theorysanjeeva/virtual_lab/virtual_laboratory/2dg_man… · fig2: levels of structural complexity...

Virtual proteomics lab

THEORY

Proteins are complex bio-molecules that have fascinated scientists for several decades due to the wide range of diverse functions they possess in all living organisms. Various techniques have been developed over the years by many researchers to study the structure and functions of proteins. Amongst all these, electrophoresis has been referred to as ‘Gold Standard’ due to its ability to provide much information about protein structure and properties. Two-dimensional gel electrophoresis (2DE) is one of the most widely used electrophoretic techniques for separation of complex mixtures of proteins due to its ease of use. Protein separation is carried out based on two different properties of proteins namely, isoelectric point and molecular weight. Several advancements have been made in the separation procedure thereby increasing the robustness and efficiency of separation.

Properties of proteins and separation

Proteins are biologically vital, complex biomolecules that possess a very wide range of functions compared to the other biomolecules. They possess dynamic physical and chemical properties which are largely dependent on their composition. Unlike DNA, they do not possess uniform negative charge, which makes the task of studying these molecules using regular, conventional biochemical techniques very challenging. Proteins are long, polymeric, macromolecules made up of repeating units of amino acid residues. Amino acids are molecules containing an amino group, a carboxylic acid group and an aliphatic or aromatic side chain that varies between different amino acids.

The amino acid side chain, referred to as the R group, varies across amino acids thereby resulting in differences in properties of the various amino acids. The presence of an acidic carboxylic acid group and a basic amino group confers amino acids with an amphoteric nature i.e. the ability to behave as acids or bases. This results in amino acids bearing a different net charge depending upon pH of the surrounding environment. This property is utilized as a separation criterion for protein mixtures. Proteins have multiple structural levels that gradually increase in complexity. The primary structure consists of the linear arrangement of amino acids that are joined together by means of peptide bonds. The polypeptide backbone of the protein undergoes folding due to formation of several internal bonds which gives rise to the secondary structure. Further interactions between the side chains of the amino acid residues and interactions between multiple polypeptide chains (subunit) lead to the tertiary and quaternary structures respectively.

©Sanjeeva Srivastava, IIT Bombay, 2010


Fig1: General structure of amino acid showing side chains

Because of their side chain they acquire a specialized property called amphoteric (reactivity as either an acid or base) nature, hence it has difference in charge depending on pH of surrounding environment; same is used as separation criterion in mixture of proteins. The primary structure of a proteins are made up of linear arrangement of amino acids, as the structural arrangement goes towards secondary, tertiary finally till quaternary, its chemical complexity increases. These properties are used up for an efficient separation on electrophoresis.

Fig2: Levels of structural complexity of proteins (Ref: Biochemistry by Lehninger, 3rd edition)

History & development of electrophoresis

Electrophoresis was invented by the Swedish scientist, Prof. Arne Wilhelm Kaurin Tiselius in 1930, for

which he received the Nobel Prize in Chemistry in 1948. The concept of electrophoresis initially

developed by him was carried out in free solution and was termed as Moving boundary electrophoresis.



Reports of using filter papers and gels in electrophoresis were first seen only in early 1950 and rapid

developments that ensued allowed electrophoresis to become a widely adopted technique by 1960.

Advantages of 2-DE over 1-DE

Initially proteins were separated by one-dimensional electrophoresis using molecular size as the

separating criterion with smaller proteins migrating further along the gel than the larger proteins.

However, this failed to provide comprehensive resolution in complex protein mixtures like serum, cell

lysates etc. due to the complexity of the sample. In early 1956, Smithies and Poulik recognized a new

approach by which a combination of two electrophoretic processes was carried out on a gel at right

angles to each other. This provided better separation than electrophoresis in a single dimension.

The concept of 2DE was developed and established by O’Farrell in the year of 1975. It became

rapidly accepted and adopted by many researchers as an extremely useful technique to study a complex

mixture of proteins. As the name suggests, proteins are separated in 2-DE based on two different

properties - isolectric point (pI) in the first dimension and molecular size in second dimension. 2-DE was

successfully employed for the first time by Anderson and Anderson for the analysis of human plasma

proteins.



Fig 3: Diagrammatic representation of isoelectric focusing

Separation across two dimensions

The first dimension of 2-DE makes use of the amphoteric nature of proteins for separation. Tube gels, in

which a pH gradient was established using a suitable ampholyte solution consisting of low molecular

weight organic acids and bases, were originally used for isoelectric focusing. These pH gradients were,

however, not very stable and had the tendency to breakdown upon application of concentrated samples

thereby leading to a lot of variation in results obtained. Reproducibility was further decreased due to

the prolonged focusing time and tube gels were eventually replaced by immobilized pH gradient (IPG)

strips to overcome the hurdles. IPG strips make use of immobulins that are co-polymerized with the gel

and therefore eliminate the errors related to IEF separation. A wide range of preformed pH gradients on

polyacrylamide gel support are available commercially that can be selected depending on the

complexity of sample to be used in the experiment. This innovative concept of IPG strips was developed

by the German scientist, Angelika Görg. During IEF, the sample proteins which possess different charges,

move across the pH gradient strip due to the applied electric field. They come to rest at a position where

their net charge is zero and they can no longer migrate in the electric field. This is known as the

isoelectric point (pI) and the final position of protein is dependent on the balance of charged groups in

the protein and independent of size.



