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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Hello everyone, Is there any way to convert XYZT image to xyt. format. I have done live cell imaging with xyzt format and and I want to export the same file to XYT.avi format. That means I want to compile the z stack so that I can get the .avi image in xyt format. I am using Leica SP5 confocal misroscope. Regards Manish Kumar Imaging facility Manager IGIB, New Delhi |
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Dear Manish, You can use Image > Stacks > Tools > Grouped Z-project... Best wishes Kees -----Original Message----- From: Confocal Microscopy List [mailto:[hidden email]] On Behalf Of Manish Kumar Sent: 03 April 2013 10:51 To: [hidden email] Subject: xyzt image ***** To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Hello everyone, Is there any way to convert XYZT image to xyt. format. I have done live cell imaging with xyzt format and and I want to export the same file to XYT.avi format. That means I want to compile the z stack so that I can get the .avi image in xyt format. I am using Leica SP5 confocal misroscope. Regards Manish Kumar Imaging facility Manager IGIB, New Delhi |
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Dear Kees, Thanks for the reply. I have tried the option. This works with the confocal software, but didnt work when i export it to .avi . Rgds Manish On Wed, Apr 3, 2013 at 3:45 PM, Straatman, Kees R. (Dr.) < [hidden email]> wrote: > ***** > To join, leave or search the confocal microscopy listserv, go to: > http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy > ***** > > Dear Manish, > > You can use Image > Stacks > Tools > Grouped Z-project... > > Best wishes > > Kees > > -----Original Message----- > From: Confocal Microscopy List [mailto:[hidden email]] > On Behalf Of Manish Kumar > Sent: 03 April 2013 10:51 > To: [hidden email] > Subject: xyzt image > > ***** > To join, leave or search the confocal microscopy listserv, go to: > http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy > ***** > > Hello everyone, > > Is there any way to convert XYZT image to xyt. format. > > I have done live cell imaging with xyzt format and and I want to export > the same file to XYT.avi format. That means I want to compile the z stack > so that I can get the .avi image in xyt format. > I am using Leica SP5 confocal misroscope. > > Regards > Manish Kumar > Imaging facility Manager > IGIB, > New Delhi > |
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** I believe Kees was referring the Grouped Z-Project in Fiji/ImageJ, not something in your confocal software. Import your Leica file into Fiji with the LOCI plugin. The Grouped Z-Project will create a new stack in which each frame is the z-projection of the corresponding time point. There are several routes to save that as an AVI from Fiji/ImageJ. It is advisable to also archive the projection movie in an uncompressed format in case you want to edit it in the future. In fact, you can add timestamps and annotations in Fiji. Please refer to documentation: http://imagej.nih.gov/ij/docs/guide/index.html Regards, Glen Glen MacDonald Core for Communication Research Virginia Merrill Bloedel Hearing Research Center Cellular Morphology Core Center on Human Development and Disability Box 357923 University of Washington Seattle, WA 98195-7923 USA (206) 616-4156 [hidden email] [hidden email] On Apr 3, 2013, at 3:44 AM, Manish Kumar <[hidden email]> wrote: > ***** > To join, leave or search the confocal microscopy listserv, go to: > http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy > ***** > > Dear Kees, > Thanks for the reply. I have tried the option. This works with the confocal > software, but didnt work when i export it to .avi . > > Rgds > Manish > > > On Wed, Apr 3, 2013 at 3:45 PM, Straatman, Kees R. (Dr.) < > [hidden email]> wrote: > >> ***** >> To join, leave or search the confocal microscopy listserv, go to: >> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy >> ***** >> >> Dear Manish, >> >> You can use Image > Stacks > Tools > Grouped Z-project... >> >> Best wishes >> >> Kees >> >> -----Original Message----- >> From: Confocal Microscopy List [mailto:[hidden email]] >> On Behalf Of Manish Kumar >> Sent: 03 April 2013 10:51 >> To: [hidden email] >> Subject: xyzt image >> >> ***** >> To join, leave or search the confocal microscopy listserv, go to: >> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy >> ***** >> >> Hello everyone, >> >> Is there any way to convert XYZT image to xyt. format. >> >> I have done live cell imaging with xyzt format and and I want to export >> the same file to XYT.avi format. That means I want to compile the z stack >> so that I can get the .avi image in xyt format. >> I am using Leica SP5 confocal misroscope. >> >> Regards >> Manish Kumar >> Imaging facility Manager >> IGIB, >> New Delhi >> |
 
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Arvydas Matiukas |
Re: colocalization between confocal and two-photon: voxel matching
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In reply to this post by manish kumar-2
