For the preparation of the data management concept of the Trends in Microscopy 2025 workshops, workshop providers were asked to report a list of ontology terms relevant to their workshop. When terms could not be found, providers were asked to specify the path where they would have expected to find the term (therefore, entries with a path in the tables are not present in the ontology).
I added links to the public data where available.
Workshop n°2: Exchangeable fluorescent probes for gentle long-time STED imaging
https://omero-tim.gerbi-gmb.de/webclient/?show=project-2952
abstract
Multi-colour imaging of cells and tissue is essential for the understanding of many biomedical research questions. Due to the small size of many subcellular organelles, there structures and dynamic interactions can be best visualized and analyzed using super-resolution light microscopy. Stimulated Emission Depletion (STED) microscopy is such technique which enables multi-colour super-resolution imaging in fixed [1,2] and in living cells [3].
Live-cell STED microscopy set high requirements on the fluorescent probes and requires gentle imaging routines. The fluorescent probes ideally should be bright and photostable to resist the high laser powers. Rhodamine dyes fulfill such properties and have been continuously optimized towards cell-permeability. To install a rhodamine label on any cellular target protein, genetically-encoded self-labeling protein tags like HaloTag, have become a powerful tool for live-cell microscopy. HaloTag undergoes a fast and specific labeling reaction with synthetic probes carrying a reactive ligand to form a covalent bond. This technology has recently been further developed by the development of exchangeable HaloTag Ligands. These probes bind to HaloTag with high affinity but can exchange regularly unlike the covalent labels [4]. In confocal and STED microscopy, this transient binding ensures that proteins can be detected in the living cell over extended periods, as bleaching processes are no longer crucial. Additionally, mutually orthogonal HaloTag7 protein/ligand pairs enabled to perform long-term dual-color imaging experiments.
In this workshop, participants will learn how sensitive samples like living cells benefit from optimized dye and imaging techniques. Participants will be introduced into novel exchangeable fluorescent probes like the abberior HaloX probes (Fig. 1). In addition, participants will get hands-on experience of how to use them for sample preparation and gentle long-time life cell imaging with reduced applied light dose.
References
[1] Hell, S.W., Wichmann, J., 1994. Opt. Lett. 19, 780
[2] Saal, S.J., Hell, S.W., Jakobs, S., 2017. Nat Rev. 18.
[3] Stockhammer, A., Bottanelli, F., 2020. J. Phys D: Appl. Phys. 54.
[4] Kompa, J., Bruins, J., Glogger M., Wilhelm, J., Frei, M.S., Tarnawski, M., D’Este, E., Heilemann, M., Johnsson, K., 2023. J. Am. Chem. Soc 145, 5.
| root term |
term label |
| resolution-enhancing method |
resolution-enhancing method stimulated emission depletion (STED) |
| visualization method |
Rhodamine |
| visualization method |
visualization method > visualization by chemical attribute > macromolecular probe > genetically encoded tag > HaloTag |
| visualization method |
visualization method > visualization by chemical attribute > macromolecular probe > genetically encoded tag > SNAPtag |
| visualization method |
visualization method > vizualization of label conjugated to probe > Exchangeable probes |
| imaged parameter |
imaged parameter > Photobleaching |
| imaged parameter |
imaged parameter > Phototoxicity |
| sample preparation method |
sample preparation method > Adherent live-cells |
Workshop n°4: Multimodal correlation spectroscopy of biomolecules in living cells
https://omero-tim.gerbi-gmb.de/webclient/?show=project-2955
abstract
Fluorescence and raster image correlation spectroscopy (FCS, RICS) are powerful tools to analyze biophysical properties of molecules in vitro as well as in living cells. The implementation of "array detector" FCS (1) and "arbitrary-region" RICS (2) on a commercial microscope (3, 4) have made the application of these techniques much more user-friendly. In or workshop we want to bring these seemingly difficult technologies closer to potential users. After theoretical introductions into the methodologies, we will assess in hands-on sessions the biophysical behavior (molecule concentration and diffuion rates) of EGFP multimeres at different locations within living cells. Benefits and potential pitfalls will be outlined in detail. After the session, users should be equipped with sufficient knowledge and starting expertise to establish the new concepts in their own labs.
(1) Scipioni et al., 2018, Nat. Comm. 9:5120, doi: 10.1038/s41467-018-07513-2
(2) Hendrix et al., 2016, Biophys. J. 111:1785, doi: 10.1016/j.bpj.2016.09.012
(3) https://www.zeiss.com/microscopy/en/products/software/zeiss-zen/dynamics-profiler.html
(4) https://www.zeiss.com/microscopy/en/products/software/zeiss-zen/spectral-rics.html
| root term |
term label |
| detection method |
photomultiplier tube (PMT) |
| illumination method |
single point scanning |
| imaged parameter |
fluorescence emission |
| imaging method |
array-scan confocal microscopy |
| visualization method |
EGFP |
| sample preparation method |
adherent cells in vitro > stably transfected |
Workshop n°8: FLIM-FRET: A powerful tool to visualize multimerization of plasticity-related gene 5 in living cells
No link to public dataset
abstract
Plasticity-related gene 5 (PRG5) is a membrane protein predominantly found in neurons and is involved in cellular processes such as growth cone guidance, migration and spine formation. Overexpression of PRG5 induces filopodia in non-neuronal cell lines, contributes to the induction of spines in immature neurons, and regulates spine density and morphology in mature neurons. Understanding the formation of spines is pivotal, as spine disruptions are associated with numerous neurological disorders. Although the importance of PRG5 in neuronal function is inevitable, the precise mechanisms how it exactly induces membrane protrusions and orchestrates cellular responses remain unresolved. We hypothesise that multimerization of PRG5 is required for its functionality. FLIM-FRET, or Fluorescence Lifetime Imaging Microscopy-Förster Resonance Energy Transfer, is an excellent approach to monitor dynamic protein-protein interaction and visualise their localisation in living cells with high tempo-spatial resolution. FRET is a physical, non-radiative process that occurs between an excited donor fluorophore and an acceptor fluorophore, while FLIM provides a sensitive approach to measure and quantify FRET. Once the acceptor is in close proximity (<10 nm) to the fluorescent donor, FRET energy transfer occurs. Due to this distance dependency, the two fluorophore labelled proteins undergoing FRET must physically interact. Here, we will use FLIM-FRET to detect PRG5 multimerization at the plasma membrane, particularly at the tips of filopodial protrusions in HEK cells. For the workshop, we will cooperate with the company PicoQuant, which will provide us with their new single photon counting confocal microscope, Luminosa, as well as with a Nikon based confocal laser scanning microscope including an upgrade kit.
