Block A Lecture 3: Protein Detection Flashcards
How can antibodies help a protein be isolated, quantified or visualised?
As the specificity of antibodies allow them to tag their specific target protein to allow it to be isolated, quantified or visualised
(Slide 7)
What is an epitope?
a small part of a protein molecule that triggers an immune response
(Slide 7)
What does the specificity of an antibody-antigen arise from?
It’s due to the complimentary shape between the 2 surfaces
(Slide 8)
Where does the antigen bind on an antibody?
The Fab domain
(Slide 9)
Do antigens have 1 or several epitopes?
They can have either, but most antibodies have several
(Slide 7)
What is the difference between polyclonal and monoclonal antibodies?
Polyclonal antibodies are heterogenous mixtures of antibodies, with each one being specific for one of the various epitopes on an antigen
Monoclonal antibodies are all identical and are produced by clones of a single antibody-producing cell. They recognise one specific epitope
(Slide 10)
How are monoclonal antibodies prepared?
Hybridoma cells are formed by the fusion of antibody-producing cells and myeloma cells.
The hybrid cells are then allowed to proliferate by growing them in selective medium.
They are then screened to determine which ones produce the antibody of the desired specificity
(Slide 11)
How can antibodies be used to quantify the amount of a protein or other antigen present in a biological sample?
As they can be used as specific analytic reagents in an enzyme-linked immunosorbent assay (ELISA)
(Slide 12)
What are 2 different types of ELISA?
Indirect and sandwich ELISA
(Slide 12)
What are the steps of an indirect ELISA?
- Well needs to be coated with antigen for the ELISA to work
- Wash
- The specific body which binds to the antigen is added
- Wash
- An enzyme-linked antibody is added, which then binds to the specific antibody
- Wash
- The substrate is added and converted by the enzyme into a colored product, with the rate of color formation being proportional to the amount of specific antibody
(Slide 12)
What are the steps of a sandwich ELISA?
- Well needs to be coated with a monoclonal antibody for the ELISA to work
- Wash
- The antigen is added and binds to the antibody
- Wash
- A second monoclonal antibody, which is linked to the enzyme is added and binds to the immobilised antigen
- Wash
- The substrate is added and converted by the enzyme into a colored product, with the rate of color formation being proportional to the amount of specific antibody
(Slide 12)
Why are electrophoretic separations nearly always carried out in porous gels?
As the gel serves as a molecular sieve which enhances separation.
(Slide 13)
What are 2 common stains which gels can have in electrophoresis?
Silver nitrate or coomassie blue
(Slide 13)
What are the steps of electrophoresis?
- Gel is mixed with buffer solution and put onto a tray
- A comb is then inserted to create wells for samples and the gel is left to solidify
- DNA or RNA samples are mixed with loading dye to track migration and increase density for proper well loading whereas protein samples are Mixed with SDS (sodium dodecyl sulfate) and a reducing agent (e.g., β-mercaptoethanol) for denaturation if using SDS-PAGE
- The solidified gel is placed into an electrophoresis chamber filled with buffer, samples are pipetted into the wells and a molecular weight marker (known as a ladder) is added for reference
- The chamber is connected to a power supply and an electric field is applied.
- Negatively charged molecules gravitate towards the positive electrode and vice versa and molecules separate via size, with smaller molecules moving slower through the gel.
- Samples are stained and proteins are transferred to a membrane for further antibody detection (if using western blotting)
(Slide 13)
What does western blotting permit?
The detection of proteins which have been separated by gel electrophoresis
(Slide 14)
What are the steps of western blotting?
- A sample is subjected to electrophoresis on an SDS-polyacrylamide gel.
- A polymer sheet is then pressed against the gel, which transfers the resolved proteins on the gel to the sheet, making proteins more accessible for reaction.
- An antibody which is specific for the protein of interest (known as the primary antibody) is added to the sheet, and reacts with the protein antigen.
- Wash
- A 2nd antibody (known as the secondary antibody) is added which is specific for the primary antibody is added, to allow detection of the antibody-antigen complex.
