Home Exam Answers:

1. How do you expect the charge distribution of a protein to change between pH 0 and 14? That is, describe qualitatively how (and why) the charge varies as a function of pH.

Answer (1).

2. For what kind of sample is it best to use differential centrifugation, Rate zonal or isopycnic centrifugation? What are the differences between these three centrifugation methods?

Answer (2). Differential centrifugation is used for the sample particles with the same mass and it is used in seperation of subcellular organelles and macromolecules.

Rate zonal centrifugation is used for separation of particles which differ in their size, but not in density so it seperates cellular organelles such as endosomes or separation of proteins, such as antibodies.

Isopycnic centrifugation is used for separation of samples particles that differ in their density, but not in size this is used to seperate nucleic acids in gradient solutions.

Differences between differential centrifugation, Rate zonal and Isopycnic centrifugation:

Differential Centrifugation

Separation is attained principally based on the size of the particles.

This separation is mostly used in simple pelleting and in locating partially-pure research of subcellular organelles and macromolecules.

Larger and more colossal components will sediment at lower speeds, while smaller components necessitate higher centrifugal force.

Rate zonal

particles differs in their size, however not in density .

Position of sample is related to the time of sedimentation (time dependent)

Its a Flat gradient.

Maximum density of gradient does not exceed that of the most dense particle that is the pellet.

Isopycnic centrifugation:

particles differ in their density, but not in size .

Here size is only affecting the rate of migration (time-independent)

It is a Steep grandient.

Separation is based on the buoyant densities of the particles, so no pellet.

3. (a) Describe the advantage of a stacking gel during denaturing electrophoresis. What is the principle behind it?

Answer 3. (a)The main advantage is that the proteins electrophorese quickly through the stacking gel and "pile" at the border between the two gels, before they move in the gels. This upturns the firmness of the proteins before they enter the running gel and increases resolution. So when the electrophoresis is started, ions from the higher reservoir enter the stacking gel since at that pH they have a middling fractional negative charge. The stacking gel buffer ions continue moving in the stacking gel, but when the ions enters the pH of the stacking gel, they are converted into zwitterions with a net charge of zero, and therefore stop motion toward the anode. The electrical resistance in the stacking gel then increases since the number of ions moving through the stacking gel decreases. To maintain constant current throughout the circuit, there will be a confined increase in the voltage in the stacking gel (from Ohms Law, V=iR). This will cause the proteins to travel quickly and all stack in a single, very thin disc right behind the ions in the stacking gel (which are in front because they have the highest charge density and electrophoretic mobility of any ion in the stacking gel). The proteins will not pass the ions since if they did, they would straightway slow down since they would no longer be in an area of reduced charged carriers and higher voltage. At the stacking gel/running gel interface, all the proteins can’t migrate at the same speed, owing to sieving effects of the more concentrated gel, and hence will be separated in the running gel.

(b) Describe the principle of a 2D-gel and discuss various staining methods for gels (sensitivity, advantages and disadvantages).

Answer 3. (b) The main principle of this technique is that it separates the proteins according to two properties in two separate steps: the first-dimension step, isoelectric focusing (IEF), separates proteins according to their isoelectric points (pI); the second-dimension step, SDS-polyacrylamide gel electrophoresis (SDS-PAGE), separates proteins according to their molecular weights (Mr, relative molecular weight). Every spot on the consequential two-dimensional array resembles to a single protein species in the sample. Many different proteins can be separated, and info such as the protein pI, the apparent molecular weight, and the amount of each protein is achieved.

Various Staining methods for Gels:1. Coomassie Brilliant Blue (CBB): It is a quick and strong way of envisaging proteins in a gel,

however they generally lack in sensitivity. CBB staining procedure is relatively quick and very easy. Since there is presence of alcohol some of the proteins release the dye during the background destaining process.

2. Silver staining is the most sensitive non-radioactive method (below 1 ng). Silver staining detects the proteins mainly on the gel surface .Silver staining is a complex, multi-step process using several reagents for which quality is critical. For the reason that it is not an endpoint procedure, the staining intensities can differ from gel to gel.

3. Fluorescent stains uses fluorescence as a detection method and the staining is an endpoint staining and is thus highly reproducible. Fluorescent protein gel stains are usually well-suited with consequent protein analysis.

4. (a) Mention two important special features of ESI.

Answer 4. (a) 1. The ability to analyze the compounds from aqueous or organic solutions by coupling with Liquid chromatography.

2. It can induce the fragmentation of small peptides typically below 3000Da thus allowing the sequencing information to be obtained.

(b) Make a simple drawing of a MALDI-TOF mass spectrometer.

Answer 4. (b)

4. (c) What simple relation is used to determine molecular masses in MALDI-TOF mass spectrometry?

