Microtubule Motility Assays
As the pivotal role of microtubule (MT) motors in
the cell cycle becomes more widely recognized, so the
demand for motility assays will increase. This article
presents a robust and straightforward set of protocols
based on experience gained in our laboratory. Readers
seeking to set up more specialised assays should refer
to one of the excellent methodological compendia that
are available (Inoue and Spring, 1997; Cross and
Kendrick Jones, 1991; Scholey, 1993).
II. MATERIALS AND
Unstained MTs are visualized most conveniently by
video microscopy using computer-enhanced differential
interference contrast (DIC). MTs assembled from
fluorescently labeled tubulin can be visualized using
epifluorescence. This article concentrates on these two
contrast modes. It is also possible to visualise microtubules
by dark field, which gives a high contrast
image of unstained microtubules, or by phase contrast,
but both modes are susceptible to dirt in the solutions
and so may be inconvenient for routine work. Interference
reflection produces higher contrast than DIC
and gives information in the Z direction, but again is
susceptible to dirt.
For all modes of contrast the main requirement of
the microscope is that it should be as simple as possible.
It should also be big and heavy to give it some
vibration resistance. Uprights are less expensive, optically
straightforward, and more convenient if the
bathing solution is to be exchanged during observation
(which is often the case). Inverted scopes are more
stable and provide much better access to the specimen
if other inputs (micromanipulators, flash-photolytic
lasers, evanescence prisms) are envisioned.
Human eyes work in colour and have a higher
dynamic range, better spatial resolution, and a bigger
view field than a video camera. It is very useful to be
able to use them to find focus. To do this the microscope
needs to incorporate some sort of device to
switch the light between the eyepieces and the camera.
One hundred Watt Hg lamps generate large
amounts of heat and it is necessary to remove this heat
from the illuminating light. Glass heat filters may be
used, but the best way to filter out heat is to use a hot
mirror, a mirror that transmits infrared but reflects
visible light (technical video).
DIC optics are optimised for visible wavelengths,
and it is conventional to use a green interference filter
to give narrow band illumination. In practice the
improvement this brings is often too slight to be
obvious by eye, and removing the green filter can be
a convenient way to brighter illumination. The UV
emitted by Hg lamps is removed substantially removed
by optical (lead) glass, but for DIC an in-line
UV filter is nonetheless a sensible precaution.
A useful improvement in both fluorescence and DIC
image quality can be achieved by sending the illuminating
light through a fibre-optic light scrambler
(Technical Video). Light from the lamp is focussed into
one end of the fibre (Fig. 1), and the other end emits a uniform disc of light (of Gaussian intensity profile),
which is used to illuminate the microscope.
3. Optical Train
For maximum image quality in all contrast modes,
it is advisable to reduce the number of optical components
between the objective and the camera to a
minimum. The most critical component is the objective.
The light intensity transmitted by a lens is proportional
to the square of its numerical aperture (NA),
whilst resolution rises linearly with the NA. It is
important therefore to use a 1.4 NA (therefore oil
immersion) 60 or 100× planapo objective in order to
maximise light-gathering power and resolution, particularly
so in epifluorescence, where the objective
doubles as the condensor.
We use a novel configuration that replaces the condensor
on a Zeiss Axiovert with a home-built fitting
that improves light collection from the Zeiss tungsten
lamp and mounts a second objective in place of the
condensor. This arrangement (Fig. 1) gives good MT
DIC with a field about 40µm and brings the considerable
advantages of the tungsten lamp, which is much
more stable than the Hg lamp. For smaller view fields,
the higher radiant intensity of the Hg lamp is
4. Antivibration Hardware
|FIGURE 1 Video microscope. An inverted microscope set up for both fluorescence and DIC microscopy.
Hg, mercury lamp; HM, hot mirror; OF, optical fibre; CCD charge-coupled device camera (for DIC); ICCD,
intensified CCD (for fluorescence); DIP, digital image processor; PC, personal computer; VCR, video cassette
Vibration will degrade the highly magnified image.
