Imaging Center

Imaging Center

Head of Imaging Center:
Naomi Kamasawa

Overview

The Max Planck Florida Institute for Neuroscience (MPFI) Imaging Center provides investigators with access to cutting-edge imaging technologies and expertise in the fields of electron microscopy, confocal, two-photon and super-resolution microscopy, to achieve tasks from ultrastructural visualization to subcellular imaging of live tissue. The Imaging Center offers training, support, and advice concerning image acquisition, experimental design, and data analysis, and collaborates with investigators to develop novel imaging approaches and techniques.

High-resolution imaging is a critical tool for neuroscience and cell biology alike, with new technologies and methods constantly pushing the boundaries of what is possible. Yet for individual laboratories, keeping pace with all the novel developments and breakthroughs, acquiring and maintaining new equipment, and understanding the intricacies of each imaging method is hard to do. Instead, the Imaging Center at the Max Planck Institute for Neuroscience (MPFI) provides access to state-of-the art-imaging capabilities in conjunction with the necessary scientific expertise and experience in the fields of electron microscopy, confocal- and super-resolution microscopy. The Imaging Center offers training in use of the microscopes, provides technical assistance for imaging, sample preparation, labeling and other techniques, and assists with data and image analysis, experimental design and the interpretation of experimental results. In collaboration with researchers, the Imaging Center develops novel techniques to overcome existing experimental limitations and to advance the field. Furthermore, the Imaging Center is actively engaged in the various education outreach initiatives at the MPFI, from training high school and undergraduate interns to teaching in advanced imaging courses and workshops. Finding the right tool for the job is difficult – we are here to help.

Services

Electron Microscopy

  • Conventional ultrathin-section TEM (transmission electron microscopy) to visualize pure morphology with high-resolution
  • Serial Block Face SEM for precise, high-throughput three-dimensional reconstruction of tissue specimens
  • Thin-section based pre- and post-embedding immuno-EM for localization of target molecules
  • Freeze-fracture replica immunogold labeling to localize target membrane proteins, including all kinds of receptors and channels in two-dimensionally visualized membrane structures
  • Negative staining to visualize small molecules and others
  • Thin-section based Array tomography with serial-SEM imaging.
  • Conventional SEM (scanning electron microscopy)
  • Correlative light and electron microscopy

Confocal Light Microscopy

  • High sensitivity point scanning confocal imaging
  • Parallel multi-color imaging
  • Multi-point time-lapse imaging
  • Ultra-fast resonant scanning for reduced bleaching and photo-damage (particularly for live-cell imaging)
  • Full microscope enclosure for live-cell imaging High-throughput generation of large 3D data stacks for use in 3D reconstructions
  • Improved signal-to-noise (SNR) and spatial resolution based on image reconstruction using the Airyscan detection system
  • Various techniques for studying molecular dynamics and interactions, such as FRET, Anisotropy, FRAP, FLIP and FCS

Super-resolution Light Microscopy

  • Fixated or live sample imaging (at room temperature or heated using a dedicated sample chamber and objective heater)
  • Labeling with fluorescent proteins, organic dyes and other fluorescent markers possible
  • Deep imaging inside thick tissue samples (<100µm works best)
  • Subdiffraction spatial resolution between 30nm–50nm (2D) and 90nm–110nm (3D)
  • Two STED lasers for imaging in the blue/green (595nm STED) and the orange/far-red spectral range (775nm STED)
  • Simultaneous, multi-channel recording of up to 4 channels by pulse-, pixel-, line-, or frame-interleaved acquisition
  • Multi-color imaging of up to 4 colors with STED resolution (or confocal, or a mixture of both)
  • Image-sectioning (for 3D-stacks) and time-lapse imaging
  • Photon-counting detection modes (FLIM, etc.)
  • RESCue mode imaging for reduced photobleaching and -toxicity

Data Analysis and Image Processing

Training and Consultation

  • Basic and advanced light microscopy training and supervision
  • Optimizing imaging parameters for confocal and super-resolution microscopy
  • Selection of fluorescent labels and sample fixation procedures
  • Electron microscopy training and supervision
  • Sample preparation for EM applications
  • General technical support and advice
  • Development of novel imaging approaches
  • Proper image and data analysis methodology, tools and software

