Among them, 10 tubes were imaged as focal pairs, so
altogether 29 images were included for analysis and reconstruction. The 29 images were
taken at different defocus values and included tubes of different lengths and helical
selection rule (Table S2).
The image of each tube was cut from the original micrograph within a box about
four times wider than that of the tube itself. The in plane curvature of the tubes were
corrected using the straightening algorithm in the Phoelix package3. After straightening
the images were cut into three parts, the tube itself and the two images of ice on both
sides of the tube. These two images were averaged and used as the estimated
background noise for resolution assessment4. Each tube image was then padded into
4
512x4096 pixels, its Fourier transform calculated and the power spectrum was analyzed
for indexing. Layer lines were extracted from the Fourier transform of each tube up to a
resolution of 8 Å. The helical selection rule for each tube differs slightly from the others
because the Z-heights of the layer lines vary in one or two reciprocal units from tube to
tube (Table S2). For the purpose of alignment and average, the layer lines were
reassigned to a common selection rule from the average of all the tubes. The same
process was followed for the noise images.
Though most of the layer lines of double-layered tubes are composed of
overlapped Bessel terms from the inner and outer layers with different Bessel orders,
those corresponding to the one-start helical track have the same order (N=1, 2, 3…)2.
These layer lines, along with the equator, can be used to determine the x-shift and outof-plane tilt of each tube. Using the same method as Amos and Klug5, the x-shift and
out-of-plane tilt of each tube were calculated and corrected separately for the inner-layer
helix and outer-layer helix of each the tube. Tubes with out-of-plane tilt bigger than 5o
were discarded.
Only data within a resolution of 23 Å were used for alignment. During the
alignment cycle focal paired images were half-weighted in the process of data merging
and averaging2. The equator was not used for the alignment. The radial scale of each
tube was also refined during the process of alignment. The iterative alignment of our 29
images of 19 tubes converged to a stable solution after 10 iterations. The alignment
parameters were then used in the reconstruction process.
CTF parameters for each image were calculated from the diffraction spectra of the
ice within the hole of Quantifoil of each micrograph using the SPIDER TF series
programs6. The B factor was set roughly at 100 Å2 based on the above analysis7. The
amplitude contrast ratio was calculated to be 7 % using the methodology of Toyoshima
5
and Unwin8. The alignment parameters were applied to the layer line data and the final
average from all the tubes was calculated using the equation2
M
Favg 
 CTF F
j
j
 CTF
2
j 1
M
j 1
(1)
j
The noise of the final reconstruction was calculated from the average of CTF
corrected noise data using the same alignment parameters and weighting factors of their
corresponding tube image. The signal to noise ratio of every point on the layer lines of
the final reconstruction was calculated. Only those points with signal to noise ratio
higher than 2.0 were considered to be significant and retained in the final dataset9. Some
of the averaged layer lines are shown in Fig S3. Layer lines corresponding to the dimer
of -tubulin are noticeable, reflecting the presence of differences between monomers
and dimers in this GDP-bound tubulin (Fig. S1b and S3).
After Fourier Bessel transformation and radial cutting in little g space2 the real
space reconstructions of inner-layer helix and outer-layer helix were separately
calculated by Fourier integration using the averaged layer line dataset up to a resolution
of 12 Å. The 3-D densities were saved in SPIDER format for the purpose of display
using the WEB and Chimera software packages10,11.
Docking of the atomic model of tubulin into the 3D density of the double-layered
GDP-tubulin tubes. Both the atomic models of tubulin obtained by electroncrystallography in a straight protofilament (1JFF.pdb) and that obtained by X-ray
crystallography with tubulin bound to RB3 and colchicine (1SA0.pdb) were used in
docking experiments into our 3D reconstructions of the inner and outer layer. Regions
missing in the model of one monomer that had a counterpart in another (long loop in the
N-terminal domain of -tubulin; M-loop in the RB3-tubulin structures) were filled in
6
with the available section and energy minimized using molecular dynamic procedures.
