Separation techniques 1+2

Question 1

What are some chromatographic methods

Answer 1

• Dialysis
• Ultrafiltration.
• Paper and thin-layer chromatography.
• Dye-ligand and metal-affinity chromatography.
• Ion exchange chromatography.
• Hydrophobic interaction chromatography.
• Gel filtration

Question 2

Separation with dialysis

Answer 2

• based on commercial membranes with pores of a pre-determined size.
• used to be sold as long, flattened tubes of ‘Visking tubing’ now usually sold in single-use cassettes.
• often pore-size is given based on the size of (roughly) spherical protein that will not pass through the holes
  – unit is molecular-weight cut-off (MWCO).
• e.g. a membrane with 20 kDa MWCO holes, if formed into a bag. If we fill that bag with a mixture of proteins e.g. cell-free extract (CFE) from Escherichia coli (1 kDa to 500 kDa) and place the bag in a very large volume of buffer, the bag will retain all proteins >20 kDa, but those smaller than 20 kDa will go through the holes into the buffer.
• dialysis membranes are made of cellophane (often sold as ‘cellulose’), cellulose nitrate.
• terminology – what goes through is the filtrate. What stays behind is
  the retentate.
• removes ALL molecules/ions below the MWCO e.g. buffers, salts so can be used for desalting, but slow and laborious.

Question 3

What is the issue with dyalisis

Answer 3

the issue is the filtrate in the bag gets diluted so you lose the high concentration of the protein.

Question 4

Ultrafiltration

Answer 4

• “posh” version of dialysis – similar membranes are filled with
  protein solutions and then a force is applied to force the proteins and water through the membrane. Does not dilute the proteins leaving through the membrane so much more convenient and now widely used. VERY expensive!
• centrifugal ultrafiltration uses a centrifuge to apply force to the solution, pulling it through – water, salts and anything below the MWCO of the filter end up pulled through, everything else is left
  behind.
• pressure ultrafiltration same sort of idea but a chamber where base is the filter and top is connected to a cylinder of argon gas –the gas pressure above the liquid pushes it through the filter. Much faster and suited to large volumes.
• can be used to ‘cut’ a protein mixture
  *used for partial purification of a protein of known
  size.

Question 5

what do you use for;
gas chromatography
paper chromatography
FPLC, fast protein-liquid

Answer 5

stream of inert gas for gas chromatography
paper and thin layer we use organic solvents
a buffer solution for FPLC

Question 6

Explain chromatography techniques

Answer 6

• can be used to separate on the basis of shape, size, ionisation (charge), polarity and hydrophobicity.
• We have a solution of solutes (the things we want to separate) dissolved in a solvent (usually a buffer at a
  specific pH.
• this solution is applied to a stationary phase and a mobile phase passed over it – the interaction of the solutes with both phases dictates how they
  separate.
• stationary phases e.g. paper (PC), silica gel (TLC) etc.
• mobile phases e.g. buffer solutions, organic solvents etc.
  hydrophilic/hydrophobic dictates how fast they move. Hydrophilic move fast, hydrophobic move slow. This is the partitioning of inks between the two phases.

Question 7

What are the types of chromatography

Answer 7

*PC, paper chromatography
*TLC, thin-layer chromatography
*HPLC, high-performance liquid chromatography
*FPLC, fast protein-liquid chromatography
*GC, gas chromatography.

Question 8

Paper and TLC chromatography

Answer 8

• paper is cheaper but slower.
• can be run ‘descending’ by hanging paper from a trough of
  solvent and letting it drip off the bottom.
• TLC is faster and precise but expensive.
• TLC plates are glass or metal foil sheets coated in the stationary phase e.g. glass coated in silica gel, cellulose or alumina
• iTLC plates are glass fibre ‘paper’ impregnated with the
  stationary phase – much faster but very fragile. Never really
  widely used –
• HPTLC uses different plates and a printer to apply samples –
  much neater! Special constant humidity/temp/pressure chamber.
• stationary phases also ion-exchange-coated celluloses.

Question 9

Chromatography: relative front values

Answer 9

• how do you know what a band is on a chromatograph?
  you use relative front value (Rf also called ‘retention
  factor’).
• in paper chromatogram shown, assume solvent went from bottom up to the top of the paper.
• we measure from the place we applied the sample to the solvent front (the line the solvent reached soaking up the paper). This value is f (mm)
• we then measure from the place we applied the sample to each band b(mm)
  Rf = b/f
• an Rf value for any given analyte is only valid for ONE matrix solvent.

Question 10

Chromatography: detection

Answer 10

plate/paper is sprayed with a reagent to detect specific groups of biomolecules.
* iodine vapour (I2 gas from crystals of iodine in a plastic bag in a warm place) detects unsaturated fatty acids as brown spots. Doesn’t last long – spots vanish so must be drawn around.
* ninhydrin detects amino acids – most are violet but some are yellow, brown, blue etc.

