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Hi Guy,
The Zeiss 510, unlike the new Zeiss 710 confocal, has three pinholes so you
can adjust the confocal pinhole for each fluorochrome/channel, assuming you
aren't using the MetaHead for a couple of channels when you are forced to
share detector gain and pinhole.
The LMS-510 software reports the Airy diameter and the optical slice
thickness for the objective in use, i.e. with Airy=1: the optical slice is
reported as 0.9um for TRITC, 0.7um for FITC and 0.6um for DAPI when using
our 63x objective [Zeiss Plan Apochromat NA 1.4 oil]. As you say the optical
slice varies for the wavelength, so as we have adjustable pinholes on the
Zeiss we can adjust the DAPI and FITC channels confocal pinholes to a
slightly higher Airy number [Airy=1.25 and 1.5] to give the same optical
slice thickness for each fluorochrome [so all fluorochrome configs are set
to a 0.9um optical slice]. We don't have to think about it too deeply as the
LMS-510 software tells us all the numbers, as it knows the objective,
fluorochrome etc... The fluorescent beads mentioned in my posting were
scanned in the TRITC red channel.
As I say the lowest limit of our Axiovert 200M z-motor step is a 0.05um
step, around 50nm. In practice for 'quality' z stack scans most Zeiss 510
users generally select the standard Zeiss LMS-510 2x oversampling option in
Z-stacks with fixed cells [as it's an easy software button], which sets the
steps to half the LMS-510 calculated optical slice while keeping the same
first/last z slice location.
Regards
Keith
---------------------------------------------------------------------------
Dr Keith J. Morris,
Molecular Cytogenetics and Microscopy Core,
Laboratory 00/069 and 00/070,
The Wellcome Trust Centre for Human Genetics,
Roosevelt Drive,
Oxford OX3 7BN,
United Kingdom.
Telephone: +44 (0)1865 287568
Email: [log in to unmask]
Web-pages: http://www.well.ox.ac.uk/molecular-cytogenetics-and-microscopy
-----Original Message-----
From: Confocal Microscopy List [mailto:[log in to unmask]] On
Behalf Of Guy Cox
Sent: 02 November 2011 22:27
To: [log in to unmask]
Subject: Re: Deconvolution of Confocal Images? Volocity Course London
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*****
I'm a bit puzzled by this "0.9µm optical slice". If this is an NA 1.4 lens
Z resolution should be 400-500nm, I've measured this FWHM on 100nm beads, so
this is practical not theoretical. So you should be sampling at 200nm or
less in Z to meet Nyquist. This is before thinking about oversampling for
decon.
Guy
Optical Imaging Techniques in Cell Biology
by Guy Cox CRC Press / Taylor & Francis
http://www.guycox.com/optical.htm
______________________________________________
Associate Professor Guy Cox, MA, DPhil(Oxon)
Australian Centre for Microscopy & Microanalysis,
Madsen Building F09, University of Sydney, NSW 2006
Phone +61 2 9351 3176 Fax +61 2 9351 7682
Mobile 0413 281 861
______________________________________________
http://www.guycox.net
-----Original Message-----
From: Confocal Microscopy List [mailto:[log in to unmask]] On
Behalf Of Keith Morris
Sent: Thursday, 3 November 2011 12:59 AM
To: [log in to unmask]
Subject: Re: Deconvolution of Confocal Images? Volocity Course London
*****
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http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
*****
On a practical note regarding whether to bother with deconvolving a confocal
z stack using our Zeiss 510 Metahead confocal and our Volocity 3D
deconvolution [Restoration] software:
The advice from one of our Perkin Elmer's Volocity software engineers was
that in practice you don't see much improvement, if any, using Volocity's
Restoration module to deconvolve a standard confocal z-stack. Traditionally
using our Zeiss 510 we double over-sample a z-stack, i.e. for a 0.9um
optical slice [63x objective, Airy=1] we take a z-slice every 0.45um in the
z direction. In order to get a significant improvement over that with
Volocity's Restoration module, Perkin Elmer support recommended oversampling
by at least 10 [perhaps even up to 100] times rather than just 2. In
practice the confocal is limited to a minimum focus z step of around 0.05um,
giving about 20 Z scans when oversampling through each '0.9um optical slice'
[although you can push up the numbers, and scan time, by applying image
averaging as well]. This increases the scan time by about 10x for each z
stack, to typically over twenty minutes with 1024x1024 image sizes. Plus you
have to make time to apply Volocity's Fast Restoration or the slower
Iterative Restoration to the image z-stack.
