CSULB Confocal Facility, http://www.cnsm.csulb.edu/departments/biology/confocal/
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User's Guide to the CSULB Olympus Fluoview 1000
Version of 4 August 2009
Emergency contacts
In case of questions or emergencies, please contact (in this order; office numbers are here, and
cell numbers are on the whiteboard in the confocal room):
1. Bruno Pernet (Dept. Biol. Sciences, 562-985-5378)
2. Kevin Sinchak (Dept. Biol. Sciences, 562-985-8649)
3. Any other confocal-experienced faculty person (see list on wall in confocal room)
4. If all else fails, an Olympus technician (see name on emergency list in confocal room)
Essential system specifications
Optics – The microscope is an Olympus IX-81 inverted microscope. Objectives include the
following (only six can be mounted at a time), some of which are equipped for DIC optics:
4x (Plan Fluor, NA 0.13, WD 17 mm)
10x (Plan S-Apo, NA 0.4, WD 3.1 mm)
20x (Plan S-Apo, NA 0.75, WD 0.65 mm) (DIC)
40x (Plan Fluor, NA 0.6, WD 2.7-4.0 mm, correction collar) (DIC)
40x water (Plan Apo, NA 1.15, WD 0.26 mm) (DIC)
40x oil (Plan Fluor, NA 1.30, WD 0.20 mm) (DIC)
60x oil (Plan Apo, NA 1.42, WD 0.15 mm) (DIC)
100x oil (Plan S-Apo, NA 1.4, WD 0.12 mm) (DIC)
Epifluorescence illumination – mercury lamp, with four filter cubes:
DAPI (Hoechst/AMCA): ex 335-375, em 440-500
FITC (EGFP/Bodipy/Fluo3/Di O): ex 465-495, em 515-555
WIGA (TRITC, Di I): ex 515-550, em 600-640
Cy5: ex 590-640, em 670-735
Confocal illumination -- four lasers, with the following excitation lines:
405 nm – blue diode laser
515 nm – argon ion laser
458 nm – argon ion laser
559 nm – green/yellow diode laser
488 nm – argon ion laser
635 nm – red diode laser
Information on fluorophore excitation and emission frequencies is available on websites listed in
the “links” section of the facility website.
Detection – The confocal has three confocal detectors (photomultiplier tubes, or PMTs) and one
transmitted light PMT. This means that one can scan up to three fluorescent and one transmitted
channel simultaneously (by using “virtual channels”, you can also scan another 12 or so
fluorescent channels sequentially). The system is equipped with a diffraction-grating based
“spectral detection” system, which allows us to do lambda (wavelength) scanning (i.e.,
spectroscopy), and manual adjustment of detection wavelengths for separation of signals from
overlapping fluorophores.
CSULB Confocal Facility, http://www.cnsm.csulb.edu/departments/biology/confocal/
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PRECAUTIONS
• MERCURY LAMP. Once you turn the lamp on, it needs to be on for at least 30 minutes
before being turned off. Once it’s turned off, it needs to be off for at least 15 minutes before
being turned on again. The mercury lamp has a 300 hr bulb, but we usually run it for at least 400
hr; if the counter passes 400 hrs, do not turn it on! Tell Bruno and he’ll get the bulb changed.
After turning off the mercury lamp, it’s still very hot; be careful not to let the microscope cover
touch it!
• TURNING LASERS ON. Turn all lasers you’ll need during initial startup. If you turn on only
a subset of the lasers, and later decide you need the one(s) you hadn’t turned on, shut the whole
system down, let the mercury lamp cool down for 15 min (make sure that it had previously been
on for 30 minutes), then turn it all on again. Don’t turn lasers on mid-run!
• MICROSCOPE CONTROLS. Once you’ve entered the Fluoview software and found a
specimen using brightfield or epifluorescence, your use of the physical controls of the
microscope should be minimal. You will need the x-y stage controls and the focus knob. Don’t
switch objectives, filter cubes, etc. manually – do it via the “Microscope Controller” window on
the desktop.
• REMOVING AND REPLACING OBJECTIVES. Don’t do this. All of our objectives are
expensive, and some are very expensive. If you know ahead of time what objectives you’ll want,
ask Bruno and he’ll swap them out for you.
• USING OIL OR WATER IMMERSION OBJECTIVES. For both types of lenses, only a very
small amount of fluid is required. Drop it on the coverslip, then flip your slide over and put it on
the stage. Both oil and water can be found in the microscope room; water must be high-quality
milli-q water. If you’ve put oil or water on a slide, be careful about changing objectives – you
can change to another oil objective (we don’t have another water objective), but if you want to
switch to a dry objective you need to clean the slide first. When finished using immersion
lenses, clean the objective using lens paper and cleaning fluid (never Kimwipes!).
