Addition Of Primary Antibody

Published Date: 02 Nov 2017

Dulbecco’s Modified Eagles Medium (DMEM) was procured from Invitrogen, Fetal Bovine Serum (FBS) was purchased from Lonza, 5-FU was obtained from Calbiochem, Curcumin and MTT and antibodies were purchased from Santa cruz Biotechnology.

Cell lines MDAMB, SKBR3, T47D, and MCF7 were obtained from National Centre for Cell Science, Pune, India.

EQUIPMENTS:

CELL CULTURING:

T25 AND T75 Culture Flask

Micropipettes (Eppendorf)

Centrifuge tubes

Cryovials (Tarsons)

Centrifuge

CO2 incubator (Sanyo)

Inverted microscope (Nikon)

-20 Deep Freezer (Evolution)

-80 Deep Freezer (Thermo Scientific)

Liquid Nitrogen container (ARPEGE 170)

MTT ASSAY

96 well Microtitre Plates (BD Falcon)

60 mm Culture Plates (BD Falcon)

ELISA Plate Reader (BIORED)

Western Blotting

Western Blotting apparatus (Bio- Red mini PROTEANIII)

METHODS

Maintenance of the cells

4.1 Revival of the cells:

Cells were collected from the -196ºC cryopreservation and thawed in 37°C water bath. Pre- warmed media was added and cells were centrifuged at 2,000 rpm for a period of 2mins. After centrifugation, supernatant was discarded and to the pellet fresh media with 20% FBS was added. The breast cancer cell lines were cultured in Dulbecco’s Modified Eagles Medium (Life Technologies) supplemented with 10% fetal bovine serum (sigma) along with 100U/ml penicillin, 50µg/ml streptomycin. The cell lines were maintained at 37oC in a humidified atmosphere of 5% CO2.

4.2 Sub Culturing of cells:

For sub culturing, the used up media was removed, and the monolayer was washed with 1X PBS-EDTA and trypsinized with 0.25% Trypsin-EDTA for 5 min at 37°C. Fresh medium (with serum) was again added to inactivate trypsin followed by centrifugation. The pellet was dispensed gently in complete media. After diluting, the cells were seeded into fresh tissue culture flasks for growth. For the experiments, cells were counted using heamocytometer and a required number of cells were seeded.

4.3 Long term storage of cells:

For future use, the cells were stored in liquid nitrogen. For this, after trypsinization, the cells were transferred to freezing vials. The cells were pelleted by spinning the vials at 2000rpm for 2 minutes. After discarding the supernatant, the pellet was resuspended in freezing mixture (1:9 ratio of DMSO to Serum) and mixed well. The vials were transferred first to -20oC freezer for overnight incubation followed by the same in -80°C. The vials were then transferred to -1960C in liquid nitrogen for long term storage.

4.4 Preparation of the Drugs.

Stock solution of curcumin and 5-FU was made in Dimethyl Sulphoxide (DMSO), the concentration of which in media should not exceed 0.2%. 25µM, 10µM of 5-FU were taken as a working solution and 10µM of curcumin was considered as a working solution.

4.5 Drug Treatment

Drug treatment was performed after 24 hr of seeding. It was divided into two stages:

Pretreatment: The cells undergoing combination studies were pre-treated with 10µM of curcumin and incubated for 6hrs.

Post-treatment: After 6hrs of incubation, all cells including those undergoing combination studies were treated with 10µM curcumin, 25µM and 10µM 5-FU and combination of both. Incubation time was determined based on the experiment.

4.6 MTT Assay

MTT assay is a colorimetric assay for measuring cell viability, cellular proliferation and activation. It is also used to determine the cytotoxicity of potential medical agents and other toxic materials. MTT was first described by Mosmann in 1983.

Principle:

MTT (3-(4,5-dimethylthiazol-2-yl)-2,5diphenyltetrazolium bromide(yellow), is reduced to formazan, a purple colored product in the mitochondria of living cells. A solubilization solution is added to dissolve the insoluble purple formazan product into a colored solution. The absorbance of this colored solution can be quantified by measuring at a certain wavelength (usually between 500 and 600 nm) by a spectrophotometer. The reduction of MTT takes place only when mitochondrial reductase enzyme is active and therefore conversion is directly related to number of viable cells. When the amount of purple formazan produced by cells treated with an agent is compared with formazan produced by untreated control cells, effectiveness of the agent in causing death of cells can be deduced.

