Organization:

The registration fee includes all scientific sessions, conference materials, morning and afternoon coffee/tea and lunch on Thursday, Friday and Saturday. Catering sites for coffee/tea are on the ground and the first floor. Catering sites for lunch are on the ground floor. Seats and tables during coffee and lunch breaks can be found on both floors. Sponsors' exhibition booths are located on both floors. Free WLAN access points (SSID: TIM2014, password LSSRCLEMBD1!) are available in the hallway and the lecture hall. One computer with stationary internet access can be found on the ground floor at the registration office. Entrance to the conference site is only possible with your name badge. No smoking inside the meeting building, please!

Organizers: Dr. Roland Nitschke Life Imaging Center (LIC) at the ZBSA Albert-Ludwigs-University Habsburgerstr. 49 D-79104 Freiburg phone: +49 (0) 761 203-2934 email: Roland.Nitschke@biologie.uni-freiburg.de

Dr. Stefan Eimer Center for Biological Signalling Studies Albert-Ludwigs-University Habsburgerstr. 49 D-79104 Freiburg phone: +49 761 203-97161 email: S.Eimer@eni-g.de

Conference Office Secretary LIC at the ZBSA Albert-Ludwigs-University Habsburgerstr. 49 D-79104 Freiburg phone: +49/761/203-2902 fax: +49/761/203-2941 email: lic@imaging.uni-freiburg.de

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Acknowledgements We gratefully thank the following sponsors for their contribution to this meeting: Academic and Scientific Institutions: Deutsche Forschungsgemeinschaft (DFG), Bonn Zentrum für Biosystemanalyse (ZBSA), Freiburg Bioss Center for Biological Signalling Studies, Freiburg Companies:

GOLD Sponsor of TIM 2014:

SILVER Sponsor of TIM 2014:

BRONZE Sponsor of TIM 2014:

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Program Thursday, 27.03.2014

09.00-09.15

Gunther Neuhaus, Vice Rector for Research Achim Tieftrunk (DFG) & Roland Nitschke (LIC)

Welcome

Superresolution I (Chair by S. Hell)

09.15 - 10.00 Stefan Hell (Göttingen, D)

Far-field optical nanoscopy: principles and recent advancements

10.00 - 10.30 Christoph Cremer (Mainz, D)

Localization Microscopy of Nuclear Nanostructures

10.30 - 11.00 Coffee Break

11.00 - 11.30 Ulrich Nienhaus (Karlsruhe, D)

Optical Nanoscopy of Biomolecular Structure and Dynamics

11.30 - 12.00 Alexander Rohrbach (Freiburg, D)

Fast, fluorescent and non-fluorescent super-resolution microscopy of living cell

12.00 - 12.30 Jörg Bewersdorf (Yale, USA)

Nanoscopy of stochastically switching molecules: How fast can we go?

12.30 - 13.45 Lunch

Correlative Microscopy (Chair: R. Wepf)

13.45 - 14.30 Roger Wepf (Zuerich, CH)

Bridging Microscopes: 3D correlative light and scanning electron microscopy of complex biological structures

14.30 - 15.00 Bram Koster (Leiden, NL)

Zooming in on cell architecture with cryo-correlative electron microscopy

15.00 - 15.30 Winfried Denk (München, D)

Towards the connectomic analysis of the whole mouse brain.

15.30 - 16.00 Coffee Break

16.00 - 16.30 Paul Verkade (Bristol, UK)

Correlative Light Electron Microscopy: 1 + 1 = 3

16.30 - 17.00 Eric Jorgensen (Salt Lake City, USA)

Super CLEM

17.30 - 18.00 Heinz Schwarz (Tübingen, D)

Affinity labeling of thin sections – still a versatile tool for correlative light and electron microscopy

18.00 - 19.00 all Wrap up and Open discussion

20:00 Speakers Dinner, location will be announced at the conference

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Program Friday, 28.03.2014

Light Sheet (Chair: E. Stelzer)

09.00 - 9.45 Ernst Stelzer (Frankfurt, D)

Minimally invasive studies of lateral root organogenesis deep inside the main root of Arabidopsis thaliana with Light Sheet-based Fluorescence Microscopy (LSFM)

09.45 - 10.15 Jan Huisken (Dresden, D)

Novel concepts in fluorescence microscopy

10.15 - 10.45 Ulrich Kubitscheck (Bonn, D)

Extended, sensitive and fast 3D-tracking of single molecules and RNA particles in living tissue

10.45 - 11.15 Coffee Break

11.15 - 11.45 Pablo Loza-Alvarez (Barcelona, E)

Light sheet microscope for fast volumetric imaging

11.45 - 12.15 Alipasha Vaziri (Vienna, A)

Towards a dynamic map of neuronal circuits

12.15 - 13.45 Lunch

Superresolution II/Techniques (Chair: M. Sauer)

13.45 - 14.30 Markus Sauer (Würzburg, D)

Eight Years of Single Molecule Localization Microscopy: From Concepts to Biological Impact

14.30 - 15.00 Kai Johnsson (Lausanne, CH)

New Fluorescent Probes for Life Cell Imaging

15.00 - 15.30 Rainer Heintzmann (Jena, D)

Structured Illumination and the Analysis of Single Molecules in Cells

15.30 - 16.00 Coffee Break

16.00 - 16.30 Rainer Uhl (Munich, D)

Hexagonal SIM with phasegratings and coherent light

16.30 - 17.00 Douglas Richardson (Cambridge, USA)

Rapid Imaging of large 3D volumes: applications of light sheet fluorescent microscopy

17.30 - 18.00 Walter Schubert (Magdeburg, D)

Imaging Cycler Microscopy (ICM): Translating the spatial protein network code into efficient therapies (Human Toponome Project)

18.00 - 18.30 Susan Gasser (Bale, CH)

Watching gene dynamics

18.30 - 20.00 all Wrap up and Open discussion with wine and snacks

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Program Saturday, 29.03.2014

Big Data (Chair: J. Swedlow)

09.00 - 09.45 Jason Swedlow (Dundee, UK)

The Open Microscopy Environment: Open Source Image Informatics for the Biological Sciences

09.45 - 10.15 Pavel Tomancak (Dresden, D)

Big Data Viewer for multiview SPIM microscopy data

10.15 - 10.45 Olaf Ronneberger (Freiburg, D)

Fast landmark detection and elastic registration for large volumetric data sets

10.45 - 11.15 Coffee Break

11.15 - 11.45 Hari Shroff (Bethesda, USA)

Sharper, faster, and gentler: improved 4D imaging for cells and embryos

11.45 - 12.15 Lars Hufnagel (Heidelberg, D)

Bioimaging across scales with light-sheet microscopy

12.15 - 12.45 Urban Liebel (Karlsruhe, D)

Social screening machines » learn from each other – challenge us..

12.45 - 13.15 Fernando Amat (J. Farms, USA)

Efficient processing and analysis of large-scale light-sheet microscopy data sets

13.15 - 14.00 Lunch and End of symposium

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Thursday 27.March 2014, 09.15 - 10.00

Far-field optical nanoscopy: principles and recent advancements Stefan W. Hell (Göttingen, D)

Throughout the 20th century it has been widely accepted that, at the end of the day, a light microscope relying on conventional lenses (far-field optics) cannot discern details that are finer than about half the wavelength of light (> 200 nm). However, in the 1990s, it was discovered that overcoming the diffraction barrier is realistic and that fluorescent samples can be resolved virtually down to molecular dimensions. Here we discuss the simple yet powerful principles that allow neutralizing the resolution-limiting role of far-field optical diffraction1,2. In a nutshell, features residing closer than the diffraction barrier are prepared in different molecular (quantum) states so that they are distinguishable for a brief detection period. As a result, the resolution-limiting role of diffraction is overcome, and the interior of transparent samples, such as living cells and tissues can now be imaged non-invasively at the nanoscale using focused light in 3D.

Besides discussing basic principles, we will show most recent advancements. In particular, we demonstrate massive parallelization of RESOLFT and STED recording using simple patterns of light, by more than 100,000 fold3. Likewise, we demonstrate the relevance of emerging ‘far-field optical nanoscopy’ to various areas, especially to the life and the material sciences.