• REMOVING IMAGE FILES FROM THE FLUOVIEW COMPUTER. Anti-virus programs
interfere with the Fluoview software. Thus we have to be careful not to introduce viruses to the
system. This is why the computer is not, and should not be, connected to the internet. The only
safe way to remove files from the computer is by copying them to a NEW, datafree CD or DVD
(this is also nice in that it creates a semi-permanent backup). CDs are available for free in the
confocal room. Copy the files, transfer them to your own computer, and eventually delete them
from the Fluoview computer. Files older than a few weeks are fair game for deletion.
• KEEP THE MICROSCOPE AND ROOM CLEAN. There is lots of scanning time in confocal
microscopy, time when you will be twiddling your thumbs or doing email or something. If you
need food or drink, do it outside while a long scan is happening – no food or drink in MLSC
201! Also, please don’t do any specimen prep in the confocal room. Make sure to leave the
room clean when you’re finished.
CSULB Confocal Facility, http://www.cnsm.csulb.edu/departments/biology/confocal/
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A. Fill out the user log sheet
We use this to keep track of usage of the microscope and lasers (e.g., to help justify our grant to
NSF in annual reports) and to keep track of use of the mercury lamp.
B. Turning on the system
You only need to turn the power on to the components that you plan on using, and turning lasers
and the mercury lamp on and off and running them affects their lifespan. So, if you are sure you
won’t be using one of those illumination components, don’t turn it on. (The exception is the
405+635 nm laser power supply – this supply also runs the laser combiner, so you NEED it if
you’re planning on doing confocal.) Make sure to turn on all the lasers you might potentially
need at the start, as you shouldn’t turn lasers on once the computer is on. The general rule is to
turn on the power supplies starting with the component that has the greatest power drain,
working down, and ending with the computer. If you just need the computer to burn a CD/DVD
or something, just turn it and the monitors on, and no other components.
Turning on goes in this order (skip components if not needed):
1. Take off the microscope cover.
2. Argon ion laser [IF NEEDED]. Turn on the power switch, then turn the key to the right.
3. 405 and 635 nm laser diodes [MUST TURN ON IF DOING CONFOCAL]. “Olympus
FV10-MCPSU” box to the left of the microscope. Turn power switch on.
4. 559 laser diode [IF NEEDED]. “NTT Electronics” box to the left of the microscope. Turn
power switch on. The “temp” LED will begin blinking. WHEN IT STOPS BLINKING (this
may take several minutes), then turn the key to the “on” position.
5. Mercury lamp [IF NEEDED]. On shelf above the computer; power switch only.
6. Scan unit and microscope. Both are just power switches. The scan unit also has a key, but
the key should always be in the “on” position.
7. Computer and monitors. Log on to the PC using your username and password (username
guest, no password), then double-click on the Olympus FV10-ASW 1.7 icon. Log into the
software using your username and password. The Fluoview software must be running before
you proceed to C!
The order of events in turning
on the system! Turn on all the
components that you need for
the session (but not those you
don’t plan on using).
CSULB Confocal Facility, http://www.cnsm.csulb.edu/departments/biology/confocal/
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C. Finding a specimen
1. Use the computer (in the “AcquisitionSetting” window) to select the appropriate objective for
your sample (start with a low-power, non-oil immersion objective!). Use the coarse focus knob
to lower the objective so there is no risk of contacting it with the slide. If your slide requires oil
immersion, focus with a low-power objective, remove the slide, change to the oil objective, put a
tiny drop of oil on the coverslip, then mount the slide on the microscope stage, coverslip down.
2. Find your specimen using the microscope oculars and either brightfield or epifluorescence.
Set up illumination in the “ImageAcquisitionControl” window on the desktop to send the right
kind of light through to the oculars. Note that if you find yourself spinning the focus knobs on
the microscope a lot to effect a small change, the focus mechanism is probably set to fine.
Change it to coarse in the MicroscopeController window on the desktop.
Brightfield (transmitted light) – Tilt the top of the microscope
forward so that you can view transmitted light. Click the
“trans lamp” button on the top left corner of the window.
Adjust lamp brightness using buttons on the front of the
microscope. Set up Kohler illumination (you need to
do this if you plan on capturing transmitted light images).
To set up Kohler:
• focus on an object on the slide using transmitted light
• close the field diaphragm so that you see the diaphragm
edges in the eyepieces
• focus the condenser so that the diaphragm edges are sharp
• if necessary, center the condenser using the centering screws
• open the field diaphragm just enough so that light fills the
eyepieces
You should set up Kohler
illumination again every time
you change objectives; this
helps optimize transmitted
light optics and is standard in
brightfield microscopy.
