00062 - Applied Biochemistry

Learning outcomes

At the end of the course the student:

- knows and understands the principles and applications of biochemical investigation methodologies, both in the research laboratory and in the field of clinical biochemistry, for the purpose of separation, identification, characterization and analysis of biomolecules , of the mechanisms of the metabolic activities of biological phenomena, also in relation to the action of drugs and diagnostic devices for biological analyses.

- will be able to design or evaluate experimental protocols used in biochemical research.

The 2 credits of exercises combined with the 2 credits of laboratory will allow the student to acquire theoretical / practical skills of some methodologies applied in biochemical and clinical biochemical experimentation with particular attention to the logic of setting up experiments and data collection.

Course contents

Applied Biochemistry – Module 1 (3 ECTS) – Prof. Cecilia PrataIntroduction

Principles of biochemical research. Biochemical experimentation. Biosensors.
Overview of microscopy techniques.

Preparative Techniques

Cell cultures: basic equipment in a cell culture laboratory (laminar flow hoods, incubators). Primary cell cultures and established cell lines. Culture media and cultivation techniques.

Biological fluids and antibodies: blood, plasma, serum, urine and other biological fluids. Immobilized enzymes and biosensors.
Antibody structure and antigen–antibody interactions. Production of antisera and monoclonal antibodies. Methods for antibody detection (e.g., serological tests) and antibody‑based antigen detection (e.g., pregnancy tests). Introduction to immunoenzymatic assays (ELISA).

Preparation of samples from biological tissues: selection of buffers. Homogenization (criteria for choosing methods and media).
Centrifugation techniques: principles of sedimentation; speed, RCF, Svedberg coefficient and sedimentation time. Types of centrifuges and rotors. Differential centrifugation: separation and analysis of subcellular fractions; differential flotation of lipoproteins. Density‑gradient centrifugation (zonal and isopycnic). Applications: separation of cells, subcellular organelles, proteins and nucleic acids. Evaluation of yield and enrichment using subcellular “markers”.

Protein purification methods: fractional precipitation with ammonium sulfate and isoelectric precipitation; precipitation with organic solvents and polymers; heat precipitation. Dialysis and ultrafiltration: principles and applications.

Chromatographic Techniques

General principles, resolution and theoretical plates. Column chromatography.
Hydrophobic interaction chromatography (HIC) of proteins.
High‑performance liquid chromatography (HPLC): main components of an HPLC system; selection of stationary and mobile phases depending on the analysis. Main types of detectors used in HPLC.
Gas chromatography (GC).

UV‑Vis Spectroscopy

Principles of UV‑Vis spectroscopy, Lambert–Beer law, instrumentation, chromophores.
Colorimetric assays: methods for determining the protein content of biological samples.

Fluorescence Spectroscopy

Basic concepts of spectrofluorimetry, quantum yield, fluorescence decay, instrumentation, intrinsic and extrinsic fluorophores.
Use of fluorimetric techniques in qualitative and quantitative analysis (linearity of fluorescence intensity vs. analyte concentration, inner‑filter effect).
Phosphorescence and chemiluminescence: basic principles and applications.
Biological applications of fluorescence methods: fluorophore localization and investigation of molecular interaction dynamics using fluorescence quenching (collisional/dynamic and static), Stern–Volmer constant, and determination of diffusion coefficients of fluorophores in solution.
Resonance energy transfer (FRET): energy‑transfer efficiency, applications such as protein oligomerization studies, protein–nucleic acid and protein–lipid interactions.
Fluorescence polarization: determination of static and dynamic anisotropy; applications to protein studies.
Basic principles of light scattering and its use in particle size analysis (flow cytometry).

Applied Biochemistry – Module 2 (2 ECTS – Classroom Practical Sessions) – Prof. Laura GiustiElectrophoretic Techniques

General principles and factors influencing electrophoretic mobility.
Gel electrophoresis: agarose, PAGE, SDS‑PAGE. Detection methods and quantitative evaluation.
Isoelectric focusing (IEF). Two‑dimensional electrophoresis. Detection methods and quantitative assessment.
Capillary electrophoresis.
Blotting – Western blotting for the study of proteins: biochemical and diagnostic applications.

ELISA: direct, indirect, sandwich (direct and indirect), competitive.
RIA.
Overview of additional immunochemical techniques: immunodiffusion, immunoelectrophoresis, immunoprecipitation.

Mass Spectrometry in the Study of Biological Macromolecules

Basic principles of mass spectrometry: ion sources (MALDI, ESI), mass analyzers (TOF, quadrupole).
Applications to peptide identification and analysis: peptide mass fingerprinting.
Use of mass spectrometry to detect protein modifications: phosphorylation stoichiometry, amino acid residue alkylation/acetylation, interactions with drugs, etc.
Protein sequencing (Edman degradation and mass spectrometry–based sequencing).
Introduction to proteomics applications.