Question 11

Chromatography: adding efficiency

Answer 11

• plates/papers are inefficient as they are only able to be run one or a few at a time and the process cannot be automated.
• column chromatography allows automation of the process
• standard liquid chromatography (LC) is uses a bed of silica gel
  and has sand on top to protect it as mobile phase is pushed
  through under a low pressure.

we use FPLC for purifying proteins and HPLC (for quantifying small analytes)

Question 12

FPLC: basics

Answer 12

• uses resin beads based on several different polymers derivatised with functional groups that proteins interact with.
  what the beads are made of dictates how much pressure they can take thus how fast the flow.
  Absorption is measured in a flow-cell, continuously and plotted on a computer or chart-recorder in real-time.
• after the flow-cell, protein-containing eluent is collected in a fraction collector in fixed volumes.

Question 13

FPLC: dye-ligand

Answer 13

• resin is derivatised with Cibacron Blue F3G-A dye.
• various resins are made with it e.g. Blue Sepharose
• binds to NAD+ using dehydrogenase enzymes and some other proteins.
• red circled bits of the dye interact with NAD+ binding-sites on the enzyme.
  Biomimetic chemistry.
• proteins washed back off column using a salt gradient (NaCl)

Question 14

FPLC: metal affinity

Answer 14

• histidine has a strong affinity for Co(II) and Ni(II) ions.
• “his tags” (6-20 histidine residues) can be added to one end of a polypeptide chain when cloning/expressing in Escherichia coli (using pHAT12 vector, for example).
• resins loaded with Ni(II) or Co(II) ions will bind his
  -tagged proteins very strongly.
• column is washed to remove all other proteins in sample.
• tagged protein recovered by flushing with imidazole (look at structure versus histidine

Question 15

FPLC: ion exchange

Answer 15

• at a specific buffer pH some proteins in a sample bind strongly to a resin with basic side-groups, some bind weakly, some don’t bind.
• column of resin is equilibrated with a buffer at that pH.
• protein mixture is run through the column – target protein usually binds strongly (column is selected on purpose!).
• column is washed in the same buffer to remove non-bound proteins.
• a salt gradient is started – buffer “A” is now mixed by a gradient mixer with buffer “B” containing e.g. 1 M NaCl.
• weak bound proteins fall off at low [NaCl], target protein at high [NaCl].
• once we know the specific [NaCl] at which it falls off, we can just flush one buffer at that [NaCl] through the column after washing it: rapid purification.

Question 16

FPLC: hydrophobic interaction

Answer 16

• works for proteins that are pretty hydrophobic.
• protein is dissolved in buffer containing enough ammonium sulfate that the protein is still in solution but only just.
• mixture is applied to a column with big, hydrophobic groups (e.g benzene rings) – A waxy column, almost. Protein sticks to the column as it wants to get out of solution (precipitate, sticking to itself).
• buffer with high ammon. sulfate “A” is now mixed with one without any “B”, and once the concentration of ammon. sulfate is low enough, the protein falls off of the column and goes back into solution.
• often used as a ‘polishing step’ on a fairly pure protein.

Question 17

FPLC: gel filtration (size exclusion chromatography.)

Answer 17

• column contains porous gel beads.
• big proteins move fastest through column as too big to fit into pores in beads and instead go around the beads (a short
  route).
• small proteins move slowest through column as pass through
  pores within beads and take a much longer route to the exit.
• big proteins come out of the column first, small ones last.
• dilutes the sample a fair amount.
• can be used to remove small molecules from a protein e.g. salt (“desalting”) or to remove one buffer when you need to use a different one (“buffer exchange”) as inorganics are so small they take ages to come through!
• column is normally calibrated with coloured proteins (etc) so
  that we know how long it takes for a protein of a specific size
  to come out.
• blue dextran (2,000 kDa) will come out long before any
  proteins so can be added to a sample to mark the ‘start’. Blue.
• vitamin B12 (1.4 kDa) is smaller than most proteins and can be used as a marker for the ‘end’ of a sample. Pink
  use calibration graph to find out unknown protein

Question 18

HPLC: key points

Answer 18

• usually uses a TLC-type partitioning across stationary and mobile phases.
• HPLC runs at a higher pressure than FPLC and uses small, metal-cased columns.
• suited to small biological molecules – amino acids, fatty acids etc.
• can be automated.
• faster than TLC but not necessarily any better!
• still need to run standards, the computer can’t work miracles.
• detection is by absorbance (“UV-Vis detector”), fluorescence or by mass spectrometry (“HPLC-MS” or “LC-MS”).
• if stationary phase is hydrophobic and non-polar (C18 or C8
  resin) and we use a polar solvent (acetonitrile, methanol, tetrahydrofuran) we call this reverse phase HPLC.
• if stationary phase is polar (-amine or –sulfonate substituted resins) and we use a non-polar solvent (n-hexane, n-heptane) we call this forward phase HPLC.
• solvents are often adjusted to extremes of pH to give the best separation.
• gradients of two solvents are also sometimes used.