| root term |
term label |
| detection method |
detection of UV/visible/IR photon |
| illumination method |
polarized light illumination |
| illumination method |
pulsed illumination |
| imaging method |
confocal microscopy |
| imaging method |
FLIM |
| imaging method |
FRET |
| source of contrast |
differences in fluorescence lifetime |
| visualization method |
EYFP |
| visualization method |
mTurquoise |
Workshop n°11: Monitoring activation of the GPCR NTSR1 in living cells by FLIM-heteroFRET and homoFRET
https://omero-tim.gerbi-gmb.de/webclient/?show=project-2961
abstract
The neurotensin receptor 1 (NTSR1) belongs to the class A of G protein-coupled receptors (GPCRs). Understanding the molecular mechanisms underlying these receptors is crucial for the development of therapeutic drugs. A central focus in the study of GPCR signaling pathways is the comparison between monomeric and oligomeric GPCRs as active signal transmitters. Our neurotensin receptor 1 (NTSR1) is fused with the bright fluorescent protein. NTSR1-mNeonGreen is expressed in living human HEK293T cells.
During the workshop, various lifetime-resolved hetero- and homoFRET imaging experiments to investigate changes in the oligomerization of NTSR1 following the addition of its ligand will be demonstrated. The technique employed for this purpose is FLIM-FRET, a powerful method used to explore molecular interactions within biological samples both in vitro and in vivo.
Monitoring oligomerization of NTSR1-mNeonGreen by homoFRET in living HEK293T cells:
Upon addition of the ligand neurotensin the oligomerization of the receptor within in plasma membrane is induced and will be monitored. This process involves the formation of receptor dimers. As dimers form, the distance between neighboring receptors decreases, leading to increased proximity between the fluorescent protein (mNeonGreen) molecules. This change can be measured using fluorescence anisotropy indicating homoFRET. Specifically, the NTSR1-mNeonGreen will be excited with a linear polarized pulsed 2-photon laser in a confocal microscope. The fluorescence is separated by polarization and simultaneously recorded by two ultra-sensitive detectors to achieve high-resolution time-correlated single photon counting and to analyze fluorescent lifetimes and time-resolved anisotropies.
Monitoring neurotensin binding demonstrated by heteroFRET:
To investigate neurotensin binding to NTSR1, a pair of donor- and acceptor-fluorophores is utilized. The donor is mNeonGreen fused to NTSR1. The acceptor-fluorophore, in this case, is a cyanine dye specifically bound to the ligand neurotensin. Changes in FRET efficiency will serve as indication for neurotensin binding to NTSR1. Additionally, it will be assessed whether the receptor-ligand complex remains intact during the internalization of receptor aggregates.
Our comprehensive approaches combine advanced microscopic imaging techniques, including FLIM-FRET, to unravel the intricate dynamics of NTSR1 oligomerization and ligand interactions.
| root term |
term label |
| contrast-enhancing method |
unprocessed raw |
| contrast-enhancing method |
fluorescence lifetime |
| contrast-enhancing method |
polarization contrast |
| detection method |
UV/visible/IR |
| detection method |
Hybrid PMT |
| detection method |
polarization-sensitive detection |
| illumination method |
coherent |
| illumination method |
pulsed |
| illumination method |
UV/visible/IR illumination |
| illumination method |
polarized light illumination |
| imaged parameter |
sub-nanosecond time-resolved fluorescence emission) |
| imaged parameter |
fluorescence polarization (polarization angle resolved fluorescence emission) |
| imaged parameter |
fluorescence lifetime FRET |
| imaging method |
single spot |
| imaging method |
FLIM |
| imaging method |
fluorescence polarization |
| imaging method |
two photon laser scanning |
| sample preperation method |
adherent living cells |
| fixation method |
ethanol fixed cells |
| sources of contrast |
differences in fluorecent lifetime |
| sources of contrast |
distribution of a specific protein |
| sources of contrast |
differences in anisotropy |
| sources of contrast |
heteroFRET |
| visualisation method |
mNeonGreen |
| visualisation method |
mScarlet |
| visualisation method |
Cy3B |
| visualisation method |
HiLyte647 |
Workshop n°26: The opportunity of a lifetime: Fluorescence Lifetime Imaging Microscopy in research, a guide from F to M
https://omero-tim.gerbi-gmb.de/webclient/?show=project-2976
abstract
Fluorescence Lifetime Imaging Microscopy (FLIM) is a well-established technique for analyzing dynamics and interactions of molecules and proteins in a wide range of samples and environments. Even if the technique is in principle available in laboratories already for quite some time, it was complicated to apply and often required the assistance of an expert to operate the device, measure the samples, and analyze the data.
With the recent releases from manufacturers of faster and easy to use devices, FLIM becomes more accessible to a larger audience. With acquiring FLIM images now being comparatively easy, the main challenge is to design and evaluate experiments in a meaningful way.
This workshop will use a Leica FALCON system to train and inform on what are the advantages and limitations of FLIM, and how to prepare experiments to obtain robust results. It will be discussed how FLIM can be a very advantageous technique over others in selected case. Several samples will be recorded and analyzed making use of the integrated software routines including phasor analysis. Interpretation of the data will be discussed.
