Methods in Cell Biology and Biochemistry

CELLULAR METHODOLOGIES
The course aims to provide the basis for methods for obtaining, growing and manipulating cellular models. The following topics will be covered in the course:

Frontal lessons
-Mammalian cell cultures
Advantages and disadvantages of cell cultures.
Primary cell lines: achievement and maintenance. Disaggregation of primary tissues and cultures. Separation and characterization of specific cell types present in a tissue.
Stabilized and / or immortalized cell lines: obtaining and maintaining. The transformation factors, the oncogene-mediated immortalization of cells, the hybrid systems obtained from cell fusions. Specific immortalization with transforming agents: direct tumorigenesis.
Pluripotent induced stem cells (iPSC). Obtaining and maintaining.
Culture methods (suspension, adhesion, monolayer and 3D culture), substrates for cell culture in adhesion, analysis of cell culture morphology. Analysis of growth curves of a cell culture, subculture, mechanical or enzymatic detachment, cell counting and seeding, freezing of cell lines.
The culture environment: substrates, culture medium, temperature, growth factors, serum types and chemical additives.
-The laboratory for mammalian cell cultures
Sterility, personal protective equipment and collective protection, biological safety hoods, incubators, small instruments and cell culture supports. Safety measures and associated risks.
Dry heat and wet heat sterilization techniques, filtrations.
-Cell cultures assays
Cell viability and mortality assays: based on membrane integrity (trypan blue staining, calcein-acetoxy methyl staining, propidium iodide staining, glucose 6P dehydrogenase enzyme release, lactate dehydrogenase enzyme release), based on redox potential (alamar-blue assay, MTT assay), based on mitochondrial functionality (ATP test).
Techniques for the study of cell migration, invasion, adhesion and proliferation (Boyden chamber, aggregates in matrigel, cell adhesion assay).
-Cellular engineering: plasmids, AON, RNAi, miRNA and genome-editing
Genetic cargoes for protein overexpression or gene silencing: DNA plasmids (classic pDNA, enhanced episomal vectors, minicircle), antisense oligonulceotides, siRNA and micro-RNA. Genome editing: zinc-finger systems, TALENs and CRISPR / Cas9 technology.
- Cell transfection techniques for gene expression study and for proteins analysis
Non-viral transfection vectors: chemical methods (calcium chloride, DEAE-dextran, lipofection) and physical methods (electroporation, microinjection and gene gun).
Viral vectors: advantages and disadvantages. Adenoviral, lentiviral, AAV, Herpes Simplex-based vectors. Production of viral vectors: modification of the viral genome, packaging cells, titration and analysis of the relevant replication revertants, main safety standards for working with viruses.
Transient and stable transfections; plasmid vectors with viral or inducible (tet on / tet off) promoters, reporter systems.
-Fluorescence and its applications for the study of cell cultures.
Basic principles of fluorescence. Fluorochromes, fluorescent proteins and fluorescent chimeras. The fluorescence and confocal microscope. Immunocytochemistry and immunofluorescence: cell fixation, permeabilization, detection using primary and secondary fluorescent antibodies. Examples of applications of using fluorescent proteins/compounds to mark organelles, to measure cellular activity.
Fluorescence microscopy techniques: FRET, FRAP, iFRAP, FLIP, FLAP.

PRACTICAL LABORATORY
During laboratory, students will be taught how to work under sterile conditions in a biological safety hood, introducing them to basic mammalian cell culture handling techniques. Students will learn how to assess the health of a cell culture, detach cells by trypsinization, count them, and perform cell seeding. On the seeded cells, each student will experiment with different methods outlined in the frontal lectures, including: transfection, a cell viability assay, a cell death assay, evaluation of transfection efficiency using beta-galactosidase enzyme activity assay. In addition, students will learn to process the data obtained using MS Excel. At the end of the laboratory, each student will have to present the data obtained in a written report, which will contribute to the final evaluation.


BIOCHEMICAL METHODOLOGIES
FRONTAL EDUCATION (including seminars and classroom exercises):
- Quantitative analysis of gene expression: definition and methodologies, basic PCR concepts, qRT-PCR operating scheme, approaches for real-time amplification detection, methods for quantitative data analysis; qRT-PCR applied to miRNAs
- Techniques for the study of macromolecule interactions: co-(immuno)precipitation with native or fusion proteins, GST-pull down and related approaches, study of interactions by FRET (in vitro and in vivo); analysis of interactions between proteins and nucleic acids by Electrophoretic mobility shift assay, analysis of interactions between proteins and DNA in the context of chromatin (Chromatin immunoprecipitation), analysis of proteins interacting with RNA (RNA immunoprecipitation)
- Approaches for the study of the proteome: general definition of proteome, proteomic analysis and related methodological approaches; application of mass spectrometry for the determination of the molecular weight of proteins (MALDI-TOF, ionization by electrospray); approaches for protein identification: 2D-gel electrophoresis, enzymatic digestion, sequencing by tandem mass
spectrometry, peptide fragmentation pattern, quantitative analysis in mass spectrometry.
- Approaches for the study of the metabolome: general definition of metabolome, metabolomic analysis and related methodological approaches; targeted metabolomics: sample preparation, instrumentation, application examples; untargeted metabolomics: sample preparation, instrumentation, software, application examples; statistical analysis of metabolomic data
- Approaches for the study of energy metabolism: methods for the analysis of mitochondrial morphology, methods for the analysis of mitochondrial density and biogenesis, methods for the analysis of respiration in cell cultures and animal models, measurement of ATP synthesis, membrane potential, reactive oxygen species, and main mitochondrial metabolic pathways.
- Next generation nucleic acid sequencing techniques: introduction to next generation sequencing techniques; technologies based on the principle "sequencing by synthesis": applications (DNA-protein interactions, RNA-seq, RNA-protein interactions, DNA methylation, whole genome sequencing, de novo sequencing); libraries preparation; sequencing (multiplex sequencing, cluster
generation); data analysis
LABORATORY PRACTICE
- Design of primers and probes for Real time PCR, quantitative analysis of qRT-PCR data and targeted metabolomics data, introduction to secondary and tertiary analysis of Next generation sequencing data (computer laboratory)
- Isolation of nucleic acids from cell cultures, preparation of the PCR reaction and analysis of amplification products by agarose gel electrophoresis; cell fractionation, analysis of the protein profile of the cell fractions by SDS-PAGE, determination of the protein concentration of the cell fractions by colorimetric method (bio-laboratory)
The contents of the lab activities will be the subject of individual tests that will contribute to the final evaluation