Protocol:

For the MTT assay, 5000 cells were seeded per well in a 96 well plate and incubated for 24 hr at 37°C.

The cells were treated with different concentration of drugs

Six wells were kept as control i.e., cells free from treatment to determine cell survival and the percentage of live cells after culture

The cells were then incubated for 72 hr at 37°C

Then the medium was removed and after a PBS wash, fresh medium was added along with 100µl of 20% MTT (5mg/ml) to each well

The cells were incubated for 2 hr at 37°C inside CO2 incubator. The yellowish colored MTT was reduced to dark blue colored formazan by the viable cells only

The crystals were solubilized with 0.1 ml lysis buffer (20% SDS in 50% Dimethylformamide)

The plates were kept for 4 hr incubation at 37ºC. The optical densities at 570 nm were measured using ELISA plate reader (Bio-Rad)

The relative cell viability in percentage was calculated by comparing the viability of the treated cells with of the control and the cell survival was expressed as:

Cell survival (%) = (Mean OD (drug exposed cells)/ Mean OD (control)) X 100

Graph was plotted by taking percentage viability on Y axis and concentration of drug on the X axis.

FLOW CYTOMETRY:

Flow cytometry is a technique based on quantitative single cell analysis. It was introduced in the 70’s and soon became an important instrument for biological sciences. The technique for analysis of individual cells in fluidic channel was first illustrated by Wallace Coulter in 1950’s. It was applied only to automated blood cell counting. Two decades later, development in various fields like laser technology, computer science, monoclonal antibody production, cytochemistry, fluorochrome chemistry etc led to the development of the flow cytometry.

Principle:

Flow cytometric analysis requires particle or single cell suspension. These proteins of interest are labeled with various immunoflurescent dyes. The single cell suspension is passed through a flow cell, surrounded by a very narrow fluid stream, where they pass through a hydrodynamically- focused laser beam one at a time. Many detectors are aimed at a single point, one parallel with the light beam i.e. FSC (Forward Scatter) and others perpendicular SSC (Side Scatter). The light on striking the cell is either scattered or absorbed. If the cell possesses a naturally fluorescent substance then the absorbed light of specific wavelength may get re-emitted as fluorescence. Light scatter mainly depends on the internal structure, size and shape of the cell. Forward Scatter tells about the cell volume while Side Scatter about the inner complexity. The fluorescent substances absorb light of specific wavelength and emit back light of different wavelength. Commonly used fluorescence dyes are Phycoerythrin, FITC and Texas red. Series of photodiodes detects the light signals and amplifies it. Optical filters block the unwanted light and permit only the light of desired wavelengths which reaches the photodetectors. These electrical pulses are digitized, the data is stored and analysed thus finally displayed through computer system.

Applications:

The chief clinical application of flow cytometry is the diagnosis of hematologic malignancy, although many other applications exist like reticulocyte enumeration and analysis of DNA ploidy, Cell cycle and Cell death, Cell function, Platelet function. It has application in the measurement of the efficacy of Cancer chemotherapy, transfusion medicine, Organ transplantation and Hematopoietic Cell therapy.

Protocol:

0.5 x 106 cells/ml were seeded in 60mm plates

Cells were treated with different concentration of drug 5-FU (10µM) alone, curcumin (10µM) and combination of 5-FU (10µM) and curcumin (10µM) followed by 24 hr incubation.

The medium was transferred into labeled eppendorf tubes (kept in ice)

Cell were trypsinized, pelletized and suspended in 700µl of 70% ice-cold ethanol followed by 30 min of incubation in cold.

Cells were pelletized and PBS serum was added into the obtained pellet

5µl of RNAse was added into the cells followed by incubation at 37°C for 30 minutes

10µl of 1mg/ml of PI was added into the solution

Cell cycle was analyzed using flowcytometer.

WESTERN BLOTTING (IMMUNOBLOTTING)

Western blotting (also called immunoblotting because an antibody is used to specifically detect its antigen) was introduced by Towbin, et al. in 1979 and is now a routine technique for protein analysis. The specificity of the antibody-antigen interaction enables a single protein to be identified in the midst of a complex protein mixture. Western blotting is commonly used to positively identify a specific protein in a complex mixture and to obtain qualitative and semi quantitative data about the protein.