Question 19

What type of FPLC is octyl sepharose used for?

Answer 19

hydrophobic interaction

Question 20

FPLC what absorbance detects;
proteins
coloured proteins like cytochrome c
nucleic acids

Answer 20

• protein coming out of the columns is detected usually by A280
• coloured proteins like cytochromes c are detected by e.g. A550.
• nucleic acids (normally a contaminant) are detected by A260.

Question 21

principles of electrophoresis

Answer 21

• ions are charged atoms/molecules e.g. Na+ Cl-
• charge depends on pH for many chemical species H3PO4 refer to Bjerrum plots
• the correct pH is required for electrophoresis to function at all as proteins have mix of base and acid aa side-chains - pH has to be such that overall protein has the charge you want it to.
• B-DNA has an overall negative charge above pH 4 [pKa = 2] so any buffers above this will work!
• type of buffer matters as much as pH – some can set your gel/paper on fire as their electrical resistance is too low!

Question 22

What gels do we use for proteins and nucleic acids?

Answer 22

• agarose gels - repeating polysaccharide from Rhodophyta- for nucleic acids.
• polyacrylamide gels – repeating acrylamide polymers cross-linked with bisacrylamide- Used mainly for proteins.

Question 23

DNA electrophoresis

Answer 23

• 0.5-2.0 % w/v agarose is dissolved in a buffer by
  heating.
• Buffers mostly used are:
  -TAE (40 mM Tris-acetate buffer pH 8.3 with 1 mM
  EDTA)
  -TBE (89 mM Tris-borate buffer pH 8.3 with 2 mM
  EDTA)
• Usually visualising agent e.g. ethidium bromide
  (EtBr) is added to the gel while still hot.
• “Safer” alternatives like Nancy-520 can also be
  added to the gel.
• Other stains (SYBR Gold, SYBR Green) are used
  to soak the finished gel and aren’t put into it.
• Gel is poured when hot into a plastic casting tray
  with the ends taped/dammed closed.
• A comb is added to the molten gel which will
  leave sample-wells once it has set. wells are marked
• DNA molecule in purified state is associated to Mg2+
• the EDTA in the buffer is a chelating agent – binds it
  and leaves the DNA negatively charged.
• DNA samples are mixed with loading buffer which
  contains EDTA and e.g. sucrose or glycerol (denser
  than buffer – makes it sink into the wells) and a
  loading dye (usually a mixture of bromophenol blue
  and either xylene cyanol FF or orange G).
• loading dyes are pulled towards positive terminal ahead
  of DNA and separate into two bands (blue and cyan or
  blue and orange) so you know the gel is working.
• cooled, set gel is put into the tank and flooded with
  same buffer it was made off (with some EtBr too).
• DNA-in-loading-buffer is pipetted into wells.
• a DNA ladder is loaded into some wells of the gel to
  help determine size of DNA bands.
  Example of DNA loading dyes xylene cyanol FF migrates the same as a 4 kbp length of DNA on this gel.
• voltage is applied – this depends on size of gel
  tank and what buffer you’ve used.
• DNA (negatively charged) moves towards the
  positively charged terminal (black) at bottom of
  the gel (meanwhile Mg(II) moves towards the top
  of the gel by the negative terminal (red).
• DNA is sieved by the gel matrix as well as moving in relation to charge – this results in large DNA molecules moving slowly and small DNA molecules moving quickly.
• Gel is removed from the tank and visualised e.g.
  (for EtBr) exposing to 290-300 nm UV light –
  DNA-EtBr-conjugate glows pink (605 nm).

Question 24

RNA electrophoresis

Answer 24

a few key differences versus DNA method
1) RNA has to be denatured to remove any hairpin loops etc – done using formaldehyde normally.
2) formaldehyde is included in the gel itself
3) every buffer etc has to be pre-treated with DEPC (diethyl
pyrocarbonate) to destroy RNase enzymes (survive autoclaving!)
this only works actually in the autoclave so unsuitable for anything that can’t be autoclaved.
4) plasticware has to be pre-treated with RNase-destroying products (e.g. RNaseZap™ by Invitrogen)
5) need to work in an ultraclean manner to avoid RNAse
contamination.

Question 25

protein electrophoresis: PAGE

Answer 25

* polyacrylamide gel electrophoresis (PAGE) * polyacrylamide is made from monomer acrylamide and crosslinker bisacrylamide dissolved in buffer, de-oxygenated in vacuo. Ammonium persulfate (APS) and TEMED catalyse the polymerisation (poisoned by O2) – takes 1-5 h for gel to polymerise. * gels come in two types: slab gels (single % acrylamide in whole gel) and gradient gels (% of acrylamide gets higher as you go down the gel). * slab gels have better resolution at the top and middle as smaller proteins separate poorly and bands are blurry. Gradient gels are used to give good resolution all the way down but fussy to make. * the higher the % of acrylamide in the gel, the harder it is and separation is suited to smaller proteins. * gel is mixed, poured into a 0.5-1.0 mm-thick casting cassette (two glass plates, disposable plastic cassette) and overlaid with water saturated n-butanol to exclude oxygen. Once set, this is removed and top of gel washed. * for most slab gels, a stacking gel (pH 6.7) is added on top of the resolving gel (pH 8.9) in which a comb is added to give sample wells.

Question 26

protein electrophoresis: SDS-PAGE

Answer 26

* uses sodium dodecyl sulfate (SDS) as a detergent to denature proteins and apply a charge to them: SDS-PAGE. Goal is to linearize proteins. * SDS is in the gel itself and in running buffers and in the sample buffer. * gels contain a Tris-HCl buffer and running buffer is Tris-glycine based. * disulfide bridges must be broken by adding a reducing agent to the loading buffer – βmercaptoethanol (BME) for SDS-PAGE. * protein mixture added to loading buffer (glycerol, buffer, BME, SDS, bromophenol blue) and boiled to assist denaturation and breaking disulfide bridges. * for normal SDS-PAGE slab gels, the gel made in Tris-HCl buffer but running buffer (at each end, connecting electrodes to gel) is Tris-glycine pH 8.3 * cooled samples in loading buffer are applied. A protein marker (proteins of known sizes) is added to some lanes ( DNA ladder). Sometimes use dye-conjugated proteins so you can see them separate in real-time. * a voltage is applied – proteins have a net negative charge because of SDS molecules bound all over them. * they initially move quickly in the stacking gel and because of the chloride-glycine buffer boundary “stack” (stop moving) at the interface between gels. * voltage now increased, they move into the resolving gel and separate. * large proteins move slowly, small ones move quickly! * gel is removed from tank and fixed and stained (normally one single step nowadays) * Coomassie brilliant blue dyes (CBB G-250 and R-250) are most common stains – used in acetic acid and methanol to fix proteins so they don’t diffuse in the gel. * excess dye is removed by destaining to give blue bands on a clear ground. * negative stains are sometimes used esp. when you need unstained bands for mass spec ID etc – zinc staining give clear bands on white ground. * silver stains and gold stains are used when v low levels of protein present or to ID protein groups. e.g. silver stains: most proteins brown/black, proteolipids are blue, glycoproteins are red.

Question 27

protein electrophoresis: Nu-PAGE

Answer 27

* much like SDS-PAGE but uses lithium dodecyl sulfate (LDS). * instead of β-mercaptoethanol, dithiothreitol is used. * a second reducing agent is present in the running buffers (sodium sulfite) to minimise re-forming of disulfide bridges. * slab Nu-PAGE gels have better resolution than gradient SDS-PAGE gels. * running buffers are usually MOPS or MES-based. * gels are Tris-acetate buffered. * time-saving, gels are pre-cast so very time-effective

Question 28

protein electrophoresis: native PAGE

Answer 28

* for multi-subunit proteins we may want to separate them “as is”, without denaturation. * many native PAGE techniques exist: * acid-native (low pH gel, separates proteins that have mainly amine-side-chain aas) * basic-native (high pH gel, separates proteins have that mainly acid-side-chain aas) * blue-native (BN-PAGE, uses neutral-ish pH and CBB G-250 itself to impart a charge on the proteins so they move at neutrality – gels are blue)

Question 29

What is the purpose of glycerol in loading buffers?

Answer 29

Makes the buffer more dense so the DNA/protein sinks into the well and doesn’t float into the buffer.

Question 30

Give two methods of polymerising a polyacrylamide gel

Answer 30

Either APS/TEMED alone or riboflavin/TEMED with UV light.

Question 31

Give the two blue dyes used to stain SDS-PAGE gels.

Answer 31

Coomassie Brilliant Blue G-250 Coomassie Brilliant Blue R-250.

Question 32

What is the advantage of silver-staining over dye-staining?

Answer 32

Can distinguish glycoproteins and proteolipids and is much more sensitive.

Question 33

What is the advantage of zinc-staining over dye-staining?

Answer 33

Doesn’t bind anything to the proteins so they can be washed out of the gel and used in other experiments.