Low frequencies (people walking across the floor) will
cause the image to bounce around, whilst high frequencies
will be averaged out by the video framing
rate and cause the image to blur. The extent of this
problem is, however, often overemphasized. Before
purchasing an expensive and awkward vibrationdamping
equipment, try placing a few layers of bubble
wrap under the microscope baseplate.
5. Temperature Control Hardware
Motility rates for several motors are extremely temperature
sensitive in the range of room temperature,
and temperature control is consequently important if measurements taken at different times are to be compared.
The best way is to temperature clamp the entire
microscope. If you have air conditioning this will
already be happening. To warm specimens above
ambient temperature, we use a homemade plexiglass
box with a warm-air blower coupled to the side.
Another option is to use water jackets for the objective
and stage. These are readily fashioned by wrapping
flexible, narrow-bore copper tubing around several
times and connecting the ends to a water bath with a
circulator (e.g., Techne) using silicon tubing. If cooling
is necessary, we find it useful to wrap the microscope
in a tent made of cling film, reducing condensation.
For fluorescence of moving objects, two types of
low-light level / high-contrast / high-framing rate
cameras are suitable: intensified charge-coupled
device (ICCD) and intensified silicon-intensified tube
(ISIT) cameras. ICCD cameras are better for current
purposes because ISIT cameras, although more sensitive,
introduce spatial and intensity distortions across
the view field that are tedious to correct for. All the
aforementioned produce an analogue video signal,
e.g., ISIT Hamamatsu C2400-08 and ICCD Hamamatsu
C2400-97E. An alternative approach that is becoming
feasible uses a digital camera to record direct to computer
hard disc in time lapse. A cooled CCD camera
coupled to a generation 4 intensifier gives excellent
sensitivity for fluorescence work, but is limited by the
quantum efficiency of the intensifier (about 30%) and
is currently very expensive and the digital data can be
awkward to archive. If considering this route, be sure
to test the software, which in our experience can place
more severe limits on performance than the specifications
of the hardware.
For DIC, a nonintensified scientific grade CCD
is fine, e.g., grey-scale CCD camera Hamamatsu
7. Camera Coupling/Magnification
The ideal magnification sets four or more camera
pixels across the width of the MT. For a 512 × 512 pixels
(2/3in.) CCD, this corresponds to a square field with
sides of 20-25µm. Zoom couplings are wonderfully
convenient but are not always a good idea because
they absorb a lot of light. Magnification must be calibrated
using a stage micrometer.
8. Image Processor
CCD cameras are often offered with a hardware box
providing real time analogue enhancement of the
video signal (any or all of gain, back-off and shading
correction). A digital video processor is better, which
is able to digitise incoming video frames, perform
frame averaging, contrast enhancement, background
subtraction, and caption overlay and then re-encode
the image as an analogue video signal, all in real time.
The Hamamatsu Argus 20 is so well thought out
that it is virtually standard equipment for video
microscopy laboratories. The best and most flexible
arrangement is to adjust the gain and backoff on an
Argus 20 or similar processor controls such that there
are no areas in the image that are completely saturated
and then apply further digital enhancement. The
resulting signal is displayed on a monitor and is fed to
the PC for direct grabbing of video clips and to the
VCR for archiving.
It is worth investing in a high-quality 14-in. multiformat
monitor. Larger monitors look impressive but
are only helpful if they have to be placed a long distance
from the operator.
10. Video Recording
At the time of writing, the most convenient and
practical way to store large amounts of video is still to
use video tape. Recording to VCRs inevitably involves
some degradation of the image (loss of spatial resolution,
noise, contrast effects). For practical purposes the
resolution loss is potentially the most serious problem.
The effective resolution following recording can be
visualised by recording and replaying a test card
image having black and white lines at various spatial
frequencies. Currently the best option is digital recording
to video tape. Digital video tape recorders input
and output an analogue signal, but encode data digitally
to tape. The digitization involves some compression
of the incoming video signal, but most of the
compression is on the chrominance rather than the
luminance, with the result that spatial information is
relatively well preserved, particularly for grey-scale
signals. Unlike other compression schemes such as
DVD, there is no compression along the temporal axis.
There are currently two sizes of tape: digital video
(DV) and mini-DV. There are also two different
formats: DVCAM (Sony) and DVCPRO (Panasonic).