This was done to be able to account for most of the mass of the proteins, and was not
intended to reproduce the real structure. The atomic resolution models were fitted
manually as rigid bodies into the OMAP map using the program O12. These manually
fitted atomic models of tubulin were then converted into a density file, low-pass filtered
to 23 Å and threshold filtered to masks. The masks were used to segment the tubulin
heterodimer densities from the inner- and outer- layer reconstructions respectively. The
resulting densities were compared by Fourier shell correlation, indicating that they were
practically indistinguishable within a resolution of 15 Å (data not shown). We tried to
perform rigid body docking into our density maps using Situs13. However, it soon
became apparent that the polymer character of the density made automatic procedures
suboptimal. The performance of the automation was further challenged by the presence
of extra density in our 3-D maps corresponding to the acidic C-terminal amino acids
missing in the crystal structure, but contributing significantly to our reconstruction.
Manual docking incorporated this information and the polymer character of the density
into the “human” search process. Helical symmetries of the two layers of tube were then
applied to the docked atomic models in the asymmetric unit of the reconstruction to
generate a docking of longer protofilaments using HLXBUILD program in Situs13.
Assembly and visualization of GMPCPP-tubulin helical ribbons/tubes. Nonhydrolysable GTP analogue guanylyl-()-methylene-diphosphonate (GMPCPP) was
purchased from Jena Bioscience Company, Germany. Bovine brain tubulin
(Cytoskeleton, Denver, CO) was incubated at 37 ˚C for 30 minutes in CBII buffer
supplemented with 2.5 mM GTP to form microtubules. Microtubules were pelleted,
then resuspended in cold CBI buffer and allowed to depolymerise in ice for 20 minutes.
The solution was centrifuged at 4oC to remove insoluble aggregates and GMPCPP was
7
added the supernatant to a final concentration of 10 mM. After 10 minutes in ice
GMPCPP-tubulin was diluted to 2-3 mg/ml in CBI buffer and MgCl2 was added to a
final concentration from 4 mM to 30 mM. The solution was incubated at different
temperatures: 4oC, 15oC, 25oC and 37oC. The helical ribbons of GMPCPP-tubulin
formed at all temperatures for magnesium concentrations equal or higher than 8 mM.
Only at temperatures higher than room temperature, microtubules were also observed.
For incubation times longer than a couple of hours, well-ordered helical tubular crystals
of GMPCPP-tubulin were obtained (Fig. S5). For the experiments showing the
conversion of GMPCPP-tubulin ribbons into microtubules, the temperature was
increased to 37oC after 4 hours incubation at 15oC. The samples were negatively stained
by 2% uranyl acetate and examined in a JEM-1200 EXII transmission microscope
(JEOL, Japan). Samples with well ordered helical tubes were quick-frozen, examined by
cryo-EM, and the best images screened and digitized following the procedures
described for the double-layered GDP-tubulin tubes (Fig. S6a).
Reconstruction of GMPCPP-tubulin tubes and docking. GMPCPP-tubulin formed
open helical ribbons as well as closed well-ordered helical tubular crystals (Fig S5; Fig.
S6a). The tubes have diameters that range from 500 to 600 Å and belong to different
helical families. The best ordered tubes, as judged by their diffraction patterns, used in
our reconstruction belong to the (11,-37) family. Diffraction patterns were indexed as
shown in Fig S6b using Toyoshima’s method14. Two images with defocus values of 1.8
m and 0.95 m respectively were straightened, and their layer lines extracted up to 12
Å. The two Fourier transforms were aligned and merged with CTF correction as in (1).
The final averaged layer lines show good amplitude and phase signals up to 18 Å (Fig
S7) and were used to calculate a reconstruction of the GMPCPP-tubulin helical tube up
to that resolution (Fig 3a).
