Download Optics in biology:

Optics in biology:
The use of optics in biology has evolved from the simple light microscope used by Darwin to
the complex cryo-electron and live cell high resolution microscopes used today. With all these
advances it can now be argued that we stand at the dawn of quantitative biology and optics
provides an essential tool in this pursuit.
This course is designed to give students a good understanding of physics involved in advanced
optics while focusing their attention on the biological problems amenable to these techniques.
Students with backgrounds in biology, chemistry or physics are equally encouraged however
knowing basic calculus is a requirement for taking this course. Each section of the course
would deal specifically with a special kind of microscopy followed with a case study in a
biological problem that is most amenable to the use of the techniques discussed.
Main Textbook:
Bioimaging: Current Techniques in Light & Electron Microscopy by Douglas
Chandler and Robert W. Roberson (2008)
Additional References:
Principles of Fluorescence Spectroscopy by Joseph R. Lakowicz (2006)
Single Molecule Techniques: A Laboratory Manual by Paul R. Selvin (2007)
Molecular Biology of the Cell, 4th edition, by Bruce Alberts (2008)
Instructor:
Saveez Saffarian; JFB 203; [email protected]
Teaching assistant:
Carl G. Ebeling; [email protected]
Class and time:
Tuesday, Thursdays; 12:25-1:45pm. JFB 101
Topics:
1. Introduction to cell biology
The animal/plant cell
Cell structure and organelles
The fungal cell, viruses and bacteria
Cellular functions
Traffic, signaling, cell division, motility
Dates
(Jan 12th, 14th, 19th)
2. Introduction to optics and electromagnetic radiation
(Jan 21, 26)
History and basics of the light microscope
Interaction of electromagnetic waves and specimens
Absorption and emission of radiation
Basic principles of fluorescence
Fluorescence resonance energy transfer (FRET), lifetime and polarization
Limits of resolution and invention of electron microscope (EM)
Development of light/electron microscopes
3. The light microscope and image formation
(Jan 28th, Feb 2nd)
Formation of image in a light microscope
Basic light path of a microscope
Kohler illumination
Phase, interference and polarization methods for optical contrast:
Phase contrast microscopy
Dark field microscopy
Differential interference contrast microscopy (DIC)
Infinity corrected optics
Lens aberrations
Case study: Cell motility
4. The human eye and image formation
The anatomy of the eye
Conversion of photons into electrons in photoreceptors
The digital resolution of the eye
Depth and stereo vision
Image processing in the human eye
Optics, transduction and optical nerve output
The neural image
How anti-Aliasing works
(Feb 4th)
5. The electron microscope and image formation
(Feb 9th, 11th)
Formation of image in an electron microscope
Basic wave path of an electron microscope
Illumination system, imaging system, spot size
System and principles of scanning electron microscopes (SEM)
Systems and principles of transmission electron microscope (TEM)
Cryogenic techniques in electron microscopy
Freeze fracture, high pressure freezing, deep etching and rotary shadowing
Crystal structures and refinements in EM images
Case study: Clathrin coated vesicles and viruses
6. Fluorescence microscopy and digital Imaging: (Feb 16th, 18th, 23rd, 25th, March 2nd)
Basics of fluorescence microscopy
Fluorescent dyes and proteins
Principles of wide field fluorescence imaging (WF)
Principles of total internal reflection microscopy (TIRF)
Principles of confocal imaging
Confocal microscopy
Scanning confocal, spinning disk confocal, deconvolution and two photon
microscopy
Digital imaging and photon counting
Basic properties of solid state cameras
Back illumination, electron multiplication and gain
Signal to noise in digital imaging
Shot noise, dark noise and read noise
Case study: Mitosis and bacterial invasion
7. Single molecule imaging and dynamics of biological molecules (March 4th, 9th, 11th, 16th)
An introduction into stochastic processes
Information content of single molecule versus ensemble experiments
Ergodicity and its implication in single molecule analysis
Detection of molecular motions and interactions
Principles of single molecule fluorescence resonance energy transfer (SM-FRET)
Principles of Fluorescence correlation spectroscopy (FCS) and photon counting
histogram (PCH)
Principles of speckle microscopy
Principles of Single particle tracking with nanometer resolution
Fluorescence imaging with one nanometer accuracy (FIONA)
Single-molecule high-resolution colocalization (SHREC)
Case study: DNA polymerase, molecular motors and actin polymerization
Midterm Exam March 18th
8. High resolution microscopy of macromolecules and super molecular complexes
(March 30th, April1st,6th, 8th)
Electron microscopy techniques for studying super molecular assemblies
Rotary platinum shadowing,
Negative staining,
Single particle analysis by cryoelectron microscopy,
Electron microscope tomography,
Near field microscopy
Fluorescence microscopy techniques for high resolution imaging
Stimulated emission depletion microscopy (STED)
Photo-activated localization microscopy (PALM)
Stochastic optical reconstruction microscopy (STORM)
Structured illumination microscopy
Differential evanescence nanometry (DiNa)
Case study: Clathrin mediated endocytosis and virus entry into cells
9. Optical traps and manipulation of molecules under the microscope (This material would be
covered at the instructor's discretion depending on time limitations)
(April 13th, 15th, 20th, 22nd)
Basic principles of optical trapping
Beam path and properties of an optical trapping apparatus
Principles of force clamp
Principles of position clamp
Principles of magnetic tweezers
Double traps and detection of sub-nanometer motion
Case study: Molecular motors
Review April 27th
Final Exam: Friday April 30th 10:30am
Homework would be assigned on a weekly or bi-weekly basis depending on the material
covered.
Exams and grading: Grades would be determined as follows:
30% midterm, 50% final and 20% homework