Fig 5: Isoelectric focusing system with IPG strips loaded.

The second dimension of separation in 2-DE makes use of SDS-Page, a commonly employed technique

that separates molecules based on their molecular weights. The Sodium dodecyl sulfate used in this

technique is an anionic detergent that denatures the proteins and confers them with a uniform negative

charge proportional to their molecular weight. This ensures that the charge-to-mass ratio of all proteins

are nearly identical thereby facilitating separation purely on the basis of molecular weight.

Fig 6: Action of SDS on proteins.

The polyacrylamide gel is prepared using acrylamide, bisacrylamide and SDS with the polymerization

reaction being aided by Ammonium persulfate (APS) and TEMED. TEMED initiates the polymerization

reaction; APS acts as a free radical supplier to facilitate the reaction and bisacrylamide functions as a



cross-linker in the gel. Samples are loaded on the gel in sample wells and the gel placed in the

electrophoresis unit containing suitable buffer solutions. Application of an electric field brings about

migration of the proteins through the gel based entirely on their molecular weight. Once separation

has taken place, the proteins are stained with various staining dyes that enable visualization. Stains

such as Coomassie blue and silver are routinely used during 2-DE. Many fluorescent stains such as

SYPRO Ruby Red, SYPRO Orange, Deep Purple etc. have been developed more recently which allow for

extremely sensitive detection of even small amounts of proteins.

Fig 7: Pictorial representation of a 2-D electrophoresis experiment.

2D-PAGE can separate around 2000 proteins simultaneously in a single run, which makes it a highly

effective and versatile tool to study relative protein concentrations in complex protein mixtures. 2-DE,

more than serving as a separation technique for resolving large numbers of proteins, acts as an effective

visualization and profiling tool for monitoring of relative abundances of the proteins through

appropriate staining procedures. Hence the technique was adopted to study maps of proteins that

portray changes in protein expression level, isoforms or post-translational modifications.



Comparative profiling of multiple samples using 2-DE proved to be a challenge since only one sample

could be resolved per gel. Gel-to-gel variations and lack of reproducibility made it difficult to compare

gel patterns of samples run on two different gels. Therefore the requirement for a stable and robust

method which could overcome this hurdle led to the development of Difference Gel Electrophoresis

(DIGE) in 1997 by Mustafa Ünlü and his group. This technique allows separation and detection up to

three distinct protein mixtures on a single 2DE gel. It enabled researchers to study the relative

expression of samples with respect to varied biological conditions such as disease. 2D-DIGE uses

fluorescent cyanine dyes, Cy2, Cy3 and Cy5, each having a specific excitation energy and wavelength

that enables the accurate detection of proteins labeled with each dye. Diseased samples, healthy

control and an internal pooled standard, consisting of equal amounts of sample and control, are labeled

with the three different Cy dyes; they are then mixed and run on a single gel. Once separation of

proteins takes place across the two different dimensions, the gel is scanned and visualized at different

wavelengths allowing relative expression patterns to be compared and quantified based on the spot

intensity. The gel images are analyzed using advanced softwares which accurately quantify the

differential expression of many proteins on the single gel. The pooled internal standard having all the

proteins of both the sample and control minimizes any variability that may arise in the gel, thereby

providing an efficient and accurate technique for protein profiling.



Fig 8: Pictorial depiction of a typical 2D-DIGE experiment.

MANUAL

2-DIMENSIONAL ELECTROPHORESIS

2-dimensional electrophoresis is a versatile tool used for the separation and visualization of complex protein mixtures. Separating proteins from mixtures such as serum, whole cell lysate etc is a daunting task, which can be efficiently carried out with the help of 2-DE. The experimental procedure employs two stages - isoelectric focusing, which separates proteins in the first dimension on the basis of their isoelectric point, and SDS-PAGE, which further separates the proteins in second dimension based on their molecular weight. Here, we focus on 3 different models, whose proteome is studied with the help of gel based proteomics involving 2-DE approach;

A)Human serum

B)Escherichia coli

C)Plant



As we know, serum is a complex mixture of few highly abundant proteins and many low abundance proteins. Studies have shown that there are around 20 proteins in serum that are found in high abundance, which usually tend to mask the presence of other low abundance proteins. Albumin and immunoglobulin G (IgG) account for a very large fraction of these 20 abundant proteins. In order to obtain effective separation on 2-DE, it is ideal to remove these proteins from serum prior to electrophoresis, a process known as serum depletion.

SERUM PROTEOMICS

Removal of these high abundance proteins is usually carried out with the help of commercially available kits, which work on the basic principle of affinity chromatography. The columns present in the commercial kits are packed with affinity beads that are capable of binding specifically to certain proteins. High abundance proteins, which need to be removed, bind to the surface of the beads, while proteins of interest (proteins other than albumin and IgG) elute out of the column. Elution is carried out using a pH gradient. Serum depletion can also be performed using size exclusion criterion wherein molecular cut-off columns are used. These columns retain the proteins that are above its cut-off range but release the lower molecular weight proteins. For example, a molecular cut-off column of 60 kDa will trap albumin having a molecular weight of 66 kda. Despite several techniques available to carry out serum depletion, it must be emphasized that the process of depletion is only considered as a subsidiary technique and it is the choice of the user whether or not to carry out depletion prior to 2-DE.

The whole procedure of 2-DE can be subdivided into the following 4 steps, each of which will be described in detail.