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Hello list, One of my Core users wants to measure colocalization between DAPI labeled and FITC-immunolabeled structures in nuclei of her fixed cells. Visual inspection shows good level of colocalization. However, what would be the correct way to quantify it. Our confocal is LSM510 META NLO. No UV or 405nm laser so two-photon mode at 780nm is used to excite blue dye while FITC is excited at 488nm in confocal mode . At default pinhole settings (1 airy unit for confocal and max=~10 A.u. for two-photon) both blue and green signals are good but the optical slices (and voxel sizes) differ by the factor of 10. Letting the software (LSM) to equalize the slices at 0.6um reduces the pinhole to 0.76 for confocal and 1.0 for two-photon. Unfortunately at such small pinhole for two-photon the blue signal is lost in noise (even at 5% laser power and 16 times averaging, further increase may bleach the dye during 15 min long scan). I can get the blue signal back to a measurable level by increasing the two-photon pinhole to 4 A.u. but this creates a four-fold mismatch in blue/green voxel size. I could boost the blue signal by increasing pixel size but the loss in resolution is not desirable. My first thought was to switch to red DNA stain and then do colocalization between two confocal signals with identical voxels. However, the user already completed experiment using DAPI staining, and it worked well. My question is what would be the correct way to acquire the confocal/two-photon z-stacks and subsequently quantify the colocalization. And related question if reducing pinhole works the same way for two-photon as for a regular confocal. Any feedback/advice/idea are very welcome, Arvydas ---------------------------- Arvydas Matiukas, Ph.D. Director of Confocal&Two-Photon Core Department of Neurosci& Physiology SUNY Upstate Medical University 766 Irving Ave., WH 3167 Syracuse, NY 13210 tel.: 315-464-7997 fax: 315-464-8014 email: [hidden email] |
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Armstrong, Brian |
Re: colocalization between confocal and two-photon: voxel matching
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Hello, when performing 2-Photon imaging keep the pinhole at maximum. In 2PE an optical slice is created via pinpoint excitation and not through the pinhole in the emission path (as is the case for Confocal imaging). The optical slice may be smaller/thinner in the 2P case. There is extensive explanation of this in Pawley's book (Handbook of Biological Confocal Microscopy) and in Diaspro's book (Confocal and 2-Photon Microscopy, Foundations Applications and Advances, Alberto Diaspro. Moreover, I have performed this same analysis on this same instrument (in press) so I may be able to answer other questions. Cheers, Brian D Armstrong PhD Assistant Research Professor Director, Light Microscopy Core Beckman Research Institute City of Hope Dept of Neuroscience 1450 E Duarte Rd Duarte, CA 91010 626-256-4673 x62872 http://www.cityofhope.org/research/support/Light-Microscopy-Digital-Imaging/Pages/default.aspx -----Original Message----- From: Confocal Microscopy List [mailto:[hidden email]] On Behalf Of Arvydas Matiukas Sent: Wednesday, April 03, 2013 10:29 AM To: [hidden email] Subject: Re: colocalization between confocal and two-photon: voxel matching ***** To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Hello list, One of my Core users wants to measure colocalization between DAPI labeled and FITC-immunolabeled structures in nuclei of her fixed cells. Visual inspection shows good level of colocalization. However, what would be the correct way to quantify it. Our confocal is LSM510 META NLO. No UV or 405nm laser so two-photon mode at 780nm is used to excite blue dye while FITC is excited at 488nm in confocal mode . At default pinhole settings (1 airy unit for confocal and max=~10 A.u. for two-photon) both blue and green signals are good but the optical slices (and voxel sizes) differ by the factor of 10. Letting the software (LSM) to equalize the slices at 0.6um reduces the pinhole to 0.76 for confocal and 1.0 for two-photon. Unfortunately at such small pinhole for two-photon the blue signal is lost in noise (even at 5% laser power and 16 times averaging, further increase may bleach the dye during 15 min long scan). I can get the blue signal back to a measurable level by increasing the two-photon pinhole to 4 A.u. but this creates a four-fold mismatch in blue/green voxel size. I could boost the blue signal by increasing pixel size but the loss in resolution is not desirable. My first thought was to switch to red DNA stain and then do colocalization between two confocal signals with identical voxels. However, the user already completed experiment using DAPI staining, and it worked well. My question is what would be the correct way to acquire the confocal/two-photon z-stacks and subsequently quantify the colocalization. And related question if reducing pinhole works the same way for two-photon as for a regular confocal. Any feedback/advice/idea are very welcome, Arvydas ---------------------------- Arvydas Matiukas, Ph.D. Director of Confocal&Two-Photon Core Department of Neurosci& Physiology SUNY Upstate Medical University 766 Irving Ave., WH 3167 Syracuse, NY 13210 tel.: 315-464-7997 fax: 315-464-8014 email: [hidden email] --------------------------------------------------------------------- *SECURITY/CONFIDENTIALITY WARNING: This message and any attachments are intended solely for the individual or entity to which they are addressed. This communication may contain information that is privileged, confidential, or exempt from disclosure under applicable law (e.g., personal health information, research data, financial information). Because this e-mail has been sent without encryption, individuals other than the intended recipient may be able to view the information, forward it to others or tamper with the information without the knowledge or consent of the sender. If you are not the intended recipient, or the employee or person responsible for delivering the message to the intended recipient, any dissemination, distribution or copying of the communication is strictly prohibited. If you received the communication in error, please notify the sender immediately by replying to this message and deleting the message and any accompanying files from your system. If, due to the security risks, you do not wish to receive further communications via e-mail, please reply to this message and inform the sender that you do not wish to receive further e-mail from the sender. (fpc5p) --------------------------------------------------------------------- |
