Midterm No. 1 Lab Techniques Flashcards
What does an SDS-PAGE / 1D gel tell us?
Shows the approximate molecular weight of a protein of interest
Steps to an SDS-PAGE / 1D gel
- SDS ionic deterget is used to make the proteins negatively charged over all surface area
- A reducing agent (mercapethanol) adds hydrogen to the proteins to break disulfide bonds and other interactions
- Proteins are heated to further denature
- Samples rune on the PAGE, a porous jelly gel. A ladder with known weights and size is run parallel to contextualize the samples
What is SDS and what is it used for?
It’s the ionic detergent. It makes proteins negatively charged over their entire surface area
What is the purpose of the reducing agent in the SDS-PAGE 1D gel?
It adds hydrogens to the proteins to break disulfide bonds and other interactions
What experimental questions (aside from determining approx. molecular weight!) can an SDS-PAGE 1D gel be used for?
How long until the protein of interest is expressed in cell extracts after I introduced it?
Did the protein get heavier or lighter after modifications?
Has the protein been degraded?
What is an Isoelectric Focusing 1D gel used to tell us?
Tells us the overall charge of a protein from its collection of amino acids, whether its basic or acidic
What charge do proteins have at low pH?
Positive
Because they tend to be negatively charged (they still have a unique charge, but most tend towards negative), they snatch up the excessive positive charges (H+) from the environment
What charge do proteins have at high pH?
Negative
Proteins tend to usually be negatively charged. At high pH they will still be negatively charged because there’s not enough H+ to go around
Steps to an Isoelectric Focusing 1D test
- Hook a pH gel to an electric field. This will force protein movement to their isoelectric points
- Load samples and run the gel
- Based on the proteins isoelectric points, we can determine their overall charge and whether they are acidic or basic
What test should you run to determine wheter a protein has become phosphorylated?
Isoelectric focusing 1D gel
Also possibly a Western Blot + 2D gel
Phosphorylation increases negative charge, affecting acidity
Steps to an SDS + Isoelectric 2D gel
- Use an isoelectric pH gel strip to separate a protein mixture by charge
- Soak the gel strip with SDS and a reducing agent. This strip will now act as wells for the electrophoresis
- This strip will now act as wells for the electrophoresis. Run the gel to separate proteins by weight
What can Western Blots NOT be used for?
To directly discover a never before seen protein
This is because Western Blots run off of antibody recognition, and in order to make the antibodies specific we need to already know the target protein
What is the job of the primary antibody in a Western Blot?
To bind to a specific protein of interest
What is the job of the secondary antibody in a Western Blot?
To bind to the primary antibody. It’s equipped with a synthetic fluorescent marker or some other visual tag
What questions does a Western Blot + SDS PAGE gel answer?
The antibodies from the Blot and the gel from the SDS PAGE gel allow us to see if our target protein is present in our sample mix
Also used to determine concentration levels. Dark bands indicate high levels, light bands indicate low levels
What questions can a Western Blot + 2D gel answer?
Did my specific protein get phosphorylated?
Is my specific protein present after a certain mutation?
What is immunofluorescence + colocalization used for?
Immunofluorescence uses fluorescently tagged antibodies to bind to proteins within a living cell. The localizations of the antibody+proteins can be seen under a microscope
Using two different fluorescence colors on two different antibodies allow us to see points of overlap, aka colocalization
What is immunoprecipitation used for?
To extract proteins from a mix
Steps to immunoprecipitation
- Antibodies specific to a protein of interest are added to a mix
- Non-bound proteins are removed
- Antibodies are removed
What is Co-Immunoprecipitation used for?
To show if two proteins actually interact
Does my target antigen bind to or associate with other proteins? Is it in a complex with other proteins?
Steps to Co-Immunoprecipitation
- Use an antibody for a known protein
- The antibody-bound protein interacts with other unknown proteins in a larger complex
- Chemical solvents or antibody breads help remove the anitobdy and whatever is attached to it from the protein mix
- Unknown proteins + known proteins are precipitated out
- Bonus: other gel techniques can then be used to learn more about the unknown proteins
Polyclonal antibodies
Several different varieties of antibodies that bind to one antigen
Made in animals when more than one B cell responds to a particular antigen
Monoclonal antibodies
Single antibody species
Harvested from myeloma cancer cultures
Pros of polyclonal antibodies
Cheap
Creates more than one variety of antibody for the same antigen
The different varieties may bind to different areas of the target antigen
The antibodies, due to their diversity, are more tolerant of small changes in protein structure, heat, shock, etc to the antigen
Cons of polyclonal antibodies
Finite supply (the animal making them will eventually die)
Pros to monoclonal antibodies
Infinite supply
Single antibody species, highly specific
Cons to monoclonal antibodies
Not cheap to make
Will bind to only one specific site on the antigen
Antibody isn’t tolerant of small changes in the target antigen
Why are co-immunoprecipitation experiments better than co-immunolocalization for showing if two proteins actually interact?
Co-immunolocalization only shows if two proteins are in the same area
Co-immunoprecipitation shows if they actually interact
Steps to X-Ray crystallography
- Obtain a protein of interest
- Purify the protein using fast protein liquid chromatography
- Crystallize the protein using a vapor diffusion technique. All the molecules will be in the same orientation, thus forming a crystal and not an aggregate
- Collect data from an x-ray diffraction experiment. The crystal must be super cold while subjected to the x-rays
- Structure determination. The X-ray diffraction pattern is analyzed to yield an electron density map. The alignment of the electron densities are compared to the known primary sequence, and the protein model is generated
How do researchers determine the structures of proteins and protein complexes?