Epifluoresecence – Close the mechanical shutter below the objective turret.
Click on the “epi lamp” button on the top left corner of the window.
Choose the appropriate filter cube for your specimen in the microscope
controller. If you now open the mechanical shutter, your specimen will
be illuminated. (If you see no light, perhaps the shutter on the mercury
lamp – see picture to the right – is closed; open it.) Find a region of
interest!
CSULB Confocal Facility, http://www.cnsm.csulb.edu/departments/biology/confocal/
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3. Once you’ve found your specimen, turn off brightfield or epifluorescent illumination. Once
you do this, the filter wheel automatically goes to position 1 for laser scanning (LSM). Change
the focus mechanism to fine.
D. Acquiring a quick and dirty confocal image
1. Pull out the Wollaston prism (the lower DIC prism, picture on previous page) from beneath
the objective turret until it comes to a stop (not completely!). The DIC prism may be out
already, since the previous user may not have used it. It interferes with confocal imaging
slightly. Leave it in if you plan on capturing DIC images as well as confocal images, though.
2. Choose the appropriate dye(s) for your sample. Click on the “Dye List” button on the left in
the ImageAcquisitionControl window (picture below). Double-click on the relevant dyes you’ve
used in your sample to add them to the “selected dyes”. Click “Apply”, and the relevant filters
will be put into place. In addition, the relevant lasers now show as “active” in the
ImageAcquisitionControl box. Close the Dye List window.
CSULB Confocal Facility, http://www.cnsm.csulb.edu/departments/biology/confocal/
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3. Check the settings in the “AcquisitionSetting” window. For an fast scan you want something
like:
• the fastest scan speed available
• an image size of 512x512
• zoom of 1
• laser powers all fairly low:
~1% for 405 nm
~5% for all other lasers
• double-check to make sure that
the correct objective is selected
4. In “ImageAcquisitionControl” (see below) do the following:
• set the relevant confocal
photomultiplier (PMT) HV’s
(high voltages) to ~700. The
transmitted light detector
PMT (TD1) needs less voltage
(~100); set that if you plan on
using it.
• turn the gain in each
confocal channel down to
minimum (1)
• make sure that both Kalman
filtering and sequential
scanning are not checked.
CSULB Confocal Facility, http://www.cnsm.csulb.edu/departments/biology/confocal/
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5. Now if you click “X-Y Repeat”, you will hopefully see an image in one or more channels
(depending on the number of dyes you selected). Click the “stop” button to stop scanning. If
you don’t see images, it might be due to one of the following:
is the the shutter below the objective turret open? (if not, open it.)
problems with staining the specimen (did you see fluorescence in widefield?)
is the relevant laser turned on?
did you select the right fluorophore in the Dye List?
does the laser actually excite the fluorophore you’ve used? (check its excitation spectrum)
or, you know, something else…
6. If you’re doing multiple labelling, make sure to make sure you’re interpreting everything
correctly, and that you are not afflicted by the following problems:
• autofluorescence? Examine a negative control.
• bleedthrough/crossover? This occurs when emission light from one fluorophore is detected
by multiple PMTs. To check for this, switch off (uncheck) all but one laser, and make sure that
there is only signal in the relevant channel. Do this for each laser in turn. If there are problems,
you can separate excitation and detection in the channels by doing sequential (line) scanning –
you should do this routinely for multiple labelling. If you must do simultaneous scanning for
some reason, you can adjust the detection wavelength windows with the “spectral” system (the
VBF icon in ImageAcquisitionControl) to control emission overlap.
7. Adjust the laser power or PMT HV in each channel to get an image of approximately the
correct brightness.
E. Capturing an XY image
1. Adjust the dynamic range of the detector PMTs to capture the maximum range of grays.
• press Ctrl-H on the keyboard to view the images in “intensity” mode.
• turn off (in AcquisitionSetting) all but one of your lasers.
• start X-Y repeat scanning. Ideally, now only one panel on the “Live” screen will show an
image – if images are showing in other panels, you’ve got bleedthrough or autofluorescence to
deal with (see above).
• for the fluorophore excited by that laser, adjust the relevant photomultiplier tube offset, gain,
and HV (in ImageAcquisitionControl).
1) offset: adjust so that the background (black parts of the image) shows a “snow” of blue
pixels. In intensity modes, blue pixels represent black (no signal) pixels.
2) HV: adjust so that a few pixels in the object image itself are red; in intensity mode, red
pixels represent completely white (saturated) pixels. HV should never go above about
700; with higher HV (high voltage) the image gets grainy or starts showing other weird
artifacts.
