Lynn & Mike Garvey established the Garvey imaging core to support research in stem cell and regenerative medicine. The Garvey core provides more than 50 laboratories access to state-of-the-art light microscopy systems, as well as training and assistance with imaging applications.

The core continues to be supported by the Garvey family gift, and recieves additional resources from the State of Washington and the UW Student Technology Fund.

Core Access and fees:

The Garvey Imaging Core provides microscopy equipment and expertise to more than 50 labs. Lynn and Mike Garvey established the core to help advance stem cell and regenerative medicine, and our goal is to facilitate your research. The core is accessible 24/7 to trained users. Between 9 AM and 6 PM you can reserve sessions up to three hours long.

We charge $15 per hour for use of the A1R, SP8, and Spinning Disk confocal systems. To use these and the wide-field systems, you need to be a full or affiliate member of ISCRM. The multiphoton system is available to any UW community researcher at no cost, and we can arrange access to researchers outside the UW system for a nominal fee.


All users must be trained on each instrument that they wish to use. To schedule a training session, contact Dale Hailey at Training sessions will teach you how to operate the equipment, demonstrate capabilities of the equipment, and help you optimize your particular work. It may help if you bring your own sample, but it’s not necessary.


Time on each microscope is booked via sign-up calendars here.

Please adhere to your scheduled time. Users who are over 30 minutes late forfeit their booking. If you need to cancel a session, remove your name from the calendar and notify the core director. Users who repeatedly don’t show may lose access to the core. When you are done, check the calendar to see if anyone is using the system after you. If you are the last person signed up for the day, make sure you turn off the system.

Handling hazardous samples:

For the safety of users, inform the director of any health risks associated with your samples. Live human cell lines, primary cultures and virally transduced cells may be considered BSL-2 specimens and require special handling specified by hazardous material protocols. Also, be sure to appropriately contain any hazardous chemicals present in your samples, including common reagents like DAPI-Fluoromount which has both sodium azide and a DNA binding agent. Seal slides to avoid getting mounting agents and other chemicals on yourself and core equipment.

Managing files:

To minimize risk to our computers, please don’t transfer files by plugging in external or USB Flash drives. We maintain a server as a file transfer point. You can write files directly to this server and then move files from it to your own storage site. You can also save files to cloud-based storage (like your UW google drive) via the web. For long experiments (multipanel fields, large Z-stacks, or overnight experiments), write files to the internal data drive. When the experiment is finished, transfer the file from the internal data drive to the server or off-site storage, and then delete it from the internal drive. (This prevents your experiment from freezing if the server loses connection.) Users are responsible for moving and backing up their data. Please don’t use drives on any of the microscope computers for long term storage.

We expect users to operate the systems appropriately and keep the equipment and space clean and organized. This includes cleaning objectives and microscope and counter surfaces, and removing any materials you have brought into the facility. We reserve the right to deny access if the equipment or space is not appropriately cared for.

Please contact the core director if you have questions about access or about any of the policies above.

To use any of the microscopes in the Garvey Core, you need to be trained on the system you want to use. Please click here for information about training and use.

Nikon A1R Confocal

The A1R is a point scanning confocal system used to image samples with up to four fluorescent labels (blue, green, red, and far-red). It is mounted on an inverted microscope with six objective lenses (10X – 60X). Users can take serial optical sections to create 3D reconstructions. With the motorized stage, it is also simple to aquire sequential panels and stitch together images of large areas. The scanning lasers can also be targeted to regions of interest for experiments that require photobleaching, photoactivation, or photo-uncaging (e.g. FRAP, FLIP, or optogenetics studies). The system can also be configured to image other fluorophores like the cyan and yellow fluorescent proteins and FRET sensors.

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Olympus FV1000 Multi-photon

The Olympus Multi-photon microscope is designed to image thick specimens. It has a tunable laser that can emit excitation wavelengths from 690 to 1020nm. These long wavelengths are better able to penetrate samples, and generate less phototoxicity. Because 2-Photon excitation only occurs at the focal point of the objective, the system acquires optical sections, and 3-D reconstructions can be made from serial sections. The system is mounted on an upright microscope with 10X, 25X, and 60X dipping lenses and a motorized stage. Using exchangeable filter sets, 2 colors can be captured simultaneously–Blue and Green, Green and Red, or Cyan and Yellow. The Multi-Photon can also be used for SHG (Second Harmonic Generation) imaging in the backscatter direction, and for photodegradation-based microengineering.

