Cell Biology and Cell Imaging Core
 Contact
John Williams, M.D., Ph.D.
Director
734-764-4376
Stephen A. Ernst, Ph.D.
Co-Director
734-763-8109
Services
- Tissue Preparation/Embedding/Sectioning
- Transmission Electron Microscopy
- Light Microscopic and Confocal Immunofluorescence Analysis
- Flourescence Analysis of Living Cells
- FRET and Other Flourescent Protein Visualization
- Image Analysis
- In situ Hybridization
- Autoradiography
- Training and Education
1. Tissue Preparation/Embedding/Sectioning
The primary services offered will be 1) fixation and cryosectioning preparatory for immunocytochemistry and 2) fixation, resin embedding and sectioning for high resolution LM and EM. Ultracyromicrotomy and embedding in special resins such as Lowicryl or LR White can also be performed. Fixation procedures will be discussed with the investigator. Instruction can be given in perfusion fixation of animals or fixation of cell subfractions in ultracentrifuge tubes. Fixatives and their recipes are available from the Core or alternatively tissue is given to the Core for fixation. Paraffin embedding will not be performed and investigators requiring this service will be directed to the Department of Pathology, which performs this routine service for a fee. Samples for resin embedding are processed with osmication as needed, dehydrated and embedded usually in Spurr resin. Each sample is initially sectioned as 1 mm sections, stained with toluidine blue and evaluated on the light level prior to electron microscopy.
2.Transmission Electron Microscopy
Blocks prepared as above will be sectioned as 60-100 nm ultrathin sections and picked up on nickel or copper grids. For routine analysis these will be stained with uranyl acetate and lead citrate and viewed and photographed on the Philips EM in the Department of Cell and Developmental Biology’s central research facility, the Microscopy and Image Analysis Laboratory (MIL). Investigators are invited to be present when specimens are viewed. Normally 5-10 images will be digitally captured and will be discussed with the Investigator. When appropriate, publication quality photographs will be prepared including digital processing. Immunocytochemistry by immunogold labeling or Protein A Gold will be carried out on cell fractions prior to embedding and for intact cells by on grid staining. Unless the investigator has a tried and true protocol this is labor intensive and requires trials with a number of fixatives, resins, and dilutions of antibodies.
3. Light Microscopic and Confocal Immunofluorescence Analysis
The emphasis here is on confocal microscopy as our experience has been that this will improve upon almost every type of immunohistochemistry, and provides ready access to digital image analyses, including 3-D reconstructions. Following conventional immunofluorescence to validate staining, sections will be analyzed on the LSCM either by Core personnel, or more generally, by users after training. All users will be provided 4 free hours of instruction. After this, involvement of Core personnel will be recharged. It is anticipated that most users will want to capture their own images but some clinical investigators may want full service with provision of hard copy images. Once trained a user will sign up and use the microscope and store their own images on CDs which can be provided by the Core at cost. Images can also be transferred to the user’s computer via the network. Both single sections and volume rendered stacks can be processed with deconvolution software to improve image sharpness. Assistance will be provided in designing multiwavelength analysis often combining two antibodies with a nuclear stain or fluorescent phalloidin to visualize actin.
4. Flourescence Analysis of Living Cells
Initial consultation with the Core Director will determine whether the study is better performed on the FRET microscope, the Attofluor, or the Olympus LSCM. Most work on cultured cells and/or where a ratioed image is desired will be performed on the Attofluor which is also easier for the user to learn. High resolution work, work on thick specimens or studies requiring high speed time resolution will be performed on the confocal microscope. We have experience to date using fura-2, fluo3, calcium green and Magfura dyes for Ca 2+, SBFI for Na +, BCECF for H+, SPQ for C1- and lucifer yellow and lissarhodamine as gap junction tracers and GFP and YFP fluorescent proteins as markers for transfected cells. Other probes are available for potential sensing, labeling of mitochondria and following endocytosis. Most projects in living cells have a developmental component and will require continued consultation with core personnel and between users. Investigators can be instructed in the use of stage mounted microinjection when it is appropriate for their work including the preparation and filling of pipettes. Both instruments can be set up to vary the frequency of image collection which effects both size of data files and bleaching of probes.
5. FRET and Other Flourescent Protein Visualization
Because all FRET work requires a large developmental component this is not offered as a routine service. The Core will provide instrumentation and training in its use. At present we are able to monitor energy transfer from CFP to YFP. Advice can be given on construction of tagged proteins but normally the investigator will prepare these. Other types of FRET will require purchase of appropriate filters by the investigator or jointly with the core if there is general interest. Because we wish to foster the development of such techniques there will initially be no charge for the use of the apparatus, although its continued use must be scheduled.
6. Image Analysis
Image analysis services include preparation of publication quality images and projectable images. This usually involves labeling and frequently production of montages. This will be done on our PC image analysis computer primarily using Photoshop. Investigators can use this workstation on an hourly recharge or this service can be provided by the core personnel. Instruction in the use of Photoshop is available from the Lab Director or the Associate Director. Quantitative analysis of static images can be performed using Image-1 software on a PC for analysis of area, size, shape, distance, and relative position of defined cellular and sub cellular structures. Relative volume densities can also be calculated by the point counting method of Weibel. Particle counting can be performed and expressed per cell area or membrane length. Quantitative analysis including 3D reconstruction, quantification of fluorescence intensity, etc., can also be performed. This software also performs transfer of video rate images to high resolution video tape. Analysis of time series data from the Attofluor is carried out on its own computer.
7. In situ Hybridization
Although not routinely offered as a Core service, the Laboratory Director, Dr. Lentz, has extensive experience with a large variety of in situ hybridization techniques for whole-mounts, tissue sections, and cultured cells from adult and embryonic tissues (mouse, rat, and zebrafish). Hybridization signals can be detected with either isotopic (35S, 33P) or non-isotopic methods (fluorescent and colorimetric). Multiple signals can be detected with double in situ hybridizations (using digoxigenin and fluorescein tagged cRNA probes) or in situ hybridizations combined with immunohistochemical staining of proteins (both peroxidase- and fluorescent-based detections of antibodies). Thus, the Core can provide help and assistance to an investigator interested in this technique.
8. Autoradiography
Although not routinely available as a Core service, the Core Director, Dr. Willaims, has experience with both light and EM autoradiography of newly synthesized protein to follow the secretory pathway and of localizing radioiodinated hormones bound to receptor on target cells. The latter can also be used in expression cloning of receptors. Thus the Core can provide help and assistance to an investigator interested in this technique.
9. Training and Education
Any investigator can be trained in any of the techniques used to prepare and analyze specimens. Training is available in all areas of Core services. The educational mission of the Core is also served in interacting with the investigators during the consultation and review and interpretation of data. To serve both the broadened GI Peptide Center and MDRTC membership and the University community the Core will also sponsor occasional lectures or workshops on new techniques and the use of Core instrumentation in which the Core directors or invited speakers will present.
Facilities
The Cell Imaging Core facilities are located in the East Ann Street Building which houses the Michigan Diabetes Research and Training Center (MDRTC) and Medical Sciences II Building which houses Drs. Williams’ and Ernst’s laboratories and the Morphology and Image Analysis Laboratory (MIL) of the Department of Cell and Developmental Biology. These two buildings are about two blocks apart, allowing easy interchange. Most of the GI Center investigators are located in the Medical Sciences and Medical Science Research Buildings, a complex of 5 attached buildings, which are also connected to the Cancer Center and University Hospital. In the next two years some investigators and the MIL Core Lab will move to the new Medical School Research Building (BSRB) scheduled to open in January 2006. It is only one block away from the Medical Sciences complex and the East Ann Street Building.
The MIAC laboratory occupies approximately 800 sq. ft. and consists of a large laboratory containing a fume hood, a bench for specimen preparation, and space for an office for the Laboratory Director as well as desk top computers and printers. Adjoining are rooms for tissue culture, the confocal microscope, and the multiwavelength fluorescence microscope. In addition, the Core has access to shared space in the MDRTC including walk-in cold storage, -70° freezer, autoclave and dishwashing facilities. A tissue culture incubator is available for short-term storage of cells under a controlled environment.
The other portion of the Core facilities are located in Room 4734 MS II assigned to Dr. Ernst where the research assistant, Bradley Nelson, is stationed. Specialized equipment there includes a cryostat, ultramicrotome and the Zeiss Attofluor workstation as well as usual laboratory equipment. It is located next door to the MIL, which contains the Phillips Electron Microscope and two confocal microscopes (Olympus Fluoview 500 and Zeiss LSM510), which can be used as back-up for our own microscopes in case of significant down-time. Dr. Williams’ office is located three floors above in MS II as is the Department of Molecular & Integrative Physiology Machine Shop, which is available on a recharge basis to develop and fabricate equipment such as observation chambers and solution reservoirs. All of the facilities are Ethernet connected by the U of M fiber optic network, which also provides high speed access to the internet.
