Notes for Zeiss LSM 510 Version 3.2

 Calendar is at Ru Yi  4-3474

Lasers (excitation wavelengths): Argon (458nm and 488nm) HeNe1(543nm) HeNe2 (633nm)
Inverted microscope; objective lenses in the preset rotation:

10 dry
20 dry
25 immersion—set correction factor to oil, glycerol, or water.
40 water
63 water; 63 oil available, but not in the rotation set.

Before turning on anything, turn on mercury arc lamp (HBO) underneath the microscope air table.
If HBO is not on, turn off computer first before turning the power on to prevent a power surge.
Switch on “remote control” in the left corner behind the microscope stand.

Push power button to power up the computer (underneath the computer monitors).
Two swing-out filters (polarizer for DIC) near top of scope should be swung out.
§ Login

Start LSM-FCS (icon is on the computer desktop)
Select ‘Scan New Images’, and ‘Start Expert Mode’
§ FileNew > Create a database for today’s session (file type *.mdb)
Make sure that you are saving to your own folder on the computer’s E drive under ‘Data’ directory.
File for future reference: load an old database and click on ‘Reuse’ button to bring up previous settings.
§ Acquire
§ Laser
Turn on the laser lines required for your experiment; Argon 488 for FITC, GFP; Helium Neon 543 for Rhodamine Red, Cy3; HeNe 633 for Cy5.
The Ar laser is switch on by clicking on ‘Standby’ for a mandatory warm up.
Set Argon laser output to ~50% (which corresponds to tube current of 5.9A~6.1A)
Point of reference: excitation wavelength is shorter than the emission wavelength of the fluorophore.
§ Microscope
Top part of scope flips back out of the way for putting sample onto the stage.
If switching between slide or dish holder, align the red dot on slide/dish holder with red dot on the stage.
Choose objective lenses.
If using 25immersion lens, remember to adjust for correction factor on the objective lens itself; line up symbol on with the type of immersion you’re using, e.g. W|| is for H2O immersion with coverslip (inverted microscope) vs. W| is for H2O immersion without coverslip on an upright microscope.
If it’s not a dry lens, add a tiny drop of H2O or glycerol to objective before loading sample onto holder.
Turn on transmitted light to align slide. Power box to right of scope should be switched on (green button).
Back to the LSM 510 software, check on ‘Light Remote’ and ‘On’; set light intensity to lowest setting, 0% is fine—anything higher may be too bright for your eyes.
To find sample under fluorescence, set reflector turret to FSet09 for FITC/GFP, FSet15 for Rhodamine.
Sliders on either side of oculars have to be pulled out to be able to see through the eyepieces.
Sliders at bottom right of the microscope base have to be set for VISible mode > top in bottom out

