Methods in Cell Biology and Biochemistry

A.Y. 2019/2020
Overall hours
BIO/10 BIO/13
Learning objectives
The aim of the course is to:
- provide an adequate description of the techniques used for the "in vitro" growth and manipulation of cells and the use of different types of cell cultures in the biotechnology field;
- present the principles and applications of basic and advanced methods and approaches for the study of biologically active molecules and biochemical and molecular processes, with particular attention to biotechnological approaches used in modern biochemical and biomedical research;
- to deepen, from a methodological point of view, the knowledge acquired in the other disciplines of the course of studies carried out in the previous semesters (General and cellular biology, Biochemistry and Molecular biology).
With the theoretical part, the course contributes to the consolidation of basic knowledge of living systems at molecular and cellular level. Through laboratory exercises, students have the opportunity to become familiar with the manipulation of cells in culture in sterile conditions and with the use of basic biochemical and bio-molecular techniques, in line with the professional profiles and employment opportunities provided for the entire training.
In addition, the teaching contributes to provide the basis for the continuation of studies in the master's degree courses in biotechnology and bio-medical area.
Expected learning outcomes
At the end of the course, the student should have acquired the theoretical and practical knowledge relating to the main cellular and biochemical methodologies used in research and analysis laboratories in the bio-medical field. In particular, the student should have acquired basic skills for the manipulation of eukaryotic cells in sterile conditions and the ability to perform standard protocols for the analysis of proteins and nucleic acids. In addition, the student should be able to: identify the most suitable techniques or experimental approaches depending on the context of application; identify the correct or most appropriate solution through simulations of experimental cases; critically analyze the results obtained after the application of the techniques studied and or put into practice during laboratory exercises.
Course syllabus and organization

Single session

Lesson period
Second semester
Course syllabus
The laboratory for eukaryotic cell cultures
Sterility, personal protective equipment and collective protection, biological safety hoods, incubators, small instrumentation 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.

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).

During laboratory practice, students will be taught to work in sterile conditions in a biological safety cabinet, introducing them to the basic techniques of manipulation of eukaryotic cell cultures. Students will learn to evaluate the health of a cell culture, to detach the cells by trypsinization, to count them and to carry out a cell seeding. On the seeded cells, each student will experiment, with different methods illustrated in the lectures, including: transfection (with two different methods), a cell vitality test, a cell mortality assay, transfection efficiency evaluation using fluorescence microscopy and assay of beta-galactosidase enzymatic activity.

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

- Design of primers and probes for Real time PCR, quantitative analysis of qRT-PCR 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)
Prerequisites for admission
The following prerequisites are recommended:
- Cell biology
- Molecular biology
- Biochemistry
Teaching methods
The teaching is articulated with frontal lessons (2.5 CFU), seminars of experts of the various topics (0.5 CFU), classroom exercises (0.5 CFU).
The strongly experimental character of the teaching is realized through different single-place laboratory experiences (2.5 CFU).
Teaching Resources
The following texts are recommended for consultation;
A.J. Ninfa, D.P. Ballou, Basic Methodologies for Biochemistry and Biotechnology, Zanichelli
Voet & Voet, Biochemistry, Zanichelli
Watson, Baker, Bell, Gann, Levine, Losic, Molecular Biology of Gene, Zanichelli
Nelson, Cox, Lehninger's Principles of Biochemistry, Zanichelli
Iconographic material in digital format provided by the lecturer via the Ariel website
Assessment methods and Criteria
The examination will be divided into two tests.
Written test:
The written test consists of a questionnaire of multiple-choice questions on topics from the teaching unit of Cellular Methodologies (15) and Biochemical Methodologies (15). Students who have correctly answered at least 8 questions for each teaching unit are admitted to the oral test.
Oral exam:
The oral test takes place immediately after the written test and is aimed at completing the assessment of the student's knowledge and ability to discuss the various topics covered during the course.
Methods in Biochemistry
BIO/10 - BIOCHEMISTRY - University credits: 0
BIO/13 - EXPERIMENTAL BIOLOGY - University credits: 0
Practicals: 8 hours
Single bench laboratory practical: 24 hours
Lessons: 8 hours
Turno 1
Professor: De Fabiani Emma Selina Rosa
Turno 2
Professor: Audano Matteo
Methods in cell biology
BIO/10 - BIOCHEMISTRY - University credits: 0
BIO/13 - EXPERIMENTAL BIOLOGY - University credits: 0
Single bench laboratory practical: 16 hours
Lessons: 16 hours
Professor: Crippa Valeria
Professor: Crippa Valeria
Turno 1
Professor: Crippa Valeria
Turno 2
Professor: Crippa Valeria
Turno 3
Professor: Crippa Valeria
Mondays, Wednesdays, and Fridays from 4:00 p.m. to 5:00 p.m. and upon request via Microsoft Teams or email
Microsoft Teams