Bending Dynamics of Fluctuating Biopolymers Probed by Automated
High-Resolution Filament Tracking
Clifford P. Brangwynne,* Gijsje H. Koenderink,* Ed Barry,yZvonimir Dogic,yFrederick C. MacKintosh,z
and David A. Weitz*§
*Harvard School of Engineering and Applied Sciences, Harvard University, Cambridge, Massachusetts;yRowland Institute at Harvard,
Cambridge, Massachusetts;zDepartment of Physics and Astronomy, Vrije Universiteit, Amsterdam, The Netherlands; and§Department
of Physics, Harvard University, Cambridge, Massachusetts
and dynamics. However, successful extraction of this information requires precise localization of filament position and shape
from thousands of noisy images. Here, we present careful measurements of the bending dynamics of filamentous (F-)actin and
microtubules at thermal equilibrium with high spatial and temporal resolution using a new, simple but robust, automated image
analysis algorithm with subpixel accuracy. We find that slender actin filaments have a persistence length of ;17 mm, and
display a q?4-dependent relaxation spectrum, as expected from viscous drag. Microtubules have a persistence length of several
millimeters; interestingly, there is a small correlation between total microtubule length and rigidity, with shorter filaments
appearing softer. However, we show that this correlation can arise, in principle, from intrinsic measurement noise that must be
carefully considered. The dynamic behavior of the bending of microtubules also appears more complex than that of F-actin,
reflecting their higher-order structure. These results emphasize both the power and limitations of light microscopy techniques for
studying the mechanics and dynamics of biopolymers.
Microscope images of fluctuating biopolymers contain a wealth of information about their underlying mechanics
Actin filaments and microtubules are semiflexible biopoly-
mers that form the elastic cytoskeletal network within cells
that controls cell migration, division, cargo transport, and
mechanosensing (1). Understanding the mechanical behav-
ior of these filaments is thus of central importance for
establishing how they function as a dynamic mechanical
scaffold within living cells. Indeed, mechanical models of
the cell require accurate measurements of the stiffness and
dynamical behavior of the component filaments (2–4).
However, the semiflexible macromolecular backbone of
microtubules and actin filaments results in physical behavior
that remains incompletely understood.
Using light microscope images to directly measure the
shape fluctuations of individual biopolymers is a powerful
technique for studying their dynamics and mechanical
behavior (5–9). By extracting a set of filament coordinates
from each image, the variance of the curvature fluctuations
induced by thermal motion can be used to obtain the bending
rigidity. The rigidity is typically expressed in terms of the
length scale beyond which the filament shows significant
curvature due to thermal forces, known as the persistence
length, lp: Changes in thermally-induced curvature occur on
a timescale that is set by viscous drag from the surrounding
fluid. This time can be obtained from the relaxation time of a
shape autocorrelation function, such as the mean-squared
difference in curvature calculated for increasing lag times.
Utilizing variations of this technique, actin filaments have
been shown to have a persistence length of ;17 mm (5,8,10),
although they were initially suggested to have a length scale-
dependent rigidity (11). Microtubules have been shown to be
orders of magnitude more stiff, with a persistence length on
the order of millimeters (8). However, recent studies have
suggested that microtubule rigidity may depend on their
growth velocity (7) and other factors related to their mac-
romolecular structure (12–14). Indeed, it was recently
suggested that inter-protofilament shearing leads to a soft-
ening of shorter microtubules (15). Other studies have
suggested that internal dissipation mechanisms may domi-
nate over hydrodynamic drag to increase the relaxation times
of microtubules on short length scales (7,16).
Successfully extracting such mechanical information from
microscope images is limited by position uncertainties re-
sulting from image noise, requiring precise filament tracking.
Despite the need for accurate filament localization, reports
utilizing this technique often use semi-manual filament
tracking methods and do not report a noise floor. Automated
video tracking of objects with spherical symmetry, such as
colloidal particles and fluorescent point-sources, is a well-
developed technique for quantifying their dynamic behavior.
Tracking of large numbers of spherical probe particles is
central to microrheological measurements of the mechanical
behavior of soft materials such as biopolymer gels and even
living cells (17–19). Particle tracking algorithms typically
employ an initial particle localization, such as intensity
Submitted September 7, 2006, and accepted for publication February 15,
Address reprint requests to David A. Weitz, Gordon Mckay Professor of
Applied Physics and Professor of Physics, Harvard University, Pierce Hall,
Rm. 321, 29 Oxford St., Cambridge, MA 02138. Tel.: 617-496-2842; Fax:
617-495-2875; E-mail: email@example.com.
Editor: Marileen Dogterom.
? 2007 by the Biophysical Society
346 Biophysical JournalVolume 93July 2007346–359
maxima, followed by particle position refinement using an
intensityweighted center of mass (20) or fitting to a Gaussian
or parabolic function. These algorithms can be used to obtain
particle positions with subpixel accuracy. Positions are then
linked in time to establish the trajectories of the particles.
Precise tracking of objects with nonspherical symmetry,
such as biopolymer filaments, presents more of a challenge.
Unlike colloidal particles, the shape of the filament is not
known a priori and thus conventional automated fitting
techniques are no longer applicable. There is an extensive
computer vision literature that describes identification of lin-
ear structures, such as roads or capillary tubes, from complex
landscapes (21). A related technique for automated tracking
of differential interference contrast (DIC) images of linear
structures such as microtubules has been described (22). A
recent study describes automated tracking of uniformly-sized
fluorescent colloidal rods (23). There are also a number of
papers describing filament-centroid tracking routines for in
vitro motility assays of fluorescent actin filaments gliding
over myosin-coated surfaces (24,25). However, to our know-
ledge there have been no reports describing automated
contour tracking of fluctuating fluorescent filaments with
In this paper, we develop and utilize a robust automated
image analysis algorithm for tracking fluorescently-labeled
biopolymer filaments with subpixel accuracy. We first dem-
onstrate the accuracy of this technique by tracking im-
mobilized fluorescent biopolymers and show that we obtain a
root mean-square precision of ;0.15 pixel (;20 nm), even
as the filaments begin to be photobleached after hundreds of
exposures. We then use this technique to study the shape
fluctuations of microtubules and actin filaments at thermal
equilibrium down to short length and time scales. This
allows us to address recent conflicting reports of the
mechanics and dynamics of biopolymer bending fluctu-
ations, particularly those of microtubules. The persis-
tence length of actin filaments obtained from our
measurements agrees well with previous experiments
(5,8,10), lp? 17 mm. We obtain microtubule persistence
lengths on the order of a few mm, also in agreement with
previous measurements (7,8,13,14). Our data show a
slight correlation between filament length and stiffness,
consistent with a recent study (15). However, we dem-
onstrate that this correlation can in principle arise from
inherent noise limitations that occur even with high precision
measurements. Thus, over the range we study, there does not
appear to be a systematic dependence of the bending rigidity
on length or wavevector, in agreement with other studies
(7,8,13,14). However, after a careful analysis of the relax-
ation times of the fluctuating normal modes, we find
evidence that microtubules do display anomalous bending
dynamics that reflects their more complex molecular struc-
ture (26) compared to actin filaments. Although the relax-
ation times of fluctuating actin filaments are consistent with
simple viscous drag, for microtubules they are much longer
than expected from hydrodynamic drag on short length
scales. This effect may be due to a surprisingly large internal
dissipation that dominates the relaxation times of microtu-
bules on biologically relevant length and time scales. Our
findings emphasize both the power and limitations of light
microscopy techniques for probing the complex mechanical
behavior of semiflexible polymers.
Filament tracking algorithm
The analysis algorithm used to track the shape of fluo-
rescently labeled biopolymers in successive images consists
of the steps detailed below. Briefly, an initial rough estimate
of the position of a filament is first determined by thresh-
olding the image. Pixels above threshold are then skeleton-
ized to obtain a 1-pixel-wide representation of the filament.
A polynomial fit to this skeleton is then used to walk along
the contour of the filament, refining the position estimate by
finding the intensity maximum along perpendicular cuts
across the filament.
Background noise reduction
We first performed an image noise reduction by convolving
each pixel of the image with a Gaussian kernel over a local
region of size w:
where Bðx;yÞ is an appropriate normalization. The long
wavelength background intensity variation is also
accounted for by averaging each pixel of the unfiltered
image over a local region of size w, to obtain Abackgroundðx;yÞ:
The final filtered image is then obtained from: Afilterðx;yÞ ¼
abs AGaussðx;yÞ ? Abackgroundðx;yÞ
localization, the size w was set to be around three times
larger than the width of the filament, ;5 pixels in our
system. For refinement of the initial filament localization, we
limit the convolution of local position information with
neighboring position information by setting the filter size to
approximately the width of the filament. An example of the
result of this procedure for a typical image of a fluorescently
labeled microtubule (Fig. 1 A) is shown in Fig. 1 B. Filters
designed specifically for highlighting linear structures are
another alternative (27).
??(20). For initial filament
Thresholding was done to separate the filament from the
remaining background intensity fluctuations. Pixels with a
value greater than the threshold value are assigned a value of
1, whereas those with a value less than the threshold value
are assigned a value of 0. The threshold value is given by
Biopolymer Bending Dynamics347
Biophysical Journal 93(1) 346–359
f ¼ ÆIijæ1msij; where ÆIijæ is the average intensity value of
all pixels ij, and sij is the standard deviation of the pixel
intensity, both of which are dominated by the nonfilament
background pixels. For high-frequency image acquisition,
the signal/noise ratio (s/n) is usually small; thus, it is critical
to determine an appropriate threshold value by determining
the number of standard deviations away from the average, m.
If m is too small, some background may be included, whereas
if it is large, the entire filament may not be included. We
typically start with a small value of m(;3) and successively
increase its value in small increments (0.2) until the backbone
of the thresholded region is well fit by a polynomial. An
example of thresholding is shown in Fig. 1 C.
For good s/n and an appropriate threshold value, pixels
above threshold will cover the fluorescent filament in a
cluster several pixels wide. This cluster can be thinned to a
1-pixel-wide line using a skeletonization routine that erodes
pixels while maintaining cluster connectedness (28). Skel-
etonization has the advantage that small clusters correspond-
ing to nonfilament background fluctuations above threshold
are typically eliminated by the erosion process. Persistent
fluorescent specks that are still not eliminated are ignored
using a simple routine to exclude objects above threshold
from specific regions of the image. If necessary, more
complex pixel clustering algorithms may be used to identify
pixels associated with the desired filament (28). An example
of skeletonization of a thresholded filament is shown in Fig.
1 D. This line, corresponding to a set of position coordinates
x ¼ i, y ¼ j such that Iij¼ 1; provides a first, imprecise,
estimate of the filament location.
Polynomial fitting and filament rotation
We refine the initial position estimate by analyzing the inten-
sity distribution of sections taken perpendicular to the local
slope of the filament (Fig. 2 A). To obtain the local slope,
we fit the xy coordinates of the skeleton with a polynomial
of order p. The polynomial degree required depends on the
amount of curvature in the filaments under study; we
typically use p ¼ 3–5 for microtubules and actin filaments.
We thus obtain a curve fðxÞ ¼ +p
ferentiated to obtain an estimate of the local slope of the
filament at position x0: uðx0Þ ¼ tan?1df
i¼0cixithat can be dif-
dxjx¼x0: To obtain the
localization algorithm for a typical fluorescence micrograph of an Alexa-488
dye-labeled microtubule constrained to move in a quasi-two-dimensional
chamber. (A) Unprocessed image, and image after bandpassing, as in Eq.
1 (B), followed by thresholding (C), and skeletonization (D). Note that the
spurious speck in the thresholded image is eliminated upon skeletonization.
Example of the initial image processing stages of the filament
intensity profiles along cuts perpendicular to the filament contour. (A)
Skeleton (see Fig. 1 D) in the xy coordinate frame with polynomial fit and
schematic perpendicular cuts. (B) The intensity profile along a perpendicular
cut of an unprocessed image is shown (solid squares). After bandpassing the
image, the intensity distribution (open squares) is fit to a Gaussian (solid
line). The filament location is taken to be the maximum of the Gaussian.
Example of filament position refinement by fitting Gaussian
348 Brangwynne et al.
Biophysical Journal 93(1) 346–359
intensity profile of the filament perpendicular to this local
slope, we rotate the original image by an angle uðx0Þ around
the position, ? xrot¼ ðxo;fðxoÞÞ; to orient the filament hor-
izontally. Successful image rotation requires the use of a
good interpolation algorithm that maps the intensity of all
pixels onto the rotated image without introducing artifacts.
Image rotation routines implementing good interpolation
algorithms accompany software packages such as IDL and
Matlab. The process is computationally intensive, and thus
we restrict image rotation to a local region of interest around
the filament with coordinates ½ðx0?bÞ:ðx01bÞ;ðfðx0Þ?bÞ:
ðfðx0Þ1bÞ?; where b is larger than the extent of the Gaussian
Because the skeletonization procedure erodes pixels at the
ends of the thresholded filament, we sample perpendicular
sections along the polynomial backbone from, typically, 4
pixels before to 4 pixels after the end points of the skeleton.
We calculate the integrated signal along the perpendicular
column of pixels k, M ¼ +kIk; and set a threshold value
(‘‘masscut’’) for identified filament positions. The first and
last filament positions with a signal mass above threshold
define the two ends of the filament. Because fluorescently
labeled biopolymers frequently display nonuniform labeling
along their length, masscut thresholding often leaves holes in
the position identification of the filament, particularly in
low-s/n images. We fill in these gaps by linear interpolation
between the nearest successfully sampled points.
Position refinement by Gaussian fitting
We proceed to analyze the intensity distribution of the
vertical column of pixels in each rotated region. An example
of such an intensity distribution is shown in Fig. 2 B (solid
squares). By applying the Gaussian convolution kernel and
background subtraction to this rotated local region, we
improve the s/n ratio by eliminating both high- and low-
frequency noise (Fig. 2 B, open squares). The center of the
filament can be identified from this intensity distribution
using either functional fitting or an intensity-weighted center
of mass. It is important to note that the Gaussian filter leads
to averaging of pixel intensities over a local region defined
by the size of the Gaussian kernel (;5–10 pixels); averaging
the filament centroid obtained from adjacent perpendicular
cuts of unbandpassed images could provide an alternative s/n
improvement, without undesirable averaging in the perpen-
dicular direction. We choose to fit the intensity distribution
to a Gaussian intensity profile (Fig. 2 B, solid line), which
yields the highest precision for tracking of spherical objects
(29). We find that the primary advantage of Gaussian fitting
over an intensity-weighted center-of-mass approach is its
relative insensitivity to any residual background signal.
The position of the center of the Gaussian, y9g; identifies
the position of the filament in the rotated frame. This center
position is then mapped back onto the unrotated full image
coordinate system to obtain the real-space position of the
mthsampled point ? xm; using ym¼fðx0Þ1f½y9g?fðx0Þ?3
cosðuðx0ÞÞg and xm¼x0?f½y9g?fðx0Þ?3sinðuðx0ÞÞg: This
procedure is repeated along the polynomial at the desired
sampling frequency, typically in steps of size Ds¼2 pixels,
until a full set of refined filament coordinates is obtained.
MATERIALS AND METHODS
Monomeric (G) actin was purified from rabbit skeletal muscle (30), with an
additional gel-column chromatography step (Sephacryl S-200) to remove
residual cross-linking and capping proteins. Actin filaments stabilized with
phalloidin were prepared by polymerizing G-actin in AB-buffer (25 mM
pH 7.4). Filaments were fluorescently labeled by using Alexa-488-modified
(Molecular Probes). We analyzed actin filaments with contour lengths
between 8 and 15 mm. Tubulin was purified from bovine brain according to
standard procedures and fluorescently labeled with Alexa-488. Microtubules
were polymerized in K-Pipes buffer (80 mM K-Pipes, 1 mM EGTA, and
1 mM MgCl2, pH 6.8) at 37?C and stabilized with 10 mM taxol.
Microtubules were imaged in a quasi-two-dimensional sample chamber
that was made by placing a small volume (typically 0.3 mL) of a dilute
solution of stabilized filaments augmented with a standard antioxidant
mixture (glucose oxidase, catalase, glucose, 2-mercaptoethanol) (31) to slow
photobleaching between a microscope slide and coverslip. The edges of
the coverslip were sealed with mineral oil to prevent fluid flow due to
evaporation. Adsorption of filaments onto the glass surfaces was avoided by
passivating the glass with adsorbed bovine serum albumin or covalently
attached poly-(ethylene glycol) chains (MPEG-silane-5000, Nektar, San
Carlos, CA). All experiments were performed at room temperature.
Fluorescent images of microtubules were acquired on a Leica DM-IRB
inverted microscope equipped with either a Hamamatsu intensified charge-
coupled device (CCD) camera C7190-21 (exposure time 33 ms, 0.136 mm/
pixel) or a Hamamatsu ORCA CCD camera C4742-95 (exposure time 59
ms, 0.128 mm/pixel) (Hamamatsu City, Japan). Images of actin filaments
and some microtubules were obtained using a Nikon inverted microscope
with Coolsnap HQ camera (exposure time of 5–100 ms, 0.129–0.194 mm/
pixel). Microtubules were also imaged with a 10-ms exposure using a high-
speed camera (Phantom V7, Vision Research, Wayne, NJ) equipped with an
image intensifier (Model VS4-1845HS, Video Scope International, Dulles,
VA). Some actin filament bending data (see Fig. 7) was obtained using a
secondmethod to slow down the filament dynamicsby adding nonadsorbing
polymer (2–3%, Dextran 500,000 g/mol). The polymer induces an attractive
depletion interaction between the coverglass and the filaments, leading to
confinement of the filaments close to the bottom surface. Using these
samples, we were able to acquire images with either epi-illumination or total
internal reflection fluorescence (TIRF) illumination. Due to the low levels of
background fluorescence,we were ableto decrease exposure time with TIRF
illumination to a few milliseconds.
The data in Fig. 11 B were fit to a line to obtain a slope of ?0.050 6
0.066, indicating no statistically significant correlation between persistence
length, lp; and wavevector, q. In addition, the correlation coefficient was
calculated usingr ¼ slpq=slpsq;whereslpqisthecovariancebetweenlpand
q, and slpand sqis the standard deviation of lpand q, respectively. The
values of r vary from 1 (strongly correlated) to ?1 (strongly anticorrelated).
We obtained a value of ?0.096, which is a statistically insignificant
correlation (p . 0.005).
Evaluation of tracking precision
As a first test to evaluate the performance of our algorithm
we determined the uncertainty in filament localization
Biopolymer Bending Dynamics349
Biophysical Journal 93(1) 346–359
arising from camera shot noise and tracking limitations. This
was accomplished by analyzing the shape fluctuations of
microtubules that were immobilized by physically adhering
them to poly-L-lysine coated surfaces. These microtubules
were locked tightly to the glass coverslip surface, so any
residual filament shape fluctuations that we measured would
be due to uncertainty in filament localization inherent in our
technique. To visualize the typical noise level, we overlay
the extracted shapes of an immobilized filament for 10
consecutive frames of a movie in Fig. 3. The insets show
higher-magnification views of two representative positions
along the filament against a pixel-sized grid. This explicitly
demonstrates that we obtained subpixel tracking accuracy
with a noise level of ;0.1–0.2 pixels (;20 nm). This degree
of accuracy is representative of the filaments we track in typ-
ical conditions, although, as we quantify below, the tracking
accuracy decreased with decreasing s/n. Since spherically
symmetric colloidal particles are much more straightforward
to track, and typically result in ;0.1-pixel accuracy, our fil-
ament tracking algorithm performed well, with a comparable
root mean-squared (RMS) noise level.