References 1. Hell, S.W. Far-Field Optical Nanoscopy. Science 316, 1153-1158 (2007).

2. Hell, S.W. Microscopy and its focal switch. Nature Methods 6, 24-32 (2009).

3. Chmyrov, A. et al. Nanoscopy with more than 100,000 'doughnuts'. Nature Methods (2013).

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Thursday 27.March 2014, 10.00 - 10.30

Localization Microscopy of Nuclear Nanostructures Christoph Cremer (Mainz, D)

The spatial organization of the genome in the interphase nucleus has far reaching functional consequences for gene regulation. Recently, various methods of superresolution light microscopy have been developed which made possible to enhance the analysis of nuclear structures far beyond the conventional limits of about 200 nm in the object plane and 600 nm along the optical axis. Here, we shall report on quantitative nuclear nanostructure analysis based on Structured Excitation Illumination/Structured Illumination Microscopy (SEI/SIM), and on Spectrally Assigned Localization Microscopy (SALM), respectively, using home-built microscope systems. Presently, these approaches allow us to optically resolve nuclear structures down to the range of ca. 120 nm laterally/350 nm axially using structured illumination, and few tens of nanometer in 3D using a special variant of localization microscopy, Spectral Precision Distance/Position Determination Microscopy (SPDM). Both SIM and SPDM techniques were also combined in a single microscope setup. Application examples will be presented on the use of such ‘nanoscopy‘ approaches to perform quantitative analyses of individual small chromatin domains labelled by Fluorescence-in situ Hybridization(FISH); of fluorescence-labelled replication units; of repair foci induced by individual accelerated heavy ions; of Fluorescent-Protein (GFP/YFP/mRFP) tagged histones and chromatin remodeling proteins; or of immunolabelled transcription/splicing related nanostructures. In addition, we report on the direct high resolution SPDM of nuclear DNA distribution, localizing up to several million individual DNA sites in optical sections of mammalian cell nuclei. Some perspectives of these novel, quantitative “superresolution” microscopy methods for deciphering the ”4D Nucleome“ will be discussed.

References S. Rossberger, G. Best, D. Baddeley, R. Heintzmann, U. Birk, S. Dithmar, C. Cremer (2013) Combination of structured illumination and single molecule localization microscopy in one setup, J. Opt. 15, doi:10.1088/2040-8978/15/9/094003.

C. Cremer, Optics far Beyond the Diffraction Limit: From Focused Nanoscopy to Spectrally Assigned Localization Microscopy (2012). In: Springer Handbook of Lasers and Optics, 2nd edition (F. Träger, Edit.), pp. 1351 – 1389.

Y. Markaki, M. Gunkel, L. Schermelleh, S. Beichmanis, J. Neumann,M. Heidemann, H. Leonhardt, D. Eick, C. Cremer, T. Cremer, Functional nuclear organization of transcription and DNA replication: a topographical marriage between chromatin domains and the interchromatin compartment (2010) Cold Spring Harbor Symposia on Quantitative Biology 75:475 – 492.

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Thursday 27.March 2014, 11.00 - 11.30

Optical Nanoscopy of Biomolecular Structure and Dynamics Gerd Ulrich Nienhaus (Karlsruhe, D)

Enormous advances have been made in fluorescence-based optical nanoscopy since the introduction of STED superresolution imaging by Stefan Hell 20 years ago. In this lecture, I shall present two recent contributions to this field from our laboratory.

Localization microscopy (PALM/STORM) is a widefield technique that is conceptually simple and easy to implement. Individual fluorescence emitters are activated stochastically by light and are subsequently localized with a precision of 10 – 30 nm. The technique enables studies of live cell dynamics by using photoactivatable fluorescent proteins (paFPs) [1]. Unlike synthetic fluorophores, paFPs deliver only a small number of photons before photobleaching occurs, so the image reconstruction software has to faithfully identify molecules in the presence of significant, spatially varying background [2]. By using GPU-based computation, we have implemented a fairly complex analysis algorithm that runs on a PC and still keeps up with the data flow from state-of-the-art CCD or sCMOS cameras [3].

STED/RESOLFT microscopy is a scanning technique that uses especially shaped focal patterns to reduce the effective point-spread function, either by using stimulated emission or photoactivation of fluorophores for spatially selective depletion of the excitation. Fast dynamics such as molecular diffusion can be probed by using fluorescence fluctuation techniques such as fluorescence correlation spectroscopy (FCS) and raster image correlation spectroscopy (RICS). For both methods, the combination with STED offers key advantages, particularly the extension of the marker concentration up to the micromolar range and the increased spatial resolution [4].

References 1. J. Fuchs, S. Böhme, F. Oswald, P. N. Hedde, M. Krause, J. Wiedenmann, and G. U. Nienhaus, Nat. Methods 7, 627-630 (2010).

2. P. N. Hedde, J. Fuchs, F. Oswald, J. Wiedenmann, and G. U. Nienhaus, Nat. Methods 6, 689-690 (2009).

3. Y. Li, P. N. Hedde, Y. Ishitsuka and G. U. Nienhaus, ACS Nano 7, 5207-5214 (2013).

4. P. N. Hedde, R. M. Dörlich, R. Blomley, D. Gradl, E. Oppong, A. C. B. Cato and G. U. Nienhaus, Nat. Commun. 4, 2093 (2013).

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Thursday 27.March 2014, 11.30 - 12.00

Fast, fluorescent and non-fluorescent super-resolution microscopy of living cell Alexander Rohrbach (Freiburg, D)

The smaller the biological structures, the less friction they produce and the faster they move due to thermal forces and active intracellular forces. The smaller the structure, the more optical resolution is required for imaging. However, the higher the required image resolution, the more photons have to interact with the structures. And this costs time and fluorophores!

In this talk I will present a fast super-resolution imaging variant based on total internal reflection fluorescence (TIRF) and structured illumination microscopy (SIM), which will be used to reveal the complicated dynamics of small MreB filaments inside living bacteria.

Second, I will show that coherently scattered laser light does not necessarily destroy an image because of too many speckles, but can provide images of very high resolution and contrast also in cell biology. This new technique, used in TIR mode, is applied to image macrophages uptaking particles.

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Thursday 27.March 2014, 12.00 - 12.30

Nanoscopy of stochastically switching molecules: How fast can we go? Jörg Bewersdorf (Yale, USA)

Nanoscopy techniques that utilize stochastic switching to achieve precise and accurate localization of a large number of single molecules achieve ~25 nm spatial resolution, far beyond the diffraction limit. However, the time resolution of these techniques has been limited to typically seconds to minutes due to the necessity of accumulating large numbers of localization estimates [1].

Using recently introduced sCMOS cameras, we have shown that much faster acquisition speeds up to 32 super-resolution images per second are possible [2]. Using a set of novel statistics-based algorithms specifically designed for sCMOS cameras allows us to perform unbiased and precise localization analysis at the Cramer-Rao lower bound.

I will present the underlying technology of ultra-fast single molecule switching nanoscopy, recent cell biological applications demonstrating its versatility as well as new quantitative data comparing image quality of conventional and high-speed imaging conditions.

Disclaimer: J.B. declares financial interest in Vutara Inc., a start-up company producing a fluorescence microscope utilizing 3D particle localization.

References [1] T.J. Gould, S.T. Hess, and J. Bewersdorf, “Optical nanoscopy: from acquisition to analysis,” Annu. Rev. Biomed. Eng. 14, 231-254 (2012).

[2] F. Huang, T.M.P. Hartwich, F.E. Rivera-Molina, Y. Lin, W.C. Duim, J.J. Long, P.D. Uchil, J.R. Myers, M.A. Baird, W. Mothes, M.W. Davidson, D. Toomre, J. Bewersdorf, “Video-rate nanoscopy using sCMOS camera–specific single-molecule localization algorithms”, Nature Methods 10(7), 653-658 (2013).