Equipment
1. Olympus FluoView 500 Laser Scanning Confocal Microscope is controlled by a 2.4 Ghz personal computer under Windows 2000 and is capable of imaging 5 separate channels simultaneously (4 fluorescence + 1 transmitted light photomultiplier detectors) offering highly efficient, maximum emission sensitivity and the ability to record scanned images in 12 bits or 4096 gray levels, thus allowing quantitative linear measurement of fluorescence within regions of low contrast as well as very high contrast. Users are able to image a wide variety of fluorophores with laser excitation that includes Blue Violet (405 nm), Multi-Line Argon Blue (458,488,515nm), Helium Neon Green (543nm) and Helium Neon Red (633nm) for standard Blue, Green, Red and Far-Red fluorochromes. The FV500’s acoustical optical tuning filter (AOTF) and adjustable scan speeds provides for minimal specimen fading, sequential scanning for reduced fluorescence cross talk, multiple regions of excitation, high resolution imaging (up to 2048 x 2048 pixels) of fixed or static samples, and rapid recording of kinetic events. Optical sections in the z plane can be collected using a step motor attached to the fine focus control of the microscope and driven by Fluoview software. The system is also equipped with Differential Interference Contrast (DIC) objectives and condensers and has the ability to capture transmitted light images with a highly sensitive photomultiplier (PMT) transmission dector.
The Images can be saved to a peripheral hard drive for later analysis. Integral software allows for analysis of saved images in 2 dimensions (e.g., brightness vs. time); confocal images obtained in a “z” series can be volume rendered and analyzed in 3 dimensions. Data can be archived on CD, DVD, or Zip disks, or transferred to an alternate image analysis platform with greater storage capacity.
2. Fluorescence Resonance Energy Transfer (FRET) System
The MIAC has recently developed a new state-of-the-art fluorescence resonance energy transfer (FRET) system. This type of microscopy will enable investigators to study a wide variety of biological events that influence the interaction between molecules. FRET technology involves the non-radioactive transfer of energy from a fluorophore in an excited state to a nearby acceptor fluorophore. For this transfer to occur, the donor and acceptor molecules must be less than 10 nanometers apart and the emission spectra of the donor fluorophore must overlap the excitation spectra of the acceptor fluorophore. The farther apart the two molecules are, the weaker the transfer efficiency. FRET has grown in popularity due to the emergence of GFP mutants with blue or yellow-shifted spectral properties. We have developed our system to take advantage of the fluorescent characteristics of enhanced cyan fluorescent protein (CFP) and yellow fluorescent protein (YFP).
FRET microscopy relies of the ability to capture weak and transient fluorescent signals efficiently and rapidly from the interactions of labeled molecules in live samples. This necessitates the use of a digital camera that can perform under these stringent conditions. Our FRET system is mounted on a Nikon Diaphot 200 inverted stage microscopy equipped with specialized Chroma CFP and YFP excitations/emission filters, Sutter excitation and emission filter wheels and controller (Lambda 10-2), a Prior automated x, y, z stage, a TE-ICV incubator with digital thermistor probe and incubator case, and a Hamamatsu ORCA extended range digital CCC camera (C4742-95-12ER). This camera rapidly captures images at rates ranging from 8.3-45 frames per second with very high quantum efficiency resulting in shorter exposures of sensitive samples to fluorescent light. Thus, the ORCA digital camera allows us to get the maximum performance and utility from FRET microscopy. The system is controlled by as Dell 2.4 GHz Pentium IV personal computer with 768 MB of RAM, Matrox Meteor II digital PCI frame grabber, and an 80 GB hard drive running Windows 2000 operating system and Metamorph software (version 6.1r3 Universal Imaging Corporation). The acquisition and analysis of FRET data is semi-automated with the use of specialized Metamorph macros/journals developed by Dr. Joel Swanson, the MIAC FRET consultant. Images and data can be exported in convenient formats including tiffs and Microsoft Excel spread sheets and archived on an internal CD-RW drive.
3. Zeiss-Attofluor Fluorescence Digital Imaging System
The instrument provides a user-friendly system for fluorescence based measurements of intracellular ion concentration in living cells and visualization of EGFP or EYFP and labeled proteins. The system configuration consists of an ATTOFLUOR Ratiovision system coupled to a Zeiss Axiovert 35 inverted microscope equipped with a 40 x high numerical aperture oil immersion lens. Various peripherals, including a color printer and video recorder are interfaced with the system. The ATTOFLUOR system exploits the optical properties of many commercially available probes by providing both single and dual excitation by means of a four position filter changer and intensified CCD camera to capture the emission defined by a dichroic filter cube. This information is subsequently digitized and stored either to RAM or hard drive. The system can be used to gain useful temporal information (video rate measurement of ion-concentration in single excitation mode; or 4 measurements per second in dual excitation/ratio mode) and important sub-cellular spatial information (allowing measurement on a pixel by pixel basis from 99 user defined regions of interest). Currently suitable probes are available for many physiological relevant ions, including Ca2+, Mg2+, Na+, K+ and Cl- , together with fluorescent tracers useful as indicators of gap-junctional permeability and EGFP and derivatives for identifying transfected cells. The microscope is mounted on a vibration isolation platform and is equipped with stage temperature control, together with a Burleigh Cell Penetrator microinjection system. A CCD camera attached to the 35 mm port is used for visual control of cell injection.
