GEM4 Summer School 2006 - August 7-18, 2006
[[ NOTE: all the "Recommended Reading" papers below can be downloaded together as a single PDF document (22MB) from this location. ]]
Handwritten notes recorded by Maxine Jonas and Jorge Ferrer.
Monday, August 7, 2006
- Parallel Tutorial Session #1: Basic Mechanics (MJ).
- Parallel Tutorial Session #2: Introduction to physiology (JF).
- General Tutorial Session #1: Introduction to infectious diseases (MJ).
- ... Slides by Geert Schmid-Schonbein: "The inflammatory cascade - Part I" (GSS).
- ... Slides by Geert Schmid-Schonbein: "The inflammatory cascade - Part II" (GSS).
- ... Slides by Rhonda O'Keefe: "Biosafety and laboratory preparedness" (RO).
Tuesday, August 8, 2006
- Parallel Tutorial Session #3: Continuum and statistical mechanics (MJ).
- Parallel Tutorial Session #4: Introduction to Molecular Biology (JF).
- Parallel Tutorial Session #5: Introduction to Immune System (JF).
- Parallel Tutorial Session #6: Cell biology lab (hands-on)
Wednesday, August 9, 2006
- Parallel Tutorial Session #7: Molecular mechanics (MJ).
- ... Slides by Ju Li: "Molecular Mechanics: The Ab Initio Foundation" (JL).
- Parallel Tutorial Session #8: Introduction to Cell Biology (JF).
- Lab demonstrations
Thursday, August 10, 2006
- General Tutorial Session #3: Experimental Methods (JF).
- ... Slides by Peter So: "Single and multiple particle tracking methods, and magnetic trap" (PS).
- ... Slides by Peter So: "3D microscopy: deconvolution, confocal, and two-photon" (PS).
- ... Slides by Taher Saif: "MEMS-based tools" (TS).
- Lab demonstrations
Friday, August 11, 2006
- General Tutorial Session #5: Space, time, and energy landscapes in mechanobiology (MJ).
- ... Slides by Ju Li: "Thermal Forces and Brownian Motion" (JL).
- Lab demonstrations
Monday, August 14, 2006
- General Tutorial Session #6: Connective tissue biomechanics (MJ).
- General Tutorial Session #7: Dynamics of infectious deseases (JF).
Tuesday, August 15, 2006
- General Tutorial Session #8: Cell biomechanics (MJ).
- General Tutorial Session #9: Continue session #8 + Case studies (JF).
Wednesday, August 16, 2006
- General Tutorial Session #10: Molecular biomechanics (JF).
- ... Slides by Ju Li: "PolymerChains" (JL).
- General Tutorial Session #11: Applications and case studies (MJ).
Thursday, August 17, 2006
LABS -- August 9-11, 2006
Tissue Biomechanics and Mechanobiology
- Mechanical testing of cartilage under static and dynamic compression and shear. We will demonstrate the use of plugs of bovine tissue run through a series of tests in the apparatus.
- Demonstrations of incubator-housed bioreactors for application of dynamic compression and shear to cartilage specimens, and the various cell- and molecular-biological outcome measures that are used to assess mechanotransduction mechanisms in cells in their native dense extracellular matrix environment.
Title: Fast fluorescence microrheology for quantitative studies of cytoskeletal mechanotransduction
Summary: We will look at cell mechanics using a device of nanometer spatial resolution and covering a five-decade range of frequencies (from 0.5 Hz to 50 kHz). Please read this handout for the lab description, background, and protocol.
