Syllabus
The Cytometry course is running in 4 subsequent cycles, each containing 3 lectures in two lecture hours (1,5 hours) and one 3-hour practical training which immediately follows the last lecture in each cycle the same day. 1. cycle: Cytometry basics LECTURES: 1. Basics of cytometry - technology, terminology and history.Classification of cytometry techniques - diverse methods of particle detection up to the cytomics (fluorescent, mass, image, -omics combinations). History, parameters measured by flow cytometry, main parts of flow cytometer - fluidics, laminar flow, flow cell, hydrodynamic focusing, principles of sorting.Electronics and optics of cytometers - light sources, mercury lamps and lasers, optical filters. Detection of light pulses, photodiodes and photomultipliers, parameters of light pulse. Signal amplifiers, digital processing, fluorescence compensation, types of plots for data display. 2. Basic application of flow cytometryIntroduction to data analysis - nuclear DNA content, analysis of ploidy, size of nuclear genome, identification of interspecific hybrids, detection of aneuploidy, determination of mode of reproduction, analysis of cell cycle kinetics, apoptosis, analysis of gene expression, study of phytoplankton. 3. PRACTICAL TRAINING: Basics of practical data analysisWhat you see is what you get: data quality, data standards, data quality, data filtering, resolution and data distribution, transformation, displaying the data - basic plots. Introduction to the open source software for basic analysis (FCSalyzer, opt. Flowing Software), complex software packages (SW: FlowJo, BC Kaluza, BD DiVa).
Course credit test #1 2. cycle: Basic cytometric applications LECTURES: 4. Fluorochrome principles and the use of cytometry in biotechnology and biomedicine: Previous cycle repetition with the focus on standardization and reproducibility of measurement. Small fluorochromes, tandem dyes, FRET, fluorescent proteins, sensors. Selection methods in biotechnology, environmental analysis, connection to the -omics methods. 5. Correlation of the measurement to the data visualizationOptics: 3 planes in the flow cell, interrogation point, laser delay, fluorochrome excitation/emission (filters, dichroic mirrors), detectors and signal amplifiers, spectra viewers.Data analysis: light pulse parameters and detection, digital signal processing, compensation of spectral overlap, 1D, 2D, 3D, 4D-kinetics, multiD analysis (incl. dimensionality reduction principles).
Functional tests in cytometry: Apoptosis, proliferation, cell signaling, mitochondria, membrane potential, kinetic tests, rare cells, metabolism. Immunopenotyping of cells and tissues: immune response, tumors, infections, microvesicles, microparticles.Opt. - cytometry methods in virology, microbiology, hydrobiology, protistology and ecology. 6. PRACTICAL TRAINING: Experimental design A design of an experiment (to be done in the 3rd practical lesson) and the limitations of cytometry methods (statistical background - rare cell, autofluorescence, compensation, basics of combinations of fluorochromes, design of antibody panels). Sample preparation theory (solid tissues, bacteria, yeast, viruses, biosafety regulations) 3. cycle: Details of analysis and data measurement LECTURES: 7. Practical data analysis tutorial (titration/voltration)An example of titrated data analysis followed by the measurement of titrated samples. 8. Working with real dataSI calculation based on the data measured last lesson, preparation of a report.
Course credit test #2 9. PRACTICAL TRAINING: Sample preparation and measurement of multiparametric dataPreparation of 8-color panel of monoclonal antibodies according to the recommendation of EUROFLOW, labeling of blood samples and measurement on a multi-parametric flow cytometer (8-color FACSVerse plus an additional one having more detectors = LSR-II, CytoFLEX S).4. cycle: Multiparametric data analysis LECTURES: 10. Basic principles of multiparametric data analysis (FlowJo introduction)Data compensation, visualization, scaling, transformation, normalization. Population identification, FMO controls and controls in general, file concatenation, graphical and statistical output in FlowJo. 11. Data compensation tutorial (FlowJo) 12. PRACTICAL TRAINING: Work with our measured dataThe use of FlowJo in supervised analysis and R tools for unsupervised analysis, data quality check, a complete process of data evaluation, reporting and general troubleshooting - exam training.
Course credit test #3
Annotation
You will be guided through the basic classification of cytometric methods in their historical context. Technological advantages and disadvantages of individual branches of cytometry will be documented using applications from various fields of science: botany, microbiology, protistology, hydrobiology, molecular biology, genetics, immunology and biomedicine in general.
Practical training is an integral part of this course and it is organized in 4 modules (3 hours each):
- Basic data analysis
- Experiment design
- Sample preparation and measurement of multi-parametric data
- Reporting and advanced data analysis
Applications of cytometry cover wide range of experimental approaches: analysis and sorting of particles of different sizes from sub-micron (chromosomes, bacteria, organelles, vesicles, exosomes) through cells (up to hundreds of micrometers) up to whole organisms of 1mm size. Analysis is not limited to static single parameter - univariate tests, but the main power of cytometry lies in multi-parametric analysis of heterogeneous samples and their functional properties. The introduction of new fluorochromes, including variants of fluorescent proteins, allows us to study functional processes in single living cells down to single protein-protein interactions. Therefore, cytometry is an essential building block of systems biology able to measure dozens of parameters in millions of cells at once.
Cytometry instruments as well as cytometry data are highly complex, fully standardized, and allow the use of robust statistical methods for the analysis. The course contains practical demonstration and detailed exercise in data analysis as well. The main emphasis is put on flow cytometry. Other variants, including mass or image cytometry, are covered theoretically but are not part of the practical exercise.
After completing the course you will have:
- Knowledge of principles of cytometry
- Knowledge of cytometry instrumentation and the use of fluorochromes and probes
- Knowledge of current applications in cytometry in the wide range of scientific fields
- Ability to assess the suitability of selected method in solving a specific problem or a question
- Ability to analyze and interpret results generated by flow cytometry
- Ability to critically assess the quality of cytometry data in the scientific publications