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Thursday 27.March 2014, 13.45 - 14.30

Bridging Microscopes: 3D correlative light and scanning electron microscopy of complex biological structures

MS Lucas, M Günthert, F Lucas, P Gasser, Anne-Greet Bitterman, Chris Buser and R Wepf (Zuerich, CH)

Correlative light and electron microscopy (CLEM) is a powerful tool in life science to combine large-scale volume imaging of cells or tissues by LM with a high-resolution description of their morphology using EM. The combination of 3D microscopy techniques such as CLSM with FIB-SEM or serial block face SEM (SBF-SEM) [1] or array tomography (AT) [2] opens up new possibilities to expand morphological context description and analysis into to the third dimension on the nm-scale.

These three approaches to correlate 3D LM data with SEM volume data have been shown to provide a valid alternative to TEM-based serial sectioning approaches [3]. Modern SEM-platforms allow imaging with an x/y-resolution of 2-3 nm, and offer the advantage of automation: e.g. routine overnight FIB-SEM volume imaging can cover cross-sections of up to 40×20 µm2. With a slice thickness of 5-10 nm, these stacks can expand to several 10 µm in the z-axis. FIB-SEM further enables precise, site-specific localisation and milling.

SBF-SEM offers more flexibility when choosing the volume size: theoretically the area to be scanned is limited only by the size of the block face. However, very large scan areas come at the cost of the x/y pixel size or extended scan time (BSE imaging). The slice thickness can be chosen from 15 - 200 nm to ideally match the biological question. Thus areas from several 100x100µm and several 100µm in depth (volumes: 10000 to 6*106 µm³) can be recorded [4].

In combination with prior screening of the specimen by confocal LM to identify the ROI, this allows targeted high-resolution 3D imaging of the structure of interest. This approach requires a sample preparation that facilitates the application of both LM and EM on a bulk specimen. To make this possible we have adapted protocols for freeze-substitution after high-pressure freezing, or chemical fixation and embedding in resin to include fluorophores in the samples [5].

Thus fluorescently labeled, resin-embedded specimens can also be used for AT. Here, ribbons of serial sections are loaded onto glass-slides, enabling even larger ROIs to be investigated by wide-field LM and subsequently by SEM. With the introduction of special surface coatings such as ITO these glass supports remain transparent for LM, but become highly conductive for SEM investigations. Such ribbons can be stained for histology (e.g. Toluidine blue or H&E) or for fluorescent immuno-labeling to

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specifically identify a ROI by F/LM prior to HR-SEM imaging. If needed, the sections can also be post-stained with uranyl acetate for (FIB)SEM-imaging.

Software solutions are available to facilitate the relocation of the ROI by a marker-based calibration of the sample in both LM and (FIB-)SEM. And automated recording of 3D volumes is possible for all three described methods. However, the FIB-SEM approach can achieve the better z-resolution, whereas the z-resolution for AT and SBF-SEM is limited by the sectioning process. FIB-SEM can therefore record isotropic voxels, which is an advantage when it comes to 3D reconstruction and modeling. SBF-SEM and AT on the other hand provide a much larger field of view. And AT even enables specific fluorescence labelling to identify the ROI combined with immune-gold labelling for HR-SEM. For SBF-SEM the ROI has to be defined using the information which can be obtained from the surface of the block face, either by LM or SEM. Furthermore, FIB-SEM and SBF-SEM are destructive methods, while the AT-ribbons can be stored and re-investigated ad libitum [6].

Resulting 3D datasets from LM and EM imaging have very different resolutions but can be merged in-silico to 3D models. By combining with other imaging modalities (e.g Xray-CT) one can reconstruct the structural hierarchical scaffold and hence better understand the complexity of living systems. Further transforming these structures into a visual perception space or haptic perception space by 3D printing the nano-world becomes better (comprehensive) accessible also for the layman. The method of choice has to be determined anew for every project, according the sample type, the size of the structure or the volume of interest and the desired imaging resolution in x,y, and z.

References [1] W Denk et al, PLoS Biol 2 (2004), p. e329.; [2] KD Micheva et al, Neuron 55 (2007), p. 25.

[3] MS Lucas et al, Methods Mol. Biol. 1117 (2014), p. 593.;[4] CJ Peddie et al, Micron (2014), p. epub ahead of print.; [5] SS Biel et al, J. Microsc. 212 (2003), p. 91.; [6] MS Lucas et al, Methods Cell Biol. 111 (2012), p. 325.

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Thursday 27.March 2014, 14.30 - 15.00

Zooming in on cell architecture with cryo-correlative electron microscopy R.I. Koning, F.G.A. Faas, M. Barcena, A.J. Koster (Leiden, NL)

In correlative light and electron microscopy (CLEM) imaging modalities are combined to study of cellular processes. Fluorescence light microscopy (FM) enables the imaging of dynamic events in relatively large fields of view exploiting a wide range of available fluorescent markers, while electron microscopy (EM) can reveal structural macromolecular arrangements in their cellular context in relatively narrow fields of view at nm-scale resolution. [1].

EM specimens prepared by conventional methods that incorporate chemical fixation and a metal staining steps can provide a wealth of information on the cellular architecture and processes. However, at the molecular level the fidelity of interpretation is limited because of effects related to the fixation and staining. Imaging of cryo-immobilized frozen-hydrated specimens excludes the use of stain and as such the molecular resolution is preserved. Unfortunately, images of frozen hydrated specimens have an inherent low contrast and signal to noise ratio because of their electron dose sensitivity and the lack of heavy atoms. These two drawbacks pose a hurdle to the possibility to identify areas of interest in specimens prepared for cryo electron microscopy. For applications of cryo electron tomography, fluorescence microscopy can be instrumental as a powerful tool for navigation, localization and selection of areas of interest..

We give an overview of on-going correlative approaches in our laboratory and report on technological developments and applications specifically related to cryo correlative microscopy. We have worked on three approaches to bridge the resolution gap between FM and EM imaging. First, we have built a cryo-immobilization device (plunger) that has a fluorescence microscope attached to it. The instrument makes it possible to perform live cell imaging of tagged structures up to seconds prior to plunging. Second, we have worked on the development of a next generation freezing stage that is suitable for upright light microscopes in collaboration with Linkam Company (CMS196). The freezing stage can be mounted on an optical system that can be chosen to be optimal for a particular type of light microscopy [2,3]. Third, we pursued the integration of a scanning FM in a TEM, allowing sequential FM and EM imaging with a single instrument (iLEM, iCorr™) [4,5].

Using these instruments a workflow can be defined. First, a series of fluorescence images are collected at liquid nitrogen temperature to acquire a map of the fluorescent signals. Next, the sample is observed in a cryo electron microscope and at

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specific areas, that are selected based upon the map of fluorescent signals, 3D electron microscopy images are recorded using cryo electron tomography.

We illustrate this workflow for cryo correlative microscopy by showing results obtained on several types of biological systems, in particular on fluorescently labelled bacteria.

References [1] Koster and Klumperman. Electron microscopy in cell biology: integrating structure and function. Nat Rev Mol Cell Biol. 2003.

[2] Briegel et al. Correlated light and electron cryo-microscopy. Methods Enzymol. 2010;481:317. [3] van Driel et al. Tools for correlative cryo-fluorescence microscopy and cryo-electron tomography applied to whole mitochondria in human endothelial cells. Eur J Cell Biol. 2009;88(11):669.

[4] Faas et al. Virtual nanoscopy: generation of ultra-large high resolution electron microscopy maps. J Cell Biol. 2012;198(3):457.

[5] Faas et al. Localization of fluorescently labeled structures in frozen hydrated samples using integrated light electron microscopy. J Struct Biol. 2013;181(3):283.

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Thursday 27.March 2014, 15.00 - 15.30

Towards the connectomic analysis of the whole mouse brain. Winfried Denk (München, D)

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Thursday 27.March 2014, 16.00 - 16.30

Correlative Light Electron Microscopy: 1 + 1 = 3 Paul Verkade (Bristol, UK)

Correlative Light Electron Microscopy combines the strengths of light and electron microscopy in one experiment and the sum of such an experiment should provide more data / insight than each technique alone (hence 1 + 1 = 3). There are many ways to perform a CLEM experiment and a variety of microscopy modalities can be combined. The choice of these instruments should primarily depend on the scientific question to be answered.

A CLEM experiment can usually be divided in 3 parts; probes, processing, and analysis. I will discuss 3 processing techniques based on light microscopy in conjunction with Transmission Electron Microscopy, each with its advantages and challenges.