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High-Resolution Widefield

The High Resolution Widefield system has a high resolution CCD camera, and is optimized for fluorescent microscopy of fixed cells and tissues. The microscope has filters for Blue, Green, Red, and Far Red fluorescent probes, and other filter sets can be swapped in. It is an inverted system with 4X to 60X lenses, a motorized filter changer, and a motorized XY stage. The system can rapidly survey samples and create mosaic images of large specimens. It can also be programmed for complex, automated image capture, and can be configured to suit a wide variety of specimens and experimental needs.

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Live-Cell Inverted Widefield/Spinning Disk Confocal

The Live-Cell Widefield System has a very sensitive EMCCD camera and is capable of imaging at 100 frames per second (with reduced field size). It has lenses from 4X to 60X (including phase and extra long working distance objectives), and has an enclosure that can control both temperature and CO2 levels. The microscope can be programmed for long-term imaging across multiple positions, and is routinely used for applications like wound healing assays, cell division assays, and calcium imaging using dyes excitable above 400nm like Fluo-4.

The Live-Cell system also supports spinning disk confocal microscopy. It has 405, 488, 561, and 640nm lasers that can excite a range of blue, green, red and far red fluorophores. The Yokogawa W1 spinning disk head on this system was specifically designed to minimize crosstalk between pinholes in the disk. It is capable of high speed volumetric imaging, even in relatively thick specimens. This spinning disk confocal is also well-suited for confocal imaging of very weak signals or capturing thin z sections at high framerates.

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2020 contest winners



200 um-thick section of human liver tissue immunostained with anti-CD32b to label sinusoidal endothelial cells, the highly specialized cells lining the fenestrated sinusoidal capillaries of the liver. [Video Courtesy of Chelsea Fortin]

Compartmentalization of hepatic and endothelial cells in close proximity using 3D bioprinting with vascular endothelial ‘cords’ shown in red and rat Hepatic aggregates in green. [Video Courtesy of Daniel Corbett]

Rainbow human iPSCs culture imaged every 12 minutes over 24 hours of culture (days 3-4 after plating) [Video Courtesy of Danny El-Nachef]

To view the 2018 image contest winners, click here.

Dale Hailey, PhD, Director

Dale Hailey is the director of the Garvey Imaging core. Dr. Hailey received his PhD in 2008 in the labs of Jennifer Lippincott-Schwartz (NIH) and Ian Mather (University of Maryland, College Park). He has used fluorescence microscopy in a wide variety of settings. From 1998 to 2001 he worked at the University of Washington Yeast Resource Center. He developed Fluorescence Resonance Energy Transfer and deconvolution-based assays to study centrosome structure with Professor Trisha Davis. From 2001 to 2008, he worked in the Lippincott-Schwartz lab. He and Dr. Holger Lorenz developed techniques to evaluate protein topology in membranes, and to discriminate freely diffusing from membrane-bound protein subpopulations. For his graduate dissertation, Dr. Hailey investigated how autophagosomes form under different induction conditions. In 2009, he joined Professor Dave Raible’s and Professor Ed Rubel’s labs to study aminoglycoside induced cell death. In humans, these antibiotics kill sensory cells and cause permanent hearing loss. Dr. Hailey studied how aminoglycosides enter and traffic through cells, and how they activate cell death pathways in mechanosensory cells in zebrafish.

Dr. Hailey trains users of the Garvey imaging core, advises conventional imaging projects, and helps ISCRM researchers with more complicated applications. Institute researchers work with microfluidic devices that mimic physiology, organoids that model disease states, and patterned cell culture environments that promote tissue maturity and development. Dr. Hailey is available to help researchers use these platforms, trouble shoots experimental design, and assists with laser-based microfabrication projects.

His full publication list is available here: