CORE B: IMAGING


Director, Bin Wu, PhD
bwu20@jhmi.edu
410-502-4201

 

 


Manager, George McNamara, PhD
gmcnamara@jhmi.edu
410-614-0134 (office)

 

 

Fatemeh Jahan Bakhsh, PhD
Technician, Fatemeh Jahan Bakhsh, PhD
Johns Hopkins University School of Medicine
Biophysics and Biophysics and Biochemistry Department
fjahanb1@jhu.edu
(410) 614-3959 (office)

GOALS

Ross Fluorescence Imaging Center core is dedicated to providing state-of-art light microscopy technology to the members of the Hopkins Basic Research Digestive Disease Development Center and to the general Hopkins scientific community.

SERVICES OFFERED

Consultation

The Core Director and Manager meet with Investigators prior to starting a new project to discuss the project goals, the specific equipment to be used for best acquisition of the data, sample preparation procedures, and image analysis. These consultations provide an opportunity to avoid costly mistakes in sample preparation, particularly if experimenting with live tissue/cells, or rare patient-derived samples, and to increase the chances for successful image acquisition from the first experiment on. After collecting the images, the Core Manager and Core Director meet with the Investigator (if necessary) to assist in data interpretation and to instruct their post-doctoral fellows or graduate students in using software for data analysis.

State-of-the-Art Imaging

See Equipment below for a comprehensive list of Imaging Core instrumentation.

Protocols

We gather or develop common protocols (ie, SOPs) for sample preparation, imaging routines, and analysis programs, and ensure that the instruments have designated configuration and stepwise routines to perform the corresponding protocols and analysis. For example, the smFISH protocol includes procedures for coverslip cleaning and coating, tissue preparation and cutting, hybridization with probes, mounting with antifade reagent, imaging on FISHscope, and finally quantification with Matlab code. The research specialist Dr. Bakhsh will assist and train new users in each of these aspects of the smFISH studies and will prepare the probes as well as supervise the experiments until the DDRCC Member or their laboratory is able to perform the studies independently.

Assistance

One of the major tasks of the Core Manager is to ensure that the DDRCC Members are adequately trained to perform the studies and then to assist users when they have difficulties, either in instrumentation setup or data analysis. The Core Manager has extensive experience in microscopy, imaging, and user training, and handles problems promptly and efficiently. Feedback from Imaging Core users indicates that availability of our imaging expert when needed is key to obtaining high-quality and reproducible data.

Formal Training

Investigators must demonstrate that they can operate the designated equipment and conduct experiments independently before making reservations. This critical training program allows us to provide 24-7 availability to the Imaging Core based on electronic ID access.

Workshops

The Imaging Core organizes demonstrations of state-of-the-art technology and instrumentation that is emerging. JHU DDRCC investigators can use these opportunities to evaluate the technologies and generate preliminary data. If the trial use indicates that there is a clear need or demand for an instrument, this initiates the process by which the Core considers obtaining the equipment.

IMAGING CORE EQUIPMENT

A. Olympus FV3000RS laser scanning confocal fluorescence microscope. This replaced our Zeiss 510 LSM that was no longer serviced by Zeiss. Our FV3000RS has seven laser lines (405, 445, 488, 561, 640, and 730 nm), four internal GaAsP PMTs, each with volume phase holographic transmission grating and monochromator-like slits for wavelength range selection, and two external GaAs PMTs for near-infrared fluorescence (including Li-Cor IRDyes). The objective lenses include 2x, 10x/0.4NA, 20x/0.75NA dry, and 30x/1.05NA, 40x/1.25, and 100X/1.35 NA silicone oil objective lenses (this oil has refractive index 1.405, closer to that of live cells; cell culture media additives are available to match R.I. to that of the oil, effectively making cells invisible; we note that standard fixed cells and tissue immunofluorescence and FISH also works well with these lenses).

B. Leica SP8 laser scanning confocal fluorescence microscope. The microscope is owned by the Department of Anesthesiology and Critical Care Medicine (ACCM) and managed and housed by the Core which makes it available 50% of the time for Core Center Investigator use.  We charge a management fee, which is added to our charge-back funds. This Windows 10 64-bit PC typically acquires 1+ gigabytes data per sessions, but occasionally 10+ gigabytes.

C. Keyence BZ-X710 all-in-one microscope. It is an automated inverted fluorescence microscope with multiple imaging modalities, such as bright field, phase contrast, oblique illumination. The microscope has excellent environmental control with CO2, humidity and media perfusion to keep the sample for long-term imaging. An important use is for tiling images for the presentation of larger fields of analysis. Another major use of the Keyence is overnight and multi-day time-lapse imaging of organoids.  We have installed a local data drive and asked users to save locally, upload it at end of their sessions.