These two formats use the same compression scheme
but DVCPRO spaces the tracks further apart on the
tape, giving (arguably) better reliability and accuracy
for editing, but with correspondingly less recording
time on the tapes. Digital video recorders tend to be
built for the pro market and are more robustly engineered
than consumer machines, but are also more
We have evolved a video-recording strategy that
offers maximum flexibility: Time-lapse digital recording of grabbed video frames to computer hard
disc (with no resolution loss) and simultaneous realtime
recording to digital VCR. The VCR runs uninterrupted
in the background and generates an archive.
The operator is free to go back to this archive at a later
date and transfer interesting sequences to the computer
for analysis. Captured digital sequences can be
archived to external hard discs. We now have a fast
digital capture and analysis application RETRAC II,
which allows batch digitisation of video clips from
tape for subsequent analysis. RETRAC II can be downloaded
from our website (http://mc11.mcri.ac.uk/
Retrac / index.html).
Analogue video recording is still in widespread use.
Different video formats unfortunately operate in different
countries. In the United States and Japan, NTSC
format applies (525 lines per frame; 30 frames/s, typically
captured at 640 × 480 pixels). European countries
use PAL, which has higher spatial resolution but lower
time resolution (625 lines per frame, 25 frames per sec,
typically captured at 768 × 576 pixels).
The most convenient way to analyse motility is to
capture a sequence of frames into computer memory
and to track objects using a mouse-driven cursor. It
helps to have a hard disc big enough to hold 2 day's
work (more is dangerous because of the temptation not
to back up) and enough hard memory to hold the stack
of captured frames. A typical 20 frame stack uses about
8 Mb of application memory; if you want to use larger
stacks then you need more memory. Processed stacks
can conveniently be archived to removable discs. We
use 100-Mb zip discs or 230-Mb magneto-optical discs.
CD writers are getting less expensive and are worth
considering if a permanent archive is required. Video
compression protocols (JPEG, MPEG) are best avoided,
as all involve some data loss. That said, Quicktime has
become a standard for digital video and can use a
variety of compressors, some of which are lossless.
12. Frame Grabber Card
Large numbers of video grabbing cards are available.
Only a few are supported by Retrac and NIH
Image, the freeware software packages that are recommend
later. Because the situation is fluid, please
check the software documentation for a list of supported
On the Mac, the best route for analysis is NIH Image,
which can be customised using macros to track objects
and output data in spreadsheet-compatible format.
Macros for basic tracking through NIH Image stacks are
available for download from our web page http://
mc11.mcri.ac.uk/retrac.html. NIH Image runs on
system 9 macs, with support for the Scion LG3 frame
capture board, or in classic in OSX but without the Scion
support. Wayne Rasband is continuing development of
ImageJ, an NIH imageminspired java programme that
runs in OSX and currently has partial support for the
LG3 board (http://rsb.info.nih.gov/ij/). NIH Image
and ImageJ are both available for the PC.
RETRAC 2 for Windows is purpose-written for the
analysis of motility assay data. The latest version supports
time-lapse frame grabbing from either VCR or
live video, autofocus, autocontrast, tracking (including
drift correction) spatial filtration, and magnification.
The programme now incorporates a powerful file
manager. Figure 2 shows a screenshot during tracking.
|FIGURE 2 A screenshot from RETRAC 2.
The type of slide used does not matter. The type of
coverslip does. The thickness of the coverslip should
be matched to the objective. The objective will
be marked appropriately [e.g., 60/planapo DIC 1.4
0.17/160 means a 60× objective selected as strain free
for DIC, aplanatic (flat field); apochromatic (low chromatic
aberration for blue yellow and green); optimised
for cover glasses 0.17mm thick and with a 160-mm
focal length]. We use Chance 22 × 22-mm No.1.5 coverslips.
