Content of Courses Under Development in Structural Biology

BCH 6740: Structural Biochemistry (formerly Advanced Physical Biochemistry) (3-hour course).  Provide an introduction to 1) biological macromolecular structures (protein and nucleic acids), 2) biophysical techniques (NMR, X-ray, Cyro-EM, CD, etc), and 3) kinetics and thermodynamics of molecular motion and interactions. The course objective is to provide the students with a background in biological structure and introduce the methods of structural biology.  This course will be a prerequisite for the following more advanced courses.

Part A: Macromolecular Structure

    1.    Peptide Bonds - Dimensions and isomerism
    2.    Dihedral Angles - Secondary Structure
    3.    Tertiary Structure
    4.    Structural Motifs
    5.    Prediction of Structure - Secondary and Tertiary
    6.    SYBYL – Computer lab experience
    7.    RNase-Experimental and Theoretical Folding
    8.    Chaperones and Chaperonins
    9.    Protein Design and Engineering
    10.  Supramolecular Structure: Membranes, Channels and Membrane-Associated Proteins
    11.  Supramolecular Structure: Nucleic Acid Structure
    12.  Supramolecular Structure: Ribosomes
    13.  Supramolecular Structure: Chromatin and Ribonucleoprotein Particles
    14.  Supramolecular Structure: Cytoskeletal Fibers
    15.  Supramolecular Structure: Viruses
 
Part B: Biophysical Techniques

    16.  X-Ray Crystallography – Principles and experimental design
    17.  X-Ray Crystallography - Active Sites
    18.  Cryo-EM – Principles and experimental design
    19.  Cryo-EM –  Structure analysis
    20.  NMR – Principles and multidimensional Spectroscopy
    21.  NMR – Nuclear Overhauser Effects and Structure Analysis
    22.  Absorption Spectroscopy Ultraviolet/Visible/Infrared; Beer-Lambert Law; Molar Absorptivity; Solvent Effects;
            Difference Spectroscopy)
    23.  Circular Dichroism
    24.  Fluorescence  (Basic Phenomena; Born-Oppenheimer Approximation; Franck-Condon Principle;  Prompt versus
            Delayed Emission;  Anisotropy; Lifetimes (including Rotation Diffusion); Recovery after Photo-bleaching; Caged
            Fluorophores; etc.)
    25.  Fluorescence
    26.  Light Scattering
    27.  Viscosity
    28.  Molecular Diffusion
    29.  Ultracentrifugation (Isopycnic Centrifugation; Sedimentation Velocity; Sedimentation Equilibrium
    30.  Ultracentrifugation
 
Part C: Kinetics & Thermodynamics

    31.  Chemical Kinetics (Time-Scales of Chemical Processes; Basic Definitions, including Reaction Rate, Reaction Order,
            Molecularity, Rate constant, Diffusion-Limited Process, etc.;  First-Order Processes;  Biomolecular Reactions;
            Relaxation Methods;  Stopped-Flow and Mix-Quench Methods)
    32.  Chemical Kinetics
    33.  Chemical Kinetics
    34.  Model Testing (Simulation & Data Analysis)
    35.  Equilibrium Processes (Ligand Binding; Scatchard Analysis; Hill Equation for Infinite Cooperativity;
            Adair-Koshland-Nemethy-Filmer Treatment; Monod-Wyman-Changeux Model)
    36.  Equilibrium Processes
    37.  Equilibrium Processes
    38.  Basic Enzyme Kinetics (Rapid Equilibrium and Steady-State Models for Enzyme-Catalyzed Processes; Equilibrium
            Exchange Behavior; Kinetic Isotope Effects)
    39.  Basic Enzyme Kinetics
    40.  Basic Enzyme Kinetics
    41.  Pulse-Chase Kinetics
    42.  Kinetics of Protein Polymerization
    43.  Kinetics of Ion Channels
    44.  Kinetics of Protein-Nucleic Acid Interaction
    45.  Kinetics of Protein-Nucleic Acid Interaction
 
 

BCH 67aa: Molecular Structure Determination (3- or 4-hour course). Provide a detailed treatment of current methods (NMR spectroscopy and X-ray crystallography) for protein structure determination and for the study of protein interactions. Half the lectures will discuss NMR and other half X-ray methods. In addition, students will have a lab for each section to provide practical experience in sample preparation, operation of the instrumentation, and preliminary data analysis. Students may attend one or both laboratory classes. Course credit will be 3 hours with 1 lab and 4 hours for both labs). The course objective is to provide the students with the knowledge necessary to apply modern methods of structure determination to solve research problems

X-ray Crystallography: A Practical and Theoretic Approach. The course is divided into two parts (theoretical and practical components) that will run concurrently. The students will learn the theory behind the technique of X-ray crystallography and will apply the knowledge obtained to the structure determination of a macromolecule. This hands-on approach will reinforce the applicability of this methodology in the analysis of the functional properties of a biological molecule.