D. Olympus FV1000MPE Multiphoton microscope is equipped with a fully motorized Prior ProScan III stage and Warner temperature-controlled perfusion chamber (apical and basolateral sides) for multiphoton imaging of live tissue and stem cell-derived enteroid/colonoid samples. The FV1000 is based on an upright BX61 WI Olympus microscope, which is connected to a Spectra-Physics DeepSee automatic dispersion compensated, mode-locking Ti:Sapphire laser with a spectral range of 690-1040nm. The system has 5 detectors (4 “epi” NDD’s for fluorescence and 1 transmitted light). Images can be obtained simultaneously with any number of detector configurations. The system computer was rebuilt in 2012 with dual i7 Pentium CPU’s and 24 GB of DDR3 RAM, a solid state drive for the Windows 7 64 bit operating system (to be updated to Windows 10 before end of 2019), and second dedicated 1.5 TB RAID data drive. The microscope utilizes two high-resolution objectives (XLPlan N 25x 1.05 W and a LUMFL N 60X NA1.2 W immersion) both for the imaging of living tissue and cells in perfusion media. The system has been optimized for deep tissue imaging (up to 1 mm) with simultaneous 3-color fluorescence and transmitted light data acquisition.

E. Zeiss AxioObserver.A1 epifluorescence microscope upgraded with a new Olympus DP80 dual monochrome + RGB CCD camera (replaced DP72 RGB only camera). The addition of the monochrome CCD capability now makes this microscope useful for Cy5 and similar near-infrared fluorophores enabling any, some, or all of simple DAPI/green/red/NIR fluorescence imaging. RGB CCD sensor still enables histology imaging (H&E histology, H&DAB immunohistochemistry). Either CCD can be used for phase-contrast imaging – users are shown how to use each CCD during training. We installed a new PC in late 2018, and have Windows 10 (64-bit) and current Cellsens (part of DP80 camera purchase).

F. “FISHscope”: Olympus IX83 wide-field automated fluorescence inverted microscope. This IX83 is equipped with motorized Z (internal) and motorized SSU “gliding” stage, Spectra III-360 light engine from Lumencor with 8 bright light sources (single-source control through CellSens), currently optimized for DAPI, Alexa Fluor 488, “TRITC” (more often Cy3), Cy5, Cy7, using a Semrock “Penta” filter cube (complete excitation, dichroic, and emission filters), Semrock single band emission filters in Sutter 10-3 emission wheel in IX83 lower deck (upper deck is the Penta cube turret), Hamamatsu ORCA-FLSH4.0LT sCMOS camera, Olympus Cellsens GPU deconvolution with fast Nvidia GPU card in a Windows 10 64-bit PC with new fast SSD and HDD arrays, HD 4K monitor. This microscope will be used primarily for single molecule RNA FISH (smFISH), immunofluorescence in fixed cell lines in culture and in fixed tissue sections, including human enteroid/organoid monolayers. Stage tiling of slides and SBS plates can be achieved. Emphasis is on high-resolution smFISH.

G. Live Cell Laser Microdissection (newly upgraded and offered as a service): Zeiss PALM MicroBeam. For Core B, this is a new service in development that includes live cell capture capability. The equipment is a Zeiss PALM Micro Beam that is a laser microdissection set up with a pressure catapulting technology donated by Dr. Lutsenko and renovated by Core Center Administrative Core funds. This is able to isolate live cells for culturing as well as conventional laser capture microdissection.

H. Zeiss LSM880 AiryScan Super-Resolution microscope We have arranged with the JHU Institute for Cellular Engineering (ICE) for access for our Core Investigators to the Zeiss AiryScan microscope (located in the  Miller Bldg which is adjacent to the  Ross Bldg) on a fee-for-service basis with backup, use of the JHU Imaging Facility Zeiss 880 AiryScan .  AiryScan has a super-resolution mode (fiber-optic array positioned as 0.2 Airy unit pinholes, 20% better spatial resolution than standard 1.0 Airy unit pinhole, which is 20% better resolution than widefield).

I. Chambers for Polarized Cells (Warner Instrument Corp.). The chambers used for perfusion in the multiphoton and confocal microscopes have a closed bath design which provides a large viewing area and good access for microscope optics. The bath volume is 260 μl, and the distance between the top and bottom coverslips is 2.5 mm. The chamber is mounted into a heater platform (Warner Instrument Corp.), which allows control of the temperature in the perfusion chamber and also provides a clamp to seal the chamber, the filter with the cells/tissue and coverslips. The samples in a chamber can be separately perfused from the apical, or apical and basolateral surfaces for studies of epithelial cells grown as monolayers on permeable supports or for studies of tissue physiology based on different luminal or serosal stimuli. These chambers are useful for enteroid cultures, intact animal intestinal tissue, or other tissues.