1) Sample collection & preparation

2) First dimension - Isoelectric focusing

3) Second dimension - Separation on SDS-PAGE

4) Gel staining and data analysis

1) Sample Collection and Preparation

1. 1.A - Sample Collection

Blood is composed of different types of blood cells and plasma. The plasma acts as a suspension liquid for blood cells and other biological molecules. Plasma that is deprived of coagulation factors is referred to as serum and contains all circulatory proteins except fibrinogen and other clotting factors. The collection of blood and separation of serum for proteomics experiments is the first crucial step that influences the result and reproducibility of the experiment. Proper care during sample handling prevents the denaturing of proteins in serum. Sample collection is carried out as described below:



1. Around 5 ml of intravenous blood is collected into a commercially available blood

collection tube, which does not contain any external anti-coagulating agent. The

Vacutainer tubes are commonly used to bring about efficient and rapid withdrawal.

2. Immediately after collection, the tube is placed on ice for 30 minutes. This incubation

time is required to bring about coagulation of the blood and formation of the fibrin

clot, which is essential to separate the serum.

3. The serum is then separated by centrifuging the coagulated blood sample at 2500 rpm

at 200C for 10 minutes. The blood clot containing the different types of blood cells and

clotting factors forms the pellet while the serum forms the supernatant.

4. The dark, yellowish, viscous supernatant is then carefully aspirated out using a

micropipette and collected into a fresh, clean, labeled microcentrifuge tube. Care

should be taken not to disturb the pellet.

5. Usually, about 1.5 -2 ml of serum can be collected per 5 ml of whole blood.

6. The tube containing the serum is then labeled appropriately and stored at -800C until

further use.

1. 1.B- Sonication of proteins

Biological fluids like serum are extremely complex and have a wide variety of proteins, which can hinder the separation in 2-DE. In order to improve the quality of separation, various physical methods and chemical methods have been developed which reduce the complexity of the crude serum sample. Sonication of proteins is one of the widely used physical methods to distort complex inter- and intra- protein interactions. In this process, high frequency sound waves are applied to the crude samples which help in disrupting the proteins. Procedure for sonication of the crude serum sample is as follows:

The sample to be sonicated may be diluted using a buffer which is of appropriated pH. Procedure is as follows

1. A fresh, clean, autoclaved microcentrifuge tube is taken and 200 µL of serum is pipette

into it.

2. The serum is then diluted five times using Phosphate buffer, pH 7.4, after which the

contents are mixed thoroughly by vortexing for 30 seconds.3. The sample is then sonicated for 6 cycles of 5 second pulse with a 59 second gap in

between each cycle at 20% amplitude.



4. Since the process of sonication generates a large amount of heat, one must ensure that

the sample tube is always kept immersed in an ice bucket to avoid any denaturation or

spoiling of the sample.

Fig 8: Sonication of serum proteins on ice

In this process, acetone is used to precipitate out proteins from solution, while ethanol is used as a washing agent. Addition of ice-cold acetone to the sample brings about precipitation of proteins from solution, while the lipids and any detergents that may be present remain in the supernatant. The precipitated proteins are then harvested by centrifugation and the pellet containing desired proteins is washed with ethanol to remove any organic contaminants that may be present in the sample. (Note: Acetone to be used for precipitation must be kept for at least 5 hours at -200C). The pellet is then rehydrated using rehydration solution having the following composition:

1. 8mM Urea: Urea, being a chaotropic agent, helps in stabilization and unfolding of

proteins so that the protein’s charged surfaces are exposed towards the solution.

2. 2% w/v CHAPS {3-[(3-cholamidopropyl) dimethylammonio]-1-propanesulfonate}: This is

a zwitterionic detergent used for solubilizing proteins.

3. 40mM DTT (Dithiothreitol): DTT is a reducing agent used for reduction of disulphide

bonds that are commonly found in a protein’s complex structure. This efficiently

reduces inter and intra molecular disulphide interactions thereby exposing the ionic

surfaces of the proteins.

4. 12 µL TBP (Tributylphosphine): TBP is a reducing agent which increases solubility of the

protein. Due to its non-ionic nature, TBP does not migrate in the Immobilized pH

gradient (IPG) strip during first dimension separation and therefore helps in

maintaining reducing conditions.



5. IPG buffer: This buffer is added in accordance with the IPG strip that is to be used for

first dimension separation. More details in this context are provided in the section

discussing isoelectric focusing in detail. All these chemicals are mixed to prepare the

rehydration solution. The detailed procedure for protein precipitation using acetone is

as follows:

6. 1 ml of pre-processed serum is first divided into 5 parts, each containing 200 µL. Each

sample is then mixed with 4 times the volume of acetone (800 µL). This turns the

solution turbid, indicating the precipitation of proteins. The tubes are vortexed for

about 15 seconds to ensure proper mixing.

7. These tubes are then stored at -200C for 6 hours.

8. After this, the tubes are centrifuged at 1000g for 15 minutes at 40C. This helps in

complete precipitation of proteins and formation of pellet.

Fig 9: TCA-Acetone precipitated proteins showing the white precipitation pattern.

1. 1.C- Precipitation of Proteins using Acetone

•Carefully discard the supernatant, and add about 200 µL of ethanol for washing the pellet. Vortex the tube for 20 sec.

• Centrifuge the tube at 5000g for 10 min at 40C. This separates the pellet of protein. The supernatant is discarded and the same step is repeated once more.