4. The MetaMorph® Imaging System
The Metamorph system is a powerful integration of software and hardware from Universal Imaging Corporation that enables core users to automatically capture and analyze microscopy images from a color video CCD camera. The system consists of a Nikon Microphot-FXA microscope capable of capturing bright field and fluorescent (rhodamine and FITC) images on a Spot-RT color CCD camera. The program is run on a Dell Optiplex computer equipped to facilitate the manipulation of large image files. Data can be easily archived with either an Iomega Zip or a CD-RW drive. High quality montages of large specimens can be rapidly acquired with the use of Metamorph’s ability to precisely control the stepping of a Ludl automated x,y,z stage. Multiple users of the core require the ability to determine the percentage of cells that have a certain characteristic (e.g. apoptotic, myelinated/unmyelinated axons, or protein expression). For these studies, it is important to collect a set of images of an entire tissue section or tissue culture plate. Once captured, the software can rapidly count objects or average pixel areas according to our researcher’s protocols. Metamorph’s sophisticated automation features such as image stacks, and a macro capability enable unattended image acquisitions and the ability to process data sets containing hundreds of images with ease. The addition of color video microscopy allows for more accurate and timely collection and analysis of chemically labeled cells and tissues compared to problems associated with black and white systems. The core has the complete version of Metamorph that comes with all the device drivers (stages, Z-focus motors, filter wheels, etc.) and all the advanced processing capabilities (motion analysis, colocalization, multi-dimensional acquisition and analysis, etc.) built in for maximum flexibility. This version also has the added advantage of providing us with access to future enhancements of acquisition and processing tools and techniques that are continually being added to MetaMorph. We get the latest developments as they become available by simply upgrading to the most recent version of the software.
5. Phillips CM-100 Transmission Electron Microscopy (TEM)
The Phillips CM-100 TEM is maintained and operated in the Morphology and Image Analysis Laboratory (MIL), part of the Department of Cell and Developmental Biology. It is equipped with a motorized stage and a Kodak 1.6 megaplus digital camera capable of capturing electron images directly from the viewing screen. This microscope will be used for the collection of digital EM images and for the direct transfer of data from the microscope to the image analysis software reducing the need for photographic film, chemicals, and paper. Use of this equipment is available to all University of Michigan investigators on a recharge basis for $40/hour.
6. Ancillary Equipment
Other equipment available in the MIAC or Dr. Ernst’s laboratory include two cryostats, a RMC MT-7 ultramicrotome and CR-21 cryosectioning attachment, knifemakers and a Kodak Digital Sciences 865OPS dye sublimation printer.
Facilities
The Cell Imaging Core facilities are located in the East Ann Street Building which houses the Michigan Diabetes Research and Training Center (MDRTC) and Medical Sciences II Building which houses Drs. Williams’ and Ernst’s laboratories and the Morphology and Image Analysis Laboratory (MIL) of the Department of Cell and Developmental Biology. These two buildings are about two blocks apart, allowing easy interchange. Most of the GI Center investigators are located in the Medical Sciences and Medical Science Research Buildings, a complex of 5 attached buildings, which are also connected to the Cancer Center and University Hospital. In the next two years some investigators and the MIL Core Lab will move to the new Medical School Research Building (BSRB) scheduled to open in January 2006. It is only one block away from the Medical Sciences complex and the East Ann Street Building.
The MIAC laboratory occupies approximately 800 sq. ft. and consists of a large laboratory containing a fume hood, a bench for specimen preparation, and space for an office for the Laboratory Director as well as desk top computers and printers. Adjoining are rooms for tissue culture, the confocal microscope, and the multiwavelength fluorescence microscope. In addition, the Core has access to shared space in the MDRTC including walk-in cold storage, -70° freezer, autoclave and dishwashing facilities. A tissue culture incubator is available for short-term storage of cells under a controlled environment.
The other portion of the Core facilities are located in Room 4734 MS II assigned to Dr. Ernst where the research assistant, Bradley Nelson, is stationed. Specialized equipment there includes a cryostat, ultramicrotome and the Zeiss Attofluor workstation as well as usual laboratory equipment. It is located next door to the MIL, which contains the Phillips Electron Microscope and two confocal microscopes (Olympus Fluoview 500 and Zeiss LSM510), which can be used as back-up for our own microscopes in case of significant down-time. Dr. Williams’ office is located three floors above in MS II as is the Department of Molecular & Integrative Physiology Machine Shop, which is available on a recharge basis to develop and fabricate equipment such as observation chambers and solution reservoirs. All of the facilities are Ethernet connected by the U of M fiber optic network, which also provides high speed access to the internet.
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