Re: Assessing phototoxicity in live fluorescence imaging

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Claire Brown Claire Brown
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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
0000001ed7f52e4a-dmarc-request 0000001ed7f52e4a-dmarc-request
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Re: Assessing phototoxicity in live fluorescence imaging

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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

*****
To join, leave or search the confocal microscopy listserv, go to:
http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
Post images on http://www.imgur.com and include the link in your posting.
*****

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
Armstrong, Brian Armstrong, Brian
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Re: Assessing phototoxicity in live fluorescence imaging

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Hi Claire, I agree with you and I am sorry but I really do not have a clever solution.
A few thoughts: I always "park the beam" for measuring power from the objective. I don't think other methods make much sense (as you already pointed out).
 
Fluorescent standards do exist; Argo Light may be one of the better options. There are other less expensive options available as well (i.e., Ted Pella). Kurt Thorn wrote a nice blog on this subject some time ago and you may be able to still read it on-line.

Cheers,

Brian Armstrong PhD
Associate Research Professor
Developmental and Stem Cell Biology
Diabetes and Metabolic Diseases
Director, Light Microscopy Core
Beckman Research Institute, City of Hope


-----Original Message-----
From: Confocal Microscopy List [mailto:[hidden email]] On Behalf Of Claire Brown
Sent: Tuesday, July 18, 2017 10:20 AM
To: [hidden email]
Subject: Re: Assessing phototoxicity in live fluorescence imaging

*****
To join, leave or search the confocal microscopy listserv, go to:
http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
Post images on http://www.imgur.com and include the link in your posting.
*****

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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Craig Brideau Craig Brideau
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Re: Assessing phototoxicity in live fluorescence imaging

In reply to this post by 0000001ed7f52e4a-dmarc-request
*****
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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:

> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> 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
>
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> 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 Steffen Dietzel
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Re: Assessing phototoxicity in live fluorescence imaging

*****
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http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
Post images on http://www.imgur.com and include the link in your posting.
*****

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:

> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> 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:
>
>> *****
>> To join, leave or search the confocal microscopy listserv, go to:
>> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
>> Post images on http://www.imgur.com and include the link in your posting.
>> *****
>>
>> 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
>>
>> *****
>> To join, leave or search the confocal microscopy listserv, go to:
>> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
>> Post images on http://www.imgur.com and include the link in your posting.
>> *****
>>
>> 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
Cole, Richard W (HEALTH) Cole, Richard W (HEALTH)
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Re: Assessing phototoxicity in live fluorescence imaging

In reply to this post by Claire Brown
*****
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*****

What I do when I REALLY need to measure/know power (not for the faint of heart).  Works on all imaging modalities, albeit w/some minor tweaks.  I detailed this in a paper which I can dig up an anyone is interested.

Basics: use match pair of immersion objectives configured such that the emitted light from one objective is collected by the 2nd and a power meter measures the light emitted from the 2nd objective.  I typical use a double coverslip dye sandwich for alignment and then remove for the final measurement.  
CLSM tweaks: bidirectional collection or park, zoom >2.5, longest dwell time/slowest scan speed
        Cautions: check all filter cubes/ AOTF settings
                   check illumination stability before starting


Richard Cole
Research Scientist V
Director: Advanced Light Microscopy & Image Analysis Core
Wadsworth Center
 
Research Assistant Professor
Dept. of Biomedical Sciences
School of Public Health State University of New York

120 New Scotland Avenue, Albany N.Y. 12208
518-474-7048 Phone
518-408-1730 Fax

Website http://www.wadsworth.org/research/cores/alm
 twitter.com/microscopejock
zdedenn zdedenn
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Re: Assessing phototoxicity in live fluorescence imaging

In reply to this post by Steffen Dietzel
*****
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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
"*****
To join, leave or search the confocal microscopy listserv, go to:
http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
Post images on http://www.imgur.com and include the link in your posting.
*****

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:
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> 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:
>
>> *****
>> To join, leave or search the confocal microscopy listserv, go to:
>> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
>> Post images on http://www.imgur.com and include the link in your posting.

>> *****
>>
>> 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
>>
>> *****
>> To join, leave or search the confocal microscopy listserv, go to:
>> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
>> Post images on http://www.imgur.com and include the link in your posting.

>> *****
>>
>> 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
"
Guillermo Marques Guillermo Marques
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Re: Assessing phototoxicity in live fluorescence imaging

In reply to this post by Craig Brideau
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Olympus Fluoview 1000 allows parking of the beam, Nikon A1R does not.
Guillermo Marqués
University of Minnesota
Twin Cities Campus
University Imaging Centers  
Nikon Center of Excellence
www.uic.umn.edu
http://uic.umn.edu/content/locations
Any work utilizing UIC equipment or staff support must acknowledge the University Imaging Centers in publications and presentations.


> On Jul 18, 2017, at 5:24 PM, Craig Brideau <[hidden email]> wrote:
>
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> 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:
>
>> *****
>> To join, leave or search the confocal microscopy listserv, go to:
>> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
>> Post images on http://www.imgur.com and include the link in your posting.
>> *****
>>
>> 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
>>
>> *****
>> To join, leave or search the confocal microscopy listserv, go to:
>> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
>> Post images on http://www.imgur.com and include the link in your posting.
>> *****
>>
>> 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
>>
Matthew Pearson-2 Matthew Pearson-2
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Re: Assessing phototoxicity in live fluorescence imaging

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I'm sure i've parked the beam on our Nikon A1, you can set a single pixel as an ROI and only image at that point, thus not scanning with the galvo's.  Its in the simple ROI editor and is a crosshair icon, can't remember what its called, bleach point or something like that.

--
Matt Pearson
Microscopy Facility
MRC Human Genetics Unit
Institute of Genetics and Molecular Medicine (IGMM)
University of Edinburgh
Crewe Road
EH4 2XU




On 19 Jul 2017, at 14:57, Guillermo Marques <[hidden email]<mailto:[hidden email]>>
 wrote:

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Olympus Fluoview 1000 allows parking of the beam, Nikon A1R does not.
Guillermo Marqués
University of Minnesota
Twin Cities Campus
University Imaging Centers
Nikon Center of Excellence
www.uic.umn.edu
http://uic.umn.edu/content/locations
Any work utilizing UIC equipment or staff support must acknowledge the University Imaging Centers in publications and presentations.


On Jul 18, 2017, at 5:24 PM, Craig Brideau <[hidden email]> wrote:

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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



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Scotland, with registration number SC005336.
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Re: Assessing phototoxicity in live fluorescence imaging

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Correct. You can select a point ROI for photostimulation. But you cannot image that point.  For measuring the laser power it may work.
Guillermo Marqués
University of Minnesota
Twin Cities Campus
University Imaging Centers  
Nikon Center of Excellence
www.uic.umn.edu
http://uic.umn.edu/content/locations
Any work utilizing UIC equipment or staff support must acknowledge the University Imaging Centers in publications and presentations.


> On Jul 19, 2017, at 9:11 AM, PEARSON Matthew <[hidden email]> wrote:
>
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> I'm sure i've parked the beam on our Nikon A1, you can set a single pixel as an ROI and only image at that point, thus not scanning with the galvo's.  Its in the simple ROI editor and is a crosshair icon, can't remember what its called, bleach point or something like that.
>
> --
> Matt Pearson
> Microscopy Facility
> MRC Human Genetics Unit
> Institute of Genetics and Molecular Medicine (IGMM)
> University of Edinburgh
> Crewe Road
> EH4 2XU
>
>
>
>
> On 19 Jul 2017, at 14:57, Guillermo Marques <[hidden email]<mailto:[hidden email]>>
> wrote:
>
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> Olympus Fluoview 1000 allows parking of the beam, Nikon A1R does not.
> Guillermo Marqués
> University of Minnesota
> Twin Cities Campus
> University Imaging Centers
> Nikon Center of Excellence
> www.uic.umn.edu
> http://uic.umn.edu/content/locations
> Any work utilizing UIC equipment or staff support must acknowledge the University Imaging Centers in publications and presentations.
>
>
> On Jul 18, 2017, at 5:24 PM, Craig Brideau <[hidden email]> wrote:
>
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> 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:
>
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> 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
>
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> 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
>
>
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> The University of Edinburgh is a charitable body, registered in
> Scotland, with registration number SC005336.
James Pawley James Pawley
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Re: Assessing phototoxicity in live fluorescence imaging

In reply to this post by zdedenn
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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 [hidden email]<mailto:[hidden email]>, Phone 1-604-885-0840, cell 1-604-989-6146

James and Christine Pawley, 5446 Burley Place, Box 2348, Sechelt BC, Canada, V0N3A0 [hidden email]<mailto:[hidden email]>, Phone 1-604-885-0840, cell 1-604-989-6146



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
"

George McNamara George McNamara
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Re: Assessing phototoxicity in live fluorescence imaging

*****
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My 2 paragraphs:

1. Confocal microscopes: measure the signal from a coverglass, using
reflection mode (low laser power!). If ambitious, measure with different
pinhole sizes. Zeiss has an RT80/20 (which I hope means 20% reflection,
80% transmission), Olympus FV3000 has 10/90 (reflection/transmission).

