MBS1002 Biomedical Approaches
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| What are the three main cell types that can be cultured in a lab and what are their differences? | 1. primary cells are directly isolated from human/animal tissue. they represent the tissue and organisms characteristics well, but they have limited proliferative capacity. 2. transformed cells are cells that have a fast growth rate and are stable in maintenance and for cloning. they are usually formed by genetic manipulation, which can lead to non-physiological phenotypes. 3. self-renewing cells that are able to differentiate into other cell types, such as pluripotent stem cells. |
| What is the difference between primary cells and cell lines? | Primary cells are directly derived from the patient and have limited proliferation cycles. cell lines are formed when normal cells are cultured and allowed to divide and expand in number, which is usually due to genetic modifications. cell lines are often derived from a tumour of the cell type of interest or by transforming and immortalising a primary cell with an oncogene. contrary to primary cells, cell lines can be maintained indefinitely in culture and are able to generate large number of cells. |
| What are the pros and cons of neuronal cell lines | Pros: can be maintained indefinitely, can generate large amount of cells, ability to manipulate gene expression and control cellular environment. cons: genetic alterations can accumulate, mature neurons are non-dividing and cannot be subcultured, cell line neurons do not represent the full phenotype of a neuron, there is loss of synaptic pathways and normal anatomic relations. |
| What are things to be considered when working with cell cultures in the lab? | - some cell lines are categorized in a high Hazard Group and are pathogenic. - lab workers should wear personal protective equipment (PPE). - surfaces that come in contact with the cell culture media should be cleaned with 70% ethanol. - an aseptic environment is created using a biosafety cabinet. |
| What are the steps of immunofluorescence? | 1. fixation to preserve morphology and cellular architecture. 2. permeabilization in case plasma membrane is not yet disrupted by fixation, to give antibodies access to intracellular epitopes. 3. embedding in paraffin to fix the tissue (not needed for cells). 4. deparaffination and rehydration. 5. antigen retrieval to restore epitope-antibody reactivity through; - protease induced epitope retrieval; enzymes cleave protein cross-links - heat-induced epitope retrieval; heat and pressure is used 6. blocking to prevent binding of antibodies to non-target epitopes. 7. primary or secondary immunofluorescence. 8. detection using fluorescent microscope. |
| What is the difference between primary and secondary immunofluorescence? | In primary fluorescence, one antibody coupled to a fluorophore that binds to the epitope is used, while in secondary fluorescence, a primary antibody is used to bind to the epitope and a secondary antibody coupled to a fluorophore is used to bind to the primary antibody. primary immunofluorescence is quicker, but secondary immunofluorescence is more widely used since it is more sensitive. |
| What is the difference between immunofluorescence and immunohistochemistry? | Immunofluorescence uses a fluorophore to detect the epitope. this means that a confocal or other fluorescent microscope has to be used for analysis. the fluorescence emission can directly be measured. immunohistochemistry does not use a fluorophore, but instead enzymes to detect the epitope. this bond is visualised by the reaction of the enzyme with the epitope and the substrate, and the change in stain/colour that produces. this means that the sample can be analysed using a light microscope. this technique is semi-quantitative since the absolute presence of the epitope cannot be measured. |
| What is transfection? what are the two subtypes? and what are the three ways of transfection? | Transfection is the process of introducing foreign nucleic acids into eukaryotic cells to genetically modify the cell. transfection can be stable, meaning that the foreign nucleic acids are integrated in the host's genome or remain present in the host cell as an extra-chromosomal element. this means that the transgene remains expressed as the cell replicates. transfection can also be transient, meaning that the foreign nucleic acids are present in the host cell as a plasmid or as oligonucleotides. after a couple of cell replications, the transgene will be lost. the three ways of transfection are viral transfection, physical transfection and chemical transfection. |
| What is the process of viral transfection (transduction)? | A viral vector, protected by an envelope, carries foreign nucleic acids into its capsid. the virus interacts with the host cell using surface proteins on its envelope. once inside the cell, the virus releases its nucleic acids. viruses that are often used for transduction are retroviruses (RNAs), adenoviruses (double-stranded DNA) and adeno-associated viruses (single-stranded DNA). retroviruses contain the enzyme integrase which allows for stable transfection. |
| What are the different ways of physical transfection? | Creating holes in the cell membrane to allow for nucleic acid entry using: - electroporation: an electrical voltage is used to increase cell membrane permeability - sonoporation: microbubbles techniques - laser irritation-assisted transfection: a laser beam magnet-assisted transfection: a magnetic force is used to guide nucleic acids across the membrane gene microinjection: a needle is used to puncture the cell membrane and inject the nucleic acids into the nucleus. |
