PHAR 503 midterm

how is the affinity of a drug on a target experimentally determined?

competition

pure target protein purification vs pure drug purification

it's difficult to isolate proteins from cell lysate, but pure drugs are easier—just get cDNAs and synthesize then purify that via chromatography

how does chromatography work?

hopefully the compounds have different characteristics, like electronegativity, size, IMF, etc. that you can use to separate them over beads

what is size exclusion chromatography

to separate POI from other proteins based on size

how does size exclusion chromatography work?

run solution containing POI over column containing porous beads. smaller proteins will get more stuck in the beads while larger ones will flow through, so you elute differently sized groups

what does a 205nm absorbance indicate?

peptide bond!

what does a 280nm absorbance indicate?

tryptophan/any aromatics with delocalized e- side chain

the e- delocalized makes it so high

what's the downside of looking for 205nm absorbance?

it's very narrow, so it's hard to produce = machines are expensive, but the upside is you can see any protein

affinity chromatography

to separate a protein from other proteins by its biophysical traits based on a specific binding interaction between an immobilized ligand and its binding partner

examples of biophysical traits

hydrophobicity, ligands (lectins, protein antibodies, collagen...), metals, protein tags

6-His tag

can bind nickel! makes separating this recombinant protein out easy

glutathione-sepharose (GST)

helps stop bacterial proteins from aggregating during elution

myc pro-oncogene

an epitope we have good antibodies for

how do you get the protein out of the affinity chromatrography column?

wash with salts after eluting everything else

what is the first challenge to determining drug-target affinity?

getting pure drug and pure target!

concept of compound competition

you increase the amount of normal ligand:radio ligand to see at what concentration of normal ligand 50% of the radio ligand is removed—this gives IC50.

IC50

the concentration at which the competing ligand displaces 50% of the radioligand

How do you do compound competition (steps)?

1. introduce known concentration of radioligand

2. wash with known concentration of regular ligand

3. increase concentration of regular ligand and repeat

4. less and less labelled ligand will show up on the filter as you wash

5. determine IC50

Ki from IC50

The inhibition constant for a drug; the concentration of competing ligand in a competition assay which would occupy 50% of the receptors if no radioligand were present. Whereas the IC50 value for a compound may vary between experiments depending on radioligand concentration, the Ki is an absolute value. It is calculated from the IC50 using the Cheng-Prusoff equation:

identification of same or different binding sites

if normal ligand B competes with radioligand A, they have the same binding site

if normal ligand C doesn't compete with radioligand A, they have different binding sites

imidazole

side chain of histidine

aromatic ring with two nitrogens

how is elution also a compound competition?

- increase ion concentration to wash proteins off beads

- compete with imidazole, glutathione, etc. to bind the compound

Surface Plasmon Resonance

Technique used to characterize molecular interactions

Binding interactions are detected by monitoring the reflection of a beam of light off the interface between an aqueous solution of potential binding molecules and a biosensor surface carrying the immobilized bait protein

how does surface plasmon resonance work?

detects changes in reflectance properties on surface of antigen coated sensor when it binds with antibody

what is the biophysical phenomenon behind surface plasmon resonance?

buffer flows over proteins bound to chip. drug binds protein. change in mass on chip surface. change in diffractive index between media. evanescent wave changes, causing a measurable change in its 'shadow' angle in diffracted light

SPR resonance signal

the change in the angle of the 'shadow' in the diffracted light

SPR sensorgram

senses change in resonance signal

SPR sensorgram graph

resonance signal vs. time

shows buffer flow, then association of drug with protein til equilibrium, then dissociation of drug as injection of drug stops

what does SPR measure?

change in resonance signal, which shows online kinetics i.e. association dissociation rates over time + equilibrium constants

low Kd means what?

high affinity (it's not dissociating!)

high Ka means what?

high affinity; Ka describes binding affinity

Ka =

= [target + drug]/[target]*[drug]

Kd =

= [target]*[drug]/[target+drug]

when a drug is given at a concentration equal to its Kd, how many receptors will be occupied?

50%

how can competition be used to show specificity on a sensorgram?

flow a soluble target protein over the chip to compete as a control, because this will eat up any unbound proteins = you make sure measured signals are only of bound proteins

IC50 stands for?

inhibitory concentration 50

mass spectrometry

a technique that separates particles according to their mass

how do you 'fish' for side-effects you like?

modify the drug at random sites and see if those sites cause GOF/LOF of your target side effect

trifunctional capture compound

1. reactive azide

2. biotin for purification

3. drug you're interested in

reactive azide

forms a covalent bond with whatever it's close to upon UV light exposure

biotin for purification

binds with 10^-15 affinity for avidin so makes purification easy

biotin avidin affinity

10^-15 = very low Kd = very high affinity

what do you do once you purify your trifunctional compound?

mass spectrometry to identify proteins that were co-purified and thus potentially drug-binding partners

trifunctional compound workflow

1. modify

2. culture cells, wash with trifunctional drug

3. shine UV 5-10min

3.5 azide binds interaction protein

4. pull out drug-protein complex with biotin-streptavidin

5. wash hard to get the complex off the streptavidin

6. anything unspecific is gone! mass-spec to find the drug + protein!

why protein digestion?

to remove protein from avidin beads via proteases

to make the protein bite-sized so you can run mass spec (too big otherwise)

importance of trypsin in mass spec

this protease only cleaves after K (lysine) and R (arginine), so you can piece the mass spec fragments together based on R and K

how does mass spec work?

negatively ionize your peptides, then accelerate by adding a positive voltage

ions fly through mass analyzer and hit the detector at different times depending on their mass

Only particles with a certain mass/charge (m/Z) ratio will NOT hit the walls and exit onto the detector at the end of the flight tube.

how to read a mass spectrum

y axis = relative intensity

x axis = mass per charge

fingerprint analysis of mass spectrum

list with exact peptide masses measured in a spectrum is compared to 'in silico' cleavage of known proteins in a database

Mascot search

can do the fingerprint analysis for you!

parts 1 and 2 of mass spec

1. generate mass spec, fingerprint analysis

2. validate

how to validate mass spec

crush peptide with air before secondary analysis to figure out where Y and B ions line up

Y and B peptide ions

most common cleavage occurs between C=O and NH group of peptide bond

MALDI-TOF

matrix assisted laser desorption ionization time of flight

what is the distance between two peaks in MALDI-TOF sequencing?

the mass of one amino acid—so you can read the sequence out like a book!

Fluorescence Anisotropy

Depolarized emission for free molecules (low anisotropy)

Anisotropy

having a different value when measured in different directions

fluorophore

molecules that absorb light and reemit it at longer wavelengths

why is emission always at higher wavelengths?

red shift due to energy being lost = higher wavelength = lower energy

this is Stoke's shift!

why are emission/excitation peaks mirror images?

The same vibrational levels being involved in absorption and emission.

Franck-Condon principle: the best overlap between discrete excited and ground state wave functions just happens to be a mirror image of the excitation

Jablonski Diagram

Shows the probabilities of going from one state to another.

An energy diagram that illustrates the electronic states of a molecule and the transitions between them; the states are arranged vertically by increasing energy and grouped horizontally by spin multiplicity.

Franck-Condon Principle

best highest overlap between discrete positions will occur, meaning it is most likely that transfer happens between the best overlapping vibrational wave functions

vibrational levels are DISCRETE

the best overlapping vibrational waves just happen to be mirror images of each other

Stoke's shift

difference between excitation and emission wavelengths

how does Stoke's shift and Jablonski diagrams explain the mirror image of emission spectra?

if you go up to v'=1, you'll go back down to v=1—due to the same vibrational levels being involved in absorption and emission.

monochromator

A device for isolating individual wavelengths or frequencies of light. the ideal fluorescence measurer because they don't need filters

why are 96-well fluorescence plate reader plates opaque black?

so fluorescence doesnt leak between samples, contaminating them

FRET

fluorescence resonance energy transfer

BRET

bioluminescence resonance energy transfer

quenching

gain fluorescence after enzymatic activity

FRAP

fluorescence recovery after photobleaching

TIRF

total internal reflection processes at the plasma membrane

when does FRET occur?

when fluorophores are close together, so it's a great way to see if your drug is binding a particular protein

how does FRET occur?

when emission spectrum of the donor overlaps with the excitation spectrum of the acceptor, the emissions of the donor will excite the acceptor

how close do fluorophroes have to be to give FRET?

10nm

how is BRET different from FRET?

the donor "fluorophore" is a bioluminescent reaction

luciferase reaction

substrate: luminol

luciferase: protease

reaction with oxygen releases light

how does quenching with proteases work?

you have a peptide that includes a fluorophore and a quencher. the quencher steals energy from the fluorophore in a FRET-like manner, cancelling out its fluorescence and keeping it dark. When a protease comes along and cleaves the peptide, the fluorophore is released from the quencher and it lights up.

how is quenching with proteases relevant?

you can tell if your drug is acting at the right protease, ex. to inhibit it, if you don't see fluorescence

importance of the dipole of fluorophores

light exactly parallel to the transition moment excites the fluorophore the best

what direction does a fluorophore emit light?

in the direction dictated by its structure, related to its transition moment

transition moment

dipole, basically

photoselection

in a solution of fluorophores, only those aligned with polarized light will get excited the best, meaning the emitted light will also be polarized

how do you polarize light?

tiny screen with slots

anisotropy

directionality of properties;

describes how much emitted light is still polarized

what does anisotropy indicate about a protein?

how fast it's moving, i.e., if it's bound to something

how does anisotropy indicate the protein is bound to something?

proteins will move slower if they're bound to something (more cumbersome) and so the polarization will be lost less fast

possible movements of a fluorophore

- rotational diffusion of the fluorophore

- mobility of the protein segment (ex. on a hinge)

- rotation of the whole protein

anisotropy emitted light polarization depends on what?

movement of the protein; transition moment

when will emitted light be polarized?

when movement is slow

when will emitted light be unpolarized?

when movement is fast

applications of anisotropy

compound screening assays to see if protein is bound to drug

label at selected proteins in target to see if a target is mobile

which amino acid is intrinsically fluorescent?

tryptophan

low anisotropy

means shit is moving fast and fluorophore is not bound to a protein = light is not polarized

high anisotropy

light is polarized because shit is not moving fast and fluorophore is probably bound to a protein

how did we used to discover drugs?

natural medicines were investigated, then refined

biologically observed agent -> asssay -> hit -> optimization -> studies -> drug

this was great because we already knew it worked!

genes-to-drugs target-based screens

in the 90s, gene sequencing revealed new targets

molecular genetics target identification -> target-based assay -> target hit -> lead optimization -> preclinical and clinical studies -> drug

this was not great because a lot of targets are not biologically activated—you can make a drug that doesn't work in vivo

phenotype-based screens

targeting a phenotype, so biology first!

human cell system -> biological hit (find something that works) -> lead optimization -> preclinical and clinical studies -> drug

this works because you're looking for something that has an effect phenotypically

cell-based assay cell lines

immortalized: human embryonic kidney, HEK293, HeLa

primary cell cultures (die quickly)

induced pluripotent stem cells

issue with primary cell cultures?

not immortalized; die quickly

target-based screen

you're targeting a specific protein—shit may not work because you don't know the full MOA of the protein, but you'd do this for example in cases you know for sure a target is bad