•The pellet obtained is allowed to dry for about 15 min in room temperature. Since acetone is a highly volatile the excess of it evaporates leaving the pellet dry.

•Rehydrate dry pellet using 100µL rehydration solution. The tubes are vortexed to ensure proper re-suspension. If required about 20µL more of rehydration solution can be added to re-dissolve all the suspended particles.



•Finally all the re-dissolved fractions of respective samples are pooled to make a single aliquot. This can be readily used for further work or can be stored in -200C.

Albumin Apolipoprotein A-II Complement C1q CeruloplasminIgG Apolipoprotein A-I Complement C4 α2-MacroglobulinIgA Apolipoprotein B Complement C3 HaptoglobinIgM Transferrin α1-Acid Glycoprotein Prealbumin

IgD Fibrinogen α1-Antitrypsin Plasminogen

Table 1: List of 20 highly abundant proteins from Human Serum

(M. Schuchard et al, Specific Depletion of Twenty High Abundance Proteins from Human Plasma. NCI Proteomic Technologies Reagents Resource Workshop, December, 12–13, 2005)

1: D: Removal of high abundant proteins:

Commercially available serum depletion kits contain affinity chromatography based columns along with relevant buffer solutions and collection tubes. The procedure for removal of high abundance proteins involves 2 major steps - column preparation and protein binding & elution

(a) Column preparation: Commercially available kits are provided along with chromatography bead slurry, which is either given separately such that the column must be carefully packed by the user; or in the form of a packed, ready-to-use column.

1. The column needs to be activated to bring in the proper charged surfaces to facilitate

the binding of desired proteins on the surface of same. This is done by passing the

column with stipulated amount of activation buffer provided with the kit.

2. Once the column is applied with the activation buffer and fixed with collection tube it is

centrifuged with recommended time and RPM.

3. The collected wash is discarded and the procedure is repeated. Care should be taken to

ensure proper activation of the affinity beads with the buffer resulting in proper

binding of the proteins.

2. b)Protein binding and Elution: In this step, the desired proteins are bound on the

column matrix by spinning the column with the protein sample. The chemistry of the

column is such that the abundant proteins bind to the surface of beads containing

specific affinity agents that capture the proteins. The unbound proteins (proteins of



our interest), on the other hand, elute out of the column. Therefore it is very crucial to

collect the unbound proteins in a fresh, clean tube and store them under proper

conditions.

3. The column is first loaded with the recommended amount of protein sample, after

which it is incubated for specified time duration in order to facilitate proper binding.

4. The column in then centrifuged with a fresh collection tube at the outlet and the

collected flow-through is stored and labeled as flow through (F1).This fraction contains

proteins of our interest i.e serum deprived of high abundance proteins.

5. Next, the column is washed with the provided wash buffer by passing a specified

volume of buffer into the column. It is centrifuged and the flow-through (F2) collected

in a separate tube. This fraction contains loosely bound proteins, which were not

eluted out on the first attempt.

6. The fractions, F2 and F1, can then be mixed to make a whole fraction of serum

containing most of the low-abundance proteins without any masking from the high

abundance proteins such as albumin and IgG.

1: D2: Binding of proteins and Elution: In this step the desired proteins are bound on the matrix surface by spinning the column with desired protein sample. The chemistry of column is so that the abundant proteins will bind on the surface of chromatographic baits and the unbound proteins (proteins of our interest) elute out of the column. Hence it is very crucial to collect the unbound proteins in a fresh clean tube and store the contents in the proper condition with labels.

1. E: Desalting:

Serum not only contains proteins but also traces of other components of blood such as nucleic acids, lipids, detergents, salts from buffers etc. These can hinder the overall process of protein separation in the first dimension, which depends on charge of the proteins, and also in the second dimension by forming streaks in the gel image. Therefore, the sample should ideally be depleted of any other charged components in order to obtain better separation of spots.

This can also be done with the help of commercially available kits. The principle involved is similar to that of acetone or TCA precipitation of proteins. The precipitating agent in the kit is added to the protein solution, which selectively precipitates out the proteins and leaves other unwanted ionic impurities in solution. The precipitated proteins can be separated from the supernatant with the help of centrifugation. The protein pellet remaining after pipetting out the supernatant is then re-suspended in a rehydration buffer.



F: Quantitation:

Quantification of proteins in a sample is a crucial step for 2-DE since every IPG strip has an optimal protein intake capacity. It is also important to load similar protein quantities on different gels so that comparison across gels is feasible. This also helps in avoiding experimental artifacts and allows analysis of the gel in a biological context. The protein estimation method should be selected in such a way that the reagents used for the method must be compatible with other added chemicals such as detergents, reducing agents etc.

The modified Bradford method is conventionally used by many researchers due to the ready availability of reagents in the market and ease of use. The Bradford reagent contains Coomassie Brillient blue G-250 dye, which is initially brownish red in color and eventually changes to a blue color after protein binding, which can be read at 595nm. The procedure for protein quantification using the modified Bradford assay is as follows:

1. The tubes are labeled suitably as standard, blank and sample. It is advisable to carry out

standard in duplicates in order to obtain high accuracy in results.

2. Bovine Serum Albumin (BSA) can be used as the standard and can be prepared for

desired range of protein concentration in respective labeled tubes.

3. The Bradford dye reagent is then prepared as per the specified concentration.

4. The recommended amount of dye is added to tubes containing sample protein and

standards. The tubes must be properly mixed for uniformity and are then incubated for

15 minutes at room temperature.