2. Widefield microscopes, excitation side, http://www.epitechnology.com 
manufactures (3D prints) filter cubes with a mirror in place of the
dichroic, and facing the 'other way', to enable imaging the lamp (or LLG
or fiber) onto the detector -- which you (or someone else) paid (a lot
of) money for, and should be quantitative. Examples of what you'll see
are shown on http://www.epitechnology.com/epitechnology-1/

enjoy,

George


On 7/19/2017 4:08 PM, JAMES B PAWLEY wrote:

> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> 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 [hidden email]<mailto:[hidden email]>, Phone 1-604-885-0840, cell 1-604-989-6146
>
> James and Christine Pawley, 5446 Burley Place, Box 2348, Sechelt BC, Canada, V0N3A0 [hidden email]<mailto:[hidden email]>, Phone 1-604-885-0840, cell 1-604-989-6146
>
>
>
> On Jul 19, 2017, at 6:55 AM, [hidden email]<mailto:[hidden email]> wrote:
>
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> 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
> "*****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> 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:
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> 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:
>
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
>
> *****
>
> 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
>
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
>
> *****
>
> 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
> "
>

--


George McNamara, PhD
Baltimore, MD 21231
[hidden email]
https://www.linkedin.com/in/georgemcnamara
https://works.bepress.com/gmcnamara/75   (may need to use Microsoft Edge or Firefox, rather than Google Chrome)
http://www.ncbi.nlm.nih.gov/myncbi/browse/collection/44962650
http://confocal.jhu.edu (as of May 22, 2017)
phil laissue-2 phil laissue-2
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Re: Assessing phototoxicity in live fluorescence imaging

*****
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*****

Dear imagers,

Claire has raised an important issue, and it's great to see the discussion
and solutions - and so many people interested in reporting power
measurements. Thanks to everyone who chipped in with great advice.
There almost seem to be as many ways to do this as there are microscopists,
so this is a tricky area. We have used references in our review we thought
are reasonably practical:

- Waters, J.C. & Wittmann, T. (eds.) Quantitative Imaging in Cell Biology.
(Academic Press, 2014).
- Cranfill, P.J. et al. Nat. Methods 13, 557–562 (2016).

As discussed in this thread, conventional power sensors, where the flat
entry window is placed against the incoming light, are useful for measuring
the power of low NA objectives. They cannot be used to accurately measure
high-NA oil and water immersion lenses, which produce a highly focussed
cone of light (Dobrucki, 2013 - see reference below). The Thorlabs S170C
slide power sensor was developed with this problem in mind, with an
index-matching layer between the protective glass window and the photodiode
to prevent reflection, and can be used for high NA oil and water objectives.
- Fluorescence microscopy. JW Dobrucki. In: Fluorescence Microscopy: From
Principles to Biological Applications. First Edition. Edited by Ulrich
Kubitscheck.  Wiley-VCH Verlag GmbH & Co. KGaA, 2013.

A practical approach using an iris, as mentioned by Zdenek, is described
here:
Grünwald D, Shenoy SM, Burke S, Singer RH. Calibrating excitation light
fluxes for quantitative light microscopy in cell biology. Nat
Protoc. 2008;3(11):1809-14. doi: 10.1038/nprot.2008.180. PubMed PMID:
18974739;

Note that there were similar threads in this forum a few years ago:
instrument to measure laser intensity on slide (05/07/2014)
Reporting laser power in publication (18/08/2015)

There is also a practical note by Vojnovic, Newman and Barber from 2007
(updated in 2011):
http://users.ox.ac.uk/~atdgroup/technicalnotes/Optical%20pow
er%20meters%20for%20fluorescence%20microscopy.pdf

@ Richard Cole: I don't know the publication you mention, but it would be
great if you could post the reference here. Thanks!

