Methods in Cell Biology and Biochemistry

A.Y. 2021/2022
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
More specific information on the delivery modes of training activities for academic year 2021/2022 will be provided over the coming months, based on the evolution of the public health situation.
Course syllabus
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:
-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, 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.
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 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: homing-endonucleases, zinc-finger systems, TALENs, 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.
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 (using two different methods), a cell viability assay, a cell death assay, evaluation of transfection efficiency using fluorescence microscopy and beta-galactosidase enzyme activity assay.

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 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)
Prerequisites for admission
The following prerequisites are recommended:
- Cell biology
- Molecular biology
- Biochemistry
Teaching methods
The teaching is articulated with: a) frontal lessons (2 CFU for the unit of Cellular Methodologies and 1.5 CFU for the unit of Biochemical Methodologies); b) seminars and classroom exercises (0.5 CFU for the unit of Biochemical Methodologies); single-place laboratory experiences (1,5 CFU for the unit of Cellular Methodologies and 1.5 CFU for the unit of Biochemical Methodologies).
Teaching Resources
Students will be provided with the slides presented during the lesson in pdf (Ariel platform); as an in-depth text is recommended: 'Colture cellulari' by Mariantonietta Meloni, Aracne editrice.
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 and other interactive platforms
Assessment methods and Criteria
The examination will be divided into two tests, a written test and an oral exam.
The written test consists of a questionnaire of multiple-choice questions on topics from both teaching units (15 concerning Cellular Methodologies and 15 concerning Biochemical Methodologies). Students who have correctly answered at least 8 questions for each teaching unit are admitted to the oral test.
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.
Specifically, the examination will be aimed at:
- assess the level of knowledge and understanding of the topics covered during the course discussing both theoretical aspects (for example the underlying the methodologies described during the course) and the most common applications in the bio-medical and biotechnological field;
- verify the ability to apply the acquired knowledge to specific cases taking inspiration both from examples carried out during the practical sessions and from studies published in scientific journals;
- verify the ability to describe the topics clearly and, where required, with specific terms pertinent to biological and biotechnological disciplines
The evaluation will be expressed in thirtieths and will result from the weighted average of the evaluations related to the two teaching units.
Unità didattica: Metodologie biochimiche
BIO/10 - BIOCHEMISTRY - University credits: 0
BIO/13 - EXPERIMENTAL BIOLOGY - University credits: 0
Practicals: 8 hours
Single bench laboratory practical: 24 hours
Lessons: 12 hours
Turno 1
Professor: De Fabiani Emma Selina Rosa
Turno 2
Professor: Audano Matteo
Turno 3
Professor: Audano Matteo
Unità didattica: Metodologie cellulari
BIO/10 - BIOCHEMISTRY - University credits: 0
BIO/13 - EXPERIMENTAL BIOLOGY - University credits: 0
Single bench laboratory practical: 24 hours
Lessons: 16 hours
Professor: Crippa Valeria
Turno 1
Professor: Crippa Valeria
Turno 2
Professor: Crippa Valeria
Turno 3
Professor: Cristofani Riccardo Maria
Mondays, Wednesdays, and Fridays from 4:00 p.m. to 5:00 p.m. and upon request via Microsoft Teams or email
Microsoft Teams