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Protein Identification and Localization Core

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Protein Identification and Localization Core


John A. Williams, M.D., Ph.D.

Stephen A. Ernst, Ph.D.

Philip C. Andrews, Ph.D.
Co-Director and Director of the Proteomics Core




  1. Standard Protein Identification by LCMSMS
  2. Global proteome analysis (2D LCMSMS)
  3. Quantitative Proteomics using iTRAQ, SILAC, and related techniques
  4. 2D Gel analysis
  5. Mapping sites of post-translational modification and global quantitative phosphoprotein analysis
  6. Tissue Preparation/ Embedding/Sectioning
  7. Confocal Fluorescence Microscopy (Microscopy Service Request Form)
  8. Immunohistochemistry
  9. Transmission Electron Microscopy (Microscopy Service Request Form)
  10. Live Cell Imaging (Microscopy Service Request Form)

1.  Standard Protein Identification by LCMSMS
Routine protein identification is performed by tandem mass spectrometry either directly from proteolytic digests or via LCMSMS of proteolytic digests. Common sources of proteins are from gel bands or spots on 2D gels, soluble proteins purified using affinity columns, or simple mixtures of proteins purified via TAP tags or similar methodologies. LCMSMS is performed on capillary reversed-phase columns either coupled directly to our Thermo Fisher Orbitrap or via collection on a large format MALDI sample plate and analyzed offline on one of our Sciex model 4800 MALDI TOFTOF tandem mass spectrometers. Sensitivities are similar between the two instruments and the specific workflow chosen is dependent on the specific needs of the project and on instrument availability.

Data are generally analyzed using one of the several search engines we have available (Sequest, Protein Pilot, Mascot, X!Tandem) depending on the instrument used, sample properties, and any protein chemistries used in conjunction with the analysis. Global false positive rates may be estimated using a reverse database. Sequence coverage is often an issue when identifying sites of chemical modification and this is obviated using multiple proteases, various modification chemistries and manual de novo analysis as needed.

2. Global proteome analysis (2D LCMSMS)
Global proteome analysis is generally performed as described above except that proteins are generally resolved by 2D LC methods (typically SCX followed by reversed-phase) prior to tandem mass spectrometry. Most global proteome analysis is performed using quantitative methods as described below, although label-free methods may be used for screening purposes. Computational methods are much as described above except that false positive rates are generally determined using the Scaffold software which is similar to Peptide Prophet in function.

3.  Quantitative Proteomics using iTRAQ, SILAC, and related techniques
The bulk of global proteomics analyses performed by the Core are quantitative in nature with isotope enrichment being the major methodology. The specific method used varies with the nature of the sample and the goals of the experiment, but the largest number have used isobaric tags (iTRAQ), while SILAC and ICAT also have significant usage. The separation methodologies are generally the same as described above for global proteome analysis. Either the Orbitrap or the model 4800 MALDI TOFTOFs may be used for quantitative analyses although isobaric tags perform considerably better on the TOFTOFs. Quantitative proteomics data is generally analyzed using Protein Pilot, Scaffold Q+, or our own algorithms, depending on the method used.

4.  2D Gel analysis
2D gel separation analysis is performed by routine methodology using immobilized pH gradients for the first dimension and SDS slab gel electrophoresis for the second dimension. Staining methods employed include Coomassie Blue, Silver, and several fluorescent methods. Image analysis is available using the Z3 software. Large, medium, and small format 2D gels are provided and sample complexity and size generally determines the gel format used.

5.  Mapping sites of post-translational modification and global quantitative phosphoprotein analysis
Quantitative phosphoproteome analysis is a new service for the core which takes advantage of the expertise in the Andrews research laboratory. Quantitation is currently only by SILAC labeling (duplex and triplex) although other quantitative methods are being explored. The methodology currently used is a modification of standard 2D LCMSMS in which one or more affinity separations are inserted between the SCX and the reversed-phase capillary HPLC columns. Phosphopeptide affinity methods utilized are zirconium dioxide and/or antiphosphotyrosine immunoaffinity chromatography. Global protein levels are determined by analysis of the flow-through from the phosphoaffinity columns to provide verification that the level of phosphorylation has changed and not just the protein level. Tandem mass spectrometry is performed on the Thermo Fisher Orbitrap and the data are analyzed using MaxQuant or the TransProteome Pipeline. Mammalian cell lines generally result in the identification of phosphorylated peptides from over one thousand proteins. Identified peptides are recategorized to correct for misassigned sites of phosphorylation. Initial evaluation of signaling pathways use GeneGo or Cytoscape with the appropriate plugins.

6.  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. Frozen sectioning can be provided by the Core or investigators may sign up and use the Core’s cryostat. Training will be provided when necessary. 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. Samples for resin embedding are processed with osmication as needed, dehydrated and embedded usually in Epon or Spurr resins. Each sample is initially sectioned as 1 mm sections, stained with toluidine blue and evaluated on the light level prior to electron microscopy. Paraffin embedding will not be performed and investigators requiring this service will be directed to the Cancer Center or the Department of Pathology, which perform this routine service for a fee.

7.   Confocal Fluorescence Microscopy
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. Users with advice from the Core will pick which confocal microscope is suitable for their needs with emphasis on location. Available scopes include a Olympus Fluoview 500 and Zeiss LSM510 in BSRB and the Olympus Fluoview 500 in the Diabetes Center. Training is arranged by the User with the selected Core. It is anticipated that most users will want to capture their own images but some clinical investigators for pilot projects 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. 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 by Dr Ernst in designing multiwavelength analysis often combining two antibodies with a nuclear stain or fluorescent phalloidin to visualize actin. Image analysis software packages including Volocity (version 5.02 64-bit, Improvision), Autoquant (version X2.1-64-bit, Media Cybernetics, Inc.) and MetaMorph (version 7.6.3, Molecular Devices) are available for use in the Diabetes Center morphology core. The PIL Core will provide subsidy of 50% of the cost (currently $50/hour) for microscope use or training. Investigators are limited to 20 hours of subsidy. Twenty users have indicated an interest in this service and we expect more than in the previous period as use is no longer restricted to a single confocal microscope. Microscope subsidy request form.

8.  Immunohistochemistry
The primary functions here are carrying out pilot studies, providing advice on fluorophore selection and providing training. These functions are carried out by Dr. Ernst and Bradley Nelson and are based on standard techniques established by the Core and used in a large number of previously published studies. We can combine one to three color immunofluorescence with DAPI staining of nuclei or fluorescent phalloidin staining of filamentous actin as well as a Nomarski image. This makes use of the 405nm laser for staining of DAPI and second antibody fluorescent conjugates such as the Alexa dyes (e.g., 488, 543, 563, 594 and 647 nm). For standard techniques such as immunolocalization of DAPI, staining for PCNA, digestive enzymes such as amylase and visualization of actin we use predetermined conditions. For novel proteins the Core can advise on antibody selection and the use of a dilution series and appropriate controls. This aspect of the Core has a significant teaching function as new investigators can come to the Core lab to carry out immunostaining or perform it in their own laboratory. We use a standard widefield fluorescence microscope to initially establish fluorescent staining and then direct the investigator to an appropriate microscope.

9.   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. Microscope subsidy request form.

10.  Live Cell Imaging
The core has considerable experience in this area and can provide advice and assistance with live cell imaging by confocal or widefield microscopy. Most of our experience is using the Olympus Confocal or the Nikon wide field microscope in the Diabetes Center Core and these will be the primary microscopes used. Hands on help with these microscopes can be provided by Dr Ernst or arranged with Dr Stephen Lentz, the Laboratory Director of the Diabetes Center Core. Three main types of live cell imaging are supported by the PIL Core. First, the observation and translocation of fluorescent tagged proteins, usually a GFP or related tagged protein (e.g., CFP, cYFP, RFP). The Molecular Biology Core can provide advice on vector construction. This is the simplest type of imaging and involves recording digital images at second or minute intervals. Secondly, the Core can assist with the use of small molecule fluorescent probes such as fura-2 or fluo3 or 4 for following changes in intracellular free Ca2+. Other dyes such as SBFI for Na+, BCECF for H+ or SPQ for Cl- can also be used. Other probes are available for potential sensing, labeling of mitochondria and following of endocytosis. These measurements are often ratiometric and carried out on the wide field scope or single wavelength and carried out on the confocal with results expressed as F/Fo The third type of study involves more advanced techniques such as FRET, FLIP and FRAP (see Past Utilization and Progress section). Dr. Ernst with advice from Dr Edward Stuenkel supervises this component of the Core. At present we are able to monitor energy transfer from CFP to YFP. Other types of FRET will require purchase of appropriate filters by the investigator or jointly with the core if there is general interest. Most projects in living cells have a developmental component and require continued consultation with core personnel. Both instruments can be set up to vary the frequency of image collection which affects both the size of data files and bleaching of probes. Specialized software is required for analysis. We use Metafluor for turnkey operation measuring Ca2+ and in house software based on Metamorph for FRET measurements. Microscope subsidy request form.



