Methodologies and Experimental Models for Therapy with Hormones

The course will present and discuss the major techniques of molecular biology, in relation to specific endocrinological questions, including the use of hormones in as therapeutic strategy and the study of their mechanisms of action. In particular, the following topic will be presented:

- Study of the genome, applied to hormones:
Extraction of nucleic acids for the study of gene expression, quantification, electrophoresis
DNA study through restriction endonucleases
PCR; RT-PCR; Real-Time PCR
Study of promoters; design of oligonucleotides for Real-Time PCR
Sequencing; micro and macroRNA

- Study of protein hormones, receptors and enzymes:
Protein extraction, quantification, immunoblotting
Immunohistochemistry
ELISA and Radio Immuno Assay
Chromatography and mass spectrometry: general concepts

- Study of protein-DNA interactions:
EMSA; DNase I Footprinting
Chromatin immunoprecipitation (Chip)
SELEX

Other methods:
LC-MS/MS, practical examples (i.e. method set up; steroid molecules and catecholamines extractions from biological tissues)

Animal models applied to the study of specific hormonal conditions:
Colony maintenance, genotyping
Smear: ovarian cycle in rodent
Animal models for the study of endocrine pathophysiology
Animal models for the study of sexual differentiation
Virus infection (ICV)
Optogenetics for hormones study
Innovative techniques for the study of hormones
Behavioral analysis in rodents
Therapeutic use of hormones

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:

The laboratory for eukaryotic 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.
The culture environment: substrates, culture medium, temperature, growth factors, serum types and chemical additives.

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, maintaining and differentiating.
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.

Use of bacterial cultures for the production of plasmids for cell cultures.
Growth conditions of E.Coli, liquid and solid media, growth curves, freezing and preservation, transformation with plasmid DNA, DNA extraction. Examples of bacterial vectors: plasmids, lambda bacteriophage and cosmids. Essential elements for a plasmid (origin of replication, multiple cloning site, promoter, selection markers, lactose operon and beta-galactosidase), cloning strategies. The lambda phage and its uses for cloning exogenous DNA. The cosmids.

Cell transfection techniques for gene expression study and for proteins analysis.
Genetic cargoes for protein overexpression or gene silencing: DNA plasmids (classic pDNA, enhanced episomal vectors, minicircle), antisense oligonulceotides, siRNA and micro-RNA.
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.
Genome editing: homing-endonucleases, zinc-finger systems, TALENs, CRISPR / Cas9 technology.

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.

Cytofluorimetry.
Introduction to cytofluorimetry, the flow cytometer, measurable parameters (density of a cell suspension, complexity, size and shape of the cells analyzed, detection of fluorescent dyes). Examples of applications: identification of different cell types within a sample, cell sorting, study of proliferation and cell cycle, measure of apoptosis.

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 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).
Examples of the use of cell cultures for the study of hormones