Background:
With the development of sensitive and highly resolving instruments, fluorescence microscopy has become an essential tool of biological imaging. Its best-known feature is the specificity of its emission signal (fluorescence) that allows structures of interest to stand out against an otherwise dark background. In addition to its spectral properties for excitation and emission, a fluorophore is defined by a third property: the lifetime of its excited state before the return to the ground state by fluorescence emission. Since the emission of a fluorescence photon normally happens in the range of nanoseconds after excitation, dedicated instrumentation is needed for lifetime measurements, but fluorescence lifetime can add significant information that helps to understand the sample better. It is affected by the environmental conditions in the sample. Spectrally overlapping signals can be separated by their lifetime, allowing a better understanding of complex samples and the removal of background autofluorescence. Changes in de-excitation pathways (e.g. by FRET or STED) change the lifetime and can be used for distance measuremens at the molecular level and for improved resolution.
| root term |
term label |
| detection method |
detection of UV/visible/IR photons |
| detection method |
spot detector/hybrid detector |
| illumination method |
coherent illumination |
| illumination method |
UV/visible/IR illumination |
| illumination method |
pulsed illumination |
| imaged parameter |
sub-nanosecond time-resolved fluorescence emission |
| imaging method |
single-spot confocal microscopy |
| imaging method |
FLIM |
| resolution-enhancing method |
stimulated emission depletion (STED) |
| sample preparation method |
dispersed cells in vitro |
| sample preparation method |
formaldehyde fixed tissue |
| sample preparation method |
detergent permeabilized |
| source of contrast |
differences in fluorescence lifetime |
| vizualization method |
primary antibody plus labeled secondary antibody |
| vizualization method |
phalloidin |
| vizualization method |
Alexa Fluor 594 |
Workshop n°28: Hands-on light sheet microscopy with Flamingo, the shareable custom research microscope platform
https://omero-tim.gerbi-gmb.de/webclient/?show=project-2978
abstract
In the early 21st century, light sheet microscopy was introduced to the biological sciences and is now accepted as the new standard for fast and gentle 3D fluorescence imaging in living multicellular specimens. The core of our lab evolves around custom-built light sheet microscopy technology for challenging biological multiscale imaging projects. To streamline custom microscope development, we developed Flamingo, a powerful and modular microscope framework that is also compact and can be quickly deployed in collaborators’ labs. Over the last few years, the Flamingo has enabled novel biological experiments inside and outside our optics lab. As a significant part of the scientific community struggles to access a light sheet microscope tailored to their individual needs, the Flamingo gives more researchers the chance to utilize this powerful technology for their research.
In this workshop, we will introduce the concept of shared custom microscopy technology and demonstrate Flamingo's modularity. We will provide insights into light sheet microscopy and give you hands-on time with the microscope to better understand both the optics and the modes of operation. We will explain the relevant optics and discuss opportunities and challenges in light sheet microscopy, illustrated with image data recorded on the Flamingo. Live zebrafish embryos and larvae will be available to demonstrate and test Flamingo’s capabilities in living specimens.
| root term |
term label |
| detection method |
complementary metal oxide semiconductor (CMOS) |
| imaged parameter |
fluorescence emission |
| imaging method |
SPIM |
| illumination method |
UV/visible/IR illumination |
| sample preparation method |
whole mounted tissue |
| source of contrast |
compartmentalisation of stain or label |
| visualization method |
fluorescent label |
Workshop n°39: Simplifying reproducibility in high sensitivity TIRF microscopy to capture dynamics of single molecules
No link to public dataset
abstract
Well-defined and controlled imaging conditions are crucial for reproducible experiments with complex biological samples, such as live cell cultures and single molecule assays. Essential parameters include the homogeneity of optical excitation, temperature, gas composition, and relative humidity. Our workshops will demonstrate methods to precisely control these parameters using advanced sensor technologies commonly employed in the semiconductor industry but not yet widely adopted in optical microscopy. Attendees will gain insights into potential pitfalls in sensitive biophysical experiments and learn strategies to avoid them when designing their experiment.
Workshop 1: Waveguide-Based TIRF Microscopy for Ultra-Large Field of View Super-Resolution Imaging.
We will introduce the fundamentals of TIRF microscopy and experimentally quantify key physical parameters such as field homogeneity and penetration depth, both of which are crucial for quantitative single-molecule studies. The experiments will be conducted using one of our QUSCITE systems, which enables large field-of-view illumination over a few square millimeters. We will benchmark the system with dynamic DNA origami and stained live cell cultures. Alongside workshop participants, we will analyze our acquired imaging data using open-source software. By extracting sub-diffraction limited features, we will demonstrate the ease of generating reproducible single-molecule super-resolution data on any microscope.
Workshop 2: Precision and Dynamic Environmental Control for Single Particle Studies and Live Cell Imaging.
We will explore temperature and gas-sensitive processes at the single particle level, investigating how slight changes in temperature and gas composition can significantly impact experimental outcomes. Additionally, we will identify and address external factors that can substantially influence the temperature within the sample volume, potentially affecting the results if not properly managed. By integrating the AIROBOX stage-top cooling and heating unit with the VAHEAT micro-heating device and the TIRF module QUSCITE, we will demonstrate precise and rapid adjustments of temperature, humidity, oxygen, and carbon dioxide concentration on-site.
| root term |
term label |
| illumination method |
darkfield illumination |
| illumination method |
widefield illumination |
| imaged parameter |
fluorescence emission |
| imaging method |
darkfield microscopy |
| imaging method |
evanescent wave scattering |
| imaging method |
evanescent wave microscopy |
| imaging method |
FLIM |
| imaging method |
FRET |
| imaging method |
FRAP |
| resolution-enhancing method |
PAINT |
| resolution-enhancing method |
PALM |
| resolution-enhancing method |
STORM |
| sample preparation method |
dispersed cells in vitro |
Workshop n°41: Adaptive Optics for Microscopy – from Technology to Applications
No link to public dataset
abstract
Shaping the wavefront of light in microscopes with adaptive optics modules has attracted increasing attention. Adaptive optics modules are used to correct optical aberrations and compensate optical scattering, in order to improve microscope image quality. Several adaptive optics technologies are available, such as special light modulators, deformable mirrors, deformable phase plates etc. In this workshop, we want to explain the principles on how different adaptive optics modules shape the wavefront. We will build a small light path including a deformable mirror, present how optical aberrations are caused by different optical elements such as objectives, lens, filters and medium with inhomogeneous refractive index, and how deformable mirrors can shape and correct the distorted wavefront. Finally, we will discuss different possibilities to correct a distorted wavefront with adaptive optics and give examples where adaptive optics has been successfully applied in light microscopy.