to send signal to the detector for LSM > top out bottom in

§ Configuration
Select single track or multi track (for two or more fluorophores).
In Single Track, to select a preset beam path setting from the pull down list, click on ‘Config’ floppy disk button on the side panel.
In Multi Track, click on ‘Config’ button and choose one of the preset configurations from the list, then click ‘Apply’; one can also choose to store/apply ‘Single Track’ settings to each of the beam paths.
User can check or uncheck the buttons to turn on or off reflector light or the detectors in each channel.
Clicking on the ‘Ch’ button itself allows user to apply a different false-color for that channel; black is designated as ‘off’.
§ Scan
Select mode, ‘frame’ or ‘line’
Frame size: 512512 pixels is typical, 10241024 pixels or higher may be suitable for some experiments
Faster scan speed = lower resolution so 8 is good to start, then 7 or 6 might be better, it’s a slower scan but perhaps it gives brighter images at this particular size of pinhole.
If images are noisy, turn on averaging, method = mean > number = 2 (or higher); scan time would be doubled likewise; averaging is an option for fixed slides and not for experiment using live samples.
8-bit is fine, but 12-bit is better, it yields higher degree of signal in each pixel. 2^8=256 vs. 2^12=4096
Scan direction  single direction; usually not loop or bi-directional.
Find the interested focal plane in specimen and adjust parameters:
Go to Channels, either hit Single to capture a snapshot of the specimen; or, hit Find (black and white icon) which will shows specimen on screen at the specified pinhole size with all sort of auto-adjustments to digital gain, amplitude offset, etc.
To align specimen in field of view, hit XY Continuous and move stage or focus in/out, then hit Stop when it’s adjusted to minimize photo-bleaching. Fast XY gives real-time response to changes, but it yields dimmer images.
Set the parameters for each channel independently; use XY Continuous to adjust
(1) Detector gain, typically ~800.
(2) Pinhole Ø should be minimized, but anything smaller than 1 Airy unit will significantly restricts light. Note: smaller pinhole Ø = more Z slices.
In Multi Track: pinholes should be aligned so that the size of optical slices match in each channel—not by matching the airy units—an important consideration for quantitative analyses later.
(3) Amplitude offset (when necessary, see notes below).
(4) Transmissions for the laser lines are typically set at:
488nm1~15%
543nm100%
633nm50%
To image in full range, use the color paletterange indicator to help adjust detector gain and/or amplitude offset. Run XY Continuous to see the effect of adjustment to gain or offset.
Red dots = upper limit > lower detector gainto reduce saturation or overexposure.
Blue dots = lower limit > move amplitude offset scale bar further right to reduce darkness.
XY Continuous is recommended over Fast XY because what you see (on screen) is what you get (in final results). With the scan speed maxed out, images inadvertently appear to be dimmer using Fast XY.
Range indicator changes image to grayscale (with red/blue) contrasting expression vs. background.
In an image with optimized range, there would be little red dots in the brightest cells, and some blue dots in the background.
Turn off color palette when finished adjusting.
There is a crop tool to resize the images, and fine-tuning on x or y-axis is possible through controls in bottom of Mode panel. The Reset button is useful to resize the cropped or zoomed-in image.
§ Z-stack
Under Z stack tab, select the Mark First/Last tab to mark the first slice and the last slice:
Hit XY Continuous and focus away from specimen to find bottom slice. Hit stop and mark as first slice.
Hit XY Continuous and focus back into specimen to find top slice, stop and mark as last.
Optimize # of slices > Hit the Z slice button
Select ‘Optimal interval’ that will yield the right # of slices to scan; ‘Optimal # of slices’ will changes with different pinhole sizes. A bigger pinhole Ø = thicker optical slices = fewer # of slices.
Can check on ‘mid’ (the middle slice)—this will help to check intensity/brightness; or activate auto Z correction [bottom quarter in Z stack tab] if working with fixed slides, not with live samples, and not for quantitative analyses afterwards.
Merging slices in a z-stack makes final picture much brighter ( # of slices, e.g. 10 slices will be ~10 times brighter on a flattened image).
When ready, hit the Start button, and it will scan the optical slices.
§ Time series
Set up either a single or a Z-stack under Scan Control
Open Acquire > TimeSeries
Under ‘End Series,’ choose the number of times you want to acquire the image(s)
For ‘Cycle Delay,’ choose time (30, 60, etc.) and unit of time (msec, sec, or min)
Hit StartT, at the bottom of the panel, it shows ‘scanning’ then a countdown to the next scan.
Cycle Delay is a timed delay after each cycle (of scan has been completed).
Time Interval is time of the interval between the beginnings of each scan (inclusive of scan time).
In other words, Cycle Delay starts the countdown only after each scan, but Time Interval starts the countdown at the beginning of each scan.

Compare these examples, where the durations of Cycle Delay and Time Interval are identical, but the results are different:

While it’s scanning, follow along through the Gallery view (shows the slices as they come up) or via the Split view (shows each channel separately, plus the composite view).
Save your newly acquired data to E:\Data\*.*

Always save acquired image(s) before you do anything else; the mantra is save to be safe.

Exporting images . . .
Export ‘Raw data’ as Tagged Image File (*.tif).
If saving a split view or adding other graphics, i.e. scale bar, export images as *.tif with ‘contents of window’ as a single or a series.
You may use ImageJ to open the *.lsm files. ImageJ is a free image-processing software for PC and Mac available from NIH website at
When you’re done…
Switch Argon laser to ‘standby’ if there is another user after you. Switch off lasers if yours is the last session of the day; remember to let Ar laser cool off (it takes ~3min). HeNe lasers do not require a cooling period. Clean the objective lens with lens paper (do not use Kimwipes).
§ Exit program.

Before you logoff, check the CDIF calendar online (refresh the browser).

Stop here if someone else is coming after you’re done.
But if you are the last one, let the computer shut down; the computer is off when the screen goes dark; then switch off the remote power control (left corner behind the microscope).

Also, switch off the HBO lamp; the sequence is not as critical as when powering up the system.

Backup and delete your data; please keep your folder under 4 gigabytes.

Some Rights Reserved. Creative Commons Attribution-NonCommercial-ShareAlike 3.0 License. These notes were adapted from notes courtesy of Mary Kinkel and Vicky Prince at Univ of Chicago (Revised 2011.10.10)

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