5. The tube containing only the dye will serve as a blank for the assay.

6. The colorimeter is set at 595 nm as reading wave length.

7. The cuvette is cleaned with distilled water, and to this, the blank solution is added and

the reading noted.

8. Next, the sample protein and standard BSA solutions are taken in the cuvette

sequentially and absorbance for each is read at the same wavelength.

9. A linear graph of standard concentrations on the X-axis is then plotted against

absorbance on the Y-axis.

Concentration of the unknown sample is determined by plotting the absorbance value on the

graph and then extrapolating it to the concentration axis.



Note: The sample concentration will depend on the range of the standards used. If the sample concentration is above the limit of reading of colorimeter, then the sample must be diluted and the same assay performed. Finally, the result must be multiplied by the dilution factor to obtain the original undiluted protein concentration. If, on the other hand, the concentration is below the lowest standard, then concentration of the standard can be reduced.

2. Iso-Electric Focusing:

Isoelectric focusing constitutes the first dimension of separation in 2-D electrophoresis. This technique separates proteins on the basis of their respective isoelectric points (pI) i.e the pH at which net charge on the protein is zero. Focusing is carried out with the help of commercially available Immobilized pH Gradient (IPG) strips on which, the proteins move under the influence of an electric field until their net charge is zero, when they stop migrating.

IPG strips are solid surfaces on which dehydrated polyacrylamide is coated. The different derivatives of acrylamide having a range of pKa values are found across the gel, which enables separation of proteins in the first dimension. These commercial strips are available in a range of pH gradients such as 4-7, 3-10 etc. as well as, in a variety of lengths. The user can choose the strip based on the protein separation requirements in a particular experiment. The strips are rehydrated with the rehydration buffer prior to protein loading. They are then soaked with protein suspension of suitable concentration in the rehydration solution. The focusing is carried out on an instrument that enables gradient supply of electric current to the IPG strips. Once the proteins have been resolved in the first dimension, the strips are loaded on the SDS-PAGE gel to proceed with second dimension separation. The detailed procedure for isoelectric focusing can be subdivided into following categories:

(Angelika Görg et al, Current two-dimensional electrophoresis technology for proteomics. Proteomics 2004.)

2A. Rehydration of IPG strips:

Commercially available IPG strips contain the polyacrylamide gel in its dehydrated form. The strips therefore need to be rehydrated before they can be loaded with the protein sample. This is done by placing the strip in the rehydration tray containing the rehydration solution, overnight, for about 10-12 hours. This enables efficient absorption of proteins on the gel. The procedure for strip rehydration is as follows:

The rehydration solution containing 8M urea and 4% CHAPS/TRITON X100 is prepared. The components are mixed well and stored at -20oC.

5 µl of the required IPG buffer and 6.2 mg of DTT are added just prior to use to 1 mL of the rehydration solution and the components mixed thoroughly by vortexing.



E)

Fig: Procedure for rehydration of IPG strips a) assembly of tray and chemicals b) Addition of rehydration solution to tray c) Placement of the strip in solution d) Addition of mineral oil and maintenance at room temperature e) Schematic representation of rehydration process.

The rehydration tray is cleaned thoroughly, and the dry lanes are loaded with the rehydration buffer.

The IPG strip to be rehydrated is selected and the protective cover that is present on it is removed carefully using forceps.

Each IPG strip has two sides; a base which is made of plastic and the other side having the gel on it. The IPG strip is then carefully placed into the lane using forceps such that the surface with gel faces downwards and gets maximum contact with the solution around it.

The set-up is then left undisturbed for about 15-20min.



Mineral oil is applied on the set-up to avoid the drying of gels. It is advisable to apply mineral oil to two adjacent lanes so as to supplement uniformity of oil levels.

This re-swelling with mineral oil is carried out for 15 hours.

The tray of the IEF instrument is then cleaned and placed on an even surface after 15 hours to prepare for focusing.

The IPG strips are removed from the rehydration tray with the help of forceps and the ends are gently dabbed over an absorbent tissue to remove any excess mineral oil.

The strip is then carefully placed in the fresh tray, with the gel side facing upwards. Wicks are then placed on each end of the gel and all the wells are filled with fresh mineral oil .

Electrodes are carefully placed over the assembly and the desired programme with suitable voltage gradient and time intervals is set and the focusing is started.

The voltage gradients and time intervals are decided based on the strip being used and the sample being run. Optimization of these critical parameters is required to get best resolution in the first dimension. The whole step can be end with a holding step which works on a low voltage for stipulated amount of time.

The separation can be monitored real-time. The wick in use may have to be changed in middle of experiment if a color change to yellow is observed. This coloring of the wick takes place because of the excess salts and impurities that may be present in the sample.

Once focusing is complete, the strips are taken out of the tray and the ends gently dabbed on the surface of an absorbent tissue to remove excess oil.



G)

Fig: Process of isoelectric focusing of proteins. a) Assembly on Instrument b) Rehydrated strip is placed in the lane c) Mineral oil is poured to prevent drying of strip d) Wicks are placed on

either side e) Electrodes are placed across the strip f) The IEF lid is closed and focusing started g) Schematic representation of IEF and separation on first dimension.

