Confocal and Fluorescence Microscopy

Location: Thomas Building, DE-512
Contact phone: (206) 667-4205
Contact e-mail:

3-D Microscopy (Deconvolution and Confocal)

Fluorescently labeled cells imaged with conventional widefield fluorescence microscopy without (left) and with deconvolution (right), showing the effect of deconvolution in removing out of focus haze and improving resolution and contrast.Deconvolution and confocal microscopy are optical sectioning techniques for capturing high resolution 2-D or 3-D images of biological specimens. Wide field deconvolution uses conventional widefield microscopy and deconvolution software to remove out of focus blur.

Effect of Deconvolution
Deconvolution (right) removes out of focus haze and improves resolution and contrast. This method provides superb results with thin specimens such as bacteria, yeasts, cell monolayers, thin sections, and other samples no more than 10-15 microns thick. Confocal microscopy uses optical techniques to remove out of focus haze and capture high contrast images of in-focus planes. This method is recommended for thicker specimens such as tissue sections, fly embryos, embryonic and adult worms, isolated tissues, cell spheroids, and other specimens. Several confocal technologies are available, including laser scanning confocal microscopy (i.e. conventional confocal), spinning disk, and swept field confocal microscopy.


(for technical details, follow the links to individual microscopes)

  • GE/Applied Precision Deltavison Elite image restoration (deconvolution) microscope. Specially suited for high resolution 3-D imaging of thin samples (bacteria, yeasts, cell monolayers, some thin sections).
  • Zeiss LSM 780 NLO spectral laser scanning confocal and multi photon microscope. Ideal for the imaging of thick specimens (worm embryos, adult worms, fly embryos, tissue sections, biopsies, cell spheroids, and more).
  • Perkin Elmer Ultraview spinning disk confocal system. Specially recommended for time lapse experiments with live cells or small model systems (such as worm and fly embryos).
  • Nikon Ti Live swept field confocal. Primarily used for widefield fluorescence imaging, but can be used in confocal mode. Suitable for imaging CFP/YFP samples.

High Content Analysis and Screening

High content analysis is a technology that combines automated, high throughput microscopy with image cytometry to extract quantitative information for a variety of parameters from a large number of samples. Typically, high content assays are performed with cells grown in monolayers in a micro-plate format and stained with fluorescently labeled antibodies and/or a variety dyes. Cellular Imaging provides support with assay planning and optimization, imaging, and image analysis including the creation of custom analysis protocols.


(For technical details, follow the links below for the instruments)

  • Molecular Devices ImageXpress Micro Confocal. The ImageXpress is a high content analysis system for image-based screening applications in microplate format. The instrument is capable of imaging in fluorescence (Blue, Green, Red and Far Red channels), and transmitted light mode. The system can image in widefield (conventional fluorescence) or confocal mode. This is specially useful for 3D cell cultures and large specimens such as spheroids. The system includes an automated incubator and robotics for automated plate imaging. It is highly recommended to contact the Resource for experiment planning and for a list of compatible plates.
  • Cellomics Arrayscan VTI high content reader. The ArrayScan is specifically designed for high content analysis and for image-based screening applications in multi-well plate format. The instrument is primarily designed for the imaging of fluorescent samples (Blue, Green, Red and Far Red channels), but is also capable of collecting transmitted light images. It is highly recommended to contact the Resource for experiment planning and for a list of compatible plates.
  • Essen Biosciences Incucyte Zoom. The Incucyte is a dedicated, easy to use incubator microscope for kinetic live cell assays. The system is capable of enhanced phase contrast and two color (green, red) fluorescence imaging. The system is ideal for monitoring cell proliferation, invasion, and apoptosis, for monitoring stem cell reprogramming, cytotoxicity assays, reporter gene expression, and more.

Multi-photon Microscopy

Two-photon microscopy uses non-linear fluorescence excitation with pulsed infra-red or near infra-red lasers, combined with sensitive non-descanned detectors to image fluorescence deeper inside biological specimens compared to conventional confocal microscopy.

In conventional fluorescence microscopy, short wavelength visible light is used to excite fluorescence of a fluorophore, which then emits light of a slightly longer wavelength. For example, blue light at 488 nm is used to excite fluorescein or green fluorescent protein, which then emit green light centered around 530 nm. In two photon microscopy, two longer wavelength photons in the near infra-red (NIR) are absorbed quasi-simultaneously, providing the same energy as a single shorter wavelength photon, while the emission wavelength remains the same. To achieve quasi-simultaneous absorption of two photons, extremely high illumination intensities are needed. This is achieved with special ultrafast NIR pulsed lasers. Since the illumination intensity required for the stimulation of fluorescence is reached only at the focal spot, other regions of the sample will not fluoresce, and therefore optical sectioning is achieved without need for a pinhole or deconvolution software. The increased imaging depth of two photon microscopy is due to the lower scatttering of the NIR excitation light, and also to the more efficient detection of scattered emitted light, compared to confocal microscopy. An added benefit of two photon imaging is better viability for certain types of specimens, due to lower toxicity of longer wavelength light.