| What is the main principle of chemical transfection? and which molecules are used? | The two main principles of chemical transfection are - that certain positively charged molecules surround the negatively charged nucleic acids, thereby shield it and allow for entry into the cell. molecules that can do this are calcium phosphate, dendrimers, cationic polymers, nanoparticles. - that positively charged lipids surround the negatively charged nucleic acids, enabling merging with the cell membrane and thereby inserting the nucleic acids into the cell. |
| How can efficiency of transfection be assessed? | - fluorescence microscopy with fluorescently tagged molecules - real time PCR to directly measure nucleic acid expression level - plasmid reporting system - flow cytometry to quantify numbers of fluorescently labelled transfected cells - immunostaining |
| What are the disadvantages of the different physical transfection techniques? | The use of high voltage in electroporation can lead to necrosis, apoptosis and permanent cell damage. both sonoporation and laser irradiation transfection carry the risk of damaging the cell membrane and leading to irreversible cell death. compared to these techniques, magnetofection is less destructive to the cell, however, this technique is less efficient. gene microinjection can prevent cell damage very precisely, but this requires trained personnel or robotic systems. |
| What are applications for bacterial and human/animal cell culture? | Bacterial cell culture: diagnosis of infections, genetic manipulation, food and beverage production, detecting food contaminants, development of vaccines and therapeutics, epidemiological studies. human/animal cell culture: gene therapy, model systems, toxicity testing, cancer research, drug screening, vaccine production, genetic engineering. |
| How can plasmids be inserted into bacteria? | 1. competent cell preparation: pick a cell colony, culture it at 37ᵒC, prepare the cells at 4ᵒC, freeze cells at -70ᵒC. 2. transformation, e.g. through chemical or physical transformation. 3. recovery in an antibiotic-free medium to allow for expression of antibiotic resistance genes. 4. cell plating on agar plate to determine transformation efficiency. |
| What is the difference between (bacterial) culture on agar plates or in liquid suspension? | A suspension is used when you want to upscale your culture. an agar plate is used to; - select individual colonies from a mixed culture - to count the colony forming units within a sample - to transport strains from one lab to another |
| What is the transformation efficiency? and what is the transfection efficiency? | Transformation efficiency: the total number of colonies divided by the amount of DNA that was used to transform the cell. transfection efficiency: the number of cells that express the antigen/protein of interest divided by the total number of cells. |
| What is the goal of transformation in bacteria compared to transfection humans? | The goal of transformation in bacteria is to produce multiple copies of a recombinant DNA molecule. the goal of transfection in humans is to introduce nucleic acids into eukaryotic cells. |
| What are the pros and cons of physical transfection? | Pros: simple to perform, good for difficult to transfect cell types, reproducible results, no vector required, less dependent on cell type, rapid transfection of large number of cells. cons: requires special instruments, parameters must be carefully optimized, potential for high toxicity as well as cell damage and mortality. |
| What are the pros and cons of viral transfection (transduction)? | Pros: high efficiency, works well in difficult cells, can be used for the generation of stable or transient cell lines. cons: immunogenicity and cytotoxicity, technically challenging and laborious, high costs, variations in infectivity, low packaging capacity. |
| What is the difference between linear and conformational epitopes? and why is this important? | Linear epitopes are epitopes that are located on the antigen in a linear fashion, while conformational epitopes are spread on the antigen, requiring a fold/3D structure of the antigen when the antibody binds this is important when you're analysing and detecting antibody-antigen reactions. e.g. in ELISA, mostly linear epitopes are used. |
| What is the difference between polyclonal and monoclonal antibodies? | Polyclonal antibodies are antibodies that are produced by different B cells and recognise multiple epitopes of a single antigen. they are cheap to produce, but generate mixed populations of antibodies and they are tolerant to small changes in the protein structure. monoclonal antibodies are antibodies that are generated by identical immune cells, which are clones of a single parent cell. this means that they are expensive to produce, but they will only bind to one epitope. monoclonal antibody production generates a single antibody species. they are senstive to protein structure. |
| What is the basic principle of immunoblot and ELISA for detecting the presence of autoantibodies in a sample? | 1. antigens are labelled on a nitrocellulose filter membrane in case of immunoblot and are coated on a 96-well plate in case of ELISA 2. you add your sample that you think contains the autoantibodies. 3. the autoantibody will bind to the antigen. 4. a secondary antibody conjugated with an enzyme is added. 5. substrate is added to visualise the reaction. |
| What are the different types of ELISA? | Direct: the use of only a primary antibody conjugated with a substrate. indirect: the use of a primary antibody and a secondary antibody conjugated with a substrate. sandwich: the use of a capture antibody that "captures" the antigen as it is added, as well as a primary antibody and a secondary antibody conjugated with a substrate that bind on top of this to visualise the reaction. competitive: the sample antigen competes with a reference antigen in binding to the capture antibody, which is indicative of the concentration of the sample antigen. |