8
The atomic model of -tubulin (1JFF.pdb) was manually docked in the monomer
densities of two adjacent protofilaments using O12. The docked atomic structures were
used to generate a full pseudo-atomic model for the whole density by applying helical
symmetry. Monomer alignment for the comparison of this structure with microtubules
used the LSQ command in O12.
Fluorescence microscopy of the conversion of GMPCPP-tubulin tubes into
microtubules. In order to prove that the temperature-driven conversion of GMPCPP
tubes into microtubules was direct, without involving a depolymerization step, we
devised the following fluorescence assay. Two populations of GMPCPP-tubulin tubes
with different fluorescent labels were prepared by incubation of two types of GMPCPPtubulin mixtures (5:1 unlabeled tubulin:fluorescein-tubulin and 5:1 unlabeled
tubulin:rhodamine-tubulin) following the procedure described above. The two
GMPCPP tube samples were pelleted by centrifugation at 30,000 rpm for 10 min below
room temperature and washed once with CBI buffer containing 15 mM magnesium to
remove any free tubulin. The two pellets were resuspended and then mixed in CBI
buffer with 15 mM magnesium. The mixture was incubated at 37 ˚C for over 3 hours to
make sure all the helical ribbons/tubes had converted into microtubules. The incubation
solution was then diluted ten times, fixed and viewed by fluorescence microscopy with
a 60X Nikon objective on a Nikon Eclipse fluorescence microscope according to the
method in Westermann et al.16. A view area was examined using two different filters in
order to record the patterns of fluorescein-tubulin (green) and rhodamine-tubulin (red).
The two patterns were then merged to show relative distributions (Fig 4c). Images
showed no overlap of the two colours, as would have been expected if the GMPCPP
tubes had depolymerised to dimers or small oligomers before microtubule formation.
Interestingly, many microtubules showed adjacent green and red regions indicating
9
annealing of the structures. As a control, the two fluorescent tubulins were mixed before
any assembly, then incubated at 37 ˚C in the presence of GMPCPP to form
microtubules, fixed and examined in the same way (Fig S8). In this case, the green and
red images showed perfect overlap, as expected from assembly of randomly mixed
tubulin dimers with the two labels.
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References for Supplementary Information
1. Ludtke, S. J., Baldwin, P. R. & Chiu, W. EMAN: Semi-automated software for high
resolution single particle reconstructions. J. Struct. Biol. 128, 82-97 (1999) .
2. Wang, H-W. & Nogales, E. An iterative Fourier–Bessel algorithm for reconstruction
of helical structures with severe Bessel overlap. J. Struct. Biol. 149, 65-78 (2005).
3. Carragher, B., Whittaker, M., & Milligan, R.A. Helical processing using PHOELIX.
J. Struct. Biol. 106, 107-112 (1996).
4. Beroukhim, R. & Unwin, N. Distortion correction of tubular crystals: improvements
in the acetylcholine receptor structure. Ultramicroscopy 70, 57-81 (1997).
5. Amos, L.D., & Klug, A. Three-dimensional reconstruction of the contractile tail of T4 bacteriophage. J. Mol. Biol. 99, 51-73 (1975).
6. Zhu, J., Penczek, P.A., Schroder, R., & Frank, J. Three-dimensional reconstruction
with contrast transfer function correction from energy-filtered cryoelectron
micrographs: procedure and application to the 70S Escherichia coli ribosome. J. Struct.
Biol. 118, 197-219 (1997).
7. Huang, Z., Baldwin, P.R., Mullapudi, S., & Penczek, P.A. Automated determination
of parameters describing power spectra of micrograph images in electron microscopy. J.
Struct. Biol. 144, 79-94 (2003).
8. Morgan, D.G., Owen, C., Melanson, L.A., & DeRosier, D.J. Structure of bacterial
flagellar filaments at 11 Å resolution: packing of the -helices. J. Mol. Biol. 249, 88110 (1995).
9. Toyoshima, C. & Unwin, N. Contrast transfer for frozen-hydrated specimens:
determination from pairs of defocused images. Ultramicroscopy 25, 279-292 (1988).