Summary: Information regarding cellular rheology is needed to understand a variety of cellular processes including intracellular transport and migration. Because changes in cellular mechanical properties trigger a cascade of biological responses, an understanding of cellular rheology has implications in understanding many biomedical problems, such as the development of cardiovascular diseases (e.g. atherosclerosis) and the development of tissue engineering constructs (such as promoting cell growth over artificial surfaces). In addition, the ability to regulate cellular mechanical properties has important implications in a wide variety of fundamental processes ranging from wound healing to cellular biosensors. Magnetic tweezers belong to a class of active microrheological techniques designed to probe the rheological properties of cells. Fibronectin coated magnetic beads are dropped on the surface of the cell and allowed to internalize overnight. The tweezers are an electromagnet that generates a magnetic field which exerts a quantifiable constant force on a paramagnetic or ferromagnetic object. By varying the current through the electromagnet, the amount of force applied to the bead may be controlled. The displacement of the bead as a function of time may be modeled as a Voigt element in series with a dashpot. Based on this analysis, the viscoelastic behavior of the cell may be characterized by three mechanical parameters. In this lab you will use magnetic tweezers to measure the viscoelastic parameters of NIH3T3 fibroblast cells and compare these obtained results with values reported in the literature.
Multiphoton Excitation Microscopy
Title: Cellular and tissue imaging with multiphoton excitation microscopy
Summary: Two-photon microscopy (TPM), sometimes called multiphoton excitation microscopy, is a three dimensional incoherent imaging technique based on the nonlinear excitation of fluorophores. Two-photon excitation occurs only at focal point where the simultaneous absorption from two photons having half of the energy each is needed for the excitation transition. It has several unique features to image cells and tissues, compared with the other 3D imaging microscopes. First, TPM uses high numerical aperture objective with tunable modelocked femtosecond pulsed laser (titanium:sapphire laser), and they enable various biological specimen imaging with sub micrometer resolution down to a depth of a few hundred micrometers using illumination of near infrared (NIR) wavelength light. Second, TPM has a little effect of photodamage on imaging of living specimens, since excitation happens only where the signal is collected without reducing cell viability. TPM can initiate photochemical reaction within a subfemtoliter volume inside cells and tissues, unlike confocal microscopy which obtains three-dimensional information by eliminating out-of-focus light through the use of a pinhole, and result in minimizing photobleaching. Fourth, TPM allows high-sensitivity imaging by eliminating background such as the contamination of the fluorescence signal by the excitation light. Last, the excitation wavelength used in confocal microscopes is typically in the UV and blue/green range which is scattered and absorbed strongly in tissues. This effect limits the depth of signal detection. In this lab, we briefly introduce the TPM instruments (lasers, intermediate optics, objective, detector, and etc) and image two sets of specimens: fluorescence-labled cells, and cartilage tissue containing chondrocytes and collagen. In the case of cells, we collect three different fluorescence signals with phtomuliplier tube (PMT), using different color filters. For the tissue imaging, we image Chondrocytes stained with green fluorescent cell tracker in the tissue. We also image SHG signal of cartilage collagen. We use ImageJ to quickly look at the 3D image stack obtained during imaging, and further image visualization.
Optical Trap: DNA
Title: Optical Trapping and Single Molecule Fluorescence
Summary: Optical tweezers are an excellent experimental tool to study the biophysics of single molecule systems including the mechanics of molecular motors (kinesin, myosin, RNA polymerase), mechanical conformations/transitions of molecules (dsDNA, RNA hairpins, filamentous proteins) and interactions of receptor-ligand systems(anitgen-antibody). In the most common assays, the mechanical state of the system is monitored by tracking the position of a handle (usually a dielectric microsphere with diameter of 0.5-2um) tethered to the molecule of interest (protein, DNA, etc), with nanometer and picoNewton resolution. The handle also serves as probe to apply force to the system to study the energetics of mechanical changes. Single molecule fluorescence allows the direct observation of the mechanical/conformational changes of the system as it is subjected to perturbations, such as force. The combination of these two techniques allows researches to study the biophysical properties of single molecules. In this lab you will learn the basics of operating a high-end optical tweezers to record mechanical transitions of single molecules. The instrument is also equipped with a novel single molecule fluorescence technique to allow simultaneous, coincident optical trapping and single molecule flourescence. In our demonstration we will measure the force required to unzip a double-stranded DNA molecule, with a resolution of ~5nm and ~0.1pN, while using single molecule fluorescence to confirm the location of the break. Alignment and calibration procedures will also be presented.
See file below for a short description of the experimental set up and what you will see in our demonstration.
"Experimental set up" by J.Ferrer
(optional) Optical Trapping Review : K.C. Neuman & S.M. Block, "Optical trapping," Rev. Sci. Instrum. 75 (2003).