PROTOCOL

Whole cell lysate preparation

0.7 X 106 cells were seeded in 60 mm plates are incubated. After treatment with 5-FU and curcumin, as needed for each experiment, the medium was removed and scrapped using 1X PBS. Centrifuged for 2 minutes at 13,000 rpm at 40C and to the obtained pellet, WCL solution containing protease inhibitors was added. Mixed well and kept in ice for 30 minutes with intermittent quick vortexing after every 5minutes. Spunned down at 13,000 rpm for 10 minutes and obtained the supernatant. The supernatant was transferred to a fresh tube. To this 5X loading dye was added and boiled for 5 minutes and stored at -700c. The tube with the pellet should be kept for protein estimation.

Protein estimation

Bradford method was used for protein estimation. It is a calorimetric method which depends on quantitating the binding of a dye, Coomassie Brilliant Blue to an unknown protein and comparing the binding to that of different amounts of a standard protein like BSA. The assay is based on the observation and the absorbance maximum for an acidic solution of Coomassie Brilliant Blue G-250 shifts from 465 nm to 965 nm when binding to protein occurs. Both hydrophobic and ionic interactions stabilize the anionic form of the dye, causing a visible color change. It’s a designed to qualify 1-10 µg protein. It is provide equal loading of the sample protein for gel electrophoresis 2.5ul of the protein sample was mixed with 122.5µl of distilled water and 50 µl from this solution was transferred to corresponding two wells of the microtitre plate. Then 200µl of Bradford’s reagent was added and the reading was taken at 570 nm. 80µg/well of protein sample was loaded on a 10% gel.

SDS-PAGE

The proteins of the sample were separated using SDS-PAGE (Sodium dodecyl sulphate polyacrylamide gel electrophoresis). Polyacrylamide gels are formed by the polymerization of monomeric acrylamide to polymeric polyacrylamide chain by cross linking of the chain by N, N’-methylene bis acylamide. The reaction is initiated by Ammonium persulphate (APS) and the reaction is accelerated by TEMED, which catalyses formation of free radicals from ammonium persulphate. The mean diameter of pores formed in these networks is determined by the concentration of acylamide monomer and the bifunctional cross linker.

One dimensional gel electrophoresis under denaturing condition (i.e, in the presence of 0.1% SDS) separates protein based on their molecular size. Polyacrylamide gel is cast as a separating gel (resolving/running gel) topped by stacking gel. The most widely used method for discontinuous gel electrophoresis is the system described by Lammili. A discontinuous buffer system uses buffers of different pH and composition to generate a discontinuous voltage gradient in the gel. In the discontinuous system the sample first passes through stacking gel which has larger pore size. The stacking gel buffer contains chloride ions (called the leading ion) whose electrophoretic mobility is less than the mobility of protein in the sample. The electrophoresis buffer contains glycine ion (called the training ion) whose electrophoretic mobility is less than the protein in sample. Upon entering the stacking gel the glycinate ion in the running buffer encounters a condition of low pH which shifts equilibrium towards the formation of zwitter ions which causes lesser electrophoretic mobility. The net result is that the faster migrating ions leave a zone of conductivity between themselves and the migrating protein. The higher voltage gradient in this zone allows the protein to move faster and to stack in the zone between the leading ion and trailing ions. After leaving the stacking gel, the protein enters the separating gel. The separating gel has a smaller pore size, high salt concentration and higher pH. In the separating gel the glycine ion migrate past the protein and thus the proteins are separated according to their molecular size.

SDS-PAGE separates proteins based on their molecular weight. Sample proteins become covered in the negatively charged SDS and move towards the anode through the acrylamide mesh of the gel. Smaller proteins migrated faster through this mesh and the proteins are thus separated according to their size (in KDa). From the lysate, required amount of protein sample was loaded on the required percentage SDS-PAGE gel. A voltage of 40 mille ampere was applied along the gel and the protein migrated at different speeds. The different rates of advancement or different mobilities separated proteins into bands within each lane formed under the wells. One lane was reserved for protein marker, which is a mixture of proteins having defined molecular weight, typically stained so as to form visible bands.

Following electrophoresis, the protein must be transferred from the electrophoresis gel on to the membrane. There are a variety of methods including diffusion transfer, capillary transfer, heat accelerated convectional transfer, vacuum blotting and electroelution.

The transfer method, are used most commonly for proteins in electroelution or electrophoretic transfer, because of its speed and transfer efficiency. The method uses the electrophoretic mobility of proteins to transfer them from one gel to the matrix.