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Julio Vazquez |
Re: colocalization between confocal and two-photon: voxel matching
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In reply to this post by Arvydas Matiukas
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Hi Arvydas, You want to keep your pinhole fully open for two-photon imaging..... in two photon mode, you are exciting one single spot in the specimen, so there is no need to have a pinhole to exclude out of focus light, since there isn't any. You want the pinhole open to maximize signal collection. Do not trust the estimate of the optical slice thickness in two photon mode that the LSM software gives you. The voxel size for two photon excitation is defined by the wavelength you use for excitation, I'm guessing around 800 nm, so the radius of your laser spot is only 1.6 times greater for DAPI compared to excitation with 488 nm. The thickness of the optical slice is fixed for two photon (size of exciting laser spot). You can open the pinhole a little bit for the green chanel to try to match (I'm guessing to maybe 1.5 Airy units), so this would give you voxels that are not so different. Keep in mind that the accuracy of your colocalization will depend on other factors: how you threshold, how you subtract background, amount of noise in your images, etc.... If you are trying to "colocalize" structures that are significantly larger than the resolution of you microscope, the error contributed by the differences in voxel size will be comparatively small. If the structures you are comparing are near or below the resolution limit, then the error will get comparatively higher. As an extreme example, if you have two single molecules 100 nm apart, their images will mostly overlap and give a high degree of colocalization using conventional colocalization methods, while in reality the two molecules do not overlap at all. So my advice is to adjust teh pinhole for 488 to try to match the voxel size for DAPI as best as you can, and then use a sound colocalization method (and good controls) for your colocalization. Deconvolving the data might be a good idea to reduce noise and tighten your PSFs before running your analysis. I found the discussions on colocalization provided by SVI (scientific volume imaging) to be very useful. You can Google colocalization + SVI. to access their documentation. Just saw Brian's response, so some of the above is redundant. -- Julio Vazquez Fred Hutchinson Cancer Research Center Seattle, WA http://www.fhcrc.org On Apr 3, 2013, at 10:28 AM, Arvydas Matiukas wrote: > ***** > To join, leave or search the confocal microscopy listserv, go to: > http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy > ***** > > Hello list, > > > One of my Core users wants to measure colocalization between > DAPI labeled and FITC-immunolabeled structures in nuclei of > her fixed cells. Visual inspection shows good level of colocalization. > > However, what would be the correct way to quantify it. > Our confocal is LSM510 META NLO. No UV or 405nm laser > so two-photon mode at 780nm is used to excite blue dye > while FITC is excited at 488nm in confocal mode . At default > pinhole settings (1 airy unit for confocal and max=~10 A.u. > for two-photon) both blue and green signals are good > but the optical slices (and voxel sizes) differ by the factor of 10. > Letting the software (LSM) to equalize the slices at 0.6um reduces the > pinhole to 0.76 for confocal and 1.0 for two-photon. > Unfortunately at such small pinhole for two-photon the > blue signal is lost in noise (even at 5% laser power and 16 times > averaging, further increase may bleach the dye during 15 min > long scan). I can get the blue signal back to a measurable level > by increasing the two-photon pinhole to 4 A.u. but this creates > a four-fold mismatch in blue/green voxel size. I could boost the > blue signal by increasing pixel size but the loss in resolution > is not desirable. > > My first thought was to switch to red DNA stain and then > do colocalization between two confocal signals with identical > voxels. However, the user already completed experiment > using DAPI staining, and it worked well. My question is > what would be the correct way to acquire the confocal/two-photon > z-stacks and subsequently quantify the colocalization. > And related question if reducing pinhole works the same > way for two-photon as for a regular confocal. > > Any feedback/advice/idea are very welcome, > Arvydas > ---------------------------- > > > > Arvydas Matiukas, Ph.D. > Director of Confocal&Two-Photon Core > Department of Neurosci& Physiology > SUNY Upstate Medical University > 766 Irving Ave., WH 3167 > Syracuse, NY 13210 > tel.: 315-464-7997 > fax: 315-464-8014 > email: [hidden email] |
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In reply to this post by manish kumar-2