Although time consuming, z-stack oversampling with deconvolution does get a
little bit more detail from our 3D z-stacks - it's just whether you want to
take another half hour to go though the acquisition and deconvolution
process for every z-stack, as the extra detail may not provide anything more
informative. Volocity Support added that Volocity's Restoration
deconvolution module predictably looks it's best when dealing with a
brighter and fuzzier z-stack from a normal wide-field microscope - where the
out of focus light can be put to use rather than be simply excluded by the
confocal iris. There is talk on the listserver of confocal deconvolution
also producing superior 3D spatial information for quantification of
sub-micron structures and well as improving resolution. Without something
like serial TEM section corroboration of the structure you can't really
allay the fear that the extra detail might include processing artefacts,
although most of this extra detail can be seen in the original fuzzier
confocal z-stack once the deconvolution has highlighted it. These days
though I expect you'd also look towards a colleagues super-resolution
STED/PALM system.
For those in the UK with an interest in using Volocity 3D software, next
week Perkin Elmer are running a two day Volocity user training course in
London on the 8th and 9th November 2011 [price £300 a day, £550 both days],
see:
http://now.eloqua.com/e/es.aspx?s=643&e=123818&elq=9c93259d524640798ee7b233e
bf0246c
Day 1: Volocity Essentials, Day 2: Volocity Quantitation
With our Volocity 3D software it tends to be the Quantitation module we use
the most, with standard Zeiss [non-de]convolved 2x over-sampled confocal
z-stacks, as it can measure intracellular structure volume and track
objects. We have the old Volocity v4.2 [latest is v6.0], although it's used
less frequently here than our 2D image analysis options, MetaMorph v7.7 and
ImageJ.
Regards
Keith
I have placed a deconvolved confocal ~20x oversampled 45 slice z-stack 3D
image at
http://www.well.ox.ac.uk/cytogenetics/deconvolved.jpg
It's a slice from the top of a couple of Invitrogen FocalCheck fluorescent
microspheres [Volocity Fast Restoration, see right image], where the
diameter of the slice shown is about 10 um and the depth of the ring about 2
um.
---------------------------------------------------------------------------
Dr Keith J. Morris,
Molecular Cytogenetics and Microscopy Core,
Laboratory 00/069 and 00/070,
The Wellcome Trust Centre for Human Genetics,
Roosevelt Drive,
Oxford OX3 7BN,
United Kingdom.
Telephone: +44 (0)1865 287568
Email: [log in to unmask]
Web-pages: http://www.well.ox.ac.uk/molecular-cytogenetics-and-microscopy
-----Original Message-----
From: Confocal Microscopy List [mailto:[log in to unmask]] On
Behalf Of John Oreopoulos
Sent: 30 October 2011 20:42
To: [log in to unmask]
Subject: Re: Deconvolution of Confocal Images? (was: Airy Units)
*****
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http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
*****
Peter, there are some advantages to deconvolving even confocal imaging data.
Until I took Pawley's course, I too had always seen these two imaging
techniques as mutually exclusive, but that's not the case.
Here is a passage from Chapter 2 of Pawley's Handbook:
"Although deconvolution and confocal imaging are often seen as competing
methods aimed at the same goal, namely producing images of 3D stain
distributions, in fact, they are not exclusive, and there is much to be said
for combining them. No only does deconvolution suppress the "single-pixel"
features created by Poisson noise, it also effectively averages the signal
in the Nyquist-sampled image of a point object. In other words, it has the
same effect of reducing the uncertainty in the estimate of the brightness in
individual voxels as Kalman averaging for 64 to 125 frames. This point is so
important that the present edition of this volume devotes an entire chapter
to it: Chapter 25."
Now, having repeated that passage, can I say that I deconvolve my confocal
data all the time? No, mainly due to lack of access to deconvolution
software, and partly due to time constraints usually. But I can assure you
that your images will look a bit better after deconvolving them. Apparently
all the best confocalists do this, but I seldom see it practiced in the
literature.
Have a look at Chapter 25 of his book. It's quite interesting.
John Oreopoulos
Research Assistant
Spectral Applied Research
Richmond Hill, Ontario
Canada
www.spectral.ca
On 2011-10-30, at 4:09 PM, Peter Werner wrote:
> *****
> To join, leave or search the confocal microscopy listserv, go to:
> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
> *****
>
> An interesting point was made here by Jim Pawley:
>
>> I agree that sampling a bit higher than Nyquist never hurts, especially
if you deconvolve (as you always should), but I think that it is a mistake
to think that one can "separate" out the noise by decon. I think that noise
is pretty fundamental.
>
> I had always heard that if you're doing confocal microscopy, at least
point-scanning confocal with a pinhole size of 1AU or smaller, that
deconvolution was superfluous, because you shouldn't be getting out of focus
light. So what is gained by deconvolution when one is sampling voxel by
voxel?
>
> Peter G. Werner
> Merritt College Microscopy Program
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