The Advanced Imaging Core Facility (AICF) provides access to a wide range of state-of-the-art optical microscopes, enabling users to perform experiments ranging from basic transmitted light observations to advanced in vivo fluorescence imaging techniques. Facility staff offer specialized technical and scientific support at every stage of imaging experiments, from experimental design to image acquisition, as well as data processing and analysis.

Facility Applications and Services

Microscopy techniques available at the Facility:

• Wide-field microscopy in reflected light (epifluorescence).
• Wide-field microscopy in transmitted light, with the following contrast methods: bright field, dark field, phase contrast, and differential interference contrast (DIC).
• Wide-field optical sectioning using structured illumination.
• Laser scanning and spinning disk confocal microscopy.
• Multiphoton microscopy, enabling imaging up to approximately 1 mm deep in 3D samples (tissues, organs, organoids, small animal models).
• FLIM (Fluorescence Lifetime Imaging Microscopy), available in both single-photon laser scanning confocal and multiphoton modes. This technique measures fluorescence decay time of fluorophores and allows, for example, the use of biosensors (to study metabolic activity, signaling mechanisms, pH changes, and microenvironment), separation of overlapping fluorophores, and discrimination of autofluorescence
• Excitation λ-scan / emission λ-scan experiments, essential for characterizing unknown samples and/or non-standard fluorochromes.
• Detection of nonlinear signals such as Second Harmonic Generation (SHG), enabling label-free imaging of structures such as collagen or tissue fibers.
• Time-lapse and in vivo fluorescence and bright-field imaging with incubation systems that allow environmental control over extended periods.
• Large-area fluorescence and bright-field imaging (mosaics/tile-scan) for large samples (e.g., mouse brain sections or histological preparations).
• Super-resolution microscopy:

  • PALM (Photoactivated Localization Microscopy), based on the stochastic activation of photoactivatable fluorophores, and dSTORM (direct Stochastic Optical Reconstruction Microscopy), which uses conventional fluorophores switched between “on” and “off” states using reducing chemical buffers; for studying membrane protein distribution and clustering, single-molecule counting in protein complexes, nanoscale subcellular architecture (nuclear pores, chromatin, cytoskeleton, synapses), and extracellular vesicle imaging.

  • SIM (Structured Illumination Microscopy), based on structured illumination and mathematical image reconstruction; for studying cytoskeleton dynamics, mitosis, vesicular trafficking, and intracellular organelles.
  • TIRF-SIM (Total Internal Reflection Fluorescence SIM), combining near-surface excitation (~100 nm) with structured illumination, improving membrane-plane resolution; used for focal adhesions, exocytosis/endocytosis, receptor dynamics, and synapses.
• FRAP (Fluorescence Recovery After Photobleaching), for studying mobility, lateral diffusion of membrane proteins, protein turnover, and cytoskeleton dynamics.
• FRET (Förster Resonance Energy Transfer) and single-molecule FRET (smFRET), for protein–protein interactions, conformational changes, and intracellular biosensors.
• Single-Particle Tracking (SPT), for membrane receptor diffusion, intracellular transport, and viral dynamics.
• TIRF and HILO (Highly Inclined and Laminated Optical sheet) illumination, for membrane events (endocytosis, exocytosis, focal adhesions) and receptor dynamics.
• Laser microdissection at the single-cell level (on fixed samples and live cultured cells).

Services provided by Facility staff:

• Technical and scientific support in experimental design.
• Technical support for microscope purchasing and configuration.
• Training for independent microscope use.
• Assisted microscopy sessions for non-independent users.
• Image acquisition services.
• Image analysis services, including development of automated analysis pipelines and data interpretation using software such as ImageJ/Fiji, QuPath, SVI Huygens Core and Essential (for deconvolution), Leica LasAF/LasX, Zeiss ZEN, and Nikon NIS-Elements.