Lectures (1-hour class): Theory

    1.    General Overview
    2.    Protein Purification
    3.    Crystallization
    4.    Specimen preparation
    5.    Diffraction theory: Braggs Law
    6.    Data Collection/instrumentation
    7.    Data Collection procedure
    8.    Space group determination:  Crystal symmetry
    9.    Data processing and reduction
    10.  Fourier transforms
    11.  Phase determination:  Heavy atom method
    12.  Phase determination:  Molecular replacement method
    13.  Model building:  Electron density map interpretation
    14.  Model refinement
    15.  Model validation and functional interpretation

    Laboratories (1-hour class; 3 hour morning or afternoon sessions): Practical Approach to crystallization and
        structure determination of Lysozyme

    1.    Crystallization
    2.    Crystal preparation and data collection
    3.    Data processing and reduction
    4.    Phasing, model building and refinement
    5.     Structure function analysis

    Facilities required:

    1.    X-ray facility (Room LG-171 in MBI) for data collection in pairs at each session
    2.    Crystal Clear software provided by Rob McKenna (PC’s in LG-191)
    3.    Multidisciplinary Simulation and Computer Laboratory  (Room L1-111 in MBI: CNS and O software
            provided by Rob McKenna.

NMR Spectroscopy: The NMR portion of the class is an introduction to the theory and practical
implementation of modern biological NMR spectroscopy.  In the lectures, students will learn the basis of
multidimensional (2D, 3D, 4D, ...) and multinuclear (1H, 15N, 13C, ...) NMR experiments.  In the lectures,
emphasis will be placed on practical applications that will involve hands-on exercises using various computer software packages.  The laboratory will complement the lectures and will provide initial training to become a regular user of the Advanced Magnetic Resonance Imaging and Spectroscopy (AMRIS) Facility of the McKnight Brain Institute (MBI) of UF.  Pairs of students will work together to perform labs by using the AMRIS 500 MHz spectrometer and MBI computer facilities.
 

Lectures (1-hour class):

    1.    Behavior of nuclear spins in a magnetic field
    2.    Product operators I (tools to describe the basics)
    3.    Data collection and the Fourier transform
    4.    Chemical shift and scalar coupling
    5.    Relaxation
    6.    Product operators II (description of multidimensional experiments)
    7.    2D NMR
    8.    Data processing
    9.    Introduction to protein data analysis: TOCSY, NOESY
    10.  Heteronuclear NMR experiments
    11.  Triple resonance data analysis
    12.  Dynamics
    13.  Oriented samples
    14.  Structure Calculations I
    15.  Structure Calculations II

Laboratory (1-hour class):

    1.  Experimental basics of NMR: Sample preparation and loading, probe tuning, locking, shimming, pulse
         width calibration, and 1D 1H NMR.
    2.  2D 1H NMR experiments: NOESY and TOCSY.  Resonance assignments with a small peptide.
    3.  Indirect observation: X-pulse width calibration while detecting 1H, 15N-edited 1H NMR experiments, 2D
         15N-HSQC.
    4. 2D 15N Relaxation experiments (2 Lab classes): 1H-15N NOE, T1, or T2.

Facilities required:

    1.    500 MHz narrow-bore NMR instrument (MBI AMRIS Room LG-123D) for data collection
    2.    CITS Computer Facility  (Room L1-111 in MBI): Mathematica, Biosym, Bruker X-WinNMR, NMRPipe  and
            NMRView

BCH 67bb: Imaging and Monitoring Biochemistry in Living Systems (3-hour course). Lectures will provide a detailed treatment of current imaging methods (MR imaging, laser/optical microscopy) for visualizing the structure of cells, tissues and whole animals. Also current methods (in vivo MR spectroscopy) will be treated for monitoring biochemistry in cell suspensions and whole animals. Half the lectures will discuss MRI and other half laser/optical methods. In addition, a lab will be taught with each section so that the students will have practical experience in sample preparation, operation of the instrumentation, and preliminary data analysis. The course objective is to provide the students with the knowledge necessary to apply modern methods of imaging to solve research problems.

MR Imaging and Spectroscopy: These lectures and labs will provide instruction in the theory and practice of modern methods used in the study of biological sample with a focus on living systems.  The goal of the course is to provide the knowledge and experience necessary for the student to apply this technology to the study of significant biological problems.

Lectures (1-hour class):
    1. Behavior of nuclear spins in a magnetic field
    2. RF coils and magnetic field gradients
    3. Data collection and Fourier transformation
    4. Product operators I (tools to describe basic theory)
    5. Relaxation and magnetic-field-strength dependence
    6. Slice selection using shaped RF pulses
    7. Basic two-dimensional Fourier MR image acquisition and processing
    8. Spin echoes, gradient-recalled echoes, and stimulated echoes for imaging
    9. Rapid imaging methods: RARE and Echo planar
    10. Weighting image intensity with relaxation, diffusion, flow or magnetization transfer
    11. Functional magnetic resonance imaging
    12. Product operators II: Chemical shift and scalar coupling
    13. Chemical-shift-selective imaging (e.g. fat suppression) and spectroscopic imaging
    14. Localized MR spectroscopy
    15. Measurement of physiological parameters; pH and reaction rates