- Wash
- Usually the secondary antibody is then fused to an enzyme which produces a chemiluminescent (colored) product or which contains a fluorescent tag, which enables the identification and quantification of the protein of interest.
(Slide 14)
What are 3 examples of uses for western blotting?
Used to test for infection of hepatitis C
Monitoring protein purification
Cloning of genes
(Slide 14)
What is able to produce a higher resolution, electron microscopy or light microscopy?
Electron microscopy
(Slide 17)
How is light focussed onto the specimen by a microscope?
Via lenses in the condenser. A combination of objective, tube and eyepiece lenses are arranged to focus on the imaged of the specimen in the eye of the microscope.
(Slide 18)
What are condenser, objective, tube and eyepiece lenses?
Condenser lenses: These lenses focus light onto the specimen. They help direct the light in such a way that it illuminates the specimen evenly.
Objective lenses: These are positioned closest to the specimen and are responsible for gathering light from the specimen and forming an initial magnified image.
Tube lenses: These work in conduction with objective lenses to focus the image and transmit it through the microscope.
Eyepiece lenses: The eyepiece further magnifies the image formed by the objective lens and allows you to view it through the eyepiece
(Slide 18)
What are 4 examples of microscopies which can be obtained by most modern microscopes by interchanging optical components?
Bright-field microscopy
Phase-contrast microscopy
Differential-interference-contrast microscopy
Dark-field microscopy
(Slide 19)
What are bright-field, phase-contrast, differential-interference-contrast and dark-field microscopy?
Bright-field microscopy: Where light is transmitted directly through the sample.
Phase-contrast microscopy: Where phase alterations of light transmitted through the specimen are translated into brightness changes
Differential-interference-contrast microscopy: Edges where there is a steep change of refractive index are highlighted
Dark-field microscopy: In which the specimen is lit from the side and only scattered light is seen
(Slide 19)
What is refractive index?
The ratio of the apparent speed of light in the air or vacuum to the speed in the medium
(Slide 19)
What is in situ hybridisation?
With in situ meaning “in its original place”, in situ hybridisation is a technique used to to detect specific nucleic acid sequences within a tissue or cell sample, allowing scientists to examine the location and expression of specific genes or RNA molecules within their natural biological context.
(Slide 20)
What are the steps of in situ hybridisation?
- The tissue or cells are fixed to preserve their structure and the nucleic acids. This is typically done by using formaldehyde or other fixatives.
- A probe is designed, which is a short sequence of nucleotides that is complementary to the target sequence of interest. The probe can be labelled with a detectable marker, such as a radioactive isotope, fluorescent dye, or an enzyme.
- The probe is applied to the sample, where it binds (hybridizes) to its complementary nucleic acid sequence. The hybridization occurs at a specific location within the tissue or cells where the target sequence is present.
- Washing
- The labelled probe is detected using various methods depending on the type of label used.
- The position and abundance of the hybridized probe reveal the location and relative expression of the target sequence in the tissue or cells.
(Slide 20)
What are the steps involved in indirect immunocytochemistry?
- Cells are fixed (e.g., with formaldehyde) to preserve their structure.
- A primary antibody (such as a rabbit antibody) binds specifically to the target antigen in the cell
- A secondary antibody (anti-rabbit here) which is conjugated to a marker is directed against the primary antibody
- The marker allows visualization under a fluorescence microscope or through enzymatic color reactions.
(Slide 21)
Why is indirect immunocytochemistry very sensitive?
As many molecules of the secondary antibody recognise each primary antibody, allowing the signal to be amplified
(Slide 21)
What are 3 examples of commonly used markers in indirect immunocytochemistry?
Answers Include:
Fluorescent probes (for fluorescence microscopy
Horseradish peroxidase enzyme (for conventional light microscopy or electron microscopy)
Colloidal gold spheres (for electron microscopy)
Alkaline phosphatase or peroxidase enzymes (for biochemical detection)
(Slide 21)
What is the theory of fluorochromes?