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Manish If you export all of your image data you should end up with a series of xy images at every z and at every time. You can then just pick out the xy images you want and for example in imagej make a sequence and generate an avi. Dr Lloyd Donaldson Project Leader - Microscopy & Wood Identification Senior Scientist - Plant Cell Walls & Biomaterials Scion - Forests, Products, Innovation 49 Sala Street, Rotorua 3010 New Zealand Ph 07 343 5581 www.scionresearch.com -----Original Message----- From: Confocal Microscopy List [mailto:[hidden email]] On Behalf Of Manish Kumar Sent: Wednesday, 3 April 2013 11:44 p.m. To: [hidden email] Subject: Re: xyzt image ***** To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Dear Kees, Thanks for the reply. I have tried the option. This works with the confocal software, but didnt work when i export it to .avi . Rgds Manish On Wed, Apr 3, 2013 at 3:45 PM, Straatman, Kees R. (Dr.) < [hidden email]> wrote: > ***** > To join, leave or search the confocal microscopy listserv, go to: > http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy > ***** > > Dear Manish, > > You can use Image > Stacks > Tools > Grouped Z-project... > > Best wishes > > Kees > > -----Original Message----- > From: Confocal Microscopy List > [mailto:[hidden email]] > On Behalf Of Manish Kumar > Sent: 03 April 2013 10:51 > To: [hidden email] > Subject: xyzt image > > ***** > To join, leave or search the confocal microscopy listserv, go to: > http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy > ***** > > Hello everyone, > > Is there any way to convert XYZT image to xyt. format. > > I have done live cell imaging with xyzt format and and I want to > export the same file to XYT.avi format. That means I want to compile > the z stack so that I can get the .avi image in xyt format. > I am using Leica SP5 confocal misroscope. > > Regards > Manish Kumar > Imaging facility Manager > IGIB, > New Delhi > This e-mail and any attachments may contain information which is confidential or subject to copyright. If you receive this e-mail in error, please delete it. Scion does not accept responsibility for anything in this e-mail which is not provided in the course of Scion's usual business or for any computer virus, data corruption, interference or delay arising from this e-mail. |
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phil laissue-2 |
Re: colocalization between confocal and two-photon: voxel matching
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In reply to this post by Arvydas Matiukas
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Hi Arvydas, further to the other two replies, I'll just add a few references for quantifying colocalisation. By no means a comprehensive list, but in my humble opinion some of the most user-friendly discussions/approaches. Really depends on the structures in question, there's not one single approach that works best. pixel-based: http://www.ncbi.nlm.nih.gov/pubmed/23026999 diva-portal.org/smash/get/diva2:563664/FULLTEXT02 coloc2: http://fiji.sc/Colocalization_Analysis http://www.ncbi.nlm.nih.gov/pubmed/21209361 http://www.ncbi.nlm.nih.gov/pubmed/15189895 object-based: http://www.ncbi.nlm.nih.gov/pubmed/20858446 http://crg.ubc.ca/moore/ http://www.ncbi.nlm.nih.gov/pubmed/19746416 http://www.ncbi.nlm.nih.gov/pubmed/23381680 (happy to send you reprint and matlab code) Also worth checking out: http://www.ncbi.nlm.nih.gov/pubmed/17210054 http://www.ncbi.nlm.nih.gov/pubmed/22086768 Hope this helps. Kind regards Philippe ____________________________________ Philippe Laissue, PhD, Bioimaging Manager School of Biological Sciences, Room 4.17 University of Essex, Colchester CO4 3SQ, UK (0044) 01206 872246 / (0044) 07842 676 456 [hidden email] privatewww.essex.ac.uk/~plaissue <http://privatewww.essex.ac.uk/%7Eplaissue> _____________________________________ Philippe Laissue, PhD, Bioimaging Manager School of Biological Sciences, Room 4.17 University of Essex, Colchester CO4 3SQ, UK (0044) 01206 872246 / (0044) 07842 676 456 [hidden email] privatewww.essex.ac.uk/~plaissue On Wed, Apr 3, 2013 at 6:28 PM, Arvydas Matiukas <[hidden email]>wrote: > ***** > To join, leave or search the confocal microscopy listserv, go to: > http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy > ***** > > Hello list, > > > One of my Core users wants to measure colocalization between > DAPI labeled and FITC-immunolabeled structures in nuclei of > her fixed cells. Visual inspection shows good level of colocalization. > > However, what would be the correct way to quantify it. > Our confocal is LSM510 META NLO. No UV or 405nm laser > so two-photon mode at 780nm is used to excite blue dye > while FITC is excited at 488nm in confocal mode . At default > pinhole settings (1 airy unit for confocal and max=~10 A.u. > for two-photon) both blue and green signals are good > but the optical slices (and voxel sizes) differ by the factor of 10. > Letting the software (LSM) to equalize the slices at 0.6um reduces the > pinhole to 0.76 for confocal and 1.0 for two-photon. > Unfortunately at such small pinhole for two-photon the > blue signal is lost in noise (even at 5% laser power and 16 times > averaging, further increase may bleach the dye during 15 min > long scan). I can get the blue signal back to a measurable level > by increasing the two-photon pinhole to 4 A.u. but this creates > a four-fold mismatch in blue/green voxel size. I could boost the > blue signal by increasing pixel size but the loss in resolution > is not desirable. > > My first thought was to switch to red DNA stain and then > do colocalization between two confocal signals with identical > voxels. However, the user already completed experiment > using DAPI staining, and it worked well. My question is > what would be the correct way to acquire the confocal/two-photon > z-stacks and subsequently quantify the colocalization. > And related question if reducing pinhole works the same > way for two-photon as for a regular confocal. > > Any feedback/advice/idea are very welcome, > Arvydas > ---------------------------- > > > > Arvydas Matiukas, Ph.D. > Director of Confocal&Two-Photon Core > Department of Neurosci& Physiology > SUNY Upstate Medical University > 766 Irving Ave., WH 3167 > Syracuse, NY 13210 > tel.: 315-464-7997 > fax: 315-464-8014 > email: [hidden email] > |