Figure 1: Deformable mirror consist of actuators which can be controlled individually to shape the incident beam wavefront on different Zernike modes. A beam profiler shows the change of the beam shape after being reflected from the deformable mirror when different Zernike modes are applied.
| root term |
term label |
| detection method |
wide-field detection |
| illumination method |
coherent illumination |
| imaging method |
recorded image |
| sample preparation method |
cryostat-sectioned tissue |
| visualization method |
fluorescent label |
Workshop n°44: FLUCS (Focused Light-Induced Cytoplasmatic Streaming) – a new method for non-invasive induction of thermoviscous flows for the characterisation of complex viscoelastic materials
https://omero-tim.gerbi-gmb.de/webclient/?show=project-2994
abstract
In this workshop, we will introduce the concept of thermoviscous flows and demonstrate how FLUCS (Focussed Light Induced Cytoplasmic Streaming) can be utilised to transport microscopic objects through media of different viscosity, from water (order of 1 mPas) to honey (order of 10 Pas), all the way up to coal tar pitch (order of 1 MPas). The unique feature of FLUCS is that it works independently of the absolute viscosity of the medium; all that is required is a fluid exhibiting strong temperature variation of its viscosity.
We will further delve into how FLUCS can transport cargoes of various sizes, ranging from 100 nm to 20 microns in diameter, and made from different materials, such as silica beads, silver/gold nanoparticles, magnetite beads, and polystyrene spheres. The absence of direct laser-particle contact in FLUCS allows for the handling of objects of any material and refractive index properties, an insurmountable task for simple optical tweezers. Moreover, this has significant implications for studies involving laser-light absorbing materials, heat-sensitive materials, or even biological entities, such as cells or cellular components.
Lastly, we will discuss force measurements and microrheology using FLUCS, which rely on fine synchronisation between the laser driver and the camera image acquisition. With this part of the workshop, we will touch on recent developments of FLUCS, where the technique can be employed not only to induce highly controlled unidirectional laminar flows with varied intensity and locality, but also to create opposing flows or dynamic flows that periodically change direction. These added capabilities enable further applications in sensing forces and material response, deemed essential for probing the mechanics of both non-living and living matter. Our FLUCS setup consists of three major components:
1.) An inverted high-resolution microscope for fluorescence measurements.
2.) An infrared laser scanning unit with user-friendly software control.
3.) A temperature-controlled stage enabling implementation in heat-sensitive samples.
Stoev et al. (2021) Highly sensitive force measurements in an optically generated, harmonic hydrodynamic trap. eLight, 1, 1–9, https://doi.org/10.1186/s43593-021-00007-7
Minopoli et al. (2023) ISO-FLUCS: symmetrization of optofluidic manipulations in quasi-isothermal micro-environments. eLight, 3, 1-16, https://doi.org/10.1186/s43593-023-00049-z
| root term |
term label |
| contrast-enhancing method |
optical method |
| illumination method |
multiple point scanning |
| illumination method |
trapping illumination |
| imaged parameter |
absorption of illumination |
| imaging method |
microscopy |
Workshop n°46: Plants on stage: tips and tricks for sample preparation and imaging of plant tissues
https://omero-tim.gerbi-gmb.de/webclient/?show=project-2996
abstract
Plant tissue is a challenging target for microscopic imaging. Autofluorescence of chlorophyll and lignin limit the use of fluorescent labels for multiplexing, the scattering of cell walls and cuticles prevents efficient imaging of deeper cell layers. In addition, some parts of the plant are not easily accessible as they are very sensitive to e.g. mechanical stress during preparation.
We will present several ways to overcome typical problems in imaging of a wide range of plant tissues including e.g. leaf and stem. Furthermore we will show ways to gently and reproducibly prepare sensitive tissues (e.g. ovules) for imaging without preparation artifacts and even for further cultivation.
The focus will be on imaging living or freshly fixed material, mainly from Arabidopsis thaliana. Participants will have the opportunity to do preparation steps on their own and are invited to discuss own techniques.
The workshop is targeted at people without any prior knowledge to get started with plant imaging as well as experienced scientists who wish to discuss techniques or "see how others are doing it".
| root term |
term label |
| detection method |
complementary metal oxide semiconductor (CMOS) |
| illumination method |
slit-scanning illumination |
| imaged parameter |
fluorescence emission |
| imaging method |
slit-scan confocal microscopy |
| whole mounted tissue |
whole mounted tissue |
| sample preparation method |
chemically fixed tissue |
| sample preparation method |
detergent permeabilized |
| source of contrast |
compartmentalization of stain or label |
| visualization method |
EGFP |
| visualization method |
SR2200 (undefined "probe for lipid"?) |
| visualization method |
chlorophyll autofluorescence (undefined) |
Workshop n°47: Optical sectioning - comparing different approaches to removing out-of-focus signal
https://omero-tim.gerbi-gmb.de/webclient/?show=dataset-25779
abstract
Since the introduction of the first commercial laser scanning confocal microscopes in the mid-1980ies, optical sectioning has become a workhorse technique for generating fluorescence images free of out-of-focus fluorescence and with inherent 3D resolution. Long a method mostly limited to investment-intensive fully integrated microscope systems, the past decade has seen a range of novel, more budget-friendly technologies become readily available. These different technologies all have their own strengths and weaknesses. While some of these can be quite obvious, others are more subtle. As a result finding the right technology for a given experiment or sample is not always easy.
In this workshop, we will provide a brief introduction into general optical sectioning, followed by an interactive discussion of several currently available techniques (e.g. spinning disk confocal, point laser scanning, slit scanning, and spinning disc confocals, as well as structured illumination and HiLo), including strengths and weaknesses. Following this, we will provide a practical hands-on session in which three modules implementing three different technologies (Slit-scanning, resonance point laser scanning, and HiLo) will each be used with several different samples. This will not just allow participants to gain first hand experience with these techniques, but also provide a more intuitive feel for which strengths and weaknesses are relevant for different sample types.