Bending rigidity of microtubules and actin
filaments from mode analysis
quasi-two-dimensional chambers. We obtain the filament
bending rigidities by analyzing the shape fluctuations using a
Fourier decomposition technique developed in previous stud-
ies (8,9). From the pixel coordinates ðxm;ymÞ of the filaments
we first calculate the tangent angle uðsÞ as a function of
arclength s, using: uðsmÞ ¼ tan?1ðym11? ymÞ= ðxm11? xmÞ;
ðxj11? xjÞ21ðyj11? yjÞ2
ðxm11? xmÞ21ðym11? ymÞ2
The tangent angle is then decomposed into a sum of cosines,
where the wave vector q is defined as q ¼ np=L; with n the
mode number and L the filament contour length (8). A cosine
expansion of the tangent angle using different normalization
prefactors has been used in other studies (9), but we find the
normalization used by Gittes et al. most natural since it re-
sults in a length-independent variance of the mode ampli-
tude. Use of this pure cosine mode decomposition assumes a
zero-curvature boundary condition at the free filament ends,
appropriate for freely fluctuating filaments.
The amplitudes, aq; of the first 24 bending modes of a
freely fluctuating microtubule are plotted as a function of
time in each of the subpanels of Fig. 4. The microtubule has a
nonzero intrinsic curvature, u0ðsÞ ¼
resulting in a nonzero mean amplitude. The variance of the
lower modes contains information about the flexural rigidity,
whereas the amplitude of the higher modes is dominated by
experimental noise. The amplitudes of these higher modes
widen with time (Fig. 4, inset) due to photobleaching and
concomitant reduction of the s/n.
A convenient feature of the cosine expansion in Eq. 2 is
that random noise due to errors in filament localization obeys
the following simple relation (8):
charged, poly-L-lysine-coated glass surface. (Insets) Higher-magnification
views of 10consecutivefilamentcontoursfrom a videoacquisition;the pixel
size is represented by the gray grid, demonstrating that the filament tracking
algorithm performs with subpixel accuracy.
Filament contours for a microtubule fixed to an oppositely
in Fig. 1, plotted as a function of time in each of the subpanels. The micro-
tubule has intrinsic curvature, resulting in a nonzero mean amplitude. The
variance of the lower modes contains information about the flexural rigidity,
whereas the amplitude of the higher modes is dominated by experimental
noise. The amplitudes widen with time (see inset) due to photobleaching and
concomitant reduction of the signal/noise ratio.
350Brangwynne et al.
Biophysical Journal 93(1) 346–359
Le211ðN ? 2Þsin2
2ðN ? 1Þ
ment’s intrinsic curvature, e2is the mean-squared error in
filament localization, and N is the number of position points
sampled along the filament. We tested the validity of Eq. 3
by performing a mode analysis on immobilized microtu-
bules. The variance in mode amplitude due to noise could be
well fit to the above expression, as shown in Fig. 5 A. From
this fit, we obtain a RMS error
agreement with the direct measurement of the noise floor
shown in Fig. 3.
later frames of a movie sequence of an immobilized micro-
algorithm to decreasing s/n caused by photobleaching. Fig. 5
B shows that, as expected, the variance increases with time,
reflecting the decreasing ability to precisely localize the
position of the filament as it photobleaches. However, even
qis the mode amplitude corresponding to the fila-
? 0:14 pixels, in good
after 900 frames, when the s/n has decreased by ;20%, the
error in pixel position is still only 0.25 pixels. With a high-
sensitivity CCD camera, the s/n and corresponding tracking
precision can be improved and maintained for many frames;
we have successfully tracked filaments for over 7000 frames.
We note that the error in filament localization introduces
an increase in the measured contour length of the filament,
DL, which is of order DL=L ? e2=Ds2(8). With the high
tracking accuracy of our algorithm, this error should be ,1%
and can thus be neglected. A more important source of error
is the fact that the ends of the filaments must be defined in
each successive image by setting a masscut value. However,
we find fluctuations in the measured length of the filament to
be of order dL ? 1 pixel. For filaments of 10 mm or longer,
this corresponds to an error in length of ,1%, which is
To investigate the time dependence of the mode ampli-
tudes, aq; we plot the mean-squared amplitudes of the first
four modes of a 30-mm-long microtubule as a function of lag
time in Fig. 6 A. The fluctuations in mode amplitude at a
given lag time display a Gaussian distribution, suggesting
that small intrinsic curvature on long length scales helps
avoid artifacts from filament rotation (8). The variance of the
distribution was therefore obtained by fitting a Gaussian to a
histogram of the data; direct calculation of the variance is
less precise due to occasional filament misidentification that
greatly exaggerates a direct calculation of the variance.
We first analyze the long-time limit, where the bending
mode amplitudes have decorrelated. To understand this limit,
we note that the bending energy U of a curved filament can
be expressed as an integral of its deviation in curvature from
the intrinsic curvature along the filament backbone (8,32):
where k ¼ kBT 3lp is the bending rigidity; this is known
as the worm-like chain Hamiltonian. By differentiation of
Eq. 2 and subsequent integration, one obtains: U ¼ ð1=2Þ3
saturating values of the mean-square difference of the
mode amplitude fluctuations are
q? aqÞ2: By the equipartition theorem, the
Æðaqðt1DtÞ ? aqðtÞÞ2æt;Dt?t¼2kBT
where kBis Boltzmann’s constant, T is the temperature, Dt is
the lag time between different images, and t is the relaxation
(correlation) time of the mode. The modes must be fully
decorrelated to accurately obtain the saturating variance,
requiring the long-time limit Dt ? t:
The saturation levels of the mean-square amplitudes
extracted from the data in Fig. 6 A, plotted as a function of
mode number, are shown in Fig. 6 B. Long-wavelength
fluctuations corresponding to the first few modes of typical
filaments are well above the noise floor and reflect curvature
amplitude plotted as a function of mode number n, for an individual
frame number (i.e., time) as the signal/noise ratio of the video images is
reduced due to photobleaching. Squares, frames 0–200; circles, frames 200–
frames 800–1000. Filaments can be tracked with subpixel accuracy for a
duration of typically 2000 frames.