Electrophoretic transfer of protein involves placing a protein containing polyacrylamide gel in direct contact with a piece of nitrocellulose or other suitable, protein binding support and sandwiching this between two electrodes submerged in a conducting solution. When an electric field is applied, the proteins move out of the polyacrylamide gel and onto the surface of the membrane, where the protein becomes tightly attached. The resulting membrane is a copy of the protein pattern that was found in polyacrylamide gel itself.

Transfer efficiency can vary dramatically among proteins, based upon the ability of a protein to migrate out of the gel and its property to bind on to the membrane under a particular set of conditions. The efficiency of transfer depends on factor such as the composition of the gel, complete contact of the gel with the membrane, the position of the electrodes, the transfer time, size and composition of proteins, field strength and the presence of detergents. Optical transfer of proteins is generally obtained in low ionic strength buffers and with low electrical current.

The resolve bands in the SDS PAGE were electro blotted on to a PVDF membrane as described by Towbin. In order to make the proteins accessible to antibody detection, they were moved from the gel onto a membrane made of nitrocellulose or PVDF. PVDF membrane was pre treated using methanol for 10 seconds followed by 5 minutes treatment with water, then by Towbin,s buffer. After electrophoresis, both the gel and PVDF membrane of same dimension were equilibrated using Towbin’s buffer. The protein bands were transferred on to the PVDF membrane using Bio-Red Mini PROTEAN III wet blot apparatus at 40V overnight in cold room. The method for transferring the protein is called electro blotting. Electrophoretic transfer of proteins involves placing a protein containing polyacrylamide gel in direct contact with a piece of PVDF and sandwiching between two electrodes submerged in a conducting solution. When an electric field is applied, the proteins moved out of the polyacrylamide gel and onto the surface of the membrane, where the protein becomes tightly attached. After the transfer process, the membrane was stained with Ponceau stain to find its transfer has taken place completely.

Blocking

Since the membrane has high affinity for protein and both antibodies and the target protein, steps had to taken to prevent interaction between the antibodies used for the detection and the membrane. Blocking non specific binding was achieved by placing the membrane in a dilute solution of protein typically BSA or non fat dry milk for 1 hours, with a minute percentage of detergent such as Tween 20. The protein in the dilute solution attaches to the membrane where the sample protein had not attached during the process of transfer. Thus when the antibody was added, there was no place on the membrane for the antibody to attach other than on the binding sites of the specific target protein. This reduced noise in the final product of the western blot, leading to clearer results, and eliminated false positive.

Washing

Washing is usually done with large volume of TBST buffer.

Addition of Primary Antibody

After blocking, a dilute solution of primary antibody against caspase 8, caspase 9, β actin (generally between 0.5 and 5 mg/ml) the concentration varies from 1:2000 to 1:1000) was added and incubated with the membrane under gentle agitation. Typically the solution was comprised of Tris buffered saline (TBS) solution with a small percentage of detergent like Tween 20 and sometimes with powered or BSA. The antibody solution and the membrane could be sealed and incubated together for overnight in cold room in order to protect the antibodies. When antibody is incubated is prepared in BSA the lifetime of the antibody is increased and hence primary antibody is usually prepared in BSA.

Secondary Antibody

After removing the primary antibody, membrane was washed thrice with 5 minutes each TBST to remove unbound primary antibody and then was exposed to another antibody, directed at specific portion of the primary antibody. This was known as secondary antibody. The secondary antibody was usually linked to biotin or to a reporter enzyme such as Alkaline Phosphates or Horse Raddish Peroxidase. It is usually made in 5% fat milk solution. The membrane was incubated with the secondary antibody for three hours with gentle agitation in cold room or incubated at 1 hr for room temperature.

Detection by Chemiluminescence

The unbound secondary antibody was washed off with TBST and detected by Enhanced Chemiluminescence method (Amersham ECL Plus kit). Chemiluminescent detection methods depend on the incubation of the western blot with a substrate that will luminescence when exposed to the reporter on the secondary antibody. ECL solution A and B mixed in a ratio of 1:1. After mixing the solution was poured over the dried blot. After this blot was wrapped with saran wrap. Then the wrapped blot was placed in a X-ray film cassettes with the protein side facing up. An X-ray film was placed over the blot and cassettes was closed to that the film does not get exposed to light. The luminescence observed was transferred to X-ray film in the dark and the film was developed and visualized. ECL detection is considered among the most sensitive methods for blotting analysis.