Grant Language: Boilerplates
The VCU Flow Cytometry Core Laboratory (updated July 2015)
"The VCU Flow Cytometry Core Laboratory was established in 1976, and has been constantly modernized since that time. It currently occupies over 1500 sq. ft. in the Molecular Medicine Research Building and the Kontos Medical Science Building, and provides a comprehensive instrumentation suite and technical service in support of both cell sorting and analysis on a fee-for-service basis. The core currently features six primary instruments. Cell sorting is conducted using two multi-laser BD Aria II sorters. One of the Aria sorters is enclosed in a Baker BioProtect III Hood, enabling the sorting of human and biohazardous tissues. The two Arias have UV and violet laser capability and up to 4 simultaneous sorts per Aria. In addition the core has a 4 laser Amnis ImageStream Mark II analyzer which acquires images of cell populations for analysis. Flow analyzers include a two laser BD FACSCanto II and as of June 2015, a 5 laser BD LSRFortessa X-20. The two analyzers are capable of 9 or 18 color simultaneous analyses, respectively and the 5 laser makes compensation with multiple colors easier. In addition to flow instrumentation, the core also maintains a Biacore T200 surface plasmon resonance (SPR) instrument for determination of protein interaction constants. The director, Daniel H. Conrad, has over 30 years of experience in flow cytometry. The laboratory manager and head technician, Julie Farnsworth, is highly trained in flow cytometry and has been with the core for more than 15 years. The other support technician, Qingzhao Zhang has more than 7 years of flow cytometry experience and assists with both flow analysis and sorting. The resource provides a wide range of services, including instrument training, routine fluorescence analysis, and development of innovative custom analysis and sorting protocols. Walk-up instrument time is also available to trained users. This core maintains a stable base of instrumentation, expertise, and trained personnel and currently provides services and training for multiple users in over 60 research groups from 21 departments at VCU. The Flow Cytometry Core Laboratory is supported, in part, by funding from the NIH-NCI Cancer Center Support Grant (P30 CA016059)"
The VCU Chemical and Proteomic Mass Spectrometry Core Facility (updated July 2015)
The Chemical and Proteomic Mass Spectrometry Core Facility offers a wide variety of services ranging from routine mass spectrometry analysis to interactive custom design of innovative analyses, addressing the specific needs of individual investigators. The resource director closely follows the analyses performed by the facility, insuring quality control and advising facility users on alternative, often novel, approaches to experimental design and data analysis. The shared resource maintains a stable base of instrumentation. Routine proteomics experiments are performed on a 5800 MALDI TOF-TOF instrument from AB Sciex, which utilizes off-line separations performed on an Easy nLC HPLC system from Thermo Electron and a spotter robot from Leap Technologies. The MALDI instrument can handle basic proteomics experiments such as analysis of the proteins in gel bands, as well as basic mass measurement. It has high mass accuracy and the ability to measure samples multiple times to increase the number of peptides identified in each experiment. More complex proteomics experiments are handled by an Orbitrap Velos mass spectrometer from Thermo Electron Corporation, which is interfaced to a nanoAcuity UPLC from Waters Corporation. Newly added in Q2 2015 is a state-of-the-art Waters Synapt G2Si mass spectrometer with electron mobility capability and M-Class UPLC front end. This instrument is ideal for the analysis of unlabelled clinical samples and highly complex proteomics samples. Finally, the Mass Spectrometry Facility provides data analysis, including database searching of the peptides detected by mass spectrometry, identification of proteins present in the samples, and mapping of the detected peptides within the protein. Verification of post-translational modifications is also provided. Data are shared with investigators using Scaffold software, which has been developed with a user interface allowing investigators to see where peptides are located within the protein, examine a list of proteins detected within the sample, look at fragmentation spectra and scoring values for each peptide, and even compare proteins and peptides across a number of samples.