| What is the principle of cell-based assay? and what is an example? what techniques are considered cell-based assays? | Cell-based assays assess the efficacy of compounds in a cellular environment, which aids to understaning of the cell. 1. plasmids that contain the gene of interest are engineerd. 2. the plasmid is transfected into the cells. 3. the corresponding protein is expressed and validated. 4. immunofluorescence is done to check for the presence of the autoantibody. cell-based assays are; flow cytometry, immunocytochemistry, immunofluorescence on cells |
| What is a tissue-based assay? and what is an example? | A tissue-based assay is an experiment that is done on a tissue and gives you insights in tissue characteristics. it is often done for detecting neuronal antibodies. 1. you have a tissue that you want to study. 2. you add a sample that contains autoantibodies. 3. if the antigen is present, the autoantibodies will bind. 4. you add a secondary antibody conjugated with an enzyme. 5. you add a substrate to visualise the reaction. |
| What is immunoprecipitation and how is it done? | Immunoprecipitation is the isolation of a specific antigen from a mixture, using the antibody-antigen reaction. 1. add the suitable antibody to the mixture. 2. the antibody binds to the antigen/protein of interest. 3. IgA or IgG is added to make the antibody-antigen complex insoluble. 4. centrifuge to form a pellet so that the supernatant can be removed. |
| What are the 3Rs? | Replacement: the use of animals should be completely (or partially) avoided. reduction: the number animals should be minimised. refinement: the pain, suffering, distress and lasting harm that animals might experience should be minimised. |
| How does CRE recombinase work? | CRE recombinase is a tool to edit DNA. it recognises loxP sequences. when these sequences are in opposite directions, CRE reverses the gene sequence. when these loxP sequences are in the same direction, CRE cuts out one of the loxP sequences, and can thereby remove the gene in between the loxP sequences. |
| What is needed/should be done before an animal experiment can be performed? | - a course certificate of an animal course. - an establishment and project licence is needed from the CCD (centrale commissie dierproeven)/article 9 WoD licence. - a master in a relevant discipline. |
| How does CRE recombinase work? | CRE recombinase is a tool to edit DNA. it recognises loxP sequences. when these sequences are in opposite directions, CRE reverses the gene sequence. when these loxP sequences are in the same direction, CRE cuts out one of the loxP sequences, and can thereby remove the gene in between the loxP sequences. |
| In which categories can animal models be divided? | 1. animals in which disease is induced, through pharmacological injection, stress induced, biological injection or lesion induction. 2. spontaneous animal models. 3. genetically modified animals. 4. negative animal models (animals that will not develop the disease, but can be used to study disease susceptibility, e.g. pigs and SARS-CoV-2). 5. healthy animals. |
| How does CRE recombinase work? | CRE recombinase is a tool to edit DNA. it recognises loxP sequences. when these sequences are in opposite directions, CRE reverses the gene sequence. when these loxP sequences are in the same direction, CRE cuts out one of the loxP sequences, and can thereby remove the gene in between the loxP sequences. |
| What are the advantages of the use of animal models? | - contribute to the understanding of pathological and biological processes. - enable development of drugs, vaccines and surgical techniques. - have a short lifespan. - can be used for animal production, to create animals that are bigger (generate more consumption meat) or produce more milk. - can be used to decrease environmental pollution (e.g. pigs that generate less phosphate). |
| How does CRE recombinase work? | CRE recombinase is a tool to edit DNA. it recognises loxP sequences. when these sequences are in opposite directions, CRE reverses the gene sequence. when these loxP sequences are in the same direction, CRE cuts out one of the loxP sequences, and can thereby remove the gene in between the loxP sequences. |
| What are the disadvantages of the use of animal models? | - high costs and time consuming. - ethics. - low translatability of findings in animals to humans. - difficult to assess behavioral and cognitive tests in animals. |
| What is the difference between active immunization and passive transfer in the generation of animal models? | In active immunization, an animal model is induced through exposure to the antigen, after which the animal develops its own antibodies to this antigen. in passive transfer, an animal model is induced through the administration of serum, purified immunoglobulins, monoclonal antibodies or antibody-producing cells, which are isolated from a diseased organism. antibodies are present directly, and this model can be used to study the acute effects of a disease. |
| How can the efficacy of a MG animal model be assessed? | - clinical score: score of animal weakness prior and after induction, scores from 0 (no weakness) through 4 (death). - measures of weakness and fatigability, through grip meter or mesh test. - electromyography: measurement of muscle action potential, and thus muscle weakness/strength. - immunofluorescence or electron microscopy of the neuromuscular junction to assess absence of AChR. - radioimmunoassay to measure AChR concentration. |
| What are recommended methods to sacrifice rodents? | - overdose of injectable agents (sodium pentobarbital) - overdose of inhalational agents (halothane) - inhalation of carbon dioxide gas (con: creates stress, increased HPA axis) - inhalation of carbon monoxide gas (con: dangerous to humans) - decapitation (con: brain activity still present after 14s) - cervical dislocation - blunt force trauma (con: skilled lab personnel needed) - focus beam microwave irradiation, heating of the rodent's brain |