The first is based on the use of coverslips with a finder pattern, it allows live cell imaging and captures an event of interest using chemical fixation, A second uses the Tokuyasu cryo immuno labelling to trace back objects of interest, this allows for relatively high immuno labelling efficiencies but is almost impossible in combination with live cell imaging. The third is based on cryo-fixation to obtain best possible preservation of ultrastructure. This allows us to capture events that would be lost because of chemical fixation, e.g. membrane tubules. It allows for live cell imaging but immuno labelling options are limited.

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Thursday 27.March 2014, 16.30 - 17.00

Super CLEM Eric Jorgensen (Salt Lake City, USA)

We are developing methods to correlate superresolution fluorescence images with electron micrographs of the same section, this technique is called ‘nanoresolution fluorescence electron microscopy’ (nano-fEM) or ‘superresolution correlative light and electron microscopy’ (super-CLEM). I will discuss improvements focusing on several aspects of our protocol. The most important change has been to use organic fluorescent dyes instead of fluorescent proteins, and using ground-state depletion instead of PALM or STED. These changes have allowed us to visualize multiple fluors. I will also discuss improved embedding resins, and the use of transmission electron microscopy instead of scanning electron microscopy. We are now working toward localizing our fluorescent dyes in 3D and correlating these images with 3D electron tomograms.

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Thursday 27.March 2014, 17.30 - 18.00

Affinity labeling of thin sections – still a versatile tool for correlative light and electron microscopy Heinz Schwarz and Bruno M. Humbel (Tübingen, D)

In correlative microscopy, light microscopy provides the overview and orientation in the complex cells and tissue, while electron microscopy offers the detailed localization and correlation to subcellular structures. High quality electron microscopical preparation methods provide optimal preservation of the cellular ultrastructure. From such preparations serial thin sections are collected and used for comparative histochemical, immunofluorescence and immunogold staining.

In light microscopy histological stains identify the orientation of the sample. Immunofluorescence labeling facilitates to identifying the region of interest, namely, the labeled cells expressing the macromolecule under investigation whereas colloidal gold visualize the label within the cellular architecture at high resolution provided by electron microscopy.

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Friday 28.March 2014, 09.00 - 9.45

Minimally invasive studies of lateral root organogenesis deep inside the main root of Arabidopsis thaliana with Light Sheet-based Fluorescence Microscopy (LSFM) Daniel von Wangenheim, Alexander Schmitz, Alexis Maizel, Ernst H.K. Stelzer (Frankfurt, D)

KEY WORDS: Plant, Arabidopsis, lateral root, lineage analysis, LSFM, SPIM, DSLM

In Arabidopsis thaliana, the root system consists of an embryo-derived primary root, from which a variable number of lateral roots evolve. The lateral root primordia originate from pericycle cells located deep within the parental root and have to pass through several tissue layers. Capturing the lateral root organogenesis from the first cell division to the emergence of a lateral root, requires non-invasive and viable imaging methods. In our light sheet-based fluorescence microscopes the plant grows in an upright position with its leaves in the air and its root in the medium. A timed light source matching the solar spectrum ensures photosynthetic activity and provides a diurnal cycle. Physiologic growth conditions enable dynamic studies of plant growth at organ, cellular and sub-cellular levels. We show insights gained by tracking all cells in time, space and angle over a period of up to three days. Our studies reveal a novel cell division pattern that has been confirmed in multiple independent investigations of different plants.

Arabidopsis lateral root grows out of the main root nuclei (red), plasma membrane (green), 3D-rendering with Arivis software

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References Vermeer, J. E. M., Wangenheim, D. von, Barberon, M., Lee, Y., Stelzer, E. H. K., Maizel, A., and Geldner, N. (2014). A spatial accommodation by neighboring cells is required for organ initiation in Arabidopsis. Science, 343(6167):178–183. PMID: 24408432.

Rosquete M R, Wangenheim D von, Marhavý P, Barbez E, Stelzer EHK, Benková E, Maizel A, and Kleine-Vehn J (2013) An Auxin Transport Mechanism Restricts Positive Orthogravitropism in Lateral Roots, Current Biology. doi:10.1016/j.cub.2013.03.064.

Lucas M, Kenobi K, Wangenheim D von, Voß U, Swarup K, Smet I D, Damme D V, Lawrence T, Péret, B, Moscardi E, Barbeau D, Godin C, Salt D, Guyomarch S, Stelzer EHK, Maizel A., Laplaze L., and Bennett M. J. (2013) Lateral root morphogenesis is dependent on the mechanical properties of the overlaying tissues, PNAS, doi: 10.1073/pnas.1210807110.

Maizel A*, Wangenheim D von, Federici F, Haseloff J, Stelzer EHK (2011) High resolution, live imaging of plant growth in near physiological bright conditions using light sheet fluorescence microscopy, The Plant Journal, 68 (2): 377-385.

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Friday 28.March 2014, 09.45 - 10.15

Novel concepts in fluorescence microscopy Jan Huisken (Dresden, D)

Light sheet microscopy has already overcome several limitations of established techniques such as confocal microscopy and offers plenty of new opportunities in fluorescence imaging. I will highlight two concepts that exploit the strengths of light sheet microscopy: (1) High-speed volumetric imaging with a deformable lens and (2) real-time data processing for data compression and visualization.

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Friday 28.March 2014, 10.15 - 10.45

Extended, sensitive and fast 3D-tracking of single molecules and RNA particles in living tissue Jan-Hendrik Spille, Tim Kaminski, Katharina Scherer, Jennifer S. Rinne, Alexander Heckel and Ulrich Kubitscheck (Bonn, D)

Tracking of fluorescently labeled molecules and particles can reveal detailed information about the kinetics of intracellular processes on the molecular level. Particle observation is usually limited to 2D projections of short trajectory segments. Here, we report real-time 3D single particle tracking in an active feedback loop with high photon efficiency. Axial coordinates were encoded in an astigmatic point-spread function and extracted from image data by a novel, fast localization scheme based on template matching. Results were obtained in real-time and used to keep a particle of interest close to the focal plane of a light sheet microscope. We were able to track particles carrying only 1-3 fluorophores more than 100 µm deep inside living tissue over an axial range of up to 10 µm with high spatio-temporal resolution. Several hundred localizations per trajectory enabled mobility analysis on the basis of individual particles and direct observation of transitions between mobility states.

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Friday 28.March 2014, 11.15 - 11.45

Light sheet microscope for fast volumetric imaging Pablo Loza-Alvarez (Barcelona, E)

Light sheet fluorescence microscopy (LSFM) is a convenient tool for bio-imaging as it efficiently collects the generated fluorescence while at the same time minimizes photobleaching.

Recently, the selective plane illumination microscopy (SPIM), being based on an intrinsic 2D illumination, has been put forward as an interesting candidate for fast volumetric imaging. In spite of this, fast SPIM implementations still rely on the movement of the sample or other optical elements to acquire the optical sections. This put constrains when imaging at high speeds, especially when working with living samples as moving artifacts could be adversely affect them. Overcoming this limitation, by scanning only the light-sheet instead, would provide real unperturbed, non-invasive imaging.

We report on the use of an alternative version of SPIM for fast 4D (3D+time) imaging where only the light sheet is scanned at speeds limited only by the integration time of the camera. This is done without any mechanical scan of the sample. The optical setup is based on the use of a deformable mirror that is placed in the detection path of the SPIM microscope.

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Friday 28.March 2014, 11.45 - 12.15

Towards a dynamic map of neuronal circuits

Alipasha Vaziri (Vienna, A)

Knowledge on structural connectivity in neuronal circuits is necessary for understanding information representation and processing in local circuits. However, as some examples of well-characterized neuronal architectures illustrate, structural connectivity alone it is not sufficient to predict how input stimuli are mapped onto activity patterns of neuronal populations and how the collective dynamics of all neurons in the network leads to behavior. Addressing this challenge has been hampered by lack of appropriate tools and methods that allow parallel and spatiotemporally specific application of excitation patterns onto neuronal populations while capturing the dynamic activity of the entire network at high spatial and temporal resolution. The combination of new optical excitation techniques, optogenetics and high speed functional imaging are providing new opportunities to address this question and move towards a dynamic map of neuronal circuits.

Most structural and functional imaging modalities, including two-photon imaging, rely on the manipulations of light fields in the spatial domain. However, pulsed excitation sources such as femtosecond lasers also possess a spectral width that provide an additional degree of freedom that can be used to “sculpt” other spatial light distributions. This has been exemplified in the technique of temporal of focusing through which sheet-like light distributions with an axial confinement close that of point scanning have been demonstrated. Using this approach we developed a two-photon technique for brain-wide calcium imaging in C. elegans [1]. The combination of this microscope with a nuclear-localized, genetically encoded calcium indicator, NLS-GCaMP5K, has allowed us to capture the activity of individual neurons within the densely packed head ganglia of C. elegans. We demonstrate near-simultaneous recording of activity of up to 70% of all head neurons. The application of this technique to other model organisms including rodents and zebrafish larvae is being currently pursued.

In order to functionally image even larger volumes and ultimately apply volumetric brain imaging in freely behaving animals we have recently established light-field microscopy in combination with 3D deconvolution [2]. Thereby we demonstrate intrinsically simultaneous volumetric Ca-imaging of neuronal population activity at single neuron resolution for the entire C. elegans organism. The simplicity of this technique and the possibility of the integration into epi-fluoresence microscopes

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make it an attractive tool for high-speed volumetric Ca-imaging. These tools combined with high speed optogentic control of neuronal circuits [3, 4] will be crucial to move from an anatomical wiring map towards a dynamic map of neuronal circuits.

References: 1. Schrodel, T., et al., Brain-wide 3D imaging of neuronal activity in Caenorhabditis elegans with sculpted light. Nature Methods, 2013. 10(10): p. 1013

2. Prevedel, R., et al., Simultaneuos whole-animal 3D-imaging of neuronal activity using light-field microscopy arXiv:1401.5333 (http://arxiv.org/abs/1401.5333).

3. Andrasfalvy, B., et al., Two-photon Single Cell Optogenetic Control of Neuronal Activity by Sculpted Light. Proceedings of the National Academy of Sciences of the United States of America, 2010. 107.

4. Losonczy, A., et al., Network mechanisms

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Friday 28.March 2014, 13.45 - 14.30

Eight Years of Single Molecule Localization Microscopy: From Concepts to Biological Impact Markus Sauer (Würzburg, D)

Super-resolution imaging by single molecule localization (localization microscopy) provides the ability to unravel the structural organization of cells and the composition of biomolecular assemblies at a spatial resolution that is well below the diffraction limit approaching virtually molecular resolution. Through constant improvements in fluorescent probes, efficient and specific labeling techniques as well as refined data analysis and interpretation strategies localization microscopy allows us today to interrogate how the distribution and stoichiometry of interacting proteins in subcellular compartments and molecular machines accomplishes complex interconnected cellular processes. Thus, it exhibits potential to address fundamental question of cell and developmental biology. Here, I briefly introduce the history and basic principles of localization microscopy methods with special focus on direct stochastic optical reconstruction microscopy (dSTORM), summarize recent key developments and application examples of two-dimensional and three-dimensional (3D) multicolor methods and correlative electron microscopy experiments, and discuss prospects of quantitative imaging to provide accurate counts of molecular copy numbers using standard fluorescent probes.

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Friday 28.March 2014, 14.30 - 15.00

New Fluorescent Probes for Life Cell Imaging Kai Johnsson (Lausanne, CH)

The ideal fluorescent probe for bioimaging is bright, absorbs at long wavelengths and can be implemented flexibly in living cells and in vivo. However, the design of synthetic fluorophores that combine all of these properties has proved to be extremely difficult. Here, we introduce a biocompatible near-infrared silicon–rhodamine probe that can be coupled specifically to proteins using different labelling techniques. Importantly, its high permeability and fluorogenic character permit the imaging of proteins in living cells and tissues, and its brightness and photostability make it ideally suited for live-cell super-resolution microscopy. The excellent spectroscopic properties of the probe combined with its ease of use in live-cell applications make it a powerful new tool for bioimaging.

I will also discuss new far-red, fluorogenic probes for imaging actin and tubulin in living cells based on the same fluorophore. By combining minimal cytotoxicity with excellent brightness and photostability, our probes facilitate fluorescence microscopy of the cytoskeleton in living cells and tissues. Applied in STED microscopy they reveal the nine-fold symmetry of the centrosome and the spatial organization of actin in the axon of cultured neurons with an unprecedented resolution of the cytoskeleton in living cells.

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Friday 28.March 2014, 15.00 - 15.30

Structured Illumination and the Analysis of Single Molecules in Cells Rainer Heintzmann (Jena, D)

The concept of a confocal microscope in which the pinhole is replaced with an array detector has been around for some time. In every position of the scanning laser beam, the image of the emitted fluorescence is detected. It has been shown that a preferable way of processing the so-obtained 4D data (scan position XY and detection position XY) is to assign the detected signal in the final image to a location placed half way between the nominal excitation and detection positions1. The reason is that this is the most probably position of the origin of the respective fluorescence. This results in an image, with enhanced resolution and detection efficiency, as compared to a standard confocal image. Recently this reassignment method has gained substantial attention2, also in the framework of multi-spot excitation3.

We here suggest ways which achieve a similar reassignment but in an all-optical way4. One possible realisation (Fig. 1) is to change the magnification between a de-scanning and a rescanning process. In stark contrast to computational

M2

BS

Large Pinhole

Scan/ Rescan

Unit

Sample

L1 L2

L3

TL

Objective

L4

L5 M4

M5 L6 Laser

M1

M3

Filte

sCMOS

Figure 1: Concept of optical reassignment: L4 and L5 have different focal length. All lenses in 4f arrangement. (M: plane mirror, L: lense, BS: dichromatic beam splitter)

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reassignment1-3, it is now not necessary to acquire an image for every scan position. A single image yields the efficient super-resolved result. Other ways to achieve a similar optical reassignment are to decouple the de-scan and re-scan process with the help of separate scan mirrors or to rescan twice. By tailoring the change in intermediate magnification, the system can be optimized for the influence of the Stokes-shift or even for realising a direct-view STED or RESOLFT microscope. We will discuss the theory of computational and optical reassignment and show experimental results. Concepts for realising optical reassignment for multi-beam scanners5 are also discussed.

References: [1] C. J. R. Sheppard, Optik 80, 53-54 (1988). [2] C. B. Müller and J. Enderlein, Physical Review Letters 104, 198101 (2010). [3] A. G. York et al., Nature Methods 9, 749-754 (2012). [4] S. Roth, et al., Optical Photon Reassignment microscopy, J. Opt. Nanosc., 2:5, doi:10.1186/2192-2853-2-5, 2013 [5] York et al. presented a talk at the Focus on Microscopy 2013 conference, which probably was using multi-beam optical reassignment as suggested here. However, no description was given what exactly was done and how.

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Friday 28.March 2014, 16.00 - 16.30

Hexagonal SIM with phasegratings and coherent light Rainer Uhl (Munich, D)

Key Words: Structured Illumination Microscopy, Superresolution SIM, confocal We will present a novel approach for Structured Illumination Microscopy (SIM), using 2-dimensional patterns with hexagonal symmetry. To obtain all necessary phase images for reconstructing sectioned superresolution images, patterns need to be shifted in one lateral direction only, no pattern rotation is required!

Our SIM approach employs a dynamic pattern which is as wide as the camera chip, but only a few lines high. It is swept over the sample synchronized with the rolling shutter window of a sCMOS camera, thus combining the advantages of structured illumination and slit-scan confocality. By exciting only a small fraction of the total field at a time and rejecting light from all other areas we gain contrast and reduce one of the major drawbacks of SIM, the pollution of small signal amplitudes by shot-noise from bright neighbors.

Superresolution SIM with a mouse antibody stained Tubulin sample. Left: a line confocal image. Right: a calculated Hexagonal Superresolution SIM image.