In the past we have used these without any
special cleaning treatment and rejected "bad" batches
of coverslips that show poor binding of motor and/or
a poor image because of surface contamination. This
is still a workable approach, but we have begun to use
a cleaning procedure that appears effective in removing
contamination and making the coverslip reproducibly
hydrophilic, as evidenced by the spreading of
a drop of buffer placed on the surface so that it wets
the entire surface. This coverslip cleaning procedure is
based heavily on that given on the Technical Video
CCP.html). Our localized variant is on our website
A. Taxol-Stabilised Microtubules
- 1M K-PIPES: PIPES dissolves around its isoelectric
point of about pH 6.5. Take 500ml water, add 65 g
solid KOH, and then, after cooling if necessary, slowly
add 302g PIPES buffer (Sigma P-6757). Once everything
is dissolved, monitor pH and roughly adjust by adding more KOH pellets as necessary. Allow the
warm solution to cool and then fine-adjust pH using
5M KOH. Be careful not to overshoot, as there is no
- 100 mM NaGTP stock solution: Because nucleoside
triphosphates such as GTP and ATP undergo rapid
hydrolysis at acidic pH, efforts should be made to
control pH when dissolving and storing them. Dissolve
1g NaGTP (Sigma G8877) in 15ml 10mM Na-
PIPES, pH 6.9, monitoring pH. Rapidly reneutralise
pH by titrating in 5M KOH. Fine adjust pH and then
make volume up to 19.11ml. Store frozen at -20°C in
aliquots of 5-2000µl. Do not add MgCl2 to the stock
solution (it precipitates).
- 100 mM MgATP stock solution: Dissolve 5.87g
NaATP (Sigma A7699 ATP ultra or Boehringer 519 987)
in 60ml 10mM K-PIPES, pH 6.9, monitoring pH continuously
and holding as close as possible to neutral
using concentrated KOH. Once the ATP is dissolved,
add 10ml of 1M MgCl2 and readjust pH to 6.9. Adjust
volume to 100.0ml and freeze in aliquots of 5-5000µl.
- Taxol stock solution: Wear gloves and work in the
fume hood. Inject 2.93ml anhydrous dimethyl sulfoxide
(DMSO, Aldrich 27685-5) into a 25-mg bottle of
taxol (Sigma T 7402). Dissolve by vortexing and store
as 2- to 20-µl aliquots at -20°C. Taxol is stable in DMSO
but unstable in water. It is insoluble in aqueous buffers
above about 18 µM. DMSO is explosive if it gets wet.
Store small volumes at room temperature over beds of
- 0.2M NaEGTA: Dissolve 15.2g EGTA (Sigma
E 4378) in 190ml water. Adjust pH to neutral by adding
concentrated NaOH and then make volume to
200.0ml. Store at room temperature.
- 1M MgCl2: 20.33 g MgCl2·6H2O to 100 ml water.
Sterile filter and store at room temperature.
- BRB 80 (Brinkley reassembly buffer): 80mM KPIPES,
1 mM MgCl2, 1 mM EGTA, pH 6.9. Make up as
a 10× stock, store at 4°C and dilute freshly for use.
Purified tubulin at about 100µM (protocol for tubulin
preparation on our web page) in BRB80 should be flash
frozen in 10- to 25-µl aliquots in the presence of 30%
glycerol by immersion in liquid nitrogen and stored
either at -70°C or preferably in liquid nitrogen.
B. Preparation of Flow Cells
- Thaw an aliquot of tubulin (typically 200µM) and
add stock 100mM NaGTP to 1 mM and MgCl2 to
2mM. Warm to 37°C and incubate for 20min.
- After 20min, add taxol from a 10mM stock in
DMSO to 20 µM final. Dilute microtubules 1000-fold
for use using BRB80 buffer supplemented with
20 µM taxol.
C. Surface Adsorption of Motor
- Apply single-sided Scotch tape to the long edges
of a microscope slide such that the strip of glass surface
between the two pieces of tape is 8-10 mm wide. Trim
away overhangs with a razor blade.
- Extrude two parallel stripes of Apiezon M grease
from a syringe with a squared-off wide-bore needle
along the inner edges of the tape strips.
- Press a clean coverslip onto the grease. The
volume of the flow cell can be adjusted by spacing the
grease strips apart and/or by placing spacers between
the coverslip and the slide. Single-sided Scotch magic
tape is about 50µm thick, giving a flow cell of about
10mm × 5mm × 50µm, or 25µl. Thinner metal or
cellophane foils can be used to make a shallower flow
cell and conserve sample. It is helpful to make the flow
cell shallow because the microtubules below the top
surface scatter light and reduce contrast. For inverted
scopes, it is convenient to arrange flow crosswise. The
inset to Fig. 1 illustrates flow cells for inverted (A) and
upright (B) microscopes.