2. B Equilibration of strips : The isoelectrically focused strips must be equilibrated before they are separated in their second dimension. This step ensures proper denaturation of proteins, which in turn enables efficient separation based on molecular weight during SDS-PAGE. Dithiothreitol (DTT) is used for reduction of inter and intra-chain disulfide bonds in proteins. Iodoacetamide, (IAA) being a potent alkylating agent, prevents the reformation of these broken disulfide bonds by alkylating the sulphur atoms.

The equilibration solution consisting of 6 M urea, 75 mM tris HCl buffer, 29.3% glycerol, 2% SDS and 0.02% bromophenol blue, made up to 20 mL using distilled water, is prepared. The contents are thoroughly mixed and can be stored at -20oC for later use.

Just before use, 10 mg of DTT is added to 10 mL of the equilibration solution to obtain solution X and 25 mg of iodoacetamide is added to the remaining 10 mL of the equilibration solution to prepare solution Y.

First, solution X is taken in a container and the strip is placed in it for around 10 minutes.

Next, the strip is transferred to solution Y where it remains for another 10 minutes, thereby completing the process of focusing. The strip is then ready for separation in the second dimension.



Fig: Equilibration of proteins on IPG strips before transfer to second dimension.

Sodium Dodecyl Sulphate (SDS)-PAGE

SDS-PAGE, which separates proteins on the basis of their molecular weight, constitutes the second dimension of 2-DE after isoelectric focusing. The gel to be used must be cast and kept ready at least half an hour before it is required so that there is sufficient time for proper gel polymerization. The concentration and size of the gel vary based on the sample being separated and the dimensions of the IPG strip used for first dimension, respectively. The commonly used and preferable gel concentration is 12.5%. The IPG strip containing the separated proteins, being very sleek and fragile, must be placed carefully and firmly in contact with the SDS-PAGE gel, such that there are no air gaps between the two surfaces that could allow the protein to escape. To facilitate this process, an agarose solution is often used as a fixing agent, which helps in removing any air bubbles that may be formed. Bromophenol blue (BPB), which is added to the agarose solution, acts as a tracking dye for the experiment to view the electrophoretic migration at any given time. Once the proteins have been separated in the second dimension, the SDS-PAGE gel is stained using a suitable staining solution in order to view the protein bands. The detailed protocol is as follows:

First, the appropriately sized gel casting apparatus is assembled together.

Next, the following components are prepared and mixed together for casting the gel (indicated contents are optimum for casting an 8cm gel):

Acrylamide bis-Acrylamide solution (29:1 ratio) -3.14 mL

Distilled water-3.685 mL

Tris-HCl (pH 8.8)-2.435 mL



SDS (10%)-93.75 µL

APS (10%)-93.75 µL

TEMED-3.75 µL

A 10X stock solution of the buffers required for the gel tanks are prepared by mixing the following:

Tris base (FW 121.1) 250 mM—30.3 g

Glycine (FW 75.07) 192 mM—144.1 g

SDS (FW 288.38) 0.1% (w/v)--10 g

Double-distilled water to--1 L

This 10X stock solution can be diluted to 1X strength before use (100 ml made up to 1000ml).

Fig 14: a) Gel casting assembly b) Casting of gel c) Pouring agarose on top of gel with strip d) Setting up for separation by pouring tank buffer e) Typical experimental set-up for SDS-PAGE.

1. Acrylamide Bis-Acrylamide solution together form polymer and form gel. SDS helps for

providing uniform negative charge on the protein surface which helps for movement

proteins in PAGE depending on molecular weight regardless of charge. APS along with

TEMED will form free radicals which help in polymerization of gels. Note that the

TEMED can be added just before pouring the gel.

2. Add all the contents mentioned above. Mix it thoroughly.



3. Slowly pour the gel through the sides of the plates by touching the sides to avoid the

formation of air bubble.

4. Keep in mind that a small space has to be left for placing the IPG strip and pouring

agarose solution on the top.

5. Once the gel is poured immediately pour a small quantity of water with 0.1% SDS on

top. This maintains an uniform surface for placing the IPG strip.

6. Gel takes approximately35-40 min to polymerize.

7. Mean while prepare 0.75% agarose solution with 0.02% of BPB in water. Mix the

content and warm to dissolve the content. After dissolution cool it a bit so that it is

optimum to pour.

8. Decant the water on gel carefully remove the remaining water with the help of

absorbent tissue paper.

9. Carefully take equilibrated IPG strip, place on the gel so that the strip ensure proper

contact of strip with gel

10. Carefully pour the agarose solution till the top mark and allowed to solidify.

11. Place the set-up inside tank of electrophoresis and pour the tank buffer into inner and

outer tank.

12. Apply constant voltage of 90 volts. The procedure approximately takes 2 hours.

13. The electric supply can be stopped when the dye front reaches the end of the gel or

slightly moves out of the gel.

4. A: Staining and destaining

Once the proteins are separated on SDS PAGE, to visualize the separation patterns, proteins must be stained. Stains are chemical components which specifically bind to proteins enabling the visualization. The property of dye should be such that it helps visualization of maximum number of protein spots on gel.

Coomassie brilliant blue is most widely used stain. There are some more varieties such as silver staining, sypro ruby etc. Depending up on sensitivity and cost effectiveness the selection can be done. Also some of the dyes are not compatible when the spots are to be identified for proteins with mass spectrometry. So a dye must be selected by user which serves the best of its purpose.

Image of stained gel can be taken using gel documentation instrument which will typically have a chamber and a ccd camera for capturing image. Comparison of the global expression profiling



or differential expression of proteins within or across the gel can be done with the help of commercially available software.