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Friday 28.March 2014, 16.30 - 17.00

Rapid Imaging of large 3D volumes: applications of light sheet fluorescent microscopy Douglas Richardson (Cambridge, USA)

Compound microscopes and their variants have a number of limitations when imaging 3D volumes in live samples, primarily: speed of acquisition, phototoxicity, photobleaching, difficulty maintaining environmental parameters such as temperature, and restrictions on sample orientation. Researchers are often presented a paradox in which they must either choose between fast imaging capabilities, while accepting poorer resolution; or increased resolution, and slow scan speeds. Fortunately, the current implementations of lightsheet fluorescence microscopy now provide researchers with a tool to image large 3D volumes, with minimal phototoxicity and bleaching, at high speed. Here, we present a number of data sets that have recently been acquired on a lightsheet microscope in the areas of small organism embryogenesis, tissue explant imaging, and clarified tissue imaging.

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Friday 28.March 2014, 17.30 - 18.00

Imaging Cycler Microscopy (ICM): Translating the spatial protein network code into efficient therapies (Human Toponome Project) Walter Schubert (Magdeburg, D)

Decoding the spatial system of biomolecular networks driving functionalities in morphologically intact cells and tissues (defined as the toponome) provides a fundamental challenge in biological sciences. It is the domain of microscopic techniques because it requires the co-mapping of large numbers of distinct biomolecules assembled as 3D structures (supermolecules and their networks). The co-mapping of > 100 distinct biomolecules has been shown by ICM (type TIS-Similarity Mapping; TIS-SIM) at the target sites of disease and at functional super-resolution. ICM is based on epifluorescence microscopy. It overcomes the spectral resolution limit by running robotically controlled incubation-imaging-bleaching cycles of one-dye-conjugated large probe libraries and thereby creates a random number of quasi-multi-fluorescence channels. This enables the collection of many thousand distinct combinatorial molecular informations per data point resulting in the detection of millions of distinct multimolecular domains in one image at 2D and 3D. The resulting parameter- and dimension-unlimited approach has been shown to allow for the simultaneous detection of: (i) the compositional spatial properties of thousands of distinct supermolecules (SM) in situ; and (ii) the spatial connectivities of such SM as networks. Surprisingly, recent higher-order data sets of SM networks uncover multicellular machines directly in human cancer tissue, which are built by many thousand distinct SM at a time. Such machines severely hamper the normal function of tissues by mechanically entrapping normal cells acting against the tumour and by introducing a novel system of SM order breaking the rules of tissue organization. This provides direct insight: (i) into SM which are components of these mechanisms; (ii) into novel drug-targets at these driving sites; (iii) into disease mechanisms that will help to explain failures of cell systems in cancers. Moreover, ICM has detected that the toponome is hierarchically organized with lead proteins controlling protein network topology and function (for overview: http://www.toposnomos.com/doku/MC2013.pdf). The corresponding HUTO initiative is an ICM-based independent collaboration with interdisciplinary activities in Europe, USA, and China.

Acknowledgement: funded by DFG ab initio in the early days in 1987 on the track to ICM invention (Schu 627/1 through 627/10); by the BMBF, the EU-FP7, and by the KTS, as well as the Land Saxony Anhalt.

Saturday 29.March 2014, 09.00 - 09.45

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Friday 28.March 2014, 18.00 - 18.30

Watching gene dynamics Susan Gasser (Bale, CH)

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Saturday 29.March 2014, 09.00 - 09.45

The Open Microscopy Environment: Open Source Image Informatics for the Biological Sciences Jason Swedlow (Dundee, UK) and the OME Consortium

Despite significant advances in cell and tissue imaging instrumentation and analysis algorithms, major informatics challenges remain unsolved: file formats are proprietary, facilities to store, analyze and query numerical data or analysis results are not routinely available, integration of new algorithms into proprietary packages is difficult at best, and standards for sharing image data and results are lacking. We have developed an open-source software framework to address these limitations called the Open Microscopy Environment (http://openmicroscopy.org). OME has three components—an open data model for biological imaging, standardised file formats and software libraries for data file conversion and software tools for image data management and analysis.

The OME Data Model (http://openmicroscopy.org/site/support/ome-model/) provides a common specification for scientific image data and has recently been updated to more fully support fluorescence filter sets, the requirement for unique identifiers, screening experiments using multi-well plates. The OME-TIFF file format (http://openmicroscopy.org/site/support/ome-model/ome-tiff) and the Bio-Formats file format library (http://openmicroscopy.org/site/products/bio-formats) provide an easy-to-use set of tools for converting data from proprietary file formats. These resources enable access to data by different processing and visualization applications, sharing of data between scientific collaborators and interoperability in third party tools like Fiji/ImageJ. The Java-based OMERO platform (http://openmicroscopy.org/site/products/omero) includes server and client applications that combine an image metadata database, a binary image data repository and visualization and analysis by remote access.

The current stable release of OMERO (OMERO-4.4; http://openmicroscopy.org/site/support/omero4/downloads) includes a single mechanism for accessing image data of all types-- regardless of original file format-- via Java, C/C++ and Python and a variety of applications and environments (e.g., ImageJ, Matlab and CellProfiler). This version of OMERO includes a number of new functions, including SSL-based secure access, distributed compute facility, filesystem access for OMERO clients, and a scripting facility for image processing. An open script repository allows users to share scripts with one another. A permissions system controls access to data within OMERO and enables sharing of data with users in a specific group or even publishing of image data to the worldwide community. Several applications that use OMERO are now released by the OME Consortium, including a

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FLIM analysis module, an object tracking module, two image-based search applications, an automatic image tagging application, and the first release of a biobanking application (http://www.openmicroscopy.org/site/products/partner). Our next version, OMERO-5 (http://openmicroscopy.org/site/products/ome5; currently available as a Release Candidate) includes updates and resources to specifically support large datasets that appear in digital pathology and high content screening. Importing these large datasets is fast, and data are stored in their original file format, so they can be accessed by 3rd party software.

OMERO and Bio-Formats run the JCB DataViewer (http://jcb-dataviewer.rupress.org/), the world’s first on-line scientific image publishing system and are used to publish 3D EM tomograms in the EMDataBank (http://emdatabank.org/). They also power several large institutional image data repositories (e.g., http://odr.stowers.org and http://lincs.hms.harvard.edu/).

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Saturday 29.March 2014, 09.45 - 10.15

Big Data Viewer for multiview SPIM microscopy data Pavel Tomancak (Dresden, D)

Handling large scale multi-dimensional and multimodal microscopy image data becomes an increasingly relevant problem as modern microscopy techniques produce amounts of data never encountered before. This applies most prominently to long-term, time-lapse datasets from light sheet microscopes, but also to other modalities such as serial section Electron Microscopy. We have developed tools to browse and interact with very large image volumes stored either locally or remotely using standalone software as well as online. The purpose of this presentation is to provide the technical details on the implementation of these new generation image browsing tools and to demonstrate their capabilities in a live demo. Depending on availability of fast internet connections this may or may not succeed. Nevertheless, I would like to use this presentation as an opportunity to introduce two active, open source projects: BigDataViewer under Fiji and CATMAID on the web, to attract more users and developers to these promising platforms.

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Saturday 29.March 2014, 10.15 - 10.45

Fast landmark detection and elastic registration for large volumetric data sets Olaf Ronneberger (Freiburg, D)

A central requirement to draw quantitative conclusions from microscopical data sets from different experiments is to map them to a common reference. Such a mapping can be achieved by an elastic registration of each dataset to a standard sample. Established registration techniques from the medical community often fail in microscopic applications due to the inhomogeneous data quality, severe deformations from different treatments and random orientation in automated high content experiments. Furthermore the runtime of available registration software packages is in the order of hours for a single volumetric image, which hinders their application for "big data" analysis.

In the talk I will present a trainable landmark detection, and a fast landmark initialized elastic registration that allow to overcome these shortcomings. The landmark detection learns the (possibly varying) appearance of certain landmarks from manually labeled training datasets. After training it is able to precisely detect these landmarks in new arbitrarily oriented datasets. The framework is based on rotation-equivariant filters, which can be computed within a few seconds on large datasets.

The obtained landmarks are then used to initialize an elastic registration algorithm. Our elastic registration uses locally normalized cross correlation combined with a fast combinatorial optimization algorithm.