| In which fields of study are different animals commonly used? | - rhesus monkeys and other nonhuman primates: emerging infectious diseases, serious (brain) diseases, life-threatening situations. - nematodes (C. elegans): metabolic diseases (esp. obesity). - zebrafish: endocrinology and metabolic diseases, neurodevelopmental conditions. - drosophila: neurobiological processes. - pigs: studying and performing animal-to-human transplants. - rodents: infection and immunity, endocrinology, metabolism, cancer, pharmacology and therapeutics, neuroscience, surgical techniques. |
| What is the definition of a transgenic animal? and what is the process of transgenesis? | Transgenic animal: an animal whose genome has been changed to carry genes from another species or to use techniques for animal genome editing for specific traits. transgenesis: introduction of foreign DNA sequences into the genome of transfected cells ensuring that the DNA sequences are integrated and transmitted to the offspring. |
| How does the DNA microinjection technique of transgenesis work? | 1. collect fertilized oocytes or embryonic stem cells (from fertilized blastocysts). 2. inject the genes of interest in the oocytes or inner cell mass of the stem cells. 3. culture the oocyte into a blastocyst or insert the cells into a blastocyst. 4. perform genetic tests, freeze the cells or transplant them into a recipient. |
| How does sperm-mediated gene transfer work? | A spermatozoon is incubated with foreign DNA, which is then used in fertilization to produce a transgenic animal. or a spermatozoon is treated with a detergent in a way that the membrane is altered so that foreign DNA can enter. this spermatozoon is then used for fertilization. |
| How does somatic cell nuclear transfer work? | Somatic cell nuclear transfer is essentially cloning: 1. one oocyte is enucleated. 2. a second enucleated oocyte is injected with isolated fibroblasts containing the gene of interest. 3. the two oocytes are merged. 4. a complete embryo is formed. |
| What is a knock out model and what has been found using knock out models? what is a knock in model? | Knock out: an animal model in which researchers have inactivated an existing gene by replacing or disrupting it with an artificial piece of DNA. it has been found that most genes are pleiotropic; they are expressed in different tissues, in different ways and at different times and that knock outs of many genes in the mouse genome can be compensated for. knock in: an animal model in which a gene sequence of interest is altered by one-for-one substitution with a transgene, or by adding gene sequences that are not found within the locus. |
| How can DNA be edited? | - via the generation of a DNA vector through homologous recombination to make sure that the vector is incorporated into the genome and random integration is prevented. - via the use of recombinases, such as the CRE recombinase - via the use of nucleases, such as CRISPR-Cas9, to induce deletions or insertions. |
| How does CRE recombinase work? | CRE recombinase is a tool to edit DNA. it recognises loxP sequences. when these sequences are in opposite directions, CRE reverses the gene sequence. when these loxP sequences are in the same direction, CRE cuts out one of the loxP sequences, and can thereby remove the gene in between the loxP sequences. |
| How to produce an animal model for McArdle disease? | 1. produce a recombination vector including the mutation. 2. make the vector into a linear DNA sequence. 3. introduce the vector into stem cells by electroporation to allow for cell permeabilization. 4. grow the cells (under neomycin selection). 5. identify the recombinant clones using PCR. 6. confirm that homologous recombination has occurred using southern blot; the lengths of the wild-type and mutant DNA should be different, as measured by a probe. 7. perform a karyotype analysis. |
| What are the general steps of mass spectrometry? | 1. ionization: the sample is ionized by a beam of electrons, because of which an electron is removed, creating cationic ions. 2. acceleration: all ions undergo acceleration through an electric or magnetic field, to have the same amounts of kinetic energy. 3. deflection: depending on the mass-to-charge ratio, the ions will undergo different amounts of deflection in the magnetic field. small ions and ions with a greater positive charge will get deflected more. 4. detection: a detector, such as an electron multiplier, will detect all cationic ions. neutral and anionic ions are removed by a vacuum. |
| What is MALDI? | MALDI means matrix assisted laser desorption ionization, which is an ion source. the principle of MALDI is as follows: 1. an organic, crystalline matrix is added in excess to the sample. 2. the sample is irradiated by a laser. 3. there is vaporization of the analyte molecules. 4. the analyte molecules are ionized. 5. single ion images are generated. |
| What is the purpose of the matrix in MALDI and which one can best be used? | The matrix layer absorbs the energy from the irradiation laser and induces desorption and ionization of the analytes. to not introduce any errors, the matrix crystal deposition should be homogenous to ensure high sensitivity, reproducibility and artefart-free imaging. for imaging proteins in the brain: 2,6-DHA matrix for imaging lipids: 1,5-DAN or CHCA. |
| What is the principle of electrospray ionization? | Electrospray ionization (ESI) is an ion source. in electrospray ionization, ions are continuously generated at atmospheric pressure by guiding a solution-based sample through a small tube, to which a voltage is connected. the sample solution is then electrostatically sprayed to generate an aerosol of charged droplets, which make their way into the mass analyser. |
| What is the principle of secondary ion mass spectrometry (SIMS)? | SIMS is an ion source that works as follows; 1. a focused primary ion beam bombards the sample with primary ions. 2. because of this, secondary ions are ejected from the sample. 3. secondary ions are collected and analysed in the mass spectrometer. SIMS is mainly used to analyse the composition of solid surfaces and thin films. the application to biological samples is limited since SIMS has insufficient sensitivity and spatial resolution. |