Protein Identification Component

  1. Three Applied Biosystems model 4800 MALDI TOF/TOF mass spectrometers for proteome mapping, identification of post-translational modifications, and iTRAQ quantitation with 10 ppm mass accuracy and sensitivity to at least one femtomole.
  2. ThermoFisher Orbitrap with an Advion Nanomate interface that improves reproducibility and sensitivity for phosphoproteome analyses.
  3. Virgen InstrumentsMiniTOF for peptide mass fingerprinting, development, and QC
  4. Two Agilent HPLC with MALDI fraction collector for interfacing LC with the TOFTO
  5. Eksigent capillary HPLC with non-ferrous solvent path coupled to the Orbitrap
  6. Michrom HPLC for SCX separations
  7. Sample handling robot to support 2D gels
  8. Laser and transmittance scanners for imaging 2D gels
  9. 2D gel apparati with capacity for up to 40 gels per day
  10. Off-gel electrophoresis system for preparative isoelectric focusing.

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. A similar microscope with slightly different peripherals but running under the identical operating system is available for use in the MIL.

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. (Microscopy Service Request Form)

2.  Zeiss LSM 510-META Laser Scanning Confocol Microscope
This micropscope is mounted on a Zeiss Axiovert 100M inverted microscope is located in the MIL. This instrument is equipped with four lasers offering a wide range of excitation wavelengths. Coherent Enterprise laser for UV (351,364 nm), Argon laser for FITC/GFP (458, 488, 514 nm), Helium Neon 1 laser for Rhodamine, Texas Red, Cy3 (543 nm), and Helium Neon 2 laser for Cy5 (633 nm). Using the META system, the researcher can easily separate highly overlapping emission peaks. (Microscopy Service Request Form)

3.  Nikon widefield inverted stage microscope equipped for FRET analysis
This instrument is located in the MDRTC Core. It is 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 acquisition and analysis of FRET data is semi-automated with the use of specialized Metamorph macros/journals. This multiwavelength fluorescence workstation is also equipped with appropriate filters and software (Metafluor) to work with fluorescent probes such as fura-2 for monitoring intracellular calcium concentrations. (Microscopy Service Request Form)

4.  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 $45/hour.(Microscopy Service Request Form)

5.  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.


The protein identification component of the PIL Core is adjacent to the National Resource for Proteomics and Pathways (NRPP) and shares some resources and personnel including the Director, Dr. Andrews.  The NRPP UM laboratory occupies approximately 800 square feet (rooms 1188 and 1195 of North Ingalls Building) of wet laboratory space and includes an ultracentrifuge, high-speed preparative centrifuge, IEF and 2D Gel equipment, gel scanner, cell culture equipment, HPLC, PCR, apparatus for organic synthesis, and other typical laboratory equipment.

The protein localization component of the PIL Core is located in the laboratories of Drs Williams and Ernst while the confocal and electron microscopes are located in the MIAC core of the MDRTC and in the MIL. Dr Williams’ lab is located on the 7th floor of Medical Sciences II. It contains general purpose equipment, a tissue culture room, a Nikon inverted fluorescence microscope with a digital camera and a computer workstation running Metamorph software for image analysis. Dr Ernst’s laboratory of about 200sq ft is located on the 3rd floor of the Biomedical Sciences Research Building (BSRB) and is devoted to the support of this core. It houses general purpose equipment plus a new Leica cryostat and a Reichert Ultracut E ultramicrotome with associated glass knifemaker and diamond knives. Bradley Nelson, the Research Assistant directed by Dr. Ernst who carries out fixation, embedding and sectioning, is located in this laboratory. The MIL which contains several confocal microscopes and the EM is located in the basement of the BSRB. The MIAC core of the MDRTC which contains confocal and widefield fluorescence microscopes is located on the 2nd floor of the BSRB but in early 2010 will move to the newly constructed Brehm Center for diabetes research located about 5 blocks away from MSII and the BSRB. Most GI Center members are located in the Medical Sciences Complex and the BSRB although some are in the Life Sciences Institute or will be in the new Brehm Center.




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