Maico
| root term |
term label |
| detection method |
photomultiplier tube (PMT) |
| illumination method |
coherent illumination |
| imaged parameter |
fluorescence emission |
| imaging method |
single spot confocal microscopy |
| sample preparation |
embedded tissue |
| visualization method |
fluorescent label |
NL5+
| root term |
term label |
| detection method |
complementary metal oxide semiconductor (CMOS) |
| illumination method |
coherent illumination |
| imaged parameter |
fluorescence emission |
| imaging method |
slit-scan confocal microscopy |
| sample preparation |
embedded tissue |
| visualization method |
fluorescent label |
SPARQ
| root term |
term label |
| contrast-enhancing method |
computational method |
| detection method |
complementary metal oxide semiconductor (CMOS) |
| illumination method |
coherent illumination |
| illumination method |
widefield illumination |
| imaged parameter |
fluorescence emission |
| imaging method |
fluorescence microscopy |
| sample preparation |
embedded tissue |
| visualization method |
fluorescent label |
Workshop n°50: Automated cell tracking and fluorescence analysis
No link to public dataset
abstract
Cell tracking and fluorescence profile analysis are crucial for studying cellular behaviors and responses. These techniques allow to monitor cell dynamics such as migration, proliferation, and differentiation, providing key insights into cell fate decisions. By analyzing fluorescence profiles, it is possible to classify cell fates based on specific markers or investigate intracellular processes like calcium signaling, vital for many cellular functions.
However, segmentation and tracking are impeded for example by cells migrating in and out of focus, debris, sub-optimal staining or dying cells showing altered fluorescence profiles. Fluorescence analysis is often made difficult by uneven illumination, image noise or segmentation issues. Moreover, integrating fluorescence profiles requires some practical (and numerical) considerations. Overcoming these issues requires robust algorithms and advanced image analysis to obtain biologically meaningful and reliable results.
In this workshop, we will discuss general principles and methods for robust segmentation and tracking, as well as techniques for extracting, analyzing, and classifying fluorescence profiles. In the first part of the workshop, we will cover general solutions and considerations about the individual analysis steps.
In the second part, we will provide a hands-on demonstration of how to track and analyze cells using ImageJ. Therefore, we will use existing tools and plugins to perform segmentation, tracking, and to extract fluorescence data.
In the third part, we will introduce MSparkles, our custom-built analysis application, highlighting its advanced capabilities for cell tracking and fluorescence analysis. MSparkles offers enhanced features and user-friendly interfaces that streamline the analysis process, providing more accurate and efficient results. Attendees will gain practical experience using MSparkles, learning how to leverage its unique tools to tackle complex analysis challenges in their own research. Through these sessions, participants will be equipped with comprehensive knowledge and hands-on skills in both ImageJ and MSparkles, empowering them to effectively analyze cell behavior and fluorescence profiles in their studies.
| root term |
term label |
| illumination method |
Polarized light illumination |
| illumination method |
UV/visible/IR illumination |
| imaging method |
fluorescence microscopy |
| imaging method |
bright-field microscopy |
| sample preparation method |
dispersed cells in vitro |
For the preparation of the data management concept of the Trends in Microscopy 2025 workshops, workshop providers were asked to report a list of ontology terms relevant to their workshop. When terms could not be found, providers were asked to specify the path where they would have expected to find the term (therefore, entries with a path in the tables are not present in the ontology).
I added links to the public data where available.
Workshop n°2: Exchangeable fluorescent probes for gentle long-time STED imaging
https://omero-tim.gerbi-gmb.de/webclient/?show=project-2952
abstract
Multi-colour imaging of cells and tissue is essential for the understanding of many biomedical research questions. Due to the small size of many subcellular organelles, there structures and dynamic interactions can be best visualized and analyzed using super-resolution light microscopy. Stimulated Emission Depletion (STED) microscopy is such technique which enables multi-colour super-resolution imaging in fixed [1,2] and in living cells [3]. Live-cell STED microscopy set high requirements on the fluorescent probes and requires gentle imaging routines. The fluorescent probes ideally should be bright and photostable to resist the high laser powers. Rhodamine dyes fulfill such properties and have been continuously optimized towards cell-permeability. To install a rhodamine label on any cellular target protein, genetically-encoded self-labeling protein tags like HaloTag, have become a powerful tool for live-cell microscopy. HaloTag undergoes a fast and specific labeling reaction with synthetic probes carrying a reactive ligand to form a covalent bond. This technology has recently been further developed by the development of exchangeable HaloTag Ligands. These probes bind to HaloTag with high affinity but can exchange regularly unlike the covalent labels [4]. In confocal and STED microscopy, this transient binding ensures that proteins can be detected in the living cell over extended periods, as bleaching processes are no longer crucial. Additionally, mutually orthogonal HaloTag7 protein/ligand pairs enabled to perform long-term dual-color imaging experiments.In this workshop, participants will learn how sensitive samples like living cells benefit from optimized dye and imaging techniques. Participants will be introduced into novel exchangeable fluorescent probes like the abberior HaloX probes (Fig. 1). In addition, participants will get hands-on experience of how to use them for sample preparation and gentle long-time life cell imaging with reduced applied light dose.
References
[1] Hell, S.W., Wichmann, J., 1994. Opt. Lett. 19, 780
[2] Saal, S.J., Hell, S.W., Jakobs, S., 2017. Nat Rev. 18.
[3] Stockhammer, A., Bottanelli, F., 2020. J. Phys D: Appl. Phys. 54.
[4] Kompa, J., Bruins, J., Glogger M., Wilhelm, J., Frei, M.S., Tarnawski, M., D’Este, E., Heilemann, M., Johnsson, K., 2023. J. Am. Chem. Soc 145, 5.