J. Core managed non-microscope equipment, including Li-Cor Odyssey CLx, PTI-QuantaMaster monochromator (2 units), MetaMorph (3 licenses) analysis software licenses.

  1. The Li-Cor Odyssey Western blot scanner is mostly self-serve. The Odyssey is equipped with two solid-state diode lasers, which simultaneously provide excitation at 680 and 780 nm of IRDye labeling reagents. Odyssey software is used for scan control and data analysis. The system provides accurate and fast quantification of immunoblots over a time range in which intensity is related to the fluorescence signal. The expression of two proteins can be quantified simultaneously on the same blot, a procedure that significantly improves the accuracy with which changes in a specific protein’s expression vs. that of a loading control can be measured. This is particularly relevant when changes in protein post-translational modifications are being measured (e.g., phosphorylation, glycosylation). The imaging system is used extensively by Center Investigators for quantitative Western blot analysis. It is multifunctional and can also be used in quantitative protein array analyses, gel-shift assays, in-cell Western and even tissue fluorescence. Secondary fluorescent antibodies are either purchased by individual investigators (non-Core function) or for a set fee from the Core.
  2. PTI-QuantaMaster monochromator (2 units) provide fast and accurate measurements of changes in pH and calcium by using either ratio-excitation or ratio-emission fluorescent dyes in living cells growing as a monolayer or in suspension. Because each fluorometer is equipped with a temperature-controlled perfusion chamber for epithelial cells growing as a confluent monolayer on permeable support or coverslip, changes in ion concentrations or application of second messengers can be made rapidly and separately to either apical or basolateral compartments. The data are collected and stored on a Windows-based PC, with 2.5 GHz Pentium Dual-Core processor, 2GB RAM, 465GB hard drive and 20 inch, flat-panel monitor, which allows instantaneous comparison of data obtained during the experiment. Detailed data analysis is performed after the experiment on a separate workstation.
  3. MetaMorph (3 licenses) analysis software licenses. Two USB license dongles move among several PC’s in image core and Donowitz lab and other Ross 9th floor labs. Dr. McNamara has a personal license, on his new Windows 10 64-bit workstation.

NEW CORE INSTRUMENTS

  • Olympus FV3000RS laser scanning confocal microscope
  • Leica SP8 laser scanning confocal microscope
  • Olympus IX83 wide-field fluorescence microscope (FISHScope)
  • Keyence BZ-X710 all-in-one microscope
  • Zeiss PALM MicroBeam Laser Microdissection microscope
  • Zeiss LSM880 AiryScan Super-Resolution microscope
  • BioTek absorbance plate reader

FEES FOR IMAGING CORE INSTRUMENTATION

InstrumentHourly rate for trained users
Olympus FV3000RS Confocal microscope$27 (50% reduction for Full Members, 75% reduction for Associate Members)
Leica SP8 Confocal Microscope$27
Olympus FV1000MPE Multiphoton Microscope$50 (*cost-offset as above under FV3000RS)
Keyence BZ-X710 all-in-one microscope$10
Zeiss AxioImager.A1 inverted microscope w/DP80 CCD$10
Olympus IX83 inverted microscope (“FISHscope”)$10
PTI fluorimeters, BioTek absorbance plate reader$4
Li-Cor Odyssey CLx western blot fluorescence scanner$4
MetaMorph Imaging System$4
Zeiss LSM880 AiryScan Super-Resolution microscope (ICE, Miller Bldg)$40 (*cost –offset as above under FV3000RS)
Training on confocal and multiphoton microscopes$250 per new user (2 x 2 hours each)
Training on epifluorescence and bright field imaging$50 per new user (1 hour)
Laser Microdissection and Pressure Catapulting (LMPC) technology via the Zeiss PALM Micro BeaTo be determined (new service 4/2021)
smFISH services probe services (probe generation, specimen preparation, imaging, user training) conducted by Research Specialist , Fatemeh Jahan Bakhsh, PhDTo be determined (new service 1/2021) $200 per 5 nmole probe set of 50 fluorescently labeled oligoes. $100 per set for additional colors.

WEBSITE

For additional information about our services and equipment, please go to our website confocal.jhu.edu.