Enzyme Purification Methods

Selection of the most appropriate purification strategy for a given sample.
Determination of specific activity and evaluation of yield.
Enzyme assays.
Overview of enzyme kinetics, steady‑state assumption and Michaelis–Menten equation.
Experimental determination of enzyme‑catalyzed reaction rates: continuous, discontinuous, direct, indirect and coupled assays.
Study of enzyme inhibition: competitive, non‑competitive and mixed inhibitors. Lineweaver–Burk plots and determination of Ki.
Immobilized enzymes.

Molecular Biology Techniques

Recombinant DNA technology: basic principles.
Polymerase Chain Reaction (PCR): principles and diagnostic, forensic and paleobiological applications.
VNTR sequences: paternity testing. Identification of specific DNA and RNA sequences: Southern blotting and applications in the diagnosis of genetic diseases.
Northern blotting and applications in gene expression analysis.
Gene expression profiling: DNA microarray technology.

Applied Biochemistry – Module 3 (2 ECTS – Laboratory) – Prof. Laura Giusti

The laboratory course will be organized in two groups: Group A / Group B.

By the end of the Applied Biochemistry laboratory course, students will have practiced and learned several biochemical techniques covered during lectures: working with cell cultures, extracting and separating proteins from cell samples, and determining the concentration and enzymatic activity of an enzyme.

The laboratory is structured into interconnected practical sessions, accompanied by explanatory lectures on the underlying principles of each technique.

• CELL CULTURE

Training in basic techniques for maintaining and culturing stable neuronal cell lines as in vitro models to study biochemical and molecular effects induced by oxidative‑stress–generating compounds. The laboratory will include cell seeding to prepare samples for subsequent experiments.

• CELL VIABILITY ASSAY• PROTEIN EXTRACTION AND COLORIMETRIC PROTEIN ASSAY (BRADFORD METHOD)

Extraction of proteins from a cell pellet and determination of protein concentration in the lysate using the Bradford colorimetric assay. The results will be used to calculate the amount of protein required for enzymatic assays and gel loading.

• DETERMINATION OF THE SPECIFIC ACTIVITY OF AN ENZYME IN CELL LYSATES

Direct enzymatic assay to determine the specific activity of lactate dehydrogenase in neuronal cell lysates.

• ENZYME KINETICS: EFFECT OF SUBSTRATE CONCENTRATION ON ENZYME ACTIVITY

Determination of how reaction velocity varies with substrate concentration and evaluation of kinetic parameters (Km and Vmax). The enzyme studied will be alkaline phosphatase.

• PROTEIN SEPARATION BY SDS‑PAGE

Separation of cellular lysate proteins under denaturing conditions using SDS‑PAGE and determination of the molecular weight of a purified protein of unknown size.

 

Readings/Bibliography

K.Wilson and J.Walker - Biochemistry and Molecular Biology: Principles and Techniques - Cortina Editore, 2006

K.Wilson e J.Walker - Biochimica e Biologia Molecolare: Principi e tecniche – NUOVA EDIZIONE (VIII) 2019

M.Maccarrone - Metodologie biochimiche e biomolecolari. Strumenti e tecniche per il laboratorio del nuovo millennio. Zanichelli 2019

 

Teaching methods

The course includes 5 frontal credits (module 2 with discussion of experimental protocols in class) and 2 laboratory credits.

During the lectures will be discussed the methods connected with the trial in both biochemical and instrumental aspects of the applications. For each technique will be presented and discussed some practical examples.

In the 2 laboratory credits problems such as: determination of the protein content of a solution, determination of enzymatic kinetic parameters, cell viability measurements will be addressed from a practical point of view.

 

*** As concerns the teaching methods of this course unit, all students (including all the international incoming exchange students, i.e. ERASMUS) must attend Module 1, 2 online [https://www.unibo.it/it/servizi-e-opportunita/salute-e-assistenza/salute-e-sicurezza/sicurezza-e-salute-nei-luoghi-di-studio-e-tirocinio], while Module 3 on health and safety is to be attended in class. Information about Module 3 attendance schedule is available on the website of your degree programme ("studiare"--"formazione obbligatoria su sicurezza e salute")***

Assessment methods

Modules 1 and 2: Final exam consists of MCQs and Open-ended questions to assess:

- Knowledge of the basic principles of the main biochemical methodologies used in the separation, identification, characterization and analysis of biomolecules;

- Abilities to design or evaluate experimental protocols used in biochemical research.

 

Laboratory Module (modules 3 and 4) : the final exam will consist in a test with 10 Quizzes/short answer questions

The The overall rating is a weighted average of the two tests.

Teaching tools

Video, PC, Overhead Projector

Office hours

See the website of Cecilia Prata

See the website of Laura Giusti

See the website of Laura Giusti

See the website of Laura Giusti