Optical Trap: cells
Title: Optical Tweezers: Membrane and Cell
Summary: Optical tweezers that exert forces up to hundreds of picoNewtons can probe the mechanical properties of membranes and cells. Standard tweezers experiments involve optically trapping small microspheres attached to specific cell membrane locations which serve as grips to deform the cell membrane locally (tether experiments) or entire cell (cell stretch experiments). Calibration of optical trap forces on the microspheres allows quantitative measurement of static and dynamic cell mechanical properties. In this laboratory module, two experiments will be performed: membrane tether pulling of B-cells and cell stretch tests of erythrocytes.
AFM and optical trapping in education
Title: Advanced Instrumentation in the teaching lab
Summary: This session makes use of the atomic force microscopy (AFM) and optical trapping experiments used to teach an undergraduate instrumentation laboratory. Using a home-built and easily-manipulated AFM, you will learn basic imaging techniques, take some simple force curves for elastic modulus, and make a measurement of thermal energy. Using similarly scaled-down optical traps, you will learn calibration methods for the optical traps and perform experiments on the molecular motors of E. coli bacteria. The documents in "Recommended Reading" below provide the essential background for both instruments and an overview of the experiments.
- The lab manuals for the teaching AFM experiments.
- A document containing a description of the optical trapping tool and associated lab procedures: Optical Trapping for the Teaching Lab.
Title: Atomic force microscopy imaging of cells
Summary: In this laboratory, you will use the atomic force microscope to image the structure and stiffness of living and chemically fixed human microvascular endothelial cells. The pN- to nN-scale mechanical force used to create these images allows you to observe both the micrometer-scale height of these cells, as well as the nanometer-scale cytoskeletal network beneath the cell surface. Because the cells are living and imaged under near in vitro conditions, it is possible to observe cell processes in real time, including migration, response to drugs added to the imaging media, and of course apoptosis. It is also possible to compare the near-surface structure of living and diseased cells. If time allows, you will also observe the near-field optical / fluorescent image of these cell surfaces.
AFM: force spectroscopy
Title: Molecular force spectroscopy on living cells
Summary: In this laboratory, you will use the atomic force microscope to acquire the mechanical interaction forces between the AFM probe and the surface of living human microvascular endothelial cells. By pushing into the cell surface, the stiffness of various points on the cell can be determined qualitatively. By pulling away from the cell surface, the adhesion force between the probe and specific points on the cell membrane can be measured, including the imaging of single cell surface molecules. Both of these loading approaches are used to infer changes in the cell surface / interior as a function of mechanical or chemical environments, and as a function of disease state.
Title: BioMEMS Force Sensor
Summary: We will be showing a bioMEMS force sensor and its application in measuring stretch and compression force response of healthy and malaria-infected human red blood cells. The bioMEMS force sensor is made from pure single crystal silicon, and consist of a probe and flexible beams. The probe is used to contact, indent and stretch the cells, and the flexible beams to measure the cell force response. The probe is about 5 µm wide and 5 µm deep. Each of the flexible beams is about 2 mm long, 1 µm wide and 5 µm deep. We will show how to manipulate the sensor and bring it in contact with the cells, and how the cell force response is measured. Every student will have the chance to try out this manipulation process. Two particular cell force response measurements will also be shown: a poly-L-lysine coated sensor probe will be used to measure (1) the stretch force response of a healthy red blood cell and (2) the compression force response of a malaria-infected red blood cell. For more information about this bioMEMS technique, please refer to the following two papers:
Summary: This laboratory will demonstrate the use of microfabricated structures to investigate the mechanical response of cells as they are deformed through narrow microfluidic channels. Specifically, the biorheological behavior of red blood cells at different stages of malaria infection will be studied. Under a known constant pressure differential, it can be seen that the entrance time and velocity through narrow channels (varying from 2 - 8 microns square cross-section) differ between the early and late stage infected and much stiffer red blood cells. It can also be shown that the sufficiently stiff cells cannot pass through the narrowest channels. Analogies can be made between this behavior and that experienced in the body as the cell passes through capillaries of comparable size.