This allows to perform a robust elastic registration within a few minutes. We used this framework successfully for registration of large collection of zebrafish larvae of different ages (48 hpf, 72 hpf and 96 hpf) with different preparation/staining procedures (Immunoflurescence, WISH).

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Figure: The landmark detection is performed in three steps. (a) Raw data. (b) Landmark probability map computed on down-sampled raw data using the trained filters (one filter per landmark type). (c,d) The best global constellation of all landmarks is found by a combinatorial optimization, using the learned pairwise distances of the landmarks. (e) In a final step local equivariant filters are applied to the high-resolution raw data to find the precise landmark positions. (f) Illustration of the underlying kernelized equivariant filter framework.

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Saturday 29.March 2014, 11.15 - 11.45

Sharper, faster, and gentler: improved 4D imaging for cells and embryos Hari Shroff (Bethesda, USA)

I will discuss our efforts to further develop high resolution optical methods that are better suited for the study of live, dynamic, and 3D samples. Structured illumination microscopy (SIM) doubles the spatial resolution of a light microscope, and requires lower light intensities and acquisition times than other super-resolution techniques, but has been almost exclusively applied to the study of single cells. I will present our multifocal SIM (MSIM) implementation that permits resolution doubling in live embryos and in volumes 5-8x thicker than possible with conventional SIM. A recent hardware modification that enables ‘instant’ SIM imaging at rates 10-100x faster than other SIM implementations will also be discussed.

The second half of the talk will focus on the development of inverted selective plane illumination microscopy (iSPIM), and subsequent application to the noninvasive study of neurodevelopment in nematode embryos. I will finish by discussing recent progress that quadruples the axial resolution of iSPIM by utilizing a second specimen view (‘diSPIM’), thus enabling imaging with isotropic spatial resolution.

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Saturday 29.March 2014, 11.45 - 12.15

Bioimaging across scales with light-sheet microscopy Lars Hufnagel (Heidelberg, D)

Selective-plane illumination microscopy has proven to be a powerful imaging technique due to its unsurpassed acquisition speed and gentle optical sectioning. We present a multiview selective-plane illumination microscope (MuVi-SPIM), comprising two detection and illumination objective lenses, that allows rapid in toto fluorescence imaging of biological specimens with subcellular resolution. The fixed geometrical arrangement of the imaging branches enables multiview data fusion in real time. However, even in the case of multi-view imaging techniques that illuminate and image the sample from multiple directions, light scattering inside tissues often severely impairs image contrast. Here we combine multi-view light-sheet imaging with electronic confocal slit detection (eCSD) on modern sCMOS sensors. In addition to improved imaging contrast, eCSD doubles the acquisition speed in multi-view setups with two opposing illumination directions as it allows for simultaneous dual-sided illumination without the otherwise inherent loss in image quality. This eliminates the need for specimen-specific data fusion algorithms and thus greatly reduces image post-processing, eases data handling and storage. The high speed of MuVi-SPIM allows faithful tracking of nuclei and cell shape changes, which we demonstrate through in toto imaging of the embryonic development of Drosophila melanogaster.

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Saturday 29.March 2014, 12.15 - 12.45

Social screening machines » learn from each other – challenge us.. Sven Eppler, David Feuerle, Lars Hufnagel, Sven Jacobi, Urban Liebel, Markus Riedesser, Paulo Queiros (Karlsruhe, D)

Modern robotic microscope platforms (High content screening platforms) are widely known for their potential in large scale cell based screens. Recent developments allow also large scale in vivo experiments e.g. in zebrafish and drosophila model organisms. These developments complement in many aspects cell based screens and open new insights e.g. in behavioral studies or complex physiological questions.

Logically not only BigData issues challenge us, but also the reproducibility of the platforms we create. Currently several experts are required to run successful screening platforms, which limits mass availability and reproducibility. Luckily we can learn a lot from existing technologies in other disciplines.

I will give an overview of our past and current developments, which improve reproducibility of smart data acquisition methods, storage and processing tricks.

This includes: precision motors with absolute encoded axis, real-time protocols for component control, software recycle methods, efficient software team developments, embedded / FPGA devices for software deployment, stable hardware design, social p2p machine networks for framework deployment, IT hardware design for storage processing and visualization of data w/o large server farms, methods to collaborate on data across different labs and “app driven” machine methods, which finally help many people in using newly generated detectors by themselves.

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Saturday 29.March 2014, 12.45 - 13.15

Efficient processing and analysis of large-scale light-sheet microscopy data sets Fernando Amat (J. Farms, USA)

The comprehensive reconstruction of cell lineages in complex multicellular organisms is a central goal of developmental biology. State-of-the-art fluorescence microscopes capture development at high spatio-temporal resolution for long periods of time. However, the complexity and size of the resulting data sets make systematic reconstructions of cell movements and divisions extremely challenging. We present a complete computational pipeline that efficiently stores, processes and analyzes such data on a single workstation, by taking advantage of multi-core and general purpose computation on graphical processing units (GPGPU). In particular, we introduce an open-source computational framework for segmentation and tracking of cell nuclei with high accuracy and speed. We demonstrate its (1) generality, by reconstructing cell lineages in terabyte-sized time-lapse data sets of fruit fly, zebrafish and mouse embryos, acquired with three different types of fluorescence microscopes, (2) scalability, by reconstructing cell lineages in advanced stages of development with up to 20,000 cells, within less time than needed for image acquisition, and (3) ease of use, by adjusting only two parameters across all data sets and providing visualization and editing tools for efficient data curation.

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List of participants Ackermann, Christian Universität Heidelberg Nikon Imaging Center BioQuant Im Neuenheimer Feld 167 Heidelberg/ Germany christian.ackermann@bioquant.uni-heidelberg.de

Amat, Fernando Howard Hughes Medical Institute Janelia Farm Research Campus 19700 Helix Drive 20147 Ashburn/ United States famat81@gmail.com

Avilov, Sergiy Max Planck Institute of Immunobiology and Epigenetics Imaging Facility Stubeweg 51 79108 Freiburg/ Germany avilov@ie-freiburg.mpg.de

Balazs, Balint EMBL Cell Biology and Biophysics Meyerhofstrasse 1 69117 Heidelberg/ Germany balint.balazs@embl.de

Bauch, Hubert Carl Zeiss Microscopy Sales Königsallee 9-21 37081 Göttingen/ Germany hubert.bauch@zeiss.com

Baumgart, Eugen University of Bonn Institute of Physical Chemistry Wegelerstr. 12 53115 Bonn/ Germany baumgart@pc.uni-bonn.de

Beaurepaire, Emmanuel Polytechnique - CNRS Lab for optics and biosciences route de Saclay 91128 Palaiseau/ France emmanuel.beaurepaire@polytechnique.edu

Bennett, Keith Hamamatsu Photonics K.K. Systems Division 2225 Webster St. 94301 Palo Alto/ United States kbennett@hamamatsu.com

Bergter, Annette Carl Zeiss Microscopy GmbH Product management Carl Zeiss Promenade 10 7745 Jena/ Germany annette.bergter@zeiss.com

Bertulat, Bianca Technische Universität Darmstadt Cell Biology & Epigenetics Schnittspahnstr. 10 64807 Darmstadt/ Germany bertulat@bio.tu-darmstadt.de

Bewersdorf, Joerg Yale School of Medicine Cell Biology 333 Cedar St. 6520 New Haven/ United States joerg.bewersdorf@yale.edu

Bourke, Steven The Friedrich Miescher Institute for Biomedical Research FAIM Maulbeerstrasse 66 4058 Basel/ Switzerland steven.bourke@fmi.ch

Brachner, Christoph PCO AG Marketing & Sales

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Donaupark 11 93309 Kelheim/ Germany christoph.brachner@pco.de

Brackmann, Maximilian Biozentrum, University of Basel Infection Biology Klingelbergstrasse 50/70 4056 Basel/ Switzerland maximilian.brackmann@unibas.ch

Braunstein, Thomas Core Facility for Integrated Microscopy Institute of Biomedical Sciences Panum Instituttet Blegdamsvej 3 Copehagen N/ Denmark thobra@sund.ku.dk