| What is a time-of-flight (TOF) mass analyser? | A TOF tube is a type of mass analyser. the ions that are created through an ion source are separated based on their velocity. the ions are accelerated by a certain voltage in the TOF drift tube. ions with lower m/z ratios will achieve a higher velocity and will be detected first by the detector. by measuring the time it takes to reach the detector, the m/z ratio can be determined and the ions can be identified. |
| What are advantages and disadvantages to mass spectrometry? | Advantages: high sensitivity and able to recognize small molecules, high selectivity, fast analysis, provides quantitative results, widely applicable (used to detect proteins, lipids, nucleic acids etc.), can be used for the identification of unknown molecules. disadvantages: expensive equipment, skilled personnel needed, sample preparation can be extensive and challenging (embedding and fixation may introduce artifacts), large proteins are difficult to analyse, ionization process can be destructive to sample. |
| What are advantages and disadvantages of MADLI specifically? | Advantages: very little sample is wasted, singly charged analytes are usually generated, high troughput, matrix facilitates ionization, allows for molecular mass determination of the analytes while keeping their spatial resolution in tact. disadvantages: difficult to couple to certain mass analysers due to its sensitivity, matrix causes chemical noise in samples with low molecular weights |
| How does mass spectrometry allow for protein detection? | - peptide mass fingerprinting: the measured proteolytic peptide masses are matched to the theorectical proteolytic peptide masses of the proteins. this can be done because you know where trypsin is supposed to cut. if you find peptides that are very rare, proteins are easily identified. but if you find peptides that are very common, many different peptides are needed to identify the protein. - labelling: the sample is labelled with stable isotopes, which are visualised after analysis. |
| What are different ion sources, mass analysers and ion detectors that can be used in mass spectrometry? | Ion sources: MALDI, (desorption) electrospray ionization ((D)ESI), direct analysis in real time (DART), fast atom bombardment (FAB), electron ionization (EI). mass analysers: time-of-flight (TOF), quadrupole, magnetic sector, ion trap. ion detectors: electron multiplier, faraday cup, array detectors, photomultiplier conversion dynode. |
| What can be seen in a mass spectrum? | The mass spectrum shows the intensity (y-axis) as a function of the mass-to-charge ratio (x-axis). the higher the peak of a certain m/z ratio, the higher the relative presence of the compound belonging to that peak in the sample. small molecules typically have a single charge, which means that their m/z ratio is the same as their mass. however, larger molecules often have a charge (z) greater than 1, which means that the m/z ratio is not the same as their mass. |
| What the differences between the genome and the proteome? | Genome: static, can be amplified with PCR, little sample complexity, good solubility, only 4 base pairs. proteome: dynamic (variable with time), cannot be amplified, high sample complexity, various solubility. |
| What is mass spectrometry? | MS is an analytical technique that separates ionized particles by using differences in the ratios of their charges to their respective masses and can be used to determine the molecular weight of particles. MS is used to identify molecules that are present in solids, liquids and gases, to determine the quantity of each type of molecule and to determine which atoms comprise a molecule and how they are arranged. |
| What are bottom up and top down approaches of sample analysis? what are the differences? | In the bottom up approach, intact proteins are digested into peptides prior to introduction into the mass spectrometer. this approach is very user friendly, generates a high throughput and gives more information about proteins with 'extreme' properties. but posttranslational modifications and isoform information is lost. in the top down approach, intact proteins are introduced into the mass spectrometer. this keeps posttranslational modifications and protein isoforms intact, but has limited sensitivity and throughput and quite pure samples are required and expensive instrumentation is needed. |
| What is 2D electrophoresis, why is it used in relation to mass spectrometry and what are some pros and cons? | 2DE is a technique that is used to separate proteins prior to mass spectrometry. proteins are separated along one axis based on isoelectric point (pH at which the net charge becomes zero) and along a different axis based on molecular weight. proteins of interest can then be 'cut' out and introduced into mass spectrometry. pros: reflects changes in protein expression, isoforms or posttranslational modifications, is able to separate many proteins, inexpensive. cons: time consuming, multiple proteins can remain present in one spot, only detects high abundance proteins, large proteins cannot be seen. |
| What occurs when proteins are digested? | 1. reduction with DTT to break the disulfide bonds 2. alkykation with IAM to block cysteines in proteins (if present) to prevent forming disulfide bonds again. 3. digestion with trypsin, which breaks the peptide bonds between the carboxyl group of arginine or lysine and the amino group of the adjacent amino acid. |
| What are different protein labelling techniques? | Stable isotope labelling metabolic (SILAC): label introduced in first stage of analysis, very accurate labelling. chemical labelling: an isotope coded affinity tag (ICAT) is added to cysteines. isobaric peptide tagging: labelling is done after protein extraction and denaturation (all tags have identical mass). |
