BIOCHIMICA APPLICATA

To confer knowledge and to acquire skills related to the rationale of the various methodologies and instrumental techniques involved in the biochemistry and biomolecular experimentation. To provide the cognitive tools to monitor developments in molecular genetics and its protocols for the identification, cloning with its vectors, sequencing and gene expression that have revolutionized the understanding of biological processes at the molecular level, contributing to the interdisciplinary character of the course.

Frontal lessons with slide projection will be held. The teaching method is developed to stimulate the students' learning ability on the course program. If the teaching is carried out in mixed mode or remotely, the necessary changes may be included with respect to what is reported in the syllabus.

Information for students with disabilities and/or DSA

To ensure equal opportunities and in compliance with current laws, interested students may request a personal interview, in order to plan any compensatory and/or compensatory measures, based on the educational objectives and specific needs. The student may also contact the CInAP (Center for Active and Participatory Integration - Services for Disabilities and/or DSA) contact teacher of the Department of Drug Sciences and Health, Prof. Santina Chiechio.

Principles of biochemical research. Preparation and handling of biological samples. Use and preparation of buffer solutions. Cell disruption; preparation and purification of cell organelles; centrifugation: centrifuges, rotors, differential and equilibrium isodensity centrifugation with application examples (separation of cell organelles, purification of nucleic acids).

Biological experimentation models: Whole animal, perfused organs, tissue sections. Buffer solutions. Cell cultures. The cell culture laboratory. Sterilization. Culture methods: primary, secondary cultures, cell lines. Culture medium. Cell population separation and analysis. Cell count. Cryo-conservation. Flow cytometry, FACS and MACS.

Microscopic techniques. Optical microscope. Optical sectioning. Imaging of living cells and tissues. Stereomicroscope. Imaging and biochemistry. Specialized imaging techniques.

Electrophoretic techniques: General principles. Zonal electrophoresis on cellulose acetate; serum proteins. Gel electrophoresis: agarose, polyacrylamide gel electrophoresis (PAGE and SDS-PAGE). Isoelectric focusing (IEF). Two-dimensional electrophoresis (applications to the study of the proteome). Methods of detection and quantitative evaluations (“gel” staining and following blotting). Capillary electrophoresis.

Chromatographic techniques for the purification, separation, identification and quantitative analysis of low (metabolites, drugs, hormones, etc.) and high (nucleic acids and proteins) molecules. PM: General principles. Notes on TLC. Ion exchange chromatography. Exclusion chromatography, affinity chromatography. HPLC and GC (stationary phases, detectors and main applications).

UV-Visible Spectrophotometry: Review of the nature of electromagnetic radiations. UV-VIS spectrophotometry (principles, instrumentation and applications). Absorption spectra. Lambert-Beer law and its quantitative applications. Calibration line concept. Spectrophotometric determination of the protein concentration. Application of spectrophotometry in enzymatic assays.

Spectrofluorimetry (general principles and applications) and Chemiluminescence. Flow cytometry and cell sorting. Hints of Luminometry.

Immunochemical techniques. Structure of antibodies, antigen-antibody reaction. Production of antisera, monoclonal antibodies. Methods of analysis: free phase and gel immunoprecipitation reaction, immunodiffusion; radioimmunological methods: RIA and IRMA; immunoenzymatic assays: EMIT, ELISA and PEIA. Hormonal dosages: methods and applications.

Electrochemical techniques. Oxygen electrode; studies on mitochondrial respiration. Biosensors.

Radioisotope methods. Principles, instrumentation and applications. Recalls on the types of radioactive decay. Energy and speed of radioactive decay. Detection and measurement of radioactivity. Counting and quenching efficiency. Autoradiography. Applications of radioisotopes in biochemistry. Application of radioisotopes in clinical analysis.

Mass spectrometry techniques for the qualitative and quantitative analysis of low molecules (metabolites, drugs, hormones, etc.) and high molecular weight (nucleic acids and proteins). General principles and instrumentation (sources and analyzers).

Techniques for the study of proteins: methods for sequencing, the determination of the secondary and tertiary structure, and of post-translational modifications and mutations. Introduction to proteomics.

Methods of labeling of biological molecules: Review of the types of radioactive decay. Energy and speed of radioactive decay. Detection and measurement of radioactivity by scintillation. Autoradiography. Marking with non-radioactive systems.

Generalities on nucleic acids. Nucleic acid manipulation: basic tools and techniques. Isolation of DNA and RNA. Automated extraction of nucleic acids by kit. Nucleic acid electrophoresis. Automated analysis of nucleic acid fragments. Elements of bioinformatics.

Molecular biology techniques. Recombinant DNA technology: general principles. Polymerase Chain Reaction (PCR): principles and diagnostic applications. Identification of specific DNA and RNA sequences: Southern hybridization (Southern blotting) and applications in the diagnosis of genetic diseases. Northern blotting and applications in the study of gene expression. Evaluation of gene expression: the DNA microarray technique.

The course includes compulsory laboratory exercises on some of the topics covered in the course.

Wilson K., Walzer J.: Biochemistry and molecular biology: principles and techniques. Ed. Raffaello Cortina Editore.

Ninfa A.J., Ballou D.P.: Basic methodologies for biochemistry and biotechnology. Ed. Zanichelli.

D.L. Nelson, M.M. Cox: Lehninger's Principles of Biochemistry, VII Edition 2018, Ed. Zanichelli.

The learning assessment consists of an oral test. Verification of learning can also be carried out electronically, should conditions require it.

For students attending the course there is an ongoing assessment test with open-ended questions. The test, once the minimum score of 18/30 has been passed, will contribute to the final evaluation.

The final evaluation (exam grade) will be carried out based on the relevance of the answers to the questions asked, the quality of the contents, the ability to connect the topics covered by the program, the ability to formulate examples, the ownership of technical language and the overall expressive ability of the student.

1. Cell cultures. Electrophoretic techniques. Basic principles of fluorescence. Gene silencing techniques. PCR.

2. Confocal fluorescence and laser scanning microscope. Flow cytometer. MALDI and MALDI-TOF. HPLC. Western Blot.

3. Stem cells. Production of polyclonal and monoclonal antibodies. Cloning techniques. Therapeutic proteins. ELISA.