Facility Equipment

Wide-field systems

• Fully motorized inverted microscope Nikon Eclipse Ti2 E, equipped with a Lumencor CELESTA multi-line laser source, two monochromatic digital cameras (sCMOS Andor Zyla 4.2 Plus and EMCCD Andor iXon 888), and an Okolab incubation system for live-cell imaging.
• Fully motorized inverted microscope Nikon Eclipse Ti2 E with Nikon D-LEDI multi-line LED source and Hamamatsu ORCA Flash 4.0 sCMOS monochromatic camera.
• Fully motorized inverted microscope Leica DMi8 equipped with Lumencor Sola white LED and Andor Zyla 4 sCMOS monochromatic camera.
• Fully motorized inverted microscope Zeiss Axio Observer Z1 equipped with two light sources (Colibri 2 multicolor LED and HXP 120 metal halide fluorescence lamp), ApoTome 2 structured illumination module, AxioCam 503 mono D monochromatic camera, and Pecon incubator for live-cell imaging.
• Fully motorized upright microscope Zeiss Axio Imager M2 with HXP 120 metal halide fluorescence lamp, Zeiss Axiocam 305 sCMOS monochromatic camera, and Zeiss Axiocam 705 sCMOS color camera.

Confocal and multiphoton systems

• Leica STELLARIS 8 laser scanning confocal system on a Leica DMi8 CS Premium inverted microscope with Okolab incubation system, integrated with DIVE multiphoton unit. Both modes support FALCON and TauSense modules for FLIM acquisition.
  • Confocal configuration: single-photon white laser (pulsed source with emission in the 440–790 nm range), single-photon laser with emission at 405 nm, AOBS beam splitter, galvanometric scanner, PMT detector for transmitted light, and Power HyD S/X detectors for reflected light. Leica LAS X software includes FRET, FRAP, and Lightning deconvolution modules.
  • Multiphoton DIVE configuration: two-photon excitation with tunable IR pulsed laser (680–1080 nm range), 4Tune spectral detection system with non-descanned detectors (Power Hybrid/PMT).
• Leica TCS SP8 laser scanning confocal system on Leica DMi8 microscope with multi-line laser bank, one HyD PMT detector, three conventional PMTs, and a transmitted light detector; prism-based spectral detection. Leica LAS X software includes FRET, FRAP, and Lightning.
• Nikon AX laser scanning confocal system on Nikon Eclipse Ti2 with multi-line laser bank, two ultra-sensitive GaAsP detectors, two PMTs, and a transmitted light detector; emission filter-based channel separation. Nikon AR software includes NIS.ai deconvolution.
• CREST X-light V3 spinning disk system on Nikon Eclipse Ti with CELESTA laser bank, Andor Zyla 4.2 sCMOS and Andor iXon 888 EMCCD monochromatic cameras, and Okolab incubation system. Nikon AR includes NIS.ai deconvolution.

Super-resolution systems

• Nikon N-SIM S system on Nikon Eclipse Ti2 E with multi-line laser bank and Hamamatsu ORCA Flash 4.0 sCMOS monochromatic camera; supports 2D, 3D, and 2D-TIRF SIM reconstruction via Nikon AR software.
• ONI Nanoimager inverted system with multi-line lasers and LED sources, Hamamatsu ORCA Flash 4.0 monochromatic camera. Techniques: wide-field, TIRF, HILO, dSTORM, PALM, SPT, smFRET, DMD-based SIM, and confocal microscopy.

Laser microdissection system

Leica LMD6500 system based on Leica DM6000 B inverted microscope.

Stereomicroscopes

Manual fluorescence stereomicroscopes Leica MZ 10F, MZ 16F, and Leica S9i.

Histological sample preparation instruments

• Leica CM 1850 UV cryostat.
• Thermo Scientific HM525 NX cryostat.
• Leica HistoCore Biocut microtome.
• Leica VT 1200 vibratome.
• Histo-Line HistoPro200 automatic tissue processor.
• Histo-Line TEC 2900 paraffin embedding station.