Posted by
Zdenek Svindrych-2 on
URL: http://confocal-microscopy-list.275.s1.nabble.com/Re-Assessing-phototoxicity-in-live-fluorescence-imaging-tp7587005p7587028.html
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Hi Andreas,
to a firsts approximation, it's not so complicated.
Assumptions: bleaching is linear with illumination intensity, "reasonable"
sampling (confocal - scanned region is bigger than Airy disk, no big gaps
between lines (or z-stack acquisition); widefield - spatially constant
illumination intensity over known field of view).
Units: J/um^2 (Joule per square micrometer) seems appropriate.
Widefield: exitation_power [Watts] * exposure_time[seconds] / illuminated_
area [um^2]
Confocal: exitation_power [Watts] * scanning_time[seconds] * duty_factor /
illuminated_area [um^2]
The power is after the objective... the 'factor' is the duty cycle of the
scanning process (assuming the 'power' is the peak power) - then power*
factor = average excitation power.
In this approximation the PSF size, pixel size and counts, dwell time, etc.
are irrelevant (other than defining the scan time and scanned area).
Beware: the "linearity of bleaching" assumption is easy to break!
zdenek
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Datum: 20. 7. 2017 17:18:47
Předmět: Re: Assessing phototoxicity in live fluorescence imaging
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Dear all,
I wanted to pick this up again and discuss a different aspect. Even when we
could measure the laser power accurately, how would one compare power
density between widefield and confocal microscopy? The widefield case seems
pretty straightforward, one would need to know the area illuminated by the
light source. Usually I bleach a part of the sample and do a larger tile
scan and can hopefully see a sharp edge to measure the area. In the confocal
case one has the Gaussian beam profile, presumably easy to measure with a
small bead and an open pinhole. One could calculate an average over the beam
profile. But how can one deal with the beam scanning and account for
different situations like undersampling or oversampling? The easiest would
be power density x pixel dwell time x number of pixels which should be fine
when the pixels are on beam diameter apart. But when we then zoom in and
undersample, the same energy will be concentrated in a smaller area,
presumably leading to higher phototoxicity? Should one multiply by an
overfill factor? Would the photoxicity in this case not be lower than when
doing the same with a higher NA objective which would have a beam size
matching the (now zoomed in) pixel spacing? When undersampling, like using a
low mag objective with 512 x 512 pixels one can actually bleach nice lines
into the sample. In this case the photoxicity in the line will be high, but
the area between will not be illuminated. How to account for this?
best wishes
Andreas
-----Original Message-----
From: Claire Brown <
[hidden email]>
To: CONFOCALMICROSCOPY <
[hidden email]>
Sent: Tue, 18 Jul 2017 18:21
Subject: Re: Assessing phototoxicity in live fluorescence imaging
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Thank you for this great article and pointing to many great resources.
I wanted to bring up one issue we have had when trying to work on different
microscope and compare light density/exposure.
For the CLSM microscopes when we use a power meter at the focal plan the
power we measure depends a lot on the scan settings.
If we park the beam as a point we get one power. If we go to a 100x100 pixel
array at zoom 1 with a 10x lens the power is different. if we change the
scan speed the power is different again. I suspect this is related to how
the power meter integrates the light over time and also how sensitive it is
spatially across the sensor. We have decide to just quote our power as the
power we measure at the power meter with set conditions and we detail those
conditions in our materials and methods section of the paper. We try to use
a 10x/0.3 planfluar lens with no phase optics if we can.
We have stayed away from trying to calculate the power at the sample because
a lot of assumptions have to be made. The assumptions may be different for
wide-field versus CLSM versus light sheet versus spinning disk and so on.
We ran into these issues when just trying to repeat measurements on two
different confocals from two different manufacturers. It can really get
quite complex.
Does anyone have thoughts on this issue? Any cleaver solutions? It is my
thought that comparing relative powers on the same instrument is okay but
comparing between systems will be very complex.
Ideally, it would be good for the manufacturers to have some kind of laser
power measurement in the instrument and software that is always monitored.
Even if this is just a relative value to the actual power at the sample it
would really improve quantitative microscopy and also help in maintenance
and trouble shooting equipment. I'm not sure about others but this kind of a
feature would really be a strong selling point for me and the core
facilities I manage. In many cases these options are already built into the
hardware for the service engineers but are not accessible to the end user.
Sincerely,
Claire
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