X-ray crystallography
Cryoelectron microscopy
NMR
What does PAGE stand for?
Polyacrylamide gel electrophoresis
Steps to cryoelectron microscopy
Start with purified protein/protein complex in solution
Dilute the purified sample
Disperse the diluted solution onto an electron microscopy grid
Freeze the grid with liquid helium and allow shapes to form
Determine structure by observing the shapes from their different angles, relative to the known primary sequence
Compare x-ray crystallography to cryoelectron microscopy
In both x-ray crystallography and cryoelectron microscopy, the protein of interest’s primary sequence must be known beforehand in order to interpret any meaningful results. The electron density map and shapes on the EM grid aren’t very useful on their own. They need to be contextualized with the known primary sequence.
You’re trying to run an SDS PAGE gel. What would happen if you forgot the SDS?
The SDS is the ionic detergent. Without it the proteins wouldn’t be coated with the negative charge and wouldn’t be kept apart, resulting in one large glob of proteins in the gel matrix because they couldn’t be separated by their molecular weight
You’re trying to run an SDS PAGE gel. What would happen if you forgot the beta mercaptoethanol?
The disulfide bonds stabilizing the proteins’ tertiary and quaternary structures wouldn’t be broken, resulting in an inaccurate sorting of proteins in the gel matrix as the proteins’ true molecular weights wouldn’t be clear (quaternary structures still together would make all proteins involved seem heavier than they are)
You’re trying to run an SDS PAGE gel. What would happen if you forgot to heat the samples to 95 C?
The proteins would not denature/unfold, and wouldn’t be able to properly sort themselves through the gel matrix
You’re trying to run an SDS PAGE gel. What would happen if you forgot to run a lane of molecular weight standards?
You’d get accurate results from your experimental lanes, but you wouldn’t be able to interpret or contextualize your results. Your experiment would be useless
Where does the positive electrode go in a gel?
For the gel to work, the positive electrode will need to go at the bottom of the gel so that the negatively charged proteins will be attracted to it and run through the gel. If the positive electrode was at the top of the gel, the negatively charged proteins would run towards the top and out of the lanes, ruining the experiment.
Steps to 2D PAGE gel
Separate proteins by their native charge using isoelectric focusing
Individual proteins will move across a charged pH gradient to their electrically neutral state, aka their isoelectric point. Different proteins have different isoelectric points, are neutral at different pHs
Incubate a tube gel with an existing stable pH gradient in the protein solution/cell lysate. This forms the first dimension
Align the isoelectrically separated tube with the negative end of an SDS PAGE gel to form the second dimension
Does it matter which method is applied first in 2D PAGE gel tests?
The end result is a 2D separation, in which proteins are separated by both their native charge AND their molecular weight. The order in which these tests are run is vital, as the SDS PAGE gel only works with the tube gel’s input, while the tube gel cannot work with the SDS PAGE gel input
Western Blot steps
Load and separate protein samples onto an SDS PAGE gel
Electrophoretically transfer the subsequent fractionated proteins onto a PVDF membrane
Block the membrane with a neutral protein, usually milk/casein powder
Incubate the membrane with the primary antibody specific to your target protein
Incubate the membrane with an HRP-labeled secondary antibody specific to the primary antibody
Incubate the blot with chemiluminescence
Steps to a DNA footprint assay
Cloned gene fragment is amplified and is radioactively labeled at one end, then incubated in samples with and without its cell extract (RTFs)
The genes are then chopped with DNase after incubation. The sample cell without the cell extract is chopped the full gradient length, while the sample with the cell extract is missing a section of the chop gradient
Fragments are run on a gel
The gel is layered with x-ray film so that only the radioactive fragments are shown
The missing chunk in the sample with cell extract shows a footprint where the RTF binds, as the DNase was blocked by the RTF
Steps to a gel shift assay
Tool used to identify RTFs and the DNA elements they bind to
A DNA fragment of interest is fractioned by some metric (weight, charge, etc), then mixed with RTF/cell extracts
The DNA is then bound to a radioactive probe
The samples are run on a gel, then overlaid with an x-ray film so that only the radioactive samples are shown
DNA bound to a RTF protein won’t travel as far as free DNA. The x-ray film looks for free probes vs bound probes
How is binding strength determined?
Use stress tests.
Ex: increase thermal energy to make it harder for proteins to bind
Ex: decrease concentration of reactants to decrease the ratio of AB dimers`
How do we know which cyclin types are present at which times and phases of the cell cycle?
Work with Western Blots
How did we learn about the different levels of DNA packaging?
We learned about histones and nucleosomes and their “beads on a string” characteristics by using transmission electron microscopy
We learned how long the strands wrapped around each nucleosome are through nuclease digestion. The nucleases chewed up the DNA but left the histones intact, and when treated with salt the DNA was released and the average number of base pairs per octamer was revealed (it’s ~147, with ~50 bps making up the linker strand, so a total of ~200 bp per nucleosome)
What is RNAseq and how is it used?
Comparison of gene expression in different cells or in different conditions
Asks how cells can change their patterns of gene expression in response to changed conditions
Reverse transcriptase transcribes mRNA molecules into DNA, the DNA is then sequenced
This technique shows both wha’ts being transcribed and relative transcription levels, i.e. how much mRNA is being produced from a single gene
This data is graphed via Log2 FoldChange