Equipment

Electron Microscopy

  • Zeiss Merlin VP Compact (FE-SEM)
  • Zeiss GeminiSEM 300 (FE-SEM)
  • Tecnai 12 G2 Spirit BioTwin (TEM)
  • 3View serial block face imaging system with OnPoint detector

Electron Microscopy Sample Preparation

  • High-pressure freezer (HPM100)
  • Automatic freeze-substitution system (AFS2)
  • Cryo-fixation and –preparation system (CPC)
  • Ultramicrotome (UC7)
  • Automated tape collecting ultramicrotome (ATUMtome)
  • Glass knife maker (KMR3)
  • Freeze-fracture and etching system (JFDII)
  • Sputtering and e-beam coater (ACE600)

Confocal Microscopes

  • Zeiss LSM 880 with Airyscan
    (inverted stage; 7 excitation lasers @405nm, 458nm, 488nm, 514nm, 561nm, 594nm & 633nm; confocal detection with high-sensitivity GaAsP detector & 2 PMTs, spectrally configurable between 390nm – 750nm; Airyscan module; motorized stage; automatic focus-correction; objectives: 10x/0.45 air, 20x/0.80 air, 40x/1.2 water, 63x/1.4 oil)
  • Zeiss LSM 780
    (upright stage; 7 excitation lasers @405nm, 458nm, 488nm, 514nm, 561nm, 594nm & 633nm; confocal detection with high-sensitivity GaAsP detector & 2 PMTs, spectrally configurable between 390nm – 750nm; photon-counting; objectives: 10x/0.40 air, 20x/0.80 air, 25x/0.8 multi-corr, 40x/1.3 Oil and 63x/1.3 multi-corr)
  • Leica TCS SP5 II with resonant scanner
    (upright stage; 6 excitation lasers @405nm, 458nm, 488nm, 514nm, 561nm & 633nm; 8kHz resonant-scanner; confocal detection: 3 hybrid- & two regular PMTs, spectrally configurable; motorized scanning stage; objectives: 10x/0.40 air, 20x/0.70 air, 40x/1.25 oil, 63x/1.40 oil and 63x /0.9 water)

Super-resolution Microscopes

  • Abberior Instruments Expert-Line 3D-STED
    (inverted stage; 4 excitation lasers @405nm, 485nm, 561nm & 640nm; 2 pulsed STED lasers @595nm & 775nm; easy3D STED depletion module; beam-scanner; motorized pinhole; four detection channels (APDs) between 425nm – 725nm; time-gated detection; single-photon counting; RESCue; objectives: 10x/0.25 air, 40x/1.3 oil, 63x/1.2 water, 100x/1.4 oil)

 

Contact

For questions about:
» Electron Microscopy services, contact Dr. Naomi Kamasawa
» Confocal Microscopy services, contact Dr. Long Yan
» Super-resolution Microscopy services, contact Dr. Nicolai Urban

 