Answer 4. (c) MALDI-TOF mass spectrometers operate on the principle relation

K.E. = [mv2]/2 = zeEs

Where K.E. = kinetic energy; m = the mass of the ion; v = velocity of the ion; z = number of charges; e = the charge on an electron in coulombs; E = electric field gradient; and s = the distance of the ion source region.

d) What is the lowest detection limit of peptides in MALDI-TOF mass spectrometry?

e) Using peptide mapping-How would you determine the identity of a protein present in a mix with other proteins?

Answer 4. (e). To identify the unidentified protein of interest in a mix the protein is first cleaved into smaller peptides, whose absolute masses can be accurately measured with a mass spectrometer such as MALDI-TOF or ESI-TOF. These masses are then related to either a database comprising of known protein sequences or even the genome databases.

f) How can you use MALDI-TOF mass spectrometry to determine if a protein is phosphorylated or not on a serine or a threonine amino acid?

Answer 4. (f).

(i) Degradation of the phosphoprotein into small peptides by specific enzymatic or chemical reactions.

(ii) Identification of the phosphopeptides by -80 (or multiples of -80)-Da mass shifts in the mass spectra after dephosphorylation with alkaline phosphatase.

(iii) Location of the phosphorylation sites by mass mapping.

(ref: Liao PC, Leykam J, Andrews PC, Gage DA, Allison J., Anal Biochem. 1994 May 15;219(1):9-20.)

5. You are trying to purify a Very Interesting Protein (VIP) with the help of ion-exchange chromatography. After the purification step you analyse the fractions by SDS-PAGE and Western Blotting. The Antibody identifies a protein of 22 kDa as VIP, but in the SDS gel you see that the fraction also contains three contaminating proteins with molecular masses 6 kDa, 28 kDa, and 60 kDa. How would you continue to purify your protein using chromatography? Discuss the advantages and disadvantages of the possible techniques.

Answer 5.We can use the gel filtration chromatography here, since it separates the molecules according to their size and shape. Hence our protein of interest can be sieved from the contaminating proteins with molecular masses 6 kDa, 28 kDa, and 60 kDa and thus can be purified. To perform this separation, all the proteins with known molecular weights are run on the column and their elution volumes noted. If the elution volumes are then plotted against the log molecular weight of the matching proteins, a straight line is obtained for the separation range of the gel being used.

The main advantage of gel filtration is that it’s the best method for separation of molecules differing in molecular weight because:

1. It doesn’t rely on temperature, pH, ionic strength and buffer composition. So separation can be carried out under any conditions.

2. There is very slight adsorption

3. The elution volume is associated to the molecular weight.

The disadvantages of the gel filtration chromatography:

1. The column should be precisely prepared to get optimum separation. 2. Any cracks or cutoffs in the column will interfere. 3. The size of the sample and the rate of buffer flow must be strictly controlled.

6. (a) Describe the principles of CD-spectroscopy.

Answer 6. (a). The physical principle of CD-spectroscopy involved states that the Chiral or asymmetric molecules produce a CD spectrum because they absorb left and right handed polarized light to different extents and thus are considered to be "optically active"The difference between the absorption of left and right handed circularly-polarized light and is measured as a function of wavelength. CD is measured as a quantity called mean residue ellipticity, whose units are degrees-cm2/dmol.

6. (b) What type of information can be obtained from a far- and near- UV CD spectrum, respectively?

Answer 6. (b).

i. The near-UV CD spectrum has very high sensitivity for the native state of a protein. It can be used as a fingerprint of the correctly folded conformation.

ii. The far –UV CD spectrum detects the occurrence of ordered secondary structure and can be used to guess the secondary structure .CD spectra of natively unfolded proteins, being measured in the far ultraviolet region, These values are discrete enough to differentiate them from ordered proteins.

7. Interactions between biomolecules can be studied using several different techniques mentioned during this course. Suppose you would like to study the interaction between a DNA-binding protein and DNA. How would you characterize this interaction in detail? Describe the experiment(s) you have in mind.

Answer 7.

1. Chromatin Immunoprecipitation (ChIP) Assays: The ChIP method can be used to screen transcriptional regulation through histone modification (epigenetics) or transcription factor-DNA binding interactions. The ChIP assay method permits study of DNA-Protein interactions in living cells by considering the cells with formaldehyde or other crosslinking reagents in order to stabilize the interactions for downstream purification and detection. It can be used to do quantitative study when coupled with qPCR investigation.

2. DNA Electrophoretic Mobility Shift Assay (EMSA): It is used to investigate the proteins binding to known DNA oligonucleotide probes and can be used to evaluate the ability of affinity or specificity of the interface. The method is built on the statement that protein-DNA complexes migrate more gently than free DNA molecules when exposed to non-denaturing polyacrylamide or agarose gel electrophoresis.

3. Reporter Assays: It delivers a real-time in vivo read-out of translational movement for a promoter of interest. Reporter genes are fusions of a target promoter DNA sequence and a reporter gene DNA sequence. Here we can incorporate the reporters such as firefly luciferase, Renilla luciferase or alkaline phosphatase. This technique detects real-time data and it’s a potent tool for mutational analysis of promoters which is adaptable to high-throughput screening.