Multi-photon microscopy uses the same principle, but in this case three (or more) photons are absorbed simultaneously. The imaging resource provides two multi-photon systems: The Zeiss LSM 780 NLO, based on an inverted Zeiss Observer Z1 stand, combines conventional confocal with multi-photon imaging.

Equipment for two-photon and multi-photon microscopy:

(For technical details, follow the links to individual microscopes)

  • Zeiss LSM 780 NLO. Laser scanning confocal and multi-photon microscope system. Recommended uses: Tissue sections, brain sections, biopsies, fly and worm embryos, adult worms, F-techniques (FRAP/FRET/FCS), deep imaging of thick specimens; live imaging, imaging of unconventional fluorescent dyes, separation of spectrally close dyes.

Special Techniques (FRAP, FRET, FCS, Ablation)

The Cellular Imaging shared resource provides instrumentation and support with a number of advanced microscopy techniques. Some examples of available techniques are listed below. We are always interested in trying out new things. If you are interested in an application that is not listed here, please contact the resource.


Cell with immunofluorescent staining for vinculin, a focal adhesion marker, imaged in conventional widefield fluorescence mode (left) and TIRF (right).

Cell with immunofluorescent staining for vinculin, a focal adhesion marker, imaged in conventional widefield fluorescence mode (left) and TIRF (right). Image courtesy of Anjali Teckchandani (Jon Cooper lab).
Power of TIRF
Total internal reflection fluorescence microscopy uses a special illumination technique to achieve selective fluorescence excitation of a very thin plane of approximately 100 nm, resulting in outstanding optical sectioning. This method is particularly powerful for the visualization of dim signals that are obscured by string fluorescence from the background. Typical applications include the imaging of molecules at the cell surface, such as coated pits, that would be obscured by strong cytoplasmic signal, or the imaging of molecules in vitro, such as reassembled microtubules.


Stochastic optical reconstruction microscopy is a recent super-resolution microscopy technique based on the principle of localization microscopy, that can provide images of cellular or in vitro components with resolutions down to 20 nm, or ten times better than conventional light microscopy. Several localization techniques have been implemented, but their basic principle is the same: a very small subset (less than 0.1 %) of the molecules in the field of view are imaged at any given time. This allows precise localization of each molecule by curve-fitting. The imaged molecules are bleached, and the cycle is repeated hundreds or thousans of times until most of the molecules have been localized. At our resource, we use Nikon's implementation (N-Storm). More details can be found at the vendor's web site.


Fluorescence recovery after photobleaching (and the related technique of fluorescence loss in photobleaching FLIP) are live-cell techniques for studying the motion of fluorescent molecules, such as protein localization or turn-over. In FRAP, fluorescent molecules are photobleached in a specified region with a pulse of intense light, and the recovery of fluorescence in the bleached region is measured over time with time lapse microscopy. The increase in fluorescence intensity over the bleached region indicates the relocalization to that region of fluorescent molecules from the unbleached portion of the cell. In FLIP, bleaching of a specified region will lead to a decrease in fluorescence in the non-bleached areas, due to re-localization of unbleached molecules. Both techniques can provide information about molecular motility (such as diffusion vs active transport), indicate whether molecular associations are stable or labile, and provide information about rates of protein synthesis and degradation. Our resource has equipment for performing FRAP and FLIP experiments, including the Ultraview spinning disk confocal system and the Zeiss LSM 780 confocal system.


Fluorescence (or Forster) resonance energy transfer is a microscopy technique for the study of molecular interactions. In FRET, if two fluorescent molecules are chosen so that the emission peak of the first molecule (donor) overlaps with the excitation peak of the second molecule (acceptor), energy transfer can happen when the two molecules are in close contact, so that when the donor molecule is excited, it transfers its energy to the donor which then fluoresces. Such effect can be obtained, for example, between a Cyan (CFP) and A Green (GFP) fluorescent protein in close contact. The technique can be used to determine whether two molecules tagged with the appropriate fluorescent labels are in close enough proximity to suggest direct molecular contact. Several implementations of FRET exist, including sensitized emission, whereby the acceptor molecule fluoresces upon excitatin of the donor molecule, and acceptor photobleaching whereby the signal from the donor molecule increases upon photobleaching of the acceptor. Our resource has equipment for the implementation of both methods.


