Posted by
James Pawley on
URL: http://confocal-microscopy-list.275.s1.nabble.com/Re-Assessing-phototoxicity-in-live-fluorescence-imaging-tp7587005p7587019.html
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Even the simplest things aren’t.
Steffen is quite right about the power meter assuming (almost) normal incidence. You might think you could reduce this problem by putting immersion oil between the high-NA objective and the sensor surface, however, in many power meters, the silicon detector is separated from the protective cover glass by an air gap, so this doesn’t work (although you might be able to get around this by using a home-made
power meter consisting of a small “oilable” silicon solar cell from the electronics store connected to a current meter. But you would have to make sure that no stray room light hits this usually rather large cell and you would also have to calibrate it against a “real” power meter while remembering to correct for their different areas…)
The official way to do this measurement is to set up the microscope optics for Kohler Illumination using an oiled condenser with an NA at least as high as that of the objective. Assuming that there is no specimen in the way, the light emerging from the condenser should be parallel to the axis. Of course, you will still have reflection and absorption losses in the condenser and if you are doing this in wide field, the setting of the field diaphragm will have a massive effect (the transmitted power will be proportional to the area of the image plane illuminated by the field diaphragm). You will also have to be sure that the ray bundle from the condenser isn’t larger than your sensor.
On the other hand, as few scopes are set up with the required high-NA oiled condenser, it is common to fall back on the “10X low-NA” option.
Although this can provide fairly consistent day-to-day readings, you should NOT assume that you can just apply this reading to what would have emerged from a high-NA objective of higher magnification. This is because the illumination system is set up to provide a beam of a certain size in the BFP of the objective (in the space between the back of an infinity-corrected objective and the tube lens). This beam is usually roughly Gaussian in shape and its size is a compromise between the need to “fill the NA” of the “best lens” with fair uniformity and the desire not to waste too much light hitting the metal mounting of the optics (and then scattering all over the place causing "stray light”).
The problem is that “filling the NA” of, say a 40x 1.3 objective requires a beam diameter that is 2.5x larger (and 6.25 larger in area) than would be required for a 100x 1.3 objective. This is why early confocals often got better resolution with higher mag objectives:The ray bundle just didn’t fill the BFP of the lower-mag objectives. Conversely, they often got better “penetration” when using oil lenses into aqueous specimens when using the low-mag objectives: spherical aberration increases rapidly with NA and by being underfilled, the lower mag objectives were actually operating at a lower NA (at least on the illumination side).
So, although you can use the "10x low-NA" trick for monitoring day-to-day performance, you will probably need to use the Kohler set-up at least once to determine how the illumination optics in your scope fills the BFP. (It won’t work just to assume that the 40x 1.3 should pass 6.25x more light than the 100x 1.3 because, even if the beam is large at the BFP, it is still a Gaussian and brighter in the centre.) Basically, you need to determine the real conversion factors between your low-NA 10x and all your other objectives.
Even using a “dry" NA 0.9 condenser is better than trying to measure the convergent/divergent spot from a high NA objective. Try it with something like a 1.3 NA 40x oil (or a 0.5NA 10x dry) and, after you have set up Kohler using the normal transmitted light optics, open the condenser aperture to 0.9, turn off the transmitted source and turn on the epi-illumination (laser or WF) and hold a piece of white paper in the beam coming out of the condenser. You should see a bright blob and this blob may not even extend as far as NA 0.9 (which you can determine by moving the condenser aperture control to see if the edges of the blob are cut off by a sharp, black edge.) Knowing the size of this blob can help you at least estimate how much of the other objectives will be "filled."
I hope that this isn’t all too complicated and therefore disappointing because I think that, particularly when working with living cells, nothing is more important than knowing how much light hit the specimen while you collected your images.
So please persist!
Jm Pawley
James and Christine Pawley, 5446 Burley Place, Box 2348, Sechelt BC, Canada, V0N3A0
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James and Christine Pawley, 5446 Burley Place, Box 2348, Sechelt BC, Canada, V0N3A0
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On Jul 19, 2017, at 6:55 AM,
[hidden email]<mailto:
[hidden email]> wrote:
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Hi Steffen,
you can improve the accuracy of the method "a)" (that is measuring the laser
power before reaching the objective) greatly by mounting an iris stop in
place of the lens, adjusting the aperture to equal the diameter of the back
focal plane (BFP) aperture of the objective lens, and measuring the power of
the light that gets through this aperture.
