proteomics

proteomics

focusing on the 1.5% of the genome that’s going to code for proteins to figure out when and where these protein are made

what is 1 SDS-PAGE used for

it separates proteins based on molecular weight after denaturation

why do protein gels run vertically instead of horizontally like DNA/RNA gels

gel composition

why are proteins denatured before SDS-PAGE

to linearize them and eliminate effects of shape and subunit interactions (like disulfide bridges)

what does SDS do to proteins

denatures protein to make them linear and gives all proteins a uniform negative charge (same charge to length ratio) so migration depends only on molecular weight

how do proteins move in SDS-PAGE

negatively charged proteins migrate toward the positive pole, smaller proteins move faster through the gel pores and larger move slower

how are proteins visualized after SDS-PAGE

using coomassie blue stain which binds to positively charged amino acids (arginine, lysine, histidine)

what is the limitation of 1D SDS-PAGE

proteins with the same molecular weight will appear as one band even if multiple proteins are present

what reagent is used to break disulfide bonds before SDS-PAGE

B-mercaptoethanol breaks disulfide bridges to fully denature proteins

why do proteins migrate based only on size in SDS-PAGE

Because SDS gives every protein the same charge-to-mass ratio, one SDS molecule binds about 2 amino acids, overriding native charges

2D-PAGE

separates proteins based on 2 properties: isoelectric point (pl) and molecular weight

what happens during the first dimension (IEF)

proteins are separated based on their isoelectric point (pl), the pH at which their net charge is zero. A pH gradient is established using ampholytes and proteins migrate through the gradient until they reach the pH where they are electrically neutral

why can’t SDS be added before the first dimensions

SDS would give all proteins a uniform negative charge, preventing separation by charge. IEF requires proteins in their native state with both positive and negative residues 

what happens during the second dimension

the IEF gel (usually a tube gel) is placed on top of an SDS-PAGE gel and proteins are now coated with SDS so denature and separate by molecular weight

what does the final result of 2D SDS-PAGE look like

instead of bands, you get spots where each spot = one unique protein defined by its pI and molecular weight

How can 2D SDS-PAGE be used in disease research?

you can compare protein extracts like Alzheimer’s tissue and identify spots that appear or disappear, showing disease-specific proteins.

after finding a unique spot how can it be identified

By sequencing its amino acids ( Edman degradation), but this method is slow, expensive, and limited to the first ~10 amino acids from the N-terminus.

why can’t all proteins be sequenced with edman degradation

some proteins have blocked N-terminus preventing sequencing from start

Matrix associated laser desorption/ionization-time of flight (MALDI-TOF)

used to identify and analyze proteins or peptides by separating them based on their mass to charge ratio

how is the sample prepared in MALDI-TOF

the protein or peptide is crystallized in a matrix, the matrix absorbs laser energy and helps ionize the peptides

what happens when the laser hits the matrix

the laser ionizes the matrix and embedded peptides and these ionized molecules are released as charged particles (ions)

how are ions separated in MALDI-TOF

ions are accelerated through a vaccum tube, smaller ions (lower m/z) travel faster and hit the dectector first and larger ion( higher m/z) take longer to reach the detector 

what does the detector measure in MALDI-TOF

the time it takes each ion to reach the detector (TOF). from that instrument calculated the mass to charge ratio (m/z) and peaks on the output graph represent different peptides or proteins

peptide mass fingerprinting 

split a protein into small peptides at specific locations, can find the m/z of each of those peptides and match them to a database of proteins broken into pieces at the same locations theoretically 

what can be used to break proteins

trypsin, chymotrypsin, pepsin, and thermolysin

what does trypsin do

hydrolyzes the peptide bonds at lysine and arginine residues (cuts there)

where does chymotrypsin cleave peptide bonds?

phe, trp, and tyr

where does pepsin cleave peptide bonds

phe, trp, tyr, and several other

thermolysin

leu, lle, and val

what order for peptide fingerprinting spectrum go

M/Z going smallest to largest 

what do you do when you have sample spectrum

see what the theoretical mass spectrum matches with the sample mass spectrum and compare

steps of peptide mass fingerprinting

genome of the organism, theoretical digestion by trypsin of the entire genome, mostly transcriptome, each small peptide would have its own fingerprint and look through them to predict the probability that the peptide comes from whatever the protein is there because they match

what does mass fingerprinting do for us

looking at overall protein

where does metal oxide affinity chromatography go when looking at phosphorylation status 

before 2D gels, picking spots, and doing MALDI-TOF

phosphorylation status

which proteins might be coming, activated or deactivate by phosphorylation or dephosphorylation

metal oxide affinity chromatography

Phosphorylated proteins have an affinity for metal oxides and can separate phosphorylated proteins from unphosphorylated through a column

how are active and inactive proteins separated  

whatever proteins are phosphorylated are gonna stick to the column and what isn’t will go through it then see how a stimulus causes activation of signaling cascades 

Ubiquitinitylation

76 amino acid peptide chain that signals for many things not just protein degradation

Mono-ubiquitylation

 ends with histone regulation so that has to do with chromatin condensation

Multi-ubiquitylation

not together in any sort of chain can trigger endocytosis

Polyubiquitylation

multiple certain numbers of them can lead to protein degradation and other is DNA repair 

yeast 2-hybrid

after doing everything to identify protein look at any direct protein-protein interaction

what are the two domains of a transcription factor 

the DNA binding domain and the activation domain

Why must the activation and binding domains be close together

so RNA polymerase can be recruited and transcription of the gene can begin

what are fusion constructs used for in the yeast two-hybrid system

to create expression vectors that produce fusion proteins

What does the cDNA of a gene provide in the fusion construct

The sequence that encodes the protein of interest (protein X or Y)

What is the bait in a yeast two-hybrid experiment

The protein fused to the GAL4 DNA-binding domain (usually protein X)

What is the target (or prey) in a yeast two-hybrid experiment

The protein fused to the GAL4 activation domain (usually protein Y)

What happens if protein X and protein Y interact

They bring the activation and DNA-binding domains close enough to activate transcription of the reporter gene.

What is the reporter gene commonly used in yeast two-hybrid assays

the lacZ gene which encodes B-galactosidase

How can you tell if the two proteins interact in the yeast two-hybrid system

Yeast cells turn blue when β-galactosidase is expressed, indicating an interaction.

What happens if protein X and protein Y do not interact

The activation and DNA-binding domains remain too far apart, and no transcription occurs

Do the activation and binding domains have natural affinity for each other

no they only come together if the two fused proteins interact

what does a blue color in the 96-well plate indicate

The proteins interacted, causing expression of β-galactosidase.

interactomics 

study of how all proteins within a cell interact with one another to better understand how changes in protein-protein interactions can manifest as disease. looking at functional genomics to see what gene products interact with other gene products