Laboratory (1-hour class):

    1. Experimental basics of MR spectroscopy; a) Sample preparation and loading, b) RF coil tuning, c) shimming, d) RF
        pulse-power calibration, and e) 1H NMR spectroscopy.
    2. Basic MR imaging; a) single- and b) multiple-slice 2D spin-echo Fourier imaging with relaxation weighting, c) 3D
        spin-echo and d) gradient-recalled-echo Fourier imaging.
    3. MR imaging of diffusion, perfusion and flow
    4. Localized MR spectroscopy
    5. RF coil construction; a) Single-turn surface coil, and b) coupled coils

Facilities required:

    1. 4.7 T animal MR facility (MBI AMRIS Room LG-123C) for data collection
    2. RF coil lab (Room LG-131 in MBI) for RF coil construction
    3. Temporary student accounts on Faraday (Room LG-152 in MBI): IDL software package
 

Optical Microscopy: The OM portion of the class is an introduction to the theory, instrumentation and practical implementation of optical microscopy to study biological problems with an emphasis on the application of fluorescence. In the lectures, students will learn the theoretic basis of the use of fluorescent probes targeting specific cellular structures in 3D and more dynamic processes such as localized ion concentrations, electrical potentials, and cellular trafficking (4D imaging adding the time domain). In the lectures the students will be prepared to do the hands on experiments in the labs of investigators and the core OM facilities of the McKnight Brain Institute. The course will provide a level of training sufficient to create independent users of the various microscopic techniques available, and a thorough understanding of the techniques that should foster the development of original applications in their own fields of research.

Lectures (1 hour class):

    1. Introduction to fluorescence microscopy, physical basis.
    2. Components of the fluorescent compound microscope, optics.
    3. Light sources and light detectors used in optical imaging.
    4. Optical Probes and their applications, properties and history.
    5. Using optical probes in cells, practicalities, problems and pitfalls.
    6. The reality of imaging, is what you see what happened?, artifacts.
    7. Optical probes for specific molecules, ions, organelles and cells.
    8. Photolabile caged compounds and their applications in signal transduction research.
    9. Technology for qualitative and quantitative detection of optical probes in cells.
    10. Application of confocal microscopy (LSCM) for optical probe imaging.
    11. LSCM: Dual excitation, high-speed, 4D imaging and kinetic studies.
    12. LSCM: multi-photon, imaging of thick living preparations and tissues.
    13. CCD cameras and low light imaging, deconvolution microscopy.
    14. Special applications: flow cytometry, biosensors, transgenic probes.
    15. Special applications: nuclear calcium signaling, gap junctions, cellular transport, cytoskeletal dynamics in mitosis and cell
          motility.

Laboratory (1 hour class)

    1. Conventional digital imaging of calcium: qualitative (CCD) vs. quantitative (photon counting) measurements.
    2. Image analysis, morphometric 2D measurements, 3D reconstruction, software packages.
    3. Confocal microscopy 1: Multi label imaging of structures, co-localization.
    4. Confocal microscopy 2: Dynamic imaging in the time domain, time-lapse, kinetics of transport and ion concentration.
    5. Deconvolution microscopy: Optical sectioning and 3D reconstruction.
 
Facilities required:

    1. OM Facility (MBI-UF Rooms LG-164 & LG-180) for data collection
    2. Dr. Knot’s Lab (ARB R5-285) for quantitative preparations.
    3. CITS Computer Laboratory (Room LG-191 in MBI-UF) for image processing
 

  • BCH 67xx: Numerical Methods in Structural Biology   (1-hour course, 15 hours of lectures and labs).  Provide an overview of mathematical and computational methods needed to understand current structural models, biophysical processes, data acquisition methods, and the analysis of data acquired with current methods in structural biology. For students without advanced course work in physical and mathematical methods, this course will be taught as a complement to the “methods” courses (BCH 67aa and 67bb).  The course objective is to provide the students with an overview of mathematical and computational methods necessary to perform complex analysis of biophysical and structural data.  Course prerequisite is undergraduate calculus or equivalent.


    • Lectures and Labs (with homework in Mathematica):

        1. Introduction to computers and programming languages (e.g. Mathematica)
        2. Geometry and vectors
        3. Matrix algebra
        4. Waves; linear, circular, and interference
        5. Wave analysis and Fourier transformations
        6. Noise and uncertainty in measurements
        7. Probability and predictions
        8. Sample-size selection and error estimation
        9. Raw data file format(s)
        10. Molecular Geometry
        11. Calculating distance between atoms (1 hour lab)
        12. Calculating Ramachandran diagrams (1 hour lab)
        13. Image geometry
        14. Image windowing and fast Fourier transformation (1 hour lab)
        15. Distance and volume measurements (1 hour lab)
     
    Facilities required:

        1. Multidisciplinary Simulation & Computer Lab (Room L1-111 in MBI): Mathematica, CNS and O software
        2. CITS Computer Laboratory (Room LG-191 in MBI): PC’s with Crystal Clear software
        3. Temporary student accounts on computer, Faraday (LG-152 of MBI); IDL software