Burger, Gabriele Leica Microsystems CMS GmbH Application Am Friedensplatz 3 68165 Mannheim/ Germany gabriele.burger@leica-microsystems.de

Büttner, Lisa Rheinische Friedrich-Wilhelms-Universität Bonn Institute for Physical and Theoretical Chemistry Wegelerstr. 12 53115 Bonn/ Germany buettner@pc.uni-bonn.de

Charlet, Anne Universitätsherzzentrum Freiburg Bad Krozingen Kardiologie und Angiologie I Breisacherstr.33 79106 Freiburg/ Germany anne.charlet@universitaets-herzzentrum.de

Chiorean, Roxana Department of Molecular Dermatology II, University of Freiburg, Germany Molecular Dermatology II

Hauptstrasse 7, 79104, Freiburg, Germany 79104 Freiburg/ Germany roxana.chiorean@yahoo.com

Cicchetti, Gregor Wiley Global Research Boschstr. 12 69469 Weinheim/ Germany grcicche@wiley.com

Colombelli, Julien IRB Barcelona Advanced Digital Microscopy C. Baldiri Reixac, 10 Barcelona/ Spain julien.colombelli@irbbarcelona.org

conrad, christian DKFZ Theoretical Bioinformatics INF 256 69120 Heidelberg/ Germany c.conrad@dkfz.de

Cremer, Christoph Institute of Molecular Biology (IMB) Superresolution Microscopy 4 Ackermannweg 55128 Mainz/ Germany c.cremer@imb-mainz.de

De Mey, Jan CNRS-Univ. de Strasbourg, UMR 7213 Lab of Biophotonics and Pharmacology Tumoral Signalling route du Rhin, 74 67401 Illkirch/ France jan.de-mey@unistra.fr

den Hoedt, Sander Delmic CEO Molengraaffsingel 12-14 0 Delft/ Netherlands hoedt@delmic.com

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DITENGOU, Franck University of Freiburg Institute of Biology II, AG Palme Schänzlestrasse 1 79104 Freiburg/ Germany franck.ditengou@biologie.uni-freiburg.de

Eckerle, Stephanie University of Freiburg Developmental Biology Hauptstr.1 79104 Freiburg/ Germany stephanie.eckerle@biologie.uni-freiburg.de

Erhard, Birgit Imtek Bio- und Nanophotonik Prof. A. Rohrbach Sonnenstr. 5 79104 Freiburg/ Germany birgit.erhard@pharmazie.uni-freiburg.de

Fahrbach, Florian Max Planck Institute of Molecular Cell Biology and Genetics (MPI-CBG) Huisken Lab Pfotenhauerstr. 108 1307 Dresden/ Germany fahrbach@mpi-cbg.de

Fay, Nicolas Novartis Pharma AG NIBR Informatics & Technologies Postfach 0 Basel/ Switzerland nicolas.fay@novartis.com

Ferrand, Alexia University of Basel Imaging Core Facility Klingelbergstrasse 50/70 4056 Basel/ Switzerland alexia.ferrand@unibas.ch

Firaha, Tatsiana Universität Bonn

Institut für Physikalische und Theoretische Chemie Wegelerstrasse 12 53115 Bonn/ Germany t.p.firaha@gmail.com

Fouquet, Wernher Leica Microsystems CMS GmbH Application Development Am Friedensplatz 3 68165 Mannheim/ Germany wernher.fouquet@leica-microsystems.com

Friedrich, Martin Wiley-VCH Verlag Business Line Laboratory & Biotechnology Boschstrasse 12 69469 Weinheim/ Germany mfriedrich@wiley.com

Friedrich, Lars Leica Microsystems R&D Am Friedensplatz 3 68165 Mannheim/ Germany lars.friedrich@leica-microsystems.com

Gandhi, Hetvi Center for Systems Biology university Hospital freiburg, Renal Division Center for Systems Biology (ZBSA) Habsburgerstr. 49 79104 Freiburg/ Germany gandhi.hetvi@gmail.com

Gaub, Aline MPI Freiburg Chromatin Regulation Stuebeweg 51 79108 Freiburg/ Germany gaub@ie-freiburg.mpg.de

Gawlik, Jennifer University Freiburg BIOSS, Developmental biology (AG Pyrowolakis)

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Schänzlestr. 18 79104 Freiburg/ Germany jennygawlik@t-online.de

Gellner, Anne-Kathrin Universitätstklinik Freiburg Neurologie Breisacher Strasse 64 79106 Freiburg/ Germany anne-kathrin.gellner@uniklinik-freiburg.de

Gelman, Laurent Friedrich Miescher Institute for Biomedical Research Facility for Advance Imaging and Microscopy (FAIM) Maulbeerstrasse 66 4058 Basel/ Switzerland laurent.gelman@fmi.ch

Glass, Markus Institute of Molecular Medicine Section for Molecular Cell Biology Heinrich-Damerow-Str. 1 6120 Halle/ Germany markus.glass@medizin.uni-halle.de

Gohn-Kreuz, Cristian University of Freiburg Lab for Bio- and Nano-Photonics Georges-Köhler-Allee 102 79110 Freiburg/ Germany cgk@imtek.de

Goldner, Michael Leica Microsystems CMS GmbH R&D - Modules and Systems Am Friedensplatz 3 68165 Mannheim/ Germany michael.goldner@leica-microsystems.com

Graewe, Walter Hamamatsu Photonics Deutschland GmbH Systems Arzbergerstr. 10 82211 Herrsching/ Germany

wgraewe@hamamatsu.de

Graff, Alexandra Friedrich Miescher Institut (FMI), Basel Microscopy and Imaging facility Maulbeerstrasse, 66 4002 Basel/ Switzerland alexandra.graff@fmi.ch

Greb, Christoph Leica Microsystems GmbH Life Science Division - Marketing Ernst-Leitz-Strasse 35578 Wetzlar/ Germany christoph.greb@leica-microsystems.com

Groot, Edwin Universität Freiburg Institut für Biologie III Schänzlestr. 1 79104 Freiburg/ Germany edwin.groot@biologie.uni-freiburg.de

Gugel, Hilmar Leica Microsystems Development Optics Am Friedensplatz 2 68165 Mannheim/ Germany hilmar.gugel@leica-microsystems.com

Gwosch, Eva University of Konstanz Bioimaging Center Universitätsstrasse 10 78457 Konstanz/ Germany eva.gwosch@uni-konstanz.de

Hainmüller, Thomas University of Freiburg Physiology I Hermann Herder Str. 7 79104 Freibur/ Germany Thomas.Hainmueller@physiologie.uni-freiburg.de

Hanulova, Maria Institute of Molecular Biology Core Facility Microscopy

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Ackermannweg 4 55128 Mainz/ Germany m.hanulova@imb-mainz.de

Harter, Klaus University of Tübingen ZMBP / Plant Physiology Auf der Morgenstelle 32 72076 Tuebingen/ Germany klaus.harter@uni-tuebingen.de

Heintzmann, Rainer IPHT Microscopy Albert Einstein Str. 9 7745 Jena/ Germany heintzmann@gmail.com

Hell, Stefan W. Max Planck Institute for Biophysical Chemistry NanoBiophotonics Am Fassberg 11 37077 Göttingen/ Germany hell-office@gwdg.de

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Hoischen, Christian Leibniz Institute for Ageing Research Molecular Biology Beutenberg Str. 11 7745 Jena/ Germany hoischen@fli-leibniz.de

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Kindle, Petra MPI-IE Imaging Facility Stuebeweg 79108 Freiburg/ Germany kindle@ie-freiburg.mpg.de

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Kobler, Oliver Leibniz Institute for Neurobiology Combinatorial Neuroimaging (CNI) Brenneckestrasse 6 39118 Magdeburg/ Germany oliver.kobler@lin-magdeburg.de

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Koster, Abraham Leiden University Medical Center Molecular Cell Biology Einthovenweg 20 0 Leiden/ Netherlands a.j.koster@lumc.nl

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Loza-Alvarez, Pablo ICFO-The Institut of Photonics Sciences Biophotonics Av. Carl Friedrich Gauss, 3 8860 Castelldefels/ Spain pablo.loza@icfo.es

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