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Jeremy Adler-4 |
Re: colocalization between confocal and two-photon: voxel matching
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In reply to this post by Arvydas Matiukas
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Surprisingly actually measuring colocalization is surprisingly tricky - there are far too many coefficients and nonsense to several decimal places is still nonsense. It is also important to appreciate that colocalization addresses two separate questions (1) cooccurence, are the molecules in the same pixels and (2) correlation, is there any relationship between the intensities. These cannot be measured with a single coefficient. It is also important to appreciate that the quality, meaning noise, of the individual images seriously alters the measured colocalization. Try taking a second image and comparing it to the first - they are unlikely to be identical and for a correlation measurement this noise creates a difference between the measured and the true (noise free) correlation. A solution to the problem of noise is available for correlation measurements. look at http://diva-portal.org/smash/record.jsf?searchId=11&pid=diva2:563664 Jeremy Adler IGP Rudbeckslaboratoriet Daghammersköljdsväg 20 751 85 Uppsala Sweden 0046 (0)18 471 4607 |
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Andreas Bruckbauer |
Re: colocalization between confocal and two-photon: voxel matching
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In reply to this post by Julio Vazquez
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Hi Arvydas, make sure that the two-photon laser is aligned well with the visible laser, otherwise there will be a shift between the two channels, this is usually done during service of the microscope but quite often not with the best accuracy. For the optical slice of the two-photon signal it is important how large the laser spot at the back-aperture of the objective is, this can vary between setups and objectives (overfilling or underfilling) Best take z-stacks of some small beads and check how well the channels are aligned in x,y and z and the real size of the sections. You can then discuss the results with the microscope vendor or someone who knows how to set up such a system. best wishes Andreas -----Original Message----- From: Julio Vazquez <[hidden email]> To: CONFOCALMICROSCOPY <[hidden email]> Sent: Wed, 3 Apr 2013 19:14 Subject: Re: colocalization between confocal and two-photon: voxel matching ***** To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Hi Arvydas, You want to keep your pinhole fully open for two-photon imaging..... in two photon mode, you are exciting one single spot in the specimen, so there is no need to have a pinhole to exclude out of focus light, since there isn't any. You want the pinhole open to maximize signal collection. Do not trust the estimate of the optical slice thickness in two photon mode that the LSM software gives you. The voxel size for two photon excitation is defined by the wavelength you use for excitation, I'm guessing around 800 nm, so the radius of your laser spot is only 1.6 times greater for DAPI compared to excitation with 488 nm. The thickness of the optical slice is fixed for two photon (size of exciting laser spot). You can open the pinhole a little bit for the green chanel to try to match (I'm guessing to maybe 1.5 Airy units), so this would give you voxels that are not so different. Keep in mind that the accuracy of your colocalization will depend on other factors: how you threshold, how you subtract background, amount of noise in your images, etc.... If you are trying to "colocalize" structures that are significantly larger than the resolution of you microscope, the error contributed by the differences in voxel size will be comparatively small. If the structures you are comparing are near or below the resolution limit, then the error will get comparatively higher. As an extreme example, if you have two single molecules 100 nm apart, their images will mostly overlap and give a high degree of colocalization using conventional colocalization methods, while in reality the two molecules do not overlap at all. So my advice is to adjust teh pinhole for 488 to try to match the voxel size for DAPI as best as you can, and then use a sound colocalization method (and good controls) for your colocalization. Deconvolving the data might be a good idea to reduce noise and tighten your PSFs before running your analysis. I found the discussions on colocalization provided by SVI (scientific volume imaging) to be very useful. You can Google colocalization + SVI. to access their documentation. Just saw Brian's response, so some of the above is redundant. -- Julio Vazquez Fred Hutchinson Cancer Research Center Seattle, WA http://www.fhcrc.org On