VCU Transgenic Mouse Core
“The VCU Transgenic Mouse Core laboratory, established in 2000, provides a comprehensive suite of services for the creation of genetically modified mice. The facility, located on the 8th floor of the Molecular Medicine Research Building occupies ~1000 sq ft of space, including a 600 sq ft laboratory, a 150 sq ft tissue culture room, and a 250 sq ft animal holding room. The facility is well-equipped (primary instrumentation includes a Olympus IX-71 inverted injection scope with DP-72 digital camera and stage cooling, Narishige micromanipulators, and a Celtram air microinjector), and offers the production of both transgenic and knock-out or knock-in mice. All stages of transgenic mouse production are offered, including consulting, creation of targeting vectors, ES cell electroporation, screening of ES cell clones, blastocyst injection, and breeding of chimeras. In addition, the core laboratory also has extensive expertise in the mouse line re-derivation by embryo transfer, embryo and sperm cryopreservation, tail DNA preparation and genotyping. The core also maintains an IVIS spectrum imager for live animal imaging. The core is directed by Dr. Jolene Windle, who has been actively involved in developing genetically engineered mouse models since 1986. She is supported by Resource Manager Greg Campbell, who has worked with transgenic animals since 1996, Laboratory Specialist Jillian Stifflinger, and Research Associate Dr. Mark Subler. The Transgenic Mouse core laboratory is supported, in part, by funding from the NIH-NCI Cancer Center Support Grant (P30 CA016059)”
VCU Structural Biology Core Laboratory (updated February 2018)
"The VCU Structural Biology Core Laboratory is a VCU institutionally supported resource that is directed by Jan Chlebowski, PhD. The core facilitates access to a comprehensive suite of instrumentation and computational resources in support of macromolecular structure determination. There are 3 main components to the core: X-ray crystallography, nuclear magnetic resonance (NMR), and molecular modeling. The X-ray crystallography component is managed by Martin Safo, PhD, and Faik Musayev, PhD. Its resources include an X-ray diffractometer installed in Q1 2018 comprising a MicroMax-007HF generator, VariMax-HF Arc Optics, Hybrid Photon Counter, Eiger R 4M Detector, AFC11 Goniometer and Oxford Cobra Cryo-system. The core has a crystallization robotics suite consisting of a Gryphon drop setter, a Minstrel/Gallery imaging and incubation system for crystallization trials, and an Alchemist liquid handling system. The NMR component is managed by J. Neel Scarsdale, PhD. The component operates a Bruker Avance III 700 MHz instrument suitable for 1D, 2D, 3D, or 4D homo- or hetero-nuclear experiments. The instrument features 3 RF channels with pulse field gradients, permitting the acquisition of data for triple or pseudoquadruple resonance experiments. The molecular modeling component is managed by Glen Kellogg, PhD. Molecular modeling is supported by a suite of 8-core ApplePro and HP linux multicore graphics workstations, supplemented with a Linux-cluster back-end with over 3,000 computer cores provided by the VCU CHiPC. Software supported by the molecular modeling includes the commercial Tripos Sybyl suite and a number of other packages including CCP4, GOLD, Dock, AutoDock, HINT, NAMD, and Hex. In addition to instrumentation, the core provides training and consultation, both through formal classes and one-on-one sessions with individual investigators.”
VCU Center for High Performance Computing (updated October 2017)
“The VCU Center for High Performance Computing (CHiPC) is located in approximately 2000 sq ft of total space, predominantly on the third floor of Harris Hall on the Monroe Park Campus. The mission of the CHiPC is to provide high performance computing services for the VCU research community. To accomplish this goal, the CHiPC maintains four major supercomputing clusters, each specialized for different computing environments. They may be summarized as follows (descriptions current as of September 2017): 1) teal.vcu.edu is the primary cluster intended for large scale parallel computing, and is especially well suited for applications such molecular dynamics simulations, quantum chemistry and other Physical Sciences jobs. Teal consists of ~4250 64 bit AMD compute cores, each with 2-4 GB RAM/core, 9.5 TB of total RAM, 36 TB of /home space, and tmp space of between 360 and 787 GB per node. High speed network infrastructure is provided by a 20 Gb/second infiniband architecture; 2) bach.vcu.edu is the cluster designated for serial and small parallel applications. Bach consists of a total of 1048 AMD 64 bit cores, each with a minimum of 2 GB/core RAM, 2 TB total RAM, 12 TB of /home space, and /tmp space of 360 GB per node. Networking infrastructure is gigabit ethernet; 3) godel.vcu.edu is a cluster optimized for bioinformatics applications, with 1432 AMD 64 bit and Intel 64 bit cores, each with at least 3 GB RAM/core, 4.5TB of total RAM, 17 TB of /home space, tmp space of at least 180 GB/node, and 40 Gb/second infiniband networking; 4) fenn.vcu.edu is a cluster designed to support research using data that must comply with federal security and privacy requirements, with 1016 Intel 64 bit cores, 2/GB of RAM/core, 420TB of GPFS storage (expandable to 2.2PB)m 54 Gb/second Infiniband networking. These clusters are collectively served by over 1.3 PB of networked nfs and GPFS high speed storage. To support this infrastructure, theCHiPC employees 4.5 FTE positions, (J. Mike Davis, Technical Director; John Noble & Carlisle Childress, Systems Analysts; John Layne, Applications Analyst; and Neel Scarsdale, Ph.D., Assistant professor and Physical Sciences Applications specialist. In addition to maintaining the hardware, the CHiPC works collaboratively with the user base to maintain and optimize a large number of applications and development tools (BLAST, R, MATLAB, NAMD, Gaussian, Gromacs, Charm, C/C++, Fortran compilers, etc.)”