That an orbital electron of a fluorochrome molecule can be raised to an excited state following the absorption of a photon. Fluorescence occurs when the electron returns to its ground state and emits a photon of light at a longer wavelength.
(Slide 23)
What can destroy a fluorochrome molecule and what is this process called?
Too much exposure to light, or too bright a light, can also destroy the fluorochrome molecule in a process called photobleaching.
(Slide 23)
Why is there a difference between excitation and emission peaks of fluorochromes?
As the photon emitted by a fluorescent molecule is necessarily of lower energy (aka longer wavelength) than the absorbed proton
(Slide 24)
What is DAPI?
It is a general fluorescent DNA probe which absorbs ultraviolet light and fluoresces bright blue
(Slide 24)
What does the structure of a fluorescent microscope consist of?
An excitation filter, emission filter, objective lens and a dichroic (beam-splitting) mirror
(Slide 26)
What does the excitation filter in a fluorescent microscope structure?
It lets through a specific wavelength of light which ensures only the designated wavelength reaches the sample.
(Slide 26)
What does the dichroic (beam-splitting) mirror do in a fluorescent microscope?
It .reflects excitation light toward the sample but allows emitted fluorescence to pass through (e.g reflects light below 510 nm but transmits light above 510 nm)
(Slide 26)
What does the objective lens do in a fluorescent microscope?
It magnifies the sample and collects the emitted fluorescence
(Slide 26)
What does the emission filter do in a fluorescent microscope?
it removes residual excitation light and allows only the emitted fluorescence to pass through, ensuring only specific fluorescence signals are detected.
(Slide 26)
How can multiple different fluorescent probes be used?
In order to visualize different structures or molecules within the same sample.
(Slide 28)
What is green fluorescent protein (GFP)?
A protein which emits green light when stimulated with blue light. It can be produced by jellyfish
(Slide 29)
What mutants of green florescent protein (GFP) have been engineered?
Mutants which can absorb and emit light over different parts of the visible spectrum, with scientists now having a protein which each colour.
(Slide 33)
How are fluorescent proteins used in FRET?
2 proteins are produced as fusion proteins attached to different colour variants of green fluorescent protein (GFP).
If the 2 proteins don’t interact, illuminating the sample with light which triggers the first protein (protein X), will only result in fluorescence from that protein only.
If the proteins interact, the resonance energy transfer (FRET) can occur. Illuminating the sample with light which triggers the first protein (protein x) will excite that, as normal, but it will now also transfer its energy to the other protein (protein Y) which results in protein Y emitting its coloured light rather than protein X emitting its.
(Slide 34)
What does FRET require?
It requires the fluorochromes to be very close together, ~ 1-5nm apart.
(Slide 34)
Why may some light from the first protein (protein X) still be detected, even if FRET occurs?
As not every molecule of protein X and protein Y are bound to each other at all time
(Slide 34)
What trend is noticed in FRET as the proteins are oved closer together / begin to interact?
Emission from the donor fluorescent protein decreases as mission from the acceptor fluorescent protein increases
Note: The pictures on these slides (especially 35) may be the best I’ve ever seen at explaining a concept ever
(Slides 34 and 35)
What is photoactivation?
The light-induced activation of an inert (chemically inactive) molecule at an active site.
(Slide 36)
How can photoactivation of fluorescent protein be used to monitor trafficking, turnover and degradative pathways of proteins?
The fluorescent protein is inserted into a certain selected target region. Once the protein becomes activated via photoactivation, the movement of the protein as it diffuses out of the selected region, can be measured, allowing us to monitor how many times a pathway etc is activated.
(Slide 36)
How can we monitor recovery using photobleaching?
A strong laser light will extinguish (bleach) the fluorescence of green florescent protein (GFP). By selectively photobleaching a set of fluorescently tagged protein molecules within a defined region of a cell, we can monitor recovery over time as the remaining fluorescent molecules move into the bleached region.
(Slide 37)