(A) Variance of the Fourier coefficients aq of each mode
Biopolymer Bending Dynamics351
Biophysical Journal 93(1) 346–359
fluctutions due to thermal energy. The saturating amplitudes
of the first few modes scale with the expected q?2depen-
dence of Eq. 5 (Fig. 2 B, solid line). Higher modes cannot be
used if their relaxation timescale, t; is similar to or shorter
than the exposure time texp(5–100 ms in our studies). The
fluctuations of these modes are effectively blurred, causing
them to appear stiffer. We were able to extend the mea-
surable q range to higher q values by using short exposure
times, either with TIRF illumination (33,34), or by using a
high-speed camera equipped with an intensifier. As ex-
pected, for data acquired using short exposure times, the
regime displaying the expected q?2dependence is extended,
as demonstrated in Fig. 7.
We obtain the persistence length of filaments by fitting the
saturating mean-square amplitude versus q to Eq. 5. The fits
were extended over the range of modes for which no sys-
tematic deviation below q?2was observed, i.e., texp? t
(typically, the first three to five modes). From these fits, we
actin filaments, in good agreement with previous studies
(5,8,10). This persistence length is independent of the length
of the actin filaments within the observed range of 8–15 mm,
as shown in Fig. 8 A. In contrast, for microtubules, we find
a small apparent correlation between lpand length: short
filaments tend to appear softer, as shown in Fig. 8 B.
Microtubules with lengths between 25 and 66 mm have an
average persistence length of 2.8 6 1 mm (mean 6 SD),
similar to previously reported values (7,8,14) (Fig. 8 B).
Shorter filaments with lengths in the range 18–25 mm have a
lower average persistence length of 1.5 6 0.7 mm. The data
are thus in qualitative agreement with a recent report
suggesting that the internal protofilament structure of micro-
tubules leads to a bending rigidity that depends on total
well to the reported functional form of this dependence lp¼
as shown by the dashed line in Fig. 8 B. However, the close
proximity of the noise floor both in our measurements (Fig. 8
B, dotted line) and in those of Pampaloni et al. (15) suggest a
cautious interpretation of these results.
Þ2??1; with lN
p¼ 4:5 mm; and Lcrit¼ 30 mm;
Bending relaxation timescales
By using a Fourier decomposition of the filament shape,
we can readily determine mode relaxation times from the
autocorrelation function, as shown in Fig. 6 A. However,
these Fourier modes are only approximations of the true
normal modes of the dynamical equation of motion (see
Appendix). Although a Fourier decomposition is suitable for
determining the static behavior, as described above, Fourier
modes are not normal modes of the system. Instead, Fourier
modes will reflect the combined behavior of different normal
modes relaxing on different timescales. Nevertheless, the
the mean-square amplitude of Fourier
coefficients aqof the first four bending
modes of a 30-mm -long microtubule,
with fits to Eq. 6. (Inset) Note that data
scale together when plotted as a func-
tion of the scaled variables MSDðaqÞ=
plotted as a function of mode number n,
for increasing lag times Dt ¼ 0.059 s,
0.0177 s, and 5.9 s. At longer lag times,
few modes approaches the expected
q2-dependence of Eq. 6, with a persis-
tence length of 4 mm (decreasing solid
line). At higher mode numbers, the am-
plitudes are dominated by a RMS noise
of 0.15 pixel (increasing solid line).
(A) Time dependence of
ing modes of 12 actin filaments imaged using different ratios of exposure
time to mode relaxation time. Light gray squares were imaged with 100 ms
exposure in aqueous buffer, gray triangles were imaged with 100 ms ex-
posure time in a viscous background solution. Black circles were imaged
with 5–10 ms exposure time using TIRF illumination. The onset of deviation
from q2-scaling (black line, slope 1/lp¼ 17 mm) corresponds to wavevectors
for which the relaxation time roughly equals the exposure time; fluctuations
for larger wavevectors are blurred and register as smaller values.
Saturation values of the mean-square amplitudes of the bend-
352Brangwynne et al.
Biophysical Journal 93(1) 346–359
Fourier modes are a convenient basis set that completely
captures the instantaneous filament shape. It is therefore
possible to express the true normal-mode amplitudes in
terms of our measured amplitudes aqðtÞ; and thus to measure
the normal-mode relaxation rates. A more complete descrip-
tion of this procedure is provided in the Appendix.
Surprisingly, we found that over all experimentally
accessible timescales, mode-mixing effects arising from the
fact that these Fourier modes are not pure normal modes
were small. This can be understood as follows. By symme-
try, the odd (even) Fourier modes decompose into only odd
(even) normal modes Jl: Strictly speaking, this means that
the amplitudes in Eq. 5 for finite Dt exhibit non-single-
exponential relaxation. For the lowest Fourier modes, n ¼ 1,
2; however, only the corresponding l ¼ 1, 2 normal modes
are apparent at later stages of relaxation, since the higher-
order normal modes relax more quickly. A comparison
between the first Fourier mode and the corresponding normal
mode of a microtubule is shown in Fig. 9 A. The relaxation
times obtained from fits to a single exponential are very
similar: 0.84 s for the Fourier mode and 0.82 s for the normal
mode. For the higher Fourier modes, n ¼ 3, 4..., the
filament end effects are less apparent, and the normal modes
are better approximated by Fourier modes. However, as
discussed in the Appendix, since these modes mix with more
slowly decaying modes the non-single-exponential behavior
may be apparent at longer times in the relaxation. Neverthe-
less, these non-single-exponentialcorrections can be shown to
beoforder10–15%.Thus,inpractice,the dynamical behavior
of the cosine modes is well approximated by the single-
exponential relaxation of the normal modes:
Æðaqðt1DtÞ ? aqðtÞÞ2æt¼ ð1 ? e?Dt=tÞ2kBT
As shown in the appendix, the mode relaxation times t are
where q?? ðn11=2Þp=L: The hydrodynamic drag coeffi-
cient for a rod confined between two surfaces is approxi-
where h is the buffer viscosity, h is the distance between the
filament and the coverslips (;1 mm), and 2a is the filament
diameter (8,10). The mean-square fluctuation of the mode
amplitudes can indeed be well fit to Eq. 6 (Fig. 6 A, lines).
The collected data for the relaxation times t of 26
microtubules and 22 actin filaments, obtained from fits to Eq.
6, are shown in Fig. 9. For actin filaments (circles), the re-
laxation times decay rapidly with q?; consistent with the
?-dependence. We fit the data to Eq. 7; using the
average persistence length, lp¼ 17:0 mm; we obtain a drag
coefficient of gactin¼ 9:4 mPa?s, in reasonable agreement
with the calculated value of 1.8 mPa?s obtained from the
approximate hydrodynamic expression. The inset shows the
normalized relaxation timescales, tkq4
constant. This indicates that the bending relaxation timescale
for actin filaments is set by hydrodynamic drag. The relax-
ation timescales of microtubules show a similar behavior
(Fig. 9, squares) at the smallest wavevectors, with the re-
laxation times decreasing rapidly as t ? q?4
larger wavectors, the relaxation times are much larger than
expected and appear to become wavevector-independent.