- Motility buffer: BRB 80 plus 1 mM MgATP. For
fluorescence work only, degas and add 1% of 100× antibleach mix (GOC), which is 100mg/ml glucose
oxidase (Sigma G7016), 18mg/ml catalase (Sigma
C100), and 300mg/ml glucose (Sigma G7528) in
BRB80 plus 50% glycerol. When aliquoting, fill tubes
to exclude oxygen, cap, and store at -20°C.
- 100× diluted MTs: either motility buffer or motility
buffer plus GOC.
D. Microscope Setup
- Place the flow cell fiat. Using a Gilson, inject into
the cell 1 chamber volume of motor solution. The solution
is drawn into the cell by capillarity. Incubate the
slide in a moisture chamber for 2-5min at 20°C to
allow the motor to adsorb to the glass.
- Wash the cell with 2 chamber volumes of assay
buffer, applying the solution to one side of the
chamber using a micropipette and drawing the solution
gently through the cell using the capillary action
of the torn edge of a strip of Whatman 3 MM, placed
at the exit of the chamber.
- Flow in 1 volume of MTs in motility buffer + taxol
and mount the slide on the microscope stage, oiling the
condensor to the bottom of the slide (it may be possible
to use a dry condensor for quick-and-dirty assays).
- Extra for fluorescence work 1. Degas some
BRB80. To 10ml, add 100µl of 100× GOC. Take another
3ml and add MgATP to 2mM. Fill and cap tubes to
exclude oxygen and hold buffers on ice. Add taxol to
20 µM freshly before use.
Before the day's work, align the microscope roughly
using a test specimen (a slide made using a suspension
of plastic beads provides a stable and realistic test
specimen). Switch on the lamp and allow a few
minutes for the arc to stabilise. Rack down the objective
and oil it to the slide. Insert some neutral density
filtration to protect your eyes from the intensely bright
light, focus roughly on the top surface of the grease at
the edge of the chamber, and then drive the stage to
centre the sample below the objective. Find some
beads attached to the undersurface of the coverslip.
Open the condensor aperture and close the field aperture.
Obtain Koehler illumination by focussing and
centring the condensor so that a sharp image of the
field diaphragm appears in the view field. Open the
field diaphragm again and adjust DIC sliders close to
Focussing on MTs in the experimental flow cell is
also best done using the grease surface as a guide.
Focus as described earlier and then remove neutral
density filters and switch in the video system. Adjust
fine focus to image the surface. Adjust light intensity
to almost saturate the camera (this is the point where
signal to noise is maximal). With the contrast on the
Argus set to maximum, microtubules should be visible
without background subtraction. Defocus slightly,
collect a background image, and subtract. Microtubules
should now be clearly visible.
A test sample of multispectral fluorescent beads is
very useful (Molecular Probes multispeck M-7900).
Switch on the arc lamp and allow a few minutes for
the arc to stabilise. Once the lamp is stable, align the
microscope for epifluorescence: Remove an objective
and place a piece of paper on the stage. Inset some
neutral density filtration. Close the field diaphragm
slightly and focus and centre the image of the lamp filament
that appears on the paper. Replace the objective.
Focussing on microtubules in the experimental cell
is much easier with dark-adapted eyes. Using the full
intensity of the mercury lamp, rack the objective down
until MTs are visible, first as a dim red glow, and then
as sharply defined bright red lines on a black background. Immediately reduce the illumination intensity
to protect against photobleaching, switch in the intensified
camera, and start recording.
E. Recording Data
The most flexible arrangement for data recording is
to set up time-lapse digital recording of video frames
to a computer hard disc (with no resolution loss) and
simultaneous recording to VCR. The VCR runs
uninterupted in the background for 3 h per tape and
generates an archive. The operator is free to go back to
this archive at a later date and recapture interesting
sequences for analysis.