VCU Tissue and Data Analysis and Acquisition Core Laboratory (updated July 2015)
“The VCU Tissue and Data Analysis and Acquisition Core (TDAAC; Director Michael Idowu, M.D.), established in 2002 and located in the 6,400 sq ft VCU Molecular Diagnostics CLIA-certified laboratory, has as its goal the acquisition of human specimens and associated clinical and pathological findings to support translational research. This is done both through the aegis of the VCU-IRB approved “Tissue Acquisition System to Support Cancer Research” (TASSCR) protocol, which supplies specimens to a biorepository supporting cancer research through acquisition of residual tumor and normal tissue samples along with informed consent from patients with cancer. In addition, TDAAC collects tissue, hematopoietic and other researcher-specific samples which support investigator-initiated IRB approved research projects. The specimen acquisition process ensures that the primary purpose of the specimen for patient care is maintained and the quality of the specimen is optimal for biomedical research. These services are achieved through leveraging a network of interdepartmental and informatics relationships within the VCU Health Systems. TDAAC staff in collaboration with Department of Pathology based Molecular Morphology Genomics Laboratory provide samples of extracted, quality controlled RNA and DNA from human tissues, frozen sections, and cryopreserved samples of viable hematopoietic neoplasias. Because TDAAC is an outgrowth of a multicenter grant that focused on gene expression microarray studies on multiple cancer phenotypes, some samples have associated gene expression data as a part of their annotation. The VCU Molecular Diagnostics laboratory is a state-of-the art CLIA-certified laboratory and performs clinical as well as research molecular testing with state-of-the-art equipment, including two bioanalyzer (Agilent 2100), six robotic systems for automated nucleic acid extraction (two Qiagen EZ1 Advanced XL, two Qiagen EZ1, and two Life Technologies MagMAX Express instruments), automated DNA sequencers (two ABI 3130), six real-time PCR instrumentation systems (four ABI 7500, and two Roche LightCycler™ Systems); a Leica CM1850 Cryostat, four Thermo Scientific Revco PLUS Ultra-Low Temperature Freezers, three Thermo Scientific Locator 4 PLUS Cryobiological Storage Vessels, three NanoDrop ND 8000 Spectrophotometer, ; a complete Affymetrix GeneChip® workstation for DNA microarray technology; as well as Next-Generation Sequencing instruments (two Ion Torrent PGMs, and two Ion Torrent Protons). All temperature-sensitive equipment is constantly monitored by an electronic system (CheckPoint: 24/7 monitoring system) and all ultra-low freezers have a built-in LN2 backup system that automatically releases LN2 into the freezer in case of temperature failure. The VCU TDAAC is supported, in part, by funding from the NIH-NCI Cancer Center Support Grant (P30 CA016059)”
VCU Mid-Atlantic Twin Registry (updated July 2015)
VCU Microscopy Core Laboratory (updated July 2015)
“The VCU Microscopy Core Laboratory (Director Scott Henderson, Ph.D.), housed within the Department of Anatomy and Neurobiology, is located in a 3000 sq ft facility spread over several rooms on the 9th floor of Sanger Hall. It is a fee-for-service core that provides the instrumentation and expertise to facilitate a comprehensive spectrum of imaging methods and techniques. Instrumentation and services include: 1) Electron microscopy (TEM, SEM) is supported with a Jeol JEM-1230 TEM equipped with Gatan UltraScan 4000SP & Orius SC1000 CCD cameras, and a Zeiss EVO 50 XVP SEM equipped with SE, VPSE & BSD detectors, extended variable pressure (up to 750 Pa), Deben coolstage, a water vapor introduction kit, and a Shuttle & Find stage adapter for correlative confocal / SEM imaging; 2) Confocal laser scanning microscopy, supported by three systems: A Zeiss LSM 710 (inverted) equipped with a 32-channel array detector plus 2 side PMT detectors, a transmitted light detector, a Becker & Hickl Fluorescence Lifetime Imaging system with 2 hybrid GaAsP detectors (for FLIM / FCS), a motorized XY stage, a PeCon low profile stage incubator, Shuttle & Find stage adapter, and 5 lasers (405 nm, multi-line Argon [458/488/514 nm], 561 nm green diode, 633 nm HeNe & a pulsed 440 nm. The system is set up for FRET, FRAP and FLIM analysis. The second system is a Zeiss LSM 700 (upright) equipped with 4 solid state lasers (405 nm, 488 nm, 555 nm, and 635 nm). The scan head has 2 confocal PMTs, a variable secondary dichroic beamsplitter and a transmitted light detector. The third