Data taken with faster acquisition time also show this trend,
although some points remain consistent with a simple t ?
tubules is heterogeneous.
?; which are roughly
?: However, at
scaling, suggesting that the dynamic behavior of micro-
plotted as a function of filament length. (A) Actin filaments show a relatively
narrow distribution of persistence lengths with an average lp¼ 17:0 6 2:8
mm (dashed line). There does not appear to be any dependence on filament
length over the range studied. (B) Microtubules display a broader dis-
tribution of persistence lengths, and there is an apparent small correlation
between the bending rigidity and filament length, with shorter filaments
appearing softer. Our data can be fit to a length-dependent expression for
the persistence length, lp¼ lN
Lcrit¼ 30 mm; as shown by the dashed line (15). However, the dotted line,
denoting a conservative estimate of the intrinsic noise limitations, suggests
that this correlation may only reflect a noise limitation.
Persistence length of 22 actin filaments and 26 microtubules
ðÞ2??1; with lN
p¼ 4:5 mm; and
Biopolymer Bending Dynamics 353
Biophysical Journal 93(1) 346–359
For microtubules, a similar deviation from q?4
dence was recently reported for the relaxation times of
thermally fluctuating microtubules clamped at one end (7). A
possible explanation for the origin of these anomalous relaxa-
tion times comes from a recent study (16) that incorporates
an additional friction term in the Langevin equation (see
Appendix), describing the bending dynamics of thermally
This additional third term represents internal friction
within the biopolymer. The mode relaxation times (Eq. 7)
where g9=a4can be considered an effective internal viscosity.
Equation 9 implies that at wavevectors larger than qc;ðg=
g9Þ1=4; internal friction will dominate and the relaxation times
correspondingly become q?-independent. The microtubule re-
the average persistence length, lp¼ 2.5 mm, we obtain a drag
coefficient of gMT¼ 8:3 mPa?s, in fair agreement with the
value of 2.4 mPa?s estimated from the approximate hydrody-
namic expression. From the fit, we also obtain an internal
dissipationofg9 ¼ 16310?25N ? m2? s,correspondingtoan
internal viscosity of g9=a4
value is similar to that obtained in the previous study,
g9 ¼ 6:9310?25N ? m2? s (7) (fit shown in Fig. 9 B, inset).
Thus, in both cases, microtubules appear to cross over to a
wavevector-independent relaxation regime at qc? 0:3mm?1;
on a length scale corresponding to l ? 20 mm.
MT? 107mPa?s. Interestingly, this
Microtubules in cells
Our tracking algorithm can also be extended to extract the
contours of fluorescently labeled microtubules within living
cells. Because the cytoskeleton of living cells is typically
packedwitha dense networkofmicrotubuleswitha meshsize
of ;1 mm, initial filament localization using a thresholding/
skeletonization routine is insufficient. However, with a graph-
of the filament position, it is possible to implement our simple
algorithm with minor modifications. Alternatively, the initial
filament localization may be accomplished with automated
edge-detection routines (21). A modification for handling
filament intersections must also be added to our algorithm for
profile of the filament cross-section. In this way, filament
intersections, for which the intensity profile is poorly fit to a
Gaussian, can be easily distinguished from nonoverlapping
segments of the filament. Such filament positions can then be
interpolated between neighboring sampled positions. In Fig.
10, we demonstrate that we can track a single microtubule
through the dense network in a fixed tissue-culture cell using
such an approach (4).
We have developed and characterized a simple but robust
algorithm for precisely tracking fluorescent filaments. To
study biopolymer bending dynamics using this algorithm,
we took care to identify and characterize the main sources of
error that arise in these measurements.
1. Intrinsic noise makes the bending fluctuations appear
larger and therefore makes the filament look softer.
2. Correlated fluctuations appear to have a smaller variance
than uncorrelated fluctuations evaluated at longer times,
and thus cause error since the filament appears stiffer.
3. Long exposure times relative to relaxation time can cause
the filament to blur, making the fluctuations appear
smaller and, thus, the filament appear stiffer.
4. Improper mode decomposition can lead to mode-mixing
effects that hinder interpretation of the dynamic behavior
the decorrelation of the first Fourier
mode (green squares) and the corre-
sponding normal mode (blue circles)
of the microtubule shown in Fig. 6 A.
The normal mode was obtained from a
sum of odd Fourier modes, as described
in the Appendix. The relaxation time
obtained from a fit to a single expo-
nential is 0.84 s for the Fourier mode
and 0.82 s for the normal mode. (B)
Relaxation times of bending modes of
actin filaments (circles) and microtu-
bules (squares) plotted as a function of
wavevector q?: Solid symbols represent
dependence (solid line) expected from simple
?). The inset shows the
(A) Comparison between
data obtained using short exposure times (5–10 ms). The data for actin filaments are consistent with a q?4
hydrodynamic drag(Eq. 7). Formicrotubules,we finda deviationfromq?4
same data normalized as tkq4
?plotted versus q?: Black crosses are data for microtubules from Janson and Dogterom (7).
?at q?. 0:3mm?1(solid line,fit to Eq.9; dottedline, fit to q?4
354 Brangwynne et al.
Biophysical Journal 93(1) 346–359
Each of these sources of error can affect measurements of
bending rigidity and/or relaxation time, and need to be
Filament bending rigidity
Using our precise tracking algorithm, we determine the
persistence length and relaxation timescales of thermally
fluctuating actin filaments and microtubules. Consistent with
previous studies, we find that microtubules are two orders of
microtubules show a wavevector dependence of their bending
chainmodel.Actinfilaments havea persistence lengthof;17
mm, independent of filament length. Interestingly, for micro-
short microtubules appearing softer than long ones.
The persistence lengths plotted in Fig. 8, A and B, are
essentially averages over multiple q-values, since they are
obtained by fitting multiple bending modes to Eq. 5 (see Fig.