Workshop n°4: Multimodal correlation spectroscopy of biomolecules in living cells
https://omero-tim.gerbi-gmb.de/webclient/?show=project-2955
abstract
Fluorescence and raster image correlation spectroscopy (FCS, RICS) are powerful tools to analyze biophysical properties of molecules in vitro as well as in living cells. The implementation of "array detector" FCS (1) and "arbitrary-region" RICS (2) on a commercial microscope (3, 4) have made the application of these techniques much more user-friendly. In or workshop we want to bring these seemingly difficult technologies closer to potential users. After theoretical introductions into the methodologies, we will assess in hands-on sessions the biophysical behavior (molecule concentration and diffuion rates) of EGFP multimeres at different locations within living cells. Benefits and potential pitfalls will be outlined in detail. After the session, users should be equipped with sufficient knowledge and starting expertise to establish the new concepts in their own labs.(1) Scipioni et al., 2018, Nat. Comm. 9:5120, doi: 10.1038/s41467-018-07513-2
(2) Hendrix et al., 2016, Biophys. J. 111:1785, doi: 10.1016/j.bpj.2016.09.012
(3) https://www.zeiss.com/microscopy/en/products/software/zeiss-zen/dynamics-profiler.html
(4) https://www.zeiss.com/microscopy/en/products/software/zeiss-zen/spectral-rics.html
Workshop n°8: FLIM-FRET: A powerful tool to visualize multimerization of plasticity-related gene 5 in living cells
No link to public dataset
abstract
Plasticity-related gene 5 (PRG5) is a membrane protein predominantly found in neurons and is involved in cellular processes such as growth cone guidance, migration and spine formation. Overexpression of PRG5 induces filopodia in non-neuronal cell lines, contributes to the induction of spines in immature neurons, and regulates spine density and morphology in mature neurons. Understanding the formation of spines is pivotal, as spine disruptions are associated with numerous neurological disorders. Although the importance of PRG5 in neuronal function is inevitable, the precise mechanisms how it exactly induces membrane protrusions and orchestrates cellular responses remain unresolved. We hypothesise that multimerization of PRG5 is required for its functionality. FLIM-FRET, or Fluorescence Lifetime Imaging Microscopy-Förster Resonance Energy Transfer, is an excellent approach to monitor dynamic protein-protein interaction and visualise their localisation in living cells with high tempo-spatial resolution. FRET is a physical, non-radiative process that occurs between an excited donor fluorophore and an acceptor fluorophore, while FLIM provides a sensitive approach to measure and quantify FRET. Once the acceptor is in close proximity (<10 nm) to the fluorescent donor, FRET energy transfer occurs. Due to this distance dependency, the two fluorophore labelled proteins undergoing FRET must physically interact. Here, we will use FLIM-FRET to detect PRG5 multimerization at the plasma membrane, particularly at the tips of filopodial protrusions in HEK cells. For the workshop, we will cooperate with the company PicoQuant, which will provide us with their new single photon counting confocal microscope, Luminosa, as well as with a Nikon based confocal laser scanning microscope including an upgrade kit.Workshop n°11: Monitoring activation of the GPCR NTSR1 in living cells by FLIM-heteroFRET and homoFRET
https://omero-tim.gerbi-gmb.de/webclient/?show=project-2961
abstract
The neurotensin receptor 1 (NTSR1) belongs to the class A of G protein-coupled receptors (GPCRs). Understanding the molecular mechanisms underlying these receptors is crucial for the development of therapeutic drugs. A central focus in the study of GPCR signaling pathways is the comparison between monomeric and oligomeric GPCRs as active signal transmitters. Our neurotensin receptor 1 (NTSR1) is fused with the bright fluorescent protein. NTSR1-mNeonGreen is expressed in living human HEK293T cells. During the workshop, various lifetime-resolved hetero- and homoFRET imaging experiments to investigate changes in the oligomerization of NTSR1 following the addition of its ligand will be demonstrated. The technique employed for this purpose is FLIM-FRET, a powerful method used to explore molecular interactions within biological samples both in vitro and in vivo. Monitoring oligomerization of NTSR1-mNeonGreen by homoFRET in living HEK293T cells: Upon addition of the ligand neurotensin the oligomerization of the receptor within in plasma membrane is induced and will be monitored. This process involves the formation of receptor dimers. As dimers form, the distance between neighboring receptors decreases, leading to increased proximity between the fluorescent protein (mNeonGreen) molecules. This change can be measured using fluorescence anisotropy indicating homoFRET. Specifically, the NTSR1-mNeonGreen will be excited with a linear polarized pulsed 2-photon laser in a confocal microscope. The fluorescence is separated by polarization and simultaneously recorded by two ultra-sensitive detectors to achieve high-resolution time-correlated single photon counting and to analyze fluorescent lifetimes and time-resolved anisotropies.Monitoring neurotensin binding demonstrated by heteroFRET:
To investigate neurotensin binding to NTSR1, a pair of donor- and acceptor-fluorophores is utilized. The donor is mNeonGreen fused to NTSR1. The acceptor-fluorophore, in this case, is a cyanine dye specifically bound to the ligand neurotensin. Changes in FRET efficiency will serve as indication for neurotensin binding to NTSR1. Additionally, it will be assessed whether the receptor-ligand complex remains intact during the internalization of receptor aggregates.
Our comprehensive approaches combine advanced microscopic imaging techniques, including FLIM-FRET, to unravel the intricate dynamics of NTSR1 oligomerization and ligand interactions.
Workshop n°26: The opportunity of a lifetime: Fluorescence Lifetime Imaging Microscopy in research, a guide from F to M
https://omero-tim.gerbi-gmb.de/webclient/?show=project-2976
abstract
Fluorescence Lifetime Imaging Microscopy (FLIM) is a well-established technique for analyzing dynamics and interactions of molecules and proteins in a wide range of samples and environments. Even if the technique is in principle available in laboratories already for quite some time, it was complicated to apply and often required the assistance of an expert to operate the device, measure the samples, and analyze the data.With the recent releases from manufacturers of faster and easy to use devices, FLIM becomes more accessible to a larger audience. With acquiring FLIM images now being comparatively easy, the main challenge is to design and evaluate experiments in a meaningful way.
