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Light microscopy

Microscopes (DWB room 201, 202 and 203)

Current equipment in the light microscope facility includes:

  1. Wide-field fluorescence/brightfield/DIC microscope (Zeiss) – “Edwina”
    • Axioplan 2 imaging upright microscope
    • Wide range of objectives
    • Filter sets for DAPI, CFP, GFP, AF 488, YFP, Texas Red, Cy5
    • Brightfield and Differential Interference Contrast (DIC)
    • Spot Insight QE color digital camera for brightfield (and fluorescence) imaging or Hamamatsu Orca ER B/W digital camera for sensitive fluorescence imaging
    • CRI Nuance camera for spectral unmixing
    • Motorized Z-drive
    • MetaVue acquisition software (Molecular Devices)
  2. Inverted fluorescence microscope for checking fluorescent protein expression levels in tissue culture cells (Nikon)
    • Nikon Eclipse TS100 inverted microscope
    • Filter sets for GFP+DsRed or CFP+YFP
    • Phase contrast
  3. Inverted microscope for long-term time-lapse, fluorescent and phase contrast imaging of cells or embryos (Olympus) – “Wendolene”
    Yeast cells labeled for actin (green), Cln2 (red) and nuclei (blue). Images were captured and deconvolved using the DeltaVision image restoration microscope, allowing fine structures to be revealed within these small cells. Mary Miller and Frederick Cross (Rockefeller University)
    • Olympus IX71 inverted microscope with phase contrast optics
    • 10x, 20x, 40x, 63x and 100x objective lenses, some suitable for use with plastic as well as glass dishes/plates
    • Filter sets for DAPI, CFP, GFP, YFP, RFP and Cy5
    • Olympus DSU white-light spinning disk head for optical sectioning
    • Uniblitz shutters
    • Hamamatsu Orca ER B/W digital camera and QI Click color digital camera
    • Prior XYZ piezo stage (200 microns z-travel) for multiple point visiting during timelapse acquisitions
    • MetaMorph acquisition software (includes MDA)
    • Solent Scientific environmental chamber for temperature regulation
  4. DeltaVision Image Restoration Microscope (Applied Precision) – “Wallace”
    • Inverted Olympus IX-70 microscope
    • 40x, 60x and 100x oil objectives, 60x water objective, 60x Silicone oil objective, 20x dry objective, plus 1.5x optovar
    • Filter sets for DAPI, CFP, GFP/FITC, YFP, Rhodamine, AlexaFluor 594/mCherry and Cy5
    • DIC
    • Separate excitation and emission filter wheels (necessary for FRET)
    • pco.edge sCOS camera
    • Extremely precise stage allowing multiple point visiting and image stitching
    • 410, 488 and 532 laser module (QLM) for FRAP and photoactivation studies
    • Insight SSI 7 color solid state illumination system
    • Live cell imaging
    • Quantitative
    • Koehler and critical illumination
    • Deconvolution using measured PSFs
    • Warner Instruments and Bioptechs heated imaging chambers
  5. 2nd DeltaVision Image Restoration Microscope (Applied Precision) – “Totty”
    • All the features of the above system including a new FRAP/photoactivation module with 405 and 488 lasers
    • “Ultimate Focus” – hardware-based autofocus system for long-term timelapse imaging without focal drift
    • Weatherstation environmental chamber – can be set at temperatures from ambient temperature to 37°C
  6. Spinning disk confocal microscope for real-time live cell imaging (Zeiss/Perkin-Elmer) – “Bunty”
    Cell outlines in the Drosophila embryo, as labeled by adherens junction markers.
    Cell outlines in the Drosophila embryo, as labeled by adherens junction markers. Images were collected using a Zeiss LSM 510 confocal microscope.
    Benjamin Boettner and Ulrike Gaul (Rockefeller University)
    • Inverted Zeiss Axiovert 200 microscope
    • Wide range of oil and water immersion objectives
    • Sutter DG4 ultra-rapid wavelength switcher and Sutter emission filter wheel for fast multicolour imaging in wide-field mode (using camera on base port)
    • Perkin-Elmer UltraView spinning disk confocal head on side-port for real-time confocal imaging
    • Solid-state 443, 491, 523, 561 and 644 lasers for excitation (Spectral Applied)
    • Photonics Instruments Digital Mosaic system with 405 nm laser for performing ROI FRAP and photoactivation;
    • Choice of Hamamatsu Flash 4.0 sCMOS camera or Andor iXon 512×512 EMCCD camera for high resolution or high sensitivity/speed imaging
    • Prior XYZ piezo stage for multiple point visiting during timelapse acquisitions
    • Image acquisition performed with MetaMorph software
    • Eppendorf microinjection system
    • Solent Scientific environmental chamber for control of temperature and CO2
    • Warner Instruments peltier-controlled chamber for piezo insert, permitting rapid temperature shifts
  7. Inverted LSM 880 NLO laser scanning confocal and multiphoton microscope (Zeiss) – “Rocky II”
    • Inverted Zeiss Axio Observer Z1 microscope
    • 405 nm, 458, 488, 514, 561 and 633 laser lines
    • Chameleon Ultra II NIR laser fully tunable between 690 and 1040 nm
    • GaAsP detector for sensitive confocal imaging
    • BiG.2 NDD detector for multiphoton imaging