Staining procedures are as follows; Preparation of coomassie dye;

Methanol, acetic acid and distilled water are mixed in 40:7:53 ratio. To same 350mg Coomassie Blue is added. The solution is mixed well and can be filtered using a filter paper to get rid of un-dissolved debris. Though the stain is mostly used once, it can be used thrice efficiently. The gel after removing and before staining has to be washed with distilled water. This helps the removal of SDS which may hinder the proper staining of the gel by forming streaks and unwanted spots all over the gels.

1. Carefully remove the gel out of gel plates.

2. Place the gel into container with distilled water; wash the gels briefly at least thrice by

changing water. Then discard the water used.

3. Now immerse the gel with stain, cover the container, and keep it on gel rocker with a

constant shacking for about 6-8 hr.

4. The stain covers the whole gel including the parts without proteins. Hence the gels have

to be distained so that only the part with proteins will retain the stain and rest of the

background is clear.

5. Take the gel out of stain and immerse in distaining solution (40:7:53 methanol, acetic

acid, water) for about 5-6 hours. Change the stain about three times for efficient

destining.

6. The stained gel can be washed with distilled water twice before imaging.

Fig 15: Image of a typical 2-DE gel showing serum proteins separated on a 4-7 pH range on the X axis and molecular weight on the Y axis



4. B: Imaging and analysis

1. The gel is placed carefully on the imaging platform. Care should be taken to ensure that

the gel does not break while transferring.

2. The gel image is captured and saved with appropriate label.

Image analysis is performed for checking the differential expression between two different samples with the help of commercially available software packages. These analysis softwares have tools which allow gels to be cropped, overlayed, zoomed into, edited etc. as well as statistical and data output tools. Although there are several tools which need to be explored in detail with the software, the basic steps for image analysis is as follows:

• A suitable labeled master folder is first created which should contain the

gels to be analyzed.

• The gel images to be analyzed are opened on the software and labeled

appropriately.

Fig 16: Representation of analysis through commercially available software.

1. The region on the gel having maximum spot density is selected and the gel is cropped

with the cropping tool so that those regions without any spots do not interfere in

analysis.



2. The spots on the gel can be selected with the help of the spot picking tool on the

software. These spots are selected by the software on the basis of certain user-defined

criteria, which gets translated by the software into pixel intensities of the spots.

3. The gel can be zoomed into and carefully analyzed to detect spots that may have been

selected based on the defined criteria but are not actually proteins. This step is very

important because the selected spots will be considered for comparison across the

gels. Therefore great care must be taken not to select a wrong spot and more

importantly, not to miss an important protein spot.

4. The selected spots are then matched across gels. This process displays the number of

spots that are matched and unmatched across the gels.

5. The extent of matching can also be seen with a 3-D graphical representation of the

spots.

6. The spot parameters such as volume, intensity, possible pI, molecular weight etc. can be

obtained through the software.

7. Statistical parameters of spot matching across different gels such as coefficient of

variation, standard deviation, t-test values etc. can also be obtained.

8. All this data can be compiled together to understand the differential expression of

proteins across the desired gels.

BACTERIAL PROTEOMICS

Escherichia coli is one of the most widely used model organisms in the world, which has paved the way for a new era of biotechnology and related sciences. E. coli is a prokaryotic, unicellular microorganism that can easily be grown, harvested and manipulated in order to understand various biological processes. Many landmark discoveries in biology have been made using E. coli as the model and the same theory has been successfully extrapolated to higher organisms.

The proteome refers to the entire protein complement expressed by the genome of an organism at a given point of time under a defined set of conditions. Since proteins are the ultimate effector molecules in all organisms, study of E. coli proteins could provide important insights into various biological functions. There are several methods to study the numerous proteins being expressed by E. coli, many of which are laborious and time consuming. The gel-based proteomics approach, which has increasingly gained popularity to study large number of proteins simultaneously, will be discussed in detail in the following sections.

Culturing of E. coli



1. All glassware and media to be used for growing the bacteria are first autoclaved at 15 lb

pressure for 15 minutes at 1210C.

2. 2% Luria broth is prepared in a clean autoclaved container.

3. The broth is then inoculated with around 100 ul of fresh culture.

4. The culture is maintained at 370C, with constant agitation in an incubator for about 6-8

hours.

5. Proper growth of the microbe can be ensured by measuring the optical density of the

culture solution at regular time intervals. The O.D value should ideally be between 0.8

and 1.0.

6. Once the culture reaches the desired O.D, the organism’s growth is stopped by

centrifuging the culture in tubes at 5000 g for 15 minutes at 40C.

Fig 17 : a) Picking organism colonies from the mother culture b) Inoculation of the culture broth c) Monitoring growth of E. coli by turbidity measurements d) Precipitation of bacterial pellet

using centrifugation.

Extraction of proteins from E. coli

Unlike serum, the entire E. coli organism is considered for studying its proteome. Hence methods are required to lyse the cells and release its cytoplasmic contents, including the various proteins to be studied. Cell lysis can be carried out at the lab scale with the help of chemical and physical methods, of which sonication is one of the most widely used and accepted techniques. The procedure for sonication and subsequent protein extraction is explained below.



♠The cells must first be harvested out of the broth, which is done by centrifuging the culture at 13,000g for 15 minutes. This causes the cells to sediment out in the form of a pellet.