Because of these differences and the technical aspects that need to be
considered, it is difficult for a routine lab to do this, and e.g. the
Thorlabs power sensor comes with a noticeable price tag. For these reasons,
it would be really *most *helpful if technology developers and commercial
microscopy manufacturers would provide data on the amounts of light
entering the sample at any given settings of the microscope, or to
incorporate tools to easily obtain such measurements, such that
non-specialists can do it without too much trouble. It's a shame that even
light transmission data for an objective should often be so hard to obtain.

So in many cases, a power measurement is clearly more of an estimate. It
might be useful for us as a community to decide on one standardised
approach. Until then, careful reporting on how it was done (while avoiding
the most prominent errors as described in this thread) will be the best way
forward.

With the best wishes,

Philippe

_________________________________________
Philippe Laissue, PhD
Royal Society Industry Fellow and MBL Whitman Center Scientist
University of Essex, Colchester CO4 3SQ, UK
(0044) 01206 872246 / (0044) 07842 676 456
[hidden email]
website <https://laissue.github.io/>

On 19 July 2017 at 22:48, George McNamara <[hidden email]> wrote:

>
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> My 2 paragraphs:
>
> 1. Confocal microscopes: measure the signal from a coverglass, using
reflection mode (low laser power!). If ambitious, measure with different
pinhole sizes. Zeiss has an RT80/20 (which I hope means 20% reflection, 80%
transmission), Olympus FV3000 has 10/90 (reflection/transmission).
>
> 2. Widefield microscopes, excitation side, http://www.epitechnology.com
manufactures (3D prints) filter cubes with a mirror in place of the
dichroic, and facing the 'other way', to enable imaging the lamp (or LLG or
fiber) onto the detector -- which you (or someone else) paid (a lot of)
money for, and should be quantitative. Examples of what you'll see are
shown on http://www.epitechnology.com/epitechnology-1/

>
> enjoy,
>
> George
>
>
> On 7/19/2017 4:08 PM, JAMES B PAWLEY wrote:
>>
>> *****
>> To join, leave or search the confocal microscopy listserv, go to:
>> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
>> Post images on http://www.imgur.com and include the link in your posting.
>> *****
>>
>> 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 [hidden email]<mailto:[hidden email]>, Phone
1-604-885-0840 <(604)%20885-0840>, cell 1-604-989-6146 <(604)%20989-6146>
>>
>> James and Christine Pawley, 5446 Burley Place, Box 2348, Sechelt BC,
Canada, V0N3A0 [hidden email]<mailto:[hidden email]>, Phone
1-604-885-0840 <(604)%20885-0840>, cell 1-604-989-6146 <(604)%20989-6146>
>>
>>
>>
>> On Jul 19, 2017, at 6:55 AM, [hidden email]<mailto:[hidden email]>
wrote:
>>
>> *****
>> To join, leave or search the confocal microscopy listserv, go to:
>> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
>> Post images on http://www.imgur.com and include the link in your posting.
>> *****
>>
>> 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 <(434)%20982-4869>
>>
>> ---------- 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
>> "*****
>> To join, leave or search the confocal microscopy listserv, go to:
>> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
>> Post images on http://www.imgur.com and include the link in your posting.
>> *****
>>
>> 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-protoc
ols/maintenance_

>> protocolls/laserpower/index.html
>>
>>
>> Cheers
>>
>> Steffen
>>
>>
>> Am 19.07.2017 um 00:24 schrieb Craig Brideau:
>> *****
>> To join, leave or search the confocal microscopy listserv, go to:
>> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
>> Post images on http://www.imgur.com and include the link in your posting.
>> *****
>>
>> 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:
>>
>> *****
>> To join, leave or search the confocal microscopy listserv, go to:
>> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
>> Post images on http://www.imgur.com and include the link in your posting.
>>
>> *****
>>
>> 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
>>
>> *****
>> To join, leave or search the confocal microscopy listserv, go to:
>> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
>> Post images on http://www.imgur.com and include the link in your posting.
>>
>> *****
>>
>> 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
>> "
>>
>
> --
>
>
> George McNamara, PhD
> Baltimore, MD 21231
> [hidden email]
> https://www.linkedin.com/in/georgemcnamara
> https://works.bepress.com/gmcnamara/75   (may need to use Microsoft Edge
or Firefox, rather than Google Chrome)
> http://www.ncbi.nlm.nih.gov/myncbi/browse/collection/44962650
> http://confocal.jhu.edu (as of May 22, 2017)
0000001ed7f52e4a-dmarc-request 0000001ed7f52e4a-dmarc-request
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Re: Assessing phototoxicity in live fluorescence imaging