| What are the differences in speed and spatial resolution between MALDI, DESI and SIMS? | The spatial resolution of SIMS is the best out of the three. then comes MALDI and then DESI. but the higher the resolution, the smaller the amount of sample that can be ionized at the time. DESI is the fastest out of the three. SIMS takes quite a lot of time, MALDI is quite fast but time is lost in preparing the matrix. |
| What are the types of optical microscopy and what are their pros and cons? | Brightfield microscopy: specimen is illuminated against bright background, used to observe stained biological samples. pros: simple, inexpensive, widespread use. cons: low contrast, limited resolution. dark field microscopy: specimen is illuminated from the side, used to observe live and stained specimens. pros: enhanced contrast, live specimen observation. cons: limited application, image interpretation. phase contrast microscopy: transparant and colorless objects can be observed. pros: no staining is required, live cell imaging. cons: complex set-up, limited to transparant specimens. confocal microscopy: a laser light is used to create a 3D image of the specimen. pros: optical sectioning, high resolution, 3D reconstruction. cons: cost and complexity, photobleaching and phototoxicity. |
| What are the two main types of electron microscopy and what are their pros and cons? | TEM (transmission electron microscopy): used to study ultra thin sections (< 100 nm) and can visualise the internal structures of cells. only samples with a limited thickness can be used and only 2D images can be generated. SEM (scanning electron microscopy): used to study the surface of solid, thick sections or objects. with this 3D images can be generated, but no internal structures can be visualised. sample preparation is also more complex. |
| What are the main differences between optical and electron microscopy? | Optical microscopy creates images in color (depending on the staining), whereas electron microscopy can only generate black/white images. however, electron microscopy can generate much more detailed and magnified images with a better resolution. |
| What are the pros and cons of optical microscopy? | Pros: affordable, easy to operate, portable, live samples can be observed, do not require radiation, minor maintenance (costs), preparation is quick and easy, natural color of the sample can be observed. cons: poor resolution at high magnification, low magnifying quality, cannot provide 3D images, staining is often needed, sample thickness is a limiting factor. |
| What are the pros and cons of electron microscopy? | Pros: great resolution (up to 0.2nm), great magnification, TEM is able to show the insides of cells clearly, SEM allows surface structures to be seen clearly, high quality images can be generated. cons: expensive, large, affected by magnetic fields, preparation of material is extensive and requires expertise, preparation may distort the sample, only dead specimens can be visualised, only black/white images can be created. |
| What is the difference between specific and non-specific staining? | Specific stainings are stainings that make use of (primary and secondary) antibodies coupled to fluorophores or enzymes. non-specific stainings are stainings that bind to all pieces of double stranded DNA and result in fluorescence. an example is SYBR green. |
| What are examples of bacterial, human and biological stains? | Bacterial stains: gram staining, endospore staining, Ziehl-Neelsen staining. human stains: haematoxylin and eosin, papanicolaou (PAP), periodic acid Schiff (PAS), silver stain. biological stains: acridine orange, DAPI, Hoechst. |
| What is OCT and what are the pros and cons of using it? | OCT (optimal cutting temperature) compound is used to preserve the structure of a sample when it is frozen and during cryosectioning. OCT is water soluble and does not penetrate the tissue, meaning it can be washed away during sample preparation. pros: preparation is quick, sample shows little autofluorescence and is sensitive to immunohistochemistry, does not require fixed tissue, allows for longitudinal studies. cons: tissue degradation over time, requires rapid processing, loss of cellular viability. |
| What is paraffin embedding and what are the pros and cons of it? | Paraffin embedding is done after the tissue is fixed (in formalin) and permeabilized. pros: better preservation of tissue architecture, long-term storage, easily obtainable for research, compatible with IHC, cost-effective, sections are physically stable. cons: preparation time takes about 2 days, shows a lot of autofluorescence, degradation of DNA and RNA, long processing time. |
| What is multiplex immunofluorescent imaging? | In multiplex immunofluorescent imaging, up to 16 different markers can be used to bind to and visualise different molecules in a single section. this can be done if markers with different excitation and emission wavelengths are chosen. |
| What are problems that can occur in (multiplex) immunofluorescent imaging and how can they be solved? | Autofluorescence; by placing the tissue in a dark, UV-lit box, which removes the autofluorescence. spectral overlap (the overlap between the emissions of the different fluorophores); this is solved by using multispectral microscopy, in which you stepwise scan the visible spectrum. this generates separate pixels for each fluorophore per scan. these pixels can be separated and you can determine from which fluorophore the pixel comes from. |
| What are the cycles of PCR? | 0. initial denaturation: uncoiling the helical structure of DNA. 1. denaturation: separation of dsDNA strands by heat (95 degrees). 2. annealing: attachment of primers at 50 to 65 degrees. 3. extension: synthesis of new DNA by DNA polymerase and nucleoside triphosphates at 72 degrees. 4. final extension |