♠The pellet containing the E. coli cells is washed 3 times with 1 mL of phosphate buffer, pH 7.4 (20mM Na-phosphate, 0.15M NaCl).

♠The washed pellet is then re-suspended in a re-suspension solution, pH 7.4 (20mM Na-phosphate, 0.15M NaCl, and 5mM MgCl2). To this, 10 uL/mL of protease inhibitor is added, which ensures that there is not degradation of the proteins when they are released along with the protease enzymes.

♠This mixture is sonicated, which breaks open the cells in the pellet, thereby releasing all the cytoplasmic contents. Sonication is performed 3 times for 30 cycles at 40% amplitude.

♠Following sonication, the supernatant is collected since the proteins are now present in their soluble form.

Trizol (contains phenol and guanidine isothiocyanate) method helps for simultaneous extraction of RNA, DNA and proteins. After addition of Trizol all 3 components separates into 3 distinct layers. Protein is extracted at the end by the addition of acetone (for precipitation) to the lower most portion.

• Add 1 ml trizol reagent to the bacterial suspension.

• Immediately add 200ul chloroform to the same mixture, shake vigorously for 15 sec and incubate for 15 min at RT.

• Centrifuge at 12,000g for 15 min.

• Carefully remove upper layer containing RNA using a micropipette.

• To the bottom layer, add 300 ul ethanol, centrifuge at 5,000g for 5 min to remove DNA.

• Separate the supernatant containing protein collect into a fresh tube. Retain the pellet of DNA.

• To the resultant supernatant, add 4 volumes of chilled acetone (acetone kept in – 200C at least for 4 hours) and incubate for 6 hrs at – 200C.

• After incubation centrifuge at 12,000g for 5 min.

• Discard the supernatant, retain the pellet of protein.



• Wash the protein pellet with 95% (95% ethanol+5% water) ethanol or acetone (4 times).

• After wash give a brief spin to settle proteins. Every time through the supernatant.

• Dry the pellet in the room temperature.

• Reconstitute the dried pellet in lysis buffer.

The remaining steps for the 2-DE procedure involving quantification, desalting, first dimension and second dimension separation are the same as those described in the serum proteomics section.

Fig 18: Image of a typical gel showing bacterial proteins separated on a 4-7 pH range over X axis and Molecular weight on Y axis.

(*Michael Hecker et al,Gel-based proteomics of Gram-positive bacteria: A powerful tool to address physiological questions. Proteomics 2008

* Nelson C Soares et al 2-DE analysis indicates that Acinetobacter baumannii displays a robust and versatile metabolism. Proteome Science 2009)

PLANT PROTEOMICS

Plants are one of the nearest relatives to higher organisms such as mammals in the phylogenetic tree and have appreciable amount of complexity with respect to proteins in their system. Studying the proteome of plants could help in providing a comparative analysis with closely related proteomes of other organisms. Some plants such as Arabidopsis thaliana, which is commonly used as a model organism, are often studied at the proteome level to gain deeper insights into various biological processes. In this section, we describe the study of a plant leaf proteome for experimental purpose using 2-DE. Experimental plan involves the collection of plant leaf, protein extraction followed by separation of proteins on first and second dimension.



Preparation of sample:

1. Around 300 mg of fresh leaf is first collected.

2. The leaf must be homogenized so as to extract proteins, which is done using a clean

mortar and pestle.

3. Liquid nitrogen is then added to the homogenized leaf, which causes it to freeze and

solidify rapidly. This solidified leaf is then ground thoroughly until a powder is

obtained.

4. To the powder, 500 µL of lysis buffer (acetone, 10% TCA, 0.07% DTT) is added and the

contents ground to prepare a homogenous mixture. More lysis buffer can be added, if

required, to make the process efficient. Once it is finely ground, the volume is made up

to 1.5 mL and this mixture is left as such at -200C for 1 hour.

5. It is then centrifuged at 14,000 rpm for 30 minutes at 40C. This enables formation of a

pellet of proteins. The supernatant can be discarded.

6. Chilled acetone with 0.07% DTT is then added to the pellet and vortexed.

7. The mixture is again centrifuged at 14,000 rpm for 30 minutes at 40C, after which the

supernatant is discarded. This step is repeated 3 more times.

Fig 19: a) Fresh leaves b) Leaves being ground with liquid nitrogen c) Fine homogenous paste prepared using buffer d) Debris precipitated using centrifugation



1. After washing, the pellet is left to dry at room temperature for about 40 minutes.

Ensure that the pellet is totally dry.

2. The dry pellet is then dissolved in 400 µL of rehydration buffer (composition same as

mentioned in section 1: C)and the contents vortexed. This is then stored overnight at

40C.

3. The mixture is centrifuged the next day at 14,000 rpm for 15 minutes at 4°C.

4. The supernatant containing the proteins are separated carefully and stored in a fresh

microcentrifuge tube at -200C until further use.

5. These proteins in the supernatant can then be quantified and separated in two

dimensions as described for serum proteomics.

Fig 20: Image of a typical gel showing plant leaf proteins separated on a 4-7 pH range over X axis and Molecular weight on Y axis.

(Walter Weiss and Angelika Görg, Two-Dimensional Electrophoresis for Plant Proteomics. Methods in Molecular Biology, 2007)

(Wei Wang et al, Protein extraction for two-dimensional electrophoresis from olive leaf, a plant tissue containing high levels of interfering compounds. Electrophoresis, 2003)


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