    • X-Y motorized stage for tiling and multiple point visiting
    • Wide range of oil and water immersion objectives
    • Simultaneous 3-channel fluorescence and DIC imaging
    • FRAP and FRET modules
    • Confocal stereology software (Visiopharm)
    • Warner Instruments heating chamber or OKO lab stage-top incubator for temperature and CO2 controlled environment
  8. Inverted LSM 780 laser scanning confocal microscope (Zeiss) – “Technotrousers
    • Inverted Zeiss Axio Observer Z1 microscope with Definite Focus
    • 405, 440, 488, 514, 561, 594 and 633 laser lines
    • 34 spectral detection channels via GaAsP detector plus two PMTs
    • Fluorescence Correlation Spectroscopy (FCS) software
    • Wide range of oil and water immersion objectives
    • Piezo scanning stage for advanced tiling capabilities
    • FRET and FRAP modules
    • Stage-top incubator for heat and CO2 control
  9. Inverted TCS SP8 laser scanning confocal microscope (Leica) – “Ira
    • Inverted Leica DMI 6000 microscope
    • Fully tunable White Light Laser (470-670 nm) with AOBS, plus 405 nm and 442 nm lasers
    • Three gated HyD detectors plus one PMT detector
    • Conventional scanner and Resonant scanner
    • Super-Z stage for rapid tiling
    • Wide range of oil and water immersion objectives including PlanApo 40x/1.10 NA water objective with motorized correction collar and water immersion micro dispenser
    • Environmental chamber for temperature and CO2 control
  10. FV1000MPE Twin upright multiphoton system (Olympus) – “Philip”
    • Coherent Chameleon Vision II IR laser with integrated dispersion compensation and a wide tuning range of 680 to 1080 nm
    • Coherent Chameleon XR laser with a tuning range of 720-950 nm for MP stimulation
    • Olympus BX61 upright microscope
    • Capability for simultaneous MP imaging and MP/visible stimulation
    • 473 nm visible laser for imaging or stimulation
    • 405 nm laser for visible stimulation
    • Range of objectives including the 25x/1.05 NA Plan objective optimized for infrared wavelength transmission and deep-tissue imaging
    • Wide range of filter sets
    • Prior motorized stage for multiple point visiting
    • Inhalation anesthesia system (VetEquip) for long term live imaging
    • Biostage 600 warming plate (20-20 Technology)
    • Olympus DP-20 camera for accurate repetitions of live sample positioning
  11. STORM/ TIRF system (Nikon) with widefield-FLIM and TIRF-FLIM – “Piella”
    • Nikon TiE inverted microscope with Perfect Focus mechanism
    • 405, 488, 561 and 647 nm laser lines for STORM and TIRF
    • 2D and 3D STORM capabilities
    • Lambert Instruments frequency domain LIFA module for Fluorescence Lifetime Imaging (FLIM), either in widefield (LED) or laser (TIRF) mode (445 and 514 nm lasers);
    • Photonics Instruments Micropoint laser for photoablation/bleaching/activation
    • DG5 for widefield illumination
    • Piezo x,y,z stage
    • Andor DU-897 EMCCD camera
    • Andor Neo sCMOS camera
    • Tokai Hit environmental chamber
    • Wide range of objectives suitable for widefield and TIRF microscopy
    • Controlled by Elements acquisition software (Nikon)
  12. OMX Blaze 3D-SIM super-resolution microscope (Applied Precision) – “Reverend Hedges”
    • 405, 445, 488, 514, 568 lasers for 3D-SIM super-resolution imaging
    • 6-line SSI module for ultra-rapid conventional imaging
    • 100x/1.40 NA UPLSAPO oil objective (Olympus)
    • Three Evolve EMCCD cameras (Photometrics) for simultaneous or sequential acquisition
    • Heating chamber for live cell imaging
  13. LCV110 “VivaView” Incubator Microscope (Olympus) – “Fluffles”
    • 37 degree incubator microscope for multi-day imaging of up to eight 35mm dishes at once
    • 5% CO2 incubator
    • O2 control for hypoxia studies
    • Multi-color, multi-position, multi-z imaging
    • Filter cubes for CFP, GFP, YFP, mCherry, TagRFP657
    • 20x DIC objective with 0.5x and 2x magnification changers
    • Controlled by MetaMorph acquisition software
  14. Laser microdissection system (MMI) – “Sadie”
    • MMI CellCut system, fitted with a 355 nm solid state laser
    • Regular brightfield and fluorescence modules (for DAPI, CFP, GFP, YFP and RFP)
    • Contamination-free dissection from paraffin samples, cryo samples, smears and cell cultures
    • Compatible with microdissection of living cells under sterile conditions
    • 4x, 10x, 20x, 40x, 60x and 100x objectives
  15. Ultramicroscope (LaVision BioTec) – “Polly
    • Light sheet illumination
    • Olympus macro zoom microscope stand with 2x/0.5 N.A. MVPLAPO objective lens
    • 445, 488, 561 640 and 785 nm lasers
    • Andor Neo sCMOS camera
    • For use with fixed and cleared specimens or transparent live specimens
  16. CellVoyager (Yokagawa/Olympus) – “HMSBeagle
    • Spinning disk confocal system
    • Two disks with different pinhole sizes or widefield illumination
    • 445, 488 and 561 nm lasers
    • Hamamatsu 512 x 512 EMCCD camera
    • 10x and 20x dry objectives, 30x, 40x and 60s silicone oil objectives
    • Incubation temperature from 18 degC to 40 degC, CO2 optional