NEW SERVICES IN DEVELOPMENT

Single-molecule RNA FISH microscope

Dr. Wu’s lab has been developing smFISH technology and has shown that it can be performed on mouse and human small intestine and human liver. Up to 5 probes can be done simultaneously, with a goal of >20 probes with quantitation (combinatorial labeling and/or sequential processing with motorized stage relocation/ tiling). The IX83 microscope, received via an NIDDK Supplement, is dedicated to this application. An example image from the new FISHscope, using three Wu lab-synthesized probe sets, is shown in Fig. 1. This technique was initially standardized in the Wu laboratory but has now been ramped up and the service is being offered to the Research Base as of January 2021. This was the new technique most likely to be used by multiple DDRCC Investigators based on our latest Member survey.

Core B Figure 1
Figure 1. 3-color smFISH images of human ileum tissue using in-house synthesized oligonucleotide probes. Green: PDGFRA, Purple: F3; Red: SMA4;. (B) is enlarged from box in (A) (Wu, Donowitz labs, unpublished).

Super-Resolution Microscopy

The Zeiss 880 AiryScan confocal microscope has been available for JHU DDRCC Investigators on a fee-for-service basis starting in funding Year 9. The equipment is part of the JHU Institute for Cellular Engineering (ICE), and JHU DDRCC Investigators are one of the JHU groups that have been allowed to sign up for use. We are encouraging usage by the JHU DDRCC Research Base by devoting several Work-in-Progress visiting professor slots to investigators who have used super-resolution microscopy to answer physiologically relevant questions. In addition, we have invited the JHU ICE microscope manager, Mr. Stewart Neifert, who supervises and teaches use of the Zeiss 880, to present the advantages and potential applications of this instrumentation. We elected to work with ICE after imaging evaluations of the Zeiss 880 with Airyscan by Imaging Core Associate Director Brian O’Rourke with cardiomyocytes; and JHU DDRCC Associate Member Helen Yu, who imaged human enteroid monolayers interacting with macrophages (Fig 2).

Core B Figure 2
Figure 2. Co-Culture shows interactions of human enteroids, ETEC, and co-cultured macrophages (MΦ). Left, upper: Schematic representation of enteroid-macrophage co-cultures established with non-differentiated (ND) enteroid monolayers. Right, upper: confocal Z-stack X-Z images of monolayers of ND enteroids with MΦ extending projections through the membrane to reach the apical surface of enteroids infected 3 h earlier with ETEC (arrowheads). Left, lower: high resolution, AiryScan image showing MΦ projections (arrowheads). Right, lower: cartoon of projections. Actin, white; CD14 (MΦ), red; nuclei, blue; filter, dashed lines. Note MΦ projections contacting ETEC. Scale bar 20 μm.

Laser Microdissection Microscope

Laser Microdissection and Pressure Catapulting (LMPC) technology via the Zeiss PALM Micro Beam is available to JHU DDRCC Members as of January 2021. There has been increasing interest and demand for isolation and analysis of individual cells or groups of cells from tissue sections, including isolation of living cells that can be cultured. One example of the need from JHU DDRCC Investigators is illustrated in Fig. 3. The Lutsenko laboratory recently discovered rare cells present in human and mouse duodenum that express very high levels of the copper transporter ATP7B (Wilson disease protein). Standard methodologies (co-localization with known cell-type markers) failed to identify the cell types (no co-localization was found with known epithelial cell types), indicating that the ATP7B-rich cells could be a novel subtype of epithelial cell.

Core B Figure 3
Figure 3. ATP7B (in green) is highly expressed in the intestine (mouse intestine shown), especially in unique epithelial cells of the duodenal villus (arrow). Dimensions 400×500 μm.

The LMPC methodology combines laser microdissection of either tissue or cultured cells grown on specific plastic membranes with a laser-assisted transfer of the cut-out material for further cell propagation or analysis (Fig. 4). Unlike the original laser dissection methodology that largely destroys cells and tissue, LMPC preserves the integrity of tissue and allows isolation of living cells. This property enables not only genomic analysis and gene expression profiling, but also proteomic and metabolite assays (10, 11), as well as cell selection for culture. Members of the JHU DDRCC are beginning to use the PALM Micro Beam (Carl Zeiss Microscopy) under supervision by the Lutsenko laboratory.

Core B Figure 4
Figure 4. Intestinal tissue mounted on a polyethylene naphthalate (PEN) membrane slide is visualized with 63x objective under transmission light (A,B) and fluorescence (C,D) before (A,C) and after (B,D) Zeiss PALM Micro Beam processing (Lutsenko lab). The PEN membrane is highly UV-A absorbing, facilitating cutting, and provides support to the cell(s)/tissue during microdissection. Dimensions: 120×80 μm.