Fluorescence correlation spectroscopy is a microscopy technique to measure rates of molecular motion. Several implementations of the technique exist. In the imaging resource, Fluorescence intensity fluctuations in a small volume can be acquired at very fast frame rates with the Zeiss LSM 780, and the data can be exported for analysis with third party software. For more details about FCS, please consult the resource.

Laser ablation:

Laser ablation is a technique that allows researchers to destroy cells or other structures under the microscope to gain insights into their role. In our resource, the Ultraview spinning disk confocal microscope is equipped with a high power nitrogen laser from Photonic Instruments (now Andor) that can be precisely targeted to a defined spot or region in the field of view. The operation is extremely simple and fully controlled by the software for precise control of the laser power and region to be targeted.

Transmitted Light and Fluorescence Microscopy

The Cellular Imaging Shared Resource provides support for both transmitted light and fluorescence microscopy. Transmitted light techniques available in the resource include brightfield, differential interference contrast (DIC, Nomarski), phase contrast, and polarized light microscopy. The resource relies on scientific grade monochrome and/or color microscopy cameras for the acquisition of high resolution digital transmitted light images suitable for publication and quantitative analysis. Instruments recommended for transmitted light microscopy include the Nikon E800, Nikon Live, Tissuefax, and Incucyte. Several other instruments are also capable of imaging in transmitted light mode. For details, follow the links to individual microscopes or services, or contact the resource.

Fluorescence Microscopy

The majority of modern microscopy techniques rely on fluorescence as a means to provide highly specific high contrast labeling of cells and biological molecules. Labeling can be direct (with fluorescent protein fusions, for example), or indirect (for instance with labeled antibodies or nucleic acid probes). Fluorescent techniques are compatible with both fixed and live specimens. All microscopes in the imaging resource are capable of fluorescence imaging. For technical details, use the information found for each instrument in the Instrument dropdown list. To find the microscope best suited to your needs, select compatible dyes and labels, or consult the resource for assistance with experiment planning.

Whole Slide Scanning and Analysis

Whole slide scanning and analysis (digital pathology) allows researchers to scan large specimens such as whole tissue sections, whole slides, and tissue micro array slides (TMAs) for archival, sharing, and analysis purposes. The acquired high resolution images provide a visual record that can be archived or shared with collaborators for evaluation and analysis. The images can also be processed via dedicated image cytometry software for unbiased quantitative analysis (scoring). The resource provides microscopy equipment for automated slide imaging (Tissuefax system), and TissueQuest and Histoquest software for scoring slides stained with immunohistochemical (H&E, DAB) or immunofluorescent methods.

Transmitted light image of whole mouse brain section acquired with 10x objective through automated image stitching (left), and full resolution detail (right). Transmitted light image - mouse brain section. Image stitching (left), full resolution detail (right).


(For technical details, follow the links to individual instruments or software below.)

  • Tissuegnostics Tissuefax system.he Tissuefax is an automated microscope for the imaging of histology slides, including large tissue sections and tissue micro-array (TMA) slides.
  • Recommended Uses: imaging of histology slides including smears, cytospins, and whole tissue sections. Imaging of tissue micro-array (TMA) slides. Slide digitization and archival; digital pathology; quantitation and scoring of IHC and IF staining.
  • Tissuegnostics TissueQuest software: Image cytometry software for the semi-automated analysis of fluorescent images acquired on the Tissuefax system.
  • Tissuegnostics HistoQuest software: Image cytometry software for the semi-automated analysis of immunohistochemistry images acquired on the Tissuefax system.

Scheduling time for instruments, training, and support:

To schedule time on any of our microscopes, or to schedule staff assistance or training, please use iLab. The resource is open to all from 9:00 AM - 5:00 PM, Monday through Friday. After hours access can be granted to experienced users and requires training by the resource's staff. Please contact Cellular Imaging for further details.

Please notify the resource if you need to cancel or change a reservation, so that we can make instruments available to others if needed. Users who are more than one hour late without notice may lose their reservation.

How to access your data:

Data acquired on Cellular Imaging instruments is transferred to the si folder in the user's Homelink account. If you do not have a Homelink account, please see Data Storage and Archiving for more information. Once you have obtained a Homelink account, please contact Cellular Imaging so that we can create an si link for you. External users without access to Homelink can obtain their data via ftp, or can download to their own data storage device from one of our dedicated computers. Please note that no portable drives are allowed on instrument computers.