system is a Leica TCS-SP2 AOBS (inverted) with a spectrophotometer scan head, a motorized XY stage and three confocal detectors (PMTs) (plus a transmitted light detector). This system has five lasers: blue diode (405 nm), Argon (458, 476, 488, 514 nm), green HeNe (543 nm), orange HeNe (594 nm) and red HeNe (633 nm); 3) Multi-photon laser scanning microscopy, supported by a Zeiss LSM 510 META NLO (fixed stage upright) equipped with a Spectra-Physics Broadband (710-990 nm) MaiTai Ti:sapphire laser and 3 visible lasers (multi-line Argon, 561 DPSS, HeNe 633), 2 non-descanned detectors, 3 descanned detectors (including a META detector for spectral scanning) a transmitted light detector, and a variety of objective lenses (dry, oil, multi-immersion, water immersion). A Luigs & Neumann 380 FM workstation (with slice chamber & in vivo bridges) and physiology equipment (stimulators, VP timing unit, A/D converters, multiclamp amplifier, line heaters, peristaltic pump, pipette puller) are linked with the system to facilitate simultaneous imaging & physiological recording; 4) Live cell confocal microscopy supported by a Zeiss Cell Observer SD spinning disc confocal microscope, equipped with a Yokogawa CSU-X1A spinning disc, 2 Photometrics Evolve 512 cameras, an Axiocam MRm camera, a high resolution piezo driven Z stage, 4 lasers (405 nm, 458/488/514 nm, 561 nm, 635 nm), DIC, phase contrast, FRAP/uncaging and a stage incubation system (to support live cell / live tissue imaging, with regulation for temperature, humidity, CO2 and O2); 5) Total internal reflection (TIRF) microscopy, supported by an Olympus cellTIRF system equipped with 3 lasers (405 nm diode, Argon [458, 476, 488, 514 nm] and 561 DPSS), AOTF, DIC, phase contrast, Andor iXon DU-897E emCCD camera, a laser-based automated focus drift compensation system and a stage incubator (temperature, humidity, CO2); 6) A Nikon N-SIM Structured Illumination Microscope (“Super-Resolution”). This system is equipped with 3 diffraction gratings, 4 lasers (405, 488, 561 & 640 nm), Andor iXon DU-897E emCCD camera, a laser-based automated focus drift compensation system, motorized XY stage, a high resolution piezo-driven Z stage, motorized TIRF optics (with independent laser input), fluorescence optics, DIC, and a Tokai Hit stage incubator (to facilitate live cell imaging); 7) seven wide-field fluorescence microscopes, including: an Olympus BX51 configured for computer-assisted stereology and equipped with a Prior Proscan XYZ motorized stage, a Heidenhain microcator, an Olympus DP72 CCD camera, fluorescence optics (DAPI / GFP / DsRed / Cy5), DIC, phase contrast, Visiopharm newCAST software and Olympus cellSens software. The second system is a Zeiss AxioImager Z2 automated microscope configured for morphometric analysis and equipped with a Q-Imaging CCD camera and a Zeiss Axiocam MRm, fluorescence optics, (DAPI / CFP / GFP / DsRed / Cy5), DIC, motorized XY stage, microcator, and Microbrightfield Neurolucida & StereoInvestigator imaging / image analysis software. The third system is a Zeiss Axiovert 200 equipped with a Hamamastu ORCA ER CCD camera, Colibri LED illumination unit (blue, green, red), fluorescence filter sets (DAPI, GFP, DsRed), white light LED and a stage heater. The fourth system is a Zeiss Axioimager A1 equipped with fluorescence optics (DAPI, GFP, DsRed), and an Axiocam MRc CCD camera. The fifth system is a Zeiss AxioObserver A1 equipped with fluorescence optics (DAPI, GFP, DsRed), and an Axiocam MRc5 CCD camera. The sixth system is a Nikon ECLIPSE E800M with fluorescence optics (DAPI, GFP, DsRed, Cy5), and a Diagnostic Instruments Spot RT CCD camera. The seventh system is a Zeiss Axiovert 40 equipped with fluorescence optics (DAPI, GFP, DsRed), and an Axiocam MRc CCD camera. A Zeiss Discovery V20 Stereo zoom microscope with transmitted, reflection and fluorescence illumination, 0.63x, 1.25x and 1.5x lenses, filter sets for GFP, CFP, YFP and DsRed and a Axiocam MRc CCD camera is available for micro/macroscopic imaging; This equipment suite (supported by sputter coaters, ultramicrotomes, critical point dryers, etc.) allows the core to offer many types of imaging methods and services including: Sample preparation; Live-cell imaging; Immuno-localization (fluorescence, EM); Fluorescence recovery after photobleaching (FRAP); Fluorescence resonance energy transfer (FRET). The core also provides multi-dimensional image analysis. A cell culture facility (with incubators and biological safety cabinet) is located on-site to support live cell imaging. Training is available for all of the imaging modalities and methods offered.”