This workshop will use a Leica FALCON system to train and inform on what are the advantages and limitations of FLIM, and how to prepare experiments to obtain robust results. It will be discussed how FLIM can be a very advantageous technique over others in selected case. Several samples will be recorded and analyzed making use of the integrated software routines including phasor analysis. Interpretation of the data will be discussed.
Background:
With the development of sensitive and highly resolving instruments, fluorescence microscopy has become an essential tool of biological imaging. Its best-known feature is the specificity of its emission signal (fluorescence) that allows structures of interest to stand out against an otherwise dark background. In addition to its spectral properties for excitation and emission, a fluorophore is defined by a third property: the lifetime of its excited state before the return to the ground state by fluorescence emission. Since the emission of a fluorescence photon normally happens in the range of nanoseconds after excitation, dedicated instrumentation is needed for lifetime measurements, but fluorescence lifetime can add significant information that helps to understand the sample better. It is affected by the environmental conditions in the sample. Spectrally overlapping signals can be separated by their lifetime, allowing a better understanding of complex samples and the removal of background autofluorescence. Changes in de-excitation pathways (e.g. by FRET or STED) change the lifetime and can be used for distance measuremens at the molecular level and for improved resolution.
Workshop n°28: Hands-on light sheet microscopy with Flamingo, the shareable custom research microscope platform
https://omero-tim.gerbi-gmb.de/webclient/?show=project-2978
abstract
In the early 21st century, light sheet microscopy was introduced to the biological sciences and is now accepted as the new standard for fast and gentle 3D fluorescence imaging in living multicellular specimens. The core of our lab evolves around custom-built light sheet microscopy technology for challenging biological multiscale imaging projects. To streamline custom microscope development, we developed Flamingo, a powerful and modular microscope framework that is also compact and can be quickly deployed in collaborators’ labs. Over the last few years, the Flamingo has enabled novel biological experiments inside and outside our optics lab. As a significant part of the scientific community struggles to access a light sheet microscope tailored to their individual needs, the Flamingo gives more researchers the chance to utilize this powerful technology for their research.In this workshop, we will introduce the concept of shared custom microscopy technology and demonstrate Flamingo's modularity. We will provide insights into light sheet microscopy and give you hands-on time with the microscope to better understand both the optics and the modes of operation. We will explain the relevant optics and discuss opportunities and challenges in light sheet microscopy, illustrated with image data recorded on the Flamingo. Live zebrafish embryos and larvae will be available to demonstrate and test Flamingo’s capabilities in living specimens.
Workshop n°39: Simplifying reproducibility in high sensitivity TIRF microscopy to capture dynamics of single molecules
No link to public dataset
abstract
Well-defined and controlled imaging conditions are crucial for reproducible experiments with complex biological samples, such as live cell cultures and single molecule assays. Essential parameters include the homogeneity of optical excitation, temperature, gas composition, and relative humidity. Our workshops will demonstrate methods to precisely control these parameters using advanced sensor technologies commonly employed in the semiconductor industry but not yet widely adopted in optical microscopy. Attendees will gain insights into potential pitfalls in sensitive biophysical experiments and learn strategies to avoid them when designing their experiment.Workshop 1: Waveguide-Based TIRF Microscopy for Ultra-Large Field of View Super-Resolution Imaging.
We will introduce the fundamentals of TIRF microscopy and experimentally quantify key physical parameters such as field homogeneity and penetration depth, both of which are crucial for quantitative single-molecule studies. The experiments will be conducted using one of our QUSCITE systems, which enables large field-of-view illumination over a few square millimeters. We will benchmark the system with dynamic DNA origami and stained live cell cultures. Alongside workshop participants, we will analyze our acquired imaging data using open-source software. By extracting sub-diffraction limited features, we will demonstrate the ease of generating reproducible single-molecule super-resolution data on any microscope.
Workshop 2: Precision and Dynamic Environmental Control for Single Particle Studies and Live Cell Imaging.
We will explore temperature and gas-sensitive processes at the single particle level, investigating how slight changes in temperature and gas composition can significantly impact experimental outcomes. Additionally, we will identify and address external factors that can substantially influence the temperature within the sample volume, potentially affecting the results if not properly managed. By integrating the AIROBOX stage-top cooling and heating unit with the VAHEAT micro-heating device and the TIRF module QUSCITE, we will demonstrate precise and rapid adjustments of temperature, humidity, oxygen, and carbon dioxide concentration on-site.
