Overview of Multiphoton/Electrophysiology Core Facility Overview

Overview of the Multiphoton/Electrophysiology Core Facility

Our Mission

The goal of the Multiphoton Imaging/Electrophysiology Core is to provide instrumentation for analyzing protein localization, protein dynamics, and protein-protein interactions with high resolution. This facility also allows users to perform time-lapse imaging of multiple fluorophores in living cells and tissues, and to combine high resolution imaging of fluorescently tagged proteins or ion indicator dyes with electrophysiological monitoring of electrical activity.

The Multiphoton Imaging Core promotes interactions among a diverse group of neuroscientists at Johns Hopkins University. This has facilitated the completion of many studies that represent advances relevant to key issues in basic and clinical neuroscience.

[Publications presenting work supported by the Multiphoton Imaging Core]

Location

The Multiphoton Imaging Core is located in Room 1008, Preclinical Teaching Building, 725 North Wolfe St. The phone number is 410 502 7858.

Support

A great deal of expertise is available to users of the Multiphoton Core Facility. Michele Pucak, PhD (Facility Manager) has experience in single and multiphoton imaging, immunohistochemistry, maintenance of animals for in vivo imaging, and electrophysiological techniques. The facility Director, Dr. Dwight Bergles, has extensive expertise in imaging and electrophysiological techniques. Furthermore, the many users of the facility each bring unique knowledge and experience, so that investigators profit from informal interaction with each other. More structured events include regular seminars in which investigators from within and outside Hopkins share their experiences with developing in vivo imaging preparations, and meetings among graduate students and postdocs with an interest in sharing knowledge regarding image processing techniques.

Core Service

The facility contains two confocal microscope systems that allow high resolution imaging of labeled cell components in three dimensional space. The first system consists of a Zeiss LSM 510 motorized upright microscope, a META scanning module, two non-descanned detectors for multi-photon work, and a motorized stage for time lapse imaging. The LSM 510 can be configured with the META detector plus 2 traditional detectors for fluorescent or reflected light imaging. The META detector can also be used as a unique spectral detector as well as a conventional "band-pass" detector. All 3 detectors can be recorded and displayed simultaneously. This system employs PC-based user interface to control the microscope, laser excitation, scanning and image acquisition, and is capable of digitally capturing up to 12 bit 2K x 2K pixel images, while the LSM software controls image data acquisition and processing. The system can perform Fluorescent Recovery after Photobleaching (FRAP), Fluorescence Resonance Energy Transfer (FRET), and quantitative co-localization. The META detector provides multi-spectral imaging.

The second system is a Zeiss AxioExaminer upright microscope with an LSM 710 scan head. Due to refinements of the light path within the scan head, the system has better detection capabilities that improve the signal-to-noise ratio of fluorescent images. In addition, the filter-free spectral detection allows the user to define emission collection parameters, rather than being limited to a particular set of installed emission filters. Furthermore, this instrument has a number of distinct advantages for multiphoton imaging, including a redesigned microscope stand that has increased objective travel, allowing the use of a new 20x high numerical aperture objective with increased transmission in the infrared. An important feature of this microscope is the addition of a low noise photomultiplier tube placed immediately adjacent to the objective for optimal light capture. The 710 can also be used with a highly sensitive GaAsP detector, which excels at collecting images of fluorescently labeled cells deep within tissue. This improvement has been particularly critical for in vivo applications, and has allowed users to monitor cell motility in intact preparations. Because the GaAsP detector is extremely sensitive to ambient light, we have constructed a light-tight enclosure around this system, so that both systems can be used simultaneously.

[click here for a list of lasers, dichroics, and emission filters for each system]

The Core possesses a near-infrared tunable pulsed femtosecond Ti:Sapphire laser (Chameleon Ultra II). The 510 and 710 systems are positioned on the same table, so that the two systems can share the infrared beam from this laser. The IR laser of the two-photon system facilitates imaging of thick specimens, and is capable of in vivo imaging of fluorescent dyes or proteins.

Custom, interchangeable temperature-controlled stages are available for use in live cell/tissue imaging and computer controlled multisite recording. Tissue superfusion is achieved using a gravity-fed system that passes through computer-controlled solenoid valves. Solutions can be oxygenated and, if necessary, heated using an in-line heater or peltier chamber heater, depending on the application. Microscopes are also equipped with DIC optics to allow visualization of cells in brain slices, and imaging can be performed using a B & W CCD camera or using laser illumination and a transmitted light detector. With the assistance of Zeiss, we have developed a custom macro which enables mapping of receptor density using 2-photon mediated photolysis. Components which are critical for supporting successful imaging of labeled cells in vivo are present in the Core. As previously mentioned, the GaAsp detector greatly facilities imaging of fluorescent cells located deep within tissue. The increased objective travel of the 710 facilitates positioning of animals beneath the objective. Finally, the core contains an adjustable, heated platform for immobilizing the head or spinal cord of animals during imaging, administering anesthesia, and maintaining animal body temperature.

The facility also has the capability to couple high resolution imaging with electrophysiological recording of currents and voltages. Electrophysiological data can be acquired with an Axon Instruments Axopatch 200B patch amplifier, Axon Instrument DigiData 1440A A-D converter coupled to a stand alone PC. In addition, there is a Master 8 timer for applying controlled stimuli, a picrospritzer for locally applying drugs, and a video camera module for IR-DIC imaging.

The Core offers a wide range of objectives (5x, 10x, and 20x air; 25x multi-immersion; 40x, 63x, and 100x oil; and 40x and 63x water immersion). In addition, we have a piezoelectric focus drive for z-focus control. Piezoelectric positioning drives provide higher focusing speed (~10 vs 100 msec) and better resolution (~1 vs. 100 nm) than stepper motor-drives, and thus have the ability to provide higher resolution 3D images in a shorter time. This is especially advantageous for applications, such as tracking mitochondrial movement and assessing process motility of labeled cells in vivo, in which the ability to assess temporal changes is limited by the time required to construct high-quality z-stack images.

In addition, the facility has two dedicated image analysis computers each running Imaris software from Bitplane for performing deconvolution, 3D reconstruction, isosurface rendering, filament tracing/particle tracking, and quantitative co-localization. The Imaris suite of programs has been designed to accept images in many different formats, include Tiff series, BMP series, and images collected with Leica, Olympus, and Zeiss acquisition software. The available modules are Imaris suite, ImarisXT, Filament tracer, and AutoDeblur. By using ImarisXT, users can develop their own, task-oriented algorithms for Imaris, or use analysis modules developed by others. AutoDeblur is a deconvolution software package that produces extremely high-quality results through image restoration.

Contact and Scheduling Information

Core investigators should contact Michele Pucak, Ph.D. (mpucak1@jhmi.edu; 410 502 7858), the facility manager, or Dwight Bergles, Ph.D. (core director; dbergles@jhmi.edu) to discuss their project and facility procedures. Use of microscope must be scheduled with Michele Pucak in advance..

Facility Manager

Michele Pucak, Ph.D.

Dwight Bergles , Ph.D.

Core Director

Click here to see the current schedule for the 510/710 microscope.

Click here to see the current schedule for the Imaris 1 System.

Click here to see the current schedule for the Imaris 2 System.

See this document describing system start-up, shut-down, and software