R A Cross

Molecular Motors Group
The Marie Curie Research Institute
The Chart
Oxted
Surrey RH8 OTL UK.

Phone 44 1883 722 306
Fax 44 1883 714 375
E mail r.cross@mcri.ac.uk
 

Last edit 06 06 2000

This is an updated version of an article I wrote for the Celis Handbook of Cell Biology, 3rd edition, (1998), which contains a number of helpful short articles on light microscopy. The Readers seeking to set up more specialised assays should refer to one of the excellent methodological compendia which are available (Inoue & Spring 1997, Cross and Kendrick Jones 1991, Scholey, 1993).
 

A. Hardware

1. Microscope

Unstained MTs are most conveniently visualised by video microscopy using computer enhanced DIC (differential interference contrast). MTs assembled from fluorescently labelled tubulin can be visualised using epifluorescence. Here I concentrate on these two contrast modes. It is possible also to visualise microtubules by dark field, which gives a high contrast image of unstained microtubules (see pic), or by phase constrast, 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 cheaper, 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, uncaging lasers, evanescence prisms) are envisioned.

Human eyes work in colour, have a higher dynamic range, better spatial resolution and a bigger viewfield 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.

2. Illumination

100W 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 which transmits infra red 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 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 (figure 1), and the other end emits a uniform disc of light (of Gaussian intensity profile) which is used to illuminate the microcope.

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) 60x or 100x planapo objective in order to maximise light gathering power and resolution; particularly so in epifluorescence, where the objective doubles as the condensor.

4. Anti-Vibration hardware

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 overemphasised. Before purchasing an expensive and awkward vibration-damping 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. We don't, and use water jackets for the objective and stage. These are readily fashioned by wrapping flexible, narrow bore copper tubing around several times, the ends of which are connected to a water bath with a circulator (eg Techne) using silicon tubing. If cooling is necessary, we found it useful to enclose the microscope in a tent made of cling film - this reduces condensation.

6. Camera

For fluorescence of moving objects, two types of low-light level / high contrast / high framing rate cameras are suitable: ICCD (intensified charge coupled device) and ISIT (intensified silicon intensified tube) cameras. ICCD cameras are better for current purposes because ISIT cameras, though more sensitive, introduce spatial and intensity distortions across the viewfield which are tedious to correct for.  All the above produce an analogue video signal.  Examples: ISIT Hamamatsu C2400-08 ; ICCD Hamamatsu C2400-97E .  An alternative approach which is becoming faesible 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 unparalleled sensitivity for fluorescence work,  but is currently very expensive (eg Princeton Instruments Pentamax about £30 000),  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 specs. of the hardware.

For DIC, a non-intensified scientific grade CCD is fine. Example: Greyscale CCD camera Hamamatsu C2400 77e.  Try also cheap surveillance cameras, which can give very good contrast.

7. Camera coupling / magnification

The ideal magnification sets 4 or more camera pixels across the width of the MT. For a 512 x 512 pixels (2/3 inch) CCD, this corresponds to a square field with sides of 20-25 microns. Zoom couplings are wonderfully convenient but are not always a good idea because they absorb a lot of light.

8. Image processor

Cameras typically come with a hardware box providing real time analogue enhancement of the video signal (any or all of gain, back-off and shading correction). Better is a digital video processor, 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 labs. The best and most flexible arrangement is to perform analogue enhancement, then feed the signal to the Argus and thence to the computer and VCR (see fig).

9. Monitor

It is worth investing in a high quality 14 inch multiformat monitor. Larger monitors look impressive but are not helpful for video.

10. Video recorder

At the time of writing, the only practical way to store large amounts of video is on 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 potentally the most serious problem. The effective resolution following recording can be visualised by recording and replaying a testcard image having black and white lines at various spatial frequencies. SVHS VCRs record considerably more faithfully than standard VHS. The more expensive domestic models are just as good as the "professional" versions, in our experience. Models with RS232 interfaces (e.g. Panasonic AG 5700) allow software control of the VCR. We have evolved a video recording strategy which offers maximum flexibility: Time lapse digital recording of video frames to computer hard disc (with no resolution loss) and simultaneous real-time SVHS recording to VCR. The VCR runs uninterupted in the background for three hours 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. We now have a fast capture application Scope which allows batch digitisation of video clips from tape for subsequent analysis.

Video discs (Optical Memory Disc Recorders, OMDRs) are convenient for data analysis because they offer random access to frames, but the media are too expensive to use for routine work. Many labs record to VCRs and then transfer interesting sequences to OMDRs because the random access to frames is useful for analysis. In our experience this is more conveniently and cheaply done now using a large computer hard disc.

Currently, different video formats unfortunately operate in different countries. In the US and Japan, NTSC format applies (525 lines per frame; 30 frames/sec, typically captured at 640x 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 x 576 pixels). It is worthwhile buying a VCR which can playback multiple formats, so that tapes provided by visiting scientists can be played. Editing VCRs which can convert formats are available but the investment is probably not worthwhile. On the rare occasions when this is necessary, video bureaux can do it (yellow pages).

11. Computing

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 two 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 100Mb Zip discs or 230Mb magneto-optical discs. CD writers are getting cheaper 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, the quicktime format is set to 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 which I recommend below. The situation is fluid, so please check the software documentation for a list of supported cards.

13. Software

On the Mac, the best route for analysis is NIH Image, which can be customised using macros to track objects and output the data in spreadsheet-compatible format. Macros for basic tracking through NIH Image stacks are available for download from our webpage http://mc11.mcri.ac.uk/retrac.html. NIH Image is now available for the PC.

RETRAC for the PC is to my knowledge the best thing available for either platform, because it is purpose-written. The programme is written in assembler and consequently runs very quickly under DOS or Windows 95/98. The latest version supports software control of a VCR, 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 file manager. Figure 2 shows a screenshot during tracking.

14.  Glassware

The type of slide used does not matter. The type of coverslip does. The thickness of the cover slip should be matched to the objective. The objective will be marked appropriately (eg 60 / planapo DIC 1.4 0.17/160 means a 60x objective selected as strain free for DIC, aplanatic (flat field); apochromatic (low chromatic aberration for blue yellow and green); optimised for cover glasses 0.17 mm thick and with a 160mm focal length). We use Chance 22mm x 22mm no.1.5 cover slips, without any special cleaning treatment. Experience suggests that "good" coverslips do not improve on cleaning, and "bad" coverslips cannot by cleaning be made "good"
 

A. Taxol-stabilised microtubules

Solutions
1M K-PIPES.
PIPES dissolves around it s isoelectric point of about pH 6.5. Take 500 mls water, add 65g solid KOH then, after cooling if necessary, slowly add 302 g 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, there is no way back.

100 mM NaGTP stock solution.
Nucleoside triphosphates like GTP and ATP undergo rapid hydrolysis at acidic pH, so efforts should be made to control pH when dissolving and storing them. Dissolve 1g NaGTP (Sigma G8877) in 15ml 10 mM NaPIPES pH 6.9, monitoring pH. Rapidly reneutralise pH by titrating in 5M KOH. Finely adjust pH, then make volume up to 19.11 ml. Store frozen at -20C in aliquots of 5-2000 µl. 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 10 mM K-PIPES pH 6.9, continuously monitoring pH and holding as close as possible to neutral using conc KOH. Once the ATP is dissolved, add 10 mls of 1M MgCl2 and readjust pH to 6.9. Adjust volume to 100.0 ml and freeze in aliquots of 5-5000 µl.

Taxol stock solution.
Wear gloves and work in the fume hood. Inject 2.93 ml anhydrous DMSO (Aldrich 27685-5) into a 25 mg bottle of taxol (Sigma T 7402). Dissolve by voretxing and store as 2-20µl aliquots at -20C. Taxol is stable in DMSO but unstable in water . It is insoluble in aqueous buffers above about 18µM. DMSO is explosive if it gets wet. Store small volumes at room temperature over beds of Sephadex G-50.

0.2 M NaEGTA
Dissolve 15.2g EGTA (Sigma E 4378) in 190 mls water. Adjust pH to neutral by adding conc NaOH, then make volume to 200.0 mls. Store at room temp.

1M MgCl2
20.33g MgCl2.6H2O to 100 ml water. Sterile filter and store at room temp.

BRB 80 (Brinkley reassembly buffer)
is 80 mM K-PIPES, 1 mM MgCl2, 1 mM EGTA pH 6.9. make up as a 10x stock, store at 4C and dilute freshly for use.

Purified tubulin at about 100 µM (Protocol for tubulin preparation on our webpage) in BRB80 should be flash frozen in 10-25µl aliquots in the presence of 30% glycerol by immersion in liquid nitrogen, and stored either at -70C or preferably in liquid nitrogen.

Steps
1. Thaw an aliquot of tubulin (typically 200µM), and add stock 100 mM NaGTP to 1mM and MgCl2 to 2mM. Warm to 37C and incubate for 20 mins.
2. After 20 minutes, add taxol from a 10 mM stock in DMSO to 20 µM final. Dilute microtubules 1000-fold for use using BRB80 buffer supplemented with 20µM taxol.

B. Preparation of Sample cells

Steps

1. 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.
2. 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.
3. Press a clean cover slip on to 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 50 µm thick, giving a flow cell of about 10mm x 5mm x 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 (see figure).  

C. Surface adsorption of motor

Solutions

Motility buffer
is BRB 80 plus 1 mM MgATP. For fluorescence work only, degas and add 1% of:

100xAntibleach mix (GOC)
which is 100 mg/ml glucose oxidase (Sigma G7016), 18 mg/ml catalase (Sigma C100), and 300 mg/ml glucose (Sigma G7528) in BRB80 plus 50% glycerol. When aliquoting, fill tubes to exclude oxygen, cap and store at -20C.

100x diluted MTs
in either motility buffer or motility buffer plus GOC.

Steps

1. Place the flow cell flat. 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 2-5 mins at 20C to allow the motor to adsorb to the glass.
2. 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 3MM, placed at the exit of the chamber
3. 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 for quick-and-dirty assays to use a dry condensor).
4. Extra for fluorescence work 1. Degas some BRB80. To 10 mls, add 100µl of 100xGOC. Take another 3 mls and add MgATP to 2 mM. Fill and cap tubes to exclude oxygen and hold buffers on ice. Add taxol to 20µM freshly before use.

D. Microscope set up (DIC)

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, then drive the stage to centre the sample below the objective. Find some beads attached to the undersurface of the cover slip. 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 viewfield. Open the field diaphragm again and adjust DIC sliders close to extinction.

Focussing on MTs in the experimental flow cell is also best done using the grease surface as a guide. Focus as above, 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 max, microtubules should be visible without background subtraction. Defocus slightly, collect a background image, and subtract. Microtubules should now be clearly visible.

D. Microscope set up (epifluorescence)

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, 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 which appears on the paper. Replace the objective.

Focussing on Mts 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 a 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 computer hard disc (with no resolution loss) and simultaneous SVHS recording to VCR. The VCR runs uninterupted in the background for three hours 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 is by imaging 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 and Y; simply rotate the camera 90 degrees. Most systems will give you a different number of pixels per micron in X and Y. Tracking 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 max 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 dmicrotubules move across the full field (22µM) 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 datestamp 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 on to the audio track of the tape. Currently, digital video clips are most conveniently archived to 230Mb magneto optical discs.

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 stretched to optimise contrast. It will be some time before we can dispense with the VCR. Real time full resolution recording 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 768x 512 pixels, which with 8 bit (256 greys) data, means each frame is 384 Kb. Real time recording to hard disc fills the disc up at about 0.5 Gb per minute, and it soon becomes necessary to archive the data to video tape.  Notwithstanding,  consumer digital video cameras,  players and desktop computer editing packages are is already with us.   It is probable the Quicktime format will become the digital video standard.
 

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 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 any 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.
 

1. Computerphilia

The most common fault in video microscopy is to over-process 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 setup.

2. Lamp intensity fluctutates

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 100 hours, because their intensity then drops fairly rapidly.

3. Microtubules fishtail or don't move at all

Some motor proteins bind better to the glass surface than others. Eratic 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 mg ml in the assay buffer efficiently protein-coats glass. Motor activity can also be sensitive to thiol oxidation, so try including 5mM DTT in your motility buffer.
 

Motility Assays for Motor Proteins Ed Jonathan M. Scholey (1993) Methods in Cell Biology no. 39

Motor Proteins Ed R.A Cross and J Kendrick Jones J Cell Sci 1991 Suppl 14

Assays for actin sliding movement over myosin-coated surfaces. Kron, S.J., Y.Y. Toyoshima, T.Q.R. Uyeda & J.A. Spudich. 1991. Meth. Enzym. 196:399-416.

Video Microscopy Inoué, S., Spring, K.R. (1997). Plenum Press, New York. ISBN 0-306-45531-5

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