Workshop n°41: Adaptive Optics for Microscopy – from Technology to Applications
No link to public dataset
abstract
Shaping the wavefront of light in microscopes with adaptive optics modules has attracted increasing attention. Adaptive optics modules are used to correct optical aberrations and compensate optical scattering, in order to improve microscope image quality. Several adaptive optics technologies are available, such as special light modulators, deformable mirrors, deformable phase plates etc. In this workshop, we want to explain the principles on how different adaptive optics modules shape the wavefront. We will build a small light path including a deformable mirror, present how optical aberrations are caused by different optical elements such as objectives, lens, filters and medium with inhomogeneous refractive index, and how deformable mirrors can shape and correct the distorted wavefront. Finally, we will discuss different possibilities to correct a distorted wavefront with adaptive optics and give examples where adaptive optics has been successfully applied in light microscopy. Figure 1: Deformable mirror consist of actuators which can be controlled individually to shape the incident beam wavefront on different Zernike modes. A beam profiler shows the change of the beam shape after being reflected from the deformable mirror when different Zernike modes are applied.Workshop n°44: FLUCS (Focused Light-Induced Cytoplasmatic Streaming) – a new method for non-invasive induction of thermoviscous flows for the characterisation of complex viscoelastic materials
https://omero-tim.gerbi-gmb.de/webclient/?show=project-2994
abstract
In this workshop, we will introduce the concept of thermoviscous flows and demonstrate how FLUCS (Focussed Light Induced Cytoplasmic Streaming) can be utilised to transport microscopic objects through media of different viscosity, from water (order of 1 mPas) to honey (order of 10 Pas), all the way up to coal tar pitch (order of 1 MPas). The unique feature of FLUCS is that it works independently of the absolute viscosity of the medium; all that is required is a fluid exhibiting strong temperature variation of its viscosity. We will further delve into how FLUCS can transport cargoes of various sizes, ranging from 100 nm to 20 microns in diameter, and made from different materials, such as silica beads, silver/gold nanoparticles, magnetite beads, and polystyrene spheres. The absence of direct laser-particle contact in FLUCS allows for the handling of objects of any material and refractive index properties, an insurmountable task for simple optical tweezers. Moreover, this has significant implications for studies involving laser-light absorbing materials, heat-sensitive materials, or even biological entities, such as cells or cellular components. Lastly, we will discuss force measurements and microrheology using FLUCS, which rely on fine synchronisation between the laser driver and the camera image acquisition. With this part of the workshop, we will touch on recent developments of FLUCS, where the technique can be employed not only to induce highly controlled unidirectional laminar flows with varied intensity and locality, but also to create opposing flows or dynamic flows that periodically change direction. These added capabilities enable further applications in sensing forces and material response, deemed essential for probing the mechanics of both non-living and living matter. Our FLUCS setup consists of three major components: 1.) An inverted high-resolution microscope for fluorescence measurements. 2.) An infrared laser scanning unit with user-friendly software control. 3.) A temperature-controlled stage enabling implementation in heat-sensitive samples.Stoev et al. (2021) Highly sensitive force measurements in an optically generated, harmonic hydrodynamic trap. eLight, 1, 1–9, https://doi.org/10.1186/s43593-021-00007-7
Minopoli et al. (2023) ISO-FLUCS: symmetrization of optofluidic manipulations in quasi-isothermal micro-environments. eLight, 3, 1-16, https://doi.org/10.1186/s43593-023-00049-z
Workshop n°46: Plants on stage: tips and tricks for sample preparation and imaging of plant tissues
https://omero-tim.gerbi-gmb.de/webclient/?show=project-2996
abstract
Plant tissue is a challenging target for microscopic imaging. Autofluorescence of chlorophyll and lignin limit the use of fluorescent labels for multiplexing, the scattering of cell walls and cuticles prevents efficient imaging of deeper cell layers. In addition, some parts of the plant are not easily accessible as they are very sensitive to e.g. mechanical stress during preparation.We will present several ways to overcome typical problems in imaging of a wide range of plant tissues including e.g. leaf and stem. Furthermore we will show ways to gently and reproducibly prepare sensitive tissues (e.g. ovules) for imaging without preparation artifacts and even for further cultivation.
The focus will be on imaging living or freshly fixed material, mainly from Arabidopsis thaliana. Participants will have the opportunity to do preparation steps on their own and are invited to discuss own techniques.
The workshop is targeted at people without any prior knowledge to get started with plant imaging as well as experienced scientists who wish to discuss techniques or "see how others are doing it".
Workshop n°47: Optical sectioning - comparing different approaches to removing out-of-focus signal
https://omero-tim.gerbi-gmb.de/webclient/?show=dataset-25779
abstract
Since the introduction of the first commercial laser scanning confocal microscopes in the mid-1980ies, optical sectioning has become a workhorse technique for generating fluorescence images free of out-of-focus fluorescence and with inherent 3D resolution. Long a method mostly limited to investment-intensive fully integrated microscope systems, the past decade has seen a range of novel, more budget-friendly technologies become readily available. These different technologies all have their own strengths and weaknesses. While some of these can be quite obvious, others are more subtle. As a result finding the right technology for a given experiment or sample is not always easy.In this workshop, we will provide a brief introduction into general optical sectioning, followed by an interactive discussion of several currently available techniques (e.g. spinning disk confocal, point laser scanning, slit scanning, and spinning disc confocals, as well as structured illumination and HiLo), including strengths and weaknesses. Following this, we will provide a practical hands-on session in which three modules implementing three different technologies (Slit-scanning, resonance point laser scanning, and HiLo) will each be used with several different samples. This will not just allow participants to gain first hand experience with these techniques, but also provide a more intuitive feel for which strengths and weaknesses are relevant for different sample types.
Maico
NL5+
SPARQ
Workshop n°50: Automated cell tracking and fluorescence analysis
No link to public dataset
abstract
Cell tracking and fluorescence profile analysis are crucial for studying cellular behaviors and responses. These techniques allow to monitor cell dynamics such as migration, proliferation, and differentiation, providing key insights into cell fate decisions. By analyzing fluorescence profiles, it is possible to classify cell fates based on specific markers or investigate intracellular processes like calcium signaling, vital for many cellular functions. However, segmentation and tracking are impeded for example by cells migrating in and out of focus, debris, sub-optimal staining or dying cells showing altered fluorescence profiles. Fluorescence analysis is often made difficult by uneven illumination, image noise or segmentation issues. Moreover, integrating fluorescence profiles requires some practical (and numerical) considerations. Overcoming these issues requires robust algorithms and advanced image analysis to obtain biologically meaningful and reliable results.In this workshop, we will discuss general principles and methods for robust segmentation and tracking, as well as techniques for extracting, analyzing, and classifying fluorescence profiles. In the first part of the workshop, we will cover general solutions and considerations about the individual analysis steps.
In the second part, we will provide a hands-on demonstration of how to track and analyze cells using ImageJ. Therefore, we will use existing tools and plugins to perform segmentation, tracking, and to extract fluorescence data.
In the third part, we will introduce MSparkles, our custom-built analysis application, highlighting its advanced capabilities for cell tracking and fluorescence analysis. MSparkles offers enhanced features and user-friendly interfaces that streamline the analysis process, providing more accurate and efficient results. Attendees will gain practical experience using MSparkles, learning how to leverage its unique tools to tackle complex analysis challenges in their own research. Through these sessions, participants will be equipped with comprehensive knowledge and hands-on skills in both ImageJ and MSparkles, empowering them to effectively analyze cell behavior and fluorescence profiles in their studies.