In reply to this post by Claire Brown
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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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http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
Post images on http://www.imgur.com and include the link in your posting.
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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
Craig Brideau Craig Brideau
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Re: Assessing phototoxicity in live fluorescence imaging

*****
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Post images on http://www.imgur.com and include the link in your posting.
*****

Well, the laser spot size will be constant for a given objective, so the
PSF will give you a good idea of the distribution of the laser energy over
the three-dimensional volume. If your sample is smaller than the PSF then
the exposure will depend on how the PSF crosses the sample as the laser
scans. Exposure time will actually be less than the pixel dwell. For a
larger object, a good approximation would be the summation of the pixel
dwell time of all pixels that comprise the object, so for instance you can
determine that a cell has received 'X' microseconds of laser based on the
total number of pixels and the dwell time. The instantaneous energy
deposition is the energy density of the PSF, but the average power
deposited will be the length of time the cell actually has the laser on it
per scan.
One thing that bothers me though is the gap between x-lines as you reduce
resolution. The pixels will be larger but the same laser PSF is used to
construct the larger pixels, so you are using the same energy flow over a
larger area. In accommodating the larger pixel size though, the system must
slew the Y galvo through a larger step, so there should be areas of the
sample the laser skips if your pixel size is much larger than the laser
PSF. I'm still mulling that one over if anyone wishes to share their
thoughts.

Craig

On Thu, Jul 20, 2017 at 3:04 PM, <
[hidden email]> wrote:

> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
>
> 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
>
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> 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
>
0000001ed7f52e4a-dmarc-request 0000001ed7f52e4a-dmarc-request
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Re: Assessing phototoxicity in live fluorescence imaging

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*****

>so there should be areas of the sample the laser skips if your pixel size is much larger than the laser PSF.

I think lavision designed their cloud scanner for the multiphoton exactly for this problem, basically extending the beam size by multiplexing the beam.
Lenses with an aperture to limit the NA might be useful too, but you would loose light in the detection, underfilling?

Best wishes

Andreas

Best wishes

Andreas



-----Original Message-----
From: "Craig Brideau" <[hidden email]>
Sent: ‎20/‎07/‎2017 22:36
To: "[hidden email]" <[hidden email]>
Subject: Re: Assessing phototoxicity in live fluorescence imaging

*****
To join, leave or search the confocal microscopy listserv, go to:
http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
Post images on http://www.imgur.com and include the link in your posting.
*****

Well, the laser spot size will be constant for a given objective, so the
PSF will give you a good idea of the distribution of the laser energy over
the three-dimensional volume. If your sample is smaller than the PSF then
the exposure will depend on how the PSF crosses the sample as the laser
scans. Exposure time will actually be less than the pixel dwell. For a
larger object, a good approximation would be the summation of the pixel
dwell time of all pixels that comprise the object, so for instance you can
determine that a cell has received 'X' microseconds of laser based on the
total number of pixels and the dwell time. The instantaneous energy
deposition is the energy density of the PSF, but the average power
deposited will be the length of time the cell actually has the laser on it
per scan.
One thing that bothers me though is the gap between x-lines as you reduce
resolution. The pixels will be larger but the same laser PSF is used to
construct the larger pixels, so you are using the same energy flow over a
larger area. In accommodating the larger pixel size though, the system must
slew the Y galvo through a larger step, so there should be areas of the
sample the laser skips if your pixel size is much larger than the laser
PSF. I'm still mulling that one over if anyone wishes to share their
thoughts.

Craig

On Thu, Jul 20, 2017 at 3:04 PM, <
[hidden email]> wrote:

> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
>
> 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
>
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> 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
>
zdedenn zdedenn
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Re: Assessing phototoxicity in live fluorescence imaging

*****
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*****

Sure, NA aperture is not very helpful here, as in fluorescence "every photon
counts".
Some microscope vendors (Olympus?) allow for underfilling (limiting the
excitation beam diameter), but it also reduces the z-resolution (apart from
XY resolution, of course).

But remember that when taking z-stacks, the localized bleaching is not an
issue, for you are bleaching the whole thickness of your sample all the
time!

zdenek

---------- Původní e-mail ----------
Od: Andreas Bruckbauer <[hidden email]>
Komu: [hidden email]
Datum: 20. 7. 2017 18:02:18
Předmět: Re: Assessing phototoxicity in live fluorescence imaging
"*****
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Post images on http://www.imgur.com and include the link in your posting.
*****

>so there should be areas of the sample the laser skips if your pixel size
is much larger than the laser PSF.