Apr 3, 2013, at 10:28 AM, Arvydas Matiukas wrote: > ***** > To join, leave or search the confocal microscopy listserv, go to: > http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy > ***** > > Hello list, > > > One of my Core users wants to measure colocalization between > DAPI labeled and FITC-immunolabeled structures in nuclei of > her fixed cells. Visual inspection shows good level of colocalization. > > However, what would be the correct way to quantify it. > Our confocal is LSM510 META NLO. No UV or 405nm laser > so two-photon mode at 780nm is used to excite blue dye > while FITC is excited at 488nm in confocal mode . At default > pinhole settings (1 airy unit for confocal and max=~10 A.u. > for two-photon) both blue and green signals are good > but the optical slices (and voxel sizes) differ by the factor of 10. > Letting the software (LSM) to equalize the slices at 0.6um reduces the > pinhole to 0.76 for confocal and 1.0 for two-photon. > Unfortunately at such small pinhole for two-photon the > blue signal is lost in noise (even at 5% laser power and 16 times > averaging, further increase may bleach the dye during 15 min > long scan). I can get the blue signal back to a measurable level > by increasing the two-photon pinhole to 4 A.u. but this creates > a four-fold mismatch in blue/green voxel size. I could boost the > blue signal by increasing pixel size but the loss in resolution > is not desirable. > > My first thought was to switch to red DNA stain and then > do colocalization between two confocal signals with identical > voxels. However, the user already completed experiment > using DAPI staining, and it worked well. My question is > what would be the correct way to acquire the confocal/two-photon > z-stacks and subsequently quantify the colocalization. > And related question if reducing pinhole works the same > way for two-photon as for a regular confocal. > > Any feedback/advice/idea are very welcome, > Arvydas > ---------------------------- > > > > Arvydas Matiukas, Ph.D. > Director of Confocal&Two-Photon Core > Department of Neurosci& Physiology > SUNY Upstate Medical University > 766 Irving Ave., WH 3167 > Syracuse, NY 13210 > tel.: 315-464-7997 > fax: 315-464-8014 > email: [hidden email] |
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Pascal Weber |
Re: colocalization between confocal and two-photon: voxel matching
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In reply to this post by Arvydas Matiukas
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** In fact for colocalization signals, DAPI and FITC, just use the laser 2P. You'll always have a perfect colocalization. The fluorescence comes as a single dot. Why would you want to use two different lasers? For the LSM510 the only way to adjust the filling of the rear lens you have to ajust the beam size with a lens. I did it and i can modulate the beam size and the uniformity of the field. Regards, Pascal |
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In reply to this post by Glen MacDonald-2
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Dear Manish, Sorry for the confusion, I did not notice carefully via which mailing list your question was posted. I indeed referred to a program called Fiji (Fiji Is Just ImageJ) which you can download from http://fiji.sc/Fiji for free. It has many powerful option for image analysis and can read most microscope file formats. It is a distribution of a program called ImageJ, so besides information on the Fiji website the manual etc can be found on the ImageJ website (http://imagej.nih.gov/ij/) under Docs. Hopes this helps to solve your problem. Best wishes Kees -----Original Message----- From: Confocal Microscopy List [mailto:[hidden email]] On Behalf Of Glen MacDonald Sent: 03 April 2013 16:01 To: [hidden email] Subject: Re: xyzt image ***** To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** I believe Kees was referring the Grouped Z-Project in Fiji/ImageJ, not something in your confocal software. Import your Leica file into Fiji with the LOCI plugin. The Grouped Z-Project will create a new stack in which each frame is the z-projection of the corresponding time point. There are several routes to save that as an AVI from Fiji/ImageJ. It is advisable to also archive the projection movie in an uncompressed format in case you want to edit it in the future. In fact, you can add timestamps and annotations in Fiji. Please refer to documentation: http://imagej.nih.gov/ij/docs/guide/index.html Regards, Glen Glen MacDonald Core for Communication Research Virginia Merrill Bloedel Hearing Research Center Cellular Morphology Core Center on Human Development and Disability Box 357923 University of Washington Seattle, WA 98195-7923 USA (206) 616-4156 [hidden email] [hidden email] On Apr 3, 2013, at 3:44 AM, Manish Kumar <[hidden email]> wrote: > ***** > To join, leave or search the confocal microscopy listserv, go to: > http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy > ***** > > Dear Kees, > Thanks for the reply. I have tried the option. This works with the > confocal software, but didnt work when i export it to .avi . > > Rgds > Manish > > > On Wed, Apr 3, 2013 at 3:45 PM, Straatman, Kees R. (Dr.) < > [hidden email]> wrote: > >> ***** >> To join, leave or search the confocal microscopy listserv, go to: >> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy >> ***** >> >> Dear Manish, >> >> You can use Image > Stacks > Tools > Grouped Z-project... >> >> Best wishes >> >> Kees >> >> -----Original Message----- >> From: Confocal Microscopy List >> [mailto:[hidden email]] >> On Behalf Of Manish Kumar >> Sent: 03 April 2013 10:51 >> To: [hidden email] >> Subject: xyzt image >> >> ***** >> To join, leave or search the confocal microscopy listserv, go to: >> http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy >> ***** >> >> Hello everyone, >> >> Is there any way to convert XYZT image to xyt. format. >> >> I have done live cell imaging with xyzt format and and I want to >> export the same file to XYT.avi format. That means I want to >> compile the z stack so that I can get the .avi image in xyt format. >> I am using Leica SP5 confocal misroscope. >> >> Regards >> Manish Kumar >> Imaging facility Manager >> IGIB, >> New Delhi >> |