3) gain: can also be adjusted to alter red (saturation) levels in the image. Turning gain up
adds lots of noise to the image, so avoid this if possible.
• turn that laser off, and turn on the next one. Repeat the previous step, then do it again for as
many lasers as you are using.
• when you’re done, stop the X-Y repeat to avoid bleaching your sample. Get out of intensity
mode by pressing Ctrl-H again.
CSULB Confocal Facility, http://www.cnsm.csulb.edu/departments/biology/confocal/
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2. Change image size in “AcquisitionSetting” to 1024x1024, a more or less standard resolution
for publishable confocal images.
3. If desired, set the optimal pixel size for the objective and wavelengths you’re using. Optimal
pixel size for resolving fine detail in the X-Y plane is estimated by the Fluoview software
automatically. If you look near the bottom of the AquisitionSetting window, there is a line that
says something like:
X:0.40 !m/pix Y:0.40 !m/pix Z:1.34 !m/slice
The X and Y numbers are the optimal (Nyquist) pixel dimensions (pixels are square in the XY
plane, so X and Y are the same); they change when you change objectives. To see if you’re at
the right zoom to make pixels that optimal size, click on the “i" button on the left side of the
ImageAcquisitionControl window. This gives you both optimal pixel dimensions (under “optical
resolution”, and actual pixel dimensions (“pixel size”). If you change the zoom, you will change
the actual pixel dimensions, and you can make them approximate optimal by doing that.
Note that you don’t necessarily need to do this. The optimal pixel size is that which is best
for resolving fine detail, but that might not be your goal or need in imaging.
4. Improve the signal/noise ratio in your image. What you want to do is to capture as many
photons that are coming from your excited fluorophores as possible, and as few stray photons as
possible. You can do this in several ways:
• slowing the scan speed increases signal (costs: longer dwell time of the laser beam on each
part of the specimen increases bleaching, and a slower scan speed also increases the total time it
takes to scan a frame).
• Kalman filtering removes noise by scanning repeatedly and averaging the scans, subtracting
the “random” noise photons (costs: repeat scanning, so more bleaching, and increases total time
it takes to scan a frame). Kalman filtering 3-4 or more times often really improves an image.
5. Set sequential scanning if desired. If doing multiple labelling and working with fixed
specimens, you probably always want to do this to ensure against bleedthrough problems.
6. The combination of the above parameters (size of the image, scan speed, filtering, sequential,
etc.) determines the time it takes to do a scan. The time is listed below the scan speed slider as P
(point), L (line), F (frame), and S (stack – for a z-series; when you’re just doing XY, S is the
same as F since you’re doing a “stack” that’s one frame deep).
7. Right below the XY button in ImageAcquisitionControl, make sure the tabs for lambda,
depth, and time are not selected (that is, only XY is bolded in the button). Click to do a scan.
8. When your scan is complete, a window will open up with the image. Right-click, or
“File:Save As” to save it to your folder on disk D (in the Users folder). Save it as an oib file –
this is an Olympus format that saves the images and other “metadata” (on acquisition settings,
etc.) in the file. You can do manipulations (e.g., pseudocoloring, merges, etc.) on another
computer or in the Fluoview software. You can also export the image as a .tiff file, which can be
opened by Photoshop and any other image manipulation software.
CSULB Confocal Facility, http://www.cnsm.csulb.edu/departments/biology/confocal/
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F. Capturing an XYZ image (a Z-series, aka a stack)
1. Right below the XY button in ImageAcquisitionControl, select the depth tab. XYZ should
now be bolded in the button.
2. Identify the start and end points for the Z-series. Begin scanning rapidly using XY Repeat.
Use the fine focus knob on the microscope to
focus on to the first XY image you want to take.
Click the Start Set button to set that as the start
position.
3. Now use the fine focus knob to move through
the sample to the last XY image you want to
take. Click the End Set button.
4. Click the “go” button next to center to go to
the middle of your Z-series.
5. Optimize the image as in section E above
(that is, just as if it were a normal XY image).
6. Set the optimal step size for most efficient sampling. This is another Nyquist calculation, and
again, the Fluoview software does it for you. Resolution in the Z-axis is always poorer than in
the XY in confocal, so if you look at the optimal slice thickness, it will always be greater than
the XY pixel dimensions:
X:0.40 !m/pix Y:0.40 !m/pix Z:1.34 !m/slice
In any case, you can simply change StepSize to the recommended size (1.34 !m above), or you
can click on the “Op.” (optimum) to the right of the StepSize box – this automatically sets slice
thickness to (Nyquist/2). This way you oversample the Z-series, which means you’ll make sure
to capture all the detail possible in the Z-axis. Or, if you want to make a nice 3D rendering, you
may want to set step size even smaller (e.g., 0.5 !m). Smaller step sizes mean longer scanning;
you need to consider this in the final settings for the image. Check the “S” time at the top of
AcquisitionSettings – it may be quite large! If it’s too long for you, you need to trade off
something to make it shorter (faster scan speed? Fewer Kalman replicates? Shorter stack?