I think lavision designed their cloud scanner for the multiphoton exactly
for this problem, basically extending the beam size by multiplexing the
beam.
Lenses with an aperture to limit the NA might be useful too, but you would
loose light in the detection, underfilling?

Best wishes

Andreas

Best wishes

Andreas



-----Original Message-----
From: "Craig Brideau" <[hidden email]>
Sent: ‎20/‎07/‎2017 22:36
To: "[hidden email]" <[hidden email]>
Subject: Re: Assessing phototoxicity in live fluorescence imaging

*****
To join, leave or search the confocal microscopy listserv, go to:
http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
Post images on http://www.imgur.com and include the link in your posting.
*****

Well, the laser spot size will be constant for a given objective, so the
PSF will give you a good idea of the distribution of the laser energy over
the three-dimensional volume. If your sample is smaller than the PSF then
the exposure will depend on how the PSF crosses the sample as the laser
scans. Exposure time will actually be less than the pixel dwell. For a
larger object, a good approximation would be the summation of the pixel
dwell time of all pixels that comprise the object, so for instance you can
determine that a cell has received 'X' microseconds of laser based on the
total number of pixels and the dwell time. The instantaneous energy
deposition is the energy density of the PSF, but the average power
deposited will be the length of time the cell actually has the laser on it
per scan.
One thing that bothers me though is the gap between x-lines as you reduce
resolution. The pixels will be larger but the same laser PSF is used to
construct the larger pixels, so you are using the same energy flow over a
larger area. In accommodating the larger pixel size though, the system must
slew the Y galvo through a larger step, so there should be areas of the
sample the laser skips if your pixel size is much larger than the laser
PSF. I'm still mulling that one over if anyone wishes to share their
thoughts.

Craig

On Thu, Jul 20, 2017 at 3:04 PM, <
[hidden email]> wrote:

> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
>
> 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
>
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> Post images on http://www.imgur.com and include the link in your posting.
> *****
>
> 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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Re: Assessing phototoxicity in live fluorescence imaging

In reply to this post by 0000001ed7f52e4a-dmarc-request
*****
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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

---------- Původní e-mail ----------
Od: [hidden email]
Komu: [hidden email]
Datum: 20. 7. 2017 17:18:47
Předmět: Re: Assessing phototoxicity in live fluorescence imaging
"*****
To join, leave or search the confocal microscopy listserv, go to:
http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
Post images on http://www.imgur.com and include the link in your posting.
*****


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

*****
To join, leave or search the confocal microscopy listserv, go to:
http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
Post images on http://www.imgur.com and include the link in your posting.
*****

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
"
0000001ed7f52e4a-dmarc-request 0000001ed7f52e4a-dmarc-request
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Re: Assessing phototoxicity in live fluorescence imaging

*****
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http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
Post images on http://www.imgur.com and include the link in your posting.
*****

Hi Zdenek, Craig

Thanks for your replies, but Zdenek's linear case doesn't account for the instantaneous power deposition Craig mentions. The high intensity of the scanned laser completely disappeared from the equation. I think in the context of phototoxicity one cannot assume this linearity. So one should monitor both, average exposure and instantaneous. The time domain will also be important, slow scanning vs fast, pulsed lasers and cw. Experiments will hopefully show. Thanks Philippe for starting this discussion.

Best wishes

Andreas

-----Original Message-----
From: "[hidden email]" <[hidden email]>
Sent: ‎21/‎07/‎2017 00:26
To: "[hidden email]" <[hidden email]>
Subject: Re: Assessing phototoxicity in live fluorescence imaging

*****
To join, leave or search the confocal microscopy listserv, go to:
http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
Post images on http://www.imgur.com and include the link in your posting.
*****

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

---------- Původní e-mail ----------
Od: [hidden email]
Komu: [hidden email]
Datum: 20. 7. 2017 17:18:47
Předmět: Re: Assessing phototoxicity in live fluorescence imaging
"*****
To join, leave or search the confocal microscopy listserv, go to:
http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy 
Post images on http://www.imgur.com and include the link in your posting.
*****


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

*****
To join, leave or search the confocal microscopy listserv, go to:
http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy 
Post images on http://www.imgur.com and include the link in your posting.
*****

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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