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In reply to this post by manish kumar-2
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Hello I did not find any differences between Images>Stacks>Z project... and Images>Stacks>Tools>Grouped Z project...". Can anybody explain why ? |
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Arvydas Matiukas |
Re: colocalization between confocal and two-photon: voxel matching
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In reply to this post by Pascal Weber
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Hi Pascal, Thanks for the suggestion to acquire both DAPI and FITC signals in 2P mode. Actually I typically use double DAPI/FITC configuration with FITC channel disabled. I agree that by exciting both DAPI and FITC at the same 2P wavelength provides a perfect colocalization. I will tweak the settings to get less noisy images. I did not initially use FITC acquisition in 2P mode because this mode yields more noisy signal compared with confocal. Additionally, LSM software was giving incorrect optical slice for 2P mode which confused me. Best regards, Arvydas >>> Pascal Weber <[hidden email]> 4/4/2013 2:10 AM >>> ***** To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** In fact for colocalization signals, DAPI and FITC, just use the laser 2P. You'll always have a perfect colocalization. The fluorescence comes as a single dot. Why would you want to use two different lasers? For the LSM510 the only way to adjust the filling of the rear lens you have to ajust the beam size with a lens. I did it and i can modulate the beam size and the uniformity of the field. Regards, Pascal |
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** This is a REALLY bad idea. Exciting FITC at a 2P wavelength that will also excite DAPI will either give you ultra-rapid fading of your FITC or pathetic signal from your DAPI or (in my experience) both. If you look a the fluorescein spectra from the Webb lab, you'll see that there is a dual peak, one at about 910-920 the other at around 760. This means that the shorter one is clearly accessing a higher excited state. If you excite FITC at the 910 peak you'll get good fluorescence that does not fade. Guy -----Original Message----- From: Confocal Microscopy List [mailto:[hidden email]] On Behalf Of Arvydas Matiukas Sent: Friday, 5 April 2013 6:26 AM To: [hidden email] Subject: Re: colocalization between confocal and two-photon: voxel matching ***** To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Hi Pascal, Thanks for the suggestion to acquire both DAPI and FITC signals in 2P mode. Actually I typically use double DAPI/FITC configuration with FITC channel disabled. I agree that by exciting both DAPI and FITC at the same 2P wavelength provides a perfect colocalization. I will tweak the settings to get less noisy images. I did not initially use FITC acquisition in 2P mode because this mode yields more noisy signal compared with confocal. Additionally, LSM software was giving incorrect optical slice for 2P mode which confused me. Best regards, Arvydas >>> Pascal Weber <[hidden email]> 4/4/2013 2:10 AM >>> ***** To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** In fact for colocalization signals, DAPI and FITC, just use the laser 2P. You'll always have a perfect colocalization. The fluorescence comes as a single dot. Why would you want to use two different lasers? For the LSM510 the only way to adjust the filling of the rear lens you have to ajust the beam size with a lens. I did it and i can modulate the beam size and the uniformity of the field. Regards, Pascal |
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In reply to this post by Olivier Bardot
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Dear Olivier, Yes in many cases they are the same. However, I think the difference is if you have a file with a combination of Z-stacks and t-series not displayed as a hyperstack but as a stack with a single slider. In this case the Grouped Z-projection allows you to make a Z-projection per timepoint while Z-projection is not able to handle this. You of course can make first a hyperstack from this and then all is the same again... Best wishes Kees -----Original Message----- From: Confocal Microscopy List [mailto:[hidden email]] On Behalf Of Olivier Bardot Sent: 04 April 2013 10:17 To: [hidden email] Subject: Re: xyzt image ***** To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Hello I did not find any differences between Images>Stacks>Z project... and Images>Stacks>Tools>Grouped Z project...". Can anybody explain why ? |
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In reply to this post by Julio Vazquez
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To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** It is more complicated than this. In MP mode you get the sqrt2 (1.414) resolution increase from the squaring of the PSF. In confocal at 1 Airy unit you don't get this. Therefore 800nm excitation in MP is equivalent to 565nm in confocal. (565 being the mean of excitation & emission). In practice, therefore, confocal and MP can be regarded as equivalent. So opening the pinhole for the confocal collection would not be a good idea - all it would do is confuse the colocalization by allowing contamination from out of focus light. Guy Optical Imaging Techniques in Cell Biology by Guy Cox 2nd edition, 2012 CRC Press http://www.guycox.com/optical.htm ______________________________________________ Associate Professor Guy Cox, MA, DPhil(Oxon) Aust. Centre for Microscopy & Microanalysis, 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:[hidden email]] On Behalf Of Julio Vazquez Sent: Thursday, 4 April 2013 5:13 AM To: [hidden email] Subject: Re: colocalization between confocal and two-photon: voxel matching ***** To join, leave or search the confocal microscopy listserv, go to: http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy ***** Hi Arvydas, You want to keep your pinhole fully open for two-photon imaging..... in two photon mode, you are exciting one single spot in the specimen, so there is no need to have a pinhole to exclude out of focus light, since there isn't any. You want the pinhole open to maximize signal collection. Do not trust the estimate of the optical slice thickness in two photon mode that the LSM software gives you. The voxel size for two photon excitation is defined by the wavelength you use for excitation, I'm guessing around 800 nm, so the radius of your laser spot is only 1.6 times greater for DAPI compared to excitation with 488 nm. The thickness of the optical slice is fixed for two photon (size of exciting laser spot). You can open the pinhole a little bit for the green chanel to try to match (I'm guessing to maybe 1.5 Airy units), so this would give you voxels that are not so different. Keep in mind that the accuracy of your colocalization will depend on other factors: how you threshold, how you subtract background, amount of noise in your images, etc.... If you are trying to "colocalize" structures that are significantly larger than the resolution of you microscope, the error contributed by the differences in voxel size will be comparatively small. If the structures you are comparing are near or below the resolution limit, then the error will get comparatively higher. As an extreme example, if you have two single molecules 100 nm apart, their images will mostly overlap and give a high degree of colocalization using conventional colocalization methods, while in reality the two molecules do not overlap at all. So my advice is to adjust teh pinhole for 488 to try to match the voxel size for DAPI as best as you can, and then use a sound colocalization method (and good controls) for your colocalization. Deconvolving the data might be a good idea to reduce noise and tighten your PSFs before running your analysis. I found the discussions on colocalization provided by SVI (scientific volume imaging) to be very useful. You can Google colocalization + SVI. to access their documentation. Just saw Brian's response, so some of the above is redundant. -- Julio Vazquez Fred Hutchinson Cancer Research Center Seattle, WA http://www.fhcrc.org On Apr 3, 2013, at 10:28 AM, Arvydas Matiukas wrote: > ***** > To join, leave or search the confocal microscopy listserv, go to: > http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy > ***** > > Hello list, > > > One of my Core users wants to measure colocalization between > DAPI labeled and FITC-immunolabeled structures in nuclei of > her fixed cells. Visual inspection shows good level of colocalization. > > However, what would be the correct way to quantify it. > Our confocal is LSM510 META NLO. No UV or 405nm laser > so two-photon mode at 780nm is used to excite blue dye > while FITC is excited at 488nm in confocal mode . At default > pinhole settings (1 airy unit for confocal and max=~10 A.u. > for two-photon) both blue and green signals are good > but the optical slices (and voxel sizes) differ by the factor of 10. > Letting the software (LSM) to equalize the slices at 0.6um reduces the > pinhole to 0.76 for confocal and 1.0 for two-photon. > Unfortunately at such small pinhole for two-photon the > blue signal is lost in noise (even at 5% laser power and 16 times > averaging, further increase may bleach the dye during 15 min > long scan). I can get the blue signal back to a measurable level > by increasing the two-photon pinhole to 4 A.u. but this creates > a four-fold mismatch in blue/green voxel size. I could boost the > blue signal by increasing pixel size but the loss in resolution > is not desirable. > > My first thought was to switch to red DNA stain and then > do colocalization between two confocal signals with identical > voxels. However, the user already completed experiment > using DAPI staining, and it worked well. My question is > what would be the correct way to acquire the confocal/two-photon > z-stacks and subsequently quantify the colocalization. > And related question if reducing pinhole works the same > way for two-photon as for a regular confocal. > > Any feedback/advice/idea are very welcome, > Arvydas > ---------------------------- > > > > Arvydas Matiukas, Ph.D. > Director of Confocal&Two-Photon Core > Department of Neurosci& Physiology > SUNY Upstate Medical University > 766 Irving Ave., WH 3167 > Syracuse, NY 13210 > tel.: 315-464-7997 > fax: 315-464-8014 > email: [hidden email] |
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