VCU Nanomaterials Characterization Core Facility (updated July 2015)
The VCU Nanomaterials Characterization Core Facility (Director Everett Carpenter, Ph.D.), The nanomaterials core (founded in 2010) provides a comprehensive instrumentation suite for VCU researchers interested in nanomaterials, or surface characterization. The facility features a ThermoFisher ESCAlab 250 X-ray photoelectron spectrometer. The ESCALab is multitechnique platform for studying the surface of materials with XPS, Auger, UPS, and ISS techniques. The sample stage is an automated 5-axis stage for angle resolved XPS with heating and cooling from 77K to 600K. The system also has a High Pressure Gas Cell for introduction of reactive gases for absorption studies. The facility also has three scanning electron microscopes and a transmission electron microscope: a Hitachi SU-70 FE-SEM, a JEOL JSM-5610 LV, Zeiss Auriga SEM/FIB, and Zeiss Libra 120 TEM. The Hitachi SEM is a field emission unit which allows for a 1nm spatial resolution. The unit also is equipped with a STEM options, a Nabity Lithography System, dedicated backscatter detector, and Genesis EDAX system with low element windows for detection of Be to Pu. The JEOL SEM has a High Vacuum option as well as a Low Vacuum option to run non-conductive samples without metal sputtering. The Libra is Zeiss workhorse microscope which features point to point resolution of 0.34 nm. Included in the scope is an in-column Omega filter which allows for energy electron lose spectroscopy (EELS) with an energy resolution of <1.5 eV. The Zeiss Auriga is a dual beam focused ion beam scanning electron microscope. The instrument features with Octane Plus EDAX system with low element window and an EBSD detector. The facility houses a VEECO ICON atomic force microscope. The ICON features VEECO's latest technology providing AFM with less than a 35 pm signal to noise operating in tapping mode. The instrument is capable of operating in multiple modes including Contact Mode, Tapping Mode, Non-contact Mode, lateral force (LFM), magnetic force (MFM), surface potential, scanning capacitance (SCM), Tunneling (TUNA or STM), and conductive (CAFM). The instrument is equipped with a variable temperature stage and a wet cell sample holder. Also, the facility is equipped with a Bruker BioScope Catalyst Atomic Force Microscope for the high-resolution study of cells in situ. This AFM resides on a Zeiss AxioObserver Z1 inverted microscope stand equipped with fluorescence optics, a Exfo X Cite fluorescence illumination system and a CCD camera.
The facility features a number of other instruments including; X’Pert PRO PANanalytical powder x-ray diffractometer, LabRam HR Evolution Raman spectrometer from Horiba, Zeiss LSM-710 confocal microscope, SkyScan 1173 µ-CT scanner, Quantum Design Versalab platform, Viscotek GPCMax, Rame-Hart Contact Angle, and PANalytical X-ray fluorescence spectrometer.
For sample preparation facility is equipped with sputter coaters, diamond cut-off saw, EcoMet 300 polishing station, VibroMet2 vibratory polisher, Tousimis Autosamdri-931 critical point dryer, Gatan reaction plasma etcher, and Boeckeler Cryo-Ultramicrotome.
Acquisition of instrumentation in the Nanomaterials Characterization Core Facility was funded, in part, by grants from the National Science Foundation.