Workstations and software for image processing

Live Xenopus embryo with a transgenic green fluorescent marker for the nervous system and a red fluorescent counterstain marking the blood vessels.
Live Xenopus embryo with a transgenic green fluorescent marker for the nervous system and a red fluorescent counterstain marking the blood vessels. A stack of images was collected using a Zeiss LSM 510 confocal microscope and 3-D reconstruction was performed using the Imaris software from Bitplane.
Ariel Levine, Alison North and Ali Hemmati-Brivanlou (Rockefeller University)

Windows (“QueenVic”, “CharlesDarwin”“Bobo”“Cooker” and “Fetcher”), and Linux (“Gromit”) high-end workstations and servers are available for processing and analyzing acquired images, including 64-bit multi-processor systems with up to 196 GB of memory for processing large multidimensional datasets using 64-bit enabled software packages (“QueenVic” and “CharlesDarwin”). Available software includes:

  • “Huygens” and “Autoquant” deconvolution softwares (modules for deconvolving confocal, spinning disk or multiphoton data);
  • “MetaMorph” (Universal Imaging) (cell counting, tracking, measurements, kymographs);
  • “Imaris” “Surpass” and “FilamentTracer” (Bitplane) (volume rendering in 3D and 4D, surface rendering, measurements, colocalization, clipping planes, 4D tracking, filament tracing);
  • “Arivis” Vision4D for interactive 3D/4D rendering and analysis of large data sets;
  • Arivis” InViewR Virtual Reality station for climbing into your 3D data sets;
  • “FIJI”/”Image J” (freely downloadable software)*;
  • “SoftWoRx” (Applied Precision / DeltaVision) (deconvolution, volume rendering, surface rendering, quantification);
  • “Zen” confocal software (Zeiss) and Leica “LAS-AF” software (projections, movies, ROI measurements, intensity profiles)

The Linux workstation “Gromit” is dedicated to deconvolving and processing images collected on the DeltaVision and OMX microscopes.

All users are urged to save their data directly to their folder on the bio-imaging server, ready for processing on the workstations. This frees up the microscopes for image acquisition by other users. To encourage this, the charge for time on the workstations is considerably lower than the microscope charges. Data on the server can be accessed directly from the user’s laboratory computers, or can be copied to external hard drives (users are responsible for bringing their own drives).

* To install “Image J” for free on your own computer, click on the following two web sites:

Additional lab equipment for live cell imaging

Two cell culture incubators are present in the general laboratory area, one with CO2 and the other without (for cells in CO2-independent media). These can be used for temporary storage of specimens during live cell imaging experiments. A class II biological safety cabinet is also available for use. Please note that the center is not designed to accommodate infectious, radioactive or otherwise hazardous specimens. If you are in any doubt as to the suitability of your specimens for live cell imaging in the center, please contact Alison. Anyone who performs unauthorized experiments that place the other users’ health at risk will be banned from further use of the center.



The Rockefeller University
1230 York Avenue, Box 209
New York, NY 10065

Bio-Imaging Resource Center
Bronk Laboratory
DWB Room 201 (office)
DWB Room 202, 203 (Lab)