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10. Frank, J., Radermacher, M., Penczek, P., Zhu, J., Li, Y., Ladjadj, M., & Leith, A.
SPIDER and WEB: Processing and Visualization of Images in 3D Electron Microscopy
and Related Fields. J. Struct. Biol. 116, 190-199 (1996).
11. Pettersen, E.F., Goddard, T.D., Huang, C.C., Couch, G.S., Greenblatt, D.M., Meng,
E.C., & Ferrin, T.E. UCSF Chimera - A Visualization System for Exploratory Research
and Analysis. J. Comput. Chem. 25, 1605-1612 (2004).
12. Jones, T.A., Zou, J.-Y., Cowan, S.W., & Kjelgaard, M. Improved methods of
building protein models in electron density maps and the location of errors in these
models. Acta Crystallog. Sect. A 47, 110-119 (1991).
13. Wriggers, W. & Birmanns, S. Using Situs for flexible and rigid-body fitting of
multi-resolution single molecule data. J. Struct. Biol. 133, 193-202 (2001).
14. Toyoshima, C. Structure determination of tubular crystals of membrane proteins. I.
Indexing of diffraction patterns. Ultramicroscopy 84, 1-14 (2000).
15. Westermann, S., Avila-Sakar, A., Wang, H.W., Niederstrasser, H., Wong, J.,
Drubin, D.G., Nogales, E., and Barnes, G. Mol. Cell 17, 277-290 (2005).
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Supplementary Figure Captions
Fig. S1 – Cryo-EM reconstruction of double-layered tubes made of GDP-bound
tubulin.
(a) Image of a frozen-hydrated tube. Scale bar corresponds to 50 nm; (b)
Diffraction pattern from (a); (c) Averaged diffraction pattern for 19 tube images
after alignment using our iterative algorithm13. Arrows point at reflections of a
given resolution to illustrate the gain of signal with averaging.
Fig. S2 - Indexing of the diffraction pattern of a double-layered 24/32 family tube
of GDP-tubulin
Indexing of both the near- and far- sides’ lattice (in red and green respectively)
are shown based on Toyoshima’s method14. The Bessel orders of the innerand outer- layer helices are shown on the right column. The two principal layer
lines (k=0, l=1) and (k=2, l=0) used for assignment of the z-heights in Table S2
are marked with circles.
Fig. S3 – Examples of averaged layer lines used in the reconstruction of the
double-layered tubes of GDP-tubulin
The averaged data of the layer lines of the inner layer tube along the reciprocal
radii are shown in each panel with phases above amplitudes. The Bessel orders
and the reciprocal Z heights of each of the layer lines are indicated. Layer lines
(-13, 0.0260 Å-1) and (10, 0.0317 Å-1) correspond to dimmer repeats along the
13
helices. Layer lines (-82, 0.0086 Å-1), (-19, 0.0827 Å-1) and (-5, 0.0961 Å-1) show
good signal up to highest resolution (10-12 Å).
Fig. S4 – Comparison of the bending at intra and inter dimer contacts between
different tubulin polymers.
Two dimers are displayed (plus end at the top,  subunit) in a front view from
the inside of a microtubule (a) that illustrates tangential bending, and in a side
view (b) that illustrates radial bending (outside of the microtubule to the left).
The coordinates used are from zinc sheets for the microtubule (MT), from the
RB3-tubulin-colchicine (ST), and from the docking into our outer and inner
layers presented in Fig. 1.
Fig. S5 – Polymers of GMPCPP tubulin observed at low temperatures at
different incubation times
GMPCPP-tubulin was in CBI buffer containing 8 mM MgCl2 . Negatively stained
samples show ribbons of different widths and fully close tubes. The lattice of
tubulin subunits and the curvature of protofilaments is the same in all cases.
The scale bar represents 100 nm.
Fig. S6 - Cryo-EM and image reconstruction of GMPCPP-tubulin tubes.