| How do the Taqman probe and SYBR green fluorophore work in qPCR? | Taqman probe: under normal conditions, the probe is coiled up, meaning that fluorescein is near the quencher of the probe, limiting the fluorescent signal. when the target sequence is found, the probe will uncoil and bind to the sequence. as new DNA is created, the probe starts being degraded, which means that fluorescein is separated from the quencher and a fluorescent signal is shown. this is very specific. SYBR green: SYBR green dye binds to the minor groove of the DNA non-specifically and emits fluorescence. this provides information on each cycle of amplification and about the melting temperature. |
| What are the differences between PCR and qPCR? | PCR is qualitative, while qPCR is quantitative. if no detection method (e.g. gel electrophoresis) is done for PCR, there is no visual output. in qPCR, a fluorphore is added to the sample and the reaction is carried out in a thermal cycler which illuminates the sample with a light beam of a specific wavelength. in qPCR, the sequence copy number can be detected in real time. |
| What are the pros and cons of PCR? | Pros: sensitive, specific, can analyse RNA and DNA, widely applicable, fast. cons: susceptible to contamination from other DNA or RNA sources, no novel information can be generated, requires expert knowlegde, base substitutions and other mutations can lead to inaccurate amplifications, qualitative results only. |
| What are the pros and cons of qPCR? | Pros: faster than PCR, real-time monitoring, efficiency of reaction can be precisely calculated, quantitative analysis, high sensitivity and specificity, widely applicable, high throughput. cons: more expensive than PCR, quality and quantity can affect results, can produce false negatives if target sequence is only present at low concentrations, no novel information, limited to detection of DNA and RNA. |
| What is the purpose of RNA sequencing? | RNA seq is a sequencing technique that determines the presence of RNA in a biological sample and quantifies this amount. it can be used to analyse the transcriptome (the complete set of transcripts in a cell). RNA seq is used to give more insight into alternative splicing, mapping genes, mapping exon boundaries and to identify novel transcribed regions. |
| What is the process of RNA sequencing? | 1. RNA isolation from sample. 2. isolate specific RNA species you want to analyse, e.g. isolate miRNAs through size selection or other RNAs through poly-A selection. 3. conversion into cDNA. 4. add adaptors to both ends of cDNA fragments. 5. align the fragments with the reference genome. 6. classify reads as either exonic reads, junction reads and poly(A) end-reads. 7. the result is quantified by counting the number of reads that align to each gene using a computer program. |
| What are the pros and cons of RNA sequencing? | Pros: high resolution (up to single bases), high throughput, low background noise, able to differentiate isoforms, able to distinguish allelic expression variants, required amount of RNA is low, can reveal SNPs or other mutations, can be used to identify new genomic sequences (novel information), can be used to map complex transcriptomes. cons: expensive (but relatively low cost for large mapping large transcriptomes), difficult to store, retrieve and process large amounts of data, time-consuming, reverse transcripton needed, complex to analyse. |
| What is RNA interference and what is its purpose? | RNAi is a phenomenon in which small pieces of RNA can shut down protein translation by binding to mRNAs that code for those proteins. it is a process ocurring naturally. its main application is gene silencing. this can be used to silence genes in certain diseases, to inibit viral RNA and prevent viral replication, to silence cancerous genes thereby preventing tumour growth or to enhance traits in crops and plants. it can also be used to create knock down models to identify functions of certain genes. |
| What is the procedure of RNA interference? | 1. the enzyme Dicer cleaves the long dsRNAs into short double-stranded fragments of 21-23 nucleotides, called small interfering RNAs (siRNAs). 2. each siRNAs is unwound into two single-stranded RNAs; the sense and antisense strand. 3. the sense strand is cleaved by the protein argonaute 2 (Ago2) and degraded, the antisense strand is incorporated into the RNA-induced silencing complex (RISC). 4. RISC then binds and degrades the target mRNA (that is complementary to the antisense ssRNA strand) through cleavage by Ago2. silencing of the gene can be confirmed by (q)PCR. |
| What is the principle of transitive RNA interference? | It is the movement of a silencing signal along a particular gene. it is the ability of the RNA interference to spread through certain organisms. it has been shown that this is possible in plants and the C. elegans. |
| What are the pros and cons of RNA interference? | Pros: high specificity, low concentrations of dsRNA/siRNA needed, effective gene suppresion, widely applicable and therapeutic potential, siRNAs can be created exogenously and made to target specific mRNAs. cons: competition with endogenous RNAs (in case of exogenous administration), stimulation of the innate immune system, silencing of off-target genes which can create a toxic phenotype. |
| What is the method of Sanger sequencing? | In Sanger sequencing, the sequencing reaction takes place in four different tubes. each tube contains a primer, dNTPs and a DNA polymerase. each tube also contains a different radiolabelled dideoxy nucleotide (either ddATP, ddTTP, ddCTP or ddGTP) (at a very low concentration). when this ddNTPs is incorporated into the sequence, the reaction stop. the incorporation of the ddNTP is a random process. this means that the mixture will contain several DNA fragments of different lengths. when the length of the fragment is known (through gel electrophoresis), the location of the radiolabelled ddNTP can be estimated and the entire strand can be build. |