Recent Publications from the MPFI Imaging Center

  1. Dong, W., Radulovic, T., Goral, R.O., Thomas, C., Suarez Montesinos, M., Guerrero-Given, D., Hagiwara, A., Putzke, T., Hida, Y., Abe, M., Sakimura K., Kamasawa N., Ohtsuka T., Young S.M. Jr. (2018). CAST/ELKS Proteins Control Voltage-Gated Ca2+ Channel Density and Synaptic Release Probability at a Mammalian Central Synapse. Cell Reports 24, 284-293.e6.
  2. Sarria, I., Cao, Y., Wang, Y., Ingram, N.T., Orlandi, C., Kamasawa, N., Kolesnikov, A.V., Pahlberg, J., Kefalov, V.J., Sampath, A.P., Martemyanov, K.A. (2018). LRIT1 Modulates Adaptive Changes in Synaptic Communication of Cone Photoreceptors. Cell Rep 22, 3562–3573.
  3. Grassi, D., Howard, S., Zhou, M., Diaz-Perez, N., Urban, N.T., Guerrero-Given, D., Kamasawa, N., Volpicelli-Daley, L.A., LoGrasso, P., and Lasmezas, C.I. (2018). Identification of a highly neurotoxic α-synuclein species inducing mitochondrial damage and mitophagy in Parkinson’s disease. PNAS, Published ahead of Print Feb 27, 2018.
  4. Lubbert, M., Goral, R.O., Satterfield, R., Putzke, T., Maagdenberg, A.M. van den, Kamasawa, N., and Young S.M. Jr. (2017). AA novel region in the CaV2.1 α1 subunit C-terminus regulates fast synaptic vesicle fusion and vesicle docking at the mammalian presynaptic active zone. eLife Sciences 6, e28412.
  5. Steinecke, A., Hozhabri, E., Tapanes, S., Ishino, Y., Zeng, H., Kamasawa, N., and Taniguchi, H. (2017).  Neocortical Chandelier Cells Developmentally Shape Axonal Arbors through Reorganization but Establish Subcellular Synapse Specificity without Refinement.  eNeuro 4, ENEURO.0057-17.2017.
  6. Neuillé, M., Cao, Y., Caplette, R., Guerrero-Given, D., Thomas, C., Kamasawa, N., Sahel, J.-A., Hamel, C.P., Audo, I., Picaud, S., et al. (2017). LRIT3 Differentially Affects Connectivity and Synaptic Transmission of Cones to ON- and OFF-Bipolar Cells. Investigative Ophthalmology & Visual Science (IOVS) 58, 1768–1778.
  7. Wang, Y., Fehlhaber, K.E., Sarria, I., Cao, Y., Ingram, N.T., Guerrero-Given, D., Throesch, B., Baldwin, K., Kamasawa, N., Ohtsuka, T., et al. (2017). The Auxiliary Calcium Channel Subunit α2δ4 Is Required for Axonal Elaboration, Synaptic Transmission, and Wiring of Rod Photoreceptors.  Neuron 93, 1359–1374.
  8. Peach, K., Koch, M.S., Blackwelder, P.L., Guerrero-Given, D., and Kamasawa, N. (2017). Primary utricle structure of six Halimeda species and potential relevance for ocean acidification tolerance.Botanica Marina – DE GRUYTER.
  9. Owens, A., Lebowitz, J.J., Richardson, B., Jagnarine, D.A., Shetty, M., Rodriquez, M., Alonge, T., Ali, M., Katz, J., Yan, L., Febo, M., Henry, L.K., Bruijnzeel, A.W., Daws, L., Khoshbouei, H. (2017). The sigma-1 receptor modulates methamphetamine dysregulation of dopamine neurotransmission, Nat Commun.;8(1):2228.
  10. Rowan, M.J.M., DelCanto, G., Yu, J.J., Kamasawa, N., and Christie, J.M. (2016). Synapse-Level Determination of Action Potential Duration by K+ Channel Clustering in Axons. Neuron, 91, 370-383.
  11. Mikuni, T., Nishiyama, J., Sun, Y., Kamasawa, N., and Yasuda, R. (2016). High-Throughput, High-Resolution Mapping of Protein Localization in Mammalian Brain by In Vivo Genome Editing. Cell, 165, 1803–1817.
  12. Montesinos, M., Dong, W., Goff, K., Das, B., Guerrero-Given, D., Schmalzigaug, R., Premont, RT., Satterfield, R., Kamasawa, N., Young, SM, Jr. (2015) Presynaptic Deletion of GIT Proteins Results in Increased Synaptic Strength at a Mammalian Central Synapse. Neuron, 88, 918–925.
  13. Cao, Y., Sarria, I., Fehlhaber, K.E., Kamasawa, N., Orlandi, C., James, K.N., Hazen, J.L., Gardner, M.R., Farzan, M., Lee, A., Baker, S., Baldwin, K., Sampath, A.P., Martemyanov, K.A. (2015). Mechanism for Selective Synaptic Wiring of Rod Photoreceptors into the Retinal Circuitry and Its Role in Vision. Neuron 87, 1248-1260.
  14. Kamasawa, N., Sun, Ye., Mikuni, T., Guerrero-Given, D., Yasuda, R. (2015) Correlative Ultrastructural Analysis of Functionally Modulated Synapses Using Automatic Tape-Collecting Ultramicrotome – SEM Array Tomography. Microscopy and Microanalysis 21, Supplement S3, 1271-1272.
  15. Liu, X..A, Kadakkuzha, B., Pascal, B, Steckler, C., Akhmedov, K., Yan, L., Chalmers, M., Puthanveettil, S.V. (2014) New approach to capture and characterize synaptic proteome. Proc Natl Acad Sci USA, 111:16154-9.