F. Analysing Data Calibration
Image a stage graticule, a slide with etched lines at
1- or 10-µm intervals (from microscope manufacturers).
It is important to calibrate both in X
rotate the camera 90°. Most systems will give a different
number of pixels per micrometer in X
software compensates for this effect.
The best way to track is to follow the tip of a moving
microtubule: tracking the centroids, as common in cell
tracking, for example, will give you the wrong answer
as soon as the microtubule bends. For maximum accuracy,
the time lapse between frames should be adjusted
to minimise the effects of operator error when tracking
using the mouse. In practice we try to collect 20 frames
and adjust the time lapse so that the microtubules move
across the full field (22 µm
) during this time.
A. Archiving Data
It is very important to have a formal system for
identifying every video frame on every tape. In this
way there is no possibility of confusing data sets. The
simplest way to do this is to time and date stamp the
frames as they are generated, using the overlay feature
of the Argus. As ever, keeping careful written notes
also helps a lot. For complex experiments it can be
useful to speak notes onto the audio track of the
tape. Digital clips are archived most conveniently on
B. Imminent Technology
As computers get quicker, it is realistic to start recalculating
images in real time. Autocontrast is one interesting possibility, whereby the pixels of each incoming
frame are parsed and the look-up table is stretched to
optimise contrast. It will be some time before we can
dispense with the VCR. Real-time recording of uncompressed
grey-scale video to disc is pushing the limits
at present, but sufficiently fast sustained data transfer
rates will soon be available. This is not the real
problem, however. One frame of PAL video is 768 x
512 pixels, which, with 8 bit (256 greys) data, means
that each frame is 384 kb. Real-time recording to hard
disc fills the disc up at about 0.5Gb per minute, and it
soon becomes necessary to archive data to video tape.
C. Workstation Ergonomics
It is worth paying some attention to the ergonomics
of your microscope workstation. Microscope focus,
mouse, keyboard video, and contrast-adjustment electronics
all need to be within easy reach of a seated
operator. Screens should be visible with only a slight
turn of the head. It is very helpful to have a foot switch
to dim the room lights and blinds on windows.
D. Best Practice
Because of inherent uncertainties about the way a
particular protein attaches to a particular glass, motility
assays are at their strongest when used to measure
the relative motility in different treatments of samples.
It is commonly assumed that motility assays measure
motor-driven microtubule sliding under zero load. It
is probably more correct to assume that an unspecified,
variable, (but low) load applies.
The most common fault in video microscopy is to
overprocess an indifferent optical image. Too much
processing can seriously degrade the amount of information
in the image. A good primary image has high
spatial resolution (sharpness), high contrast, and low
background noise. Obtaining one is partly a function
of specimen preparation and partly of microscope
B. Lamp Intensity Fluctuates
In DIC, a troublesome problem is sudden variations
in light intensity caused by the arc of the mercury lamp
wandering. These are not noticeable in normal modes of microscopy, but with electronic amplification of contrast
they become annoying. The only solution is to
change the lamp. Cooling the lamp using a fan may
help. Mercury lamps typically need changing after
100h, because after that their intensity drops fairly
C. Microtubules Fishtail or Do
Not Move At All
Some motor proteins bind better to the glass surface
than others. Erratic motility may be due to your
protein denaturing on the glass or binding in such a
way that its force-generating conformational change is
inhibited. Areas of uncoated glass can also bind microtubules
and inhibit sliding. Increase motor concentration
if possible or try infusing the motor twice over and/or reducing or eliminating the wash step prior to
infusing microtubules. Including casein at 0.1-1 mgml
in the assay buffer efficiently protein coats glass.
Because motor activity can also be sensitive to thiol
oxidation, try including 5mM
DTT in your motility
Cross, R. A., and Kendrick, Jones J. (1991). Motor proteins. J. Cell.
Inoué, S., and Spring, K. R. (1997). "Video Microscopy."
Kron, S. J., Toyoshima, Y. Y., Uyeda, T. Q. R., and Spudich, J. A.
(1991). Assays for actin sliding movement over myosin-coated surfaces.
Methods Enzymology 196
Scholey, J. M. (1993). Motility assays for motor proteins. Methods Cell