Bigger step size?).
7. Start the scan with the XYZ scan button at the top of ImageAcquisitionControl.
8. When the scan is finished, a new window will appear containing the final image; save to your
folder (on drive D, in the Users folder), ideally as an .oib file.
9. You can play with the Z-series in the Fluoview software (e.g., pseudocolor the channels,
generate a 3D rendering, make a little movie of the 3D rendering rotating around some axis you
determine, make merges, etc.) or in other software (e.g., ImageJ).
CSULB Confocal Facility, http://www.cnsm.csulb.edu/departments/biology/confocal/
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L. Turning the system off
1. If the argon laser (#1) is on, turn it’s key to the upright position. The fan will continue
blowing. DO NOT TURN OFF THE POWER SWITCH UNTIL THE FAN TURNS ITSELF
OFF. While you’re waiting, do everything below…
2. Clean any oiled objectives. Remove the slide holder and turn the objective turret manually
until the relevant objectives are available. Dab oil off of the objective using lens paper – never a
Kimwipe or any other material! – and a tiny amount of lens cleaning fluid. Lens paper and fluid
should always be present in MLSC 201; if not, please tell Bruno and he’ll restock.
3. Change the objective to the 4X (using the Microscope Controller on screen)
4. Exit the Fluoview software.
5. Turn the scan unit and the microscope (#5) power off
6. Turn the mercury lamp (#4) off.
7. Turn the 405 and 635 nm laser diodes (#2): switch only.
8. Turn off the 559 laser diode (#3): turn the key to off, then turn the power switch off.
9. Transfer your data to a CD or DVD (optional). If you want to do this now, open up “Creator
Classic”, whose icon is on the desktop. Pop a NEW CD or DVD into the computer. Find the
files you’d like to burn onto disc in D: FV10-ASW:Users:yourname:image. Select the files to
burn, and drag them into the “data disc project” box. Name the disc. Click the “burn” icon.
Confirm that your files have transferred by checking the disk on another computer. Once sure
the files are not corrupted, you can delete the originals from the Fluoview computer (you can do
this on your next session).
10. Shut down the computer.
11. Cover the microscope (make sure the cover doesn’t get near the hot mercury lamp on the
back right side of the scope! It will burn!). Clean up any mess you’ve left in the room.
12. Log out on the user log sheet.
13. As soon as the fan on the argon ion laser turns itself off, turn the power switch on that laser
off.
14. Make sure to close the door when you leave.
CSULB Confocal Facility, http://www.cnsm.csulb.edu/departments/biology/confocal/
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Postscript: other capabilities of our system
• capturing XYTime (timelapse) images
1. If you need to do XYT imaging (presumably on live material!), either talk to Bruno or try to
figure it out yourself. It’s basically like XYZ, except clicking Time except Depth (unless you
want to do XYZTime, which you can do by clicking both). There are some additional
constraints – for example, you can’t ask the software to capture an XY image every 5 seconds if
it takes 7 seconds to capture a single image.
It is also possible to do much more complicated time series, using the TimeController. This
allows you to program a macro, basically, and change all kinds of parameters over time in
whatever way you wish. For example, you can set the software to change PMT HV, or laser
power, or even where in the field of view you are sampling, in whatever time pattern you like
(over timescales of seconds, minutes, or even many hours). One reason you might want to do
something like this is to automatically alter laser power or PMT settings as you image deeper and
deeper into a specimen. If you want to do this, ask Bruno – there’s an easier way to do it than
using the TimeController.
• capturing XYLambda images
1. You might want to do this if your samples are stained with two dyes with very similar
emission spectra, or if your main fluorophore signal is similar to that of specimen
autofluorescence. You might also want to do it if you want to use the confocal as a fluorescent
spectroscope. Whatever you want to use this for, if you want to do XYLambda scanning, talk to
Bruno!
• other capabilities
Talk to Bruno if you want to do any of these (the first you can probably figure out easily
yourself):
• simple image analysis (e.g., measuring pixel intensities across an XY transect)
• FRET (fluorescent resonant energy transfer)
• FRAP (fluorescence recovery after photobleaching)
• photoactivation