6 B). Thus, the apparent length dependence of the bending
rigidity of microtubules in Fig. 8 B could actually reflect a
q-dependence. To test this, we plot the persistence length
obtained from the variance of each mode as a function of the
corresponding wavevector, as shown in Fig. 11 A. Although
there does not appear to be a strong correlation at small
wavevector, high-wavevector fluctuations indeed appear to
be softer. These soft high-wavevector modes would tend to
make shorter filaments appear softer, since their relative
contribution to the average stiffness (obtained from a fit of
multiple modes to Eq. 5) would be larger. One possible
explanation for this wavevector dependence is that at
sufficiently high wavevectors, mode amplitude fluctuations
due to filament tracking noise dominate over thermal
bending fluctuations. This is unlikely, however, since modes
in the noise regime obey the noise relationship (Eq. 3) that is
easily identifiable, as illustrated in Fig. 6 B. Moreover, we
only considered modes that clearly decorrelated with lag
time, which is never true for modes in the noise regime.
Thus, the high wavevector fluctuations we measure are not
directly affected by intrinsic noise fluctuations.
Although intrinsic noise does not directly contribute to the
measured fluctuations, it is possible that noise still indirectly
affects the measurements, since it introduces a bias: high
wavevector fluctuations can be detected above the noise floor
only if the filament is sufficiently soft. This effect is likely to
arise for microtubules in particular because of the large
filament-to-filament variability of the persistence length.
Indeed, using conservative values for the noise floor (taking
an RMS noise of 0.5 pixel), an estimate of the location of this
bias agrees with the trend in Fig. 11 A (solid line). Further
support for this comes from the fact that within individual
filaments there is no detectable wavevector dependence of
the bending rigidity, as can be seen in the rescaled data in
Fig. 11 B. As expected from the plot, linear regression shows
no statistically significant correlation. Interestingly, although
our data are consistent with the findings of Pampaloni et al.
(15), intrinsic noise biasing likely contributes to the small
correlation we find between total filament lengthand rigidity,
as shown by the corresponding noise boundary in Fig. 8 B.
Indeed, such artifacts introduced by noise limitations could
has been fixed and stained, showing microtubules near the cell edge. We
track one microtubule by combining the automated filament tracking
algorithm with a manual initial filament identification. On the right is the
fluorescence image with the filament coordinates overlaid in red.
Fluorescence image of a monkey kidney epithelial cell that
to a bias caused by intrinsic noise
limitations. The dotted line denoting
the noise-limited regime was obtained
using conservative values for the noise
floor (RMS error of 0.5 pixel) in Eq. 3.
(B) Within individual filaments, there is
no dependence of the persistence length
obtained from each mode (normalized
by the persistence length averaged over
all modes) on wavevector (normalized
by the wavevector averaged over all
modes). Each filament is shown with a
Biopolymer Bending Dynamics355
Biophysical Journal 93(1) 346–359
also contribute to the bending rigidity/length correlation
found in this recent study. Further experimental work in-
corporating a careful analysis of the noise will be required to
determine whether there is indeed a significant dependence
of the rigidity on microtubule length, and to elucidate the
precise origin of this potentially interesting finding (15).
Bending relaxation timescales
By studying the time evolution of the mode amplitudes, we
found that actin filaments display relatively simple bending
dynamics, whereas microtubule bending reflects a more
complex mechanical behavior. The relaxation times of actin
filaments are consistent with the q?4
dence expected from hydrodynamic drag. Microtubules,
however, appear to cross over from a hydrodynamically-
dominated relaxation regime at long length scales, l.20mm
to a wavevector-independent relaxation regime at shorter
length scales, l,20mm:
These anomalous relaxation timescales do not appear to be
due to any of the sources of measurement error we identified.
In particular, if high-wavevector relaxation timescales were
affected by the exposure time of the camera, then the
relaxation timescales would become smaller and not larger,
as we have observed. We addressed the possibility of ex-
posure time effects by obtaining data using much shorter
exposure times (Fig. 9 B, solid symbols). These data show
a similar trend, although some points remain consistent with
a simple q?4
scale data show a very slight deviation from a perfect
?-dependence possibly due to measurement errors, for
microtubules the marked spread in the data showing devi-
ation from the expected q?4
?-dependence does not appear to
result from measurement error, and highlights the heteroge-
neity of the dynamic behavior of microtubules.
As discussed above, it was recently suggested that
structural changes in the form of interprotofilament shearing
lead to a filament-length-dependent stiffness (15). Interest-
ingly, the model described there would seem to lead to a
reduced effective stiffness, and therefore reduced relaxation
rate for short wavelength modes. However, we find no
evidence for a true wavevector-dependent rigidity (Fig. 11).
Moreover, the strong trend in our microtubule relaxation
time data could not be explained by the small stiffness bias
we observe, since the normalized relaxation times (Fig. 9,
inset) remain consistent with a wavevector-independent
relaxation regime. The anomalous microtubule relaxation
times thus appear to reflect some form of viscous dissipation
within a heterogeneous population of microtubules.
To estimate the dissipative parameter g9 in Eq. 8, we adopt
a simple physical picture for the microtubules (16). For an
elastic filament composed of a porous or viscoelastic gel-like
material, internal dissipation results from the flow of a fluid
of viscosity h9 through pores of size j: This leads to
g9 ¼ h9a6=j2; where 2a is the filament diameter. Wave-
scaling. Although the actin relaxation time-
vector-independent relaxation times for bulky mitotic chro-
mosomes appear to be described by such a picture (16).
Because the diameter of microtubules is ;25 nm, whereas
actin filaments are a more slender 7 nm, similar internal
dissipation should be more apparent for microtubules; this is
qualitatively consistent with our findings. However, the
high-frequency rheology of actinsolutions (35–37) shows no
evidence of deviations from pure hydrodynamic drag up to
frequencies as high as 100 kHz, corresponding to relaxation
times up to four orders of magnitude smaller than for
microtubules in this study. This fact cannot be explained by
the difference between microtubule and actin diameters
alone. Specifically, the bending moduli of both microtubules
and actin are consistent with the simple prediction k ¼ Ea4
of elasticity theory, where the Young’s modulus E ? 1 GPa
for both. Thus, the model above for porous filaments would
result in a relaxation rate of order h9ða=jÞ2=E: It seems more
likely that anomalously large values of g9 for microtubules,
as compared with actin, are due to slow structural fluctua-
tions or conformational changes (16).
We have demonstrated that bending fluctuations of fluo-
rescently-labeled biopolymers can be tracked with high
spatial and temporal resolution using a simple and robust
automated subpixel tracking algorithm. Using this algorithm,
we confirmed previous studies reporting a persistence length
of actin filaments of ;17 mm, and microtubule persistence
lengths on the order of a few millimeters. For microtubules,
we found that a correlation between persistence length and
filament length can arise due to biasing from intrinsic noise
limitations. By studying the time evolution of actin bending
fluctuations, we found that their relaxation times display a
appear to transition into a high-wavevector regime domi-
nated by surprisingly large internal viscous dissipation.
