Syllabus
Entrance test, introductions to individual practical tasks, independent measurements on fluorometer, evaluation of experimental data, preparation of protocols.
Practical tasks: a) Fluorometer calibration, excitation and emission spectra, fluorescence intensity, polarity of environment. Artifacts (inner filter effect, background correction, Raman scattering). Design of experiments and data evaluation. b) Tryptophan and tyrosine as intrinsic fluorophores, their emission spectra alone and in proteins, spectral shift. c) Polarized fluorescence, fluorescence anisotropy, membrane fluidity and phase transition. d) Time-resolved measurement: determination of concentration of Cl- ions (quenching). e) Evaluation of the data from the confocal microscope: GP of Laurdanu (membrane fluidity); ratiometric fluorescence measurements (changes in calcium concentration in the cells).
Software used: MS Excel or Gnumeric (GNU) spreadsheet, Fityk (data anlysis and fitting, non-linear regression; http://fityk.nieto.pl/), WCIF ImageJ (microscopic image processing; http://fiji.sc/Fiji, http://imagej.net/Welcome).
Annotation
If possible, a full-time exercise will take place in 2020/2021. In case of a change to the online version, the enrolled students will be informed in time.
Laboratory course in fluorescence spectroscopy with biological and biochemical applications. The course covers introductory lecture about fluorescence methods and introductions into individual practical works covering: spectra measurements, fluorescence quenching, polarized fluorescence, FRET, fluorescence of proteins, dynamics of biological membranes, physiological characteristics of cells measured by fluorescence techniques, digital image processing.