(a) Frozen-hydrated tube (scale bar 100 nm); (b) Diffraction pattern from (a) and
indexing. Layer line (2, 0; 22) corresponds to the lateral repeat between
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protofilaments in microtubules, while layer line (0, 1; -37) corresponds to the
axial subunit repeat along the protofilament.
Fig. S7 – Averaged layer lines used in the reconstruction of the GMPCPPtubulin helical tubes. The nomenclature is the same as in Fig. S3.
Fig. S8 – Fluorescence microscopy of GMPCPP-tubulin microtubules
assembled from a mixture of fluorescein- and rhodamine-tubulin dimers. (a)
Image of fluorescein-tubulin. (b) The same area for rhodamine-tubulin. The two
tubulin patterns show perfect overlap. This serves as a control for the direct
conversion experiment shown in Fig. 4c where the lack of superposition of
rhodamine- and fluorescein-tubulin strongly supports the idea that there is not
depolymerization of GMPCPP tubes previous to the temperature-induced
formation of microtubules.
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Supplementary Tables
Table S1. Different helical families of GDP-tubulin tubes
Number of layers
Outer diameter of every Number of monomers per
Number of images
layer (Å)1
turn in every layer
analyzed
Double
370, 480
22, 30
6
Double
400, 510
24, 32
54
Double
430, 540
26, 34
10
Triple
340, 450, 560
20, 28, 36
4
Triple
370, 480, 590
22, 30, 38
44
Triple
400, 510, 620
24, 32, 40
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1. The image was projected along the tube axis to give a one-dimensional profile that clearly
showed the number of layers and the diameter of each layer.
2. These are approximate numbers based on the Bessel order of layer line (k=2, l=0; see table
S2 and Fig. S2).
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Table S2. Details of individual images used in the reconstruction of the
GDP-tubulin tubes
Image ID
Defocus (m)1
Layer line
Tube
Out-of-plane
assignments2
length (Å)3
tilt
rscale
00941
0.58
202,140
810
-0.5
1.002
08392
1.15
199,136
1570
0.3
1.010
08463
1.19
198,136
620
1.1
1.020
07891
0.75
198,134
1540
0.9
1.018
03121
1.43
197,136
1680
-2.0
1.020
14701
0.86
204,140
930
1.5
1.000
18162
0.75
204,142
620
2.0
1.000
19281
0.70
196,134
1330
-1.3
1.010
30391
0.78
201,142
960
-4.6
1.018
30451
0.61
201,140
2040
-1.2
1.030
30461
1.29
-4
2110
-2.0
1.032
30491
0.74
201,140
2330
4.7
1.025
30501
1.46
-
2250
4.7
1.025
30531
0.65
200,140
1430
-0.6
1.022
30541
1.32
-
1180
-0.3
1.030
30571
0.62
202,142
790
-3.0
1.030
30581
1.28
-
800
-4.5
1.030
30611
0.66
201,142
1410
-1.4
1.020
30621
1.32
-
1270
-1.6
1.020
117261
0.80
198,140
2380
0.5
1.030
117271
1.55
-
2380
0.7
1.030
117301
0.86
199,140
810
-0.9
1.024
117311
1.64
-
610
-0.4
1.024
a2961
0.66
199,142
1450
-0.7
1.028
a2971
1.47
-
1400
-0.9
1.030
a3041
0.75
198,140
840
1.2
1.014
a3051
1.60
-
820
0.9
1.016
a3401
0.57
200,140
980
0.4
1.028
a3411
1.35
-
1000
0.2
1.030
1. All the numbers denote levels of underfocus.
17
2. The image of the tube was interpolated along the tube axis before being padded into a
512x4096 rectangle for Fourier transform in order to best fit the layer lines to the transform grid
in the Z direction. The numbers are for the Z-height of the principal (0,1; -1) and (2,0; -24,-32)
layer lines (see Fig. S2).
3. The length of tube used for reconstruction.
4. These images have the same selection rules as their defocus pair. Other parameters were
determined independently.