These results emphasize the heterogeneous mechanical be-
havior of microtubules, and suggest that microtubules
display anomalous bending dynamics that reflect their com-
plex molecular structure.
hydrodynamic scaling, whereas some microtubules
To understand the time dependence of the mode amplitude fluctuations (Fig.
6 A), we consider the following Langevin equation for changes in the
transverse position u(s,t) of the filament as a function of arclength, s, and
time, t (5,8,10,38):
The first term accounts for the elastic restoring force prescribed by Eq. 4, the
second term is the hydrodynamic drag on the filament as it moves through
the viscous buffer, and the third term is the d-function correlated random
thermal noise acting along the filament.
356 Brangwynne et al.
Biophysical Journal 93(1) 346–359
Following Aragon and Pecora (38), we find solutions to the equation of
motion (Eq. A1) in terms of normal modes jlðsÞ; which are solutions to the
with the boundary conditions corresponding to no net forces or torques on
Here, the eigenvalues are given by ll¼ kðal=LÞ4; where cosðalÞcoshðalÞ ¼
1: To within better than 0.4%, the solutions of the latter are approximately
given by alffi ðl 1 1=2Þp for l $ 1; where this approximation becomes in-
creasingly accurate for the higher modes. Thus, even though the al will
appear raised to the power of four in the expressions below, we will make
no more than ;1.5% error using the approximation above.
The solutions to Eq. A2 are orthogonal. Thus, we can construct an
orthonormal basis for functions on ð?L=2;L=2Þ; given by
For l ¼ 1;3;5...
For l ¼ 2;4;6...
The solutions we seek can be expressed as
uðs;tÞ ¼ +
The Langevin equation can also be written now in terms of the amplitudes
The solutions to this are of the form
JlðtÞ ¼ Jlð0Þe?vlt1e?vlt
for t $ 0: This allows us to calculate the correlation functions
ÆJlðtÞJmð0Þæ ¼ ÆJlð0ÞJmð0Þæe?vlt;
since ÆflðtÞJmð0Þæ ¼ 0: However, given the time translation invariance of
this result, as well as the nondegenerate spectrum of relaxation rates
vl¼ ll=g; this only makes sense if ÆJlðtÞJmð0Þæ ¼ 0 for all l 6¼ m: This
can also be seen from the fact that the bending energy
which results in Gaussian distributed amplitudes
Here, we have used partial integration for the bending energy, together with
the force- and torque-free boundary conditions.
Of course, we could have also used the more familiar Fourier modes
but only for the statics. These are not normal modes for the dynamics. The
Fourier amplitudes BnðtÞ ¼Rbnðs9Þuðs9;tÞds9 now satisfy
where this is an equal-time correlation function and fn¼ kðpn=LÞ4:
We note that the last two terms of Eq. A1 both depend on the local
position/velocity of the polymer. As these variables are nonlocal in the
tangent angle variables, it is not possible to write this Langevin equation in
terms of u(s). However, for weakly bending filaments, the local slope of the
filament uðs;tÞ ¼ duðs;tÞ=ds can also be used to describe the shape. This
shape can be further described by the Fourier amplitudes above:
In terms of the normal modes, the amplitudes anðtÞ can be expressed as
anðtÞ ¼ +
We define the matrix
Thus, the correlation of amplitudes anðtÞ can be found in terms of the normal
ÆanðtÞanð0Þæ ¼ +
We note that the functions gnðsÞ are alternately odd and even, whereas the
functions jlðsÞ are alternately even and odd. Thus, Mnl¼ 0; unless n and l
are either both odd or both even. For example, LM1l¼ 7.05, 0, 5.69, 0, 5.66,
0, for l ¼ 1, 2, ... 6, whereas LM3l¼ ?0.377, 0, 12.29, 0, 6.21, 0, for l ¼ 1,
2, ... 6.
Alternatively, the matrix inverse M?1can be used to express the normal
mode amplitudes Jlin terms of the measured amplitudes an:
JlðtÞ ¼ +
ÆDJlðtÞ2æ [ Æ½JlðtÞ ? Jlð0Þ?2æ ¼ +
llð1 ? e?vltÞ:
Since the matrices describing the linear transformations between Jland an
are easily evaluated, this procedure provides a practical way to determine the
normal-mode relaxation spectrum in terms of the anthat can be measured as
Biopolymer Bending Dynamics357
Biophysical Journal 93(1) 346–359
For large l, the dominant term in this sum is for m ¼ l, since the effect of
the ends of the filament become negligible in this limit. Thus, we expect that
for large enough n, the relaxation of anis approximately single-exponential
with relaxation rate
Attheother endof the spectrum,for n¼1, 2,the relaxationof anwill also be
approximately single-exponential with relaxation rate vn for long times,
since the most slowly relaxing mode in Eq. A17 is for l ¼ n when n ¼ 1, 2.
The corrections to this approximate single-exponential behavior, due
to modes l ¼ n 1 2, both have amplitudes smaller by approximately
ln=ln12¼ ½ðn11=2Þ=ðn15=2Þ?4and relax substantially faster (specifi-
cally, by a factor of ln12=ln). Using the matrix entries Mnlabove, we find
that the second term in Eq. A17 is smaller than the first by a factor of 0.022
(0.036) for n ¼ 1 (2). Perhaps surprisingly, even though the n ¼ 3, 4 modes
mix with the slowly relaxing l ¼ 1, 2 normal modes in Eq. A17, using the
Mnlabove, we find the relative contribution of these slow modes to be 0.027
and 0.048, respectively. Including other subdominant terms in Eq. A17, we
explains why we find little evidence, in practice, for non-single-exponential
relaxation for any of the modes we examined. We have directly implemented
the procedure described above for several data sets for n ¼ 1–6 and find that
the correction is within our error bars. However, accounting for the correct
normal mode relaxation using Eq. A20 is essential for interpretation of the
dynamics of both microtubules and F-actin in our experiments.
We thank T. Mitchison and Z. Perlman for their kind donation of tubulin
and their assistance with fluorescent labeling. We also thank L. Mahadevan
for helpful discussions.
This work was supported by the National Science Foundation (DMR-
0602684 and CTS-0505929), the Harvard Materials Research Science and
Engineering Center (DMR-0213805), the Harvard Integrative Graduate
Education and Research Traineeship on Biomechanics (DGE-0221682),
and the Stichting voor Fundamenteel Onderzoek der Materie/Nederlandse
Organisatie voor Wetenschappelijk Onderzoek. G.H.K. is supported by a
European Marie Curie Fellowship (FP6-2002-Mobility-6B, Contract No.
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