| What is the method of next generation sequencing (NGS)? | NGS is also called second generation sequencing. 1. isolate DNA and generate smaller fragments. 2. DNA fragments are coupled to different adaptors on both ends. 3. DNA fragment strands are then denatured. 4. separate DNA strands then bind with their adaptors to complementary sequences on the flow cell. 5. DNA is then amplified through PCR (while on the flow cell). 6. DNA is then denatured again and the strand that is not attached to the flow cell is washed away. 7. bridges are formed through the binding of the second adaptor to the cell. 8. bridges are amplified and both strands will stick to the flow cell. 9. bridge formation and bridge amplification is repeated several time to generate many copies. 10. once amplification is done, the reverse strand is cleaved and sequencing can start (the primer binds to the adaptors). 11. fluorescently labelled nucleotides are added and when the complementary nucleotide binds, it is excited by a laser and the fluorescent signal is obtained and the nucleotide is identified. |
| When would you perform Sanger sequencing? when would you perform next generation sequencing? | Sanger sequencing is nowadays only done when you know what you are expecting in a locus with very low diversity. in this case you have limited resources and only need a small result, and this will be cost effective. NGS is the gold standard for human DNA sequencing in diagnostics. it is low cost, very fast (compared to Sanger) and allows for multiplexing; different samples can be mixed and sequenced. |
| What are pros and cons of NGS? | Pros: generates data from hundreds of sequences simultaneously, more sensitive to low frequency variations, little amount of DNA needed (because amplification is done), fast (human genome can be sequenced overnight), allows for multiplexing, cost effective, high reproducibility, a priori knowledgde of the genome is not required (adaptors do not bind complementary). cons: large infrastructure needed (sequencer, computer, storage, expertise), cheap per base but it generates so many bases that it is overall still expensive. |
| What are examples of third generation sequencing techniques? | PacBio: wells that have a DNA polymerase attached to their bottom are used. fluorescence nucleotides are added to the wells and the unknown DNA strand is extended by a polymerase. each time a fluorecent nucleotide is added, a signal is emitted. while the strand is elongated, you know the sequence. Oxford nanopore: native DNA is pulled through a very small channel, for each nucleotide that exists the channel, a little current flows through and is detected. each nucleotide will produce a different current, by which the nucleotide can be identified. |
| What are advantages and disadvantages of third generation sequencing techniques over NGS and Sanger sequencing? | Advantages - no amplification is needed, which prevents PCR bias. - haplotying can be done; the determination of haplotypes (DNA variants of a single chromosome that are inherited together) from unordered DNA. - can generate very long sequences (200kb). disadvantages - higher error rate. - higher cost per kb compared to NGS. |
| What are the different forms of DNA damage; depurination, deamination and alkylation? | Depurination: removal of the purine base (either guanine or adenine) of DNA deamination: the removal of the amino group of the DNA base alkylation: the addition of an alkyl group to the DNA, which has implications for DNA pairing (this the not the same as methylation, in which methyl groups are added to the sugar backbone of the DNA) |
| What are errors that can be made in the DNA? what are the results of these errors? what is their repair mechanism? | - replication errors > base mismatch and bulges > mismatch repair (MMR) - ROS > single strand breaks (SSB) > base excision repair (BER) - oxidation, alkylation, hydrolysis > single base damage > base excision repair - ionizing radiation > double strand breaks (DSB) > non homologous endjoining (NHEJ) and homology directed repair (HDR) - chemotherapeutics > interstrand crosslinks > non homologous endjoining (NHEJ), homology directed repair (HDR) and nucleotide excision repair (NER) - UV light, radicals > intrastrand crosslinks and bulky adducts > nucleotide excision repair (NER) |
| How are single-stranded breaks, double-stranded breaks, DNA adducts (crosslinks and oxidized bases) and mutations repaired? | SSBs: base excision repair (BER) DSBs: non homologous endjoining (NHEJ) or homologous recombination (HR) DNA adducts, crosslinks and oxidized bases: nucleotide excision repair (NER) mutation: mismatch repair (MMR) |
| What were the precursors of Crispr/Cas9 and what were their disadvantages? | Meganucleases; only bind to a very specific DNA stretch and is only useful if this specific stretch is your gene of interest, cannot be targeted. zinc finger nucleases; can be targeted, but can only target specific sites (e.g. only 1000 genes can be targeted). TALEN; can be designed to any target site of the genome, but they are not very efficient and it takes a long time (weeks/months). |
| How does Crispr/Cas9 work? | 1. identify the sequence that has to be edited in the genome. 2. create a guide RNA that is complementary to the target sequence. 3. attach the guide RNA to Cas9 (a DNA cutting enzyme). 4. introduce the complex into the target cells. 5. the complex will recognise the target sequence and cut the DNA. 6. the cell will try and repair these breaks by non homologous endjoining. 7. NHEJ often makes mistakes, when these are not removed, the gene is knocked out. 8. when you want to create a knock in, you have to include a donor sequence that is complementary to the target site, except for the mutation that you want to introduce. for this process the repair mechanism of homologous recombination is needed. |