Most confocal microscopes 'overfill' the BFP greatly (which is good for
resolution) and you can get an order of magnitude difference when the laser
beam is > 15 mm 'diameter' (it's gaussian profile) and the BFP diameter is
just 4 mm (e.g. some high-magnification lenses).
You can determine the BFP diameter by looking at the back of the lens, or by
dong some simple math (I guess diameter = 2 * NA * focal_length; focal_
length = tube_lens_focal_lenght / magnification).
This way, your a) and b) results should be much closer to each other and to
the real value in between...
Best, zdenek
--
Zdenek Svindrych, Ph.D.
W.M. Keck Center for Cellular Imaging (PLSB 003)
Department of Biology,University of Virginia
409 McCormick Rd, Charlottesville, VA-22904
http://www.kcci.virginia.edu/tel: 434-982-4869
---------- Původní e-mail ----------
Od: Steffen Dietzel <
[hidden email]>
Komu:
[hidden email]
Datum: 19. 7. 2017 7:03:20
Předmět: Re: Assessing phototoxicity in live fluorescence imaging
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Unfortunately not a solution, but a further complication: As far as I
know, power meters are made to detect light that hits the sensor
orthogonally. So with a high NA lens, a lot of the incident light won't
be even detected.
I guess what one could do is to measure
a) the power without objective with a parked beam, focusing on a spot in
the center of the field of view. This would give the upper estimate but
not the true intensity since some of it is absorbed by the objective
itself. Transmittance is never 100%.
b) doing the same with the objective that is to be used. This will give
the lower estimate. Too low, since part of the light won't be measured
due to the incident angle.
The truth then is somewhere between the two values. With modern high NA
objectives which should have a high transmission my gut feeling is that
the truth would be closer to (a) than to (b).
You could take value (a) and correct it the transmission of the
objective at the given wavelength published by the manufacturer, if that
is available. But I don't think I have ever seen a paper that actually
did all that. Whatever value you take, as Andreas suggested you then
would have to relate it to the true pixel dwell time, i.e. disregarding
dead time of the scanner.
To get the exact value at the focal point in the sample also would
require to take the losses due to reflection at the coverslip into
account. In essence, I am definitely with Claire when she says:
It is my
thought that comparing relative powers on the same instrument is okay but
comparing between systems will be very complex.
The Leica SP8 systems do allow to park the beam, as Craig suspected.
Since I always forget how to do that I put the procedure on our web
site, where I can easily find it :-)
http://www.bioimaging.bmc.med.uni-muenchen.de/manuals-protocols/maintenance_protocolls/laserpower/index.html
Cheers
Steffen
Am 19.07.2017 um 00:24 schrieb Craig Brideau:
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I've noted the effect Andreas mentions quite often. I usually set the
pixel
dwell time to the maximum such that the laser will spend the longest time
possible scanning out a single line, but even then the flyback can disturb
the reading. The best way is to park the mirrors in the center position,
although not all systems allow you to do that. Nikon's old C1 platform
allows for it, and ThorLabs' ThorScanLS has a park button as well. I'm not
sure about other vendors, but I'm sure others can chime in with their
experiences.
Craig
On Tue, Jul 18, 2017 at 11:44 AM, Andreas Bruckbauer <
[hidden email]> wrote:
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Hi Claire,
Confocals usually blank (switch off) the beam on the return and the power
meter averages between the on and off phases. Very slow scans are more
accurate an I usually use high zoom. Parking the beam is the better
option.
Best wishes
Andreas
-----Original Message-----
From: "Claire Brown" <
[hidden email]>
Sent: 18/07/2017 18:21
To: "
[hidden email]" <
[hidden email]>
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
--
------------------------------------------------------------
Steffen Dietzel, PD Dr. rer. nat
Ludwig-Maximilians-Universität München
Biomedical Center (BMC)
Head of the Core Facility Bioimaging
Großhaderner Straße 9
D-82152 Planegg-Martinsried
Germany
http://www.bioimaging.bmc.med.uni-muenchen.de"